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REDOX REGULATION OF PROTEIN SERINE/THREONINE PHOSPHATASE CALCINEURIN

DISSERTATION

Zur Erlangung des Grades eines Doktors der Naturwissenschaften des Fachbereichs für Biologie der Universität Konstanz

Dmitry Namgaladze

Konstanz, Februar 2002

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Dedicated to Lena for her love,

support and belief

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Volker Ullrich.

Parts of this work are published:

Bogumil, R., Namgaladze, D., Schaarschmidt, D., Schmachtel, T., Hellstern, S., Mutzel, R., and Ullrich, V. (2000) Inactivation of calcineurin by hydrogen peroxide and phenylarsine oxide. Evidence for a dithiol-disulfide equilibrium and implications for redox regulation. Eur.

J. Biochem. 267, 1407-1415.

Namgaladze, D., Hofer, H.W., and Ullrich, V. (2002) Redox control of calcineurin by

targeting the binuclear Fe2+-Zn2+ center at the enzyme active site. J. Biol. Chem. 277, 5962- 5969.

Further contributions:

Daiber, A., Herold, S., Schoneich, C., Namgaladze, D., Peterson, J.A., and Ullrich, V. (2000) Nitration and inactivation of cytochrome P450BM-3 by peroxynitrite. Stopped-flow

measurements prove ferryl intermediates. Eur. J. Biochem. 267, 6729-6739.

Daiber, A., Frein, D., Namgaladze, D., and Ullrich, V. (2002) Oxidation and nitrosation in the nitrogen monoxide/superoxide system. J. Biol. Chem. Papers In Press, published online ahead of print January 22, 2002.

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In the first place, I would like to thank my supervisor Prof. Volker Ullrich who let me join his group as a PhD student and provided outstanding working conditions. His constant encouragement and enthusiasm, excellent expertise and stimulating discussions proved invaluable in accomplishing this work.

I am grateful to Prof. Werner Hofer for providing facilities in his laboratory, sharing his experience in phosphatase field and being a second referent of this thesis.

Particularly I would like to thank Dr. Galina Sud’ina, without whose help I would not be able to start my work here.

My special thanks to Dr. Ralf Bogumil who introduced me to calcineurin and supported me throughout my study. His thoughtful and thoroughful approach to biochemical research was a very good school for me.

I would also like to thank...

...Andrea Höngen and Ivanna Shcherbyna for their contribution and support in calcineurin studies, creating a good working atmosphere and helping me whenever it was necessary.

...my colleagues of past and present Markus Bachschmid, Dr. Andreas Daiber, Dr. Martin Mehl, Dr. Ming-Hui Zou, Patrick Schmidt, Kai Schuler, Elena Dormeneva, Masha Kozlova, Svenia Thurau for fruitful cooperation and helpful discussions and together with Tanja Stengele, Regina Baudler, Gudrun von Scheeven, Elsiabeth Müssig, Barbara Herte, Vera Lorenz, and Marie Luise Endele for the good working atmosphere and help.

...Gisela Naschwitz for her help with official things and encouragement.

...Prof. Dieter Malchow for providing facilites in his laboratory for molecular biology work and Dr. Rupert Mutzel, Dr. Ursula Kessen, Dr. Anette Aichem and other members of AG Malchow for teaching and helping me.

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experiments with the reporter assay. Timo Buetler, Alexandra Krauskopf, Philip Lhose and other members of AG Ruegg for their help.

... Isolde Kuhn, Michael Safinowski and Magnus Bach for their engagement during labcourses and all other people who helped me during this study.

... my parents for their support and belief in me.

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AMPA α-amino-3-hydroxy-5-methyl-4- isoxazolepropionate

ATP adenosine triphosphate BCA bicinchoninic acid BSA bovine serum albumin

CAMK calmodulin-dependent protein kinase

CD circular dichroism

CHP calcineurin-homologous protein CREB cAMP-response element binding protein

CsA cyclosporin A

DARPP dopamine- and cyclic AMP- regulated phosphoprotein

DEAE diethylaminoethyl

DEA-NO 1-diethyl-2-hydroxy-2-nitroso- hydrazine

DMNQ 2,3-dimethoxy-1-naphthoquinone DMPS 2,3-dimercapto-1-propane sulfonic acid

DMSO dimethylsulfoxide DPI diphenyleniodonium DTP 2,2’-dithiodipyridine

ECL enhanced chemiluminescence EDTA ethylenediaminetetraacetic acid EGTA ethylene glycol-bis(β-aminoethyl ether)-N,N,N’,N’-tetraacetic acid

EPR electron paramagnetic resonance ER endoplasmatic reticulum

ERK extracellular response kinase FALS familial amyotrophic lateral sclerosis

FCS foetal calf serum

FKBP FK506-binding protein

FPLC fast protein liquid chromatography GABA γ-aminobutyric acid

GPx glutathione peroxidase GR glutathione reductase Grx glutaredoxin

GSH glutathione

HETE hydroxyeicosatetraenoic acid ICP-MS inductively coupled plasma mass spectrometry

IGF insulin-like growth facor IL interleukine

IP3R inostol-1,4,5-triphosphate receptor JNK c-jun N-terminal kinase

LTD long-term depression LTP long-term potentiation

MAPK mitogen-activated protein kinase MCIP myocyte-enriched calcineurin interacting protein

MEF myocyte enhancing factor MEL melarsen oxide

MUG 4-methylumbelliferin-β-D- galactoside

NADPH reduced nicotinamide adenine dinucleotide phosphate

NEM N-ethyl maleimide NES nuclear export signal NFκB nuclear factor kappa-B

NFAT nuclear factor of activated T-cells NHE Na+/H+ exchanger

NLS nuclear localization signal NMDA N-methyl-D-aspartate NOS nitric oxide synthase PAO phenylarsine oxide

PBS phosphate-buffered solution PCR polymerase chain reaction PCS prostacyclin synthase PDI protein disulfide isomerase PHA phytohemagglutinin PKA protein kinase A PKC protein kinase C

PMA phorbol 12-myristate 13-acetate PMSF phenylmethylsulfonylfluoride PN peroxynitrite

pNPP p-nitrophenylphosphate PP protein phosphatase

PTP protein tyrosine phosphatase ROS reactive oxygen species SDS sodium dodecyl sulfate

SDS-PAGE SDS polyacrylamide gel electrophoresis

SELDI-MS surface-enhanced laser desorption/ionization mass spectrometry SOD superoxide dismutase

