Interaction of Nitric Oxide with Human Heme Oxygenase-1*

Jinling WangDagger , Shen Lu§, Pierre Moënne-Loccoz§, and Paul R. Ortiz de MontellanoDagger

From the Dagger  Department of Pharmaceutical Chemistry, University of California, San Francisco, California 94143-0446 and the § Department of Biochemistry and Molecular Biology, Oregon Graduate Institute School of Science and Engineering at Oregon Health Sciences University, Beaverton, Oregon 97006-8921

Received for publication, October 30, 2002, and in revised form, November 13, 2002

    ABSTRACT
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

NO and CO may complement each other as signaling molecules in some physiological situations. We have examined the binding of NO to human heme oxygenase-1 (hHO-1), an enzyme that oxidizes heme to biliverdin, CO, and free iron, to determine whether inhibition of hHO-1 by NO can contribute to the signaling interplay of NO and CO. An Fe3+-NO hHO-1-heme complex is formed with NO or the NO donors NOC9 or 2-(N,N-diethylamino)-diazenolate-2-oxide·sodium salt. Resonance Raman spectroscopy shows that ferric hHO-1-heme forms a 6-coordinated, low spin complex with NO. The nu (N-O) vibration of this complex detected by Fourier transform IR is only 4 cm-1 lower than that of the corresponding metmyoglobin (met-Mb) complex but is broader, suggesting a greater degree of ligand conformational freedom. The Fe3+-NO complex of hHO-1 is much more stable than that of met-Mb. Stopped-flow studies indicate that kon for formation of the hHO-1-heme Fe3+-NO complex is ~50-times faster, and koff 10 times slower, than for met-Mb, resulting in Kd = 1.4 µM for NO. NO thus binds 500-fold more tightly to ferric hHO-1-heme than to met-Mb. The hHO-1 mutations E29A, G139A, D140A, S142A, G143A, G143F, and K179A/R183A do not significantly diminish the tight binding of NO, indicating that NO binding is not highly sensitive to mutations of residues that normally stabilize the distal water ligand. As expected from the Kd value, the enzyme is reversibly inhibited upon exposure to pathologically, and possibly physiologically, relevant concentrations of NO. Inhibition of hHO-1 by NO may contribute to the pleiotropic responses to NO and CO.

    INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

Nitric oxide (NO)1 functions as a signaling molecule in a diversity of physiological responses, including vasodilation and regulation of normal vascular tone, neuronal signal transmission, cytotoxicity against pathogens and tumors, and regulation of cellular respiration (1-3). Most of these responses result from interaction of NO with the heme group of the receptor guanylyl cyclase (1). A role akin to that of NO in signaling pathways has also been postulated for CO (4). CO is produced in mammals from heme by two heme oxygenases, HO-1 and HO-2 (5-8). The involvement of CO has been invoked as a factor in atherosclerosis (9), psoriasis (10), vascular constriction (11), chronic renal inflammation (12), cellular protection (13), hyperoxia-induced lung injury (14), and other physiological situations. The role of CO as an NO-like signaling molecule has received strong support from studies of heme oxygenase and nitric-oxide synthase knockouts (15), but much of the evidence, particularly that which depends heavily on inhibition of heme oxygenase by metalloporphyrins such as tin protoporphyrin IX, is tainted by ambiguities concerning the specificity of the inhibitors (16). Nevertheless, the collective evidence makes a persuasive case for at least a limited role for CO in mammalian signaling systems.

Evidence has accumulated that interactions of CO and NO may influence the physiological responses to each of these agents through interactions at the level of the biosynthetic enzymes. Thus, NO has been shown to elevate the levels of heme oxygenase-1 mRNA and protein (17-25), and this response appears to be mediated by a guanylate cyclase-independent mechanism that may subserve a more generalized antioxidant response (18). Conversely, CO has been reported to elevate the steady state level of NO (26), but increased levels of heme oxygenase have been shown to decrease NO concentrations, possibly by consuming the heme required for assembly of the nitric-oxide synthases (27-29).

As both NO and CO are small molecules that can coordinate to the iron in heme proteins, it is possible that at physiological concentrations NO may directly inhibit the heme oxygenases, and conversely, CO may inhibit the nitric-oxide synthases. In studies of the identity of the proximal ligand in HO-1, NO has been shown by resonance Raman, and EPR to bind to the ferrous heme iron atom (30, 31). In HO-2, a heme regulatory motif binds a secondary non-catalytic heme that binds NO and, through an undetermined mechanism, inactivates the protein (32). Indirect evidence also exists for the inhibition of heme oxygenase in tissue homogenates by endogenously formed NO (33), but no focused study has been carried out of the inhibition of HO-1 by NO.

    EXPERIMENTAL PROCEDURES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

Materials-- NADPH, ampicillin, heme, bovine serum album, glucose, horse myoglobin, MAHMA NONOate (NOC9), and NO gas (98.5%) were purchased from Sigma. High purity argon (99.98%) was from Matheson (Newark, CA). 2-(N,N-Diethylamino)-diazenolate-2-oxide·sodium salt (Dea/NO) was obtained from Alexis Corp. (San Diego, CA). All chemicals were used without further purification. The NO donors have the following half-lives in 0.1 M potassium phosphate buffer at pH 7.4: NOC9, 3 min at 22 °C and 1-2 min at 37 °C; Dea/NO 16 min at 22 °C and 2-4 min at 37 °C.

