Vanadium and proteins: Uptake, transport, structure, activity and function

Coordination Chemistry Reviews 301–302 (2015) 49–86

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Coordination Chemistry Reviews journal homepage: www.elsevier.com/locate/ccr

Review

Vanadium and proteins: Uptake, transport, structure, activity and function João Costa Pessoa a,∗ , Eugenio Garribba b , Marino F.A. Santos c , Teresa Santos-Silva c a

Centro de Química Estrutural, Instituto Superior Técnico, Universidade de Lisboa, Av. Rovisco Pais, 1049-001 Lisboa, Portugal Dipartimento di Chimica e Farmacia and Centro Interdisciplinare per lo Sviluppo della Ricerca Biotecnologica e per lo Studio della Biodiversità della Sardegna, Università di Sassari, Via Vienna 2, I-07100 Sassari, Italy c UCIBIO, REQUIMTE, Departamento de Química, Faculdade de Ciências e Tecnologia, Universidade Nova de Lisboa, 2829-516 Caparica, Portugal b

Contents 1. 2.

3.

4.

5.

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.1. Vanadate(V)-phosphate analogy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Uptake and transport . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1. Vanabins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2. Vanadium in polychaetes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3. Storage and blood proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3.1. Transferrins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3.2. Albumin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3.3. Immunoglobulins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3.4. Hemoglobin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3.5. Ferritins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Vanadium for protein activity and function . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1. Nitrogenases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2. Haloperoxidases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.3. Mechanism of halide oxidation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.4. Nitrate reductases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Vanadium as substrate analogue or inhibitor . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.1. Phosphatases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2. Transferases and kinases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.3. EctoNTPDases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.4. Phosphodiesterases and phosphomutases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.4.1. Phosphodiesterases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.4.2. Phosphoglycerate mutases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.5. ATPases (myosins and transporters) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.5.1. ATPases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.5.2. Myosins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.5.3. Transporters . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.6. ATP synthases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.7. DNA binding proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.7.1. Topoisomerases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.7.2. Other DNA binding proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.8. RNA binding proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.8.1. Ribonucleases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.8.2. Ribozymes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.8.3. Helicases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Miscellaneous . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.1. Phosphonoacetate hydrolase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.2. Chymotrypsin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.3. PhoX . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.4. Lysozyme . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

∗ Corresponding author. Tel.: +351 218419268. E-mail address: [email protected] (J. Costa Pessoa). http://dx.doi.org/10.1016/j.ccr.2015.03.016 0010-8545/© 2015 Elsevier B.V. All rights reserved.

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6. 7.

J. Costa Pessoa et al. / Coordination Chemistry Reviews 301–302 (2015) 49–86

Vanadium oligomers and proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Final remarks . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

a r t i c l e

i n f o

Article history: Received 10 November 2014 Received in revised form 6 March 2015 Accepted 11 March 2015 Available online 25 March 2015 Keywords: Vanadium Vanadates Coordination geometry Vanadium-Protein structure Enzyme inhibition Transition state analogue

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a b s t r a c t Vanadium is an element ubiquitously present in our planet’s crust and thus there are several organisms that use vanadium for activity or function of proteins. Examples are the vanadium-dependent haloperoxidases and the vanadium-containing nitrogenases. Some organisms that use vanadium have extremely efficient and selective protein-dependent systems for uptake and transport of vanadium and are able to accumulate high levels of vanadium from seawater, vanabins being a unique family of vanadium binding proteins found in ascidians involved in this process. For all of the systems a discussion regarding the role of the V-containing proteins is provided, mostly centered on structural aspects of the vanadium site and, when possible or relevant, relating this to the mechanisms operating. Phosphate is very important in biological systems and is involved in an extensive number of biological recognition and bio-catalytic systems. Vanadate(V) is able to inhibit many of the enzymes involved in these processes, such as ATPases, phosphatases, ribonucleases, phosphodiesterases, phosphoglucomutase and glucose-6phosphatase, and it appears clear that this is closely related to the analogous physicochemical properties of vanadate and phosphate. The ability of vanadium to interfere with the metabolic processes involving Ca2+ and Mg2+ , connected with its versatility to undergo changes in coordination geometry, allow V to influence the function of a large variety of phosphate-metabolizing enzymes and vanadate(V) salts and compounds have been frequently used either as inhibitors of these enzymes, or as probes to study the mechanisms of their reactions and catalytic cycle. In this review we give an overview of the many examples so far reported, also disclosing that vanadate(IV) may also have an equally efficient inhibiting effect. The prospective application of vanadium compounds as therapeutics has also been an important topic of research. How vanadium may be transported in blood and up-taken by cells are particularly relevant issues, this being mainly dependent on transferrin (and albumin) present in blood plasma. The thousands of studies reported on the effects of vanadium compounds reflect the complexity of the interactions occurring. Although it is not easy to anticipate/determine if a particular effect observed in a test tube or in vitro is also going to take place in vivo, it is clear that vanadium ions may interfere with many metabolic processes at many distinct levels. Emphasis is given on structural and functional aspects of vanadium–protein interactions relevant for vanadium binding and/or for clarification of role of the metal center in the reaction mechanisms. The additional knowledge that the presence of vanadium can change the action of a protein, other than simply inhibiting it, may also be important to understand how vanadium affects biological systems. This possibility, together with the vanadate–phosphate analogy further potentiates the belief that vanadium probably has relevant functions in living beings, which may involve interaction or incorporation of the metal ion and/or its compounds with several proteins. © 2015 Elsevier B.V. All rights reserved.

1. Introduction Many metal ions have a general tendency to interact with biomolecules, therefore it is not surprising that natural evolution has incorporated some of them into performing a wide variety of tasks and playing crucial roles in organisms [1,2]. Vanadium is a transition metal that is widely distributed in soil, crude oil, water and air and its compounds may have oxidation states ranging from −3 to +5. Being ubiquitously present in our planet’s crust it is thus not surprising that it also found roles in biological systems namely as an essential element for many living beings. Only VIII -, VIV - and VV -species are of biological relevance and nature has evolved several enzymatic systems using vanadium in their active sites as relevant components for their function including vanadium-dependent haloperoxidases, nitrogenases and vanabins. 1.1. Vanadate(V)-phosphate analogy Phosphate is very important in biological systems and is involved in an extensive number of biological recognition and bio-catalytic systems. It is known that vanadate(V) and phosphate participate in similar reactions. Vanadium inhibits several ATPases with different efficiency [3]. Moreover, vanadium and vanadium compounds also inhibit different proteins such as phosphatases (alkaline phosphatases, acid phosphatases and tyrosineprotein phosphatases) [4,5], ribonucleases, phosphodiesterases,

phosphoglucomutase and glucose-6-phosphatase [5]. Inhibition can be very strong and the enzyme-inhibitor constant (Ki ) values range from 10−5 to 10−7 M [6]. The ability of vanadium to inhibit these enzymes is closely related to the physicochemical properties of vanadate(V) and phosphate (see below), with vanadate(V) showing a greater flexibility in this coordination geometry. In most cases, enzyme activation by vanadium follows indirect mechanisms. Vanadium can stimulate the enzyme activity through the formation of complexes with ligands that resemble the structure of physiological substrates. For instance, the glucose6-phosphate dehydrogenase is activated by vanadate(V). Another mechanism of activation or inhibition involves the phosphorylation of tyrosine residues. Vanadate(V) also forms esters with Tyr residues mimicking their phosphorylation process, with a great impact in several biological events and similar actions have been proposed also for VIV O2+ . Monovanadate(V) and phosphate are structural analogues (see Fig. 1). Besides monovanadate(V) being structurally similar to phosphate, the acid–base equilibria operating and other types of reactions (e.g. V and/or P ‘ester’ formation) are also comparable. Nevertheless, there are some structural and pKa differences, namely the relative stability of 5-coordinate trigonal–bipyramidal structures or intermediates, bound in protein active sites, differs for vanadate and phosphate [5]. From a geometrical point of view, the two anions are not much different and vanadate(V) is a competitor in sites commonly occupied by phosphate. However, there are also significant differences:

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Fig. 1. Analogy between phosphate and vanadate(V): monovanadate(V) and phosphate are structurally very similar sharing similar acid–base equilibria and other types of reactions (e.g. V and/or P ‘ester’ formation). Some structural and pKa differences occur as well [5], namely the relative stability of 5-coordinate trigonal–bipyramidal structures or intermediates, bound in protein active sites, differs for vanadate(V) and phosphate.

(i) due to the different pKa , at physiological pH and ionic strength, vanadate(V) is mostly present in the form of H2 VO4 − , while HPO4 2− and H2 PO4 − exist in approximately equal amounts; (ii) phosphorus can only attain the coordination number 5 in transitional states, of relatively high energy, while vanadium easily forms stable complexes with coordination numbers 5 or 6. Therefore, once incorporated into the active site of a phosphate-dependent enzyme, taking the place of phosphate, the activity of this enzyme may be inhibited [5]; (iii) vanadium is considerably more electrophilic than phosphorous and V O bonds are less covalent than P O bonds, with the VO bond about 15% longer than the corresponding PO bond [7]. As these chemical entities are 3-dimensional and making simple geometrical calculations this would imply for “vanadate(V)’s” volume an increase of 25–40% relative to that of phosphate (assuming a similar geometry). Of course the protein may adapt to this distinct size, but the resulting displacements will be energetically unfavorable, being partly compensated by changes in the ideal structure of the transition state in an extent that will depend on each individual protein.

To evaluate what are the “volumes” of phosphate and vanadate(V) anions in proteins, searching the Protein Data Bank (PDB) a few examples were chosen where both the vanadate(V)-containing and the phosphate-containing structures have been determined with good, or at least medium resolution and both anions are in similar sites. The volume and area calculations were carried out using UCSF Chimera [8]. One first example is PhoX, an alkaline phosphatase. For the enzyme containing phosphate (PDB: 4ALF) the volume and area are 52.4 A˚ 3 and 73.0 A˚ 2 , respectively, while for the vanadate(V)containing structure (PBD: 3ZWU), these are 55.6 A˚ 3 and 81.4 A˚ 2 , respectively. Thus, the increase in volume is ∼6% and that for the area is ∼12% when binding vanadate(V). Another example is Nacetylneuraminate-9-phosphate phosphatase (HDHD4), where for the phosphate-containing structure (PBD: 4KNV) the volume is 51.5 A˚ 3 and the area 71.9 A˚ 2 , while for the vanadate(V)-containing structure (PBD: 4KNW) the volume is 58.8 A˚ 3 and the area 87.7 A˚ 2 , thus, when binding vanadate(V) the increase in volume is ∼14% and that for the area is ∼22%. These calculations are a rough approximation as for the vanadate(V)-containing structures the coordination geometry is trigonal bipyramidal and for phosphate it is not, but the numbers give an idea of the stress imposed on vanadate(V) when this

metal ion binds in the phosphate sites of enzymes. Additionally, since the inhibitor (vanadate(V) anion) does not perfectly match the active site, changes in hydrogen bonds and electrostatic interactions within the protein are necessary when accommodating it, this most probably being an energetically unfavorable process for the protein when compared with the binding of phosphate. (iv) both vanadate(V) and phosphate readily form linear oligomers in the tetrahedral state, in contrast to other oxyanions such as molybdate and tungstate. However, condensation occurs spontaneously for vanadates(V) in acidic solution, with the consequence that lower oxidation states could be accessible, whereas oligophosphates are (kinetically) significantly less stable than oligovanadates and phosphorous is most likely to be found in the +V state under physiological conditions [9]. The competitive behavior of vanadate(V) with respect to phosphate is likely the clue for the insulin-mimetic/insulin-enhancing effect of vanadium compounds, as well as inhibition of the activity of phosphatases. Moreover, in dilute solutions with pH > 6, VV exists as H2 VO4 − or HVO4 2− , while free VIV O2+ exists significantly as VIV O(OH)3 − , being structurally similar to VV O(OH)3 . Inter-conversion between VIV and VV may thus be fast and easy in physiological conditions. It was first suggested by Crans et al. [5] that, besides monovanadate(V), VIV O(OH)3 − may also mimic phosphate [5,10,11]. Therefore, the fact that VIV O-species also inhibit different enzymes (in some cases with a greater potency than vanadate(V)) may be based on the formation of VIV O(OH)3 − and its resemblance to vanadate(V), namely for cysteine-based phosphatases. The generation of a stable transition state analogue (TSA) between vanadate(V) and an enzyme has been considered the reason for its inhibitory effect on the catalytic activity and studies by X-ray diffraction (XRD) showed that vanadate(V) often binds in the active site of most phosphatase hydrolases with a trigonal–bipyramid geometry. However, other geometries exist such as tetrahedral [12–14], namely when in the phosphatase–vanadium complex vanadate(V) is acting as a product inhibitor. Several reviews have been published concerning the bioinorganic chemistry of vanadium or the role of vanadium in biological systems, namely since the year 2000 [5,6,15–35], but after the review of Crans in 2004 [5], no other work focused on a global overview of interactions of vanadium and vanadium compounds with proteins appeared in the literature. In this work, we present

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Fig. 2. Cartoon representation of vanabin2 structure solved by NMR methodologies. In addition to the four ␣-helices, the nine disulfide bonds (yellow) are depicted in a stick representation. By the observation of the disulfide bridges, it is possible conclude on their importance for the maintenance of the correct tridimensional structure [44].

a comprehensive overview of the interactions of vanadium with proteins focusing on the currently available structural data. The discussion addressing the information of V-protein interactions is organized in three main sections, specifically: (i) V uptake and transport, (ii) role of V in the protein activity and function and (iii) V as substrate analogue or inhibitor.

2. Uptake and transport 2.1. Vanabins Related to this topic several reviews have been published, e.g. [5,36–39] and for more detailed aspects the reader should address to them. In the living cells, metal ions, required for physiologically essential functions, are homeostatically maintained at low concentrations in the sub-␮M to ␮M range. Some organisms, called hyper-accumulators, collect very high levels of metal ions, thus requiring special systems to ensure their selective accumulation. Typical of such organisms are the ascidians, also known as sea squirts or tunicates. In fact, about one century ago, Henze published a paper describing the occurrence of such feature in the blood cells of ascidians [5]. Besides the exceptional high levels of V in the organisms, this finding led to the hypothesis of vanadium participating in oxygen transport, thus being a putative prosthetic group in respiratory pigments [5]; later it was demonstrated that V does not play such a role. In addition, these variable-size marine beings, belonging to the phylum Chordata, are important as they share some characteristics with the vertebrates allowing their use as a study model organism [5,38]. In natural environments, vanadium is usually in the +5 state (HVO4 2− or H2 VO4 − , here designated by VV , monovanadate(V) or V1 ) but in several species of ascidians, known to accumulate high levels of vanadium in their blood cells, most of the vanadium is in the +3 state (VIII ), VV being reduced via the +4 state (VIV O2+ , here designated by VIV ) during its assimilation and accumulation.

The highest amount of V in blood cells was found for Ascidia gemmata, where the vanadium concentration can reach 350 mM [40], which is 107 times that found in seawater (35 nM) [41,42]. The concentration of metal ions in the cytoplasm is strictly regulated by membrane transporters involved in uptake and export. Their compartmentalization into organelles decreases the concentration of free metal ions in the cytoplasm, hence also acting as reservoirs. The specific blood cells involved in the vanadium accumulation are named vanadocytes. These cells contain vacuoles with a quite low pH (≈2) and a high concentration of sulfate, where vanadium is stored. Although vanadium uptake occurs as VV , in the vanadocytes it is reduced to VIV , using NADPH and/or glutathione from the pentose phosphate pathway, and in the vacuoles it is further reduced to VIII [5,43]. Consequently, it is clear the existence of several proteins (and of reducing agents) that take part in the process of storage and reduction of vanadium in the vanadocytes. Vanabins are a family of proteins located in the vanadocytes’ cytoplasm and are involved in the VIV uptake. They act as a metallochaperones, responsible for the transport of the metal from the cytoplasm to the vacuoles [5,43,44]. The characterization of these proteins shows that vanabin1 (12.5 kDa) and vanabin2 (15 kDa) are able to bind up to 10 and 20 VIV ions with dissociation constants of 2.1 × 10−5 and 2.3 × 10−5 M, respectively [44]. Despite their biological importance in the vanadium accumulation by ascidians, there is not a great range of structures deposited in the PDB. In fact, there is only one entry of a vanabin structure (PDB: 1VFI) [33,44]. This corresponds to a vanabin2 protein obtained from cytoplasm of vanadocytes of the Ascidia sydneiensis samea species and it was solved by 15 N HSQC (1 H–15 N heteronuclear single-quantum coherence) NMR methodologies [44]. Even with the absence of VIV bound to the protein, valuable information is provided by this structure. Vanabin2, presenting four ␣-helices in a bow-shaped conformation, is composed by 91 amino acids, including 18 cysteines which form nine disulfide bonds (Fig. 2) [33,43,45]. The disulfide bonds are essential in the stabilization of the structure and for the binding of VIV ions. This

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53

Fig. 3. Surface representation of vanabin2 structure (gray) with the respective residues involved in the vanadium binding (red). NMR experiments with vanabin2 incubated with VIV were carried out enabling the identification and localization of such aminoacids. The protein is shown in different perspectives (rotation of 90◦ , 180◦ and 270◦ ) [44].

was demonstrated when the protein was incubated with dithiothreitol (DTT). In the presence of this strong reducing agent, the 15 N HSQC spectrum showed the disappearance of the structural elements and moreover the binding to VIV decreases significantly [45]. Furthermore, 1–4 mM of GSH is enough to partially cleave the disulfide bonds of vanabin2, thereby forming intermediate structures [46]. Titration experiments by sequential addition of VIV OSO4 to vanabin2 were also carried out in order to obtain details on the residues involved in the binding process or located next to the binding site. The 15 N HSQC spectra provided information regarding the amino acids implicated in the VIV binding as their resonances are missing or altered when compared to the native spectrum (Fig. 3) [47]. In fact, the obtained results show that VIV ions, which are exclusively localized on the same face of the molecule, are coordinated by amine-N donors from lysines, arginines and histidines, corroborating the EPR and ESEEM data previously collected [44,45]. The mechanism of metal selectivity in vanabins is not yet fully understood [36], although the effects of acidic pH on selective metal binding and on the secondary structure of vanabin2 have been extensively studied [48]. Besides VIV , vanabin2 also binds FeIII and CuII ions under acidic conditions. Additionally, CD spectral studies showed no changes on the secondary structure of Vanabin2 upon pH changes [48]. Two other vanabins, designated as vanabin3 and vanabin4, have primary structure similar to those of vanabin1 and vanabin2 and have also been recognized as VIV binding proteins [47]. Structural modeling of vanabins 1, 3, and 4, based on the NMR

data of vanabin2, resulted in fairly good homologous structures. The results suggest that the nine disulfide bridges derived from the 18 cysteines present are well conserved among the different members of the family [36]. Although vanadium is presumably taken up from seawater through the branchial sac or alimentary canal, transferred to the coelom and concentrated in vanadocytes, it is not known if carrier proteins are involved in the transport of V from the coelomic fluid (blood plasma) into the vanadocytes. A few V-associated proteins were identified, and it was shown that the major contributor is vanabinP [48]. VanabinP binds a maximum of 13 VIV ions per molecule with a Kd of 2.8 × 10−5 M and acts as a VIV -carrier. The NMR structure of vanabin2 was also used to model VanabinP and the obtained structure differs quite significantly from the original, especially in the long loop between the 3rd and 4th helices [36]. More recently, other studies pointed to the possibility of a VV reductase action in addition to the VIV transport function [43,49,50]. In parallel, in attempts to obtain more insights on their role in the specificity of vanadium accumulation, some investigations were done to understand if vanabins are able to bind and reduce other metals [43,46–48]. The phosphate anion transporter is one possible way for the transport of VV ions [51]. Another possible pathway is a transferrin (TF)–transferrin receptor (TFR)–mediated process. TF homologs were cloned from three ascidian species, A. sydneiensis samea [52], Halocynthia roretzi [53] and Ciona intestinalis [54], although the TFR gene was not found in ascidians [55]. Functional analyses suggested that the H. roretzi TF binds to FeIII [53], while one

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Fig. 4. A possible cascade for the thiol–disulfide exchange reactions conjugated with NADPH, GR, GSH, vanabin2, and vanadium ions. Adapted from ref. [47].

of the two C. intestinalis TFs binds to both FeIII and VV [49,50]. These are monolobal proteins that are unique to ascidians. In A. sydneiensis samea, besides vanabinP, another V-binding protein, VBP-129, was identified. VBP-129 is a 129 residues-long protein that binds to VIV ions [52]. The mechanism by which vanabinP and VBP-129 capture V in the blood plasma and subsequently transfer the vanadium ions, remains to be understood [36]. NADPH is most probably involved as reducing agent in the reduction of VV to VIV ; as mentioned above, thiols, such as glutathione (GSH) and cysteine, are also candidates for the direct reduction of VV to VIV [56–58]. Michibata and coworkers showed that vanabin2 catalyzes the reduction of VV to VIV in the presence of NADPH, glutathione reductase (GR) and GSH [37]. This suggests that electrons may be transferred from the donor NADPH to the acceptor VV ions via thiol–disulfide exchange reactions. In this process, the disulfides are converted to thiols by GSH and the obtained oxidized glutathione (GSSG) is then re-reduced by GR [37]. The GR disulfides produced are then reduced to thiols by NADPH [37] (Fig. 4), and these processes may be linked to the pentose-phosphate pathway. VIV is further reduced to VIII and XAS studies suggested that globally the reduction of VV to VIII takes place in a single coordination environment [37,59]. According to the obtained data, the two reduction steps VV → VIV → VIII occur sequentially in an EDTA-like 7-coordinate ligand array under mild cytoplasm-like conditions. In the proposed model, a proton electrochemical gradient generated by V-ATPase is the driving force for VIV transport from the cytoplasm into the vacuoles [37], and several other pathways may exist that function together. 2.2. Vanadium in polychaetes The accumulation of V is also found in other marine organisms namely in polychaetes, with special emphasis on tube-dwelling fan worms of the Sabellidae family [60–64]. Vanadium contents are in the range of 320–1350 ␮g/g of dry weight of the whole body. Unlike in the case of Chordates where the highest level of V is found in the blood (coelomic) cells, in the polychaetes members, higher amounts of the metal were measured in branchial crowns of Pseudopotamilla occelata and Perkinsiana littoralis, where vanadium is stored in vacuoles of the epithelial cells, associated with high amounts of sulfur, predominantly in the form of sulfate [36]. Amounts of vanadium existing in marine organisms, with particular emphasis on hyper-accumulation of V in polychaetes, were recently revised by Fattorini and Regoli [65] and previously by Ishii [60]. Particularly elevated concentrations of vanadium were measured in branchial crowns of P. littoralis. More than 50% of V was associated to the heavy pellet, 35% to cytosol and 10% to mitochondria and lysosomes. Within the heavy pellet fraction, the separation of the cartilaginous-like structures by enzymatic digestion demonstrates that vanadium is mostly present, >95%, in epithelial tissues, in agreement with results obtained in P. occelata, concentrating V in vacuoles of the epithelial cells [62,63]. In the cytosol fraction of P. littoralis, ca. 45% of vanadium is bound to proteins with a MW ranging within 3–10 kDa, the remaining being uniformly

distributed between soluble compounds (<3 kDa), and in proteins with MW of 10–30 and >30 kDa [65]. XAS analysis on living specimens of P. occelata is consistent with vanadium being predominantly as VIII ; these results, together with the existence of sulfate in the V-containing epidermis vacuoles [38], corroborate the hypothesis of comparable speciation to V in ascidians and that low MW vanadium binding proteins, similar to those described for ascidians, modulate transport, biotransformation and storage of vanadium in polychaete tissues [65]. As little is presently known about the interactions of vanadium with proteins in these organisms, we will not extent this subject further here and we recommend the review of Fattorini and Regoli [65] to the readers interested in this field. 2.3. Storage and blood proteins 2.3.1. Transferrins The transferrins are a group of single-chain glycoproteins containing ca. 700 amino acids (∼80 kDa) [66]. The most important of them are serum transferrin (TF), found in blood serum (human serum transferrin will be indicated as hTF), lactoferrin (LTF), found in milk and in many other biological secretions (tears, saliva) and ovotransferrin or conalbumin (OTF), isolated from avian egg white [67]. The main function of hTF is the transport of iron in the organism [67]. Transferrins are composed of two globular lobes, connected by a short peptide linker and designated as N-terminal (TFN ) and Cterminal (TFC ) lobes (or domains) [66]. They bind reversibly two Fe3+ ions in the two sites in the N- and C-terminal region and each Fe3+ ion is bound in an octahedral environment to one AspO, two Tyr-O, one His-N and two O-atoms from a carbonate anion, anchored in place by an Arg residue [67–70]. Even if human serum transferrin and lactoferrin are quite similar in sequence and structure and coordinate iron in the same manner, they differ in their affinities for Fe3+ as well as in their receptor binding properties [71]. The binding of two Fe3+ forming (FeIII )2 TF causes the change of conformation from the “wide-open” to the “closed” form [68,69,72], but only the “closed” form of transferrin can be recognized by the hTF-cell receptors and internalized by the cell through a receptormediated endocytosis process [67]. The binding of Fe3+ is promoted by carbonate anions, which are anchored by electrostatic and hydrogen bonding interactions with positively charged arginine residues; (hydrogen)carbonate can be replaced by other anions, called synergistic [68,73]. Depending on the Fe3+ saturation degree, TF is indicated as apo-TF (no iron bound), monoferric hTF, (Fe)hTFC or (Fe)hTFN , and diferric hTF, (Fe)2 hTF; this last case, corresponding to the iron saturated protein is also designated as holo-hTF. The concentration of hTF in the blood serum is 37 ␮M [74]. In normal plasma, only 30% of transferrin is bound to Fe3+ ions with a distribution of approximately 27% Fe2 hTF, 23% Fe-hTFN , 11% Fe-hTFC and 40% apo-hTF [75]; these values correspond to a concentration of ca. 50 ␮M of available hTF binding sites. Therefore, hTF can also bind other metal ions, including Bi3+ , Ga3+ , In3+ , Al3+ , Cu2+ , Mn2+ , Zn2+ , Ni2+ , Ru3+ , as well as vanadium in its three oxidation states (+3, +4, +5) [25].

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Fig. 5. Proposed coordination environment of V bound to hTFN site of transferrin: (a) VIII ; (b) VIV O2+ and (c) VV O2 + . The proposed networks of hydrogen bonds involving carbonate and V O groups is also shown. Adapted from Ref. [25].

The system VIII /apo-hTF has been studied by HPLC and FPLC coupled with ICP-MS [76–78], by MALDI-TOF [78], by spectroscopic techniques [79–81], by gel electrophoresis and ICP [81] and by CD spectroscopy [80,81]. In blood serum samples, the behavior of VIII towards apo-hTF is similar to that of FeIII and AlIII [76,78]. In all cases, VIII associates to hTF with the synergistic effect of carbonate ions. This experimental evidence suggested a coordination of VIII in the two sites hTFN and hTFC of transferrin analogous to FeIII [82] (Fig. 5). However, in contrast with the VIII species formed with human LTF which are rapidly oxidized by O2 [82], (VIII )2 hTF species is quite stable to atmospheric oxygen and is not easily substituted by FeIII . Gel electrophoresis studies have confirmed that the binding of VIII to apo-hTF is quite similar to FeIII , indicating that (VIII )2 hTF also assumes the ‘closed’ conformation similarly to (FeIII )2 hTF, thus it is probable that it is recognized by TFR [81]. The only attempt to quantify the value of the VIII -hTF binding constant was carried out by Costa Pessoa and Tomaz, who predicted a value of log K1 = 20 ± 1.5 (Table 1) [25]. The order of affinity of the three V oxidation states towards hTF is therefore: VIII  VIV  VV , in the presence of carbonate [25] and VIII ∼ VIV > VV in its absence [77]. The systems formed by VIV O2+ with apo-hTF [30,73,83,90–100], apo-LTF [101] and apo-OTF have been extensively studied in the literature [102,103]. The system VIV O2+ /apo-hTF has been described postulating the formation of (VIV O)(apo-hTF) and/or (VIV O)2 (apo-hTF), depending on the ratios VIV O2+ /apo-hTF, in which the VIV O2+ ion is bound to one or both the Fe3+ sites. Spectroscopic results are compatible with the coordination environment shown in Fig. 5b, and a recent theoretical study indicates that the most stable structure for the hTFN ˚ site is characterized by the following bond distances: V O 1.59 A, ˚ V O(Asp63) 1.92 A, ˚ V N(His249) 2.13 A˚ and V O(Tyr188) 1.92 A, suggests that the protonated O(Tyr95) occupies the axial position and interacts very weakly at 3.28 A˚ from V [104]. This study also indicated that the structure with CO3 2− ion is slightly preferred over that with HCO3 − , the first being favored by a hydrogen bond with OH group of Tyr95 and Ala126 [104]. The plane of the carbonate ligand appears to be essentially parallel to the V O direction (dihedral angle of −5.6◦ ) with the distance V O(carbonate) of 1.79 A˚ [104]. Costa Pessoa et al. reported recently gel electrophoresis data which indicate that the conformation of apo-hTF closes upon formation of (VIV O)2 (apo-hTF) [81]. The (hydrogen)carbonate can be replaced by other biologically relevant synergistic anions [105], even if under physiological conditions its concentration (∼27 mM) should be high enough to supply this role. Among the biologically relevant anions, oxalate, phosphate, lactate (only at the N-terminal site) and citrate are synergistic. Only lactate seems to be capable of replacing (hydrogen)carbonate at the hTFN site to form (VIV O)hTF(lact) (Table 1). VIV O2+ has been used as a spin-probe to determine the coordination environment of metal ions in proteins by EPR spectroscopy [93,106] and it was also applied in the study of the

VIV O2+ /transferrin systems. EPR spectra recorded in the solutions containing VIV O2+ and apo-hTF at physiological pH are composed by two sets of resonances (A and B) [73,90,93]. The B resonances can be further separated into two components, indicated as B1 and B2 , which can be resolved in the Q-band spectra [92] and in the Xband spectra using D2 O [99]. The A resonances correspond to the binding to the hTFC site and those B to the binding to the hTFN site [92]. They are countersigned by specific EPR parameters (Table 2) [93]. The rather high value of the 51 V hyperfine coupling constant (Az ) is consistent with one of the two Tyr-O ligands occupying the axial position trans to the oxido ligand (Fig. 5). ESEEM spectroscopy confirmed the equatorial coordination of a His-N, the coordination of a (hydrogen)carbonate and the non-presence of water or hydroxide ligand bound to vanadium [107], whereas UV difference spectra suggest the interaction of V with Tyr-O [82]. The first values reported in the literature for the binding constants to the two lobes of human serum transferrin were obtained by Kiss et al., and the values of 13.2 ± 1.6 and 12.2 ± 1.6 were obtained for log K1 and log K2 , with a rather high uncertainty [108]. Subsequently, partial displacement reactions between (VO)hTF and [VOL2 ] were used to estimate the stability constants [84,98,109] (Table 1). The values reported indicate that the affinity of the two sites, hTFC and hTFN , for VIV O2+ ion is similar. Aisen et al. have shown that hTFC (or A) binds FeIII more strongly than the hTFN site (or B) by a factor of about 20 (1.3 in log units) at pH 7.4 with a physiological (hydrogen)carbonate concentration [110]. The system VIV O2+ /holo-hTF was also investigated [88]. Inside cells iron is released from the N-lobe of human serum transferrin around pH 5.7, whereas the C-lobe retains Fe3+ ion up to pH 4.8 [67], due to the protonation of some basic residues. The results of the experiments depend on the pH at which the protein is added to the solution containing vanadium. When holo-hTF is added at pH ca. 4.0 (Fig. 6b), there is an exchange between Fe3+ ions released from two hTF binding sites and VIV O2+ , which can occupy both the hTFN and hTFC sites, with the ratio Fe3+ /VIV O2+ in the sites depending on the ratio log K1 (Fe(hTF))/log K1 (VO(hTF)). The part of VIV O2+ not coordinated to the binding sites can interact with non-specific binding sites (named sites C), identifiable by the appearance of the lower field resonances indicated by II in Fig. 6. When holo-hTF is added at pH 5.2, Fe3+ is released mainly from hTFN and most of VIV O2+ can interact with this site; comparing with the experiment at pH 4.0 a larger amount of vanadium remains in solution and, as a consequence, the relative concentration of the EPR signal bound to the non-specific sites C increases. Finally, when holo-hTF is put in contact with a VIV O2+ solution at pH 7.4 (Fig. 6d) the exclusive binding to the non-specific sites C is observed because hTFN and hTFC sites are occupied by Fe3+ [88]. EPR parameters, gz = 1.944 and Az = 165.4 × 10−4 cm−1 (Table 2), are compatible with the coordination of His-N/Asp-COO/Glu-COO/H2 O donors. The coordination of Fe3+ in the two lobes of OTF is identical to hTF [111]. Room and liquid nitrogen temperature EPR spectra of the

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Table 1 Stability constants (log K) for the species formed by VIII , VIV O2+ and VV ions with blood proteins. Protein

Species

Log K

hTF hTF hTF hTF hTF hTF hTF hTF hTF

(VIII )hTF (VIV O)hTF (VIV O)hTF (VIV O)2 hTF (VIV O)2 hTF (VIV O)hTF(lact) (VV )hTF (VV )hTF (VV )2 hTF

20.0 13.0 13.4 12.5 11.8 14.5 6.0 7.5 5.5

± ± ± ± ± ± ± ± ±

1.5 0.5 0.2 0.5 0.4 0.8 0.1 0.2 0.2

Ref.