SR sarcoplasmatic reticulum TCEP tris-carboxyethylphosphine TCR T-cell receptor

TR thioredoxin reductase Trx thioredoxin

XO xanthine oxidase

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1. INTRODUCTION...1

1.1. REDOX REGULATION OF CELLULAR SIGNALING...1

1.1.1. SOURCES OF REACTIVE OXYGEN SPECIES (ROS)...1

1.1.2. CHEMISTRY OF ROS...3

1.1.3. PHYSIOLOGICAL TARGETS OF ROS; IMPLICATION FOR ROS INVOLVEMENT IN INTRACELLULAR SIGNALING...5

1.1.4. CELLULAR ANTIOXIDANT DEFENSES...8

1.2. CALCINEURIN, BIOCHEMISTRY AND CELL BIOLOGY...10

1.2.1. HISTORY OF CALCINEURIN...10

1.2.2. SUBUNIT STRUCTURE AND ISOFORMS...11

1.2.3. DOMAIN STRUCTURE...11

1.2.4. CALCIUM DEPENDENCE...12

1.2.5. MODULATION OF CALCINEURIN ACTIVITY IN VITRO...13

1.2.6. CALCINEURIN X-RAY STRUCTURE...15

1.2.7. CALCINEURIN CATALYTIC MECHANISM...18

1.2.8. PHYSIOLOGICAL FUNCTIONS. ROLE OF CALCINEURIN IN YEAST...20

1.2.9. CALCINEURIN IN T-CELL ACTIVATION...22

1.2.10. CALCINEURIN IN CARDIAC AND SKELETAL MUSCLE...23

1.2.11. CALCINEURIN IN NERVOUS SYSTEM...24

1.2.12. CALCINEURIN AND APOPTOSIS...27

1.2.13. OTHER CELLULAR FUNCTIONS OF CALCINEURIN...28

1.2.14. CALCINEURIN-INTERACTING PROTEINS...29

1.2.15. CALCINEURIN REGULATION BY REDOX PROCESSES...30

AIMS OF THIS STUDY...33

2. MATERIALS AND METHODS...34

2.1. REAGENTS...…...34

2.2. PLASMIDS...34

2.3. PRIMERS...35

2.4. ISOLATION OF BOVINE BRAIN CALCINEURIN...35

2.5. DOT-BLOT ANALYSIS OF CALCINEURIN...36

2.6. MEASUREMENTS OF CALCINEURIN PHOSPHATASE ACTIVITY TOWARDS PNPP...37

2.7. CALCINEURIN TREATMENT WITH OXIDANTS...37

2.8. REACTIVATION OF INHIBITED CALCINEURIN...38

2.9. DETERMINATION OF FREE THIOL GROUPS...38

2.10.CD SPECTROSCOPY OF CALCINEURIN...38

2.11.IDENTIFICATION OF CALCINEURIN OXIDATIVE MODIFICATIONS……...39

2.12.SITE-DIRECTED MUTAGENESIS OF DICTYOSTELIUM DISCOIDEUM CALCINEURIN...39

2.13.STANDARD MOLECULAR BIOLOGY METHODS...40

2.14.PURIFICATION OF RECOMBINANT DICTYOSTELIUM DISCOIDEUM CALCINEURIN A...40

2.15.MICROPLATE PHOSPHATASE ACTIVITY ASSAY USING PNPP...41

2.16.TREATMENT OF RECOMBINANT CALCINEURIN A WITH OXIDANTS...41

2.17.EXPRESSION AND PURIFICATION OF RECOMBINANT RAT CALCINEURIN...41

2.18.SENSITIVITY OF RECOMBINANT RAT CALCINEURIN TO PAO...42

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2.20.PREPARATION OF CELL AND TISSUE EXTRACTS FOR PHOSPHATASE ACTIVITY ASSAYS...43

2.21.MEASUREMENTS OF CALCINEURIN PHOSPHATASE ACTIVITY TOWARDS 32P-LABELED RII PHOSPHOPEPTIDE...43

2.22.TREATMENT OF CELL AND TISSUE EXTRACTS WITH OXIDANTS...44

2.23.ISOLATION OF PORCINE BRAIN CALCINEURIN...44

2.24.LIMITED PROTEOLYSIS OF CALCINEURIN...45

2.25.QUANTITATION OF SUPEROXIDE PRODUCTION...45

2.26.METAL ANALYSIS...46

2.27.SELDI-MS ANALYSIS...46

2.28.EPR SPECTROSCOPY...46

2.29.NFAT PHOSPHORYLATION ANALYSIS...46

2.30.β-GALACTOSIDASE REPORTER GENE ASSAY...47

3. RESULTS...48

3.1. INHIBITION OF ISOLATED BOVINE CALCINEURIN BY OXIDANTS VIA DITHIOL-DISULFIDE TRANSITION...48

3.1.1. INHIBITION OF CALCINEURIN BY PAO. REVERSIBILITY BY DISULFIDE REDUCING AGENTS...48

3.1.2. INACTIVATION OF CALCINEURIN BY H2O2...50

3.1.3. ROLE OF THIOL OXIDATION IN H2O2-MEDIATED CALCINEURIN INACTIVATION...51

3.1.4. CALCINEURIN OXIDATION DOES NOT AFFECT THE PROTEIN SECONDARY STRUCTURE, BUT CAUSES PARTIAL DIMERIZATION...52

3.2. SITE-DIRECTED MUTAGENESIS OF CYSTEINES IN DICTYOSTELIUM DISCOIDEUM CALCINEURIN...54

3.2.1. EXPRESSION AND PURIFICATION OF WILD-TYPE AND MUTATED RECOMBINANT PROTEINS...54

3.2.2. REDOX-SENSITIVITY OF WILD-TYPE AND MUTATED PROTEINS...56

3.2.3. REDOX SENSITITIVITY OF CYSTEINE-TO-ALANINE MUTANTS OF RAT CALCINEURIN A....57

3.3. INHIBITION OF CALCINEURIN BY SUPEROXIDE IN TISSUE AND CELL HOMOGENATES...59

3.3.1. CHARACTERIZATION OF CALCINEURIN ACTIVITY IN BOVINE BRAIN HOMOGENATE...59

3.3.2. SUPEROXIDE-MEDIATED INHIBITION OF CALCINEURIN ACTIVITY IN TISSUE AND CELL HOMOGENATES BY XANTHINE OXIDASE/HYPOXANTHINE SYSTEM...60

3.3.3. CALCIUM DEPENDENCE OF CALCINEURIN ACTIVITY INHIBITION BY SUPEROXIDE...62

3.3.4. Modulation of superoxide inhibitory effect by NO and reducing agents...63

3.4. REDOX SENSITIVITY OF PORCINE CALCINEURIN ISOLATED UNDER REDUCING CONDITIONS. INVOLVEMENT OF FE2+ IN THE BINUCLEAR METAL CENTER...66

3.4.1. ISOLATION AND CHARACTERIZATION OF PORCINE BRAIN CALCINEURIN...66

3.4.2. PHOSPHATASE ACTIVITY OF PORCINE CALCINEURIN. INHIBITION BY OXIDANTS...68

3.4.3. CALCIUM/CALMODULIN DEPENDENCE OF CALCINEURIN REDOX SENSITIVITY...71

3.4.4. NO ANTAGONISM OF CALCINEURIN INHIBITION BY XO/HYPOXANTHINE...72

3.4.5. EVIDENCE FOR CALCINEURIN BINUCLEAR FE2+-ZN2+ CENTER AS REDOX TARGET...73

3.4.6. EPR-SPECTROSCOPY AND METAL ANALYSIS OF CALCINEURIN...76

3.5. EFFECTS OF OXIDANTS ON CALCINEURIN-DEPENDENT SIGNALING IN INTACT CELLS....79

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3.5.2. CALCINEURIN-DEPENDENT NFAT TRANSCRIPTIONAL ACTIVITY IN JURKAT CELLS IS

INHIBITED BY OXIDANTS...80

3.5.3. STIMULATION OF RESPIRATORY BURST IN MACROPHAGES CAUSES INHIBITION OF ENDOGENOUS CALCINEURIN ACTIVITY...82

4. DISCUSSION...84

4.1. ROLE OF CALCINEURIN THIOLS IN REDOX REGULATION OF THE PHOSPHATASE ACTIVITY..84

4.2. SUPEROXIDE INHIBITION OF CALCINEURIN BY TARGETING ITS FE2+-ZN2+ BINUCLEAR CENTER...88

4.3. REDOX REGULATION OF CALCINEURIN IN VIVO...95

5. SUMMARY...97

6. ZUSAMMENFASUNG………..…....……99

7. REFERENCES………....……..101

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1. INTRODUCTION:

1.1. REDOX REGULATION OF CELLULAR SIGNALING.

Molecular oxygen (O2) is an essential element of life, and all aerobic organisms do not survive without it. Oxygen is consumed primarily via oxidative phosphorylation, a process whereby the energy of four-electron reduction of O2 to H2O within the mitochondrial electron transport chain is converted to the high-energy phosphate bond of ATP. Additionally, numerous oxidations involving O2 provide a basis of cellular metabolism. These reactions produce also partially reduced O2 metabolites as side products. These include superoxide anion-radical (⋅O2-), hydrogen peroxide (H2O2) and hydroxyl radical (⋅OH), collectively called reactive oxygen species (ROS). In addition, another free radical molecule of biological origin, nitric oxide (⋅NO) reacts with superoxide to form a potent oxidant, peroxynitrite (ONOO-).

The reactivity of ROS results in their general toxicity for the cell, and excessive ROS formation causes oxidation of proteins, lipids and DNA. Therefore, cells developed several levels of defense against toxic effects of ROS, from low-molecular weight reducing compounds (glutathione, ascorbate) to specific ROS-consuming enzymes (SOD, catalase, glutathione peroxidase). Recently, it has been recognized that ROS are not only toxic by- products of oxidative metabolism, but also important regulators of intracellular signaling.

Thus, intracellular ROS formation in response to changes of cellular environment is able to modulate various signaling mechanisms within the cell, particularly those governed by protein phosphorylation/dephosphorylation. The level of intracellular ROS participating in cell signaling is finely balanced by varying activities of ROS generating and antioxidant systems.