Enzymes-- Catalase and glucose oxidase were from Sigma. Rat biliverdin reductase and rat cytochrome P450 reductase were purified by published procedures (34-36). The hHO-1 construct used encoded human heme oxygenase-1 lacking the 23 C-terminal amino acids (37). Oligonucleotide synthesis was carried out by Invitrogen, through the Cell Culture Facility of the University of California, San Francisco. Mutants of hHO-1 were generated with a Quick-change site-directed mutagenesis kit (Stratagene). Plasmid purifications and bacterial transformations were performed by standard procedures (38). Transformants were screened initially by ampicillin resistance and confirmed by sequence analysis. Wild type hHO-1 and its E29A, G139A, S142A, D140A, G143A, G143F, and K179A/R183A mutants were expressed, purified, and reconstituted with heme as reported previously (36, 39, 40). All experiments using these proteins were carried out in 0.1 M potassium phosphate buffer at pH 7.4 (standard buffer), unless otherwise stated.

NO Treatments-- The concentrated NO donor solution stocks were prepared fresh in 0.01 N NaOH. NOC9 and Dea/NO are stable under alkaline conditions, but they decompose spontaneously when small amounts of the stock solutions are added to the standard pH 7.4 buffer. Concentrated stock solutions of the NO donors were used to minimize the changes in pH. NOC9 treatment was carried out at room temperature (~25 °C) and that with Dea/NO at 37 °C. These temperatures were chosen so that the two NO donors had similar rates of NO release. All treatments with NO donors were performed aerobically. The NO gas solutions used in stopped-flow experiments were made anaerobic by bubbling NO into argon-equilibrated standard buffer.

The concentration of the NO donors in the enzyme binding assays was varied at a fixed enzyme concentration of 4 µM. NO binding was monitored at wavelengths between 250 and 700 nm. Formation of the hHO-1-heme Fe3+-NO complex was determined from the ratio of the 416 to 404 nm absorbance. Formation of the corresponding met-Mb Fe3+-NO complex was estimated from the ratio of the absorbance at 417 to 408 nm.

Spectroscopic Characterization-- The UV-visible spectra of the hHO-1 proteins were recorded in standard buffer on a Cary Varian model 1E spectrophotometer.

The enzyme concentration for RR experiments was ~125 µM in 0.1 M potassium phosphate buffer at pH 7.4. 14NO and 15NO gas purchased from Aldrich was bubbled through a 0.1 M KOH solution to remove higher nitrogen oxides. Formation of the NO adduct was achieved by addition of an NO-saturated buffer solution to an argon-purged hHO-1 solution in the Raman capillary cell to reach a final concentration of ~1 mM NO. Before freezing the samples, the completion of the reaction was confirmed by UV-visible spectroscopy in the same Raman capillary cell using a Cary 50 spectrophotometer. Once the RR experiments were completed, the sample was thawed to obtain its UV-visible spectrum and confirm the stability of the complex during the laser illumination at 90 K.

RR spectra were obtained on a custom McPherson 2061/207 spectrograph (set at 0.67 m with variable gratings) equipped with a Princeton Instruments liquid N2-cooled CCD detector (LN-1100PB). Kaiser Optical supernotch filters were used to attenuate Rayleigh scattering. Excitation sources consisted of an Innova 302 krypton laser (413 nm). Spectra were collected on frozen samples kept at ~90 K with N2 cold finger in a backscattering geometry (41). Frequencies were calibrated relative to indene and CCl4 standards and are accurate to ±1 cm-1. CCl4 was also used to check the polarization conditions.

In the FTIR experiments, the enzyme concentration was increased to ~1 mM using a Microcon 10 ultrafiltration device (Amicon). The concentrated hHO-1 solution was made anaerobic in a vial before exchanging the head space with pure NO gas to reach a final concentration of ~2 mM NO in solution. The protein sample was then injected into an IR cell consisting of CaF2 windows separated by a 50-µm Teflon spacer. The formation of the NO adduct was confirmed by UV-visible spectroscopy in the IR cell using a Cary 50 spectrophotometer.

FTIR spectra were obtained at room temperature on a PerkinElmer Life Sciences system 2000 equipped with a liquid N2-cooled MCT detector. Sets of 20-min accumulations were acquired at a 2-cm-1 resolution on the samples and the identical cell filled with buffer for background subtraction.

Stopped-flow Kinetic Analyses-- Kinetic studies were carried out with an SF-61 DX2 double mixing stopped-flow system (Hi-Tech Scientific). The stopped-flow instrument was made anaerobic by first rinsing with argon equilibrated standard buffer, followed by an overnight incubation with anaerobically prepared catalase/glucose/glucose oxidase solution. Kinetic traces were taken at room temperature at wavelengths between 380 and 700 nm. The data were analyzed with KinetAsyst2 software and were fit to a first-exponential expression. In this experiment, 3 µM ferric hHO-1-heme and 1.5 µM horse met-Mb were used. The solubility of NO gas under 1 atmosphere at 20 °C is ~2 mM. Dilution of the NO-saturated solution was made with argon-equilibrated standard buffer in gas-tight syringes (Hamilton).

