CHEMICAL BIOLOGY
Nitric Oxide, Biological Targets of
Emily R. Derbyshire and Michael A. Marietta, University of California, Berkeley
doi: 10.1002/9780470048672.wecb387
Nitric oxide (NO) is an essential signaling molecule for many eukaryotic organisms. NO is produced in vivo by the enzyme nitric oxide synthase (NOS) from the amino acid t-arginine. The apolar gas readily diffuses across cell membranes, where it binds to the heme of soluble guanylate cyclase (sGC), the principle NO receptor. Once activated, sGC converts GTP to cGMP at a rate that is several-hundred-fold above the basal level. This NO/cGMP signaling cascade modulates several physiologic processes including vasodilation, platelet aggregation, and neurotransmission. Although the cGMP-dependent affects of NO remain active areas of research, additional cGMP-independent responses to NO also are being investigated. Endogenous levels of NO can modulate protein function by S-nitrosation, a covalent modification that has been implicated in the transcriptional regulation of genes involved in the immune response and in apoptosis.
In biologic systems, nitric oxide (NO) functions as both a critical cytotoxic agent and an essential signaling molecule. The toxicity of the diatomic gas has long been accepted; however, nitric oxide was not known to be a physiologically relevant signaling molecule until it was identified as the endothelium-derived relaxing factor (EDRF) (reviewed in Reference 1). Since this discovery, the enzymatic synthesis of NO and the signaling pathways that it regulates have been the focus of many studies. In higher eukaryotes, nitric oxide synthase (NOS) produces NO from L-arginine (reviewed in References 2-4). Despite several years of research, the NOS catalytic mechanism remains a topic of investigation, but commonly it is accepted that NO is essential for several physiologic processes. Many signaling responses that NO modulates are mediated by the NO-induced activation of the heme protein soluble guanylate cyclase (sGC). NO binds to sGC at a diffusion-controlled rate and leads to a several- hundred-fold increase in the synthesis of the second messenger cGMP from GTP (5, 6). Other diatomic gases either do not bind (dioxygen) or do not activate sGC significantly (carbon monoxide). This characteristic provides selectivity and efficiency for NO even in an aerobic environment, which is critical because of the high reactivity of NO. The NOS/sGC pathway is important for maintaining homeostasis, and many diseases have been linked to the dysfunction in NO signaling (reviewed in Reference 7). Studies on cGMP-dependent NO responses continue to expand, and other roles for the gas are emerging in both prokaryotic and eukaryotic organisms. In higher eukaryotes, protein S-nitrosation, an oxidative modification of cysteine residues, is implicated in an increasing number of cGMP-independent signaling systems (reviewed in Reference 8), whereas in bacteria, a class of potential heme-based NO sensors recently has been identified and proposed to participate in two-component signal transduction pathways (9).
Background and Significance
NOS is regulated highly to ensure that NO concentrations do not reach toxic levels and to control properly the processes that respond to the signaling molecule. At high concentrations (low mM), NO in an aerobic environment can damage DNA, oxidize critical heme proteins, and covalently modify essential biologic molecules (10-12). To deter these events, the expression, cellular localization, and activity of NOS is regulated highly (reviewed in Reference 3). All three isoforms, endothelial, neuronal, and inducible NOS (eNOS, nNOS, and iNOS), are regulated at the transcriptional level. Both eNOS and nNOS are expressed constitutively, and iNOS is induced with the appropriate immunostimulatory signals. The isoforms critical to signal transduction pathways (eNOS and nNOS) generate low nanomolar levels of NO and are regulated in vivo by the binding of calcium and calmodulin. NO produced by NOS can diffuse rapidly across a cell membrane to activate sGC, a hemoprotein that has evolved to bind NO selectively even in the presence of oxygen (μM) (Fig. 1). In addition to NO, cGMP production by sGC is regulated by the nucleotides GTP and ATP (13, 14), which are present at ~0.2 and ~1.7 mM, respectively, in vivo (15). The efficient binding of NO by sGC allows for the rapid production of cGMP, which then binds to phosphodiesterases (PDE), ion-gated channels, and cGMP-dependent protein kinases (cGK) to regulate several physiologic functions including vasodilation, platelet aggregation, and neurotransmission (16-18). The amplitude and duration of these cGMP effects are regulated additionally by the activity of PDE, the enzyme that hydrolyzes cGMP (reviewed in Reference 19).
The importance of this signaling pathway has been demonstrated in mouse models in which the triple NOS knockouts exhibit characteristics consistent with nephrogenic diabetes insipidus (20). Knockouts of the sGC β1 subunit exhibit elevated blood pressure, reduced heart rate, and dysfunction in gastrointestinal contractility (21), and studies on mice deficient of the sGC α1 subunit indicate that the protein is essential for NO-mediated pulmonary vasodilation (22). Additionally, several diseases have been linked to defects in the NO signaling pathway. Independent of cGMP production by sGC, NO also can affect biologic processes by covalently modifying and/or oxidizing proteins. Here, we summarize the best-characterized physiologic responses to NO.