Protein

Species

Log K

Ref.

[25] [84] [30] [84] [30] [84] [30] [83] [30]

hTF HSA HSA HSA HSA HSA HSA IgG Hb

(VV )2 hTF (VIV O)x HSA (VIV O)x HSA (VIV O)2 HSA (VIV O)2 HSA (VV )HSA (VV )HSA (VIV O)IgG (VIV O)Hb

6.6 ± 0.3 9.1 ± 1.0 9.1 ± 0.4 20.9 ± 1.0 20.6 ± 0.4 1.8 ± 0.3 3.0 10.3 ± 1.0 10.4 ± 1.0

[83] [84] [85] [84] [85] [86] [87] [89] [89]

Table 2 EPR parameters of VIV O species bound to the binding sites of transferrins.a Protein

Site

gz

Az

Ref.

gz

Az

Ref.

Apo-hTF

A B1 B2

1.938b 1.941b 1.937b

168.0b 170.3b 172.4b

[92,93] [92,93] [92,93]

1.937c 1.941c 1.935c

168.3c 170.5c 171.8c

[99] [99] [99]

Holo-hTF

C

1.944

165.4

[88]

Apo-OTF

A B C

1.940 1.937 1.939

171.2 170.7 163.9

[102] [102] [102]

Apo-LTF

A B

1.944 1.938

169.6 168.9

[103] [103]

a b c

All Az values reported in 10−4 cm−1 . Data obtained from a Q-band spectrum. Data obtained from a X-band spectrum.

system VIV O2+ /apo-OTF showed that VIV O2+ ions bind in three magnetically distinct environments (A, B, and C) [102]; the three signals were attributed to different metal site configurations associated to several conformations of the protein. The coincidence of Az for the resonances A, B, C of OTF with those A, B1 , B2 of hTF suggests that the V environments are also similar (Table 2). Human LTF binds two VIV O2+ ions in two specific metalbinding sites, characterized by a similar but not equivalent chemical environment [101,112]. VIV O2+ binding is promoted by carbonate/bicarbonate and oxalate which play the role of synergistic

Fig. 6. High field region of the anisotropic X-band EPR spectra recorded at pH 7.4 on frozen solutions (120 K) containing: (a) VIV O2+ /apo-hTF 2/1 (VIV O2+ 5 × 10−4 M); (b) VIV O2+ /holo-hTF 2/1 (VIV O2+ 5 × 10−4 M), in which holo-hTF was put in contact with VIV O2+ ion at pH ca. 4.0 and then pH raised to 7.4; (c) VIV O2+ /holo-hTF 2/1 (VIV O2+ 5 × 10−4 M), in which holo-hTF was put in contact with VIV O2+ ion at pH 5.2 and then pH raised to 7.4; (d) VIV O2+ /holo-hTF 2/1 (VIV O2+ 5 × 10−4 M), in which holo-hTF was put in contact with VIV O2+ ion at pH 7.4. With I and II are indicated the resonances MI = 7/2 of the species in which the VIV O2+ ion is bound to the Fe specific sites and to the nonspecific sites C, respectively. Adapted from Ref. [88].

anion [112]. Two different sites have been identified, that at the N-terminal lobe (B site) and that at the C-terminal lobe (acid-stable or A site) [101]. Analogously to hTF, human LTFN is acid-labile, whereas LTFC is acid-stable. This has allowed the preparation of mixed Fe3+ /VIV O2+ species with VIV O2+ bound to site B or A. Vanadium species have been demonstrated to have antidiabetic [113,114] and other therapeutic properties [35,114]. For example, VIV O(maltolato)2 (or BMOV) became the benchmark compound for the new molecules with anti-diabetic action [113,114] and its derivative VIV O(ethylmaltolato)2 (or BEOV) arrived to phase IIa of clinical trials [115]. Binding of VIV O compounds to hTF may have an important role in the transport and delivery of vanadium based drugs [26,30,84,88,97,116–121]. Two different and alternative binding modes of a VIV O-compound to apo-hTF, with L being a bidentate monoanionic ligand, have been proposed in the literature. The binding designated by Type 1 with cis-VIV OL2 (apo-hTF) stoichiometry [84,117–119,122] (Fig. 7) has been suggested by the comparison of Az values measured with solutions containing cis-VIV OL2 complexes and either apo-hTF or 1-methylimidazole (1MeIm), a model for the coordination of an imidazole-N of a His residue [84,97,117,118,123]. Type 2 corresponds to the binding of VOL+ unit at iron sites and has been suggested for L = ma, dhp and pyrimidinonato derivatives [109,120,121] (Fig. 7), based on the circular dichroism and EPR spectra, as well as ICP data, measured with solutions containing VIV OL2 , including cases where excess of complex and/or ligand is present. At the moment it is not clear if Type 1 and Type 2 co-exist, or if one of them is favored by the specific conditions. Recent experiments have demonstrated that the formation of mixed VIV O species is possible also with holo-hTF, for which the two metal binding sites are blocked by Fe3+ . In this case, the formation of cis-VIV OL2 (holo-hTF) is possible through the replacement of the equatorial water molecule by an accessible His-N in a Type 1 binding (Fig. 7) [88]. The analysis of surface His residues of apohTF and holo-hTF (PDB: 2HAU [124] and 3V83 [125]) has shown that the most accessible residues for this unspecific holo-hTF coordination are His289, His349, His473 and His606 (for apo-hTF they are two more, His14, His289, His349, His350, His606 and His642)

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Fig. 7. Binding mode for VIV O–-complexes to transferrins; example of BMOV and BEOV. In the case of Type 1 binding, histidine residue may be His14, His289, His349, His350, His473, His606 and His642, in the case of Type 2 binding, the VIV O:carrier stoichiometry is 1:1 and actual donor atoms of the Fe-binding site coordinated to VIV may be two of the four residues Asp63, Tyr95, Tyr188 and His249 of the N-terminal lobe of hTF, or two of residues His585, Asp392, Tyr426 and Tyr517 of the C-terminal lobe. Adapted from Ref. [81].

[88]. Since the conformation of holo-hTF upon the formation of cis-VIV OL2 (holo-hTF) remains ‘closed’, these mixed species can be recognized by the hTF receptor and this mechanism could be an alternative to the postulated passive diffusion through the cell membrane of neutral VOL2 complexes [126]. The binding of vanadate(V) to apo-hTF has been studied by difference UV spectroscopy [127,128], 51 V NMR [86,129,130], 14 N and 35 Cl NMR [94], electrophoresis [77], calorimetry [131] and recently by CD spectroscopy and gel electrophoresis [81]. Such techniques allow the distinction of the VV ions coordinated to the two different lobes of hTF. It has been suggested that VV occupies the same pockets as FeIII , or at least binds close to these sites. 51 V NMR chemical shifts at −529 and −531 ppm were assigned to the coordination of VV at C- and N-terminal sites [129]. UV difference spectroscopy measurements on human LTF suggested that V is present in the two sites in the VV O2 + form and coordinated by two Tyr-O, one His-N and one Asp-COO (Fig. 5c) [82]. One important difference with respect to VIII and VIV is that VV is able to bind to transferrins in the absence of (hydrogen)carbonate and of any synergistic anion [86,127]. VV can be displaced from the protein by the subsequent addition of ions with greater affinity towards apo-hTF, such as FeIII and GaIII [81,127]. The existence of the cationic form VV O2 + ion rather than the anionic vanadate(V) has been suggested based on the stability constants of VV with apo-hTF, which are ∼3–4 orders of magnitude higher than those with inorganic anions (phosphate, sulfate, hydrogencarbonate) [128]. However, by itself this argument is not really valid, as it is known that vanadate(V) anions bind several phosphatases and phosphorylases much strongly than phosphate. Additionally, it was also found that significant amounts of VV bound to hTF are detected by both 51 V NMR and ICP in solutions containing (FeIII )2 hTF, suggesting that at least some VV could be either bound to a distinct site other than the Fe3+ ones, or that it may act as a synergistic anion [81]. NMR results suggest that there is no competition between HCO3 − and H2 VV O4 − for the anion binding site of apo-hTF [86]. An interesting experimental evidence is that the fraction of free VV increases with (hydrogen)carbonate concentration; since (H)CO3 − cannot displace vanadate(V) from its apo-hTF complexes, this fact could be explained assuming the formation of (hydrogen)carbonate–VV adducts, similarly to the phosphate–VV compounds [86]. Recent CD evidence in the ternary AlIII /VV /apo-hTF system indicate the co-existence of a VV –hTF species with (AlIII )2 hTF. Since VV

is not able to displace AlIII from Fe-binding sites, either vanadate(V) may act as a synergistic anion instead of (hydrogen)carbonate promoting AlIII coordination or that it binds at a different site close to the iron sites [81]. Even in the system FeIII /VV /apo-hTF, CD spectra indicate the presence of VV –apo-hTF; therefore, it has been argued the probable binding of VV to residues close or belonging to the Fe-binding sites [81]. HPLC/high-resolution ICP-MS studies, that suggested that VV is bound to the ‘open’ form of hTF [77], was recently confirmed through gel electrophoresis experiments [81]. The consequence is that these species cannot be recognized by hTF receptors of the cellular membrane. However, VV can bind to holo-hTF this being a plausible form for the uptake of vanadium by cells [81].

2.3.2. Albumin Human serum albumin (HSA) is a large globular protein with several physiological roles ranging from transport of hydrophobic metabolites, such as fatty acids and bilirubin, to the maintenance of the blood osmotic pressure. Its average concentration in blood serum is ca. 0.63 mM, being the most abundant protein in the blood [74]. HSA, bovine serum albumin (BSA) and rat serum albumin (RSA) share similar structural motifs in the N-terminal part of the protein, specifically AspAlaHis for HSA, AspThrHis for BSA and GluAlaHis for RSA. The His residue in the third position is replaced by Tyr in dog serum albumin (DSA, GluAlaTyr) and porcine serum albumin (PSA, AspThrTyr), and Glu in chicken serum albumin (CSA, AspAlaGlu) [132]. HSA plays a major role in the transport of physiological CuII [132], ZnII [133,134] CoII and CaII , and toxic NiII and CdII [135]. Albumins may contain a primary metal binding site, the N-terminal site (NTS or site I) [132], and a secondary site, the multi-metal binding site (MBS or site II) [136]. The NTS is present in the albumins with a His in the third position such as HSA, BSA and RSA and is the specific site CuII and NiII ions. It supplies a square planar geometry through the donor set (NH2 , N− , N− , His-N), where NH2 represents the terminal amino, N− the deprotonated amide groups of residues 2 and 3 and His-N the imidazole nitrogen of His3. The MBS site is located at the interface between domains I and II and is the primary binding site for ZnII [137], residues His67, His247, Asn99 and Asp249 being involved in the metal coordination. It also binds CuII and NiII with similar affinity, and CdII with an affinity that is one order of magnitude lower [136]. In addition, other two sites have been identified: site B (or CdB), specific for cadmium, provided with one His and

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several carboxylate residues [137–139], which is the strongest site also for CoII [135], and the site represented by the SH of Cys34, the only thiol group of HSA not involved in intramolecular disulfide bridges. The accessibility of Cys34 for most metal ions is quite restricted; this results in a high specificity in the metal interaction, demonstrated unequivocally for Au- and Pt-compounds [140,141]. The binary system VIV O2+ /HSA has been studied by several techniques [84,96,97,99,100,142–145]. From an examination of the EPR spectra measured at room temperature at pH ∼ 5, Chasteen and Francavilla distinguished one “strong” (recently designed with VBS1 [145]) and at least five “weak” binding sites (named VBS2 [145]) having different EPR parameters [142]. These results were subsequently confirmed by Willsky et al. [96] and Liboiron et al. [97]. VBS1 binds 1 mol of VIV O2+ and it was initially affirmed that it may be the NTS [142]. However, the coincidence of the CD spectra of BSA (with His in the third position) and PSA (with Tyr in the third position) allowed Yasui et al. to rule out the coordination at NTS with (NH2 , N− , N− , Nimid ) binding mode [146]. A further proof of this insight is the comparison of the experimental 51 V Az measured from EPR spectra in the system VIV O2+ /HSA (around 165 × 10−4 cm−1 ) with that of the model complex of the N-terminal site with (NH2 , N− , N− , Nimid ) coordination (GlyGlyHis), which is significantly lower (153 × 10−4 cm−1 ) [147]. Costa Pessoa and Kiss recently observed only a decrease of the EPR intensity when ZnII was added to the VIV O2+ -HSA solution and replaces VIV O2+ ion in the MBS [85]. Competition studies between ZnII and VIV O suggest that VIV O2+ has two types of binding sites, one of them corresponding to the MBS (VBS1). ZnII is able to displace VIV O2+ from this site, but not from the other sites (VBS2) [85]. EPR spectra of an equimolar solution VIV O2+ /HSA are characterized by the presence of the very weak signals attributable to a dinuclear species, denoted as (VIV O)2 d HSA [99], in which the two VIV O2+ ions interact with each other and the spectrum is characteristic of a S = 1 spin state. The low value for Az for the corresponding mononuclear species, obtained as twice that measured in the spectrum of dinuclear one (∼160 × 10−4 cm−1 ), indicates that the ligands are rather strong. It was suggested that Glu252, at ∼5 A˚ distance from Asp249 (one of the donors of the MBS), might be involved in V coordination in the dinuclear form [85]. When the metal:protein ratio is increased to 2:1 or higher, the EPR resonances are attributable to a multinuclear complex, (VIV O)x m HSA with a S = 1/2 state, indicating that the metal ions coordinated by albumin are not interacting [85,99]. For all these species, only one set of EPR resonances is observed at physiological pH (with Az of 165 × 10−4 cm−1 ), suggesting that at least five different metal ions are coordinated by albumin and have similar binding mode. These sites may coincide with those indicated by Chasteen and Francavilla as “weak” sites [142]; the exact number of such non-specific VIV O binding sites is not known exactly, being between five and twenty [142,144]. The value of Az is consistent with nonspecific interactions with carboxylate and/or imidazole side chains of Asp/Glu and His accessible residues. VIV O2+ remains bound to defatted BSA (dBSA) when ZnII was added and this confirms the presence of another type of binding site besides MBS; Costa Pessoa and co-workers suggested that also the amino acid side chains of the NTS site may be involved in the metal coordination, but with a distinct binding set from that of CuII and ZnII [145]. Therefore, when VIV O2+ is added to HSA the first two equivalents bind to MBS (VBS1) site at a distance of magnetic interaction, whereas further added moles bind to non-specific sites (VBS2), including NTS, without magnetic interaction. This is consistent with the results obtained by Orvig [97]. The progress in the knowledge of the VIV O2+ /HSA system allowed to determine the values of log ˇ for (VIV O)2 HSA (VBS1) and (VIV O)HSA (VBS2), although with a rather high uncertainty. The values reported for (VIV O)2 HSA are log K = 20.9 ± 1.0 [84] and 20.6 ± 0.4

[85], and for (VO)HSA are log K = 9.1 ± 1.0 [84] and 9.1 ± 0.4 [85]. Log K for (VIV O)HSA can be considered as a mean value for the coordination of VIV O2+ to “weak” binding sites VBS2, since the values of log Kx for the five-six equilibrium steps should differ only slightly from each other (not more that 0.1–0.2 log units) and from the mean value (around 0.3–0.4 log units) [84]. The behavior of the system VIV O2+ /BSA is similar to VIV O2+ /HSA. Az and gz values (167.6 × 10−4 cm 1 and 1.940, respectively) are close to those obtained for fatted/defatted HSA by Orvig [97], Sanna [99] and Costa Pessoa [85]. At pH 7.4, not all VIV O2+ are displaced by ZnII and the EPR signals of VIV O2+ bound to VBS2 are observed; the donors involved in the V binding may be accessible N-imidazole of His residues or O-carboxylate of Asp or Glu residues [145]. Yasui et al., based on CD spectra recorded on the iodoacetamide-modified BSA, which is a partially thiolate masked protein, suggested that VIV O2+ binds also at the thiolate group of BSA [146], but this observation requires confirmation. The different results of CD experiments in the systems VIV O2+ /PSA confirm that NTS site may be involved in the V binding. EPR spectroscopy suggests that in PSA the VBS2 is not exactly the same as in HSA and BSA (and this may reflect either variations in the amino acid residues in the V environment or changes in their orientation) and the presence of a third binding site (different from NTS and MBS) [146], already proposed for CuII [136]. Since specific coordination sites like those of transferrin for iron are lacking, HSA and BSA form mixed complexes with vanadium insulin-enhancing compounds after the interaction with accessible residues. Orvig and co-workers showed that maltol (Hma) can also form mixed ligand complexes VIV O(ma)(HSA) and VIV O(ma)2 (HSA), that, in authors’ opinion, could be so important in the transport of an insulin-enhancing agent to be considered as the pharmacologically active species [97]. The formation of cis-octahedral species with maltol and its derivatives such as etylmaltol (Hema) and kojic acid (Hkoj), with composition cisVIV O(ma)2 (HSA), cis-VIV O(ema)2 (HSA), cis-VIV O(koj)2 (HSA) was recently confirmed through EPR and DFT studies (Fig. 8) [118,119]. In these species the equatorial water molecule is replaced by an accessible His-N; therefore, their composition should be more correctly described as (VIV OL2 )n HSA, where n is the number of amino acid residues that bind VIV O2+ . The formation of these types of complexes was demonstrated for both picolinate (pic) and 1,2-dimethyl-3-hydroxy-4(1H)-pyridinonato (dhp) [84,117]. The values of log ˇ are listed in Table 3. Recently, it was shown that at physiological pH the main (VIV OL2 )n HSA species can co-exist with a minor (VIV OL)n HSA mixed complex, and CD spectra even suggested more than two types of binding sites [145]; this appears to be in agreement with the observation of Orvig and co-workers [97]. Makinen et al. reported that [VO(acac)2 ] exhibits the greatest capacity to enhance insulin receptor kinase activity in cells compared to other organic VIV O compounds, and a dose-dependent capacity to lower plasma glucose in diabetic laboratory animals, also exhibiting a sufficiently long lifetime in the blood stream to allow correlation with blood vanadium content. This was also correlated to its binding to HSA [149]. The results obtained for the VV -HSA interaction are not so clear and straightforward as for the VV -apo-hTF system. However, all of the studies published till now agree that the interaction is weak and unspecific, some of them suggesting that the binding sites probably involve surface carboxylic functions. The first study of Crans and co-workers with BSA assumed 1:1 complexes and allowed to the authors to find a value of 3.0 for log K [87]. Subsequently, Heinemann et al. studied the VV –HSA interaction by ultrafiltration, concluding that VV is bound to HSA only in very low concentration relative to the total HSA (maximum 0.3–0.4%) [150]. Kiss and co-workers re-considered the system VV -HSA; as no interaction was detected by 51 V NMR spectroscopy or fluorimetry, the binding

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Fig. 8. Mixed species formed in aqueous solution at pH 7.4 by HSA, IgG and Hb: (a) cis-VIV O(ma)2 (Protein), (b) cis-VIV O(ema)2 (Protein) and (c) cis-VIV O(koj)2 (Protein). Ma, ema and koj indicate maltolate, ethylmaltolate and kojate anions, protein indicates albumin, immunoglobulin G and hemoglobin. Adapted from Ref. [119]. Table 3 Stability constants (log ˇ) of the ternary VIV O complexes formed by insulin-enhancing species with HSA, IgG, Hb and the model 1-MeIm. Species IV

Maltol a

cis-[V OL2 (1-MeIm)] cis-VIV OL2 (HSA)b cis-VIV OL2 (IgG)b cis-VIV OL2 (Hb)b a b

19.12 19.5 19.6 19.6

± ± ± ±

0.01 1.0 1.0 1.2

Ethylmaltol

Kojic acid

dhp

19.32 ± 0.01 19.7 ± 1.0 19.8 ± 1.0 –

17.40 ± 0.02 17.5 ± 1.0 17.6 ± 1.0 –

25.4 25.9 25.6 25.8

Ref. ± ± ± ±

0.2 0.6 0.6 1.0

[84,118,119] [84,118,119] [118,119,148] [89]

Measured by pH-potentiometry. Measured by EPR spectroscopy.

properties of HSA were quantified on the basis of literature data: they estimated a value not higher than 1.8 ± 0.3, the log K of 3.0 reported by Crans being considered an overestimation [86]. Recently Castro and co-workers have put in evidence by 1 H saturation transfer difference (STD) NMR spectroscopy and computational docking studies that [VV O2 (dhp)(OH)(H2 O)]− , [VV O2 (dhp)2 ]− and [VV O2 (ma)2 ]− can bind to drug site I [151]. For [VV O2 (dhp)(H2 O)(OH)]− species, the model shows a large number of hydrophobic interactions with the planar ring, particularly with Leu238, Ala291, whereas the two O-oxido interact with Lys199 and His242. With [VV O2 (dhp)2 ]− Trp214 interacts with the aromatic ring, whereas other interactions involve Arg222, Leu238 and Lys195. The molecular docking and NMR studies indicate a preference of [VV O2 (dhp)(H2 O)(OH)]− compared to [VV O2 (dhp)2 ]− for the binding to drug site I [150]. For [VV O2 (ma)2 ]− , both maltolato ligands are stabilized by hydrophobic interactions with residues Tyr214, Phe223 and Leu238, conferring a higher affinity of this V species for the binding pocket with respect to uncoordinated maltol [150]. On the basis of these results, His242 is a plausible candidate for the coordination to cis-[VIV OL2 (H2 O)] species with L = ma, ema, koj, dhp and pic (see above). 2.3.3. Immunoglobulins Immunoglobulins (divided into the classes IgA, IgD, IgE, IgG and IgM) are glycoproteins having the capability to react in an immune response to foreign bodies introduced or inoculated into an organism. IgG is a “Y”-shaped protein and represents 75% of the total immunoglobulins. The concentration range of the IgG is 7.7–20 mg/mL, which corresponds to a mean value of 84 ␮M [152]. Till 2011, several studies of the interaction of IgG with vanadium in the three oxidation states were carried out with different techniques: the system VIII /IgG was studied by FPLC–ICP-MS methods [78], the system VIV O/IgG by EPR and FPLC–ICP-MS [78,96] and the system H2 VV O4 − /IgG by gel filtration and anion-exchange chromatography [153]. In all cases no interaction between vanadium and IgG was found. Two reasons can hinder to observe any interaction in these systems: the instrumental response which depends significantly on the experimental procedure used to prepare the solutions, and the non-specificity of the metal sites which results in very weak spectroscopic signals. To obtain an interpretable EPR spectrum, indeed, signal averaging was required which involves

repeatedly acquisition and subsequent sum of the individual spectra: the signal increases proportionally to N, where N is the number √ of scans, and the noise to N, owing to its random nature. Therefore, √ the enhancement of the signal-to-noise ratio is proportional to N [99]. The system VIV O2+ /IgG (human) was recently revisited [148]. The EPR spectra recorded at physiological pH shows that the complexation of the metal ion by IgG takes place (even if it is weak) and indicate that the VIV O2+ ion distributes on three distinct coordination sites, whose spectroscopic parameters are listed in Table 4. The three sites were named 1, 2 and 3 and the stoichiometry was indicated as (VIV O)x IgG with x = 3. On the basis of the 51 V Az value, the donors of site 1 are stronger than those of site 2, and the donors of site 2 are stronger than those of site 3 (Table 4). Most likely, the coordinating residues involved in the vanadium binding are highly solvent exposed, located at the protein surface [148,154]. The resonances of site 2 closely resemble those of VBS2 of HSA, for which the non-specific coordination of His-N and Asp/Glu-COO was proposed. The most probable candidate for VIV O2+ binding in site 1 appears to be a Ser-O− /Thr-O− rather than a Tyr-O− or Cys-S− . The three sites of IgG were modeled through DFT calculations according to the procedures established for the determination of the environment of other V-proteins [155]. This approach was used to characterize the active site, the catalytic cycle and the structure of other metal proteins [156] and consists in the simulation of the 51 V hyperfine coupling tensor A of a VIV O complex which can be considered a model for the active site. When 51 V A is calculated using the functional BHandHLYP and the basis set 6-311g(d,p), a good model of the V sites should give Az values in agreement with the experimental one with a deviation around 2–3% [157–163]. Therefore, DFT methods may be used to predict Az for a VIV O2+ species, and vice versa, to predict an unknown structure from its Az value. The comparison between the experimental and calculated values of Az is given in Table 4 and confirms the results discussed above. 2.3.4. Hemoglobin Hemoglobin (Hb) is the iron-containing protein involved in the transport of oxygen in all vertebrates [164]. In adult humans, the most common hemoglobin type is a tetramer consisting of two ␣ and two ␤ structurally similar subunits non-covalently bound, each composed of 141 and 146 amino acid residues, respectively.

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Table 4 EPR parameters of VIV O species bound to the three binding sites of IgG [148]. exptl a

a, b

Site

gz

Az

Acalcd z

3 2 1

1.947 1.951 1.960

167.4 163.6 158.8

166.7 163.6 159.9

a b

Probable donors N(His), COO− (Asp, Glu), H2 O N(His), COO− (Asp, Glu) N(His), COO− (Asp, Glu), O− (Ser/Thr)/O− (Tyr)/S− (Cys)

exptl

Experimental (Az ) and calculated (Acalcd ) Az values reported in 10−4 cm−1 . z Values calculated at the level of theory BHandHLYP/6-311g(d,p) with Gaussian software.

Hemoglobin is found in the erythrocytes or red blood cells with a concentration of ∼5.1 mM [164]. All studies on the interaction of V with hemoglobin concern the oxidation state IV, because VV is reduced to VIV in the reducing environment of the red blood cells [89,153,165–173]. In particular, vanadate(V) crosses the cell membrane via anion channels [169] and initially binds to the phosphate-binding site of (Na,K)ATPase on the cytoplasmatic side of the membrane [168]. Inside the erythrocytes, VV is reduced to VIV at the expense of cellular glutathione [167], as demonstrated by the fact that the GSH depletor diethyl maleate blocks the process [167,169]. Most of the experimental studies indicate that, inside the erythrocytes, VIV O2+ ion is bound mainly to hemoglobin (Hb) [89,153,166,167,172,173]. The level of VIV O2+ bound to Hb was diminished by addition of adenosine 5 -triphosphate (ATP) and 2,3-diphosphoglycerate, suggesting that other intracellular bioligands can compete with hemoglobin [167]. The study of the VIV O2+ /Hb system is possible by EPR spectroscopy even if in the presence of Fe2+ in the four heme sites because the iron signals do not superimpose much. Three unspecific Hb sites were considered, named ␣, ␤ and ␥ with formation of (VO)x Hb, with x = 2,3, with x determined by the variation of EPR signals for the resonances MI = 5/2 and 7/2 as a function of the molar ratio VIV O2+ :Hb [89]. The V environment in these three sites depends on pH. The experimental evidence indicates that in site ␣ only carboxylate groups of Asp or Glu residues are bound to V, whereas in the sites ␤ and ␥ the binding of His-N adds to Asp/GluCOO− . At pH 7.4, only sites ␤ and ␥ are occupied. The participation of histidine, aspartate and glutamate is in line with the V coordination environment of V–carboxypeptidase [174] and V–albumin [85,99] and compatible with the high number of surface His residues in hemoglobin [175]. The value of the stability constant for (VIV O)Hb (log ˇ = 10.4) (Table 1) is significantly lower than for hTF (log ˇ1 = 13.0), due to the coordination of VIV O2+ to the iron specific sites in apo-transferrin. Log ˇ for (VIV O)Hb is comparable with that of IgG (log ˇ = 10.3) one order of magnitude larger than that of albumin (log ˇ = 9.1). Therefore, the strength order of the blood proteins for VIV O2+ ion is hTF  Hb ∼ IgG > HSA. The ternary systems containing hemoglobin and the antidiabetic compounds [VIV O(ma)2 ], [VIV O(dhp)2 ], [VIV O(acac)2 ] (acac = acetylacetonato) show analogies with what was published for the corresponding systems with hTF, HSA and IgG. In particular, at physiological pH, VIV O(ma)2 (Hb) and VIV O(dhp)2 (Hb) are formed with log ˇ of 19.6 and 25.8 (Table 3), comparable with those of the analogous species of albumin and immunoglobulin G. In these species, a donor from the side-chain of an amino acid residue would replace the equatorial position occupied by water in cis[VIV O(ma)2 (H2 O)] and cis-[VIV O(dhp)2 (H2 O)]; the most probable candidate are His-N and Asp/Glu-COO. By using EPR spectroscopy Sanna at al. concluded that VIV O(ma)2 (Hb), VIV O(dhp)2 (Hb), VIV O(pic)2 (Hb) and VIV O(3mepic)2 (Hb), with 3-mepic = 3-mehylpicolinate anion, are found during experiments of uptake of [VIV O(ma)2 ], [VIV O(dhp)2 ], [VIV O(pic)2 (H2 O)] and [VIV O(3-mepic)2 ] by the red blood cells. In particular, as determined by EPR, [VIV O(dhp)2 ]

transforms quantitatively in VIV O(dhp)2 (Hb), whereas [VIV O(ma)2 ], [VIV O(pic)2 (H2 O)] and [VIV O(3-mepic)2 ] in cis-VIV O(carrier)2 (Hb) and cis-VIV O(carrier)2 (Cys-S− ) with the equatorial coordination of a thiolate S− stemming from GSH or, most probably, from a membrane protein (Fig. 9) [89,173]. 2.3.5. Ferritins Ferritins are proteins that store iron as a hydrous ferric oxide mineral core (with approximate composition (FeOOH)8 ·FeO·H2 PO4 ) of ∼80 A˚ diameter. They consist of 24 subunits of two types, H and L, which assemble to form the protein shell with a molecular mass of ca. 480 kDa. The H- and L-subunits share about 50% in sequence homology and are structurally similar [176]. A number of iron binding sites have been identified and these include Asp and Glu residues of the threefold channels through which Fe2+ is thought to pass during its path towards the interior of the protein shell. Vanadium is a natural constituent of horse spleen ferritin, being present at a level of 5–10 vanadium/protein [177]. Much or all of the vanadium in ferritin is present as VIV O2+ and VIV EPR signals are observed in rats fed vanadium-supplemented diets [178]. VIV O2+ was also used as spin probe to establish which specific sites bind the natural substrates Fe2+ and Fe3+ as well as Zn2+ and Tb3+ , known inhibitors of iron deposition. EPR experiments were carried out on horse spleen apo-ferritin (HoSF) and recombinant human H-chain and L-chain apo-ferritins (HuHF and HuLF) [177]. A specific VIV O2+ −apo-ferritin interaction was observed, but in contrast with other proteins for which the VIV O2+ binding suppresses V hydrolysis, for apo-ferritin the hydrolysis decreases the concentration of the metal-protein complex in the pH range 6.0–7.0 [179]. The VIV O2+ –apo-ferritin species has an average stoichiometry of ca. 0.5–0.6 VIV O2+ /subunit, corresponding to 12–16 VIV O2+ ions bound per 24-subunit protein shell [179,180]. The amount of EPR-active VIV O2+ is 61, 36 and 27% at pH 6, 7 and 8, respectively, using a ratio VIV O2+ /protein of 16 [177]. The two distinct coordination environments observed were assigned to ␣ (further composed by ␣ and ␣ signals) and ␤ species; their EPR parameters are gz = 1.940, Az = 173.0 × 10−4 cm−1 and gz = 1.945, Az = 167.0 × 10−4 cm−1 , respectively [179]. Comparison of EPR and optical spectral data for the protein complex with inorganic VIV O-complexes suggested that COO− donors take part in the metal complexation and there is no direct evidence for a vanadium-vanadium interaction. Only the ␣ species show a pHdependence in binding in the pH 6–7 range, while the ␤ species do not. The differences were related to the deprotonation of a H2 O molecule coordinated in the equatorial position in species ␣ to give a hydroxide OH− ion in the ␤ species. The 14 N superhyperfine coupling constant in the range 6.7–7.0 MHz, measured by ENDOR and ESEEM spectroscopy, is characteristic of the coordination of an equatorial imidazole N belonging to a His residue [181,182]. ESEEM data suggest the presence of a H-atom attached to an equatorial donor which can be exchanged with an aqueous solvent [182]; this probably corresponds to the deprotonation of the equatorial water molecule with a pK of ca. 6.5. Competition between Cd2+ and VIV O2+ indicated that Cd2+ may displace VIV O up to 0.3 mol of Cd2+ /subunit [180].

