Thermodynamic Characterization of Five Key Kinetic Parameters that Define Neuronal Nitric Oxide Synthase Catalysis

Abstract

NO synthase (NOS) enzymes convert L-arginine to NO in two sequential reactions whose rates (k_cat1 and k_cat2) are both limited by the rate of ferric heme reduction (k_r). An enzyme ferric heme-NO complex forms as an immediate product complex and then undergoes either dissociation (at a rate that we denote as k_d) to release NO in a productive manner, or reduction (k_r) to form a ferrous heme-NO complex (Fe^IINO) that must react with O₂ (at a rate that we denote as k_ox) in a NO dioxygenase reaction that regenerates the ferric enzyme. The interplay of these five kinetic parameters (k_cat1, k_cat2, k_r, k_d, and k_ox) determine NOS specific activity, O₂ concentration response, and pulsatile versus steady-state NO generation. Here we utilized stopped-flow spectroscopy and single catalytic turnover methods to characterize the individual temperature dependencies of the five kinetic parameters of rat neuronal NOS (nNOS). We then incorporated the measured kinetic values into computer simulations of the nNOS reaction using a global kinetic model to comprehensively model its temperature-dependent catalytic behaviors. Our results provide new mechanistic insights and also reveal that the different temperature dependencies of the five kinetic parameters significantly alter nNOS catalytic behaviors and NO release efficiency as a function of temperature.

Introduction

Nitric oxide (NO) is a biological mediator produced in animals by three NO synthase isozymes (NOS, EC 1.14.13.39): inducible NOS (iNOS), neuronal NOS (nNOS), and endothelial NOS (eNOS) [1;2]. NOS enzymes catalyze a stepwise oxidation of L-arginine (Arg) to form nitric oxide and citrulline [3–5]. In the first reaction Arg is hydroxylated to form N^ω-hydroxy-L-arginine (NOHA), and in the second reaction the NOHA intermediate is oxidized to form NO and citrulline (Scheme 1) [1]. Both steps consume one molecule of O₂ and utilize NADPH-derived reducing equivalents. NOS proteins are comprised of an N-terminal oxygenase domain and a C-terminal flavoprotein domain, with a calmodulin (CaM)-binding site connecting the two domains [6] and each NOS is only active as a homodimer [6;7]. The oxygenase domain binds Fe-protoporphyrin IX (heme), the substrate Arg, and the essential cofactor (6R)-tetrahydrobiopterin (H₄B) [8;9], while the flavoprotein domain binds FAD, FMN, and NADPH [10–12]. During catalysis, the flavoprotein domain provides electrons to the heme in the NOS oxygenase domain, which enables the heme to bind O₂ and initiate oxygen activation in both reactions of NO synthesis [13–15]. Heme reduction also requires that CaM be bound to NOS, and is rate-limiting for the NO biosynthetic steps.

Scheme 1.

Biosynthesis of nitric oxide from L-arginine.

NOS enzymes operate under the constraint of being heme proteins that generate NO, a good heme-binding ligand. Indeed, their newly-made NO binds to the ferric heme many times before it can exit the enzyme [16]. How this intrinsic heme-NO binding event impacts NOS catalytic cycling is shown in Fig. 1, and has been previously discussed in detail [16–20]. The Arg to NO biosynthetic reactions (Fe^III to Fe^IIINO steps in Fig. 1) involves two sequential oxidation reactions whose rates are both limited by the rate of ferric heme reduction (k_r), because all subsequent catalytic steps in either reaction (termed k_cat1 and k_cat2 in Fig. 1) occur faster than k_r. Once the ferric heme-NO product complex forms in NOS, it can either dissociate and enable release of NO into the medium (at a rate k_d as shown in Fig. 1), or become reduced by the flavoprotein domain (at a rate k_r′ in Fig. 1; thought to be equal to k_r) [21] to form the enzyme ferrous heme-NO species (Fe^IINO), which releases NO extremely slowly [18;19]. Consequently, the Fe^IINO species must undergo reaction with O₂ at a rate k_ox (Fig. 1) to regenerate the ferric enzyme and return to an active form. Thus, two cycles exist during steady state NO synthesis (Fig. 1): NO dissociation from the enzyme (k_d) is part of a “productive cycle” that is essential for NOS bioactivity, whereas reduction of the Fe^IIINO product complex (k_r′) channels NOS into a “futile cycle” that actually represents a NO dioxygenase activity. The rate constants for each step of NOS catalysis are shown in Supplemental Fig. S1.