The elucidation of the mechanisms by which redox processes control cellular signaling thus presents a major challenge in unraveling the complex picture of cellular response to environmental factors. On the other hand, uncontrolled ROS production accompanying many pathological conditions overwhelms cellular antioxidant defenses and ultimately leads to cell and tissue degeneration or death. In this case, finding the critical targets of ROS and clarifying mechanisms of cellular oxidative damage would accelerate the design of drugs and therapies able to cope with oxidative stress-accompanied pathologies.

1.1.1. SOURCES OF REACTIVE OXYGEN SPECIES (ROS).

Several major enzymatic and non-enzymatic sources of ROS are present in the cell [Thannical and Fanburg, 2000]. Steady-state ROS production is mostly attributed to the

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mitochondrial electron transport chain, where approximately 1-2 % of O2 consumption results in ⋅O2- generation, mostly through side reaction of complexes I, II and III with molecular oxygen [Cadenas and Davies, 2000]. In addition, monoamine oxidase of the outer mitochondrial membrane could contribute to mitochondrial H2O2 generation [Cadenas and Davies, 2000]. In the endoplasmic reticulum (ER) the enzymes of the cytochrome P450 family produce ⋅O2- and H2O2 as by-products in the processes of fatty acid oxidation and xenobiotic metabolism [Goeptar et al., 1995]. The enzymes of the lipoxygenase family convert polyunsaturated fatty acids to lipid signaling molecules, leukotrienes and HETEs, and they also generate ROS.

Plasma membrane-associated oxidases appear to be the key enzymes involved in superoxide generation in response to extracellular signals. The best-characterized oxidase is the phagocytic NADPH oxidase, the enzyme critical for the defense against microorganism invasions [Wientjes and Sigal, 1995]. NADPH oxidase is a multiprotein enzyme complex, which is assembled on the plasma membrane of phagocytes upon cell stimulation. NADPH oxidation at the cytochrome b558 part (composed of gp91phox and p22phox proteins) of the complex generates ⋅O2-, which is subsequently released into phagocytic vacuoles or the extracellular space. Several gp91phox homologues in non-phagocytic cells have been cloned recently [Lambeth et al., 2000]. It becomes clear that these oxidases play an important role in generation of ROS upon activation of various cell surface receptors, and help shaping a cell mitogenic response through activation of MAPK kinase cascades [Suh et al., 1999].

Xanthine oxidase (XO) is a membrane-associated enzyme formed from xanthine dehydrogenase by thiol oxidation or proteolysis, and it generates ⋅O2- and H2O2 during enzymatic turnover [Fridovich, 1970]. It is widely used as a superoxide source in vitro and is involved in endothelial dysfunction in vivo.

Another recently acknowledged source of ⋅O2- could be the enzymes of the NO synthase (NOS) family. Several conditions including arginine and tetrahydrobiopterin (BH4) deficiency promote superoxide generation by NOS isozymes [Groves and Wang, 2000].

When speaking about ROS generation one should also mention the generation of NO, since NO functions both as a scavenger of superoxide and as a precursor of peroxynitrite (PN), which is together with ROS considered a major oxidant formed in the cell. NO formation occurs mainly via arginine oxidation catalyzed by NOS. Three NOS isoforms (neuronal nNOS, endothelial eNOS and inducible iNOS) have been cloned, eNOS and nNOS are constutively present in several cell types, and iNOS is induced under inflammatory conditions. eNOS and nNOS are activated by calcium/calmodulin, and iNOS is constitutively

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active. NO generation by an eNOS and nNOS plays important role in maintenance of vascular tone and neurotransmission, respectively, while NO production by iNOS contributes to bacterial killing by phagocytes.

Phagocytes produce also a specific oxidant, hypochlorous acid (HOCl) by the myeloperoxidase-catalyzed oxidation of chloride anion [Hampton et al., 1998]. HOCl is involved in bacterial killing by phagocytes, and is unlikely to have a general role in intracellular signaling except for extreme inflammatory conditions.

1.1.2. CHEMISTRY OF ROS.

The enzymatic systems described above produce O2- as a primary ROS. This species is not very stable in solution at physiological pH and undergoes a dismutation reaction:

O2- + HO2 → O2 + HO2- (1) k=7.3×105 M-1s-1

In an intracellular milieu this reaction is catalyzed by SOD enzymes, with second order rate constants 2×109 M-1s-1 for Cu, Zn-SOD and 1×108 M-1s-1 for Mn-SOD [Lee et al., 1998].

Only one reaction competes with SOD-catalyzed superoxide dismutation under physiological conditions, namely the reaction with NO:

O2- + NO → ONOO- (2) k=6.9×109 M-1s-1

Superoxide can participate in redox reactions acting either as an oxidant or as a reductant.

Thus, the reduction of Fe3+ in ferricytochrome c by superoxide or reduction of nitro tetrazolium blue provides a basis for reliable tests of superoxide production [Fridovich, 1995].

Another reaction with iron and hydrogen peroxide generates a very reactive hydroxyl radical by the Haber-Weiss cycle:

Fe3+ + O2- → Fe2+ + O2 (3)

Fe2+ + H2O2 → Fe3+ + ⋅OH + OH- (4) Reaction (4) is also called Fenton reaction.

Superoxide in its protonated form also reacts with thiols:

HO2 + RSH → H2O2 + RS⋅ (5) 2RS⋅ → RSSR (6)

Thiol oxidation by superoxide is, however, limited at physiological pH, since the pKa of the superoxide conjugated acid, hydroperoxyl radical, is 4.8. Superoxide reacts also with several low molecular weight compounds of biological significance, including ascorbate, catecholamines, polyphenols and tetrahydrobiopterin, but does not readily react with polyunsaturated fatty acids or DNA [Fridovich, 1997].

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H2O2 formed upon superoxide dismutation or univalent reduction is relatively stable.

Its dismutation occurs at appreciable rates only in the presence of metal ions or when catalyzed by catalase:

H2O2 + H2O2 → 2H2O + O2 (7) k=0.8-2×109 M-1s-1

H2O2 reacts with thiols, but otherwise it is considered not very reactive. Much of the damage inflicted by H2O2 is attributed to a metal-catalyzed ⋅OH formation (reaction (4)). However, in contrast to superoxide H2O2 penetrates cell membranes and is widely used as an extracellular inducer of oxidative stress.

Reaction (4) produces the very reactive ⋅OH radical. Apart from iron ions, copper is also an efficient catalyst in the Fenton reaction. ⋅OH radical reacts with a broad variety of functional groups on proteins, lipids and DNA. ⋅OH reactivity also limits the spatial range of its action to the sites of its formation, e.g. transition metals bound to DNA or lipids.

NO, in addition to the already mentioned reaction with superoxide, participates in other chemical reactions with biological substances. Its reaction with molecular oxygen:

2NO + O2 → 2NO2 (8) k=2-6×106 M-2s-1

is bimolecular with respect to NO. Therefore, it contributes insignificantly to the fate of NO under physiological conditions where NO concentration reaches at maximum several µM [Beckman and Koppenol, 1996]. The reaction of NO and NO2 produces N2O3, a nitrosating species:

NO + NO2 ' N2O3 (9)

Transitional metals could oxidize NO:

NO + Fe3+ → Fe2+

⋅⋅⋅

NO+ (10)

The resulting nitrosonium cation could nitrosate different compounds, including thiols NO+ + RSH → RSNO + H+ (11)

The other possibility to form nitrosothiols is through a radical mechanism:

⋅NO + RS⋅ → RSNO (12)

The product of reaction of NO with O2-, PN, is a reactive molecule with a broad range of chemical targets. PN is rather stable at alkaline pH, but its conjugated peroxonitrous acid rapidly isomerizes to nitrate:

ONOO- + H+ ' ONOOH → NO3- + H+ (13)

PN nitrates phenols and phenol moeties of different biomolecules by a radical mechanism, which could be efficiently catalyzed by iron in heme-thiolate proteins [Mehl et al., 1999]. In addition, the reaction of PN with phenols results in formation of hydroxylation, nitrosation

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and dimerization products. Reaction of PN with thiols yields disulfides, sulfenic, sulfinic, and sulfonic acids as well as nitroso- or nitro-thiols [Pryor and Squadrito, 1995]. PN also targets DNA, polyunsaturated fatty acids and such low molecular weight compounds as ascorbate or uric acid. In addition, PN forms an adduct with CO2:

ONOO- + CO2 → ONOOCO2- → [NO2+ CO32-] ↔ [⋅NO2 ⋅CO3-]

This adduct is likely to form under physiological conditions and is in several cases more reactive than PN itself [Denicola et al., 1996].