Bilirubin Activity Assay-- The reaction mixture contained ferric hHO-1-heme (1 µM), hemin (30 µM), bilirubin reductase (4 µM), and cytochrome P450 reductase (0.4 µM) in the standard buffer. The reaction was initiated by the addition of NADPH (400 µM). The production of bilirubin at room temperature was monitored at 468 nm for 100 s. The initial rate of the reaction was calculated using the value epsilon 468 = 43.5 mM-1 cm-1 for the bilirubin product. To study the effects of NO on hHO-1 activity, two sets of experiments were carried out. In one set, biliverdin reductase, cytochrome P450 reductase, hemin, and hHO-1 were incubated with various concentrations of NOC9 for different times before NADPH was added to measure the bilirubin forming activity. In the other set, hHO-1 was incubated with 1 mM NO donor and then the rest of the assay mixture was added for the activity assay. Separately, hemin (30 µM) and bilirubin reductase (4 µM)-cytochrome P450 reductase (0.4 µM) were separately incubated with the NO donors before the other components of the reaction system were added, and bilirubin formation was quantitated.

    RESULTS
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

Absorption Spectrum of the hHO-1-Heme Fe3+-NO Complex-- Ferric hHO-1-heme has a Soret maximum at 405 ± 1 nm in 0.1 M phosphate buffer at pH 7.4. Upon addition of a 1 mM concentration of the NO donor NOC9 or Dea/NO, the ferric hHO-1-heme Soret maximum immediately shifted to 416 nm (Fig. 1). This spectroscopic shift was reversible, suggesting that the NO species generated by the NO donors interacted with the heme in hHO-1. Ferric heme alone has a very broad Soret absorption at about 380 nm, and its incubation with the NO donors caused a sharpening of the Soret band with a decrease in its absorption intensity (not shown). In contrast to the reaction with ferric hHO-1-heme, the spectral change observed with free hemin was not reversible.


View larger version (25K):
[in this window]
[in a new window]
 
Fig. 1.   Spectral change of ferric hHO-1-heme (4.6 µM) in the presence of either 1.0 mM NOC9 or Dea/NO. The spectra were recorded in the standard buffer. Arrows indicate the direction of spectral change over time. The Soret band at time 0 was taken immediately after adding the NO donor and is already that of the NO complex.

RR and FTIR Spectroscopy-- The ferric hHO-1-heme NO complex prepared by bubbling NO through a solution of the ferric hHO-1-heme complex was observed to be very photolabile in the resonance Raman experiments, but at 90 K the efficiency of this process was sufficiently diminished to allow the experiments to be carried out. As shown previously (31), the spectrum of ferric hHO-1 reveals a hexacoordinated high spin/hexacoordinated low spin equilibrium with nu 3, nu 2, and nu 10 modes at 1482/1508, 1566/1585, and 1608/1640 cm-1, respectively (Fig. 2A). After exposure to NO, the porphyrin skeletal modes are observed at higher frequencies with nu 4, nu 3, nu 2, and nu 10 at 1378, 1511, 1588, and 1645 cm-1, respectively (Fig. 2B). These frequencies identify the NO adduct as a hexacoordinated low spin complex, although a minor hexacoordinated high spin population revealed by the weak nu 3 at 1483 cm-1 is assigned to some photodissociation.


View larger version (24K):
[in this window]
[in a new window]
 
Fig. 2.   High frequency region of the resonance Raman spectra of the ferric (A) and ferric-nitrosyl (B) hHO-1-heme complex. Spectra were obtained at 90 K using 413-nm excitation.

In heme ferric nitrosyl complexes, the nu (N-O) is not resonance-enhanced with Soret excitation, but it is easily observed in the FTIR spectrum. This vibration is detected at 1918 cm-1 in ferric hHO-1-heme, only 4 cm-1 lower than that in met-Mb (Fig. 3). Such nu (N-O) frequencies are characteristic of linear six-coordinated {Fe(NO)}6 complexes (42). A significant difference between these two signals resides in the ~20-cm-1 half-width of this stretching mode in hHO-1 compared with the 9-cm-1 half-width observed in met-Mb. In met-Mb the configuration of the nitrosyl group is clearly defined by the presence of the imidazole ring from the distal histidine, but the absence of distal polar side chains above the heme iron of hHO-1 and a greater solvent accessibility may permit greater fluctuation of the NO ligand, resulting in substantial inhomogeneous broadening of the nu (N-O).


View larger version (19K):
[in this window]
[in a new window]
 
Fig. 3.   High frequency region of the FTIR spectra of the ferric nitrosyl complex in met-Mb and hHO-1-heme.

In the low frequency region of the RR spectra (Fig. 4), the identification of the Fe-N-O vibrational modes was facilitated by the use of isotopic labeling and the close similarity of these frequencies with those observed in other hemoproteins (43). In the met-Mb ferric-nitrosyl complex, the nu (Fe-NO) and delta (Fe-N-O) are observed at 595 (-6) and 573 (-11) cm-1, respectively. In the ferric nitrosyl complex of hHO-1, two bands at 596 and 588 cm-1 that shift to 590 and 573 cm-1 with 15NO are assigned to the nu (Fe-NO) and the delta (Fe-N-O), respectively.


View larger version (25K):
[in this window]
[in a new window]
 
Fig. 4.   Low frequency RR spectra of the ferric hHO-1-heme nitrosyl complex.

Kinetic Analyses of NO Binding to Horse Met-Mb and Ferric hHO-1-Heme-- To study further the formation of the hHO-1-heme (Fe3+-NO) complex, stopped-flow experiments were employed to determine the kon, koff, and Kd values for the binding of NO. The experiments in this instance were performed anaerobically with NO gas instead of NO donors. Our experimental conditions in the case of met-Mb gave kon, koff, and Kd values similar to those in the literature (49), validating the methodology that was employed (Table I). The kon for formation of the hHO-1 Fe3+-NO complex was found to be ~50-fold faster, and koff 10-fold slower, than for horse met-Mb, resulting in an ~500-fold tighter binding of NO. The resulting calculated Kd value for the binding of NO to ferric hHO-1-heme is 1.4 µM.