Fig. 1. Global kinetic model for NOS catalysis.

NOS enzymes start with their heme in the ferric state (left side, yellow Fe^III). The FMN-to-heme electron transfers (designated k_r) allow O₂ to bind to heme to start catalysis, and are rate-limiting for the two sequential oxidation reactions (horizontal line). k_cat1 and k_cat2 are the conversion rates of the enzyme Fe^IIO₂ species (green) to products in the L-Arg and N^ω-hydroxy-L-arginine (NOHA) oxidation reactions, respectively. A ferric heme-NO product complex (Fe^IIINO, blue) can either release NO (k_d) or become reduced (k_r′) to a ferrous heme-NO complex (Fe^IINO, red), which reacts with O₂ (k_ox) to regenerate ferric enzyme through a futile cycle that destroys the NO. The five kinetic parameters and reactions we studied are designated by the red boxes.

Observed rates for k_r, k_cat, k_d, and k_ox have been measured in NOS enzymes in single catalytic turnover reactions [14;18;19;22–25]. However, because most of the measures have been obtained at only a single temperature (10 °C, or less often at 25 °C), little or nothing is understood regarding the temperature dependencies and thermodynamic properties of the individual steps, and how they combine to govern the catalytic behavior and overall activities of NOS enzymes. This information would improve our understanding NOS catalysis and enable accurate modeling of its catalytic cycle under physiologic temperatures and conditions. In this report, we poised rat nNOS at different points in the cycle, and utilized stopped flow spectroscopy to characterize the temperature dependence of five individual kinetic parameters (k_r, k_cat1, k_cat2, k_d, and k_ox). The thermodynamic parameters of these individual reactions provide new insights regarding NOS structure-reactivity relationships and provide a better understanding of its physiological relevance.

Results and Discussion

Temperature Dependence of Heme Reduction (k_r)

We directly assessed electron transfer from FMN hydroquinone to heme using stopped-flow spectroscopy under anaerobic conditions. Heme reduction was monitored via the formation of the enzyme ferrous heme–CO complex at 444 nm. Once ferrous heme is formed by electron transfer from the FMN, the fast and stable binding of CO prevents accumulation of the ferrous species and makes the electron transfer reaction essentially irreversible. This mimics the normal condition of having O₂ present to react with ferrous heme, and insures that the observed rate of heme reduction does not have a significant back reaction component. The reactions were initiated by mixing CaM-bound nNOS with excess NADPH [26], and heme reduction rates (k_r) were measured at 5, 10, 15, 20, 25, 30 and 37 °C. Representative diode array spectra (Fig. 2, upper panel) and the resulting kinetic trace (Fig. 2, upper panel inset) are shown. We fit the kinetic traces to a bi-exponential function, with the initial fast phase representing an absorbance decrease due to flavin reduction by NADPH and the subsequent slower phase representing the absorbance increase due to heme reduction in the presence of CO. The observed rates obtained for heme reduction at each temperature are listed in Table 1. The heme reduction rate increased approximately 1.5-fold with every 10 degrees rise in temperature. Fitting the experimental data to the linearized form of the Arrhenius equation yielded an activation energy E_a = 31.5 kJ mol⁻¹. A linear Eyring plot (Fig. 2, middle panel) of the data yielded the following values for the enthalpy and entropy of activation for the heme reduction step: ΔH^‡ = +29.1 kJ mol⁻¹; ΔS^‡ = −128 J mol⁻¹K⁻¹ (Table 2). The derived Gibbs free energy of activation ΔG^‡ at 25 °C was calculated to be 67.2 kJ mol⁻¹. The ΔG^‡ increases with temperature, which is expected, given the negative entropic contribution. The activation enthalpy and entropy values for heme reduction that we determined for nNOS are similar to values recently reported for human iNOS (36.9 kJ mol⁻¹ and −89.7 J mol⁻¹K⁻¹ respectively), which were derived using a different measurement method (flash photolysis) [27]. This suggests similar mechanisms may govern heme reduction among NOS isozymes.

Fig. 2. Temperature-dependence of ferric heme reduction (k_r) in nNOS.