1.1.3. PHYSIOLOGICAL TARGETS OF ROS; IMPLICATION FOR ROS INVOLVEMENT IN INTRACELLULAR SIGNALING.

The multitude of chemical reactions involving ROS results in their targeting various functional groups on proteins, lipids or DNA. Under pathological conditions the oxidative damage to the cell could be non-specific and lead to necrotic cell death. If redox reactions are to play a role in cell signaling mechanisms, they should target a restricted set of functional groups on specific protein targets, and these modifications should be generally reversible by intracellular antioxidative mechanisms.

Protein cysteine residues are common targets of oxidants. Thiol groups (-SH) can be oxidized to sulfenic (-SOH) or higher valence sulfinic (-SO2H) or sulfonic (-SO3H) derivatives. Two thiol groups can also build a disulfide bridge (-S-S-) upon oxidation or form a mixed disulfide with glutathione (GSH). Formation of the sulfenic acid derivative of active site cysteine was found to be a mechanism whereby ROS inactivate protein tyrosine phosphatases, e.g. PTP1B [Denu and Tanner, 1998]. This derivative can react with reduced glutathione forming mixed disulfides (a process called S- glutathionylation), which could be reduced to the active thiol state of the cysteine by glutaredoxins [Barrett et al., 1999a]. Kinetic studies have shown that superoxide could be a more potent and specific oxidant for PTP1B, providing one of the first evidence for a direct role of O2- in oxidative modification of the signaling proteins [Barrett et al., 1999b]. Reversible S-glutathionylation was also shown to be involved in redox regulation of c-jun DNA binding [Klatt et al., 1999].

Dithiol-disulfide transition appears to be a common modification of proteins, and in several cases it was shown to play a key regulatory role for intracellular signaling. Thus, in E.coli OxyR transcription factor is activated via formation of a disulfide bond [Zheng et al., 1998] resulting in the induction of a set of antioxidant genes. Similarly, dithiol-disulfide transition activates bacterial chaperone Hsp33 as a part of a cell-protective mechanism [Jacob

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et al., 1999]. Dithiol-disulfide equilibrium provides the functional basis for thiol reductase activities of thioredoxin or glutaredoxin [Holmgren, 1989] or in catalysis of disulfide bond formation like for protein disulfide isomerase (PDI) [Ferrari and Soling, 1999].

Another sulfur-containing target of ROS represent iron-sulfur clusters, which can be found in many enzymes. The [4Fe-4S] cluster is particularly sensitive to O2-, although the other oxidants and NO can also react with it. [4Fe-4S] clusters of aconitase and related enzymes are major superoxide targets in bacteria [Fridovich, 1997]. Redox reactions of [4Fe- 4S] cluster in iron regulatory protein-1 (IRP-1) control intracellular iron metabolism [Hentze and Kuhn, 1996].

NO-linked redox modifications of proteins include protein nitrosation and tyrosine nitration. Nitrosation of cysteine residues was proposed as a mechanism regulating the activity of several important proteins, including NMDA receptor or ryanodyne receptor [Stamler et al., 1997]. Recently, a proteomic approach revealed a set of endogenously nitrosated proteins, suggesting a critical role of nitrosation in regulation of cellular functions [Jaffrey et al., 2001].

Nitration of protein tyrosine residues, if catalyzed by metals, can occur at low PN concentrations expected to form in vivo. One particular example is prostacyclin synthase (PCS), which could be nitrated by submicromolar PN concentrations [Zou et al., 1997].

Nitration of PCS was implicated in a variety of pathophysiological conditions, including atherosclerosis or ischemia [Zou et al., 1999; Zou and Bachschmid, 1999]. Other proteins, including Mn-SOD [MacMillan-Crow et al., 1996] or tyrosine hydroxylase [Blanchard-Fillion et al., 2001] were also shown to be nitrated in vivo, and proteomic analysis revealed that many proteins, especially in mitochondria, undergo nitration under inflammatory conditions [Aulak et al., 2001]. In addition to nitration, tyrosine dimerization occurs under oxidative conditions, and this modification could also play a role in regulation of enzyme function [MacMillan- Crow and Thompson, 1999].

Almost every protein is able to undergo oxidative modifications under specific conditions. Redox processes modulating intracellular signaling involve, however, a limited number of targets, which determine the cellular response to environmental changes. Signaling pathways from the plasma membrane to the nucleus involve redox-sensitive components at many stages. Protein tyrosine phosphatases (PTPs) belong to a recognized class of ROS targets, since all of them contain an active site cysteine sensitive to oxidation as described above. Inactivation of PTPs appears to be primarily responsible for the increase of growth factor receptor tyrosine phosphorylation and activation after oxidative stress [Knebel et al.,

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1996]. This increase of tyrosine phosphorylation can in turn activate downstream signaling cascades leading to activation of the Ras-ERK pathway. JNK and p38 MAPK kinase cascades also contribute to the signaling downstream of ROS. Another signaling system regulated by ROS could involve the increase of intracellular calcium after oxidative stress. It seems that Ca2+-ATPase of sarcoplasmic reticulum (SR) is the redox-sensitive target in this case, since it could be inhibited by O2- and other ROS [Suzuki and Ford, 1991]. Much attention was paid also to the role of ROS in the regulation of transcription factors, particularly NFκB and AP-1.

NFκB regulates the expression of genes involved in the inflammatory response and cell survival, and ROS were shown to be involved in NFκB activation, although the exact target was not identified [Li and Karin, 1999]. The picture of redox regulation of NFκB is, however, contradictory, since NFκB itself contains an oxidation-prone cysteine critical for DNA- binding [Kumar et al., 1992]. In addition, IκB kinase-β (IKK-β), a kinase necessary for NFκB activation, contains a redox-sensitive cysteine targeted by arsenite [Kapahi et al., 2000]. AP-1 transcription factor family consists of homo-and heterodimers of Fos and Jun proteins. These proteins contain redox-sensitive cysteines in their DNA-binding domains, and Ref-1 antioxidant protein is necessary for maintenance of the reduced state of this cysteine in the nucleus [Xanthoudakis and Curran, 1992]. On the other hand, oxidative stress seems to be also involved in the activation of the AP-1-dependent transcription through activation of the upstream signaling components. In yeast, an AP-1 homologue Yap1 is activated by oxidative stress through the formation of the disulfide bridge causing Yap1 nuclear accumulation [Toone et al., 2001]. Several other transcription factors (p53, c-myb) contain cysteines in their DNA-binding domains, mainly as components of zinc finger structural motifs, and are presumed to be redox-regulated.

There is no consensus on the question, which species is mostly responsible for redox regulation of intracellular signaling. Recent evidence, however, points to involvement of O2-

in oxidative modification of key signaling proteins [Ullrich and Bachschmid, 2000]. Although O2- is considered relatively unreactive, in several situations it could selectively target different functional groups on proteins. A first example of such a regulation came from bacterial aconitase, which has a [4Fe-4S] cluster sensitive to O2- inhibition [Gardner and Fridovich, 1991]. This kind of modification can have a regulatory function as shown for a bacterial SoxR transcription factor [Fridovich, 1997] or for IRP-1 protein (above). Latest studies indicated that isolated cysteine residues could be also selectively modified by O2- like in PTP1B [Barret et al., 1999b], and this case has profound implications for intracellular signaling. Another key signaling molecule regulated by superoxide is PKC, which is activated following oxidation of

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cysteines and zinc release from its zinc finger motif [Knapp and Klann, 2000]. Zinc release was also reported to accompany superoxide-mediated inhibition of endothelin-converting enzyme [Lopez-Ongil et al., 2000], although in this case no cysteines seemed to be involved.

Superoxide was also shown to inhibit soluble guanylyl cyclase by an as yet unidentified mechanism [Brune et al., 1990]. In summary, these findings suggest that superoxide is suitable for a role as a direct modulator of intracellular signaling processes.

1.1.4. CELLULAR ANTIOXIDANT DEFENSES.

Small amounts of ROS generated upon activation of cellular signaling cascades mediate cell responses to environmental changes, but in many pathophysiological situations excessive ROS generation is damaging to the cell. To protect itself from oxidative damage the cell developed several levels of defense mechanisms. They include the enzymes involved in ROS degradation, the enzymes taking part in repair of the damaged biomolecules and small substances maintaining general reductive environment of the cell.