Figure 1. Nitric oxide signal transduction pathway. NO synthesized by NOS diffuses across cell membranes to a target cell. NO activates sGC, which leads to an increase in cGMP synthesis. The oxidation products of NO also can react with protein thiols, which leads to protein 5-nitrosation.
cGMP-dependent signaling
NO is important for the function of the cardiovascular system and is critical for blood pressure regulation. In vascular smooth muscle cells, cGMP can bind to and activate cGK, specifically the type Iα and Iβ isoforms. These isoforms are splice variants of the same gene that has different sensitivities to cGMP. During activation, cGK phosphorylates the large conductance Ca2+-activated K+ channel (23) and IRAG (IP3 receptor associated cGMP kinase substrate) (24), which are involved in the regulation of extracellular Ca2+ entry and intracellular Ca2+ release, respectively. The release of Ca2+ into the cytosol leads to smooth muscle contraction by the activation of a myosin light chain kinase (MLCK) that phosphorylates the myosin light chain (MLC) (reviewed in Reference 25). Smooth muscle contraction also is regulated by myosin light chain phosphatase (MLCP), the protein that dephosphorylates MLC. cGKI phosphorylates and inhibits Rho A, a GTPase that activates Rho kinase. Rho kinase inhibits the activity of MLCP, and therefore the cGMP-dependent inhibition of Rho A contributes to smooth muscle relaxation (26). Vasodilation also is modulated by PDE5, a major cGMP-hydrolyzing PDE. This protein is important for inducing relaxation under low Ca2+ conditions and has become an important drug target because the inhibition of PDE5 leads to increased levels of cGMP after NO-induced sGC stimulation (19).
sGC activation also is important for the immune response. Human platelets generate cGMP after NO activation of sGC, which leads to the inhibition of platelet activation or aggregation. This effect is mediated primarily by cGMP activation of cGKI. Many targets for activated cGKI have been proposed, including the vasodilator stimulated phosphoprotein (VASP). Phosphorylation of VASP correlates with the binding of fibrinogen to glycoprotein IIb/IIIa, expression of P-selectin, and platelet adhesion (reviewed in References 16 and 27). Small molecule sGC activators (28-30) have been shown to inhibit platelets and are potential antithrombic agents that could be used to treat cardiovascular diseases.
cGMP-independent signaling
The oxidative addition of NO to a thiol, termed S-nitrosation, is a posttranslational modification that can modulate protein function. With high concentrations of NO, this modification can alter protein function indiscriminately; however, only a limited number of proteins are S-nitrosated in vivo (8). This selectivity of nitrosothiol formation suggests that a mechanism of regulation of SNO formation and/or decay exists; however, the details of this regulation are unknown.
The S-nitrosation of proteins has been implicated in regulating apoptosis, protein expression, and tissue oxygenation (31-33). For example, low levels of NO can inhibit apoptosis via the S-nitrosation of caspase proteases, which contain a cysteine residue that is essential for catalytic activity (33). Furthermore, this process may be regulated by the protein thioredoxin, the primary intracellular oxidoreductase that may function as a nitrosotransferase (34). S-nitrosation also inhibits the DNA binding activity of NF-kB transcription factors, which effects protein expression (31), and S-nitrosohemoglobin has been implicated in the regulation of blood flow and tissue oxygenation (reviewed in References 32 and 35).
Interestingly, both NOS and sGC have been shown to be S-nitrosated by low levels of NO (36-38). In NOS, this nitrosation occurs at zinc tetrathiolate cysteines that are critical for maintaining a functional dimer. Modification of these cysteines leads to the formation of inactive monomers, which could be a means of regulating NO production in vivo (37). S-nitrosation of sGC results in the inhibition of NO-stimulated activity (38). This mechanism of desensitization may account for the clinical condition known as NO tolerance, which is an ongoing problem in the treatment of heart disease.
No Chemistry
NOS
Mammalian NOS is a P-450-like enzyme that catalyzes the oxidation of L-arginine to L-citrulline and NO. This process is a two-step reaction that leads to a five-electron oxidation of L-arginine. The enzyme requires NADPH and O2 as substrates for both reaction steps, and iron protoporphyrin IX (heme), FMN, FAD, and tetrahydrobiopterin (H4B) as protein-bound cofactors. NOS is active as a homodimer and contains an N-terminal oxygenase (or heme) domain, a C-terminal flavopro- tein reductase domain, and a central calmodulin binding region (4) (Fig. 2a). The heme domain of NOS (NOSheme) can be isolated, and it binds heme, H4B, and L-arginine. This domain is functional if provided with reducing equivalents such as sodium dithionite (39, 40). The crystal structures of eNOSheme(41) and iNOSheme (42) show how substrate and cofactors bind within the active site and identify residues that are important for H4B binding and dimerization, including a zinc tetrathiolate at the bottom of the dimer interface that stabilizes subunit binding and is involved in maintaining the integrity of the H4B binding site (43). The reductase domain binds to NADPH, FMN, and FAD and provides electrons to the heme active site for catalysis, a process that is controlled by Ca2+/calmodulin binding.
In the first step, L-arginine is hydroxylated to form N“-hydroxy-L-arginine (NHA) (Fig. 2b). This reaction mechanism is analogous to those catalyzed by cyctochrome P-450s, which involves a proposed high-valent oxo-iron intermediate that could transfer an activated oxygen species to a substrate. In the second reaction step, the 3-electron oxidation of NHA produces L-citrulline and NO. It has been proposed that this step involves the attack of a ferric peroxide intermediate on the guanido carbon; however, experimental evidence is not sufficient to distinguish between this and other proposed mechanisms (reviewed in Reference 2). The most controversial questions about the NOS mechanism concern the source of the electrons in each reaction step and cofactor stoichiometry.