                              
View this table:
[in this window]
[in a new window]
 
Table I
Kon, Koff, and Kd for formation of the ferric hHO-1-heme and horse met-Mb Fe3+-NO complexes
The incubation conditions in Ref. 49 were 0.05 M phosphate buffer, pH 7.4, 25 °C, and in this study 0.1 M potassium phosphate buffer, pH 7.4, 25 °C.

NO Binding to Mutant Ferric hHO-1-Heme Complexes-- As shown in Fig. 5, the binding of NO to ferric hHO-1-heme can also be seen with the NO donors NOC9 and Dea/NO. The concentrations of NOC9 and Dea/NO that cause half-maximal binding of NO to ferric hHO-1-heme are ~80 and 100 µM, respectively, although the actual concentration of NO in these experiments is much lower. Under the same conditions the binding of NO to horse met-Mb did not reach a plateau even at much higher concentrations of the NO donors, as expected from the higher Kd value for this protein (Table I and Fig. 5).


View larger version (27K):
[in this window]
[in a new window]
 
Fig. 5.   Binding of NO to 4 µM ferric hHO-1-heme and 4 µM horse met-Mb. The binding assay was performed in the standard buffer. The data points are the averages of two independent determinations, and the bars indicate the positions of the two averaged values.

The high affinity of the ferric hHO-1-heme complex for NO is unusual for a ferric hemoprotein, although the binding of NO to ferric catalase reportedly occurs with a comparably low Kd of ~0.5 µM (45, 46). In our efforts to identify specific residues that contribute to this high binding affinity, we investigated the binding of NO to hHO-1 in which individual active site amino acids had been mutated. hHO-1 residues that could stabilize distal iron ligands by hydrogen bonding or other interactions include Glu-29, Gly-139, Asp-140, Ser-142, Gly-143, Lys-179, and Arg-183. Glu-29 is close enough to form a hydrogen bond with His-25, the proximal iron ligand (47). Gly-139, Asp-140, and Gly-143 are part of a hydrogen bonding network on the distal side that interacts with distal ligands, and mutation of any one of them to an alanine results in dissociation of the distal water ligand (36, 39). Mutation of Ser-142 shift the pKa value of the distal water ligand toward more basic values.2 Lys-179 and Arg-183 appear to interact with the propionate carboxyl groups of the heme and may be important for proper orientation of the heme (47, 48). We have therefore examined the binding of NO to the E29A, G139A, D140A, S142A, G143A, G143F, and K179A/R183A mutants, all of which were heterologously expressed in Escherichia coli, purified, and found to have appropriate Soret maxima (Table II). The values for half-saturation of NO binding using NOC9 as the NO donor show that none of the mutations altered the NO binding affinity by more than a factor of ~2 (Table II). Thus, the high NO affinity of ferric hHO-1-heme is not very sensitive to the identities of these active site residues despite the fact that some of them interact strongly with the distal water ligand that is replaced by NO. Indeed, the high affinity for NO appears to be insensitive to the presence or absence of the distal water ligand, as the water ligand is absent in at least the G139A, G143A, and D140A mutants (36, 39).

                              
View this table:
[in this window]
[in a new window]
 
Table II
IC50 for NO binding to wild-type and mutant ferric hHO-1-heme with NOC9 as the NO donor

Inhibition of Bilirubin Formation-- Incubation of a system consisting of ferric hHO-1-heme, biliverdin reductase, cytochrome P450 reductase, and hemin with increasing concentrations of NOC9 for various times, followed by addition of NADPH to assay bilirubin formation, clearly demonstrated that the protein is reversibly inhibited by NO (Fig. 6). Little inhibition was observed with a 1 µM concentration of the NO donor at any time point, but inhibition was observed when the donor concentration was raised to 10 µM. With a 50 µM concentration of NOC9, bilirubin formation was completely suppressed when the assay was carried out without NOC9 preincubation, but activity was recovered when the assay was carried out after 10 min or longer of preincubation. Similar results were obtained with 100-600 µM NOC9, except that inhibition was observed after even longer preincubation periods (Fig. 6). A 100 µM concentration of NOC9 is required to half-saturate the active site of ferric hHO-1-heme with NO, and this concentration therefore must roughly correspond to the Kd of 1.4 µM for NO itself. The fact that complete inhibition is observed with half of this NOC9 concentration, and some inhibition even at lower concentrations, suggests that inhibition may also reflect some binding of NO to the ferrous intermediate obtained when the enzyme complex is reduced by cytochrome P450 reductase. Furthermore, the inhibition, like formation of the spectroscopically determined Fe3+-NO complex, is reversible. Inhibition is therefore lost with time as the NO donor is exhausted and the NO concentration decays. This is most clearly seen in Fig. 7, in which the recovery of bilirubin forming activity has been measured as a function of the time a fixed concentration of the NO donor is preincubated with the enzyme prior to carrying out the activity assay. In this case, two different NO donors were employed, NOC9 and Dea/NO. As the figure shows, the activity of the enzyme is completely inhibited when there is no preincubation and for NOC9 even after a 20-min preincubation, but the activity with both NO donors recovers as the preincubation time is prolonged. However, only ~80% of the activity was recovered in these experiments. To determine whether stable NO donor decomposition products inhibit hHO-1 activity, ferric hHO-1 was incubated with solutions of decomposed NO donors for 40 min, and the bilirubin forming activity was then assayed. As shown in Fig. 8, the products of decomposition of the NO donors have some hHO-1 inhibitory activity. Only 80% of the control activity was observed in the presence of the decomposition products from a 1.0 mM concentration of the NO donors, readily explaining the recovery of only 80% of the enzyme activity in the incubations with the NO donors (Fig. 7). Control experiments in which NOC9 was added to an incubation only containing biliverdin reductase and P450 reductase, followed by addition of hHO-1, heme, and NADPH resulted in negligible inhibition, confirming that hHO-1-heme is the site of inhibition. High concentrations of NOC9 added to heme alone prior to addition of the other components of the catalytic system also inhibited turnover (results not shown), in accord with the finding that the spectrum of heme in solution was altered by the NO donors.