The enzymes of the superoxide dismutase (SOD) family constitute the first level of antioxidant defenses [Fridovich, 1995; Culotta, 2000] catalyzing superoxide dismutation to H2O2 and O2. In mammalian cells there are two SODs, Cu, Zn-SOD and Mn-SOD, and an additional Cu, Zn-SOD is associated with the cell surface. Mn-SOD is localized in mitochondria, and Cu, Zn-SOD is predominantly cytosolic. The critical role of SOD enzymes was revealed by genetic studies showing that mice deficient in Mn-SOD die within the first 10 days of life with a dilated cardiomyopathy and different metabolic abnormalities [Li et al., 1995]. Mice lacking Cu, Zn-SOD are viable, but are increasingly sensitive to oxidative stress, especially in neurons [Reaume et al., 1996]. Cu, Zn-SOD is also implicated in the pathogenesis of familial amyotrophic lateral sclerosis (FALS), a fatal neurodegenerative disease. Several Cu, Zn-SOD mutations are associated with this disease, but the exact mechanism whereby these mutations cause neurodegeneration has not been clarified.

H2O2 generated after O2- dismutation is removed mainly by two enzymes, catalase and glutathione peroxidase (GPx). Catalase is a cytosolic and mostly peroxisomal enzyme [Michiels et al., 1994]. Its localization in mitochondria is tissue-specific. Catalase catalyzes H2O2 conversion into H2O and O2, and it is effective at high concentrations of H2O2. GPx enzyme family consists of selenoproteins with both cytosolic and mitochondrial localization catalyzing reduction of H2O2 and phospholipid/fatty acid hydroperoxides by GSH [Michiels

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et al., 1994]. Mitochondrial GPx are thought to play a major role in detoxification of H2O2

produced upon superoxide dismutation by mitochondrial SODs. In addition, enzymes of the peroxiredoxin family reduce H2O2 and organic peroxides with their active site thiols [Butterfield et al., 1999].

The enzymes that restore function of oxidized proteins constitute another level of cellular antioxidative defenses. Protein thiols are most sensitive to oxidation, and the cell developed two major disulfide-reducing systems, thioredoxin (Trx) and glutaredoxin (Grx, also called thioltransferase) systems. Trx is a protein of ~12 kDa containing a CGPC active site motif, and it catalyzes the reduction of disulfide bonds in a broad variety of proteins [Holmgren, 1985]. It forms a coupled system with its reducing enzyme, thioredoxin reductase (TR), a selenocysteine-containing protein catalyzing reduction of Trx by NADPH [Mustacich and Powis, 2000]. In addition, TR is able to reduce non-disulfide substrates like alloxan or S- nitrosoglutathione. In the nucleus the thioredoxin system is coupled to a specific disulfide reductase for transcription factors, Ref-1. Ref-1 has two cysteines critical for its reductase activity, and after oxidation it can be reduced by Trx [Powis et al., 1997]. The glutaredoxin system consists of Grx, GSH, NADPH and glutathione reductase [Holmgren, 1989]. Like Trx, Grx are small proteins containing a CPYC active site motif, and they structurally belong to the thioredoxin fold superfamily. Grx preferably catalyze the reduction of mixed disulfides with GSH, although they can also act as disulfide reductases.

Several low molecular weight compounds maintain the general reductive state of the intracellular compartments and provide reducing equivalents for various redox reactions in the cell. Particularly important are ascorbate (vitamin C) and GSH. Ascorbate (anion at physiological pH), chemically 2,3-endiol-L-gulonacid-γ-lactone, can be easily oxidized to semidehydroascorbate and dehydroascorbate, which in turn can be reduced to ascorbate by GSH, Grx or thioredoxin reductase. Antioxidative actions of ascorbate include radical scavenging, scavenging of non-radical oxidants, inhibition of lipid peroxidation and maintenance of the reduced state of iron and copper [Halliwell, 1999]. Ascorbate serves also as a co-factor for several enzymes, e.g. prolyl/lysyl hydroxylases involved in collagen synthesis. The human organism does not synthesize ascorbate, and it must be taken up from diet. Ascorbate is accumulated in areas of enhanced energy metabolism, particularly in neurons, where its intracellular concentration can reach 10 mM [Rice and Russo-Menna, 1998].

GSH is a thiol-containing tripeptide (γ-L-glutamyl-L-cysteinylglycine) [Dringen, 2000]. It is synthesized in the cell from aminoacid constituents by consecutive action of

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γGluCys synthetase and glutathione synthetase. Its cellular concentration is in the millimolar range, and it is present in most intracellular compartments. GSH plays a major role in antioxidant mechanisms as an electron donor in GPx-catalyzed peroxide reduction. It also forms mixed disulfides with oxidized proteins, which could be further reduced by the Grx system. GSH can also directly scavenge free radicals. Oxidized GSH can be reduced by enzymes of the glutathione reductase (GR) family, selenoproteins similar to thioredoxin reductase. In addition, GSH is important for detoxification of xenobiotica (reactions catalyzed by glutathione transferases) and as a cofactor in isomerization reactions.

All these enzymatic and non-enzymatic defenses help to maintain the overall reduced state of cellular biomolecules and provide protection against the deleterious side effects of aerobic metabolism. Changing the balance of pro-oxidant and anti-oxidant factors can lead to severe impairment of cellular function as happens in many pathological situations.

1.2. CALCINEURIN, BIOCHEMISTRY AND CELL BIOLOGY. 1.2.1. HISTORY OF CALCINEURIN.

Calcineurin was first detected in 1976 as a brain fraction, which inhibited calmodulin- dependent cyclic nucleotide phosphodiesterase [Wang and Desai, 1976]. It was later purified to homogeneity by Klee and Krinks [Klee and Krinks, 1978]. Klee et al. called the newly discovered protein “calcineurin” on the basis of calcium-binding properties and localization in neural tissue [Klee et al., 1979], the name that is until now generally accepted in the literature.

Its function as a phosphatase was recognized only in 1982 by the group of Cohen [Stewart et al., 1982], which also termed it protein phosphatase 2B [Stewart et al., 1983]. Molecular cloning of calcineurin followed in 1989 [Guerini and Klee, 1989; Ito et al., 1989]. The discovery of calcineurin as target of the immunosuppressive drugs cyclosporin A (CsA) and FK506 [Liu et al., 1991a] helped to elucidate its role in T-lymphocyte activation and provided pharmacological tools for studying calcineurin involvement in various signaling pathways.

Recently, major achievements included resolving calcineurin three-dimensional structure [Griffith et al., 1995; Kissinger et al., 1995] and generation of transgenic animal models, including knock-outs [Zhang et al., 1996; Winder et al., 1998; Molkentin et al., 1998; Graef et al., 2001], which greatly advanced our understanding of calcineurin biochemistry and physiological significance.

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1.2.2. SUBUNIT STRUCTURE AND ISOFORMS.

Calcineurin has a unique two-subunit structure, which is conserved among all eukaryotes. Calcineurin is a heterodimer of a 58 to 69-kDa catalytic subunit (calcineurin A), and a regulatory 16 to 19-kDa subunit (calcineurin B). Three isoforms of mammalian calcineurin A termed α, β, and γ were discovered, with some additional variety through alternative splicing. α- and β-isoforms are ubiquitous, whereas the γ-isoform is testis-specific [Muramatsu and Kinkaid, 1992]. In human, corresponding genes PPP3CA, PPP3CB, and PPP3CC are located on chromosomes 4, 10 and 8 [Muramatsu and Kinkaid, 1992; Wang et al., 1996a]. Two calcineurin A isoforms are present in Saccharomyces cerevisiae [Cyert et al., 1991; Liu et al., 1991b], one in Schizosaccharomyces pombe [Yoshida et al., 1994]. Two genes correspond to calcineurin A subunit in Drosophila melanogaster [Guerini et al., 1992;

Brown et al., 1994], and one isoform was found in Caenorhabditis elegans [Wheelan et al., 1999], Neurospora crassa [Higuchi et al., 1991], Aspergillus nidulans [Rasmussen et al., 1994], and Dictyostelium discoideum [Dammann et al., 1996]. In some lower eukaryotes the size of the catalytic subunit can be up to 20% larger than in mammals (e.g. 71kDa in Dictyostelium discoideum) with N- and C-terminal extensions, but the catalytic core is conserved in all species. Recently, a novel mammalian splice variant of calcineurin Aα, which lacked catalytic and calcineurin B-binding domains was found [Reuter et al., 2001].