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Fig. 9. Interaction of [VIV O(ma)2 ], [VIV O(dhp)2 ] and [VIV O(acac)2 ] with the red blood cell membrane and their transformation into the cytosol of erythrocytes. Ma, dhp and acac indicate maltolate, 1,2-dimethyl-3-hydroxy-4(1H)-pyridinonate and acetylacetonate anions. Adapted from Ref. [89].

It was initially suggested that VIV O2+ binds in the three-fold channels [180]. Subsequent EPR measurement on HuHF variants of several residues conserved in H- and L-chain ferritins, potentially able to interact with VIV O2+ (Asp131, Glu134, His118 and His128), suggested that His118 near the outer opening of the three-fold channel is probably a ligand for V and is responsible for the ␤ signals. The data on the variants Asp131Ala and Glu134Ala indicated that VIV O2+ does not bind to the Asp131 and Glu134 residues within the threefold channels [177]. In contrast, the spectrum of His128Ala reveals a diminution but not a loss of the ␤ signal, indicating that His128 is not involved in the metal coordination; the mutation at residue His128 – located on the inner surface of the protein shell facing the threefold channel – causes, however, structural changes in the region with His118 which partially affect signals ␤ in the EPR spectrum [177]. This experimental evidence indicates long-range interactions in the protein which cause significant perturbations in the metal binding site.

3. Vanadium for protein activity and function 3.1. Nitrogenases Nitrogen fixation is the biogenic, as well as the non-biogenic, transformation of N2 into nitrogen compounds. The biogenic fixation corresponding to the transformation of N2 into NH4 + , may be carried out by the nitrogen-fixing bacteria (Azotobacter) and cyanobacteria (“blue-green algae”, Anabaena) and by other organisms. The electrons required for the reduction of N2 to NH4 + are provided by the respiratory oxidation of organic carbon to CO2 . The reducing equivalents are supplied via iron proteins, the enzymes catalyzing the processes being designated by nitrogenases (Nases).

The nitrogenases are responsible for cycling about 108 tons of N2 per year from the atmosphere to the soil [19]. The Nases contain two major protein components and each of them is comprised of multiple subunits and/or metal clusters. One protein complex is a Fe-containing protein and another is the heterometallic protein containing either Mo (MoFe, Mo-Nase), V (VFe, V-Nase), or Fe (FeFe). Both bacteria and cyanobacteria can use vanadium in the active center of ‘alternative’ vanadium-dependent Nases for the conversion of N2 into NH4 + ions, H2 being also produced. V-Nases are found in free-living nitrogen-fixing bacteria of the genus Azotobacter (chroococcum, vinelandii and amylobacter) [32]. The enzyme Mo-nitrogenase (Mo-Nase) consists of two components: the homo-dimeric Fe-protein, and the ␣2 ␤2 tetrameric Fe–Mo protein [183]. The Fe-protein contains a single [4Fe–4S] cluster at the interface of the two protein sub-units, and two ATP (adenosine triphosphate) linked to the protein. Electrons, normally supplied by NADH, are transferred to the Fe–Mo protein in processes which are driven by the hydrolysis of ATP. The Fe–Mo protein contains two M and two P clusters. The P cluster is a double cubane with a Fe8 S7 core, consisting of two subclusters [4Fe–4S] bridged by cysteinate, in an oxidation-state dependent overall arrangement. The central core of the M cluster, where the final reduction of N2 takes place, is a Fe7 MtS9 double cubane, where Mt is either Fe (Fe-only Nase), V (V-Nase) or Mo (Mo-Nase, the most common) (Fig. 10) [19]. The topic of Vnitrogenases has been addressed in previous reviews [5,19] and chapters of books [38,184–186], which we recommend for more detailed information. To the best of our knowledge, no XRD structure of a Vnitrogenase is currently known. In the V-Nase, which has a ␣2 ␤2 ␦2 substructure, expressed by Azotobacter, the V replaces Mo in

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Fig. 10. Adapted from Ref. [186]; the iron protein component contains two ␣ subunits, which are bridged by a [4Fe-4S] cluster and the FeV component has a ␣2 ␤2 ␦2 subunit ˚ the distance between the two M clusters is ∼70 A. ˚ The structure, and contains two P clusters at the ␣ and ␤ subunit interface. The P and M clusters are separated by ∼20 A; P cluster is believed to transfer electrons to the FeM-cluster in all three nitrogenases systems. The M cluster in the V-Nase is commonly referred to as FeVco (co, cofactor) and is the proposed binding site for N2 . Two FeVco clusters are located in the ␣ subunits, and can be described as the catalytic cofactors since they are responsible for the conversion of N2 to NH3 . The reducing equivalents are supplied via the Fe-proteins, gated by the energy released by hydrolysis of Mg2+ -activated ATP to ADP (adenosine diphosphate) and inorganic phosphate [19,187].

Mo-depleted environments or at low temperatures [187,188]. The V-Nase is both genetically and biochemically similar to the MoNase. It contains P cluster redox centers and a catalytic FeVco center, in which V is in a polynuclear cluster with Fe, S and homocitrate with a chemical environment similar to Mo in Mo-Nases. Despite biochemical similarity, the V-system does not arise from the simple substitution of V for Mo in the Mo-protein. EPR and XAS/EXAFS investigations indicated that FeVco is similar to FeMoco, yet distinct from this latter in electronic properties and structural topology, this probably also accounting for the differences in the reactivity of the two cofactors [187]. In fact, the VFe-protein and the MoFe-proteins are biochemically similar but they possess different substructures: ␣2 ␤2 in MoFe-protein and ␣2 ␤2 ␦2 in VFe-protein. The ␣ subunits in V-Nases contain the M cluster (also designated as FeVco). The central core of this cluster, where the final reduction of N2 takes place, is a double cubane of composition (VFe7 [␮6 -C]S9 ) and the active center – vanadium, which is bound to a histidine and the vicinal hydroxide and carboxylate of homocitrate [187] (Fig. 10). This M cluster is responsible for the reductive protonation of N2 to NH4 + , coupled to some hydrogenase activity and several additional reductive protonations. For the Azotobacter vinelandii, the Fe-V cofactor was characterized by XAS/EXAFS [187]. The EPR recorded for V-nitrogenases are complex but a spin state of S = 3/2 was associated to vanadium. The V-Nases A. chroococcum [189] and A. vinelandii [190] were studied by X-ray absorption near edge structure (XANES) in the EXAFS region and the data considered consistent with VII or VIV in a distorted octahedral coordination environment. On the other hand, from the synthesis of clusters modeling nitrogenases it was concluded that the oxidation state of vanadium would be VIII [191], although the assignment of oxidation state in these types of clusters is difficult because of electron delocalization. The active center of the nitrogenase contains a cluster of composition X ⊂ Fe7 VS9 (the M cluster or FeVco) with vanadium bound to histidine and homocitrate, and to an adjacent iron through three ␮3 -S2− . X is probably ␮6 -carbide (C4− ), with the methyl group of adenosyl-methionine as the source [192]. An interesting observation [193] was that the Azotobacter enzyme in a total atmosphere of CO and with no nitrogen present, is able to reduce CO with the production of C2 H4 , C2 H6 and C3 H8 mimicking the Haber–Bosch and the Fischer–Tropsch syntheses. As the reduction of both N2 and CO are industrially important processes, this makes V-Nases as prospectively applicable in the future.

The enzymes carry out their function under anaerobic conditions and developed sophisticated mechanisms to exclude oxygen from the active site and the redox active cofactors. V-Nases are less stable and less efficient than Mo-Nase because the reduction equivalents used for N2 reduction are 50%, whereas for Mo they are 75% [193,194]. 3.2. Haloperoxidases Haloperoxidases are enzymes that catalyze the two electron oxidation of halides, by hydrogen peroxide (see Eq. (1)), with not much substrate specificity. This reaction may also be regarded as an oxygen transfer reaction from the peroxide to the halide. The hypohalous acids (HOX) and related halogenating intermediates, such as OX− , X2 , or X3 − , are produced during turn over and react non-enzymatically and nonspecifically with a variety of organic compounds (RH) (Eq. (2)), producing halogenated compounds RX. H2 O2 + H+ + X− → HOX + H2 O

(1)

HOX + R-H → RX + H2 O

(2)

In the absence of the nucleophilic acceptor RH, HOX may react with H2 O2 this resulting in the formation of singlet oxygen (1 O2 ) (Eq. (3)): H2 O2 + HOX → 1 O2 + H2 O + HX

(3)

There are three classes of haloperoxidases known, either cofactor free, heme-dependent or with vanadium as a relevant entity in the active site, the vanadium-containing haloperoxidases (VHPOs) [5]. The naming of the VHPOs is based on the most electronegative halide that the enzyme can oxidize: the chloroperoxidases (VCPO) can catalyze the oxidation of Cl− , Br− and I− , the bromoperoxidases (VBPO) can catalyze the oxidation of Br− and I− , and iodoperoxidases are specific for I− . However, e.g. bromoperoxidases can also oxidize Cl− but much less efficiently. In the absence of halides, the VHPOs are able to catalyze the enantioselective sulfoxidation of organic sulfides. The discovery of VBPO in 1984 [195], and later VCPO, the first enzymes using vanadium as a cofactor, gave rise to an enormous increase of interest in vanadium chemistry and biochemistry. Moreover, the practical applications of VBPO and VCPO in the halogenation of organic substrates under mild conditions raised the

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interest of the chemical industry; the enzymes catalyze the conversion of X− which are strong nucleophiles, into the very electrophilic “X+ ” (where “X+ ” is in the form of HOX, X2 , etc.) followed by nonenzymatic halogenation of the organic substrate. Importantly, the catalyzed reaction is the major source of halogenated organic natural products namely with antibiotic activity. Since the discovery of VHPO enzymes, thousands of studies of structural, spectroscopic or functional modeling of VHPOs have been published as well as multiple reviews on various aspects of this area. We refer the readers namely to the reviews of Wever [196–198] Vilter [199], Rehder [38,200], Crans et al. [5], and to the more recent ones from Wever [201], Licini et al. [202], Plass [29], Hartung [27] and several other miscellaneous publications [203–206]. It is known that upon addition of H2 O2 to solutions containing monovanadate(V), the peroxido may bind to VV , thereby increasing its coordination number, forming mono- and/or diperoxido vanadate(V)s, with distorted pentagonal bipyramidal geometries. This suggests that they are not good inhibitors for sites with trigonal–bipyramidal transition states, such as those present in many phosphatases [207]. However, peroxides are more potent oxidizing agents than vanadate(V), and this is a relevant characteristic in the action of VHPOs. In sea water, at the pH where marine bromoperoxidases have maximum activity (pH 6–7), vanadate(V) exists mainly as H2 VO4 − , which shows strong affinity to bind the enzyme at the remote imidazole nitrogen of a His residue, thereby providing the active site for Br− oxidation. The vanadate(V) cofactor and proximate amino acids provide a 3D structure for setting water molecules into a supramolecular network to facilitate H2 O2 access and binding for subsequent Br− oxidation. From a chemical point of view, Br− oxidation mediated by H2 O2 at pH 6–7 yields HOBr. In an aqueous solution containing Br− , such as sea water, hypobromous acid yields an equilibrium mixture of HOBr, Br2 and Br3 − [27,208]. According to the general mechanistic scheme for bromoperoxidase-catalyzed oxidation, carbon–bromine bond formation occurs in a non enzymatic reaction between an organic acceptor and HOBr, Br2 , or Br3 − [209], with bromoperoxidases depicting little to no organic substrate specificity. The first XRD structure of a vanadium-containing haloperoxidase was determined in 1996 by Messerschmidt and Wever at 2.1 A˚ resolution (PDB: 1VNC) [210]. It corresponds to a chloroperoxidase from the fungus Curvularia inaequalis (CiVCPO) where V is bound to His496, three O-atoms and to an azide ligand (used during the crystallization trials) in a trigonal–bipyramidal geometry (Fig. 11). The enzyme is a monomer and consists mainly of ␣-helices with two four-helix bundle motifs. The vanadate(V) is deeply buried in the protein, accessible via a highly positively charged funnel-shaped channel. Following this first example, several other structures of CiVCPO were solved either in the apo form (PDB: 1IDQ) or in the presence of peroxide (PDB: 1IDU) [211] (Fig. 12) allowing a better understanding of its mode of action. During the catalytic reaction, V adopts a reasonably stable five-coordinated state upon binding of H2 O2 (Fig. 12) which, in turn, is attacked by the halide. The hypohalide species formed is very susceptible to nucleophilic attack producing halogenated compounds and singlet oxygen (Eq. (3)). Probably the halide never strongly binds to the metal (see below). Furthermore, heterologous expression of the apo (PDB: 1VNS) and holo-enzyme (PDB: 1VNI) in yeast proved to be also successful allowing for single site mutagenesis studies [212], and establishing the role and the importance of some of the residues in the active site. Alanine scanning mutagenesis revealed that: (i) in the absence of His496 (PDB: 1VNH) the enzyme does not bind V; (ii) His404 (PDB: 1VNG) activates the axial OH/OH2 ligand of the metal prior to hydrogen peroxide attack; (iii) Arg360 (PDB: 1VNF) and Arg490

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Fig. 11. Structural representation of the adduct formed between the CiVCPO and the vanadate(V) moiety (PDB: 1VNC). The vanadium atom is bound to the His496 residue and to an azide ligand from the crystallization conditions. The residues hydrogen ˚ bonded to the vanadate(V) moiety are also shown (bond lengths are indicated in A) [210].

compensate the negative charge of the cofactor; (iv) Asp292 (PDB: 1VNE) orients Arg490 through salt bridge interaction. Lys353 is also part of the active site and is responsible for the activation of the Vbound peroxide through charge separation enabling nucleophilic attack of the halide [212]. As mentioned, the peroxido intermediate of the vanadium peroxidase was also structurally characterized [211] (Fig. 12). The vanadium coordination geometry is that of a distorted tetragonal pyramid with only four O-atoms and one N-atom, and the peroxide is bound side-on with a distance of 1.47 A˚ between the two peroxido O-atoms. There are a few changes in the hydrogenbonding network when the peroxido is bound. His404 is no longer bonded to any of the vanadate(V) O-atoms, but there are still two water molecules in the active site. Lys353 forms a hydrogen bond to one of the O-peroxido atoms, polarizing the O O bond (see Figs. 11, 12 and 16). This charge separation is very important in the catalytic process (see below). Several bacterial VCPOs have also been identified along the years although only VCPO from Streptomyces sp. CNQ525 (named S525-VCPO) was structurally characterized both in the apo (PDB: 3W35) and holo form (PDB: 3W36) at 2.4 A˚ and 1.97 A˚ resolution, respectively. A manuscript describing the structure is not available yet but superposition of the crystallographic models with CiVCPO (Fig. 13) shows that the enzymes are very similar, with the same overall fold, although S525-VCPO is proposed to be a homodimer in solution. In the bacterial enzyme, a vanadate(V) anion is also found in the active site and is involved in interactions similar to those previously described. However, a major difference exists, probably with important implications for the reaction mechanism. In S525-VCPO, the His404 residue (in the CiVCPO numbering) is replaced by a serine (Ser427 in the S525V-ClPO numbering) (Fig. 14). This residue is at hydrogen bonding distance to the axial OH/OH2 ligand of vanadium, but its nature and electrochemical behavior hinders the allegedly charge separation of the peroxido when bound to the metal. If this difference alone accounts for the

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Fig. 12. Structural representation of the adduct formed between the CiVCPO and the vanadate moiety in the absence (right, PDB: 1IDQ) and in the presence (left, PDB: 1IDU) of hydrogen peroxide [211]. It is possible to observe the change in the vanadium coordination due to the binding of peroxide.

Fig. 13. Superposition of CiVCPO (gray, PDB: 1VNC) [210] with one molecule in the asymmetric unit of S525-VCPO (orange, PDB: 3W36) [213]. The proteins present a similar structural organization namely by the presence of several ␣-helices.

stereoselectivity observed in the chlorination-cyclization of napyradiomycin antibiotic biosynthesis is yet to be disclosed [213]. Three XRD structures of V-bromoperoxidases (VBPOs) from red and brown algae have been also determined and are structurally very interesting: Corallina officinalis (CoVBPO, PDB: 1QHB) [214]; Corallina pilulifera (CpVBPO, PDB: 1UP8) [215] and Ascophyllum nodosum (AnVBPO, PDB: 1QI9) [208]. Even though the sequence homology is low (<30%) and the oligomeric states vary between homodimers (in AnVBPO) and homododecamer (in CoVBPO and CpVBPO), the enzymes share the same structural motif as CiVCPO (Fig. 15). The arrangement in the active site is also akin in the two families of enzymes and so is the suggested reaction mechanism. The first structure of a bacterial V-iodoperoxidase from Zobellia galactanivorans was released recently: PDBs 4USZ (native at 2.0 A˚ resolution) and 4CIT (Se-Met at 1.8 A˚ resolution) [216]. Not surprisingly, the overall structure is quite comparable to the structures of VCPOs and VBPOs. A vanadate moiety sits in the active center in a similar organization to that observed in the CiVCPO (namely the covalent bond to a histidine residue, His416). Again, site-directed mutagenesis was used to elucidate the role of different residues in the protein catalytic activity [216].

Fig. 14. Structural representation of the adduct S525-VCPO-vanadate(V) (orange, PDB: 3W36) superposed with the respective adduct in the CiVCPO structure (gray, PDB: ˚ The His404 residue present in the CiVCPO structure 1VNC) [210]. The vanadium atom is located next to a His residue (His494 or His496) in both proteins (bond lengths, in A). is replaced by a serine (Ser427) in the S525-VCPO structure.

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Fig. 15. Structural representation of the adduct AnVBPO-vanadate(V) (orange, PDB: 1QI9) [208] superposed with the respective adduct in the CiVCPO structure (gray, PDB: 1VNC – to allow a better visualization the azide ligand is not shown) [210]. The vanadate(V) is placed in the same site and the vanadium atom is coordinated to His residues (His486 in AnVBPO and His496 in CiVCPO) in both structures. Some bond ˚ are indicated. lengths, in A,

As mentioned, common to this enzyme family is the low substrate specificity since chloroperoxidase activity has been observed in VBPO and vice versa. They can even accommodate tungstate instead of vanadate(V) although in the case of W-CiVCPO, the distance between His496 and the metal is very long suggesting weak or no covalent binding [217]. Moreover, apo-haloperoxidases may have phosphatase activity as the vanadate(V)-binding site can also easily harbor phosphate (see below). 3.3. Mechanism of halide oxidation A considerable number of experimental [218–220] and theoretical [205,221–223] studies have led to a significant advance in the understanding of the catalytic cycle of VHPO enzymes, although some details of the vanadate(V) moiety in the active site (namely its protonation state, the protonation sites and O-atom involved in the attack of the halide) are not totally settled. The proposed catalytic cycle by VHPOs of the oxidation of halides is outlined in Fig. 16. In the presence of H2 O2 and its binding to vanadium (step c), due to its Lewis acid characteristics, electron density is withdrawn from the bound peroxide towards the V center, presumably also assisted by two positively charged arginine residues (not shown in Figs. 11, 12 and 15). The conserved Lys that forms a hydrogen bond to one of the O-atoms of the peroxide further polarizes and activates the bound peroxido (steps c, d, e). One of the O-atoms of the peroxido is protonated to allow oxidation of X− . After formation of the protonated peroxido intermediate, the next step is the nucleophilic attack of the X− on the electrophilic hydrogen peroxide (Fig. 16). The breaking of the peroxido bond occurs (step f), resulting in the formation of HOX or XO− . If the VHPO involved is either a chloroperoxidase or bromoperoxidase, the peroxido protonation is considered not a requirement [201,204,205]. Additionally, some authors state that the halide attack, thus the oxo-transfer reaction, takes place at the protonated O-atom of the peroxido [201], while others consider that it involves the nonprotonated axial O-atom of the peroxido [204]. In model systems of

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sulfoxidations, the transition states corresponding to the attack of the sulfur atom have also been theoretically predicted to take place at the non-protonated axial O-atom of the coordinated peroxide [204]. Detailed structural data are now available to allow the explanation of the difference in reactivity between the vanadium iodo-, bromo- and chloroperoxidases and the effects of mutations on the activity. This issue was discussed by Wever in [201]. Some of the main roles of vanadium in biological systems are associated with the analogy between vanadate(V)s and phosphates. The amino acid sequence of the active site of VHPOs is conserved within three families of acid phosphatases, including several lipid phosphatases, the mammalian glucose-6-phosphatase and bacterial nonspecific acid phosphatases [224–226]. Structural similarities also exist between the VHPOs and phosphatases [224–230]. The amino acids that are involved in phosphate binding in the acid phosphatase enzymes and also those that are coordinated to vanadium in the VHPOs are also conserved [201,231]. Vanadate(V) as the active site species of VHPOs is covalently bound to a His residue, but the reactivity is in fact determined by the existence of an extensive hydrogen-bonding network. In the case of e.g. the rat prostatic acid phosphatase, the motif is very similar and His12 is the residue bound to vanadate(V), as discussed in Section 4.1 [232]. Since similar supra-molecular environments are present in the two classes of enzymes, it is likely that they exhibit dual enzymatic activities [5]. Phosphatase activity may indeed be exhibited by apohaloperoxidases [5] and the possibility that peroxidase activity could be observed for vanadate(V)-inhibited phosphatases has also been investigated for several systems [224,226,227,231,233,234]. The first observation of chloroperoxidase activity was reported with phytase, upon addition of vanadate(V) and H2 O2 [5,233], and there were several other reports demonstrating that other phosphatases can act as peroxidases in the presence of VV and H2 O2 [224,226,227,231,233,234]. When vanadate(V) is substituted in the active site of acid phosphates they exhibit VHPO as well as enantioselective sulfoxidation activity. Thus, the analogy between these classes of enzymes encompasses both structural and catalytic aspects. However, the turnover is much lower than that of native bromoperoxidase from A. nodosum. When vanadate(V) is substituted in the phosphatase phytase, sulfoxidation [233,235] and bromoperoxidase [233] activity are also observed. Nevertheless, phytase does not belong to the class of acid phosphatases and the active site of phytase has an architecture that differs from the acid phosphatases. 3.4. Nitrate reductases Rehder [236], in a review focusing the knowledge of the interplay of bacteria and other primitive forms of life (cyanobacteria, algae, fungi and lichens) with vanadium, critically discussed this topic, and we recommend it for more details. Nitrate reductases are enzymes that commonly catalyze the two electron reduction of nitrate to nitrite (Eq. (4)). They normally contain Mo as a constituent of a cofactor designated by molybdopterin, in which Mo (with oxidation states varying between IV and VI) is coordinated to a dithiolene moiety of a tetrahydropterin derivative. Nitrate reductases lacking the Mo cofactor and containing vanadium, were isolated and characterized from the bacterium Thioalkalivibrio nitratireducens [237]. The corresponding enzyme from T. nitratireducens is a homotetramer of molecular mass 195 kDa, which contains vanadium and iron in a molar ratio 1:3, with a type c heme group identified by UV-visible spectroscopy. NO3 − + 2e− + 2H+ → NO2 − + H2 O

(4)

In vitro, this enzyme also promotes the reduction of other substrates, e.g. nitrite, bromate, selenate, and it further exhibits

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Fig. 16. Proposed catalytic cycle for the oxidation of halides by vanadium haloperoxidases (adapted from Ref. [205]). The role of the amino acid residues Lys353, Ser402 and His404, His496 (bound to V) is emphasized. Two positively charged arginine residues are not shown; Arg360 (PDB: 1VNF) and Arg490 compensate the negative charge of the cofactor; Asp292 (PDB: 1VNE) orients Arg490 through electrostatic interactions.

haloperoxidase activity, although it is not yet clear if this haloperoxidase activity is associated with heme-c, or with the presence of vanadium [236]. The pterin cofactor is also lacking in the nitrate reductase from the facultative anaerobic bacterium Pseudomonas isachenkovii. The enzyme from this source does not contain a heme-c, presupposing that V is directly coordinated to side-chain functions of the protein, e.g. a N-atom from a His residue as in the case of VHPOs. As emphasized by Rehder [236], P. isachenkovii, when grown in a culture medium containing lactate as an electron donor and VV as primary electron acceptor, secretes a 14 kDa protein into the culture medium which coordinates up to 20 vanadium ions per monomer [238], resembling, in this respect, the vanabins, and possibly classifying it as a storage protein. The presence of a periplasmatic V-containing nitrate/ vanadate(V) reductase, on the other hand, appears to indicate that P. isachenkovii also employs vanadium in its dissimilatory functions. The reductase, a tetramer with a molecular mass of 220 kDa, reduces nitrate to N2 . Vanadate(V) reduction starts only when denitrification has been almost complete. Moreover, for Shewanella, the reduction of VV does not depend on the induction of specific proteins [239].

4. Vanadium as substrate analogue or inhibitor There are many classes of proteins where vanadium(IV) behaves as competitive binder of other metal ions e.g. of Ca2+ , Mg2+ , Mn2+ , Zn2+ . VIV O2+ has been used frequently as a spin probe and the complexes formed studied by EPR and/or by ESEEM spectroscopy. Some of these studies are mentioned throughout this review; for more detailed discussions, we recommend the reports from Chasteen [93,106]. There are also many classes of proteins (in most cases enzymes) where vanadium, mainly as vanadate(V), behaving as a substrate analogue or an inhibitor, has contributed either to allow easier crystallization of VV –protein complexes or to the elucidation of the catalytic reaction mechanism. In many cases, 51 V and 1 H NMR spectroscopy were used for this purpose. These enzymes can be grouped as those mainly involved in phosphate binding or release (Sections 4.1–4.6), proteins responsible for DNA binding (Section 4.7) and RNA binding (Section 4.8). In Table 5, we depict a summary of the function and/or of the reactions of the proteins discussed in this section as well as a summary on the proteins already addressed in the review.