Anaerobic ferric nNOS (~10 μM) was mixed at various temperatures with NADPH (100 μM) in CO-saturated buffer in the stopped flow apparatus and buildup of the Fe^IICO absorbance peak (444 nm) was used to measure the rate of heme reduction. Upper panel- Rapid scanning spectra recorded during a representative reaction run at 10 °C and the accompanying kinetics of spectral change (inset). Middle panel- Eyring plot of the measured heme reduction rates, with slope = −ΔH^‡/R and the intercept = ΔS^‡/R + ln(k_B/h). Lower panel- Temperature dependence of the nNOS heme reduction rate in the context of the Marcus ET theory. The solid line represents a nonlinear least-squares fit of the data using equation 1 as described in the text. The derived Marcus parameters (λ and H_DA) giving the best fit are reported in the Figure.

Table 1.

Measured rates (± SD) at different temperatures.

Temperature (°C)	kr (s−1)	kcat1 (s−1)	kcat2 (s−1)	kd (s−1)	koxa (s−1)	NO synthesis (s−1)
5	4.2 ± 0.3	9.2 ± 0.8	16.4 ± 1.9	1.6 ± 0.2	0.063 ± 0.002	0.12 ± 0.01
10	5.3 ± 0.2	13. 5 ± 1.3	29 ± 3	3.7 ± 0.3	0.081 ± 0.005	0.27 ± 0.02
15	6.5 ± 0.4	21 ± 2	52 ± 5	5.9 ± 0.2	0.102 ± 0.002	0.35 ± 0.03
20	8.1 ± 0.6	31 ± 3	93 ± 4	10.7 ± 0.1	0.130 ± 0.001	0.55 ± 0.05
25	10.4 ± 0.5	46 ± 3	162 ± 15	20.1 ± 1.5	0.161 ± 0.003	0.82 ± 0.04
30	12.7 ± 1.1	68 ± 5	278 ± 18	31 ± 1	0.211 ± 0.004	1.12 ± 0.10
37	17.2 ± 0.6	113 ± 9	572 ± 36	58 ± 6	0.254 ± 0.002	1.40 ± 0.08

^aDetermined at ~ 202 μM O₂, 180 μM O₂, 162 μM O₂, 145 μM O₂, 135 μM O₂, 125μM O₂, and ~108 μM O₂ at 5, 10, 15, 20, 25, 30 and 37 °C, respectively (i.e. half air saturated concentration at respective temperature).

Table 2.

Thermodynamic parameters for k_r, k_ox, k_d, NO synthesis, k_cat1 and k_cat2. Errors reported are SD from at least three measurements

Reaction	Ea (kJ.mol−1)	ΔH‡ (kJ.mol−1)	ΔS‡ (J.mol−1K−1)
kr	31.5 ± 0.5	29.1 ± 0.5	−128 ± 3
kcat1	57.1 ± 2.3	54.7 ± 2.3	−29 ± 1
kcat2	79.5 ± 2.7	77 ± 3	+56 ± 2
kd	83.1 ± 1.7	80.7 ± 1.7	+50 ± 1
kox	31.9 ± 1.0	29.4 ± 1.0	−161 ± 14
NO synthesis	54.1 ± 4.5	51.7 ± 4.5	−74 ± 9

Ea values are calculated from Arrhenius plot

Activation parameters of heme reduction have been determined for other heme proteins such as azurin, cytochrome c oxidase and cd₁ nitrite reductase [28–31]. Particularly, activation parameters determined for interprotein electron transfer reactions, such as in cytochrome c-plastocyanin [32–34], indicate modest enthalpies and high negative entropies, similar to the values that we found for nNOS, for its putative inter-domain FMN-to-heme electron transfer. The high negative activation entropy determined for nNOS suggests that the rate-limiting elementary step of the heme reduction process is characterized by a relatively ordered transition state.