There are two mammalian isoforms of regulatory B subunit, one of which is testis- specific [Ueki et al., 1992]. A single gene was identified in Saccharomyces cerevisiae [Kuno et al., 1991] as well as in Neurospora crassa [Kothe and Free, 1998] and Dictyostelium discoideum [Aichem and Mutzel, 2001] (the later gives rise to two mRNA and protein species due to unconventional processing of the cnbA gene). The B subunit is highly conserved throughout evolution, allowing interspecies substitution in reconstitution of functionally active holoenzyme [Ueki and Kincaid, 1993].

1.2.3. DOMAIN STRUCTURE.

The functional domain structure of calcineurin A subunit is evolutionally conserved (Fig.

1.1). A core catalytic domain with high homology to other serine/threonine phosphatases is followed by three regulatory domains: calcineurin B-binding domain [Sikkink et al., 1995;

Watanabe et al., 1995], calmodulin-binding domain [Kincaid et al., 1988], and autoinhibitory

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100 200 300 400 500

Catalytic CnB

binding CaM AI

domain [Hashimoto et al., 1990]. These domains were identified by biochemical methods and later confirmed in the crystal structures of calcineurin. A C-terminal part of the enzyme Fig. 1.1. Domain structure of calcineurin A. Abbreviations: CaM – calmodulin binding, AI – autoinhibitory.

molecule containing calmodulin-binding and autoinhibitory domain is readily susceptible to proteolysis [Manalan and Klee, 1983; Hubbard and Klee, 1989]. The resulting truncated form of calcineurin is no longer dependent on calmodulin and is active in the presence of low calcium concentrations.

1.2.4. CALCIUM DEPENDENCE.

The phosphatase activity of calcineurin is dependent on both Ca2+ binding to calcineurin B and Ca2+-dependent binding of one molecule of calmodulin. Calcineurin B contains four characteristic EF-hand Ca2+-binding motifs [Aitken et al., 1984]. They have different affinities for calcium: a carboxy-terminal pair of sites with high (nanomolar) affinity, and an amino-terminal pair with (micromolar) affinity [Feng and Stemmer, 1999;

Gallagher et al., 2001]. The carboxy-terminal sites also exhibit very slow rates of calcium- exchange, retaining calcium constitutively and presumably playing a structural role [Feng and Stemmer, 1999]. In contrast, binding of calcium to low-affinity sites evokes conformational changes in both calcineurin subunits [Yang and Klee, 2000], modulates the enzyme affinity for calmodulin [Feng and Stemmer, 2001] and calcineurin activation by calmodulin [Stemmer and Klee, 1994; Feng and Stemmer, 2001]. Calmodulin binds calcineurin in the presence of calcium with very high affinity (Kd<10-10M) [Hubbard and Klee, 1987]. Calcium-bound calmodulin increases calcineurin phosphatase activity 10-100 fold [Stewart et al., 1982;

Tallant and Cheung, 1984] by increasing Vmax of the enzyme. Calcium-dependence of calcineurin activation by calmodulin is highly cooperative (Hill coefficient 2.8-3) [Stemmer and Klee, 1994], reflecting binding of calcium to more than two sites on calmodulin. This

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cooperativity allows a narrow threshold for calcium stimulation of calcineurin. Binding to calcineurin also greatly increases calmodulin affinity for calcium [Stemmer and Klee, 1994], and allows optimal stimulation by levels of calcium achieved in stimulated cells.

1.2.5. MODULATION OF CALCINEURIN ACTIVITY IN VITRO.

The phosphatase activity of calcineurin was initially assayed by measuring release of radioactive phosphate from 32P-labeled protein substrates (α-subunit of phosphorylase kinase, myosin light chain, histone H1) [Stewart et al., 1983; King and Huang, 1983]. Calcineurin was later found to hydrolyze a range of low-molecular range substrates, including p- nitrophenylphosphate (pNPP) [Pallen and Wang, 1983; Li, 1984; Pallen et al., 1985]. The latter became the main substrate in studies of calcineurin catalytic mechanism. Subsequently, the serine-phosphorylated RII regulatory subunit of protein kinase A was shown to be a good calcineurin substrate [Blumenthal et al., 1986]. A 19-aminoacid part of RII, which contained phosphorylated residue and retained substrate properties of the entire protein, provided conventional substrate for determination of calcineurin serine/threonine phosphatase activity.

Use of 32P-labeled RII peptide allowed for a reliable assay of calcineurin activity in cell and tissue extracts.

Apart from calcium, other divalent metal ions can modulate calcineurin phosphatase activity. The best activators at physiological pH values were shown to be Mn2+ and Ni2+

[King and Huang, 1983; Pallen and Wang, 1984], and Mg2+ at pH 8 [Li, 1984]. In initial work on calcineurin isolation Cohen and co-workers showed that Mn2+-dependence is acquired only after a calmodulin affinity chromatography step [Stewart et al., 1982]. Later, King and Huang [King and Huang, 1984] described rapid inactivation of calcineurin in the presence of calcium and calmodulin. Mn2+ and Ni2+ prevented and reversed this inactivation.

More detailed studies showed some difference in the activation mode between these two metals [Pallen and Wang 1984; Pallen and Wang 1986] and found up to two binding sites for each of the metal ions, one of which was competitive.

Phosphorylation affects the functional properties of many cellular proteins.

Calcineurin was shown to be phosphorylated in vitro by protein kinase C and calmodulin- dependent kinase II [Hashimoto and Soderling, 1989; Martensen et al., 1989] and casein kinase I [Singh and Wang, 1987]. The kinetic properties of phosphorylated and unphosphorylated forms are similar [Hashimoto and Soderling, 1989], and the relevance of calcineurin phosphorylation for its regulation in vivo is not clear.

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Calcineurin B is N-terminally myristoylated [Aitken et al., 1982]. A mutant, in which myristoylation is abolished retains the major properties of wild-type enzyme, except for decreased thermal stability [Kennedy et al., 1996]. It has been shown recently that myristoylation is also required for calcium-dependent calcineurin binding to phosphatidylserine vesicles, indicating contribution of myristoylation to membrane-binding properties of calcineurin [Perrino and Martin, 2001].

Phospholipids could modulate calcineurin activity with both activation and inhibition reported for different lipid types and substrates [Huang et al., 1983; Politino and King, 1987].

In addition, arachidonic acid and other unsaturated fatty acids activated recombinant calcineurin from Dictyostelium discoideum with no effect on bovine enzyme [Kessen et al., 1999]. Since part of calcineurin in vivo is localized to cell membranes this kind of modulation is probably of physiological significance.

Some natural compounds exhibit potent inhibition of calcineurin activity. The most potent inhibitors are the immunosuppressive drugs CsA and FK506 [Liu et al., 1991a;

Swanson et al., 1992]. They are effective only when bound to their respective binding proteins, cyclophilin and FKBP, and do not inhibit activity towards low molecular weight substrates like pNPP. Okadaic acid, a potent inhibitor of related phosphatases PP1 and PP2A, can also inhibit calcineurin with IC50~4 µM compared to 300 nM for PP1 and 1 nM for PP2A [Bialojan and Takai, 1988]. Pyrethroid insecticides (cypermethrin, deltamethrin) potently inhibited calcineurin activity towards pNPP [Enan and Matsumura, 1992], but later proved ineffective towards RII and phosphoproteins [Enz and Pombo-Villar, 1997; Fakata et al., 1998]. A 25-residue peptide corresponding to the sequence of calcineurin autoinhibitory domain (residues 457-481) also potently inhibits the enzyme activity [Hashimoto et al., 1990].

Similar to other phosphatases, calcineurin is inhibited by orthovanadate [Morioka et al., 1998], phosphate and pyrophosphate [King and Huang, 1984] and fluoride [Tallant and Cheung, 1984]. In addition, calmodulin antagonists such as trifluoroperazine [Stewart et al., 1983], W-7 and calmidazolium [Mukai et al., 1991] prevent calcineurin activation by calmodulin and thus serve as calcineurin inhibitors. Recently, calcineurin activity towards both RII and pNPP has been found to be inhibited by the polyphenolic aldehyde gossypol and its analogues with effective concentrations in the micromolar range [Baumgrass et al., 2001].