NO can have a short half-life in aqueous solution, which may seem problematic for it to reach its intracellular target. A second-order dependence exists on NO autoxidation shown in the rate law below (reviewed in Reference 44).
Consequently, at nanomolar signaling concentrations, the lifetime of NO is sufficient for it to reach sGC. The end-products of NO decomposition are nitrite (NO2-) and nitrate (NO3-). NO and reaction intermediates along the decomposition pathway can react with several intracellular molecules, but reactions with heme cofactors and cysteines are the most relevant to its function as a signaling agent and also contribute to its toxicity.
Figure 2. Nitric oxide synthase. (a). Domain architecture of NOS. The heme domain binds Zn2+ (gray box), heme (gray parallelogram), and H4B (white box). The reductase domain binds FMN, FAD, and NADPH (white boxes). CaM (white box) is between the heme domain and the reductase domain. (b). Two-step reaction scheme for NO synthesis by NOS.
Reactions with heme
Perhaps the best-characterized interactions are between NO and the heme proteins hemoglobin and myoglobin. NO binds to FeII-unligated globins on the order of 107 M-1s-1 and leads to the formation of a stable 6-coordinate FeII-NO complex (45). These 6-coordinate complexes can be very stable and thereby inhibit the function of heme proteins by blocking the coordination site. NO also can react with FeII-O2 complexes, which often leads to heme oxidation (FeIII-heme) and the formation of NO3-. With hemoglobin and myoglobin, this reaction occurs on the same order of magnitude as simple NO binding to the ferrous heme (10).
The affinity of NO for ferric heme is significantly lower than for the ferrous heme (46), but reductive nitrosylation of proteins has been observed (47). In this reaction, one equivalent of NO reduces the heme to FeII, and a second equivalent of NO rapidly binds to the unoccupied coordination site. This reaction also can generate the nitrosating agent NO+, which can react subsequently with free thiols. In fact, the S-nitrosation of hemoglobin and nitrophorin, a protein involved in NO storage and delivery in some bloodsucking insects, has been observed after exposing NO to the ferric form of the proteins (48, 49). As the ferric heme is the physiologically relevant state for nitrophorins, it is likely that this reaction occurs in vivo.
It is evident from these reactions that O2 binding to sGC would reduce the ability of the enzyme to function as a selective NO sensor. The mechanism of sGC activation by NO and the ability of NO to discriminate against O2 currently are under investigation.
sGC activation and ligand discrimination
sGC is a heterodimeric protein that consists of two homologous subunits, α and β. The most commonly studied isoform is the α1β1 protein; however, the α2 and β2 subunits also have been identified (50, 51). sGC contains an N-terminal heme binding region, a predicted PAS-like region, and a C-terminal catalytic domain (reviewed in Reference 52) (Fig. 3a). Truncations of the sGC heme domain as well as sGC-like homologues discovered in bacteria (9) have facilitated the study of the heme environment and ligand binding (53-55).
NO binds to the heme of sGC at a diffusion-controlled rate to form an initial 6-coordinate complex, which rapidly converts to a 5-coordinate ferrous nitrosyl complex (Fig. 3b) (52). The breaking of the Fe-His bond is thought to be critical to the activation of sGC by NO; however, recent data has shown that the NO coordination to the heme is not sufficient for full activation (13, 56). A low-activity FeII-NO complex can be formed in the presence of stoichiometric amounts of NO, and this species is identical spectroscopically to the highly active form of the enzyme that is formed in the presence of substrate or excess NO. Based on these observations, two mechanisms of NO activation have been proposed. One proposal is that excess NO activates the ferrous nitrosyl complex by binding to nonheme sites on the protein (13). The second proposal involves excess NO binding to the heme to form a transient dinitrosyl complex, which then converts to a 5-coordinate complex with NO bound in the proximal heme pocket (56).
The ability of sGC to select against O2 binding is important for it to function as a NO sensor because O2 is present at much higher levels than NO in vivo and FeII-O2 and FeIII proteins react rapidly with NO. Interestingly, some bacterial sGC-like homologues also bind O2 and NO. These proteins were named heme nitric oxide/oxygen binding (H-NOX) proteins (reviewed in Reference 57). The crystal structure of the O2-binding H-NOX from Thermoanaerobacter tengcongensis (Tt.) with O2 bound was reported recently (54, 55). This structure shows that a distal pocket tyrosine interacts with bound O2though an H-bond. Whereas the crystal structure of the O2, excluding H-NOX from Nostoc sp., shows that no hydrogen bond donor exists in the distal heme pocket (53), sequence analysis predicts that polar residues capable of interacting with O2 are absent in sGC. Mutagenesis studies that introduced a Tyr into the distal pocket of the β1 H-NOX domain produced a protein that was capable of binding O2 (58); however, the same mutation in full-length sGC did not facilitate O2 binding (59, 60). This finding indicates that the presence of a distal pocket Tyr may be involved in stabilizing FeII-O2complexes in H-NOX proteins, but other factors are involved in ligand discrimination in sGC. The size and overall polarity of the heme distal pocket and the strength of the proximal Fe-His bond have been proposed as mechanisms of discriminating against O2 binding (57). The crystal structures of the H-NOX proteins also have facilitated the study of sGC activation. Specifically, the differential pivoting and bending in the H-NOX heme during NO or CO binding may account for the varying degree of activation induced by the two ligands (200-fold versus 4-fold, respectively) (53). However, details about how movement in the H-NOX domains affect the catalytic domain may remain unresolved until the full-length structure is elucidated.