Because heme reduction in nNOS involves an electron transfer (ET) reaction between the FMN and heme prosthetic groups, we analyzed the temperature dependence data of heme reduction in the framework of the semi-classical Marcus theory describing nonadiabatic electron transfer reactions [35]. This approach has been successfully applied to several biological electron transfer systems in order to shed light on quantitative aspects of the ET step [36;37]. It is important to note that heme reduction in NOS enzymes is a relatively complex multi-step process. A protein conformational change that involves both long- and short-range motions are expected to bring the FMN domain into close proximity to the heme center in the NOSoxy domain [38;39]. Mutational and kinetic studies suggest that the FMN domain first undergoes a long range motion to swing away from the FNR domain and then undergoes short-range sampling motions to productively dock on the NOSoxy domain, which is in part guided by complementary charged residues on the domain-domain interface [11;39–41]. These conformational change(s) precede the actual ET and could conceivably influence the kinetics and thermodynamic behavior of the heme reduction that we observe. The ET process would be gated if the conformational change is rate-limiting, whereas if the actual FMN-to-heme ET step is rate-limiting, the process could be either true-ET or coupled-ET. In true-ET, the electron transfer step is rate-limiting and not kinetically influenced by preceding steps, while in coupled-ET the electron transfer reaction is still rate-limiting but is preceded by a relatively fast and thermodynamically unfavorable non-ET reaction such as conformational change. Although the FMN to heme ET in NOS enzymes has been reported to be conformationally-gated [42], a detailed study on this aspect is lacking. Application of the Marcus ET formalism to our temperature dependence data provides an opportunity to quantify parameters such as the required reorganization energy as well as electronic coupling between the FMN electron donor and heme electron acceptor during ET. The rate constant for nonadiabatic electron transfer, k_ET, is generally described by equations 1 and 2 which respectively, derive from the prediction that rates of ET exhibit Gaussian free-energy dependence, and from the fact that electronic coupling drops exponentially with distance between redox partners.

$$k_{ET}=\frac{4{\pi }^2{H}_{AB}^2}{h{(4\pi \lambda RT)}^{1/2}}exp\left[-\frac{{(\mathrm{\Delta }G°+\lambda )}^2}{4\lambda RT}\right]$$ (Equation 1)

$$k_{ET}=k_{0}exp[-\beta (r-r_0)]exp\left[-\frac{{(\mathrm{\Delta }G°+\lambda )}^2}{4\lambda RT}\right]$$ (Equation 2)

Where: λ is the reorganization energy, ΔG° is the thermodynamic driving force determined from the redox potential difference for the redox partners (FMN and Heme in this case). The other parameters have their usual meanings: Planck’s constant (h), the gas constant (R), and absolute temperature (T). H_DA describes the electronic coupling between electron donor (FMN) and electron acceptor (Heme). In equation 2: k_o =10¹³ s⁻¹ is the characteristic maximum limit for k_ET when donor and acceptor are in van der Waals’ contact (i.e. at r_o = 3 Å) and when no activation is involved (i.e. when and λ = −ΔG°). The value of parameter β reflects the nature of the intervening medium in mediating the ET, and a value of 1.4 Å⁻¹ is widely used for proteins [36].

In Fig. 2 (lower panel) the heme reduction data as a function of temperature were fit using equation 1 to obtain estimates of the reorganization energy (λ) and the electronic coupling element (H_DA). The ΔG° value used for the fitting was −0.030 eV (or −2.9 kJ mol⁻¹). This driving force is determined from the difference of midpoint values of the FMN hydroquinone/semiquinone couple and the heme Fe^III/Fe^IICO couple in nNOS [38;43–46]. Using this ΔG° value, the non-linear fitting yielded best-fit values for the reorganization energy λ = 139 kJ mol⁻¹ (or ~1.4 eV) and an electronic coupling factor H_DA = 0.2 cm⁻¹. We have varied ΔG° to see if the values determined by fitting are affected by the driving force used. Changes as much as ±60 mV in ΔG° do not result in significant changes in the fitted values of λ or H_DA. Fitting the same data to equation 2 gave a similar value for the reorganization energy λ = 135 kJ mol⁻¹ (Supplemental Fig. S2, Upper panel), which is not significantly different from λ = 139 kJ mol⁻¹ determined using equation 1.

The estimate for H_DA in nNOS is within the range known for nonadiabatic electron transfer [35], while the estimated reorganization energy (λ = 1.4 eV) lies on the higher end but is still within the range of reorganizational energies reported for most physiological ET reactions (i.e. λ = 0.5 to 2.3 eV) [36]. The relatively high value of λ in our case reflects a possible coupling of ET to the pre-requisite conformational change needed to bring the FMN domain within a distance required for ET to the heme in the nNOSoxy domain. Whether an ET reaction is gated or coupled depends on the relative rates of the ET and prerequisite non-ET steps. An estimated value of K_eq = 9.0 was reported for the FMN-FNR domain off/on conformational equilibrium for the reduced, CaM-bound nNOS at 10 °C [38], indicating that, once reduced, the FMN domain is mostly away from the FNR and thus in a conformation that could interact with nNOSoxy for ET. However, the corresponding k_off value associated with the FNR-FMN equilibrium under this condition is not yet known, and in our experimental design the FMN domain must first receive electrons from the FNR to become reduced. Also, the K_eq for the corresponding FMN-NOSoxy equilibrium has been studied in the CaM-bound nNOS poised at different redox states. The data indicate a low value (Keq ≤ 0.2) [47;48] exists in all cases. This suggests that very little FMN-nNOSoxy complex is present at equilibrium and that their interaction is intermittent and/or transient. Given the relatively slow rates for heme reduction that we observe in nNOS, the relatively high corresponding reorganization energy, and the unfavorable on/off thermodynamic equilibrium for the FMN-nNOSoxy interaction, ET from FMN to heme in nNOS is likely coupled to the pre-requisite conformational movement of the FMN domain. Similar ET reactions that are coupled to conformational equilibria are known. An example is the electron transfer reaction from methanol dehydrogenase to the heme in cytochrome c-551i where the electron transfer is rate-limiting but still influenced by a pre-requisite rapid but unfavorable conformational arrangement [49].