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1.2.6. CALCINEURIN X-RAY STRUCTURE.

Based on homology of serine/threonine phosphatases to purple acid phosphatases, enzymes with well-characterized catalytic binuclear metal center, it was predicted that serine/threoninie phosphatases might also contain this center at their active site [Vincent and Averill, 1990]. Later, sequence alignments of serine/threonine phosphatases identified a

“phosphodiesterase motif” conserved in calcineurin, PP1, PP2A and in other enzymes cleaving phosphoester bonds such as bacterial exonucleases, alkaline and acid phosphatase, λ- phosphatase, phosphodiesterase and 5´-nucleotidase [Koonin, 1994; Lohse et al., 1995] (Fig.

1.2).

Figure 1.2. Phosphodiesterase motif in calcineurin and purple acid phosphatases.

Residues identified as metal ligands are shown in bold, and conserved non-ligand residues of the active site are underlined.

The phosphodiesterase motif presumably provides a scaffold for the binuclear metal center in each member of the phosphodiesterase family, and the existence of such a center was proved for λ-phosphatase [Rusnak et al., 1999b] and 5´-nucleotidase [Knöfel and Ströter, 1999]. The conservancy of this motif suggests a common catalytic mechanism for the enzymes involved in phosphotransfer reactions.

The list of metallophosphatases whose X-ray structures have been solved up to date includes not only calcineurin [Griffith et al., 1995; Kissinger et al., 1995], but also PP1 [Egloff et al., 1995; Goldberg et al., 1995], kidney bean purple acid phosphatase [Strater et al., 1995], mammalian purple acid phosphatase [Guddat et al., 1999; Lindquist et al., 1999;

Uppenberg et al., 1999] and λ-phosphatase [Voegtli et al., 2000]. In these structures the phosphodiesterase motif forms a β-α-β-α-β scaffold for a binuclear metal center. The three β- strands form a parallel pleated sheet capped by intervening α-helices. Two metal ions are

Phosphoesterase Consensus: DXH (X)n GDXXDR (X)m GNHD/E

Calcineurin and PP1: DIH (X)23 GDYVDR (X)27 GNHE

Purple Acid Phosphatase: DXG (X)n GDXXYD (X)m GNHD/E

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situated at the apex of this fold with a distance between them 3-4 Å. The residues in loops between β-strands and α-helices provide metal ligands.

The crystal structure of recombinant human calcineurin [Kissinger et al., 1995] is shown on Fig. 1.3. The calcineurin B-binding domain forms an amphipatic α-helix, whose top nonpolar face contacts calcineurin B subunit. The autoinhibitory domain folds into an α-helix that occupies the substrate-binding cleft. The calmodulin-binding domain is disordered and not visible on the electron density map, consistent with the sensitivity of this region to proteolytic degradation. Therefore, the structural details of calmodulin regulation remain obscure.

Figure 1.3. Crystal structure of recombinant human calcineurin. Diagram showing the three-dimensional structure of calcineurin using the X-ray coordinates of Kissinger et al.

([Kissinger et al., 1995], protein data bank file 1AUI). The figure was created using public domain software Cn3D (http://www.ncbi.nlm.nih.gov/Structure/CN3D/cn3d.shtml).

Calcineurin A catalytic domain is shown in cyan, calcineurin B-binding domain and part of the autoinhibitory domain are in gray, and N- and C-terminal lobes of calcineurin B are in dark blue and green, respectively. Spheres at the catalytic domain represent iron and zinc, and those at B subunit bound calcium ions.

Calcineurin B contacts calcineurin A by a groove formed by its carboxy- and amino-terminal lobes and its C-terminal strand. In addition to amphipatic α-helix residues 14-23 of

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calcineurin A provide a contact surface with carboxy-terminal lobe of calcineurin B. In the other structure [Griffith et al., 1995] the myristoyl group of calcineurin B lies in the hydrophobic cleft between amphipatic α-helices in the N-terminal part of calcineurin B and forms multiple hydrophobic contacts with the protein.

The active site of calcineurin is formed around a binuclear metal center (Fig. 1.4). The presence of Fe and Zn in the center has been verified by atomic absorption spectrometry [King and Huang, 1984; Yu et al., 1995]. A similar metal coordination and environment was found in PP1 [Goldberg et al., 1995], but the nature of metals in its active site remains unclear. The positions of Fe and Zn were attributed according to the positions of Fe and Zn in kidney bean purple acid phosphatase, where a tyrosine residue coordinates Fe3+ at the M1 site (left on the Fig. 1.4) and provides a characteristic charge transfer band at 510-550 nm in the enzyme optical spectrum [Strater et al., 1995]. In calcineurin this tyrosinate is replaced by a histidine, and the enzyme is colorless.

O O Asp 121

H His 151 N

N

H O

Fe N

His 92 N

OH OH

O

Zn O

Asp 90O Asp 118

N N

His 199

N N

His 281 O

N Asn 150

O

Fig. 1. 4. Schematic view of the active site of human calcineurin [Kissinger et al., 1995].

In addition to a histidine (His92 in human calcineurin) the M1 iron is coordinated by two aspartates, Asp90 and Asp118. The latter is a bridging ligand also coordinating the M2 zinc atom. This atom is additionally coordinated by side chains of Asn150, His199 and His281. Apart from protein aminoacid residues solvent molecules coordinate the metals of the active center, one of them in a µ-bridging mode. His151 and Asp121, although not binding metals directly, participate in catalysis as discussed below.

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1.2.7. CALCINEURIN CATALYTIC MECHANISM.

Two major mechanisms of phosphate ester hydrolysis are discussed in literature: one involving formation of the phosphoenzyme intermediate and the other involving direct attack of the solvent nucleophile on the ester bond (Fig.1.5). The former occurs in catalysis by alkaline phosphatase, where an active site serine accepts a phosphoryl group during enzymatic turnover [Coleman, 1992]. Another example is provided by tyrosine phosphatases, where an active site cysteine plays the analogous role [Zhang 1998].

Kinetic experiments on calcineurin gave evidence supporting direct transfer of the phosphoryl group to water without formation of the phosphoenzyme intermediate. First, a direct relationship

Fig.1.5. Mechanisms of phosphate ester hydrolysis.

between log (V/K) and the pKa of the calcineurin substrates was observed [Martin et al., 1985] with increase of the rate following decrease of the leaving group pKa. Second, there was no phosphotransferase activity towards alternative nucleophiles [Martin et al., 1985].

Both phosphate and phenol were competitive inhibitors of p-NPP hydrolysis by calcineurin, consistent with a random uni-bi mechanism (in contrast to an ordered uni-bi mechanism for phosphoenzyme intermediates) [Martin and Graves, 1986]. Most convincing evidence for the direct phosphoryl group transfer to water came from the work on spleen purple acid phosphatase [Mueller et al., 1993]. Using a chiral [18O, 17O]phosphorothiolate ester the authors showed that catalytic hydrolysis of the ester bond occurred with a net inversion of configuration at the phosphorus atom, directly indicating the phosphoryl transfer to water without formation of the phosphoenzyme intermediate. Since the active site of PAPs is similar

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to calcineurin, it is possible to presume that catalysis by calcineurin follows the same mechanism.

The role of metal ions in the catalytic mechanism of calcineurin is still not clear. Metal ions could serve as Lewis acids in activating a solvent nucleophile like in carbonic anhydrase [Hakansson et al., 1992]. They could neutralize the negative charge on the phosphate oxygen atoms making the phosphorus atom more suitable for nucleophilic attack, and also stabilize the developing charge on the leaving group during the ester bond cleavage. Additionally, metals could help orient the substrate for in-line attack of the nucleophile.