View larger version (43K):
[in this window]
[in a new window]
 
Fig. 6.   Effects of NO donors on the hHO-1 bilirubin forming activity. The reaction system consisting of hHO-1, ferric heme, biliverdin reductase, and cytochrome P450 reductase was preincubated with increasing concentrations of NOC9 for the indicated times, after which NADPH was added, and the formation of bilirubin was measured. The preincubation and activity assays were both carried out at room temperature. ND indicates no bilirubin formation was detected. The values shown are the mean of duplicate determinations. The bars indicate the range of the two values.


View larger version (49K):
[in this window]
[in a new window]
 
Fig. 7.   Recovery of hHO-1 bilirubin forming activity over time. A ferric hHO-1-heme (1 µM) solution was preincubated with 1.0 mM NOC9 or Dea/NO after which the bilirubin forming activity of the enzyme was determined by adding the required additional reaction components.


View larger version (45K):
[in this window]
[in a new window]
 
Fig. 8.   Effects of decomposed NO donors on the bilirubin activity of hHO-1. NO donors (1 mM) were allowed to decomposed at 37 °C for ~4 h, after which 1 µM ferric hHO-1-heme was incubated with the decomposed NO donor solutions for 40 min prior to adding the necessary additional components and assaying bilirubin formation.


    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

NO, unlike O2 and CO, can bind to the iron atom of hemoproteins in both the ferric and ferrous states, although the binding affinity is generally much higher for the ferrous state (49, 50). For example, at pH 7.4 and 25 °C, the Kd values for the binding of NO to ferrous deoxymyoglobin and ferric met-Mb are 7 × 10-6 and 905 µM, respectively, a difference of roughly 108 (49, 51). The binding of NO to ferric hemoproteins, however, occurs over a considerable range, the tightest binding reported being Kd = 0.5 µM for catalase (45, 46). The binding of NO to the ferric hHO-1-heme complex with Kd = 1.4 µM thus approaches the tightest binding so far observed for any ferric hemoprotein. Detailed analysis shows that this high affinity, when compared with the ~500-fold lower affinity for met-Mb, is due to a 50 times faster kon and a 10 times slower koff for NO (Table I).

The basis for this difference in binding affinity is unclear. The overall vibrational characterization of these {Fe(NO)}6 structures in hHO-1 and met-Mb demonstrates that these complexes share the same bonding geometry and strength. Resonance Raman studies show that the hHO-1-heme Fe3+-NO complex is 6-coordinated low spin (Fig. 2), as is the met-Mb complex (30). Furthermore, FTIR shows that the nu (N-O) band of the hHO-1 Fe3+-NO complex is at 1918 cm-1, a value only 4 cm-1 lower than for the met-Mb complex (Fig. 3) (52). This suggests that coordination of the NO to the heme iron is similar in both hemoproteins, although the broader bandwidth observed with the hHO-1 complex suggests that the NO has greater mobility in that protein. Generally, as is the case for the binding of NO to ferric hemoproteins, these results strongly argue that the NO is bound perpendicular to the heme face rather than at an angle (53-56), in contrast to its orientation when bound to ferrous hemoproteins. Furthermore, in both met-Mb and ferric hHO-1-heme, the iron in the absence of NO is coordinated to a water molecule. The pKa values for deprotonation of these iron-coordinated water molecules are similar in both proteins (30, 57), again indicating similar coordination states and environment and suggesting that the energy cost of displacing the water ligand should be similar in both proteins.

In view of the similarities in coordination properties, the differences in the Kd for binding of NO to ferric hNO-1-heme and met-Mb presumably stem from other differences in the active sites of the two proteins. The ~50-fold increase in kon rate for NO in hHO-1 compared with met-Mb may reflect a higher steric hindrance in the distal pocket of met-Mb. In both these ferric proteins the sixth iron coordination site is occupied by a water molecule that must be displaced for NO to bind, but displacement of the water molecule in hHO-1 may require relatively little protein side chain rearrangement relative to that which occurs in met-Mb (58). In an attempt to identify residues that might contribute to the unique binding affinity of NO for ferric hHO-1-heme, we have mutated seven residues that could contribute to this affinity. However, in no instance did the mutation cause more than a 2-3-fold change in the observed Kd value. Mutations of Asp-140 and Gly-143 slightly decreased the affinity; those of Gly-139 slightly increased it; and those of Glu-29, Ser-142, and Lys-179/Arg-183 did not significantly alter it (Table II). Even though these residues, particularly Asp-140, Gly-139, Ser-142, and Gly-143, have been shown by mutagenesis and the crystal structure to interact with the distal water ligand (36, 39, 47), they do not appear to be key determinants of the high NO affinity of ferric hHO-1-heme.