Our fit of the heme reduction versus temperature data to equation 2 also yielded an estimate for the distance ‘r’ (FMN-to-Heme distance) (Supplemental Fig. S2). The estimated distance at the time of the ET event is r ‘ 13.4 Å. It is important to note that this distance does not change significantly (r =13.1 Å) when the data is fit with equation 2 using the λ value found in nonlinear fitting with equation 1 (λ = 139 kJ mol⁻¹) (Supplemental Fig. S2, Lower panel). This estimate is in excellent agreement with 9–15 Å FMN-to-heme distances proposed based on docking models for FMN-NOSoxy complexes [40;50]. The value is also consistent with the proposed range (12–13 Å) suggested for nNOS based on the kinetics of heme-to-FMN back electron transfer determined by pulse radiolysis [42] and the recent studies on FMN-heme coupling by pulsed EPR for human iNOS [51]. In this case a 18.8 Å distance between the FMN-N5 atom and the heme iron was determined, that could correspond to a FMN-heme distance as short as 13.1 Å [51]. Comparable distances have been reported in related flavocytochrome systems. For instance, a value of 18.4 Å has been reported for the FMN to heme iron distance in the complex between the heme and FMN-binding domain of cytochrome P450BM-3 [52].

Temperature dependence of the oxygenation reactions (k_cat1 and k_cat2)

The k_cat1 and k_cat2 parameters represent the conversion rates of the heme Fe^IIO₂ species to products in the Arg hydroxylation and NOHA oxidation reactions, respectively. We utilized stopped-flow spectroscopy to compare the nNOS heme transitions that occur during the single turnover reactions as reported in previous studies [53;54]. The reactions were initiated by rapid-mixing an oxygen-containing buffer with pre-reduced ferrous nNOS proteins that contained H₄B and either Arg or NOHA as substrate. For the Arg reactions we observed two consecutive heme transitions: Fe^II → Fe^IIO₂ → Fe^III and in the reaction with NOHA we observed three transitions: Fe^II → Fe^IIO₂ → Fe^IIINO → Fe^III. Global analysis of the spectral data according to these models yielded the spectrum of each species (Fig. 3A & B, upper panels). The two middle panels of Fig. 3 show how the concentration of each species changed with time during the reactions. Linear Eyring plots of the observed rates (Fig 3A & B, lower panels) show that both k_cat1 and k_cat2 increase as the temperature increases (Table 1), but do so according to significantly different ratios, such that the slope of k_cat2 is 1.4 times higher than that of k_cat1 (Fig 3A & B, lower panels). Fitting the experimental data to the Arrhenius equation yielded an activation energy E_a = 57.1 kJ mol⁻¹ for k_cat1 and 79.5 kJ mol⁻¹ for k_cat2. The linear Eyring plots gave the following values for the enthalpy and entropy of activation were derived for k_cat1; ΔH^‡ = +54.7 kJ mol⁻¹; ΔS^‡ = −29 J mol⁻¹K⁻¹ (Table 2), whereas the values for the enthalpy and entropy of activation derived for k_cat2 are: ΔH^‡ = +77 kJ mol⁻¹; ΔS^‡ = +56 J mol⁻¹K⁻¹ (Table 2). The thermodynamic parameters derived for k_cat1 are distinguished from k_cat2 by a negative entropic component. The relative entropic cost observed in the first catalytic step may be attributed to the necessary change in the hydrogen-bonding network expected at the active site upon the transformation of L-arginine to the enzyme bound N-hydroxy-L-arginine [55].

Fig. 3. Temperature dependence of the Arg and NOHA oxidation reactions (k_cat1 and k_cat2) measured by single catalytic turnover reactions.