Several conserved aminoacid residues are situated within 4-9 Å of the binuclear metal center of calcineurin. The contribution of these residues to calcineurin catalysis was investigated by site-directed mutagenesis. These studies highlighted the importance of the conserved histidine residue (His151) (Fig.1.4) in a couple with an equally conserved aspartate (Asp121). These residues form part of a phosphodiesterase motif shown in Fig. 1.2 and are present in all metallophosphatases. Mutation of His151 causes ~103-fold reduction of rate constant (kcat) [Mertz et al., 1997], and a similar decrease occurred after mutation of Asp121 analogue in PP1 [Zhang et al., 1996; Huang et al., 1997] without greatly affecting KM in both cases. His151 could assist in substrate binding or orient the nucleophilic solvent molecule, as well as participate in acid/base catalysis. Similar differences in kcat for wild-type and His151 mutant enzyme with substrates having very different pKa of their leaving groups [Mertz et al., 1997] argue against a role of that residue as acid catalyst. On the other hand, this histidine could serve as a general base deprotonating nucleophilic water molecule. With Asp121 it might form an Asp-His-HO triad similar to the Asp-His-Ser triad of serine proteases. Apart from His-Asp pair two conserved arginines, Arg122 and Arg254, are in the vicinity of the calcineurin active site. Mutation of these arginines caused a 102-103-fold reduction of kcat and a 2-10-fold KM increase for Arg254 [Mondragon et al., 1997]. These residues contact oxygen atoms of phosphate and may provide stabilization for binding negatively charged phosphoester and also neutralize developing charge in the transition state. In purple acid phosphatases these arginines are substituted by histidines, probably explaining the lower pH optimum of PAPs [Klabunde et al., 1996].

The model for the phosphate ester hydrolysis by calcineurin proposed by Rusnak and Mertz [Rusnak and Mertz, 2000] is presented in Fig. 1.6. According to this model upon substrate binding the negative charge on phosphate is neutralized by Arg122 and Arg254 together with Zn2+, while His151 removes the proton from the metal-bound water and orients the solvent nucleophile for optimal attack on the phosphate ester (Fig. 1.6.A). A dissociative

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transition state then forms (Fig. 1.6.B), where bond cleavage to the leaving group occurs before bond formation to the solvent nucleophile. The negative charge on the leaving group is neutralized by His151 and Zn2+. After bond cleavage and proton transfer to the leaving group the result is the product-inhibited state with phosphate bridging two metals as was shown for the X-ray structure of calcineurin [Griffith et al., 1995] (Fig. 1.6.C). Exchange of the phosphate for solvent molecule restores the active enzyme form.

Fig.1.6. Mechanism of calcineurin catalysis. In this model His151 acts as a general base and a general acid. A. Substrate binds to the active site and His151 assists in deprotonating water bound to iron with hydroxide formation. B. The transition state is dissociative, zinc helps to neutralize charge on the leaving group, which is protonated by His151. C. The product-inhibited state with phosphate bridging two metal ions. D. A water molecule displaces the phosphate, the enzyme is ready for the next catalytic turn.

1.2.8. PHYSIOLOGICAL FUNCTIONS. ROLE OF CALCINEURIN IN YEAST.

Saccharomyces cerevisiae provides an important model system for studying calcineurin function, especially by using genetic manipulation of its expression.

Saccharomyces cerevisiae have two isoforms of calcineurin catalytic subunit (CNA1/CMP1 and CNA2/CMP2) and one gene encoding regulatory subunit (CNB1) [Cyert et al., 1991;

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Kuno et al., 1991; Liu et al., 1991b]. Strains deficient in both isoforms of calcineurin A or in calcineurin B failed to recover from α-factor-induced growth arrest [Cyert et al., 1991; Cyert and Thorner, 1992] and were hypersensitive to salt stress (induced by Na+ and Li+) [Mendoza et al., 1994] as well as to Mn2+ [Farcasanu et al., 1995]. Similar phenotypic responses were observed after treatment of the cells with the immunosuppressants FK506 and CsA [Foor et al., 1992; Nakamura et al., 1993]. Calcineurin was shown to influence the expression of several yeast genes, including vacuolar and secretory Ca2+-pumps (Pmc1p and Pmr1p), β-1,3 glucan synthase (Fks2p) and a plasma membrane Na+ pump (Pmr2p) [Mendoza et al., 1994;

Matheos et al., 1997; Zhao et al., 1998]. The latter is involved in salt stress resistance [Mendoza et al., 1994], while Pmr1p is linked to Mn2+-homeostasis [Matheos et al., 1997], and Fks2p is the putative calcineurin-responsive element in the response to α-factor [Zhao et al., 1998]. The transcription factor Crz1p/Tcn1p was identified as a mediator of calcineurin- dependent gene expression [Matheos et al., 1997; Stathopolous and Cyert, 1997]. Mechanism of Crz1p/Tcn1p activation involves calcineurin-mediated dephosphorylation of the Crz1p/Tcn1p region having similarity to the mammalian NFAT transcription factor family (reviewed later in the text) followed by the nuclear translocation mediated by the nuclear import protein Nmd5p [Stathopoulos-Gerontides et al., 1999; Polizotto and Cyert, 2001].

Apart from the Crz1p/Tcn1p vacuolar H+/Ca2+ exchanger Vcx1p [Cunningham and Fink, 1996] as well as Hsl1, a kinase participating in yeast cell cycle regulation [Mizunuma et al., 2001] appeared to be directly modified by calcineurin.

Although biochemical properties of the yeast calcineurin seem to be identical to those of mammalian calcineurin, the later mutagenesis work identified a new region essential for the activity of yeast calcineurin [Jiang and Cyert, 1999]. This region is situated between catalytic and calcineurin B-binding domains, and mutation of several residues within this domain (S373P, H375L, and L379S) dramatically decreased the enzyme phosphatase activity.

It is not clear whether the same mutations impair the activity of mammalian calcineurin.

In Schizosaccharomyces pombe calcineurin serves functions distinct of those in budding yeast. Calcineurin mutants are defect in cytokinesis, polarity, mating and spindle body positioning [Yoshida et al., 1994] and are Cl--hypersensitive [Sugiura et al., 1998].

Among other lower eukaryotes, calcineurin is essential for growth of Aspergillus nidulans and Neurospora crassa [Higuchi et al., 1991; Rasmussen et al., 1994].

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1.2.9. CALCINEURIN IN T-CELL ACTIVATION.

The role of calcineurin in T-cell signaling was thoroughly investigated during the last 10 years and is discussed in several specialized reviews [Rao et al., 1997; Crabtree, 1999;

Crabtree, 2001; Macian et al., 2001]. T-cell activation cascade is initiated upon the binding of antigen-MHC complexes on the surface of the antigen-presenting cell to the T-cell receptor (TCR) on the reactive T-lymphocyte, and results in the complex response marked by the up- regulation of expression levels of a plentitude of lymphocyte genes, including IL-2. IL-2 then activates lymphocyte proliferation. The involvement of calcineurin in T-cell activation was revealed by the discovery of its inhibition by immunosuppressants CsA and FK506 [Liu et al., 1991]. Another critical component of the activation cascade was identified as transcription factor NFAT (nuclear factor of activated T-cells) [Shaw et al., 1988; Flanagan et al., 1991;

Clipstone and Crabtree, 1992]. NFAT comprises a family of 5 proteins (NFAT1-5), four of which are regulated in a calcineurin-dependent manner [Macian et al., 2001]. Two of them, NFAT1 and NFAT2, are critical for the T-lymphocyte cytokine production [Peng et al., 2001], with NFAT4 also contributing to T-cell function [Oukka et al., 1998]. The activation of the calcineurin-NFAT pathway begins with a receptor-initiated release of calcium from intracellular stores followed by calcium influx through store-operated channels (capacitative calcium entry). Patients having a defect of capacitative calcium entry suffer from severe immunodeficiency associated with a failure of NFAT activation [Feske et al., 2000]. Low sustained increases of intracellular calcium preferably activate NFAT, while shorter calcium spikes could activate other signaling components (e.g. NFκB) without affecting NFAT [Dolmetsch et al., 1997]. These calcium signals cause optimal activation of the calcineurin phosphatase activity. Calcineurin is constitutively bound to NFAT through the PxIxIT consensus motif in the regulatory domain [Aramburu et al., 1998], and peptides modeled on this site are potent inhibitors of NFAT activation by calcineurin [Aramburu et al., 1998;

Aramburu et al., 1999]. Some other parts of NFAT could also contribute to the binding [Liu et al., 1999; Park et al., 2000]. Calcineurin activation results in dephosphorylation of multiple serine residues in the NFAT regulatory domain [Ruff and Leach, 1995; Park et al., 1995].

Detailed analysis of NFAT1 [Okamura et al., 2000] identified 14 phosphoserine in a resting state, 13 of which are dephosphorylated upon stimulation. In NFAT1 first removal of five phosphates from a serine-rich sequence adjacent to PxIxIT motif exposes nuclear localization signal (NLS) in the NFAT regulatory domain and renders other eight phosphoserines more accessible to calcineurin. Dephosphorylation causes NFAT translocation into the nucleus

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