The unusually low Kd value for the binding of NO to ferric hHO-1-heme suggests that the catalytic turnover of the enzyme should be inhibited by NO. Furthermore, as the catalytic cycle of hHO-1 traverses the ferrous state, NO could bind not only to the ferric but also to the ferrous protein, again inhibiting the enzyme. The Kd value for the binding of NO to the ferrous protein is expected to be much lower than that for the ferric enzyme, but the Kd value for binding to the ferric enzyme is already low enough that significant inhibition of the protein could occur at physiological or pathological NO concentrations, which range from 50 nM to 5 µM (59-65). Indeed, inhibition of heme oxygenase was observed when the enzyme was incubated with 100 µM concentrations of either NOC9 or Dea/NO (Fig. 7). These concentrations of NO donors provide a sufficient flux of NO to inhibit completely the enzyme at short periods (10-20 min) of incubation, but the inhibition is lost at longer incubation times as the NO donors are exhausted and the NO concentration falls. Inhibition by NO is reversible (Fig. 7), although a full recovery of the hHO-1 activity is not observed due to a residual inhibition caused by persistent NO donor decomposition products distinct from NO (Fig. 8).

The inhibition of heme oxygenase by NO and NO donors has received little attention (32, 33). A possible role for NO as an in vivo inhibitor of heme oxygenase is suggested by the report that L-arginine, the substrate of the nitric-oxide synthases, inhibits heme oxygenase activity when added to spleen or brain homogenates, whereas analogous addition of L-NAME, an inhibitor of the nitric-oxide synthases, stimulates the heme oxygenase activity (33). In the only molecular level study prior to this work, Ding et al. (32) reported that the NO donors 3-morpholinosydnonimine (SIN-1), S-nitroso-N-acetylpenicillamine (SNAP), and sodium nitroprusside inhibit HO-2 but not HO-1. The study focused on the possible role of the non-catalytic heme binding domains that are only present in HO-2. However, in that study the NO donors were removed by dialysis prior to measuring the residual catalytic activity, so that the reversible inhibition by NO reported here would not have been detected because the NO would be removed at the same time as the NO donors. Furthermore, Ding et al. (32) reported that enzyme inactivation was not observed with HO-2 when the cysteine residues within the two heme-binding motifs had been mutated to alanines. The irreversible inhibition they observed therefore reflects modifications related to the presence of the two cysteines, neither of which is present in HO-1. This is consistent with the present findings that HO-1 is not irreversibly inactivated by NO or NO donors.

In sum, the present results clearly establish that NO inhibits HO-1 by binding to the catalytic iron atom and show that this inhibition can involve binding to even the ferric heme complex due to its unusually high affinity for NO. The inhibition of heme oxygenase by NO occurs at concentrations of this agent that are pathologically and possibly physiologically accessible. The cross-talk between the nitric-oxide synthase and heme oxygenase systems may play some role in the pleiotropic responses that are associated with these two hemoprotein signaling systems.

    ACKNOWLEDGEMENTS

We thank Yi Liu for preparing the E29A, G139A, and S142A hHO-1 constructs and Luke Lightning for construction of the G143A, G143F, and K179A/K183A mutant plasmids.

    FOOTNOTES

* This work was supported by National Institutes of Health Grants DK30297 (to P. R. O. M.) and GM18865 (to P. M. L.).The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

To whom correspondence should be addressed: University of California School of Pharmacy, San Francisco, CA 94143-0446. Tel.: 415-476-2903; Fax: 415-502-4728; E-mail: ortiz@cgl.ucsf.edu.

Published, JBC Papers in Press, November 13, 2002, DOI 10.1074/jbc.M211131200

2 Y. Liu and P. R. Ortiz de Montellano, unpublished data.

    ABBREVIATIONS

The abbreviations used are: NO, nitric oxide; heme, iron protoporphyrin IX regardless of oxidation and ligation state; HO-1, heme oxygenase-1; hHO-1, truncated human HO-1; met-Mb, metmyoglobin; FTIR, Fourier transform infrared spectroscopy; RR, resonance Raman spectroscopy; NOC9, (Z)-1-[N-methyl-N-[6-(N-methylammoniohexyl)amino]]diazen-1-ium-1,2-diolate; Dea/NO, 2-(N,N-diethylamino)-diazenolate-2-oxide·sodium salt.