Ferrous nNOSoxy containing H₄B and either Arg or NOHA was mixed at various temperatures with air-saturated buffer in the stopped flow apparatus and diode array spectral data were collected. Upper panels, A & B show the spectrum of each heme species that was detected during each reaction as calculated by global analysis. Middle panels, A & B show the concentration versus time profiles for each heme species detected in each reaction. Lower panels, A & B show Eyring plots (ln(k_cat/T) vs 1/T) for the k_cat1 and k_cat2 transitions.

Temperature Dependence of NO dissociation (k_d)

A variety of studies [14;25;56;57] indicate that newly-formed NO binds to the NOS ferric heme before it can exit the enzyme. This causes a ferric heme–NO product complex to form at the end of each catalytic cycle in all NOS enzymes examined to date [19]. In the NOHA single turnover reactions there was a clear buildup of this immediate Fe^IIINO product complex (Fig. 3B, upper and middle panels), which allows us to determine the macroscopic off-rate of NO by monitoring its transition to ferric enzyme (k_d parameter in Fig. 1). The macroscopic, observed dissociation rate (k_d) of the ferric heme-NO product complex is an important parameter, because it impacts NOS enzyme distribution during catalysis and consequently, the steady-state NO synthesis activity [19;23;25]. Linear Eyring plots of the observed rates (Table 1 and Fig. 4, upper panel) revealed that the k_d parameter has a relatively large temperature dependence, increasing more than three times every 10 degrees. The thermodynamic parameters of the reaction were derived from the linear Eyring plot and Arrhenius equation, and gave the following values: E_a = 83.1 kJ mol⁻¹; ΔH^‡ = 80.7 kJ mol⁻¹; ΔS^‡ = 50 J mol⁻¹K⁻¹ (Table 2). The activation energy for NO dissociation from Fe^III-nNOS (k_d) is quite close to the activation energy for citrulline formation by nNOS as reported earlier by Iwanaga (~103 kJ mol⁻¹) [58]. Also, the relatively large positive activation entropy we calculate for the k_d in nNOS is similar to values measured for NO dissociation from other ferric heme proteins like metmyoglobin (ΔH^‡ = 78 kJ mol⁻¹; ΔS^‡ = 46 J mol⁻¹K⁻¹) [59] and the camphor-bound cytochrome P450_cam (ΔH^‡ = 84 kJ mol⁻¹; ΔS^‡ = 41 J mol⁻¹K⁻¹) [60]. Overall, the process of NO release from ferric heme in nNOS exhibits similar thermodynamic behavior to that of other heme proteins [59;60] as indicated by comparing Eyring plots for NO release data for the different heme proteins across a range of temperatures (Fig. 4, lower panel). Their similar thermodynamic behaviors is indicative of a common mechanism regulating NO dissociation from the ferric heme and its escape into solution.

Fig. 4. Temperature dependence of NO release (k_d) from the Fe^IIINO product complex of nNOS.

NOHA single turnover reactions were run with nNOSoxy at different temperatures as described in Fig. 3. Upper panel- Eyring plot of measured NO release rates that were determined from the indicated spectral transition. Lower panel- Eyring plots of the NO release rates of nNOS (this work), metmyoglobin [59], and cytochrome P450cam proteins [60].

What is the basis for the relatively large temperature effect on k_d? Binding and dissociation of diatomic ligands such as NO, CO, and O₂ to heme proteins is generally accompanied by conformational changes that sometimes extend far beyond the local heme binding site [61–63]. This is a consequence of the disparate geometric, electronic, and electrostatic requirements imposed by ligand binding and dissociation. The local and global conformational changes and dynamic electrostatic interactions in the binding pocket modulate not only ligand affinities but also the discrimination between similar diatomic ligands [64;65]. The large temperature dependence of the observed rate of NO dissociation from ferric-heme in nNOS and other heme proteins may reflect the control of NO binding at multiple levels. At the heme level, heme distortions have been shown to be critical for stabilizing the Fe^IIINO complex, particularly in heme proteins whose physiological functions depend on controlling NO release from Fe^III-heme [63]. In NOS proteins, the observed out-of-plane modes in the low frequency regions of Raman spectra of the Fe^IIINO species (iNOSoxy and eNOSoxy) have been assigned to heme distortions and have been shown to be critical in increasing d_xy-dπ orbital overlap, which stabilizes the Fe^III-heme-NO species [62]. At the global protein level, it has been suggested that inter-subunit interactions in the NOS dimer may be important in modulating heme distortion modes [62], which in turn would affect the heme-NO dissociation rate k_d. Also, point mutagenesis studies on NOS enzymes identified amino acids near the opening of the heme pocket that help to kinetically gate release of NO [25;66;67]. Thus, the large dependence of k_d on the temperature is likely due to a general temperature effect on both the heme and protein structural fluctuations and dynamics.