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Table 5 Summary of function and/or reaction of the proteins discussed in this review. Family of protein

Function/reaction

PDB codes for V-protein interactions

Vanabins Nitrogenases Haloperoxidases

Vanadium storage/carrier 16 ATP + 8 H+ + 16 H2 O + N2 + 8 e− → 2 NH3 + H2 + 16 ADP + 16 Pi H2 O2 + X− + H+ → H2 O + HOX (X = halogen)

V-Nitrate reductases Phosphatases

NO3 − + NADH → NO2 − + NAD+ + OH− aa-OPO3 2− + H2 O → aa-OH + Pi

Kinases Other Transferases

ATP + substrate-OH → ADP + substrate-OPO3 2− Transfer of functional groups between different molecules

1VFI – 1VNC, 1IDQ, 1IDU, 1VNI, 1VNH, 1VNG, 1VNF, 1VNE, 3W36, 1QI9, 4USZ, 4CIT – 4KKZ, 2D1G, 3F9B, 1B8J, 1H2F, 3ZWK, 4QIH, 4HGP, 4ERC, 3ZX5, 4KNW, 3QKQ, 3ZWU, 3QCC, 3QCD, 3S3F, 3OMX, 3I7Z, 3I80, 2I42, 2HY3, 2I4E, 1Z12, 1RPT, 1J9L 1M7G, 3GQI, 1DKT, 4DZ6, 3Q8Y 1UZI, 1RXS, 1C4G, 4HGO, 3E81, 2RAR, 2RBK, 2AZD, 1BO6

EctoNTPDases

4BRE, 4BRH, 4BRL, 3ZX2, 3QVF

Phosphodiesterases

1JH7, 2GSO, 1RFF, 1RFI, 1RG1, 1RG2, 1RGT, 1RGU, 1RH0, 1NOP, 1MU9

Phosphomutases ATPases

HO-substrate-OPO3 2− → PO4 2− -substrate ATP + H2 O → ADP + Pi

ATP synthases Topoisomerases Ribonucleases Ribozymes DNA binding (aprataxin) (Protelomerases)

ADP + Pi → ATP Regulation of the overwinding or underwinding of DNA Degradation of RNA Catalysis of biochemical reactions by RNA Removal of AMP from DNA ends Generation of closed hairpin ends in linear DNA

4.1. Phosphatases Phosphorylation is one of the major signaling pathways and is commonly observed in all organisms. Kinases and phosphatases are the key enzymes in this reversible reaction, where addition or removal of a phosphate group is achieved. The enzymes activate or inhibit intracellular signaling pathways triggering a cascade of different physiological effects. In biological systems, the level of phosphorylated proteins is a balance resulting from the action of kinases and phosphatases, thus both types of enzymes have important roles in the regulation of cellular processes. Protein phosphorylation mainly occurs in hydroxyl-containing side chains of amino acids such as Ser, Thr and Tyr; de-phosphorylation is achieved by hydrolysis of the ester bond with the formation of a high-energy five coordinate phosphorous transition state. The phosphatases that hydrolyze phosphoproteins may be classified in two groups: (i) those with affinity for serine-threonine phosphate proteins (PS/TPases or PSPases) and (ii) those with affinity to tyrosine-phosphate proteins (PTPases). All of them catalyze the hydrolysis of phosphate esters. The PTPases have a cysteine residue at the active site and in their catalytic mechanism a phosphocysteine is formed as an intermediate. Moreover, these enzymes are structural and mechanistically different from the acid and alkaline phosphatases as well as those hydrolyzing phosphateesters of small molecules such as glucose-6-phosphate, discussed in the following sections [240]. Vanadium compounds can oxidize this cysteine residue e.g. by the formation of free radicals, regulating in that way many biological events [241–243]. Phosphatases can also be divided according to the nucleophile present in the active site responsible for the dephosphorylation and involved in the phosphate-phosphatase adduct formation. In this

1E59, 3GW8, 3GP5 1L7V, 3PUV, 3B5Z, 2XEL, 2XO8, 2X9H, 3MJX, 3MNQ, 2JHR, 2JJ9, 4BYF, 4AE3, 2YCU, 4DBR, 4E7S, 4E7Z, 3MKD, 3MYH, 3BZ7, 3BZ8, 3BZ9, 2V26, 1YV3, 1VFZ, 1QVI, 1LKX, 1DFL, 1VOM 2F43, 3P20 3IGC, 2B9S, 3MGV, 2XEL, 2XO8, 2X9H 1RUV, 6RSA, 2G8H, 3RNT 1M5O, 2P7E 4NDG 4E0G, 2V6E

regard, four families have been identified, namely alkaline phosphatases, which have serine in the active site, acid phosphatases, which have histidine in the active site, cysteine based phosphatases and metal-dependent phosphatases, where a dinuclear M1/M2 center is present. In the corresponding vanadate(V)–protein complex, besides the distinct amino acid residues involved in the metal binding (e.g., Ser, His, Cys), the binding modes may be either 4- or 5-coordinate as well. The alkaline phosphatases (AP) are membrane bound glycoproteins codified by different genes and classified as specific tissue phosphatases and non-specific tissue phosphatases. These enzymes hydrolyze phosphate monoesters from small molecules and proteins and catalyze the transfer of phosphate to hydroxyl groups of organic molecules. The mechanism of action occurs through the phosphorylation of the serine residue of the active site, followed by the transfer of the phosphate to a water molecule or to an organic acceptor [244]. Acid phosphatases (AcP) also hydrolise monoester phosphate bonds from a broad variety of molecules but, unlike the previous group, under acidic conditions. In humans they are classified as prostatic, lysosomal, macrophage, erythrocytic and osteoclastic and are widely used as serological and histological markers of disease [245]. In general, the histidine at the active site is phosphorylated by the substrate and the phosphate group is further released, restoring the enzyme in the active form. The reaction mechanism of hydrolysis carried out by phosphatases involves the formation of 5-coordinate high-energy transition state. These enzymes are generally inhibited by vanadate(V) (and some other oxometalates), which is often considered to act as a transition state analogue (TSA) of the phosphatasecatalyzed reaction (Fig. 17). The ability of vanadium to inhibit these

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Fig. 17. Probable catalytic intermediates depicting the vanadate transition state analogs (TSAs) formed, similar to possible transition states corresponding to phosphate ester hydrolysis. The lower apical linkages depicted correspond to either side groups of Ser, Thr or Tyr residues (left), or to side groups of His, Lys or Arg residues (center), or to side groups of Cys (right).

enzymes is again closely related to the analogous physicochemical properties of vanadate(V) and phosphate (see above), and the generation of stable complexes between the vanadate(V) and the enzyme at the active site has been considered the reason for its inhibitory effect to the enzyme activity. Vanadate(V) is not a specific inhibitor of all phosphatases, but can be a potent inhibitor of such activity [207]. Vanadate(V) may indeed mimic the 5-coordinate transition state of phosphate formed during the phosphatase catalytic cycle, by its ability to form a stable 5-coordinate complex at the active site of the enzyme. However, it is also possible that vanadate(V) might inhibit these enzymes by forming some other type of covalently bound enzyme complex. Vanadate(V) may also cause Cys oxidation at the active site of cysteine. Several PTPases require thiol-reducing agents for optimal activity [246–248] and there are enzymes where the thiol group is not oxidized, but vanadate(V) still inhibit their activity. The VIV O2+ cation also inhibits different enzymes and in some cases with a greater potency than vanadate(V), thereby ruling out the simple explanation that the inhibition by VIV -compounds is preceded by oxidation of VIV to VV . One proposed mechanism for this inhibition is the possibility of VIV to form a VIV -anion structurally similar to monovanadate(V) (see above), thus able to adopt a trigonal–bipyramidal geometry when binding at the active site and inhibit the activity of the enzymes [5,207,249]. Further supporting this possibility, it is known that in dilute neutral aqueous solution at pH ∼ 7, VIV O2+ exists mainly as anionic [VIV O(OH)3 (H2 O)n ]− [250,251]. Therefore, both speciation considerations (in dilute neutral aqueous solution at pH ∼ 7 [VIV O(OH)3 (H2 O)n ]− and monovanadate(V) are expected to predominate) and the structural similarity of VIV - and VV -anions, help explaining the potent phosphatase inhibition observed for aqueous VIV O-solutions [5]. These observations also reinforce the idea that the structural details of the vanadate(V) binding at the active site of these enzymes have some degree of freedom. The first crystal structure of a phosphatase was determined in 1980s by Wyckoff and co-workers at 2.8 A˚ resolution [252]. ˚ strucLater, the same authors reported a higher resolution (2.0 A) ture [253]. Currently there are over 1690 entries in the PDB of phosphatases from eukaryotes. In 1994, the crystal structure of the human PTP1B was determined (PDB: 2HNP): a cysteine-based phosphatase that effectively removes phosphate from tyrosine phosphorylated proteins (PTPs). The structure was a landmark in the field of phosphatases representing the first tyrosine phosphatase to be structurally characterized [254]. In fact, a tungstate PTP1B derivative was used to solve the crystallographic phase problem and the ion binds at the active site of the enzyme, sitting in the substrate binding site (PDB: 2HNQ). This active site is located within a crevice at the molecular surface of the protein, containing the nucleophilic cysteine (Cys215) and the phosphate-binding loop (P-loop) [254]. The Arg close to the catalytic cysteine is very important for the two-step mechanism, and, even though it is not directly

involved in bond breaking or formation, it stabilizes the phosphoenzyme complex of the intermediate step and its mutation results in complete loss of activity [12,240]. It is well accepted that during the dephosphorylation process, the cysteine of the active site is in the thiolate state and is very susceptible to oxidation. This can occur by reaction with neighboring peptide backbone atoms, inducing important conformational changes in the active site or by hydrogen peroxide. In both cases the outcome is the inactivation of the enzyme. PTPs have been implicated in inflammation, atherosclerosis, diabetes and cancer. This family of enzymes is considered a good target for cancer therapy as they are usually overexpressed in tumor cells. Design of selective PTP inhibitors is a challenging task since their active site is quite conserved and PTPs are able to dephosphorylate a large variety of tyrosine-phosphoryl bonds, independently of the overall structure of the substrate protein. Vanadate(V) has been widely used for studying the reaction mechanism of this class of enzymes. Even though some authors claim that it is not a true substrate analogue [255,256], several crystal structures of vanadate(V)–PTP complexes have been determined providing important information regarding transition state conformation and structural determinants for catalysis [12,256–259]. For PTP1B, the two relevant TSAs have been obtained by crystallizing the enzyme with a short Tyr-containing peptide that was previously incubated with vanadate(V) (different ratios of vanadate(V) and peptide were used to obtain the two TSAs). The models are considered fair structural TSAs of the reaction mechanism (Fig. 18) [12]. In the first TSA the vanadate(V)–tyrosyl peptide is bound to the active site (PDB: 3I7Z) similar to the phosphorylated tyrosine substrates (PDB: 1PTU) [260]. The vanadium adopts a trigonal–bipyramidal geometry with the nucleophylic cysteine and the tyrosyl oxygen in apical positions. The remaining O-atoms of the complex are hydrogenbonded to amide groups of the flexible P-loop. The second TSA corresponds to the Cys-bound vanadate(V) species (PDB: 3I80) [12], similar to the phosphoenzyme (PDB: 1A5Y) [261] prior to inorganic phosphate release [240]. These models represent snapshots of the reaction mechanism of PTP. The same authors reported the binding of a divanadate complex to a protein molecule. In the structure of the Yersinia PTP YopH, the oxoanion exhibits a double distorted trigonal–bipyramid containing a cyclic [VO]2 core [256]. Structural characterization of alkaline phosphatases (AP) was also achieved in the presence of vanadate(V) [262]. In 1999, a 1.9 A˚ resolution structure was determined where a five-coordinated vanadium ion was located in the active site of Escherichia coli alkaline phosphatase (PDB: 1B8J). The position of V was compared with that of the structure obtained using inorganic phosphate [253] and, in both cases, the ions are located between the metal site 1 (M1) and metal site 2 (M2). Ser102 had been previously suggested to be responsible for the nucleophilic attack at the phosphorous center due to its proximity towards the phosphate ion and its coordination to M2. In the V coordination sphere, a water molecule that coordinates M1 occupies one of the axial positions, while Ser102 occupies the other one. The vanadate(V) is mimicking the 5-coordinate transition state. In fact, the vanadate(V)–AP complex structure revealed to be a clear intermediate between the phosphorylated enzyme complex (E–P) and the enzyme–inorganic phosphate complex (E–Pi) [262]. Acid phosphatases have also been structurally described and vanadate(V) complexes have been determined with this family of enzymes allowing the characterisation of the intermediate states of the reaction. The XRD structures of rat prostate acid phosphatase bound to vanadate(V) (and molybdate) were determined (PDB: 1RPT) [232]. The 3 A˚ model shows the vanadate(V) anion with a trigonal–bipyramidal geometry, coordinated by His12 in the apical position previously implicated in the hydrolysis [263]. The structure, together with the mutagenic assays, was a breakthrough in the

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˚ (A) In the first TSA (PDB: 3I7Z) [12], the vanadate(V) moiety adopts a Fig. 18. Structural TSA models of the reaction mechanism of PTP1B. Some distances are indicated in A. trigonal–bipyramidal geometry with Cys215, the tyrosine of the peptide DADEYL (a fragment of the epidermal growth factor receptor) and three O-atoms. These atoms are hydrogen bonded (not shown) to the backbone N-atoms of the P-loop (residues 215–222) and to one N-atom of the side chain of Arg221. This residue is hydrogen bonded to the backbone carbonyl of Trp179 allowing Asp181 to be positioned next to the oxygen of the leaving group. Gln262 is not interacting with the vanadate(V) moiety. (B) In second TSA (PDB: 3I80) [12], the structure simulates the protein prior to the inorganic phosphate release; the vanadate(V) moiety remains in a trigonal–bipyramidal geometry. Gln262 is rotated when compared with the first TSA being hydrogen bonded to one of the O-atoms of the vanadate(V).

understanding of this class of phosphatases. Furthermore, hydrogen bonds between vanadate(V) O-atoms and His257 and Asp258 suggest that the former is involved in the stabilization of the negatively charged transition state intermediate while the latter is assuring protonation of the substrate during reaction mechanism. Acid phosphatase from the bacterial pathogen Francisella tularensis was also crystallized in the presence of vanadate(V) (PDB: 2D1G) [264]. The anion is bound in the active site, 5-coordinated to the nucleophilic Ser175 in the apical position and four more Oatoms, hydrogen bonded to side chain residues of the active site trough. Next to the vanadate(V), a second metal, interpreted as Ca2+ , was located in the electron density maps, also coordinated to Ser175. The Ca2+ adopts an octahedral geometry, coordinated to several side chains and to one of the equatorial O-atoms of vanadate(V). The location of the vanadate(V) in the active site and the presence of several water molecules in the large trough resemble the binding site of the enzyme substrates. VV -compounds are compatible with acid and alkaline phosphatases and many of them act as enzyme inhibitors. In some cases, loss of ligand may occur, but in others the inhibition is associated to the intact V-compound [5]. VIV -compounds are also able to inhibit phosphatases [5,34]. Vanadium compounds have also been used as models for phosphate inhibition; most of these are 5-coordinate complexes that have distorted trigonal–bipyramidal or square–pyramidal geometries [5]. Although the distortion from the perfect trigonal–bipyramidal geometry may diminish the structural analogy with the phosphate ester hydrolysis transition state, all V-compounds tested were inhibitors for alkaline and acid phosphatases [87,265–271] emphasizing that a perfect transition state geometry is not necessary for potent inhibition of phosphatases. In two recent reviews McLauchlan et al. [34] emphasized that a trigonal–bipyramidal geometry is normally anticipated and has been demonstrated for the vanadate(V) binding to phosphatase active sites. Moreover, it might also have happened that several authors made assumptions restricting the geometry refinement, namely imposing a VO3 plane in the final calculation, likely introducing bias in the final geometry of the vanadate(V)–protein complex [34,35]. In the above mentioned studies, the authors compiled the known structural information for 5-coordinate vanadium compounds coordinated with four O atoms and one X atom, VO4 X, X being O (from Ser), N (from His), S (from Cys) and others, comprising phosphates, haloperoxidases, phosphorylases, ATPases. Both square–pyramidal and trigonal–bipyramidal

geometries can support the transfer of the phosphoryl group in phosphatases and other phosphorylases, but, in the reported structures, there are much more examples of trigonal–bipyramidal complexes in the active site than square–pyramidal, while for small molecule complexes the square–pyramidal geometry is more common [34,35]. Thus, although both types of geometries are found in protein–vanadium complexes, apparently most proteins prefer the trigonal–bipyramidal geometry, a relevant conclusion but probably not very surprising, at least for phosphatases, as the role of most of these enzymes is indeed to support the highly energetic 5-coordinate phosphate transition state.

4.2. Transferases and kinases Transferases are a family of proteins capable of transfering functional groups between molecules. There are different subclasses of transferases depending on the functional group transferred and, among these subclasses, kinases are one of the most important because these enzymes, responsible for the phosphorylation reactions, are involved in several biological processes. Multiple investigations, including structural approaches, focused on kinases namely their use as a target for inhibitors against different types of cancer [272,273]. The relevance of vanadium in these studies has again been related to its similarity to phosphate. A first structure of a protein related to kinases was released in 1996: a 2.9 A˚ structure of a human regulatory subunit (CksHs1) of the cyclin-dependent kinases complexed with vanadate(V) (PDB: 1DKT) [274]. However, the first kinase vanadium-containing XRD structure was deposited in the PDB only in 2002: a 1.4 A˚ structure of an adenosine 5 -phosphosulfate kinase (APS kinase) from the fungus Penicillium chrysogenum (PDB: 1M7G) [275]. The protein is involved in the two-step ATP-dependent conversion of sulfate to 3 phosphoadenosine-5 -phosphosulfate (PAPS). In the first step, ATP sulfurylase catalyzes the formation of adenosine 5 -phosphosulfate (APS) which is, in the second step, converted by APS kinase into PAPS – important for the normal cellular function as a sulfonate donor molecule [275]. Originally, the study intended to obtain a vanadate(V) transition state analogue with the substrate APS and the ADP molecule, but that was not achieved. Instead, vanadate(V) was not bound to the protein but only to the ribose ring of the ADP molecule. Nevertheless, the study gave insights on the APS

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binding site and vanadium appeared to stabilize the crystal packing allowing a better resolution of the structure [275]. Several years later a transition state vanadium-containing kinase structure was finally made available: a 2.7 A˚ structure of a nucleoside diphosphate kinase (NDK) from Staphylococcus aureus (PDB: 3Q8Y). The enzyme transfers phosphate between nucleoside diphosphate molecules from nucleoside triphosphate molecules. In this experiment, the apo form and different substrate-bound structures were also obtained in addition to the vanadate(V) transition state with ADP, contributing to the understanding of the reaction mechanism. Mg2+ ion is present as a cofactor and the vanadate(V) moiety is found at the active site which is formed by three conserved residues (His115, Ser117 and Glu51). These residues are responsible for the stabilization of the ADP molecule, close to vanadate(V). Apart from the coordination to four O-atoms, the V-atom is merely bound to the histidine but not to the other residues, forming a phosphohistidine intermediate [276]. This corroborated the hypothesis by which the His115 is the residue responsible for the catalytic activity acting as a nucleophile. Previous mutagenesis studies with Ser117 and Glu51 also supported this theory as NDK mutants conserved – total or partially – their activity [277,278]. A more recent 2.2 A˚ structure of a transition state nucleoside diphosphate kinase from Borrelia burgdorferi bound to vanadate(V) and ADP was deposited (PDB: 4DZ6) but, so far, the corresponding manuscript is not available. Another kinase vanadium-containing structure was determined: a 1.85 A˚ structure of fosfomycin resistance kinase FomA from Streptomyces wedmorensis complexed with MgADP and fosfomycin vanadate(V) (PDB: 3QVF) [279]. FomA and FomB are recognized as kinases capable of inactivating the antibiotic fosfomycin by adding a phosphate group to its phosphonate group [280,281]. Several structures were obtained in the presence of the ATP analogue illustrating the different reaction steps [282] including the enzyme bound to MgATP, to ATP and fosfomycin, to MgADP and fosfomycin monophosphate and to ADP alone, in addition to the already mentioned MgADP and fosfomycin vanadate(V) [279]. The 3QVF structure (Fig. 19) shows vanadate(V) covalently bound to the phosphonate group of the antibiotic simulating the final reaction product (fosfomycin monophosphate), and the three O-atoms in the vanadate(V) moiety are hydrogen-bonded to the Mg2+ ion and to side chain N-atoms from Lys18 and Lys9 and His58. Sitedirected mutagenesis and respective kinetic studies were carried out successfully to further confirm the involvement of the aforementioned residues in the catalytic activity and in the stabilization of the transition state [279]. Remarkably, even mimicking the final reaction product instead of the transition state, MgADP and fosfomycin vanadate(V) structure shows noteworthy differences from the MgADP and fosfomycin monophosphate structure in which the active site is more disordered (namely the residues involved in the catalytic steps and the nearby loops). MgADP and fosfomycin vanadate(V) structure adopts a closed form with well-ordered residues. Such findings contribute for the proposed associative mechanism (SN2-like) in which a penta-coordinate bipyramidal transition state is achieved [279]. Some other transferase vanadium-containing structures are available in the PDB. In 1998, Pederson and coauthors deposited a 2.1 A˚ resolution structure of a mouse estrogen sulfotransferase (PDB: 1BO6) [283]. The protein is responsible for the sulfuryl transfer from PAPS to estrogenic steroids. To get information about the transition state, vanadate(V) was used along with the inactive cofactor adenosine 3 ,5 -diphosphate (PAP). The vanadate(V) moiety is located in the active center adopting a trigonal bipyramidal geometry with the O-atom of PAP and a water molecule as apical ligands and three O-atoms from Lys48, Lys106 and His108 as equatorial ligands. These residues are involved in the catalytic

Fig. 19. Structural representation of the active center of fosfomycin resistance kinase FomA (PDB: 3QVF) [279]. The V-atom is covalently bound to fosfomycin and the O-atoms from vanadate(V) are hydrogen bonded to Lys9, Lys18 and His58 residues and to a Mg2+ ion. The Mg2+ ion is coordinated by the non-bridging O-atoms of the ␣- and ␤-phosphate groups of ADP and to three water molecules (not shown). ˚ All indicated distances are in A.

process and mutations lead to a significant loss of the activity, confirming their importance in the stabilization of the transition state [283,284]. Over the years, other transferase structures complexed with vanadium compounds have been described. In the 1.9 A˚ resolution structure of a C3 exoenzyme from Clostridium botulinum (PDB: 1UZI), Na3 VO4 was used in the crystallization medium and the formed tetravanadate and vanadate(V) ions participate in the crystal-packing stabilization [285]. Similarly, Na3 VO4 was also used during the crystallization procedures of a 2.8 A˚ resolution structure of a uridine phosphorylase complexed with 2 -deoxyuridine/phosphate from E. coli (PDB: 1RXS) [286]. Vanadate(V) was also studied as a transition state analogue in diverse transferases. Different structural studies were done including a 2.2 A˚ resolution structure of a maltodextrin phosphorylase from E. coli (PDB: 2AZD) [287] and four 1.0, 1.5, 1.6 and 2.1 A˚ resolution structures of phosphotransferases belonging to the haloalkanoate dehalogenase superfamily from Bacteroides thetaiotaomicron (PDBs: 2RBK, 2RAR, 3E81 and 4HGO) [13,288,289]. These studies, along with some other experimental approaches namely site-directed mutagenesis, provided valuable information regarding the mechanism of action of the proteins elucidating the geometry of the complex and the residues involved in the transition state, as described in other sections of this review. 4.3. EctoNTPDases Nucleoside triphosphate diphosphohydrolases (NTPDases) are ecto-nucleotidases responsible for the hydrolysis of extracellular nucleotide triphosphates or diphosphates to nucleotide monophosphates (commonly designated by NTP, NDP and NMP, respectively) [290,291]. These small molecules act as mediators in intercellular signaling pathways and they are recognized by

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purinergic receptors distributed at the cell surface [292]. Additionally, these enzymes are implicated in inflammation, vasodilation and tumor progression [293,294]. Therefore, NTPDases have been the focus of intense research in the last decade due to their effect on cellular metabolism, proliferation, differentiation and apoptosis. In vertebrates, there are 8 types of NTPDases constitutively expressed in different tissues from which four – NTPDase1 (previously known as Cluster of Differentiation 39, CD39, in humans), NTPDase2, NTPDase3 and NTPDase8 – are found as membranebound enzymes on the cell surface [295]. NTPDases were also identified in some pathogen organisms acting as virulence agents [296] being used as targets for the design of efficient inhibitors [297]. The first crystal structures of a member of this family of enzymes was published in 2008 [298]. Several structures of the non glycosylated Rattus norvegicus NTPDase2 were obtained in the presence and absence of AMPPNP (5 -adenylyl-␤,␥-imidodiphosphate, a non-hydrolysable ATP analogue), AMP and Ca2+ (important for the protein activity) – PDBs: 3CJ1, 3CJ7, 3CJ9, and 3CJA. The cation and the substrate analogue bind in the hydrolytic site in a cleft formed by two lobes. Several highly conserved residues, belonging to the 5 apyrase conserved regions (ACR), are implicated in the binding and stabilization of the phosphorylated nucleotide via H bonds and ␲ stacking interactions. Importantly, the study points the importance of the enzyme flexibility: in the rest state, NTPDase2 is in an open form that closes upon substrate binding favoring hydrolysis [298]. To further understand NTPDases inhibition mechanisms, R. norvegicus NTPDase1 crystals were soaked with sodium vanadate(V) solutions (this condensed to decavanadate during the crystallization procedure), and ammonium heptamolybdate (PDBs: 3ZX2 and 3ZX0) [299]. In the obtained 1.8 A˚ resolution structure, decavanadate(V) is interacting with Lys406 and Lys408 close to the nucleotide binding site. This conserved site is located in the Cterminal domain, in the loop containing the base-stacking Tyr409 (involved in the NTPDase3 activity [300]). These structural data suggest that the inhibitory effect of the metal ion is due to restrictions of the flexibility of the enzyme that prevent substrate binding [299]. The structures of a few other R. norvegicus and Legionella pneumophila NTPDases were released including 3 new soluble L. pneumophila NTPDase1, namely 1.6–1.7 A˚ resolution structures in complex with adenosine 5 -phosphovanadate(V) (PDB: 4BRE), thiamine-5 -phosphovanadate(V) (PDB: 4BRH) and guanosine-5 phosphovanadate(V) (PDB: 4BRL). Common to the three structures is the in situ resulting adduct located in the active site mimicking the putative transition-state of the enzymatic reaction. The V atom assumes a trigonal–bipyramidal geometry with two well-defined apical ligands: the reaction product and a nucleophilic water, previously activated by the Glu159 residue [301]. This structural study was useful for a detailed characterization of the mode of action of L. pneumophila NTPDase1. Briefly, the leaving phosphorus atom moves in the direction of the nucleophilic water, which remains in its position next to the Glu159 residue. In addition, Arg56, bound to the ␣-phosphate, stabilizes the phosphorus atom and promotes the formation of the described trigonal–bipyramidal transition state (PDBs: 4BRE, 4BRH and 4BRL) [301]. 4.4. Phosphodiesterases and phosphomutases Phosphodiesterases, a class of phosphorylases, are enzymes that catalyze the hydrolysis of phosphoester bonds in substrates that contain two phosphoester bonds, frequently of cyclic nucleotides. If the enzyme catalyzes the transfer of a phosphate from a OH to another OH within the same substrate molecule, the enzyme is designated as a phosphomutase.

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Fig. 20. Structural representation of the vanadate(V) moiety present in the Tdp1, DNA and human topoisomerase I-derived peptide complex structure (PDB: 1NOP) [303]. With a trigonal–bipyramidal geometry, the V-atom interacts with the His263 of Tdp1 but also with the DNA and the peptide (Tyr723) and with two O-atoms hydrogen bonded to different Tdp1 residues (some distances are depicted in the ˚ figure in A).

4.4.1. Phosphodiesterases A few vanadium-containing phosphodiesterases structures are deposited in the PDB, namely human tyrosyl-DNA phosphodiesterase (Tdp1) (PDBs: 1MU9 [302] and 1NOP [303]). This protein, member of the phospholipase D superfamily, hydrolyzes tyrosineDNA-3 -phosphate phosphodiester bonds playing a role in DNA repair by removing undesired topoisomerase I–DNA complexes from the DNA strand [304]. In the 1MU9 structure, at 2.05 A˚ resolution, vanadate(V) is located in the active site with a trigonal-bipyramidal geometry mimicking the transition state of the 1st step of the catalytic reaction with the vanadium covalently bound to His263 (responsible for the nucleophilic attack on phosphate). Vanadate(V) is also hydrogen-bound to two lysine residues (Lys265 and Lys495) and to a glycerol molecule from the cryoprotectant solution. Interestingly, another structure was solved using tungstate rather than vanadate(V); similar results were obtained but tungstate adopts an octahedral geometry [302]. The 1NOP structure was obtained at 2.3 A˚ resolution containing not only the vanadate(V), but also a DNA molecule and a small human topoisomerase I-derived peptide which simulates the enzyme substrate. In this complex, the vanadate also adopts a trigonal–bipyramidal geometry where the V atom is bound to the His263 of Tdp1, the Tyr723 residue of the small peptide and the 3 hydroxyl of the DNA molecule. The remaining two O-atoms of the vanadate moiety are hydrogen bonded to four residues of Tdp1 (Lys265, Asn283, Lys495 and Asn516) [303] (Fig. 20). The structure supports the suggested enzymatic mechanism providing some new insights into conformational changes of the substrate occurring before the Tdp1 activity [303] as previously hypothesized [304]. The same group also investigated the binding ability of the Tdp1 using vanadate(V) with a large range of DNA nucleotides and different peptides or octopamine (a tyrosine analogue). Several 1.7–2.3 A˚ resolution structures were deposited (PDBs: 1RFF, 1RFI, 1RG1, 1RG2, 1RGT, 1RGU and 1RHO) attesting the capacity of

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where one of the axial positions corresponds to the nucleophilic residue Thr90. The reported results were useful to elucidate the catalytic mechanism as well as to clarify some evolutionary aspects of both proteins [309].

Fig. 21. Structural representation of the semi-reduced CPDase complexed with 2 ,3 -cyclic uridine vanadate(V) (PDB: 1JH7) [306]. The vanadate(V) moiety is found in the active site bound by His42, Thr44, His119, Ser121 and Tyr124. Some distances ˚ are shown in A.

Tdp1 to bind to several complexes potentially usable as inhibitors [305]. Two more vanadium-containing phosphodiesterase structures are available in the PDB. One is a cyclic nucleotide phosphodiesterase (CPDase) from Arabidopsis thaliana (PDB: 1JH7) [306]. This enzyme acts in the tRNA splicing process catalyzing the conversion of ADP-ribose 1 ,2 -cyclic phosphate into ADP-ribose 1 -phosphate. Additionally, CPDase also hydrolyzes nucleoside 2 ,3 -cyclic phosphates to nucleoside 2 -phosphates [306,307]. Following the oxidized native structure (PDB: 1FSI) and the respective mechanism of action [308], the inhibitor-bound semi-reduced CPDase structure was obtained at 2.4 A˚ resolution using 2 ,3 -cyclic uridine vanadate(V) as inhibitor (PDB: 1JH7) [306]. Both forms share a common overall structure despite some minor differences due to the absence of a Cys104-Cys110 disulfide bridge in the semi-reduced protein. The vanadate(V) moiety sits in the active site hydrogen bonded to His42, Thr44, His119, Ser121 and Tyr124 (Fig. 21); some other residues are also interacting with the uridine moiety (Trp12, Ser10 and Thr163) contributing to the adduct stabilization [306]. Due to similarities between 2 ,3 -cyclic uridine vanadate(V) and the cyclic phosphorylated substrates of CPDase, the obtained structural characterization confirms the previous proposed mechanism where His119 acts as the nucleophilic residue involved in the attack to the substrate [306,308]. The last structure to be deposited in the PDB was a nucleotide pyrophosphatase/phosphodiesterase (NPP) from Xanthomonas axonopodis pv. citri at 1.3 A˚ resolution (PDB: 2GSO) [309]. This study presents a comparative analysis with a related alkaline phosphatase which has a preference for phosphate monoesters, while NPP prefers phosphate diesters. Vanadate(V) is found in the active site of NPP exhibiting a trigonal–bipyramidal geometry

4.4.2. Phosphoglycerate mutases Mutases belong to the family of isomerases and are enzymes responsible for changing the position of functional groups within the same biological active molecule. Phosphoglucomutase and phophosglycerate mutase (PGM) are two examples of mutases. Phosphoglucomutase catalyzes the conversion of glucose-6phosphate to fructose-6-phosphate and the dephospho form is potently inhibited by glucose-6-phosphate and glucose-1phosphate in the presence of vanadate(V) [310]. The inhibition was attributed to the formation of 1-phosphoglucose-6-vanadate(V) and 1-vanadofructose-6-phosphate, which form stable complexes with the protein [5]. An inhibition constant of 2 × 10−12 M was determined for 1-phosphoglucose-6-vanadate(V) at pH 7.4 [311]. The effects of vanadate(V) on phosphoglycerate mutase were also described and some structural studies are available. The results were fully consistent with non-cooperative binding of vanadiophosphoglycerate to the two active sites of phosphoglycerate mutase; 2-vanadio-3-phosphoglycerate binds to the dephospho form of phosphoglycerate mutase with a dissociation constant of ∼1 × 10−11 M at pH 7 and 7 × 10−11 at pH 8.4. The two forms of phosphoglycerate mutase present in mammals – the 2,3phosphoglycerate-dependent (also known as cofactor-dependent, dPGM) and 2,3-phosphoglycerate-independent – have different affinities for vanadate(V), the cofactor dependent form responding more potently to vanadate(V) [312]. PGM participates in glycolysis and, due to its synthase, phosphatase and mutase activity, it catalyzes the interconversion of 3-phosphoglycerate (3-PGA) into its isomer 2-phosphoglycerate (2-PGA). This reaction requires an enzyme phosphorylation prior to substrate binding. A histidine residue located in the active site is phosphorylated, further transferring the phosphate group to 3-PGA and producing the 2,3-PGA intermediate; the intermediate is later hydrolyzed with the concomitant release of 2-PGA. In the case of dPGMs, 2,3-PGA is, in fact, responsible for the phosphorylation of the active site, restoring the active form of the enzyme [312–315]. The first dPGM crystal structures were determined in the active form where the enzyme is either interacting with the substrate or oxoanions like phosphate or sulphate [316,317]. The crystal structure of E. coli dPGM complexed with vanadate(V) was determined to a resolution of 1.30 A˚ (PDB: 1E59) [318]. Four vanadate(V) ions are bound in the highly basic active site, mainly as divanadate(V) in the center, but with evidence of one additional vanadate(V) at each end modeled with half occupancy. The contacts between vanadate(V) and the protein consist of hydrogen-bonds and electrostatic interactions. The authors discuss the enzyme-ligand interactions involved in inhibition of the mutase activity by vanadate(V) and identified a water molecule, observed in the native E. coli dPGM which, once activated by vanadate(V), may dephosphorylate phosphohistidine of the active protein, producing inorganic phosphate and the inactive (dephosphorylated) protein [318]. Later the structure of the bacterial dPGM from Burkholderia pseudomallia was also solved complexed with vanadate(V) (PDB: 3GW8 and 3GP5) [319]; in the 3GW8 structure at 1.9 A˚ resolution, glycerol, from the crystallization conditions, reacted with vanadate(V) forming a species quite similar to the enzymatic substrate. The O-atom of glycerol is one of the apical ligands of vanadate(V) together with the N-atom of the His9 residue and the complex perfectly fits the substrate-binding pocket, indicating the important residues for catalysis. A similar result was obtained in the 3GP5 structure at 2.25 A˚ resolution where 3phosphoglycerate was used [319].