    REFERENCES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

1. Wink, D. A., and Mitchell, J. B. (1998) Free Radic. Biol. Med. 25, 434-456[CrossRef][Medline] [Order article via Infotrieve]
2. Murad, F. (1999) Angew. Chem. Int. Ed. Engl. 38, 1857-1868
3. Zhang, J., and Snyder, S. H. (1995) Annu. Rev. Pharmacol. Toxicol. 35, 213-233[CrossRef][Medline] [Order article via Infotrieve]
4. Snyder, S., Jaffrey, S., and Zakhary, R. (1998) Brain Res. Rev. 26, 167-175[Medline] [Order article via Infotrieve]
5. Maines, M. D., Trakshel, G. M., and Kutty, R. K. (1986) J. Biol. Chem. 261, 411-419[Abstract/Free Full Text]
6. Shibahara, S., Müller, R. M., Taguchi, H., and Yoshida, T. (1985) Proc. Natl. Acad. Sci. U. S. A. 240, 7865-7869
7. Rotenberg, M. O., and Maines, M. D. (1990) J. Biol. Chem. 265, 7501-7506[Abstract/Free Full Text]
8. Maines, M. D. (1992) Heme Oxygenase: Clinical Applications and Functions , CRC Press, Inc., Boca Raton, FL
9. Ishikawa, K., Sugawara, D., Wang, X., Suzuki, K., Itabe, H., Maruyama, Y., and Lusis, A. (2001) Circ. Res. 88, 506-512[Abstract/Free Full Text]
10. Hanselmann, C., Mauch, C., and Werner, S. (2001) Biochem. J. 353, 459-466[CrossRef][Medline] [Order article via Infotrieve]
11. Duckers, H. J., Boehm, M., True, A. L., Yet, S.-F., San, H., Park, J. L., Webb, R. C., Lee, M.-E., Nabel, G. J., and Nabel, E. G. (2001) Nat. Med. 7, 693-698[CrossRef][Medline] [Order article via Infotrieve]
12. Nath, K. A., Vercellotti, G. M., Grande, J. P., Miyoshi, H., Paya, C. V., Manivel, J. C., Haggard, J. J., Croatt, A. J., Payne, W. D., and Alam, J. (2001) Kidney Int. 59, 106-117[CrossRef][Medline] [Order article via Infotrieve]
13. Clark, J. E., Foresti, R., Green, C. J., and Motterlini, R. (2000) Biochem. J. 348, 615-619[CrossRef][Medline] [Order article via Infotrieve]
14. Otterbein, L. E., and Choi, A. M. K. (2000) Am. J. Physiol. 279, L1029-L1037
15. Zakhary, R., Poss, K. D., Jaffrey, S. R., Ferris, C. D., Tonegawa, S., and Snyder, S. H. (1997) Proc. Natl. Acad. Sci. U. S. A. 94, 14848-14853[Abstract/Free Full Text]
16. Grundemar, L., and Ny, L. (1997) Trends Pharmacol. Sci. 18, 193-195[CrossRef][Medline] [Order article via Infotrieve]
17. Liang, M., Croatt, A. J., and Nath, K. A. (2000) Am. J. Physiol. 279, F728-F735[Abstract/Free Full Text]
18. Alcaraz, M. J., Habib, A., Lebret, M., Creminon, C., LevyToledano, S., and Maclouf, J. (2000) Br. J. Pharmacol. 130, 57-64[Abstract/Free Full Text]
19. Hartsfield, C. L., Alam, J., Cook, J. L., and Choi, A. M. K. (1997) Am. J. Physiol. 273, L980-L988[Abstract/Free Full Text]
20. Chen, K., and Maines, M. D. (2000) Cell. Mol. Biol. 46, 609-617
21. Polte, T., Abate, A., Dennery, P. A., and Schroder, H. (2000) Arterioscler. Thromb. Vasc. Biol. 20, 1209-1215[Abstract/Free Full Text]
22. Datta, P. K., and Lianos, E. A. (1999) Kidney Int. 55, 1734-1739[CrossRef][Medline] [Order article via Infotrieve]
23. Kitamura, Y., Furukawa, M., Matsuoka, Y., Tooyama, I., Kimura, H., Nomura, Y., and Taniguchi, T. (1998) Glia 22, 138-148[CrossRef][Medline] [Order article via Infotrieve]
24. Takahashi, K., Hara, E., Ogawa, K., Kimura, D., Fujita, H., and Shibahara, S. (1997) J. Biochem. (Tokyo) 121, 1162-1168[Abstract]
25. Foresti, R., Clark, J. E., Green, C. J., and Motterlini, R. (1997) J. Biol. Chem. 272, 18411-18417[Abstract/Free Full Text]
26. Thom, S. R., Xu, Y. A., and Ischiropoulos, H. (1997) Chem. Res. Toxicol. 10, 1023-1031[CrossRef][Medline] [Order article via Infotrieve]
27. Cavicchi, M., Gibbs, L., and Whittle, B. J. R. (2000) Gut 47, 771-778[Abstract/Free Full Text]
28. Datta, P. K., Koukouritaki, S. B., Hopp, K. A., and Lianos, E. A. (1999) J. Am. Soc. Nephrol. 10, 2540-2550[Abstract/Free Full Text]
29. Turcanu, V., Dhouib, M., and Poindron, P. (1998) Res. Immunol. 149, 741-744[CrossRef][Medline] [Order article via Infotrieve]
30. Sun, J., Wilks, A., Ortiz de Montellano, P. R., and Loehr, T. M. (1993) Biochemistry 32, 14151-14157[Medline] [Order article via Infotrieve]
31. Ishikawa, K., Takeuchi, N., Takahashi, S., Matera, K. M., Sato, M., Shibahara, S., Rousseau, D. L., Ikeda-Saito, M., and Yoshida, T. (1995) J. Biol. Chem. 270, 6345-6350[Abstract/Free Full Text]
32. Ding, Y., McCoubrey, W. K., and Maines, M. D. (1999) Eur. J. Biochem. 264, 854-861[Abstract/Free Full Text]
33. Willis, D., Tomlinson, A., Frederick, R., Paul-Clark, M. J., and Willoughby, D. A. (1995) Biochem. Biophys. Res. Commun. 214, 1152-1156[CrossRef][Medline] [Order article via Infotrieve]