Figure 3. Soluble guanylate cyclase. (a). Domain architecture of sGC. sGC consists of two homologous subunits, α1 and β1. Each subunit contains an N-terminal H-NOX domain, a central predicted PAS-like region, and a C-terminal catalytic domain. Heme (gray parallelogram) binds to the H-NOX domain on the pi subunit. (b). NO activation of sGC. NO binds to the sGC heme, which leads to the formation of a 5-coordinate ferrous nitrosyl complex and activates the protein several-hundred-fold above the basal level.
Reactions with cysteine
Many pathways exist to generate a nitrosothiol in vitro by the 1-electron oxidation of NO. Nitrosothiols can be formed via the reaction of a thiol with N2O3, a nitrosating agent that is an intermediate in the decomposition of NO in aerobic solution, or via the direct reaction of NO with a thiol to form an addition complex (SNO-) followed by a 1-electron oxidation. S -nitrosation of a protein thiol also can occur by a trans-S-nitrosation event from a low molecular weight nitrosothiol, such as S-nitrosoglutathione, or from a nitrosated protein cysteine (8). Whereas the in vivo mechanism of protein S-nitrosation is unknown, a protein-mediated trans-S-nitrosation mechanism is an attractive possibility because of the specificity it could impart on the reaction. Additionally, the same protein could catalyze both the nitrosation and denitrosation of a specific cysteine. A report showing that the protein thioredoxin can transnitrosate caspase-3 selectively and reversibly lends support to this proposal (34).
Chemical Tools and Techniques
Enzymology
The study of an enzyme requires a functional assay, or a quantitative method for measuring the conversion of substrate to product. To develop this method the enzyme substrate(s) and product(s) must be identified. Isotopically or radiolabeled compounds can facilitate this identification. NO was identified first as an intermediate in the L-arginine to NO2-/NO3- pathway by incubating macrophage cells with L-[guanido-15N2]arginine (61). Similar experiments showed that L-arginine was the precursor in this pathway (62, 63), that L-citrulline was an additional product (63), and that L-arginine conversion to L-citrulline was coupled with NO formation (64). Chromatographic methods to separate substrate from product, radioimmunoassays (RIA), or enzyme-linked immunosorbent assays (ELISA) can be used for the sensitive detection of reaction products and/or substrate. Reaction stoichiometry, turnover number (kcat), and KM for substrate can be determined. Additionally, reaction intermediates and possible transition states can be investigated by rapid quench methods, design of rational based inhibitors, and isotope exchange experiments.
Spectroscopy of heme proteins
Iron porphyrin complexes, or heme, have diverse functions in biologic systems. Specifically the iron(II) complex of protoporphyrin type IX is a cofactor critical to the production of NO by NOS and the subsequent activation of sGC by NO. The highly conjugated n-system of these porphyrins gives these proteins their characteristic color and facilitates the study of the biologic systems they regulate. Several methods can be used to study the ligation state and heme environment of these proteins. Briefly, electronic absorption, resonance Raman, and electron paramagnetic spectroscopy will be discussed; however, several useful techniques are used to investigate heme proteins (reviewed in Reference 65).
Electronic absorption spectroscopy can be used to determine the general structure of porphyins and their derivatives. The oxidation and coordination state of the iron and the identity of the amino acid that ligates the heme can be examined by comparing the absorption spectrum of the protein of interest with the spectra of known heme proteins (45, 65). General characterization with electronic absorption spectroscopy indicated that NO and CO, but not O2, bind to the sGC heme moiety (66). Dynamic studies of ligand binding and dissociation also can be examined with this technique on psec-msec time scales with standard stopped-flow systems.
Heme proteins have been studied extensively with resonance Raman (RR) spectroscopy, a method that can be used to examine the environment around a heme cofactor and to confirm the atoms that coordinate to the metal center. A Raman spectrum contains peaks of scattered light where the observed frequency shifts correspond to the various vibrational frequencies of the scattering molecules. Specifically, the oxidation state of the heme is indicated by the electron density marker, v4, in the 1350-1380 cm-1 region, and v2, v3, and v10 are the spin and coordination state markers (65). Different ligation states of sGC (67) and NOS (68, 69) have been characterized with this technique. Interestingly, the resonance Raman spectra of sGC FeII-NO and FeII-CO complexes are influenced by the presence of GTP and known sGC allosteric activators (70, 71). This influence indicates that conformational changes exist at the heme pocket during substrate and activator binding that may correlate with activation.