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4.5. ATPases (myosins and transporters) 4.5.1. ATPases ATPases are enzymes that catalyze the hydrolysis of phosphateanhydride bonds with many important roles in biology namely in cellular energy metabolism. A wide range of affinities for vanadate(V) are observed depending on the type of ATPases [320–325]. The inhibitory effect of vanadate(V) on some ATPases may vary from those corresponding to nM inhibition constants for the Na+ , K+ -ATPases [326], to those of mM inhibition constants as for F1ATPases. This subject was reviewed by Crans [5] and the discussion made continues actual. The ATPase enzymes include many membrane enzymes and inhibitory studies with vanadate(V) normally assume that it acts as a phosphate analogue and presumably inhibits the enzymes as a transition state analogue for the phosphoryl group transfer [327–329]. One of the most extensively studied example is the inhibition of the Na+ ,K+ -ATPase by vanadate(V) [326,330]. Vanadate(V) binds very tightly with an association constant of 2.4 × 108 M−1 [331], and this strong binding has been attributed to the formation of stable 5-coordinate complexes [22] involving a bidentate coordination with the carboxylate group of an Asp residue. The discovery of the inhibitory effect of vanadium on the Na+ ,K+ ATPases [326] has encouraged the investigation of similar effects on other ion pumps such as the H+ /K+ -ATPase, or Ca2+ -ATPase [326]. These enzymes are classified as P-type, or E1-E2, ATPases in which the ion pump process is coupled to ATP hydrolysis. It is well-known that the process of Ca2+ transport and ATP hydrolysis can be inhibited by monomeric vanadate(V), particularly under conditions where the E2 conformation is favored, leading to the formation of a conformer analogous to E2P, but involving vanadate(V) instead of phosphate [332,333]. Therefore, the vanadate(V) mode of action may involve formation of an E2 V analogue after binding of vanadate(V) to the enzyme phosphorylation site. Interestingly, decavanadate (V10 ), [V10 O28 ]6− , is a more potent SR Ca2+ ATPase inhibitor than monomeric vanadate(V) [332] and interacts with other conformations besides E2. In this case, the binding site is not the same as the ATP binding site [334]. Moreover, during the mechanism of decavanadate-mediated SR Ca2+ ATPase inhibition, a protein cysteine residue is oxidized through reduction of the vanadate(V), although the full implications of this process are not yet clear [335]. Decavanadate despite interacting most strongly with the E2 conformation of the Ca2+ -ATPase, can also interact with other conformations [332]. The mode of action of decavanadate and of other oxometalates implies that the main focus of the interaction is on the cytoplasmic domain of the protein, more specifically at the nucleotide or phosphorylation domains or at a pocket involving several cytoplasmic domains [9]. Additionally, in vitro studies suggest that the effects of V10 are particularly relevant in mitochondrial depolarization and oxygen consumption, and in vivo, the administration of different vanadadate species affect the V distribution in mitochondria [336,337]. Interaction of decavanadatevanadate(V)s with biologically relevant molecules, namely proteins, was reviewed by Aureliano et al.; this topic is briefly addressed below and for more details we recommend references [9,336,337]. The mechanism of inhibition and interaction of decavanadate on the sarcoplasmic reticulum Ca2+ -ATPase differs from those of other polyoxometalates or oxometalates. It involves protein cysteine oxidation and vanadate(V) reduction processes, leading to non-competitive ATPase inhibition. This inhibition is not prevented by antioxidant agents that can reverse the cysteine oxidation [336]. Moreover, decavanadate interacts with ATPase in the same fashion for all the conformations which occur during the ATP-hydrolysis coupled Ca2+ translocation.

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Heteronuclear adducts of vanadate(V) with phosphate have also been studied extensively and applications in biology could be anticipated for substrate analogues for ATPases. In fact, V-phosphate bonds are of wide interest in industry since there are important catalyst applications involving such type of species and/or intermediates, as e.g. in the large scale preparation of maleic anhydride [338,339]. Additionally structural characterization exists for a wide range of materials which contain moieties that can be classified as vanadate(V)-phosphate anhydrides; in fact, many examples of several types of On POVOm units have been characterized testifying the interest in materials containing these types of moieties and their bonding patterns. However, almost no studies addressed the importance of the vanadate(V)-phosphate anhydride in biology [5,34]. In addition to the biological applications of vanadate(V) to probe ATPases, another area of biomimetic chemistry of great interest is the vanadate(V)-induced photolytic cleavage of V-protein complexes. The initial studies were carried out with dynein and myosin, which are ATPases providing energy for intracellular transport which cleave at one specific amino acid when exposed to UV light [340–342]. Since this early discovery, the photolytic cleavage of dynein, myosin, and adenosine kinase has been extensively studied, providing information on the phosphate–vanadate(V) binding site in the presence of free or bound nucleotides. The review of Crans [5] and the chapter by Gibbons and Mocz [343] summarize most of the relevant studies in this field.

4.5.2. Myosins Myosins are recognized as molecular motor proteins participating in several mobility mechanisms such as muscle contraction and cytokinesis, upon ATP consumption. Myosins encompass three domains, arranged in a conserved architecture: head, neck and tail. The head corresponds to a motor domain comprising two binding sites responsible for interacting with the actin and ATP hydrolysis. The neck or level arm increases the conformational change caused by ATP hydrolysis and is responsible for the binding of regulatory light chains like calmodulin. The tail contains a coiled coil and a targeting domain contributing for the enzyme specificity [344,345]. Myosins, widely spread in nature, have been extensively studied and divided into different classes according to their function and location. Traditionally, myosins were organized in 18 classes albeit some more recent studies point for the existence of 35 classes [346–348]. From a structural point of view, the first XRD myosin structure was published by Rayment, Holden and co-workers in 1993: a 2.8 A˚ structure of the head domain of chicken skeletal muscle myosin (PDB: 2MYS) [349]. Several vanadium-containing myosin structures from different types were deposited in the PDB. The first structure, released in 1996, was a truncated fragment (residues 2–762) corresponding to the head section of Dictyostelium discoideum myosin II complexed with magnesium, ADP and vanadate(V) at 1.9 A˚ resolution (PDB: 1VOM) [350]. The adduct sits in the nucleotide binding site and the vanadate(V) depicts a trigonal-bipyramidal geometry. The equatorial positions are occupied by three oxygen atoms while the axial positions are occupied by an O-atom of the ␤-phosphate moiety and a nucleophilic water molecule. The vanadate(V) moiety is stabilized by several hydrogen bonds between the O-atoms and different residues namely Lys185, Ser181, Asn233 and Ser236 in addition to a Mg2+ ion [350]. Experiments involving different myosin inhibitors were also carried out. Blebbistatin is recognized as a myosin II inhibitor by decreasing the affinity towards actin [351] and, in 2005, a 2.0 A˚ resolution structure of D. discoideum motor domain of myosin II containing ADP, vanadate(V) and blebbistatin (PDB: 1YV3) was released [352].

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Blebbistatin, via hydrophobic interactions, is bound to the intermediate state of the protein in a hydrophobic pocket located between the upper and lower 50-kDa domains of the head portion (known as 50-kDa cleft). Vanadate(V) and ADP moiety are placed next to the blebbistatin holding the open state of the binding cleft. Therefore, blebbistatin allows the open state of the cleft which explains the inhibitory effect since the phosphate release occurs with a progressive closure of the 50-kDa cleft [352]. In some related studies, blebbistatin was chemically modified to evaluate the biological effect of its derivatives and three new protein-inhibitor structures were obtained (PDBs: 3BZ7, 3BZ8 and 3BZ9) [353]. Similarly, other inhibitors were also studied such as pentabromopseudilin (PDB code: 2JHR) [354], pentachloropseudilin (PDB: 2XEL) [355], tribromodichloropseudilin (PDB: 2XO8) [356] and pentachlorocarbazole (PDB: 2X9H). Other vanadium-containing myosin type II structures were deposited focusing, for example, on the importance of Ser236 in the protein activity (PDB: 3MYH) [357]. Other V-related myosin types were also structurally characterized helping to understand the respective function. That is the case of myosin I (PDBs: 1LKX and 4BYF) [358,359] and myosin VI (PDBs: 2V26, 4E7S and 4E7Z) [360,361]. For instance, the 2.7 A˚ resolution structure of human myosin 1c (PDB: 4BYF), which participates in the insulin-stimulated glucose uptake, encompasses the head and the neck in complex with calmodulin. In this complex, Ca2+ is also involved, highlighting the role of this cation in the myosin regulation [359]. Within the context of potential physiological roles of vanadate(V)’s interaction with proteins and cell membranes, besides the ability of decavanadate to interfere with the Ca2+ homeostasis in muscle tissues by inhibition of the SR Ca2+ -ATPase, it also reduces actomyosin ATPase activity [362]. Myosin is a protein that plays a pivotal role in ATP-driven Ca2+ -dependent muscle contraction and it was found that docking of decavanadate stabilizes it against break-down to monovanadate(V). In vitro experiments showed that V10 inhibits myosin ATPase, SR Ca2+ -ATPase, and SR Ca2+ release induced by the second messenger inositol triphosphate (IP3) [363–365]. Moreover, decavanadate inhibits both the contractile system and the regulation of muscle contraction. In fact, by blocking the Ca2+ release the contraction of the calcium pump and/or the actomyosin release of the metabolites maintain the proteins in a “pre-rigor” like state and prevents the relaxation of the muscle [362,363,366]. Monomeric vanadate(V) mimics the transition state for the phosphate hydrolysis [367], blocking myosin by mimicking the ADP–phosphate intermediate state, while decavanadate induces the formation of the intermediate myosin–MgATP–V10 complex blocking the contractile cycle, most probably in the pre-hydrolysis state [362]. However, whereas the former complex is destabilized by F-actin, in comparison to in vivo conditions, the latter is not, suggesting that V10 is the only species capable of preventing the release of the products during the mechanism of ATP hydrolysis by the actomyosin complex [363]. Therefore, depending on their oligomerization state, vanadate(V)s are able to populate different conformational states of the myosin–ATPase cycle. 4.5.3. Transporters ATP-binding cassette transporters (abbreviated ABC transporters), widely spread in nature, are membrane proteins found in prokaryotes and eukaryotes. ABC transporters hydrolyze ATP into ADP and phosphate. The resultant energy is used to promote the transport across cell membranes allowing either the entrance or the exit of several substances (importers and exporters, respectively). ABC transporters have been extensively studied due to their biological relevance and several reviews are available including their structural characterization [368–371].

Three vanadium-containing ABC transporters XRD structures were determined in the last decade. The first one deposited, in 2002, was a 3.2 A˚ resolution structure of BtuCD from E. coli (PDB: 1L7V) [372]. The complex is involved in the vitamin B12 uptake enclosing the two ATP binding subunits (BtuD) and the two transmembranar subunits (BtuC) composed by 20 helices each. Vanadate(V) is known to inhibit ABC transporters [373–375] and two cyclotetravanadate moieties were identified in the BtuD subunits located in the so-called P-loop or Walker A motif. P-loop is responsible for the nucleotide binding containing a conserved lysine residue with a pivotal role by interacting with the ␤- and ␥-phosphate of the ATP molecule [372]. The oligovanadate(V)-protein structure adduct was relevant for the proposal of the transport mechanism of vitamin B12 and it was the starting point for further related XRD investigations [376–378]. The second structure was released in 2007: a 4.2 A˚ resolution structure of a lipid A and lipopolysaccharide transporter MsbA from Salmonella typhimurium (PDB: 3B5Z) [379]. Despite the low resolution by comparison with the also obtained structure with AMPPNP (PDB: 3B60), this ADP–vanadate(V) structure confirms the positioning of the nucleotide and emphasizes the importance of the protein flexibility [379]. The last structure reported was a 2.4 A˚ resolution study of a maltose transporter complex (MBP-MalFGK2 ) from E. coli formed by four proteins: a periplasmic maltose-binding protein (MBP), two transmembranar proteins (MalF and MalG) and two subunits of the ATPase protein (MalK) (PDB: 3PUV) [380]. A structural approach using vanadate(V), BeF3 − and AlF4 − (other phosphate mimicking molecules) and AMPPNP was followed. The ADP–vanadate(V) adduct mimics the transition state exhibiting the expected trigonal–bipyramidal in the active site. The axial positions are occupied by the ␤-phosphate from ADP and a nucleophilic water molecule. Different residues from the conserved motifs of the ABC transporter and a Mg2+ ion stabilize the adduct. Particular attention was given to Glu159 residue: hydrogen bonded to the nucleophilic water, the residue is responsible for its activation and mutagenesis experiments proved that it is essential for the ATP hydrolysis [380]. 4.6. ATP synthases ATP synthases are responsible for the synthesis of ATP from ADP, inorganic phosphate (Pi ) and Mg2+ using an electrochemical gradient of protons from the electron transport chain. The complex encloses a soluble unit (designated by F1 in Bacteria and eukaryotes or A1 in Archaea) and a transmembrane unit (Fo or Ao , respectively) [381–383]. Multiple biochemical and structural studies were focused on this protein due to its essential cellular roles and two vanadium-containing ATP synthase XRD structures are available in the PDB. The first structure to be released was a 3.0 A˚ resolution XRD structure of the subunit F1 from the mitochondrial rat ATP synthase (PDB: 2F43) [384] representing a transition state containing ADP, vanadate(V) and Mg2+ . The vanadate moiety is located next to the P-loop (156 GGAGVGKT163 ) and several residues, not only from the P-loop, are involved in its stabilization, namely Glu188 that acts as the catalytic base [384]. Interestingly, compared to the previously obtained F1 ATP synthase in the ground state with phosphate (PDB: 1MAB) [385], a rearrangement of the P-loop is observed due to the presence of the monovanadate(V). This rearrangement involves the movement of Ala158 closer to the O-atom of the vanadate in the transition state (3.8 A˚ distance) compared to the ground state ˚ [384,385]. This study allowed a better characterization of (5.2 A) the transition state confirming the previous findings on the role of vanadate(V) on the process [386,387]. The other structure was deposited in 2011: a 2.85 A˚ XRD resolution structure of the catalytic A subunit of A1 Ao ATP synthase from

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Pyrococcus horikoshii (PDB: 3P20) [388]. This protein, an example from Archaea [389], was previously structurally characterized in complex with sulfate, ADP and AMPPNP [390]. The modeled structure contains two vanadate(V) moieties, placed next to the P-loop (7.94 A˚ apart from each other), hydrogen bonded to Ser238 and Leu417 residues (1st and 2nd vanadate(V), respectively) and also depicting weak non-polar interactions with several other protein residues (but not with water molecules). The significance of the second vanadate(V) (found in the transient binding site) and the importance of the Lys240 and Thr241 residues was discussed using a combined approach which includes mutagenesis, ITC and X-ray crystallography techniques [388]. 4.7. DNA binding proteins Vanadium ions can interact with DNA binding proteins in a similar way to what has been described for the previous families of enzymes. 4.7.1. Topoisomerases Besides phosphoglycerate mutases, there are a few vanadiumcontaining isomerase (topoisomerases) structures available in the PBD where vanadate(V) was used to obtain information regarding the transition state of the catalytic reaction. In 2006, a 2.3 A˚ resolution transition state structure of topoisomerase I from Leishmania donovani (PDB: 2B9S), a potential clinical target against leishmaniases, was deposited in the data bank [391] encompassing a nicked double-stranded DNA and vanadate(V). The protein controls the topological state of the DNA (important in several cellular mechanisms such as replication and transcription) and is able to produce DNA supercoils by cleaving one DNA strand before re-linking it without using ATP [392,393]. Following some other previously deposited human topoisomerase I structures [394–396], the 2B9S structure contains a large subunit (TOP1L) and a small subunit (TOP1S). The vanadate(V) moiety presents a distorted trigonal–bipyramidal geometry with the 5 -hydroxyl end of the DNA nick and the nucleophilic Tyr222 residue in the apical positions. His453, Lys314 and Lys352 residues are hydrogen bonded to the apical or equatorial ligands, contributing for the stabilization of the transition state analogue. Furthermore, the Arg410 residue and a water molecule are next to the phenolic oxygen of Tyr222 being implicated in the respective nucleophile activation process [391]. In 2010, another topoisomerase structure was determined: a 2.1 A˚ resolution structure of the variola virus topoisomerase IB complexed with DNA and vanadate(V) (PDB: 3IGC) [397]. Complementing the previously available information, the 3IGC structure mimics the transition state and is quite akin to that portrayed for the L. donovani [391]. The vanadate(V) moiety, once more adopting a trigonal–bipyramidal geometry is also bound to the DNA and a tyrosine residue (Tyr274). Some other residues and water molecules involved in the catalytic process were identified and their implications to the mechanism of action of the protein were thoroughly discussed [397]. Another vanadium-containing isomerase XRD structure is a 2.3 A˚ resolution structure of a phage P1 Cre recombinase complexed with DNA and vanadate(V) (PDB: 3MGV) [398]. Belonging to the tyrosine recombinase family, the enzyme rearranges the DNA similarly to that described for topoisomerases IB. The vanadate(V) moiety is also bound to a nucleophilic tyrosine residue (Tyr324) in addition to the DNA molecule corroborating such similarity. Mutagenesis experiments were done with the relevant residues for the catalytic activity (namely Arg173, Arg292, Lys201, Trp315, His289 and Glu176) followed by in vivo and in vitro recombination activity tests. The importance of Try324 and Lys201 (responsible for the activation of the O5 leaving group)

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was identified supporting the structural insights from the transition state model with vanadate(V) [398]. 4.7.2. Other DNA binding proteins DNA-binding proteins have DNA binding domains with a specific or general affinity for either single or double stranded DNA. Sequence-specific DNA-binding proteins generally interact with the major groove of B-DNA because it exposes more functional groups that identify a base pair. However there are also some known minor grove DNA-binding ligands. In accordance to what has been described, vanadate(V) was also used as a transition state intermediate in studies on DNA binding proteins, e.g. aprataxin and protelomerase. Aprataxin, involved in DNA repair mechanisms, is able to reverse DNA adenylation which consists in the removal of a 5 -adenylate molecule (adenosine monophosphate, AMP) from DNA breaks. The protein is an important target for drug design as its malfunction is responsible for ataxia oculomotor apraxia, a neurodegenerative illness [399–401]. Aprataxin has a histidine triad HHH ( are hydrophobic residues) as a catalytic domain participating in a twostep deadenylation process along with a zinc finger domain [399]. The human XRD structure of the adenosine–vanadate(V) intermediate (PDB: 4NDG) shows His260, the nucleophilic residue in the first step of the deadenylation process, in one of the axial positions of the adduct, mimicking the covalent bond established between the protein and the AMP in the catalytic reaction. In addition, the transition state stabilization is assured by several other residues namely Ser255, Met256 and His 262 [399]. Protelomerases, or telomere resolvases, are DNA cleavagerejoining proteins involved in the replication of linear DNA of prokaryotes namely in some human pathogens such as the genus Borrelia. Protelomerases, acting as dimers, form hairpin telomere – structures of covalently closed DNA present at the end of chromosomes and are important for their protection against degradation – by a two-step transesterification involving a 3 -phosphotyrosylenzyme intermediate [402]. There are two vanadium-containing protelomerase-DNA structures: TelK (PDB: 2V6E) at 3.20 A˚ resolution [403] and TelA (PDB: 4E0G) at 2.20 A˚ resolution [404]. In both cases, vanadate(V) was used with a DNA nicked at the scissile phosphate and the moiety is located in the protein active site, bound to a nucleophilic tyrosine residue (Tyr425 and Tyr405, respectively) and also to the 5 - and 3 OH groups of the DNA (Fig. 22); some other residues are involved in the hairpin stabilization like Arg205 in TelA [403,404]. These structures present new insights on the hairpin telomeres formation, and a multistep DNA refolding mechanism was proposed [404]. 4.8. RNA binding proteins 4.8.1. Ribonucleases Ribonucleases, also designated as RNases, catalyze the cleavage of RNA and vanadate(V), VIV O2+ and some of their complexes are strong inhibitors of the hydrolysis of the phosphodiester bond in RNA (and DNA as already discussed). Vanadium-nucleoside complexes are inhibitors of ribonuclease catalysis and for several years vanadate(V) was the reagent used to prevent ribonuclease cleaving action of RNA in experimental cellular isolations [5]. As in other enzymatic cleavages of the P O bond, the transition state postulated is a pentavalent P in a trigonal–bipyramidal geometry [22]. Comprehensive data on characterization of model compounds for inhibition of ribonucleases may be obtained in the review of Crans et al. [5]. The first vanadium-related ribonuclease deposited in the PDB was RNase A (bovine pancreatic ribonuclease A). RNase A is involved in two reactions: the cleavage of the 3 –5 phosphate bond (by the transphosphorylation of the RNA) and the

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Fig. 22. Structural representation of the vanadate(V) moiety bound to TelA and DNA (PDB: 4E0G). A 5-coordinated adduct is formed next to the nicked DNA and the nucleophilic tyrosine residue [404].

hydrolysis of the produced 2 ,3 -cyclic phosphate intermediate to form a 3 -nucleotide [405,406]. Uridine vanadate(V) was used as a 2 ,3 -cyclic phosphate analogue due to their structural similarities; the first complex with RNase A was solved by a combined X-ray and neutron diffraction methods [407] and it was complemented by NMR studies (PDB: 6RSA) [408]. Later, a higher XRD resolution structure at 1.3 A˚ was obtained (PDB: 1RUV) [409]. In these first studies a characterization of the adduct in the active site of the protein, with a 5-coordinate trigonal–bipyramidal geometry, was obtained (Fig. 23). As expected, the reaction involves the formation of a cyclic ester with V coordinated across the 2 and 3 positions of ribose. Several residues are hydrogen bonded to the O-atoms of vanadate(V) namely His12, His119, Lys41 and Gln11 [406,409]. Mutagenesis studies were also carried out pointing to the role of each residue in the RNase A action, particularly the His12 and His119 residues [406,410]. The structure was also compared with a free-phosphate RNase A (PDB: 7RSA) [411] in order to evaluate the possible structural changes. Apart from some minor changes (namely the side chains of residues Val43 and Arg85), the overall structure is maintained upon ligand binding, as observed by the low RMSD value of the superposition models (0.18 A˚ for all C␣ atoms) [411]. A RNase T1 vanadium-containing structure was also solved at 1.8 A˚ resolution (PDB: 3RNT) [412]. This ribonuclease, from the fungus Aspergillus oryzae, degrades the RNA at the 3 -phosphate of guanine residues presenting two very similar isoforms which differ only in the residue 25: glutamine (Gln25-RNase T1 ) or lysine (Lys25-RNase T1 ) [413,414]. Using guanosine vanadate(V) the obtained Lys25-RNase T1 structure is not mimicking the transition-state of the reaction. Instead, an adduct was obtained corresponding to a H2 VO4 − hydrogen bonded in the active site as a 4-coordinate ground-state analogue. The 3RNT structure was also compared with a previous RNase T1 -2 -GMP structure (PDB: 1RNT) [415] identifying the structural alterations caused upon the binding of the substrate analogue [412]. Another vanadium-containing ribonuclease XRD structure was deposited in 2006: a D192N mutant of Bacillus halodurans RNase H at 1.8 A˚ resolution structure complexed with vanadate(V) and a RNA/DNA hybrid with a nonphosphorylated nick (PDB: 2G8H)

[416]. RNase H, widely spread in Bacteria, Archaea and eukaryotes, binds to RNA/DNA hybrids cleaving the RNA strand in an unspecific mode. Therefore, RNase H plays an important role in the DNA replication process by removing the RNA primers [416,417]. In addition, the protein occurs also as a reverse transcriptase domain making it a possible target for anti-viral drugs [418]. RNase H has four conserved residues in the active center (Asp71, Asp132, Asp192 and Glu109) preferentially coordinated to two Mg2+ (essential for the protein function and commonly designated by metal ions A and B). Different mutant Bacillus halodurans RNase H structures with an RNA/DNA substrate were released (including structures with the reaction product) [416,417] and vanadate(V) was used to characterize the transition state. However, the vanadate(V) was not found in the phosphate binding site; instead, a vanadate(V) moiety was located next to His172 and Trp168 residues providing some insights on the catalytic mechanism, namely in the action of the metal ions A and B [416]. 4.8.2. Ribozymes Discovered in the 80s, ribozymes are RNA molecules capable of catalyzing some reactions in a similar way to protein enzymes. There are different types of ribozymes – namely hammerhead, hairpin, hepatitis delta virus (HDV) and glmS ribozymes – and a considerable amount of information has been compiled over the years [419–425]. Structural characterization was not obliterated during the investigations on ribozymes using mainly XRD but also solution NMR and fluorescence transfer spectroscopies. In fact, fluorescence transfer was used to obtain the first ribozyme structure deposited in the PDB (PDB: 1RMN) [426]. Currently, more than 220 structures are available [422,427,428] including two vanadiumcontaining entrances which mimic the transition state of these peculiar enzymes. One, deposited in 2002, is a 2.2 A˚ resolution structure of a hairpin ribozyme in the presence of vanadate(V) (PDB: 1M5O) [429]. The hairpin ribozymes act similarly to the ribonucleases A by cleaving the phosphodiester bonds of the RNA molecule in a transesterification producing a 2 ,3 -cyclic phosphate and a free 5 -hydroxyl [430]. In this case, monovanadate(V), placed in the active site, assumes the expected trigonal–bipyramidal geometry coordinating three RNA O-atoms (2 - and 3 -oxygen of nucleotide −1 and 5 -oxygen of nucleotide +1) and other two O-atoms. Vanadate(V) makes five

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Fig. 23. Structural representation of the vanadate(V)-RNase A adduct (PDB: 1RUV) [409]. The structure was important to fully understand the uridine vanadate(V) binding ˚ to the protein. Some distances are shown in A.

hydrogen bonds to the nucleobases of G8, A9 and A38, more than the interactions observed in the pre-cleavage and post-cleavage state, confirming the importance of the transition state stabilization. The role of these purines as base (G8) and acid (A38) in the catalytic process was discussed [429]. Five years later, a new vanadium-containing hairpin ribozyme structure at 2.05 A˚ resolution was released (PDB: 2P7E) [431]. The vanadate(V) moiety and its surroundings are quite similar to those described for the structure 1M5O, but present two water molecules in the active center interacting by hydrogen bonds with the Oatoms of vanadate(V) (Fig. 24). This finding led the authors to discuss their role in the catalytic mechanism concluding that, similarly to what is observed in the proteins, in addition to the RNA molecule itself the water molecules help to stabilize the transition state [431]. 4.8.3. Helicases Helicases, present in all forms of life, bind double stranded nucleic acids, either DNA or RNA, separating them into single chains in a process associated with the ATP hydrolysis [432,433]. Helicases participate in different processes as replication, DNA repair, recombination, transcription and have been considered as putative targets of anti-cancer or anti-viral drugs [434–436]. The NS3 helicase from the flavivirus dengue was structurally characterized using XRD [434–437]. This enzyme combines a protease activity in the N-terminal domain and a RNA-stimulated nucleoside triphosphatase activity in the C-terminal region. The hydrolytic activity is necessary for the unwinding of the viral RNA prior to amplification [438]. High resolution structures of the catalytic domain were obtained in the presence of single-stranded RNA, AMPPNP and ADP derivatives with vanadate(V) and/or Mn2+ . In the presence of AMPPNP (PDB: 2JLR) [439], the enzyme is trapped in an enzyme-substrate ternary complex showing important features of the active site, namely the presence of a water molecule relevant for catalysis. This water is hydrogen bonded to Glu285 and Gln456 of two conserved motifs and is suggested to attack the ␥-phosphate during dephosphorylation [439].

When ADP–vanadate(V)-Mn2+ system was studied, a ternary complex was also obtained (PDB: 2JLX) mimicking the enzyme transition state [439]. In this 2.2 A˚ resolution structure, one of the apical O-ligands of the vanadate(V) occupies exactly the same

Fig. 24. Structural representation of the vanadate(V) adduct formed with a hairpin ribozyme (PDB: 2P7E) [431]. The two water molecules are represented by red ˚ spheres. Some bond lengths are indicated in A.

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position as the catalytic water. Furthermore, the 5-coordination shell is further fulfilled with three O-atoms in the equatorial plane and ADP in the remaining apical position. In the same study other complex structures provide experimental evidence for the structural determinants of NS3 in virus assembly opening the way towards the design of new inhibitors as drug candidates to fight viral propagation [439]. 5. Miscellaneous 5.1. Phosphonoacetate hydrolase Phosphonates, widely found in biological systems, are used as carbon and phosphorus sources for several microorganisms. The breaking of the carbon–phosphorous bond is accomplished via enzymatic action either through hydrolase or lyase activity [440]. The difference between the two distinct classes is that, in the first, an electron withdrawing group is required next to the C P bond for activity. Phosphonoacetate hydrolase (PhnA) belongs to this class, converting phosphonoacetate into acetate and inorganic phosphorus [441]. The high degree of sequence homology of PhnA with alkaline phosphatases is in agreement with the structural similarities found in the available 3D models. In fact, apart from some structural determinants that account for substrate specificity, PhnA can be considered a member of the alkaline phosphatase superfamily [442–444]. Following a PhnA structure from Pseudomonas fluorescens (PDB: 1EI6) [444], other similar structures from Sinorhizobium meliloti 1021 were obtained including a PhnA complexed with vanadate(V) structure at 1.8 A˚ resolution (PDB: 3T00) [445]. The PhnA active site comprises two Zn2+ ions, similar to alkaline phosphatases, coordinated to His215, His377 and Asp211 for metal 1 (M1), and Asp29, Asp250, and His251 in the case of metal 2 (M2). The metals are 4.6 A˚ apart and, even though other divalent cations can be incorporated and promote catalysis, S. meliloti PhnA has higher specificity towards zinc [445]. For catalysis, phosphonoacetate sits in the active site, making hydrogen bond interactions with the two zinc ions (M1 and M2). The 3T00 structure represents the transition state during catalysis and a vanadate(V) moiety is located in the active site [445]. Vanadate(V) presents a distorted trigonal–bipyramidal geometry, interacting with the two metal sites via the oxygen ligands. In the apical positions V is coordinated to an O-atom (at 1.9 A˚ from M1) and the hydroxyl group of Thr68 (the nucleophile that attacks the phosphate group of the substrate). In equatorial positions, the three water molecules are not in the same plane as the V-atom but tilted towards M2. Furthermore, two of these waters are hydrogen bonded to other water molecules that are mediating their contact with the conserved residues Asn89 and Asp29, which are important for the stabilization of the ions in the active site. These watermediated interactions are not seen in the vanadate(V)-alkaline phosphatase adduct structures available, where the vanadate(V) O-atoms are directly hydrogen bonded to the protein residues. In the same work, PhnA was also crystallized in the presence of substrate and the structure of the variant T68A, confirm the importance of these residues and the geometrical constrains crucial for the hydrolysis [445]. 5.2. Chymotrypsin Proteases hydrolyze peptide bonds and are classified according to the catalytic mechanism into different groups: metallo, serine, cysteine, aspartic and threonine proteases [446,447]. Abnormal levels or mutations of such enzymes are implicated in several diseases and a lot of effort has been given to the design of efficient and selective inhibitors [448].