34. Shen, A. L., Christensen, M. J., and Kasper, C. B. (1991) J. Biol. Chem. 266, 19976-19980[Abstract/Free Full Text]
35. Liu, Y., and Ortiz de Montellano, P. R. (2000) J. Biol. Chem. 275, 5297-5307[Abstract/Free Full Text]
36. Lightning, L. K., Huang, H., Moenne-Loccoz, P., Loehr, T. M., Schuller, D. J., Poulos, T. L., and Ortiz de Montellano, P. R. (2001) J. Biol. Chem. 276, 10612-10619[Abstract/Free Full Text]
37. Wilks, A., Black, S. M., Miller, W. L., and Ortiz de Montellano, P. R. (1995) Biochemistry 34, 4421-4427[Medline] [Order article via Infotrieve]
38. Sambrook, J., Fritsch, E. F., and Maniatis, T. (1989) Molecular Cloning: A Laboratory Manual , Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY
39. Liu, Y., Lightning, L. K., Huang, H.-W., Moënne-Loccoz, P., Schuller, D. J., Loehr, T. M., Poulos, T. L., and Ortiz de Montellano, P. R. (2000) J. Biol. Chem. 275, 34501-34507[Abstract/Free Full Text]
40. Wilks, A., and Ortiz de Montellano, P. R. (1993) J. Biol. Chem. 268, 22357-22362[Abstract/Free Full Text]
41. Loehr, T. M., and Sanders-Loehr, J. (1993) Methods Enzymol. 226, 431-470[Medline] [Order article via Infotrieve]
42. Scheidt, W. R., and Ellison, M. K. (1999) Acc. Chem. Res. 32, 350-359[CrossRef]
43. Benko, B., and Yu, N. T. (1983) Proc. Natl. Acad. Sci. U. S. A. 80, 7042-7046[Abstract]
44. Laverman, L. E., and Ford, P. C. (2001) J. Am. Chem. Soc. 123, 11614-11622[CrossRef][Medline] [Order article via Infotrieve]
45. Brown, G. C. (1995) Eur. J. Biochem. 232, 188-191[Abstract]
46. Cooper, C. E. (1999) Biochim. Biophys. Acta 1411, 290-309[Medline] [Order article via Infotrieve]
47. Schuller, D. J., Wilks, A., Ortiz de Montellano, P. R., and Poulos, T. L. (1999) Nat. Struct. Biol. 6, 860-867[CrossRef][Medline] [Order article via Infotrieve]
48. Zhou, H., Migita, C. T., Sato, M., Sun, D. Y., Zhang, X. H., Ikeda-Saito, M., Fujii, H., and Yoshida, T. (2000) J. Am. Chem. Soc. 122, 8311-8312[CrossRef]
49. Laverman, L. E., Wanat, A., Oszajca, J., Stochel, G., Ford, P. C., and van Eldik, R. (2001) J. Am. Chem. Soc. 123, 285-293[CrossRef][Medline] [Order article via Infotrieve]
50. Hoshino, M., Laverman, L., and Ford, P. C. (1999) Coord. Chem. Rev. 187, 75-102[CrossRef]
51. Moore, E. G., and Gibson, Q. H. (1976) J. Biol. Chem. 251, 2788-2794[Abstract]
52. Miller, L. M., Pedraza, A. J., and Chance, M. R. (1997) Biochemistry 36, 12199-12207[CrossRef][Medline] [Order article via Infotrieve]
53. Addison, A. W., and Stephanos, J. J. (1986) Biochemistry 25, 4104-4113[Medline] [Order article via Infotrieve]
54. Shimizu, H., Obayashi, E., Gomi, Y., Arakawa, H., Park, S. Y., Nakamura, H., Adachi, S., Shoun, H., and Shiro, Y. (2000) J. Biol. Chem. 275, 4816-4826[Abstract/Free Full Text]
55. Roberts, S. A., Weichsel, A., Qiu, Y., Shelnutt, J. A., Walker, F. A., and Montfort, W. R. (2001) Biochemistry 40, 11327-11337[CrossRef][Medline] [Order article via Infotrieve]
56. Weichsel, A., Andersen, J. F., Roberts, S. A., and Montfort, W. R. (2000) Nat. Struct. Biol. 7, 551-554[CrossRef][Medline] [Order article via Infotrieve]
57. Asher, S. A., and Schuster, T. M. (1979) Biochemistry 18, 5377-5387[Medline] [Order article via Infotrieve]
58. Spinger, B. A., Sligar, S. G., Olson, J. S., and Phillips, G. N. (1994) Chem. Rev. 94, 699-714
59. Malinski, T., and Taha, Z. (1992) Nature 358, 676-678[CrossRef][Medline] [Order article via Infotrieve]
60. Malinski, T., Bailey, F., Xhang, Z. G., and Chopp, M. (1993) J. Cereb. Blood Flow Metab. 13, 355-358[Medline] [Order article via Infotrieve]
61. Malinski, T., Radomski, M. W., Taha, Z., and Moncada, S. (1993) Biochem. Biophys. Res. Commun. 194, 960-965[CrossRef][Medline] [Order article via Infotrieve]
62. Malinski, T., Taha, Z., Grunfeld, S., Patton, S., Kapturczak, M., and Tomboulian, P. (1993) Biochem. Biophys. Res. Commun. 193, 1076-1082[CrossRef][Medline] [Order article via Infotrieve]
63. Tsukahara, H., Gordienko, D. V., and Goligorsky, M. S. (1993) Biochem. Biophys. Res. Commun. 193, 722-729[CrossRef][Medline] [Order article via Infotrieve]
64. Shibuki, K. (1990) Neurosci. Res. 9, 69-76[Medline] [Order article via Infotrieve]
65. Shibuki, K., and Okada, D. (1991) Nature 349, 326-328[CrossRef][Medline] [Order article via Infotrieve]


Copyright © 2003 by The American Society for Biochemistry and Molecular Biology, Inc.