Electron paramagnetic resonance (EPR) is a powerful tool for studying radicals such as NO. This method specifically detects molecules with unpaired electrons. The g-value, a dimensionless parameter determined from an EPR spectrum, is influenced by the spin and orbital angular momentum of the unpaired electron (65, 72). The application of EPR to the structural study of metalloproteins and electron transfer systems has expanded because of the development of pulsed-EPR techniques and high-field/high-frequency spectrometers coupled with advances in rapid-freeze quench systems. Typically, EPR experiments are performed around 9 GHz (X-band), but studies also are done at 2 GHz (L-band), 4 GHz (S-band), 24 GHz (K-band), and 35 GHz (Q-band) frequencies (72). EPR studies with sGC confirmed that the nitric oxide radical was binding to the heme moiety and that the sGC FeII-NO complex was 5-coordinate (73). EPR also provided the first direct evidence that the H3B• radical is formed during the NOS reaction and supports the involvement of the cofactor in the electron transfer mechanism (40).
Future Directions for the Field of No Signaling
Despite significant advances in our understanding of NO as a signaling agent, many questions remain unanswered. Details about the mechanism of NO production by NOS, sGC activation by NO, and downstream effects of cGMP are active and relevant areas of research. In addition to these areas of research, the number of studies on cGMP-independent NO effects has increased significantly (8). Studies have shown that several proteins are modified by NO in vivo and in vitro, and future experiments may reveal how this modification is regulated. Progress in these fields can lead to the development of drugs to treat several maladies ranging from cardiovascular and neurodegenerative diseases to asthma.
Research into the function of NO in bacterial systems also is expanding. NOS-like proteins have been identified in several prokaryotic organisms including Bacillus subtilis and Streptomyces turgidiscabies. These NOSs lack the flavoprotein reductase domain and calmodulin binding motif seen in mammalian NOS but have been shown to generate NO under single turnover conditions (74). A wide range of bacteria also has nitrite reductases, which generate NO as part of denitrifying, as- similatory, and dissimilatory pathways (75). This endogenously produced NO is known to regulate several transcription factors via S-nitrosation (76, 77); however, the existence of heme-based physiologic receptors for NO has yet to be proven. Recently, two classes of potential NO receptors have been identified in bacteria. These classes include globin-like proteins and sGC-like homologues called H-NOX proteins (9, 78). Some microbial globin-like proteins bind O2 and are thought to be involved in the nitrosative stress response (78). Interestingly, the H-NOX proteins from obligate aerobic bacteria bind NO but not O2 (58). The genes that code for these proteins are found in the same operons as those that predicted histidine kinases or diguanylate cyclases. This finding suggests that a functional interaction between the H-NOX and kinase or cyclase may be mediated by NO (9).
References
1. Moncada S, Palmer RM, Higgs EA. Nitric oxide: physiology, pathophysiology, and pharmacology. Pharmacol. Rev. 1991; 43:109-142.
2. Alderton WK, Cooper CE, Knowles RG. Nitric oxide synthases: structure, function and inhibition. Biochem. J. 2001; 357:593-615.
3. Dudzinski DM, Igarashi J, Greif D, Michel T. The regulation and pharmacology of endothelial nitric oxide synthase. Annu. Rev. Pharmacol. Toxicol. 2006; 46:235-276.
4. Stuehr DJ. Structure-function aspects in the nitric oxide synthases. Annu. Rev. Pharmacol. 1997; 37:339-359.
5. Stone JR, Marletta MA. Spectral and kinetic studies on the activation of soluble guanylate cyclase by nitric oxide. Biochemistry 1996; 35:1093-1099.
6. Zhao Y, Brandish PE, Ballou DP, Marletta MA. A molecular basis for nitric oxide sensing by soluble guanylate cyclase. Proc. Natl. Acad. Sci. U. S. A. 1999; 96:14753-14758.
7. Bredt DS. Endogenous nitric oxide synthesis: biological functions and pathophysiology. Free Radical Res. 1999; 31:577-596.
8. Stamler JS, Lamas S, Fang FC. Nitrosylation: the prototypic redox-based signaling mechanism. Cell 2001; 106:675-683.
9. Iyer LM, Anantharaman V, Aravind L. Ancient conserved domains shared by animal soluble guanylyl cyclases and bacterial signaling proteins. BMC Genom. 2003; 4:5.
10. Eich RF, Li TS, Lemon DD, Doherty DH, Curry SR, Aitken JF, Mathews AJ, Johnson KA, Smith RD, Phillips GN, Olson JS. Mechanism of NO-induced oxidation of myoglobin and hemoglobin. Biochemistry 1996; 35:6976-6983.
11. Nguyen T, Brunson D, Crespi CL, Penman BW, Wishnok JS, Tannenbaum SR. DNA damage and mutation in human-cells exposed to nitric-oxide in vitro. Proc. Natl. Acad. Sci. U. S. A. 1992; 89:3030-3034.
12. Wink DA, Kasprzak KS, Maragos CM, Elespuru RK, Misra M, Dunams TM, Cebula TA, Koch WH, Andrews AW, Allen JS, Keefer LK. DNA deaminating ability and genotoxicity of nitric- oxide and its progenitors. Science 1991; 254:1001-1003.
13. Cary SP, Winger JA, Marletta MA. Tonic and acute nitric oxide signaling through soluble guanylate cyclase is mediated by nonheme nitric oxide, ATP, and GTP. Proc. Natl. Acad. Sci. U. S. A. 2005; 102:13064-13069.