Fig. 25. Structural representation of the bovine chymotrypsin bound to vanadate(V) and benzohydroxamic acid (PDB: 2P8O) [449]. The three residues of the catalytic triad (Ser195, His57 and Asp102) are represented, but only the Ser195 is covalently ˚ The vanadium (dark green) coordinating bound to vanadium (bond length: 2.0 A). sphere is completed by two O-atoms of the benzohydroxamate ion and three Oatoms from vanadate(V). Ser195 and Gly193 are interacting with one of vanadate(V) ˚ O-atoms through hydrogen bonds represented by black dashes (distances in A).

Vanadate(V) is useful for such purpose mimicking the reaction transition state. In fact, a 1.5 A˚ resolution structure of bovine chymotrypsin, a serine protease, complexed with vanadate(V) and benzohydroxamic acid (PDB: 2P8O) was reported [449] corroborating the previous findings on the inhibition of these proteins by 1:1 complexes of vanadate(V) and hydroxamic acids [450]. The complex formed by vanadate(V) and benzohydroxamic acid is in the active center and the V-atom is covalently bound to the O-atom of Ser195, one of the residues of the so-called catalytic triad, along with His57 and Asp102 (Fig. 25). The V-atom presents a distorted octahedral geometry with six ligands: the mentioned serine, the carbonyl and hydroxyl oxygens of the benzohydroxamate and three O-atoms, one of which is hydrogen bonded to the backbone N-atom of Gly193 and Ser195 residues [449]. The localization of the phenyl ring of the hydroxamate in the S1 binding pocket and the hydroxamate oxygen was discussed by the authors. Globally, the study concludes that chymotrypsin and vanadate(V) are good models for studies involving protease inhibitors due to their ability to unravel the structural details of the catalytic transition states [449]. 5.3. PhoX PhoX is a phosphate monoesterase enzyme also part of the alkaline phosphatase family [451–455]. The active site of this enzyme is quite unique with two iron and three calcium ions. These ions are allegedly involved in an almost exclusive inorganic dephosphorylation mechanism. The atomic resolution structure of the native protein and complexed with ATP analogues or vanadate(V) (PDB: 3ZWU at 1.4 A˚ resolution) was recently determined from P. fluorescens [456]. The two iron and one of the calcium ions form a triangle at the bottom of the active-site cavity. The two remaining Ca2+ form a plane almost perpendicular to the former. Vanadate(V) mimics the

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and kept at 4–6 ◦ C, confirms that the VIV species detected is the protein-bound complex and not free [VIV O(pic)2 (H2 O)]. Indeed, in the absence of any interaction with HEWL, the oxidation of VIV O(pic)2 complex would be relatively fast in these conditions [5,38]. The EPR spectrum of VIV O(pic)2 -HEWL adduct is well reproduced by DFT calculations indicating that the binding of HEWL to the VIV O(pic)2 fragment makes the complex rather stable to both hydrolysis and oxidation. The very weak CD signal detected in the range 400–900 nm in the solutions of [VIV O(pic)2 (H2 O)] and HEWL confirms the binding of VIV O to chiral donor groups of the lysozyme polypeptide chain; the pattern of the CD spectrum is very similar to that measured for the monodentate coordination to VIV O2+ of a COO− belonging to an amino acid, such as L-aspartic acid [462].

6. Vanadium oligomers and proteins

Fig. 26. Structural representation of the VIV O(pic)2 –HEWL adduct [461]. A distorted octahedral geometry is observed and the V-atom, is bound to Asp52, two picolinate anions and an Ooxido atom.

five-coordinate transition state of phosphate, close to the active site. Interestingly, even though one of the irons is coordinated to a cysteine residue (Cys179), this has no redox activity. The metal content of these enzymes is still a debate and expression might be regulated by the metal availability in the environment [456]. 5.4. Lysozyme Hen egg white lysozyme (HEWL) has been used as a model of metal complexes–protein interaction studies; in particular, the complexation by HEWL of several metal-based drugs, such as Pt [457], Ru [458] and Au compounds [459], was examined in the literature. Recently, the interaction between [VIV O(pic)2 (H2 O)], a V compound inhibitor of fatty acid mobilization and active in the treatment of STZ-induced diabetic rats [460], was studied by X-ray crystallography and spectroscopic techniques (PBD: 4C3W) [461]. The COO– group of the Asp52 residue replaces the equatorial water ligand forming a distorted octahedral geometry (Fig. 26). Asn46 residue is close to the complex, hydrogen bonded to the Ooxido atom. This was the first example of VIV –protein binding confirmed by single-crystal XRD [461]. The relatively long distance obtained for V O bond in the refined XRD structure (1.82 A˚ in contrast with 1.57–1.65 A˚ for an usual VIV O bond [250]), is the result of the exposure of the crystals to the very intense X-ray beam, which induced the reduction of VIV to VIII and the associated V O bond elongation. Similar reduction processes might explain anomalous structural data previously reported in VV -containing proteins [211]. DFT results confirm the binding of the COO− of Asp52 residue of HEWL to the VIV O(pic)2 moiety and predicted a VIV O bond of 1.60 A˚ in the soaked crystals. The presence of the intense EPR signal several days after the preparation of a solution of [VIV O(pic)2 (H2 O)]-HEWL, at pH ∼4

In aqueous solutions, different vanadate(V) species may occur simultaneously, either in true equilibrium or in a metastable state namely, monovanadate(V) (V1 ), divanavadate (V2 ), tetravanadate (V4 ) and decavanadate (V10 ). Additionally, each of them has different states of protonation and/or conformation. Even when it is recognized that each of these vanadate(V) species may affect proteins differently, namely influencing distinctively enzyme activities, often this variety of species is not accounted for in biological studies, neither the studies are done in exactly reproducible conditions. As the chemical equilibrium between the vanadate(V) oligomers may take place in a time scale of milliseconds (or not take much longer), it may be difficult to properly identify the vanadate(V) species responsible for the bio-activity measured. Even when techniques such as 51 V NMR are used, which allow separate detection of V1 , V2 , V4 and V10 , it may happen that some of these are not observable, but still be present and influence the activity observed. It may also happen that a certain species does not form in the absence of a protein, but in its presence some type of particular interaction established might stabilize it. In solutions containing vanadium(V) at pH 7, decavanadate (V10 ) is not expected to be thermodynamically stable, but because it decomposes very slowly, depending on how the solution was prepared, significant amounts may be present for many hours. Di- (V2 ) and tetravanadates (V4 ) may form at pH 7, but at low total V concentrations they will be much less important than monovanadate(V) (V1 ). Below 10 ␮M, concentrations rarely surpassed in vivo, monovanadate(V) is expected to be, by far, the major VV -species. Notwithstanding, there are many examples in the literature showing that several types of biological effects may indeed be due to vanadate(V) oligomers, namely to decavanadates [207,337,340,463–470]. This subject has been extensively addressed by the group of Aureliano, and particularly we recommend the reviews from 2009 [336] and 2014 [337], where an account of the interactions of proteins with vanadate(V) oligomers is also made, encompassing V2 , V4 and V10 structures [470]. We also recommend the chapters in books by Stankiewicz et al. [207,468]. There are indeed many studies reporting the interaction of V-oligomers with proteins, a few examples were described throughout the text, and we will only give here a short account of a few others. With divanadate, there are reports of e.g. mild competitive inhibition of glucose-6-phosphate dehydrogenase [471,472] and 1,6-bisphosphate fructose aldolase from rabbit muscle [473] and a non-competitive inhibiton of rabbit muscle glycerol-3-phosphate dehydrogenase [474]. As mentioned above, phosphoglycerate mutase (PGM), one of the enzymes involved in glycolysis, is potently inhibited by glucose-6-phosphate and glucose-1-phosphate in the presence of vanadate(V) [310]. The structure of the E. coli cofactor dPGM

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(dephospho form of PGM) complexed with vanadate(V) was determined (PDB: 1E59) [318]: vanadate(V) is bound in the highly basic active site, mainly as V2 , but with evidence of additional vanadate(V) moieties at either end. The contacts between the vanadate(V) and the protein consist of hydrogen-bonds and electrostatic interactions. The dissociation constant of V2 from PGM was determined by 51 V NMR as 4.0 ␮M [475]. Concerning tetravanadate, more than 20 years ago it was shown that it is a more potent inhibitor than V1 for 6-phosphogluconate dehydrogenase from several sources [472,476]. This investigation was followed up with a study of glucose-6-phosphate dehydrogenase [472] and of several other enzymes [207,474,473,477,478] such as phosphoglycerate mutase [479,480]. Since then many observations have followed, reporting the specific interactions between various proteins and different vanadate(V) oligomers, namely cases of inhibition of some of these enzymes [481,482]. 51 V NMR studies revealed tight binding of two V anions to super4 oxide dismutase (SOD), where probably Lys residues are involved in binding [483]. Decavanadate is a closed ellipsoidal cluster anion that forms in mildly acidic medium, kinetically stable for several hours at pH ∼7. Because of its large size and negative charge, it is likely, and it was confirmed, that it will bind at polyanionic (e.g. polyphosphate) binding sites of enzymes or receptors, either in the substrate domain or in an allosteric effector sites, rather than at the active site of enzymes. These interactions differ from those occurring for monovanadate(V) [336,337]. A 5 -nucleotidase from rat kidney is 16-fold activated by 0.10 ␮M of V10 , and half-maximal activation is observed at 1.4 nM of decavanadate; it was postulated that V10 binds at the ATP/2 ,3 biphosphoglycerate binding site [484]. Decavanadate has also been considered to top activate several G-protein dependent systems [468]. Ribonuclease A is competitively inhibited by decavanadate [467]; probably by interacting with the complex MutSADP–Mg, decavanadate acts by producing a steric hindrance of the protein, blocking ATP/ADP exchange [485]. V1 and V10 appear to act through distinct inhibition mechanisms with respect to the actomyosin ATPase activity and V10 was suggested for use as a biochemical tool in studies of muscle regulation [336,362,363,366]. The interaction of decavadates with kinases was reviewed by Stankiewicz et al. [207], and also discussed in the reviews of Aureliano [336,337]. Globally, unlike monovanadate(V), V10 is a good competitive inhibitor of several kinases, e.g. hexokinase, adenylate kinase, bacterial phosphofructokinase [486], mammalian phosphofructokinase [487,488] and fructose-6-P 2-kinase; however, V10 does not inhibit several other kinases such as e.g. galacto kinase, glycerol kinase and pyruvate kinase [486]. Further evidence for the interaction of decavanadate(V) at polyphosphate binding sites is the allosteric activation of 5 -nucleotidase at the site which recognizes ATP and 2,3diphosphoglycerate [484] and the inhibition of the SR ATPase [489,490]. Decavanadate induces the formation of the intermediate myosin-MgATP-V10 complex blocking the contractile cycle, probably in the pre-hydrolysis state [362], and apparently V10 is the only species capable of preventing the release of the products during the mechanism of ATP hydrolysis by the actomyosin complex [363]. Decavanadate(V) may also interact with F-actin filaments [463]; while V1 may stabilize them through the formation of F-actinADP-V1 complexes and induce actin polymerization by inhibiting specific tyrosine phosphatases [491], in contrast, decavanadate by interacting with actin (thereby stabilizing V10 species) inhibits Gactin polymerization [463]. In addition to V1 , it was also reported that other vanadate(V) oligomers, such as V10 interact with the SR Ca2+ -pump [489], but at a site distinct from the phosphorylation

site [9]. In fact it was shown that while most ion pump inhibitors bind mainly to the E2 ion pump conformation within the membrane domain from the extracellular side, thereby blocking the Ca2+ release, in contrast, decavanadate(V) interacts with Ca2+ -ATPase near the nucleotide binding site domain or at a pocket involving several cytoplasmic domains. For this purpose V10 needs to cross through the membrane bilayer [9]. In contrast to V1 , which only binds to the E2 conformation, V10 binds to all protein conformations, i.e. E1, E1P, E2 and E2P. Additionally, the specific interaction of decavanadate with SR Ca2+ -ATPase is non-competitive with respect to ATP and induces protein cysteine oxidation with concomitant vanadium reduction; this observation might explain the high inhibitory capacity of V10 . Vanadate(V) oligomers, e.g. V4 and V10 , prevent G-actin polymerization more potently than V1 [362]. Namely, decavanadate(V), at concentrations of ∼70 ␮M, inhibits 50% of the extension of actin polymerization, whereas no effects were observed for V1 up to 2 mM [362]. Moreover, monomeric actin (G-actin) stabilizes V10 species by increasing its half-life time against decomposition, from 5 to ∼25 h [362]. Globally, the biological role of decavanadate may partly result by its capacity to interact with actin and to affect several biological processes where actin may be involved. It was concluded that [492]: (i) V10 and VIV O2+ inhibit actin polymerization at ␮M concentrations; (ii) only interaction of V10 with actin induces cysteine oxidation and formation of VIV , these effects being prevented by ATP; (iii) decavanadate and VIV O2+ induce actin conformational changes affecting the protein ATP binding site; (iv) actin has high affinity binding sites for VIV O2+ . Although there is some support for the in vivo action of decavanadates [337], this subject is still controversial. Namely, it is not clear if for higher organisms such as mammals the given orally decavanadate will reach the blood stream, and if administered by injection it can be up-taken by the cells.

7. Final remarks Vanadium-containing proteins and interaction of vanadium compounds with proteins are extensively studied topics described and discussed in several previous reviews over the past two decades. Throughout this review the most relevant aspects in the various areas are discussed and the readers are referred elsewhere if they require more detailed information. As general reference reports we recommend the review by Crans et al. [5] for the work reported before 2004, and reviews [34] for structural discussions mainly concerning phosphatases. In the present review the objective was to give an overview of vanadium–protein interactions, but it was beyond the scope of this work to give a systematic account of all studies published. Vanadium is a transition metal widely distributed in soil, crude oil, water and air, thus quite available for living beings. Therefore, it is not surprising that natural selection made use of it for relevant roles in biological systems. It is not proven, but is highly probable that vanadium is an essential element for humans; however, it was shown that it is essential to several high organisms, namely some mammals [493]. There are several examples of organisms, e.g. bacteria, fungi and macroalgae which contain haloperoxidases that require vanadate(V) for their function, using vanadium for activity or function of proteins. In these enzymes vanadate(V) is in a coordination geometry akin of those found when inhibiting acid phosphatases. In contrast, in vanadium-containing nitrogenases, which exist in a range of microorganisms and catalyze the conversion of atmospheric N2 to ammonia, the active center where the final reduction of N2 takes place, vanadium is inserted in a double cubane, VFe7 [␮6 C]S9 , bound to a histidine residue.

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Other organisms have extremely efficient and selective systems for uptake and transport of vanadium. Well known examples discussed are the ascidians which accumulate high levels of vanadium from seawater, reducing it to VIII and store it in the so-called vanadocytes. Several proteins that take part in the processes of storage and reduction of vanadium, vanabins being a family of vanadium binding proteins involved in these processes. Polychaetes are also marine organisms and particularly the tube-dwelling fan worms of the Sabellidae family also accumulate high amounts of vanadium in vacuoles of the epithelial cells. High amounts of vanadium were shown to bind to several protein fractions, but not much is known about these proteins or the function of vanadium in these organisms. One of the most relevant features of vanadate(V), and the most thoroughly studied, has its origin on the chemical similarities between vanadate(V) and phosphate, and the ability of vanadate(V) to interfere with the metabolic processes involving Ca2+ and Mg2+ . This, in connection with the ability of vanadium to undergo changes in coordination geometry, allows V to influence the function of a large variety of phosphate-metabolizing enzymes. Due to the ease in forming low energy phosphate-transition state analogues, vanadates(V) are able to inhibit several types of enzymes such as ribonucleases, mutases and phosphatases, as well as many ATPases, kinases, lyases and synthases. Thus, vanadate(V) salts and compounds have been frequently used either as inhibitors of these enzymes, or as probes to study the mechanisms of their reactions and catalytic cycle. In this review we made a comprehensive overview of the many examples so far reported and, as described throughout the text, vanadate(V) is in a key position to interact effectively with many enzymes, thus enabling a wide range of bioactivity for this anion. Although, as discussed, vanadate(V) is not a perfect transition state analogue, few or no other suitable alternatives exist, namely with similar high binding affinities. Thus, enzymologists and bioinorganic chemists have been exploring this analogy for many years with success, obtaining relevant structural and mechanistic data. Besides the discovery of several other activities of V complexes, such as that observed in the photochemically induced cleavage of proteins at one specific amino acid residue by vanadate(V), studies have also demonstrated that many enzymes may have dual functions in biological systems, for example, haloperoxidase activity for other enzymes, e.g. phosphatases, to which vanadate(V) was added. The fact that the addition of vanadium can change the action of a protein, other than simply inhibiting it, may have far-reaching implications regarding how vanadium affects biological systems. This possibility, together with the vanadate(V)phosphate analogy, further potentiates the idea that what is known is only the tip of the iceberg and that vanadium may probably have relevant functions in living beings that involve interaction or incorporation of the metal ion and/or its compounds with several proteins. Which are exactly the most relevant roles of vanadium compounds in higher organisms are areas requiring better understanding. Studies of the effects of vanadium compounds on living systems have been abundant in the last years, reflecting the interest that vanadium ions might have for their environment impact and prospective applications in therapeutics [494]. Prediction of the in vivo effect of these compounds is not straightforward as their interactions with enzymes disseminate to so many distinct systems and at so many levels of the cell cycle [31]. The prospective application of vanadium compounds as therapeutics also gave rise to many publications namely addressing the interactions of these compounds with many proteins [5,31]. How vanadium may be transported in blood and up-taken by cells are particularly relevant issues in this respect, and this deserved a throughout discussion (section 2.3).

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There are promising clinical applications of vanadium compounds [496]. However, the complexity of their effects and the lack of knowledge in some areas such as their effects on e.g. the immune system and inflammatory processes, or in their use in longterm administration therapies, and of the risks associated with their potential carcinogenic effect and disorders in the free-radical background in cells, require a much deeper understanding and evaluation of the balance between beneficial and toxic effects of their use [31]. Nevertheless, the fact that many vanadium compounds are able to model enzyme reactions and show potential as therapeutic agents [28,31,494–496], is encouraging many biological investigations, potentiating the discovery of new types of reactivity. Thus, investigations into vanadium chemistry in biological systems are likely to continue active in the future. Acknowledgements The authors would like to thank Fundac¸ão para a Ciência e Tecnologia (FCT) for funding and Investigador FCT; grants (M.F.A.S.), PEst-C/EQB/LA0006/2013, SFRH/BD/77894/2011 and UID/QUI/00100/2013]. E. Garribba thanks Fondazione Banco di Sardegna for the financial support (project Prot. U924.2014/AI.807.MGB; Prat. 2014.0443). The authors also want to thank Hugo D. Correia for his help in figures preparation. References [1] R.J.P. Williams, J.J.R. Fraústo da Silva, The Natural Selection of the Chemical Elements – The Environment and Life’s Chemistry, Oxford University Press, Oxford, 1996. [2] R.J.P. Williams, J.J.R. Fraústo da Silva, The Chemistry of Evolution – The Development of our Ecosystem, Elsevier, Amsterdam, 2006. [3] B.R. Nechay, J.P. Saunders, J. Environ. Pathol. Toxicol. 2 (1978) 283–290. [4] C.C. McLauchlan, J.D. Hooker, M.A. Jones, Z. Dymon, E.A. Backhus, B.A. Greiner, N.A. Dorner, M.A. Youkhana, L.M. Manus, J. Inorg. Biochem. 104 (2010) 274–281. [5] D.C. Crans, J.J. Smee, E. Gaidamauskas, L. Yang, Chem. Rev. 104 (2004) 849–902. [6] J. Krzystek, A. Ozarowski, J. Telser, D.C. Crans, Coord. Chem. Rev. (2014), http:// dx.doi.org/10.1016/j.ccr.2014.1010.1014 [7] A. Tracey, G.R. Willsky, E.S. Takeuchi, Vanadium: Chemistry, Biochemistry, Pharmacology and Practical Applications, CRC Press, Boca Raton, 2007. [8] E.F. Pettersen, T.D. Goddard, C.C. Huang, G.S. Couch, D.M. Greenblatt, E.C. Meng, T.E. Ferrin, UCSF chimera – a visualization system for exploratory research and analysis, J. Comp. Chem. 25 (2004) 1605–1612. [9] M. Aureliano, G. Fraqueza, C.A. Ohlin, Dalton Trans. 42 (2013) 11770–11777. [10] D. Rehder, Future Med. Chem. 4 (2012) 1823–1837. [11] Y. Yoshikawa, H. Sakurai, D.C. Crans, G. Micera, E. Garribba, Dalton Trans. 43 (2014) 6965–6972. [12] T.A.S. Brandao, A.C. Hengge, S.J. Johnson, J. Biol. Chem. 285 (2010) 15874–15883. [13] Z. Lu, D. Dunaway-Mariano, K.N. Allen, Proc. Natl. Acad. Sci. U. S. A. 105 (2008) 5687–5692. [14] L.L. Madan, B. Gopal, Biochemistry 50 (2011) 10114–10125. [15] K.H. Thompson, J.H. McNeill, C. Orvig, Chem. Rev. 99 (1999) 2561–2571. [16] K.H. Thompson, C. Orvig, J. Chem. Soc. Dalton Trans. (2000) 2885–2892. [17] H. Sakurai, Y. Kojima, Y. Yoshikawa, K. Kawabe, H. Yasui, Coord. Chem. Rev. 226 (2002) 187–198. [18] H. Sakurai, J. Fugono, H. Yasui, Mini-Rev. Med. Chem. 4 (2004) 41–48. [19] R.R. Eady, Coord. Chem. Rev. 237 (2003) 23–30. [20] P. Hulley, A. Davison, J. Trace Elem. Exp. Med. 16 (2003) 281–290. [21] Y. Shechter, I. Goldwaser, M. Mironchik, M. Fridkin, D. Gefel, Coord. Chem. Rev. 237 (2003) 3–11. [22] D. Rehder, Angew. Chem. Int. Ed. Engl. 30 (1991) 148–167. [23] H. Sakurai, Y. Yoshikawa, H. Yasui, Chem. Soc. Rev. 37 (2008) 2383–2392. [24] G.R. Willsky, L.H. Chi, M. Godzala, P.J. Kostyniak, J.J. Smee, A.M. Trujillo, J.A. Alfano, W.J. Ding, Z.H. Hu, D.C. Crans, Coord. Chem. Rev. 255 (2011) 2258–2269. [25] J. Costa Pessoa, I. Tomaz, Curr. Med. Chem. 17 (2010) 3701–3738. [26] T. Kiss, T. Jakusch, D. Hollender, A. Dornyei, E.A. Enyedy, J. Costa Pessoa, H. Sakurai, A. Sanz-Medel, Coord. Chem. Rev. 252 (2008) 1153–1162. [27] D. Wischang, O. Brucher, J. Hartung, Coord. Chem. Rev. 255 (2011) 2204–2217. [28] D. Gambino, Coord. Chem. Rev. 255 (2011) 2193–2203. [29] W. Plass, Coord. Chem. Rev. 255 (2011) 2378–2387. [30] T. Jakusch, J. Costa Pessoa, T. Kiss, Coord. Chem. Rev. 255 (2011) 2218–2226. [31] J. Korbecki, I. Baranowska-Bosiacka, I. Gutowska, D. Chlubek, Acta Biochim. Pol. 59 (2012) 195–200.

82

J. Costa Pessoa et al. / Coordination Chemistry Reviews 301–302 (2015) 49–86

[32] D. Rehder, Dalton Trans. 42 (2013) 11749–11761. [33] S.R. Akabayov, B. Akabayov, Inorg. Chim. Acta 420 (2014) 16–23. [34] D.C. Crans, M.L. Tarlton, C.C. McLauchlan, Eur. J. Inorg. Chem. (2014) 4450–4468. [35] C.C. McLauchlan, B.J. Peters, G.R.Willsky, D.C. Crans, (2014), http://dx.doi.org/ 10.1016/j.ccr.2014.12.012 [36] T. Ueki, H. Michibata, Coord. Chem. Rev. 255 (2011) 2249–2257. [37] H. Michibata, T. Ueki, High levels of vanadium in ascidians, in: H. Michibata (Ed.), Vanadium: Biochemical and Molecular Biological Approaches, Springer, New York, 2012, pp. 51–71. [38] D. Rehder, Bioinorganic Vanadium Chemistry, John Wiley & Sons, Ltd., Chichester, 2008. [39] T. Ueki, N. Yamaguchi, Romaidi, Y. Isago, H. Tanahashi, Coord. Chem. Rev. (2015), http://dx.doi.org/10.1016/j.ccr.2014.09.007 [40] H. Michibata, Y. Iwata, J. Hirata, J. Exp. Zool. 257 (1991) 306–313. [41] P.C. Cole, J.M. Eckert, K.L. Williams, Anal. Chim. Acta 153 (1983) 61–67. [42] R.W. Collier, Nature 309 (1984) 441–444. [43] H. Kitayama, S. Yamamoto, H. Michibata, T. Ueki, Dalton Trans. 42 (2013) 11921–11925. [44] T. Hamada, M. Asanuma, T. Ueki, F. Hayashi, N. Kobayashi, S. Yokoyama, H. Michibata, H. Hirota, J. Am. Chem. Soc. 127 (2005) 4216–4222. [45] K. Fukui, T. Ueki, H. Ohya, H. Michibata, J. Am. Chem. Soc. 125 (2003) 6352–6353. [46] T. Ueki, T. Adachi, S. Kawano, M. Aoshima, N. Yamaguchi, K. Kanamori, H. Michibata, BBA-Gene Struct. Expr. 1626 (2003) 43–50. [47] N. Kawakami, T. Ueki, Y. Amata, K. Kanamori, K. Matsuo, K. Gekko, H. Michibata, BBA – Proteins Proteomoics 1794 (2009) 674–679. [48] T. Ueki, N. Kawakami, M. Toshishige, K. Matsuo, K. Gekko, H. Michibata, BBA Gen. Subj. 1790 (2009) 1327–1333. [49] A.D. Tinoco, C.W. Peterson, B. Lucchese, R.P. Doyle, A.M. Valentine, Proc. Natl. Acad. Sci. U. S. A. 105 (2008) 3268–3273. [50] J.P. Gaffney, A.M. Valentine, Biochemistry 48 (2009) 11609–11611. [51] A.L. Dingley, K. Kustin, I.G. Macara, G.C. Mcleod, BBA – Proteins Proteomics 649 (1981) 493–502. [52] M. Yoshihara, T. Ueki, N. Yamaguchi, K. Kamino, H. Michibata, BBA Gen. Subj. 1780 (2008) 256–263. [53] Y. Abe, R. Nagata, Y. Hasunuma, H. Yokosawa, Comp. Biochem. Physiol. B 128 (2001) 73–79. [54] L.A. Lambert, H. Perri, T.J. Meehan, Comp. Biochem. Physiol. B 140 (2005) 11–25. [55] L.A. Lambert, S.L. Mitchell, J. Mol. Evol. 64 (2007) 113–128. [56] D.C. Crans, B. Zhang, E. Gaidamauskas, A.D. Keramidas, G.R. Willsky, C.R. Roberts, Inorg. Chem. 49 (2010) 4245–4256. [57] J. Costa Pessoa, I. Tomaz, T. Kiss, P. Buglyó, J. Inorg. Biochem. 84 (2001) 259–270. [58] J. Costa Pessoa, I. Tomaz, T. Kiss, E. Kiss, P. Buglyó, J. Biol. Inorg. Chem. 7 (2002) 225–240. [59] P. Frank, E.J. Carlson, R.M.K. Carlson, B. Hedman, K.O. Hodgson, J. Inorg. Biochem. 102 (2008) 809–823. [60] T. Ishii, I. Nakai, C. Numako, K. Okoshi, T. Otake, Discovery of a new vanadium accumulator, the fan worm pseudopotamilla-occelata, Naturwissenschaften 80 (1993) 268–270. [61] J.D. Popham, J.M. Dauria, Mar. Pollut. Bull. 13 (1982) 25–27. [62] T. Ishii, T. Otake, K. Okoshi, M. Nakahara, R. Nakamura, Mar. Biol. 121 (1994) 143–151. [63] D. Fattorini, A. Notti, M. Nigro, F. Regoli, Environ. Sci. Pollut. Res. 17 (2010) 220–228. [64] T. Ishii, Characterization of vanadium in the fan worms, in: J.O. Nriagu (Ed.), Vanadium in the Environment, John Wiley & Sons, New York, 1998, pp. 199–216. [65] D. Fattorini, F. Regoli, Hyper-accumulation of vanadium in polychaetes, in: H. Michibata (Ed.), Vanadium: Biochemical and Molecular Biological Approaches, Springer, Dordrecht, 2012, pp. 73–92. [66] R.T. MacGillivray, E. Mendez, J.G. Shewale, S.K. Sinha, J. Lineback-Zins, K. Brew, J. Biol. Chem. 258 (1983) 3543–3553. [67] R. Crichton, Iron Metabolism – From Molecular Mechanisms to Clinical Consequences, 3rd ed., John Wiley & Sons, Ltd., Chichester, 2009. [68] H. Sun, M. Cox, H. Li, P. Sadler, Struct. Bond. 88 (1997) 71–102. [69] H.Z. Sun, H.Y. Li, P.J. Sadler, Chem. Rev. 99 (1999) 2817–2842. [70] P.T. Gomme, K.B. McCann, J. Bertolini, Drug Discov. Today 10 (2005) 267–273. [71] J. Wally, S. Buchanan, Biometals 20 (2007) 249–262. [72] B.F. Andersen, H.M. Baker, G.E. Morris, S.V. Rumball, E.N. Baker, Nature 344 (1990) 784–787. [73] N.D. Chasteen, Coord. Chem. Rev. 22 (1977) 1–36. [74] W.R. Harris, Clin. Chem. 38 (1992) 1809–1818. [75] B.E. Eckenroth, A.N. Steere, N.D. Chasteen, S.J. Everse, A.B. Mason, Proc. Natl. Acad. Sci. U. S. A. 108 (2011) 13089–13094. [76] M.H. Nagaoka, H. Akiyama, T. Maitani, Analyst 129 (2004) 51–54. [77] M.H. Nagaoka, T. Yamazaki, T. Maitani, Biochem. Biophys. Res. Commun. 296 (2002) 1207–1214. [78] K.G. Fernandes, M. Montes-Bayon, E. Blanco Gonzalez, E. Del Castillo-Busto, J.A. Nobrega, A. Sanz-Medel, J. Anal. At. Spectrom. 20 (2005) 210–215. [79] I. Bertini, G. Canti, C. Luchinat, Inorg. Chim. Acta 67 (1982) L21–L23. [80] I. Bertini, C. Luchinat, L. Messori, J. Inorg. Biochem. 25 (1985) 57–60. [81] J. Costa Pessoa, G. Gonc¸alves, S. Roy, I. Correia, S. Mehtab, M.F.A. Santos, T. Santos-Silva, Inorg. Chim. Acta 420 (2014) 60–68.