Reaction of O₂ with the Enzyme Ferrous Heme-NO Complex (k_ox)

During NO synthesis a portion of the ferric heme-NO product complex is reduced by the NOS reductase domain and the resulting ferrous heme-NO complex must then react with O₂ to return to the catalytic cycle. The rate of this reaction, termed k_ox, is a kinetic parameter (Fig. 1) that impacts NOS enzyme distribution during catalysis and, consequently, its steady state behavior and NO synthesis activity [19;68–70]. We determined k_ox values by mixing anaerobic samples of the nNOSoxy ferrous heme-NO species with an air-saturated solution in the stopped-flow spectrophotometer at 5, 10, 15, 20, 25, 30 and 37 °C and then following the reactions using a diode array detector. Figure 5 (upper panel) contains representative spectral data collected during the reaction of the nNOSoxy Fe^IINO complex with air-saturated buffer in the presence of both H₄B and L-Arg at 10 °C. The spectral changes are consistent with conversion of the Fe^IINO complex into the Fe^III high-spin form of nNOSoxy. The reaction was a single-step process with no discernible accumulation of intermediate species, as indicated by the several isosbestic points in the spectra (Fig. 5, upper panel) and by the single-exponential decay of the absorbance signal at two different wavelengths (Fig. 5, upper panel, inset), that followed the rate of Fe^IINO complex disappearance (436 nm) and the rate of ferric enzyme formation (393 nm). The spectral and fitting results obtained for reactions run at other temperatures were highly similar (data not shown), and the observed k_ox rates are listed in Table 1. The corresponding Eyring plot (Fig. 5, lower panel) and Arrhenius equation were used to calculate the following thermodynamic parameters: E_a = 31.9 kJ mol⁻¹; ΔH^‡ = 29.4 kJ mol⁻¹; ΔS^‡ = −161 J mol⁻¹K⁻¹) (Table 2).

Fig. 5. Temperature dependence of the O₂ reaction with the Fe^IINO species (k_ox).

Anaerobic Fe^IINO nNOSoxy was reacted at various temperatures with air-saturated buffer in the stopped flow instrument. Upper panel- rapid scanning spectra recorded during a representative reaction, with arrows indicating the direction of the absorbance changes. Inset, absorbance changes versus time at 436 nm (Fe^IINO disappearance) and 393 nm (Fe^III buildup). Lower panel- Eyring plot of the measured k_ox values.

The activation enthalpy for oxidation of the nNOS ferrous-heme complex is much lower than the values reported for oxidation of ferrous heme-nitrosyl hemoglobin and myoglobin or for their various mutants (ΔH^‡ = 110–120 kJ mol⁻¹) [61;71]. Their higher activation enthalpies have been rationalized in terms of a rate-limiting NO dissociation step governing the kinetics of the reversible ligand exchange reaction with O₂ that precedes the irreversible oxidation [71]. In contrast, the relatively low ΔH^‡ (29.4 kJ mol⁻¹) that we measured for oxidation of the nNOS ferrous heme-nitrosyl species effectively argues against a NO dissociative mechanism being involved in this case, because it is insufficient for breaking the strong heme-Fe^II-NO bond. The relatively low activation barrier for the nNOS reaction supports our recent mechanistic investigation of nNOS and iNOS heme-nitrosyl oxidation reactions, which suggested it involves a comparatively fast and direct reaction between the ferrous heme-nitrosyl and dioxygen [24].

Steady-state NO synthesis activity

The steady state NO synthesis activity is a combined reflection of all the individual steps in Fig. 1, thus its temperature dependence provides apparent thermodynamic parameters for the global process. Table 1 contains NO synthesis activities we measured across the temperature range for this study (5 to 37 °C). The activity increased about 12-fold over this range. The activity data gave a linear Eyring plot (Fig. 6), from which we determined the following activation parameters: E_a = 54.1 kJ mol⁻¹; ΔH^‡ = 51.7 kJ mol⁻¹; ΔS^‡ = −74 J mol⁻¹K⁻¹ (Table 2). Our E_a value differs somewhat from an earlier study that reported an activation energy of 79.4 kJ mol⁻¹ for the steady-state activity of nNOS, which was derived using measures of citrulline formation from [14C]L-Arg [58].