14. Ruiz-Stewart I, Tiyyagura SR, Lin JE, Kazerounian S, Pitari GM, Schulz S, Martin E, Murad F, Waldman SA. Guanylyl cyclase is an ATP sensor coupling nitric oxide signaling to cell metabolism. Proc. Natl. Acad. Sci. U. S. A. 2004; 101:37-42.
15. Traut TW. Physiological concentrations of purines and pyrimidines. Mol. Cell. Biochem. 1994; 140:1-22.
16. Munzel T, Feil R, Mulsch A, Lohmann SM, Hofmann F, Walter U. Physiology and pathophysiology of vascular signaling controlled by cyclic guanosine 3’,5’-cyclic monophosphate-dependent protein kinase. Circulation 2003; 108:2172-2183.
17. Sanders KM, Ward SM, Thornbury KD, Dalziel HH, Westfall DP, Carl A. Nitric-oxide as a nonadrenergic, noncholinergic neurotransmitter in the gastrointestinal-tract. Jpn. J. Pharmacol. 1992; 58:220-225.
18. Warner TD, Mitchell JA, Sheng H, Murad F. Effects of cyclic GMP on smooth muscle relaxation. Adv. Pharmacol. 1994; 26:171-194.
19. Rybalkin SD, Yan C, Bornfeldt KE, Beavo JA. Cyclic GMP phosphodiesterases and regulation of smooth muscle function. Circ. Res. 2003; 93:280-291.
20. Morishita T, Tsutsui M, Shimokawa H, Sabanai K, Tasaki H, Suda O, Nakata S, Tanimoto A, Wang KY, Ueta Y, Sasaguri Y, Nakashima Y, Yanagihara N. Nephrogenic diabetes insipidus in mice lacking all nitric oxide synthase isoforms. Proc. Natl. Acad. Sci. U. S. A. 2005; 102:10616-10621.
21. Friebe A, Mergia E, Dangel O, Lange A, Koesling D. Fatal gastrointestinal obstruction and hypertension in mice lacking nitric oxide-sensitive guanylyl cyclase. Proc. Natl. Acad. Sci. U. S. A. 2007; 104:7699-7704.
22. Vermeersch P, Buys E, Pokreisz P, Marsboom G, Ichinose F, Sips P, Pellens M, Gillijns H, Swinnen M, Graveline A, Collen D, Dewerchin M, Brouckaert P, Bloch KD, Janssens S. Soluble guanylate cyclase-alpha1 deficiency selectively inhibits the pulmonary vasodilator response to nitric oxide and increases the pulmonary vascular remodeling response to chronic hypoxia. Circulation 2007; 116:936-943.
23. Fukao M, Mason HS, Britton FC, Kenyon JL, Horowitz B, Keef KD. Cyclic GMP-dependent protein kinase activates cloned BKCa channels expressed in mammalian cells by direct phosphorylation at serine 1072. J. Biol. Chem. 1999; 274:10927-10935.
24. Schlossmann J, Ammendola A, Ashman K, Zong XG, Huber A, Neubauer G, Wang GX, Allescher HD, Korth M, Wilm M, Hofmann F, Ruth P. Regulation of intracellular calcium by a signalling complex of IRAG, IP3 receptor and cGMP kinase I beta. Nature 2000; 404:197-201.
25. Hofmann F. The biology of cyclic GMP-dependent protein kinases. J. Biol. Chem. 2005; 280:1-4.
26. Sauzeau V, Le Jeune H, Cario-Toumaniantz C, Smolenski A, Lohmann SM, Bertoglio J, Chardin P, Pacaud P, Loirand G. Cyclic GMP-dependent protein kinase signaling pathway inhibits RhoA-induced Ca2+ sensitization of contraction in vascular smooth muscle. J. Biol. Chem. 2000; 275:21722-21729.
27. Reinhard M, Jarchau T, Walter U. Actin-based motility: stop and go with Ena/VASP proteins. Trends Biochem. Sci. 2001; 26:243-249.
28. Hobbs AJ, Moncada S. Antiplatelet properties of a novel, non-NObased soluble guanylate cyclase activator, BAY 41-2272. Vasc. Pharmacol. 2003; 40:149-154.
29. Ko FN, Wu CC, Kuo SC, Lee FY, Teng CM. YC-1, a novel activator of platelet guanylate cyclase. Blood 1994; 84:4226-4233.
30. Stasch JP, Alonso-Alija C, Apeler H, Dembowsky K, Feurer A, Minuth T, Perzborn E, Schramm M, Straub A. Pharmacological actions of a novel NO-independent guanylyl cyclase stimulator, BAY 41-8543: in vitro studies. Brit. J. Pharmacol. 2002; 135:333-343.
31. Matthews JR, Botting CH, Panico M, Morris HR, Hay RT. Inhibition of NF-kappa B DNA binding by nitric oxide. Nucleic Acids Res. 1996; 24:2236-2242.
32. McMahon TJ, Stamler JS. Concerted nitric oxide oxygen delivery by hemoglobin. Method Enzymol. 1999; 301:99-114.
33. Rossig L, Fichtlscherer B, Breitschopf K, Haendeler J, Zeiher AM, Mulsch A, Dimmeler S. Nitric oxide inhibits caspase-3 by S-nitrosation in vivo. J. Biol. Chem. 1999; 274:6823-6826.