[82] C.A. Smith, E.W. Ainscough, A.M. Brodie, J. Chem. Soc. Dalton Trans. (1995) 1121–1126. [83] D.C. Harris, Biochemistry 16 (1977) 560–564. [84] D. Sanna, P. Buglyó, G. Micera, E. Garribba, J. Biol. Inorg. Chem. 15 (2010) 825–839. [85] I. Correia, T. Jakusch, E. Cobbinna, S. Mehtab, I. Tomaz, N.V. Nagy, A. Rockenbauer, J. Costa Pessoa, T. Kiss, Dalton Trans. 41 (2012) 6477–6487. [86] T. Jakusch, A. Dean, T. Oncsik, A.C. Benyei, V. Di Marco, T. Kiss, Dalton Trans. 39 (2010) 212–220. [87] D.C. Crans, R.L. Bunch, L.A. Theisen, J. Am. Chem. Soc. 111 (1989) 7597–7607. [88] D. Sanna, G. Micera, E. Garribba, Inorg. Chem. 52 (2013) 11975–11985. [89] D. Sanna, M. Serra, G. Micera, E. Garribba, Inorg. Chem. 53 (2014) 1449–1464. [90] J.C. Cannon, N.D. Chasteen, Biochemistry 14 (1975) 4573–4577. [91] N.D. Chasteen, L.K. White, R.F. Campbell, Biochemistry 16 (1977) 363–368. [92] L.K. White, N.D. Chasteen, J. Phys. Chem. 83 (1979) 279–284. [93] N.D. Chasteen, Vanadyl(IV) EPR spin probe. Inorganic and biochemical aspects, in: L.J.J. Berliner, J. Reuben (Eds.), Biological Magnetic Resonance, Plenum Press, New York, 1981, pp. 53–119. [94] N.D. Chasteen, J.K. Grady, C.E. Holloway, Inorg. Chem. 25 (1986) 2754–2760. [95] D. Mustafi, E.V. Galtseva, J. Krzystek, L.-C. Brunel, M.W. Makinen, J. Phys. Chem. A 103 (1999) 11279–11286. [96] G.R. Willsky, A.B. Goldfine, P.J. Kostyniak, J.H. McNeill, L.Q. Yang, H.R. Khan, D.C. Crans, J. Inorg. Biochem. 85 (2001) 33–42. [97] B.D. Liboiron, K.H. Thompson, G.R. Hanson, E. Lam, N. Aebischer, C. Orvig, J. Am. Chem. Soc. 127 (2005) 5104–5115. [98] T. Kiss, T. Jakusch, S. Bouhsina, H. Sakurai, É.A. Enyedy, Eur. J. Inorg. Chem. 2006 (2006) 3607–3613. [99] D. Sanna, E. Garribba, G. Micera, J. Inorg. Biochem. 103 (2009) 648–655. [100] D. Sanna, G. Micera, E. Garribba, Inorg. Chem. 48 (2009) 5747–5757. [101] J. Mazurier, J.-M. Lhoste, J. Montreuil, G. Spik, BBA-Protein Struct. Mol. Enzymol. 745 (1983) 44–49. [102] J.D. Casey, N.D. Chasteen, J. Inorg. Biochem. 13 (1980) 111–126. [103] J.D. Casey, N.D. Chasteen, J. Inorg. Biochem. 13 (1980) 127–136. [104] G. Justino, E. Garribba, J. Costa Pessoa, J. Biol. Inorg. Chem. 18 (2013) 803–813. [105] R.F. Campbell, N.D. Chasteen, J. Biol. Chem. 252 (1977) 5996–6001. [106] N.D. Chasteen, Vanadium–protein interactions, in: H. Sigel, A. Sigel (Eds.), Vanadium and its Role in Life, Marcel Dekker, New York, 1995, pp. 231–248. [107] S.S. Eaton, J. Dubach, K.M. More, G.R. Eaton, G. Thurman, D.R. Ambruso, J. Biol. Chem. 264 (1989) 4776–4781. [108] T. Kiss, E. Kiss, E. Garribba, H. Sakurai, J. Inorg. Biochem. 80 (2000) 65–73. [109] T. Jakusch, D. Hollender, E.A. Enyedy, C.S. Gonzalez, M. Montes-Bayon, A. SanzMedel, J. Costa Pessoa, I. Tomaz, T. Kiss, Dalton Trans. (2009) 2428–2437. [110] P. Aisen, A. Leibman, J. Zweier, J. Biol. Chem. 253 (1978) 1930–1937. [111] H. Kurokawa, B. Mikami, M. Hirose, J. Mol. Biol. 254 (1995) 196–207. [112] A. Carmichael, J.S. Vincent, FEBS Lett. 105 (1979) 349–352. [113] K.H. Thompson, C. Orvig, Coord. Chem. Rev. 219–221 (2001) 1033–1053. [114] K.H. Thompson, C. Orvig, J. Inorg. Biochem. 100 (2006) 1925–1935. [115] K.H. Thompson, J. Lichter, C. LeBel, M.C. Scaife, J.H. McNeill, C. Orvig, J. Inorg. Biochem. 103 (2009) 554–558. [116] T. Kiss, T. Jakusch, D. Hollender, A. Dornyei, in: K. Kustin, J. Costa Pessoa, D.C. Crans (Eds.), Vanadium: The Versatile Metal, ACS Symp. Ser. 974, Washington, DC, 2007, p. 323. [117] D. Sanna, G. Micera, E. Garribba, Inorg. Chem. 49 (2010) 174–187. [118] D. Sanna, L. Biro, P. Buglyo, G. Micera, E. Garribba, Metallomics 4 (2012) 33–36. [119] D. Sanna, L. Bíró, P. Buglyó, G. Micera, E. Garribba, J. Inorg. Biochem. 115 (2012) 87–99. [120] S. Mehtab, G. Gonc¸alves, S. Roy, A.I. Tomaz, T. Santos-Silva, M.F.A. Santos, M.J. Romão, T. Jakusch, T. Kiss, J. Costa Pessoa, J. Inorg. Biochem. 121 (2013) 187–195. [121] G. Gonc¸alves, A.I. Tomaz, I. Correia, L.F. Veiros, M.M.C.A. Castro, F. Avecilla, L. Palacio, M. Maestro, T. Kiss, T. Jakusch, M.H.V. Garcia, J. Costa Pessoa, Dalton Trans. 42 (2013) 11841–11861. [122] D. Sanna, V. Ugone, G. Micera, E. Garribba, Dalton Trans. 41 (2012) 7304–7318. [123] T. Koleˇsa-Dobravc, E. Lodyga-Chruscinska, M. Symonowicz, D. Sanna, A. Meden, F. Perdih, E. Garribba, Inorg. Chem. 53 (2014) 7960–7976. [124] J. Wally, P.J. Halbrooks, C. Vonrhein, M.A. Rould, S.J. Everse, A.B. Mason, S.K. Buchanan, J. Biol. Chem. 281 (2006) 24934–24944. [125] N. Noinaj, N.C. Easley, M. Oke, N. Mizuno, J. Gumbart, E. Boura, A.N. Steere, O. Zak, P. Aisen, E. Tajkhorshid, R.W. Evans, A.R. Gorringe, A.B. Mason, A.C. Steven, S.K. Buchanan, Nature 483 (2012) 53–58. [126] X. Yang, K. Wang, J. Lu, D.C. Crans, Coord. Chem. Rev. 237 (2003) 103–111. [127] W.R. Harris, C.J. Carrano, J. Inorg. Biochem. 22 (1984) 201–218. [128] W.R. Harris, Biochemistry 24 (1985) 7412–7418. [129] A. Butler, H. Eckert, J. Am. Chem. Soc. 111 (1989) 2802–2809. [130] J.A. Saponja, H.J. Vogel, J. Inorg. Biochem. 62 (1996) 253–270. [131] A.-K. Bordbar, A.L. Creagh, F. Mohammadi, C.A. Haynes, C. Orvig, J. Inorg. Biochem. 103 (2009) 643–647. [132] C. Harford, B. Sarkar, Acc. Chem. Res. 30 (1997) 123–130. [133] E.L. Giroux, M. Durieux, P.J. Schechter, Bioorg. Chem. 5 (1976) 211–218. [134] J.W. Foote, H.T. Delves, Analyst 109 (1984) 709–711. ´ [135] M. Sokołowska, M. Wszelaka-Rylik, J. Poznanski, W. Bal, J. Inorg. Biochem. 103 (2009) 1005–1013. [136] W. Bal, J. Christodoulou, P.J. Sadler, A. Tucker, J. Inorg. Biochem. 70 (1998) 33–39. [137] A.J. Stewart, C.A. Blindauer, S. Berezenko, D. Sleep, P.J. Sadler, Proc. Natl. Acad. Sci. U. S. A. 100 (2003) 3701–3706.

J. Costa Pessoa et al. / Coordination Chemistry Reviews 301–302 (2015) 49–86 [138] E.O. Martins, T. Drakenberg, Inorg. Chim. Acta 67 (1982) 71–74. [139] P.J. Sadler, J.H. Viles, Inorg. Chem. 35 (1996) 4490–4496. [140] J. Lu, A.J. Stewart, P.J. Sadler, T.J.T. Pinheiro, C.A. Blindauer, Biochem. Soc. Trans. 36 (2008) 1317–1321. [141] J. Christodoulou, P.J. Sadler, A. Tucker, Eur. J. Biochem. 225 (1994) 363–368. [142] N.D. Chasteen, J. Francavilla, J. Phys. Chem. 80 (1976) 867–871. [143] K. Fukui, H. Ohya-Nishiguchi, M. Nakai, H. Sakurai, H. Kamada, FEBS Lett. 368 (1995) 31–35. [144] M. Purcell, J.F. Neault, H. Malonga, H. Arakawa, H.A. Tajmir-Riahi, Can. J. Chem. 79 (2001) 1415–1421. [145] E. Cobbina, S. Mehtab, I. Correia, G. Gonc¸alves, I. Tomaz, I. Cavaco, T. Jakusch, E. Enyedi, T. Kiss, J. Costa Pessoa, J. Mex. Chem. Soc. 57 (2013) 180–191. [146] H. Yasui, Y. Kunori, H. Sakurai, Chem. Lett. 32 (2003) 1032–1033. [147] E. Garribba, G. Micera, E. Lodyga-Chruscinska, D. Sanna, G. Sanna, Eur. J. Inorg. Chem. 2005 (2005) 4953–4963. [148] D. Sanna, G. Micera, E. Garribba, Inorg. Chem. 50 (2011) 3717–3728. [149] M.W. Makinen, M. Salehitazangi, Coord. Chem. Rev. 279 (2014) 1–22. [150] G. Heinemann, B. Fichtl, M. Mentler, W. Vogt, J. Inorg. Biochem. 90 (2002) 38–42. [151] D.M. Dias, J.P.G.L.M. Rodrigues, N.S. Domingues, A.M.J.J. Bonvin, M.M.C.A. Castro, Eur. J. Inorg. Chem. 2013 (2013) 4619–4627. [152] R.G. Hamilton, Clin. Chem. 33 (1987) 1707–1725. [153] K. De Cremer, M. Van Hulle, C. Chéry, R. Cornelis, K. Strijckmans, R. Dams, N. Lameire, R. Vanholder, J. Biol. Inorg. Chem. 7 (2002) 884–890. [154] S. Nishihara, A. Shimizu, Y. Arata, Mol. Immunol. 23 (1986) 285–290. [155] D. Sanna, V. Pecoraro, G. Micera, E. Garribba, J. Biol. Inorg. Chem. 17 (2012) 773–790. [156] A.J.M. Ribeiro, M.E. Alberto, M.J. Ramos, P.A. Fernandes, N. Russo, Chem.-Eur. J. 19 (2013) 14081–14089. [157] G. Micera, E. Garribba, Dalton Trans. (2009) 1914–1918. [158] G. Micera, V.L. Pecoraro, E. Garribba, Inorg. Chem. 48 (2009) 5790–5796. [159] G. Micera, E. Garribba, Eur. J. Inorg. Chem. (2010) 4697–4710. [160] E. Lodyga-Chruscinska, G. Micera, E. Garribba, Inorg. Chem. 50 (2011) 883–899. [161] G. Micera, E. Garribba, J. Comput. Chem. 32 (2011) 2822–2835. [162] G. Micera, E. Garribba, Eur. J. Inorg. Chem. (2011) 3768–3780. [163] D. Sanna, K. Varnagy, N. Lihi, G. Micera, E. Garribba, Inorg. Chem. 52 (2013) 8202–8213. [164] J. Schaller, S. Gerber, U. Kämpfer, S. Lejon, C. Trachsel, Human Blood Plasma Proteins: Structure and Function, John Wiley & Sons Ltd., Chichester, 2008. [165] L.C. Cantley, M.D. Resh, G. Guidotti, Nature 272 (1978) 552–554. [166] L.C. Cantley, P. Aisen, J. Biol. Chem. 254 (1979) 1781–1784. [167] I.G. Macara, K. Kustin, L.C. Cantley Jr., BBA Gen. Subj. 629 (1980) 95–106. [168] A. Heinz, K.A. Rubinson, J.J. Grantham, J. Lab. Clin. Med. 100 (1982) 593–612. [169] T.V. Hansen, J. Aaseth, J. Alexander, Arch. Toxicol. 50 (1982) 195–202. [170] M. Bruech, M.E. Quintanilla, W. Legrum, J. Koch, K.J. Netter, G.F. Fuhrmann, Toxicology 31 (1984) 283–295. [171] M. Garner, J. Reglinski, W.E. Smith, J. McMurray, I. Abdullah, R. Wilson, J. Biol. Inorg. Chem. 2 (1997) 235–241. [172] T.C. Delgado, A.I. Tomaz, I. Correia, J. Costa Pessoa, J.G. Jones, C.F.G.C. Geraldes, M.M.C.A. Castro, J. Inorg. Biochem. 99 (2005) 2328–2339. [173] D. Sanna, M. Serra, G. Micera, E. Garribba, Inorg. Chim. Acta 420 (2014) 75–84. [174] R.J. DeKoch, D.J. West, J.C. Cannon, N.D. Chasteen, Biochemistry 13 (1974) 4347–4354. [175] T.-Y. Fang, M. Zou, V. Simplaceanu, N.T. Ho, C. Ho, Biochemistry 38 (1999) 13423–13432. [176] P.M. Harrison, P. Arosio, BBA – Bioenergetics 1275 (1996) 161–203. [177] J.K. Grady, J. Shao, P. Arosio, P. Santambrogio, N.D. Chasteen, J. Inorg. Biochem. 80 (2000) 107–113. [178] N.D. Chasteen, E.M. Lord, H.J. Thompson, J.K. Grady, BBA-Gen. Subj. 884 (1986) 84–92. [179] N.D. Chasteen, E.C. Theil, J. Biol. Chem. 257 (1982) 7672–7677. [180] J.G. Wardeska, B. Viglione, N.D. Chasteen, J. Biol. Chem. 261 (1986) 6677–6683. [181] P.M. Hanna, N.D. Chasteen, G.A. Rottman, P. Aisen, Biochemistry 30 (1991) 9210–9216. [182] G.J. Gerfen, P.M. Hanna, N.D. Chasteen, D.J. Singel, J. Am. Chem. Soc. 113 (1991) 9513–9519. [183] O. Einsle, F.A. Tezcan, S.L.A. Andrade, B. Schmid, M. Yoshida, J.B. Howard, D.C. Rees, Science 297 (2002) 1696–1700. [184] R.R. Eady, Vanadium nitrogenases, in: N.D. Chasteen (Ed.), Vanadium in Biological Systems, Kluwer, Dordrecht, 1990, pp. 99–128. [185] R.R. Eady, in: H. Sigel, A. Sigel (Eds.), Vanadium and its Role in Life, Marcel Dekker, Inc., New York, 1995, p. 363. [186] C. Rüttimann-Johnson, R. Chatterjee, V.K. Shah, P.W. Ludden, in: A.S. Tracey, D.C. Crans (Eds.), Vanadium Compounds: Chemistry, Biochemistry, and Therapeutic Applications, ACS, Washington, 1998, p. 228. [187] A.W. Fay, M.A. Blank, C.C. Lee, Y.L. Hu, K.O. Hodgson, B. Hedman, M.W. Ribbe, J. Am. Chem. Soc. 132 (2010) 12612–12618. [188] D. Rehder, J. Inorg. Biochem. 80 (2000) 133–136. [189] V.K. Shah, W.J. Brill, Proc. Natl. Acad. Sci. U. S. A. 74 (1977) 3249–3253. [190] G.N. George, C.L. Coyle, B.J. Hales, S.P. Cramer, J. Am. Chem. Soc. 110 (1988) 4057–4059. [191] J.L. Zuo, H.C. Zhou, R.H. Holm, Inorg. Chem. 42 (2003) 4624–4631. [192] J.A. Wiig, Y.L. Hu, C.C. Lee, M.W. Ribbe, Science 337 (2012) 1672–1675.

[193] [194] [195] [196] [197] [198]

[199] [200] [201] [202] [203]

[204]

[205]

[206]

[207] [208] [209] [210] [211] [212] [213] [214] [215] [216]

[217] [218] [219] [220] [221] [222] [223] [224] [225] [226] [227]

[228] [229] [230] [231] [232] [233] [234] [235] [236] [237] [238] [239] [240] [241]

83

Y.L. Hu, C.C. Lee, M.W. Ribbe, Science 333 (2011) 753–755. M.J. Dilworth, R.R. Eady, Biochem. J. 277 (1991) 465–468. H. Vilter, Phytochemistry 23 (1984) 1387–1390. R. Wever, B.E. Krenn, in: N.D. Chasteen (Ed.), Vanadium in Biological Systems, Physiology and Biochemistry, Kluwer, Dordrecht, 1990, p. 81. R. Wever, W. Hemrika, in: J.O. Nriagu (Ed.), Vanadium in the Environment, Jonh Wiley and Sons, New York, 1998, p. 285. A. Messerschmidt, L. Prade, R. Wever, in: A.S. Tracey, D.C. Crans (Eds.), Vanadium Compounds: Chemistry, Biochemistry, and Therapeutic Applications, ACS, Washington, 1998, pp. 186–201. H. Vilter, in: H. Sigel, A. Sigel (Eds.), Vanadium and its Role in Life, Marcel Dekker, New York, 1995, p. 325. D. Rehder, G. Santoni, G.M. Licini, C. Schulzke, B. Meier, Coord. Chem. Rev. 237 (2003) 53–63. R. Wever, in: H. Michibata (Ed.), Biochemical and Molecular Biological Approaches, Springer, Dordrecht, 2012, p. 95. G. Licini, V. Conte, A. Coletti, M. Mba, C. Zonta, Coord. Chem. Rev. 255 (2011) 2345–2357. J. Littlechild, E. Garcia-Rodriguez, E. Coupe, A. Watts, M. Isupov, in: K. Kustin, J. Costa Pessoa, D.C. Crans (Eds.), Vanadium: The Versatile Metal, ACS Symp. Ser. 974, Washington, DC, 2007, p. 136. C.J. Schneider, G. Zampella, L. DeGioa, V.L. Pecoraro, in: K. Kustin, J. Costa Pessoa, D.C. Crans (Eds.), Vanadium: The Versatile Metal, ACS Symp. Ser. 974, Washington, DC, 2007, p. 148. W. Plass, M. Bangesh, S. Nica, A. Buchholz, in: K. Kustin, J. Costa Pessoa, D.C. Crans (Eds.), Vanadium: The Versatile Metal, ACS Symp. Ser. 974, Washington, DC, 2007, p. 163. T. Polenova, N. Pooransingh-Margolis, D. Rehder, R. Renirie, R. Wever, in: K. Kustin, J. Costa Pessoa, D.C. Crans (Eds.), Vanadium: The Versatile Metal, ACS Symp. Ser. 974, Washington, DC, 2007, p. 178. P.J. Stankiewicz, A.S. Tracey, D.C. Crans, in: H. Sigel, A. Sigel (Eds.), Vanadium and its Role in Life, Marcel Dekker, New York, 1995, p. 287. M. Weyand, H.J. Hecht, M. Kiess, M.F. Liaud, H. Vilter, D. Schomburg, J. Mol. Biol. 293 (1999) 595–611. D. Wischang, J. Hartung, Tetrahedron 67 (2011) 4048–4054. A. Messerschmidt, R. Wever, Proc. Natl. Acad. Sci. U. S. A. 93 (1996) 392–396. A. Messerschmidt, L. Prade, R. Wever, Biol. Chem. 378 (1997) 309. S. Macedo-Ribeiro, W. Hemrika, R. Renirie, R. Wever, A. Messerschmidt, J. Biol. Inorg. Chem. 4 (1999) 209–219. P. Bernhardt, T. Okino, J.M. Winter, A. Miyanaga, B.S. Moore, J. Am. Chem. Soc. 133 (2011) 4268–4270. M.N. Isupov, A.R. Dalby, A.A. Brindley, Y. Izumi, T. Tanabe, G.N. Murshudov, J.A. Littlechild, J. Mol. Biol. 299 (2000) 1035–1049. E. Garcia-Rodriguez, T. Ohshiro, T. Aibara, Y. Izumi, J. Littlechild, J. Biol. Inorg. Chem. 10 (2005) 275–282. J.B. Fournier, E. Rebuffet, L. Delage, R. Grijol, L. Meslet-Cladiere, J. Rzonca, P. Potin, G. Michel, M. Czjzek, C. Leblanc, Appl. Environ. Microbiol. 80 (2014) 7561–7573. A. Messerschmidt, R. Wever, Inorg. Chim. Acta 273 (1998) 160–166. R. Renirie, W. Hemrika, S.R. Piersma, R. Wever, Biochemistry 39 (2000) 1133–1141. C. Kimblin, X.H. Bu, A. Butler, Inorg. Chem. 41 (2002) 161–163. N. Tanaka, Z. Hasan, R. Wever, Inorg. Chim. Acta 356 (2003) 288–296. M. Bangesh, W. Plass, J. Mol. Struct.-Theochem. 725 (2005) 163–175. J.Y. Kravitz, V.L. Pecoraro, Pure Appl. Chem. 77 (2005) 1595–1605. S. Raugei, F.L. Gervasio, P. Carloni, Phys. Status Solidi B 243 (2006) 2500–2515. A.F. Neuwald, Protein Sci. 6 (1997) 1764–1767. J. Stukey, G.M. Carman, Protein Sci. 6 (1997) 469–472. W. Hemrika, R. Renirie, H.L. Dekker, P. Barnett, R. Wever, Proc. Natl. Acad. Sci. U. S. A. 94 (1997) 2145–2149. W. Hemrika, R. Renerie, H. Dekker, R. Wever, in: A.S. Tracey, D.C. Crans (Eds.), Vanadium Compounds: Chemistry, Biochemistry and Therapeutic Applications, ACS, Washington, 1998, p. 216. W. Hemrika, R. Wever, FEBS Lett. 409 (1997) 317–319. K. Ishikawa, Y. Mihara, K. Gondoh, E. Suzuki, Y. Asano, EMBO J. 19 (2000) 2412–2423. C.J. Pan, K.J. Lei, B. Annabi, W. Hemrika, J.Y. Chou, J. Biol. Chem. 273 (1998) 6144–6148. J. Littlechild, E. Garcia-Rodriguez, A. Dalby, M. Isupov, J. Mol. Recognit. 15 (2002) 291–296. Y. Lindqvist, G. Schneider, P. Vihko, Eur. J. Biochem. 221 (1994) 139–142. F. van de Velde, I.W.C.E. Arends, R.A. Sheldon, J. Inorg. Biochem. 80 (2000) 81–89. N. Tanaka, V. Dumay, Q.N. Liao, A.J. Lange, R. Wever, Eur. J. Biochem. 269 (2002) 2162–2167. R. Renirie, W. Hemrika, R. Wever, J. Biol. Chem. 275 (2000) 11650–11657. D. Rehder, Org. Biomol. Chem. 6 (2008) 957–964. A.N. Antipov, D.Y. Sorokin, N.P. L’Vov, J.G. Kuenen, Biochem. J. 369 (2003) 185–189. A.N. Antipov, N.N. Lyalikova, N.P. L’vov, IUBMB Life 49 (2000) 137–141. W. Carpentier, L. De Smet, J. Van Beeumen, A. Brige, J. Bacteriol. 187 (2005) 3293–3301. Z.Y. Zhang, Crit. Rev. Biochem. Mol. Biol. 33 (1998) 1–52. D.C. Crans, in: A.S. Tracey, D.C. Crans (Eds.), Vanadium Compounds: Chemistry, Biochemistry, and Therapeutic Applications, ACS, Washington, 1998, p. 82.

84

J. Costa Pessoa et al. / Coordination Chemistry Reviews 301–302 (2015) 49–86

[242] B.I. Posner, C.R. Yang, A. Shaver, in: A.S. Tracey, D.C. Crans (Eds.), Vanadium Compounds: Chemistry, Biochemistry, and Therapeutic Applications, ACS, Washington, 1998, p. 316. [243] L.F. Chang, M. Karin, Nature 410 (2001) 37–40. [244] V.O. Van Hoof, M.E. De Broe, Crit. Rev. Clin. Lab. Sci. 31 (1994) 197–293. [245] H. Bull, P.G. Murray, D. Thomas, A.M. Fraser, P.N. Nelson, J. Clin. Pathol.-Mol. Pathol. 55 (2002) 65–72. [246] K.H.W. Lau, J.R. Farley, D.J. Baylink, Biochem. J. 257 (1989) 23–36. [247] N.K. Tonks, C.D. Diltz, E.H. Fischer, Adv. Proteins Phosphatases 4 (1987) 5. [248] C.L. Shriner, D.L. Brautigan, J. Biol. Chem. 259 (1984) 1383–1390. [249] K.G. Peters, M.G. Davis, B.W. Howard, M. Pokross, V. Rastogi, C. Diven, K.D. Greis, E. Eby-Wilkens, M. Maier, A. Evdokimov, S. Soper, F. Genbauffe, J. Inorg. Biochem. 96 (2003) 321–330. [250] L.F. Vilas Boas, J. Costa Pessoa, Vanadium, in: G. Wilkinson, R.D. Gillard, J.A. McCleverty (Eds.), Comprehensive Coordination Chemistry, Pergamon Press, Oxford, 1987, pp. 453–583. [251] (a) J. Costa Pessoa, L.F.V. Boas, R.D. Gillard, R.J. Lancashire, Polyhedron 7 (1988) 1245–1262; (b) J. Costa Pessoa, (2015) http://dx.doi.org/10.1016/j.jinorgbio.2015.03.004. [252] J.M. Sowadski, M.D. Handschumacher, H.M.K. Murthy, B.A. Foster, H.W. Wyckoff, J. Mol. Biol. 186 (1985) 417–433. [253] E.E. Kim, H.W. Wyckoff, J. Mol. Biol. 218 (1991) 449–464. [254] D. Barford, A.J. Flint, N.K. Tonks, Science 263 (1994) 1397–1404. [255] H. Deng, R. Callender, Z.H. Huang, Z.Y. Zhang, Biochemistry 41 (2002) 5865–5872. [256] T.A.S. Brandão, H. Robinson, S.J. Johnson, A.C. Hengge, J. Am. Chem. Soc. 131 (2009) 778–786. [257] J.M. Denu, D.L. Lohse, J. Vijayalakshmi, M.A. Saper, J.E. Dixon, Proc. Natl. Acad. Sci. U. S. A. 93 (1996) 2493–2498. [258] A.G. Evdokimov, M. Pokross, R. Walter, M. Mekel, B. Cox, C.Y. Li, R. Bechard, F. Genbauffe, R. Andrews, C. Diven, B. Howard, V. Rastogi, J. Gray, M. Maier, K.G. Peters, Acta Crystallogr. D 62 (2006) 1435–1445. [259] M. Zhang, M. Zhou, R.L. VanEtten, C.V. Stauffacher, Biochemistry 36 (1997) 15–23. [260] Z.C. Jia, D. Barford, A.J. Flint, N.K. Tonks, Science 268 (1995) 1754–1758. [261] A.D.B. Pannifer, A.J. Flint, N.K. Tonks, D. Barford, J. Biol. Chem. 273 (1998) 10454–10462. [262] K.M. Holtz, B. Stec, E.R. Kantrowitz, J. Biol. Chem. 274 (1999) 8351–8354. [263] K. Ostanin, E.H. Harms, P.E. Stevis, R. Kuciel, M.M. Zhou, R.L. Vanetten, J. Biol. Chem. 267 (1992) 22830–22836. [264] R.L. Felts, T.J. Reilly, J.J. Tanner, J. Biol. Chem. 281 (2006) 30289–30298. [265] A.P. Bevan, P.G. Drake, J.F. Yale, A. Shaver, B.I. Posner, Mol. Cell. Biochem. 153 (1995) 49–58. [266] D.C. Crans, A.D. Keramidas, C. Drouza, Phosphorus Sulfur 109 (1996) 245–248. [267] B.I. Posner, R. Faure, J.W. Burgess, A.P. Bevan, D. Lachance, G.Y. Zhangsun, I.G. Fantus, J.B. Ng, D.A. Hall, B.S. Lum, A. Shaver, J. Biol. Chem. 269 (1994) 4596–4604. [268] D.C. Crans, M.S. Gottlieb, J. Tawara, R.L. Bunch, L.A. Theisen, Anal. Biochem. 188 (1990) 53–64. [269] P.A.M. Williams, D.A. Barrio, S.B. Etcheverry, J. Inorg. Biochem. 75 (1999) 99–104. [270] S.B. Etcheverry, D.A. Barrio, P.A.M. Williams, E.J. Baran, Biol. Trace Elem. Res. 84 (2001) 227–238. [271] A.S. Tracey, J. Inorg. Biochem. 80 (2000) 11–16. [272] M. Vieth, R.E. Higgs, D.H. Robertson, M. Shapiro, E.A. Gragg, H. Hemmerle, BBA – Proteins Proteomics 1697 (2004) 243–257. [273] I. Akritopoulou-Zanze, P.J. Hajduk, Drug Discov. Today 14 (2009) 291–297. [274] A.S. Arvai, Y. Bourne, M.J. Hickey, J.A. Tainer, J. Mol. Biol. 249 (1995) 835–842. [275] E.B. Lansdon, I.H. Segel, A.J. Fisher, BBA – Proteins Proteomics 1814 (2011) 1349–1357. [276] S.K. Srivastava, K. Rajasree, B. Gopal, Biochim. Biophys. Acta 1814 (2011) 1349–1357. [277] H.H. Dar, P.K. Chakraborti, Biochem. J. 430 (2010) 539–549. [278] J. Janin, C. Dumas, S. Morera, Y.W. Xu, P. Meyer, M. Chiadmi, J. Cherfils, J. Bioenergy Biomembr. 32 (2000) 215–225. [279] S. Pakhomova, S.G. Bartlett, P.A. Doerner, M.E. Newcomer, Biochemistry 50 (2011) 6909–6919. [280] S. Kobayashi, T. Kuzuyama, H. Seto, Antimicrob. Agents Chemother. 44 (2000) 647–650. [281] R.D. Woodyer, Z.Y. Shao, P.M. Thomas, N.L. Kelleher, J.A.V. Blodgett, W.W. Metcalf, W.A. Van der Donk, H.M. Zhao, Chem. Biol. 13 (2006) 1171–1182. [282] S. Pakhomova, S.G. Bartlett, A. Augustus, T. Kuzuyama, M.E. Newcomer, J. Biol. Chem. 283 (2008) 28518–28526. [283] Y. Kakuta, E.V. Petrotchenko, L.C. Pedersen, M. Negishi, J. Biol. Chem. 273 (1998) 27325–27330. [284] Y. Kakuta, L.G. Pedersen, C.W. Carter, M. Negishi, L.C. Pedersen, Nat. Struct. Biol. 4 (1997) 904–908. [285] H.R. Evans, D.E. Holloway, J.M. Sutton, J. Ayriss, C.C. Shone, K.R. Acharya, Acta Crystallogr. D 60 (2004) 1502–1505. [286] T.T. Caradoc-Davies, S.M. Cutfield, I.L. Lamont, J.F. Cutfield, J. Mol. Biol. 337 (2004) 337–354. [287] M. Campagnolo, C. Campa, R. De Zorzi, J. Wuerges, S. Geremia, Arch. Biochem. Biophys. 471 (2008) 11–19. [288] Z.B. Lu, L.B. Wang, D. Dunaway-Mariano, K.N. Allen, J. Biol. Chem. 284 (2009) 1224–1233.