34. Mitchell DA, Marletta MA. Thioredoxin catalyzes the S-nitrosation of the caspase-3 active site cysteine. Nat. Chem. Biol. 2005; 1:154-158.
35. Gladwin MT, Lancaster JR, Freeman BA, Schechter AN. Nitric oxide’s reactions with hemoglobin: a view through the SNO-storm. Nat. Med. 2003; 9:496-500.
36. Erwin PA, Mitchell DA, Sartoretto J, Marletta MA, Michel T. Subcellular targeting and differential S-nitrosylation of endothelial nitric-oxide synthase. J. Biol. Chem. 2006; 281:151-157.
37. Mitchell DA, Erwin PA, Michel T, Marletta MA. S-Nitrosation and regulation of inducible nitric oxide synthase. Biochemistry 2005; 44:4636-4647.
38. Sayed N, Baskaran P, Ma X, van den Akker F, Beuve A. Desensitization of soluble guanylyl cyclase, the NO receptor, by S-nitrosylation. Proc. Natl. Acad. Sci. U. S. A. 2007; 104:12312-12317.
39. Ghosh DK, Abusoud HM, Stuehr DJ. Reconstitution of the 2nd step in NO synthesis using the isolated oxygenase and reductase domains of macrophage NO synthase. Biochemistry 1995; 34:11316-11320.
40. Hurshman AR, Krebs C, Edmondson DE, Huynh BH, Marletta MA. Formation of a pterin radical in the reaction of the heme domain of inducible nitric oxide synthase with oxygen. Biochemistry 1999; 38:15689-15696.
41. Raman CS, Li H, Martasek P, Kral V, Masters BS, Poulos TL. Crystal structure of constitutive endothelial nitric oxide synthase: a paradigm for pterin function involving a novel metal center. Cell 1998; 95:939-950.
42. Crane BR, Arvai AS, Ghosh DK, Wu C, Getzoff ED, Stuehr DJ, Tainer JA. Structure of nitric oxide synthase oxygenase dimer with pterin and substrate. Science 1998; 279:2121-2126.
43. Li H, Raman CS, Glaser CB, Blasko E, Young TA, Parkinson JF, Whitlow M, Poulos TL. Crystal structures of zinc-free and -bound heme domain of human inducible nitric-oxide synthase. Implications for dimer stability and comparison with endothelial nitric-oxide synthase. J. Biol. Chem. 1999; 274:21276-21284.
44. Wink DA, Mitchell JB. Chemical biology of nitric oxide: Insights into regulatory, cytotoxic, and cytoprotective mechanisms of nitric oxide. Free Radical Biol. Med. 1998; 25:434-456.
45. Antonini E, Brunori M. Hemoglobin and myoglobin in their reactions with ligands. 1971. North-Holland, Amsterdam, the Netherlands.
46. Hoshino M, Ozawa K, Seki H, Ford PC. Photochemistry of nitric-oxide adducts of water-soluble iron(III) porphyrin and ferrihemoproteins studied by nanosecond laser photolysis. J. Am. Chem. Soc. 1993; 115:9568-9575.
47. Hoshino M, Maeda M, Konishi R, Seki H, Ford PC. Studies on the reaction mechanism for reductive nitrosylation of ferrihemopro- teins in buffer solutions. J. Am. Chem. Soc. 1996; 118:5702-5707.
48. Weichsel A, Maes EM, Andersen JF, Valenzuela JG, Shokhireva T, Walker FA, Montfort WR. Heme-assisted S-nitrosation of a proximal thiolate in a nitric oxide transport protein. Proc. Natl. Acad. Sci. U. S. A. 2005; 102:594-599.
49. Luchsinger BP, Rich EN, Gow AJ, Williams EM, Stamler JS, Singel DJ. Routes to S-nitroso-hemoglobin formation with heme redox and preferential reactivity in the beta subunits. Proc. Natl. Acad. Sci. U. S. A. 2003; 100:461-466.
50. Yuen PS, Potter LR, Garbers DL. A new form of guanylyl cyclase is preferentially expressed in rat kidney. Biochemistry 1990; 29:10872-10878.
51. Harteneck C, Wedel B, Koesling D, Malkewitz J, Bohme E, Schultz G. Molecular cloning and expression of a new alpha- subunit of soluble guanylyl cyclase. Interchangeability of the alpha-subunits of the enzyme. FEBS Lett. 1991; 292:217-222.
52. Cary SP, Winger JA, Derbyshire ER, Marletta MA. Nitric oxide signaling: no longer simply on or off. Trends Biochem. Sci. 2006; 31:231-239.
53. Ma X, Sayed N, Beuve A, van den Akker F. NO and CO differentially activate soluble guanylyl cyclase via a heme pivot-bend mechanism. EMBO J. 2007; 26:578-588.
54. Nioche P, Berka V, Vipond J, Minton N, Tsai AL, Raman CS. Femtomolar sensitivity of a NO sensor from Clostridium botulinum. Science 2004; 306:1550-1553.