[289] K.D. Daughtry, H. Huang, V. Malashkevich, Y. Patskovsky, W.F. Liu, U. Ramagopal, J.M. Sauder, S.K. Burley, S.C. Almo, D. Dunaway-Mariano, K.N. Allen, Biochemistry 52 (2013) 5372–5386. [290] H. Zimmermann, M. Zebisch, N. Strater, Purin. Sign 8 (2012) 437–502. [291] G.G. Yegutkin, BBA-Mol. Cell Res. 1783 (2008) 673–694. [292] S.C. Robson, J. Sévigny, H. Zimmermann, Purin. Sign 2 (2006) 409–430. [293] G. Kauffenstein, A. Drouin, N. Thorin-Trescases, H. Bachelard, B. Robaye, P. D’Orleans-Juste, F. Marceau, E. Thorin, J. Sevigny, Cardiovasc. Res. 85 (2010) 204–213. [294] M. Mandapathil, M.J. Szczepanski, M. Szajnik, J. Ren, D.E. Lenzner, E.K. Jackson, E. Gorelik, S. Lang, J.T. Johnson, T.L. Whiteside, Clin. Cancer Res. 15 (2009) 6348–6357. [295] A.F. Knowles, Purin. Sign 7 (2011) 21–45. [296] F.M. Sansom, S.C. Robson, E.L. Hartland, Microbiol. Mol. Biol. Rev. 72 (2008) 765. [297] M. al-Rashida, J. Iqbal, Med. Res. Rev. 34 (2014) 703–743. [298] M. Zebisch, N. Strater, Proc. Natl. Acad. Sci. U. S. A. 105 (2008) 6882–6887. [299] M. Zebisch, M. Krauss, P. Schafer, N. Strater, J. Mol. Biol. 415 (2012) 288–306. [300] M.N. Munkonda, J. Pelletier, V.V. Ivanenkov, M. Fausther, A. Tremblay, B. Kunzli, T.L. Kirley, J. Sevigny, FEBS J. 276 (2009) 479–496. [301] M. Zebisch, M. Krauss, P. Schafer, P. Lauble, N. Strater, Structure 21 (2013) 1460–1475. [302] D.R. Davies, H. Interthal, J.J. Champoux, W.G.J. Hol, J. Mol. Biol. 324 (2002) 917–932. [303] D.R. Davies, H. Interthal, J.J. Champoux, W.G.J. Hol, Chem. Biol. 10 (2003) 139–147. [304] S.W. Yang, A.B. Burgin, B.N. Huizenga, C.A. Robertson, K.C. Yao, H.A. Nash, Proc. Natl. Acad. Sci. U. S. A. 93 (1996) 11534–11539. [305] D.R. Davies, H. Interthal, J.J. Champoux, W.G.J. Hol, J. Med. Chem. 47 (2004) 829–837. [306] A. Hofmann, M. Grella, I. Botos, W. Filipowicz, A. Wlodawer, J. Biol. Chem. 277 (2002) 1419–1425. [307] P. Genschik, J. Hall, W. Filipowicz, J. Biol. Chem. 272 (1997) 13211–13219. [308] A. Hofmann, A. Zdanov, P. Genschik, S. Ruvinov, W. Filipowicz, A. Wlodawer, EMBO J. 19 (2000) 11. [309] J.G. Zalatan, T.D. Fenn, A.T. Brunger, D. Herschlag, Biochemistry 45 (2006) 9788–9803. [310] W.J. Ray, C.B. Post, Biochemistry 29 (1990) 2779–2789. [311] M.D. Percival, K. Doherty, M.J. Gresser, Biochemistry 29 (1990) 2764–2769. [312] H.I. Fraser, M. Kvaratskhelia, M.F. White, FEBS Lett. 455 (1999) 344–348. [313] L.A. Fothergill-Gilmore, H.C. Watson, Relat. Areas Mol. Biol. 62 (1989) 227–313. [314] R.A. Walter, J. Nairn, D. Duncan, N.C. Price, S.M. Kelly, D.J. Rigden, L.A. Fothergill-Gilmore, Biochem. J. 337 (1999) 89–95. [315] M.J. Jedrzejas, Prog. Biophys. Mol. Biol. 73 (2000) 263–287. [316] D.J. Rigden, R.A. Walter, S.E.V. Phillips, L.A. Fothergill-Gilmore, J. Mol. Biol. 286 (1999) 1507–1517. [317] C.S. Bond, M.F. White, W.N. Hunter, J. Biol. Chem. 276 (2001) 3247–3253. [318] C.S. Bond, M.F. White, W.N. Hunter, J. Mol. Biol. 316 (2002) 1071–1081. [319] D.R. Davies, B.L. Staker, J.A. Abendroth, T.E. Edwards, R. Hartley, J. Leonard, H. Kim, A.L. Rychel, S.N. Hewitt, P.J. Myler, L.J. Stewart, Acta Crystallogr. F 67 (2011) 1044–1050. [320] K. Kustin, G.C. Mcleod, T.R. Gilbert, L.R. Briggs, Struct. Bond. 53 (1983) 139–160. [321] R. Wever, K. Kustin, Adv. Inorg. Chem. 35 (1990) 81. [322] N.D. Chasteen, Struct. Bond. 53 (1983) 105–138. [323] K. Kustin, W.E. Robinson, Metal Ions Biol. Syst. 31 (1995) 511–542. [324] B.R. Nechay, L.B. Nanninga, P.S.E. Nechay, R.L. Post, J.J. Grantham, I.G. Macara, L.F. Kubena, T.D. Phillips, F.H. Nielsen, Fed. Proc. 45 (1986) 123–132. [325] D.W. Boyd, K. Kustin, Adv. Inorg. Chem. 6 (1984) 5. [326] L.C. Cantley, L. Josephson, R. Warner, M. Yanagisawa, C. Lechene, G. Guidotti, J. Biol. Chem. 252 (1977) 7421–7423. [327] M. Aureliano, J. Inorg. Biochem. 80 (2000) 145–147. [328] L. Beauge, Methods Enzymol. 156 (1988) 251–267. [329] Z.E. Sauna, M.M. Smith, M. Muller, S.V. Ambudkar, J. Biol. Chem. 276 (2001) 21199–21208. [330] L. Josephson, L.C. Cantley, Biochemistry 16 (1977) 4572–4578. [331] L.C. Cantley, L.G. Cantley, L. Josephson, J. Biol. Chem. 253 (1978) 7361–7368. [332] M. Aureliano, V.M.C. Madeira, BBA-Mol. Cell. Res. 1221 (1994) 259–271. [333] P.L. Pedersen, E. Carafoli, Trends Biochem. Sci. 12 (1987) 146–150. [334] G. Fraqueza, C.A. Ohlin, W.H. Casey, M. Aureliano, J. Inorg. Biochem. 107 (2012) 82–89. [335] G. Fraqueza, L.A.E.B. de Carvalho, M.P.M. Marques, L. Maia, C.A. Ohlin, W.H. Casey, M. Aureliano, Dalton Trans. 41 (2012) 12749–12758. [336] M. Aureliano, D.C. Crans, J. Inorg. Biochem. 103 (2009) 536–546. [337] M. Aureliano, C.A. Ohlin, J. Inorg. Biochem. 137 (2014) 123–130. [338] V.V. Guliants, Catal. Today 51 (1999) 255–268. [339] B.K. Hodnett, Catal. Rev. 27 (1985) 373–424. [340] C. Slebodnick, B.J. Hamstra, V.L. Pecoraro, Struct. Bond. 89 (1997) 51–108. [341] A. Leeeiford, R.A. Ow, I.R. Gibbons, J. Biol. Chem. 261 (1986) 2337–2342. [342] A. Muhlrad, I. Ringel, Metal Ions Biol. Syst. 31 (1995) 211–230. [343] I.R. Gibbons, G. Mocz, Vanadate sensitized photocleavage of proteins, in: N.D. Chasteen (Ed.), Vanadium in Biological Systems, Physiology and Biochemistry, Kluwer, Dordrecht, 1990, pp. 143–152. [344] H.L. Sweeney, A. Houdusse, Annu. Rev. Biophys. 39 (2010) 539–557. [345] S. Loubery, E. Coudrier, Cell. Mol. Life Sci. 65 (2008) 2790–2800.

J. Costa Pessoa et al. / Coordination Chemistry Reviews 301–302 (2015) 49–86 [346] B.J. Foth, M.C. Goedecke, D. Soldati, Proc. Natl. Acad. Sci. U. S. A. 103 (2006) 3681–3686. [347] F. Odronitz, M. Kollmar, Genome Biol. 8 (2007). [348] A. Sebe-Pedros, X. Grau-Bove, T.A. Richards, I. Ruiz-Trillo, Genome Biol. Evol. 6 (2014) 290–305. [349] I. Rayment, W.R. Rypniewski, K. Schmidtbase, R. Smith, D.R. Tomchick, M.M. Benning, D.A. Winkelmann, G. Wesenberg, H.M. Holden, Science 261 (1993) 50–58. [350] C.A. Smith, I. Rayment, Biochemistry 35 (1996) 5404–5417. [351] M. Kovacs, J. Toth, C. Hetenyi, A. Malnasi-Csizmadia, J.R. Sellers, J. Biol. Chem. 279 (2004) 35557–35563. [352] J.S. Allingham, R. Smith, I. Rayment, Nat. Struct. Mol. Biol. 12 (2005) 378–379. [353] C. Lucas-Lopez, J.S. Allingham, T. Lebl, C.P.A.T. Lawson, R. Brenk, J.R. Sellers, I. Rayment, N.J. Westwood, Org. Biomol. Chem. 6 (2008) 2076–2084. [354] R. Fedorov, M. Bohl, G. Tsiavaliaris, F.K. Hartmann, M.H. Taft, P. Baruch, B. Brenner, R. Martin, H.J. Knolker, H.O. Gutzeit, D.J. Manstein, Nat. Struct. Mol. Biol. 16 (2009) 80–88. [355] K. Chinthalapudi, M.H. Taft, R. Martin, S.M. Heissler, M. Preller, F.K. Hartmann, H. Brandstaetter, J. Kendrick-Jones, G. Tsiavaliaris, H.O. Gutzeit, R. Fedorov, F. Buss, H.J. Knolker, L.M. Coluccio, D.J. Manstein, J. Biol. Chem. 286 (2011) 29700–29708. [356] M. Preller, K. Chinthalapudi, R. Martin, H.J. Knolker, D.J. Manstein, J. Med. Chem. 54 (2011) 3675–3685. [357] J.J. Frye, V.A. Klenchin, C.R. Bagshaw, I. Rayment, Biochemistry 49 (2010) 4897–4907. [358] M. Kollmar, U. Durrwang, W. Kliche, D.J. Manstein, F.J. Kull, EMBO J. 21 (2002) 2517–2525. [359] S. Munnich, M.H. Taft, D.J. Manstein, J. Mol. Biol. 426 (2014) 2070–2081. [360] J. Menetrey, P. Llinas, M. Mukherjea, H.L. Sweeney, A. Houdusse, Cell 131 (2007) 300–308. [361] J. Menetrey, T. Isabet, V. Ropars, M. Mukherjea, O. Pylypenko, X.Y. Liu, J. Perez, P. Vachette, H.L. Sweeney, A.M. Houdusse, Mol. Cell 48 (2012) 75–86. [362] T. Tiago, P. Martel, C. Gutierrez-Merino, M. Aureliano, BBA – Proteins Proteomics 1774 (2007) 474–480. [363] T. Tiago, M. Aureliano, C. Gutierrez-Merino, Biochemistry 43 (2004) 5551–5561. [364] M. Aureliano, V.M.C. Madeira, Biochem. Biophys. Res. Commun. 205 (1994) 161–167. [365] K.J. Fohr, Y. Wahl, R. Engling, T.P. Kemmer, M. Gratzl, Cell Calcium 12 (1991) 735–742. [366] T. Tiago, M. Aureliano, J.J.G. Moura, J. Inorg. Biochem. 98 (2004) 1902–1910. [367] C.C. Goodno, Proc. Natl. Acad. Sci. U. S. A. 76 (1979) 2620–2624. [368] A.L. Davidson, E. Dassa, C. Orelle, J. Chen, Microbiol. Mol. Biol. Rev. 72 (2008) 317–364. [369] D.C. Rees, E. Johnson, O. Lewinson, Nat. Rev. Mol. Cell Biol. 10 (2009) 218–227. [370] M.L. Oldham, A.L. Davidson, J. Chen, Curr. Opin. Struct. Biol. 18 (2008) 726–733. [371] J. ter Beek, A. Guskov, D.J. Slotboom, J. Gen. Physiol. 143 (2014) 419–435. [372] K.P. Locher, A.T. Lee, D.C. Rees, Science 296 (2002) 1091–1098. [373] I.L. Urbatsch, B. Sankaran, J. Weber, A.E. Senior, J. Biol. Chem. 270 (1995) 19383–19390. [374] S. Sharma, A.L. Davidson, J. Bacteriol. 182 (2000) 6570–6576. [375] J. Chen, S. Sharma, F.A. Quiocho, A.L. Davidson, Proc. Natl. Acad. Sci. U. S. A. 98 (2001) 1525–1530. [376] R.N. Hvorup, B.A. Goetz, M. Niederer, K. Hollenstein, E. Perozo, K.P. Locher, Science 317 (2007) 1387–1390. [377] V.M. Korkhov, S.A. Mireku, K.P. Locher, Nature 490 (2012) 367. [378] V.M. Korkhov, S.A. Mireku, R.N. Hvorup, K.P. Locher, FEBS Lett. 586 (2012) 972–976. [379] A. Ward, C.L. Reyes, J. Yu, C.B. Roth, G. Chang, Proc. Natl. Acad. Sci. U. S. A. 104 (2007) 19005–19010. [380] M.L. Oldham, J. Chen, Proc. Natl. Acad. Sci. U. S. A. 108 (2011) 15152–15156. [381] A.I. Jonckheere, J.A.M. Smeitink, R.J.T. Rodenburg, J. Inherit. Metab. Dis. 35 (2012) 211–225. [382] D. Okuno, R. Iino, H. Noji, J. Biochem. (Tokyo, Jpn.) 149 (2011) 655–664. [383] C. von Ballmoos, G.M. Cook, P. Dimroth, Annu. Rev. Biophys. 37 (2008) 43–64. [384] C. Chen, A.K. Saxena, W.N. Simcoke, D.N. Garboczi, P.L. Pedersen, Y.H. Ko, J. Biol. Chem. 281 (2006) 13777–13783. [385] M.A. Bianchet, J. Hullihen, P.L. Pedersen, L.M. Amzel, Proc. Natl. Acad. Sci. U. S. A. 95 (1998) 11065–11070. [386] Y.H. Ko, M. Bianchet, L.M. Amzel, P.L. Pedersen, J. Biol. Chem. 272 (1997) 18875–18881. [387] Y.H. Ko, S.J. Hong, P.L. Pedersen, J. Biol. Chem. 274 (1999) 28853–28856. [388] M.S.S. Manimekalai, A. Kumar, J. Jeyakanthan, G. Gruber, J. Mol. Biol. 408 (2011) 736–754. [389] G. Gruber, M.S.S. Manimekalai, F. Mayer, V. Muller, BBA – Bioenergetics 1837 (2014) 940–952. [390] A. Kumar, M.S.S. Manimekalai, A.M. Balakrishna, J. Jeyakanthan, G. Gruber, J. Mol. Biol. 396 (2010) 301–320. [391] D.R. Davies, A. Mushtaq, H. Interthal, J.J. Champoux, W.G.J. Hol, J. Mol. Biol. 357 (2006) 1202–1210. [392] J.J. Champoux, Annu. Rev. Biochem. 70 (2001) 369–413. [393] N.M. Baker, R. Rajan, A. Mondragon, Nucleic Acids Res. 37 (2009) 693–701. [394] M.R. Redinbo, L. Stewart, P. Kuhn, J.J. Champoux, W.G.J. Hol, Science 279 (1998) 1504–1513.

85

[395] L. Stewart, M.R. Redinbo, X.Y. Qiu, W.G.J. Hol, J.J. Champoux, Science 279 (1998) 1534–1541. [396] M.R. Redinbo, J.J. Champoux, W.G.J. Hol, Biochemistry 39 (2000) 6832–6840. [397] K. Perry, Y. Hwang, F.D. Bushman, G.D. Van Duyne, Structure 18 (2010) 127–137. [398] B. Gibb, K. Gupta, K. Ghosh, R. Sharp, J. Chen, G.D. Van Duyne, Nucleic Acids Res. 38 (2010) 5817–5832. [399] P. Tumbale, J.S. Williams, M.J. Schellenberg, T.A. Kunkel, R.S. Williams, Nature 506 (2014) 111–115. [400] U. Rass, I. Ahel, S.C. West, J. Biol. Chem. 283 (2008) 33994–34001. [401] P. Tumbale, C.D. Appel, R. Kraehenbuehl, P.D. Robertson, J.S. Williams, J. Krahn, I. Ahel, R.S. Williams, Nat. Struct. Mol. Biol. 18 (2011), 1189-U. [402] K. Kobryn, G. Chaconas, Curr. Opin. Microbiol. 4 (2001) 558–564. [403] H. Aihara, W.M. Huang, T. Ellenberger, Mol. Cell 27 (2007) 901–913. [404] K. Shi, W.M. Huang, H. Aihara, PLoS Biol. 11 (2013). [405] R.T. Raines, Chem. Rev. 98 (1998) 1045–1065. [406] C.M. Cuchillo, M.V. Nogues, R.T. Raines, Biochemistry 50 (2011) 7835–7841. [407] A. Wlodawer, M. Miller, L. Sjolin, Proc. Natl. Acad. Sci. U. S. A. 80 (1983) 3628–3631. [408] B. Borah, C.W. Chen, W. Egan, M. Miller, A. Wlodawer, J.S. Cohen, Biochemistry 24 (1985) 2058–2067. [409] J.E. Ladner, B.D. Wladkowski, L.A. Svensson, L. Sjolin, G.L. Gilliland, Acta Crystallogr. D 53 (1997) 290–301. [410] J.E. Thompson, R.T. Raines, J. Am. Chem. Soc. 116 (1994) 5467–5468. [411] A. Wlodawer, L.A. Svensson, L. Sjolin, G.L. Gilliland, Biochemistry 27 (1988) 2705–2717. [412] D. Kostrewa, H.W. Choe, U. Heinemann, W. Saenger, Biochemistry 28 (1989) 7592–7600. [413] H. Yoshida, Ribonucleases A 341 (2001) 28–41. [414] C.N. Pace, U. Heinemann, U. Hahn, W. Saenger, Angew. Chem. Int. Ed. Engl. 30 (1991) 343–360. [415] R. Arni, U. Heinemann, R. Tokuoka, W. Saenger, J. Biol. Chem. 263 (1988) 15358–15368. [416] M. Nowotny, W. Yang, EMBO J. 25 (2006) 1924–1933. [417] M. Nowotny, S.A. Gaidamakov, R.J. Crouch, W. Yang, Cell 121 (2005) 1005–1016. [418] S.J. Schultz, J.J. Champoux, Virus Res. 134 (2008) 86–103. [419] M.J. Fedor, J.R. Williamson, Nat. Rev. Mol. Cell Biol. 6 (2005) 399–412. [420] P.C. Bevilacqua, R. Yajima, Curr. Opin. Chem. Biol. 10 (2006) 455–464. [421] M.J. Fedor, Annu. Rev. Biophys. 38 (2009) 271–299. [422] A. Wochner, J. Attwater, A. Coulson, P. Holliger, Science 332 (2011) 209–212. [423] N.J. Reiter, C.W. Chan, A. Mondragon, Curr. Opin. Struct. Biol. 21 (2011) 319–326. [424] J.A. Liberman, J.E. Wedekind, Curr. Opin. Struct. Biol. 21 (2011) 327–334. [425] B.L. Golden, Biochemistry 50 (2011) 9424–9433. [426] T. Tuschl, C. Gohlke, T.M. Jovin, E. Westhof, F. Eckstein, Science 266 (1994) 785–789. [427] D.M.J. Lilley, Curr. Opin. Struct. Biol. 15 (2005) 313–323. [428] P.B. Rupert, A.R. Ferre-D’Amare, Nature 410 (2001) 780–786. [429] P.B. Rupert, A.P. Massey, S.T. Sigurdsson, A.R. Ferre-D’Amare, Science 298 (2002) 1421–1424. [430] M.J. Fedor, J. Mol. Biol. 297 (2000) 269–291. [431] A.T. Torelli, J. Krucinska, J.E. Wedekind, RNA 13 (2007) 1052–1070. [432] F. Bleichert, S.J. Baserga, Mol. Cell 27 (2007) 339–352. [433] K. Buttner, S. Nehring, K.P. Hopfner, Nat. Struct. Mol. Biol. 14 (2007) 647–652. [434] J.H. Wu, A.K. Bera, R.J. Kuhn, J.L. Smith, J. Virol. 79 (2005) 10268–10277. [435] T. Xu, A. Sampath, A. Chao, D.Y. Wen, M. Nanao, P. Chene, S.G. Vasudevan, J. Lescar, J. Virol. 79 (2005) 10278–10288. [436] T.L. Yap, T. Xu, Y.L. Chen, H. Malet, M.P. Egloff, B. Canard, S.G. Vasudevan, J. Lescar, J. Virol. 81 (2007) 4753–4765. [437] D. Luo, T. Xu, C. Hunke, G. Gruber, S.G. Vasudevan, J. Lescar, J. Virol. 82 (2008) 173–183. [438] E. Jankowsky, M.E. Fairman, Curr. Opin. Struct. Biol. 17 (2007) 316–324. [439] D.H. Luo, T. Xu, R.P. Watson, D. Scherer-Becker, A. Sampath, W. Jahnke, S.S. Yeong, C.H. Wang, S.P. Lim, A. Strongin, S.G. Vasudevan, J. Lescar, EMBO J. 27 (2008) 3209–3219. [440] S.V. Kononova, M.A. Nesmeyanova, Biochemistry-Moscow 67 (2002) 184–195. [441] J.W. Mcgrath, G.B. Wisdom, G. Mcmullan, M.J. Larkin, J.P. Quinn, Eur. J. Biochem. 234 (1995) 225–230. [442] M.Y. Galperin, A. Bairoch, E.V. Koonin, Protein Sci. 7 (1998) 1829–1835. [443] A.N. Kulakova, L.A. Kulakov, J.P. Quinn, Gene 195 (1997) 49–53. [444] A. Kim, M.M. Benning, S. OkLee, J. Quinn, B.M. Martin, H.M. Holden, D. Dunaway-Mariano, Biochemistry 50 (2011) 3481–3494. [445] V. Agarwal, S.A. Borisova, W.W. Metcalf, W.A. van der Donk, S.K. Nair, Chem. Biol. 18 (2011) 1230–1240. [446] L. Hedstrom, Chem. Rev. 102 (2002) 4501–4523. [447] M.J. Page, E. Di Cera, Cell. Mol. Life Sci. 65 (2008) 1220–1236. [448] P.K. Madala, J.D.A. Tyndall, T. Nall, D.P. Fairlie, Chem. Rev. 110 (2010) Pr1–Pr31. [449] A. Moulin, J.H. Bell, R.F. Pratt, D. Ringe, Biochemistry 46 (2007) 5982–5990. [450] J.H. Bell, K. Curley, R.F. Pratt, Biochem. Biophys. Res. Commun. 274 (2000) 732–735. [451] S.C.L. Kamerlin, P.K. Sharma, R.B. Prasad, A. Warshel, Q. Rev. Biophys. 46 (2013) 1–132. [452] J.K. Lassila, J.G. Zalatan, D. Herschlag, Annu. Rev. Biochem. 80 (2011) 669–702.

86

J. Costa Pessoa et al. / Coordination Chemistry Reviews 301–302 (2015) 49–86

[453] H.W. Luo, R. Benner, R.A. Long, J.J. Hu, Proc. Natl. Acad. Sci. U. S. A. 106 (2009) 21219–21223. [454] M. Sebastian, J.W. Ammerman, ISME J. 3 (2009) 563–572. [455] A. van Mourik, N.M.C. Bleumink-Pluym, L. van Dijk, J.P.M. van Putten, M.M.S.M. Wosten, Microbiology-SGM 154 (2008) 584–592. [456] S.C. Yong, P. Roversi, J. Lillington, F. Rodriguez, M. Krehenbrink, O.B. Zeldin, E.F. Garman, S.M. Lea, B.C. Berks, Science 345 (2014) 1170–1173. [457] A. Casini, G. Mastrobuoni, C. Temperini, C. Gabbiani, S. Francese, G. Moneti, C.T. Supuran, A. Scozzafava, L. Messori, Chem. Commun. (2007) 156–158. [458] T. Santos-Silva, A. Mukhopadhyay, J.D. Seixas, G.J.L. Bernardes, C.C. Romão, M.J. Romão, J. Am. Chem. Soc. 133 (2011) 1192–1195. [459] C. Gabbiani, A. Casini, G. Kelter, F. Cocco, M.A. Cinellu, H.-H. Fiebig, L. Messori, Metallomics 3 (2011) 1318–1323. [460] H. Sakurai, K. Fujii, H. Watanabe, H. Tamura, Biochem. Biophys. Res. Commun. 214 (1995) 1095–1101. [461] M.F.A. Santos, I. Correia, A.R. Oliveira, E. Garribba, J. Costa Pessoa, T. SantosSilva, Eur. J. Inorg. Chem. (2014) 3293–3297. [462] J. Costa Pessoa, R.L. Marques, L.F.V. Boas, R.D. Gillard, Polyhedron 9 (1990) 81–98. [463] S. Ramos, M. Manuel, T. Tiago, R. Duarte, J. Martins, C. Gutierrez-Merino, J.J.G. Moura, M. Aureliano, J. Inorg. Biochem. 100 (2006) 1734–1743. [464] I. Bougie, M. Bisaillon, Biochem. J. 398 (2006) 557–567. [465] K.J. Fohr, J. Scott, G. Ahnerthilger, M. Gratzl, Biochem. J. 262 (1989) 83–89. [466] G.R. Willsky, D.A. White, B.C. Mccabe, J. Biol. Chem. 259 (1984) 3273–3281. [467] J.M. Messmore, R.T. Raines, Biochem. Biophys. 381 (2000) 25–30. [468] P.J. Stankiewicz, A.S. Tracey, in: H. Sigel, A. Sigel (Eds.), Vanadium and its Role in Life, Marcel Dekker, New York, 1995, p. 249. [469] M. Aureliano, R.M.C. Gandara, J. Inorg. Biochem. 99 (2005) 979–985. [470] M. Aureliano, V.M.C. Madeira, in: J.O. Nriagu (Ed.), Vanadium in the Environment, John Wiley & Sons, New York, 1998, p. 333. [471] D.C. Crans, S.M. Schelble, Biochemistry 29 (1990) 6698–6706. [472] D.C. Crans, E.M. Willging, S.R. Butler, J. Am. Chem. Soc. 112 (1990) 427–432. [473] D.C. Crans, K. Sudhakar, T.J. Zamborelli, Biochemistry 31 (1992) 6812–6821.

[474] D.C. Crans, C.M. Simone, Biochemistry 30 (1991) 6734–6741. [475] P.J. Stankiewicz, M.J. Gresser, A.S. Tracey, L.F. Hass, Biochemistry 26 (1987) 1264–1269. [476] D.C. Crans, C.D. Rithner, L.A. Theisen, J. Am. Chem. Soc. 112 (1990) 2901–2908. [477] D.C. Crans, M. Mahrooftahir, O.P. Anderson, M.M. Miller, Inorg. Chem. 33 (1994) 5586–5590. [478] D.C. Crans, C.M. Simone, A.K. Saha, R.H. Glew, Biochem. Biophys. Res. Commun. 165 (1989) 246–250. [479] A.S. Tracey, M.J. Gresser, S. Liu, J. Am. Chem. Soc. 110 (1988) 5869–5874. [480] M.J. Gresser, A.S. Tracey, K.M. Parkinson, J. Am. Chem. Soc. 108 (1986) 6229–6234. [481] A.S. Tracey, M.J. Gresser, Can. J. Chem. 66 (1988) 2570–2574. [482] P.J. Stankiewicz, L.F. Hass, Fed. Proc. 45 (1986) 1649. [483] L. Wittenkeller, A. Abraha, R. Ramasamy, D.M. de Freitas, L.A. Theisen, D.C. Crans, J. Am. Chem. Soc. 113 (1991) 7872–7881. [484] M. Lehir, Biochem. J. 273 (1991) 795–798. [485] R.J. Pezza, M.A. Villarreal, G.G. Montich, C.E. Argarana, Nucleic Acids Res. 30 (2002) 4700–4708. [486] D.W. Boyd, K. Kustin, M. Niwa, Biochim. Biophys. Acta 827 (1985) 472–475. [487] G. Choate, T.E. Mansour, J. Biol. Chem. 254 (1979) 1457–1462. [488] G.L. Choate, T.E. Mansour, Fed. Proc. 37 (1978) 1433. [489] P. Csermely, A. Martonosi, G.C. Levy, A.J. Ejchart, Biochem. J. 230 (1985) 807–815. [490] C. Coan, D.J. Scales, A.J. Murphy, J. Biol. Chem. 261 (1986) 394–403. [491] E. Brener, S. Rubinstein, G. Cohen, K. Shternall, J. Rivlin, H. Breitbart, Biol. Reprod. 68 (2003) 837–845. [492] S. Ramos, J.J.G. Moura, M. Aureliano, Metallomics 4 (2012) 16–22. [493] K. Schwarz, D.B. Milne, Science 174 (1971) 426. [494] J. Costa Pessoa, S. Etcheverry, D. Gambino, Coord. Chem. Rev. (2015), http:// dx.doi.org/10.1016/j.ccr.2014.1012.1002 [495] K.H. Thompson, C. Orvig, Dalton Trans. (2006) 761–764. [496] E. Kioseoglou, S. Petanidis, C. Gabriel, A. Salifoglou, (2015) http://dx.doi.org/ 10.1016/j.ccr.2015.03.010.