55. Pellicena P, Karow DS, Boon EM, Marletta MA, Kuriyan J. Crystal structure of an oxygen-binding heme domain related to soluble guanylate cyclases. Proc. Natl. Acad. Sci. U. S. A. 2004; 101:12854-12859.
56. Russwurm M, Koesling D. NO activation of guanylyl cyclase. EMBO J. 2004; 23:4443-4450.
57. Boon EM, Marietta MA. Ligand specificity of H-NOX domains: from sGC to bacterial NO sensors. J. Inorg. Biochem. 2005; 99:892-902.
58. Boon EM, Huang SH, Marietta MA. A molecular basis for NO selectivity in soluble guanylate cyclase. Nat. Chem. Biol. 2005; 1:53-59.
59. Martin E, Berka V, Bogatenkova E, Murad F, Tsai AL. Ligand selectivity of soluble guanylyl cyclase - Effect of the hydrogen-bonding tyrosine in the distal heme pocket on binding of oxygen, nitric oxide, and carbon monoxide. J. Biol. Chem. 2006; 281:27836-27845.
60. Rothkegel C, Schmidt PM, Stoll F, Schroder H, Schmidt HHHW, Stasch JP. Identification of residues crucially involved in soluble guanylate cyclase activation. FEBS Lett. 2006; 580:4205-4213.
61. Marletta MA, Yoon PS, Iyengar R, Leaf CD, Wishnok JS. Macrophage oxidation of L-arginine to nitrite and nitrate:nitric oxide is an intermediate. Biochemistry 1988; 27:8706-8711.
62. Hibbs JB Jr, Taintor RR, Vavrin Z. Macrophage cytotoxicity: role for L-arginine deiminase and imino nitrogen oxidation to nitrite. Science 1987; 235:473-476.
63. Iyengar R, Stuehr DJ, Marletta MA. Macrophage synthesis of nitrite, nitrate, and N-nitrosamines: precursors and role of the respiratory burst. Proc. Natl. Acad. Sci. U. S. A. 1987; 84:6369-6373.
64. Bredt DS, Snyder SH. Isolation of nitric-oxide synthetase, a calmodulin-requiring enzyme. Proc. Natl. Acad. Sci. U. S. A. 1990; 87:682-685.
65. Dolphin D. The Porphyrins. 1978. Academic Press, New York.
66. Stone JR, Marletta MA. Soluble guanylate-cyclase from bovine lung - Activation with nitric-oxide and carbon-monoxide and spectral characterization of the ferrous and ferric states. Biochemistry 1994; 33:5636-5640.
67. Deinum G, Stone JR, Babcock GT, Marletta MA. Binding of nitric oxide and carbon monoxide to soluble guanylate cyclase as observed with resonance Raman spectroscopy. Biochemistry 1996; 35:1540-1547.
68. Wang JL, Rousseau DL, Abusoud HM, Stuehr DJ. Heme coordination of NO in NO synthase. P. Natl. Acad. Sci. 1994; 91:10512-10516.
69. Wang JL, Stuehr DJ, Rousseau DL. Interactions between substrate analogues and heme ligands in nitric oxide synthase. Biochemistry 1997; 36:4595-4606.
70. Pal B, Kitagawa T. Interactions of soluble guanylate cyclase with diatomics as probed by resonance Raman spectroscopy. Journal of Inorganic Biochemistry 2005; 99:267-279.
71. Tomita T, Ogura T, Tsuyama S, Imai Y, Kitagawa T. Effects of GTP on bound nitric oxide of soluble guanylate cyclase probed by resonance Raman spectroscopy. Biochemistry 1997; 36:10155-10160.
72. Ubbink M, Worrall JA, Canters GW, Groenen EJ, Huber M. Paramagnetic resonance of biological metal centers. Annu. Rev. Bioph. Biom. 2002; 31:393-422.
73. Stone JR, Sands RH, Dunham WR, Marletta MA. Electron paramagnetic resonance spectral evidence for the formation of a pentacoordinate nitrosyl-heme complex on soluble guanylate cyclase. Biochem. Bioph. Res. Co. 1995; 207:572-577.
74. Adak S, Aulak KS, Stuehr DJ. Direct evidence for nitric oxide production by a nitric-oxide synthase-like protein from Bacillus subtilis. J. Biol. Chem. 2002; 277:16167-16171.
75. Brittain T, Blackmore R, Greenwood C, Thomson AJ. Bacterial nitrite-reducing enzymes. Eur. J. Biochem. 1992; 209:793-802.
76. Hausladen A, Privalle CT, Keng T, DeAngelo J, Stamler JS. Nitrosative stress: activation of the transcription factor OxyR. Cell 1996; 86:719-729.
77. Ding H, Demple B. Direct nitric oxide signal transduction via nitrosylation of iron-sulfur centers in the SoxR transcription activator. P. Natl. Acad. Sci. 2000; 97:5146-5150.
78. Poole RK, Hughes MN. New functions for the ancient globin family: bacterial responses to nitric oxide and nitrosative stress. Mol. Microbiol. 2000; 36:775-783.
See Also
Enzyme Kinetics
Hemes in Biology
Post-Translational Modifications to Regulate Protein Function
Signal Cascades, Protein Interaction Networks in
Spectroscopic Techniques: Proteins