Fig. 6. Eyring plot of nNOS steady-state NO synthesis activities recorded at different temperatures.

Activities were measured in a conventional spectrophotometer using the oxyhemoglobin assay for NO as described in Experimental Methods.

Implications for nNOS catalytic behavior

Because the five kinetic parameters studied here differed in their temperature dependencies, this means that the distribution pattern of nNOS enzyme species during steady-state NO synthesis, and thus nNOS catalytic behaviors, will be temperature-dependent. We utilized the kinetic values we measured for the five parameters at three different temperatures (10, 25, and 37 °C) in computer simulations of the global kinetic model (Fig. 1) [21] to explore this concept. The model reports on the five main enzyme species that are present during steady-state NO synthesis, namely the ferric, ferrous, ferrous-O₂ (or ferric-superoxy), ferric-NO, and ferrous-NO forms (Fig. 1). Any additional kinetic values for associated reactions (i.e., like O₂ binding) that we needed for the simulations, besides the five kinetic parameters measured here, are described in Experimental procedures.

At 10 °C, the simulation showed that the fast k_r relative to the slower k_d and k_ox values causes the nNOS enzyme to exist predominantly as the ferrous-NO species during the steady state [19;21]. The modeled enzyme distribution indicated that ~10 % of nNOS is present as ferric enzyme and about 86% as the ferrous heme-NO species (Fig. 7) at 10 °C, with the remaining ~4% of nNOS mainly distributed between the Fe^IIO₂ and Fe^IIINO species. This falls close to estimates reported in literature [16;19;21;72;73], which range between 67 and 85% ferrous heme-NO complex during steady state catalysis at 10 °C. However, our experimental data reveal that at higher temperatures the rate of ferric heme-NO dissociation (k_d) increases significantly faster than the k_r and the other kinetic parameters. The simulations indicate how this will alter the steady-state enzyme distributions: At 25 °C ~16% of nNOS is present as ferric enzyme and about 81% as the ferrous heme-NO species (Fig. 7), with the remaining ~3% of nNOS mainly distributed between the Fe^IIO₂ and Fe^IIINO species, whereas at 37 °C, which is near the physiologic temperature of rats (~38 °C) [74;75], only about 73% of nNOS is present as the ferrous-NO species and ~24% is present as ferric enzyme. These simulations clearly show that rat nNOS gains in ferric character and has less ferrous heme–NO complex buildup during NO synthesis as the temperature increases. Overall, the different thermodynamic behaviors of the individual kinetic steps will cause nNOS to become more and more efficient at NO release as the temperature increases, while diminishing the enzyme cycling into the non-productive NO dioxygenase pathway (see Fig. 1).

Fig. 7. Temperature effect on the distribution of the five principal enzyme species during steady-state NO synthesis by nNOS.

Distributions were calculated by computer simulating a global kinetic model similar to the one shown in Fig. 1. Rate constants used for the calculations are given in Table 3. Steady state values were determined after 50s of simulated reaction although a steady state distribution is reached within 5–10 seconds in all cases.

A related consequence of the different temperature dependencies is that the relative contributions of the individual kinetic steps toward limiting the NO synthesis activity, and in determining the enzyme’s catalytic behaviors, will change with temperature. For example, at low temperatures like 10 °C, the k_d and k_r have similar magnitudes. This equivalence diminishes the release of NO due to its causing about half of the Fe^IIINO product complex to be diverted into the futile cycle. Consequently, this increases the impact of k_ox on the observed enzyme NO synthesis activity and on its apparent oxygen concentration dependence (i.e., apparent K_mO₂). Specifically, NO synthesis activity becomes much more dependent on the oxygen concentration relationship of the NO dioxygenase reaction described by k_ox [72], which displays a linear relationship with O₂ concentration across the entire physiologic range [24]. However, at higher temperatures the k_d becomes increasingly faster than the k_r, which enables a quicker and greater release of NO from the Fe^IIINO product complex, consequently diminishing enzyme partitioning into the futile cycle. This in turn diminishes the role of k_ox in limiting the overall NO synthesis activity and diminishes its role in determining the apparent K_mO₂ for NO synthesis activity, such that the apparent K_mO₂ becomes more reflective of the oxygen concentration dependence of the biosynthetic reactions, which depend on the ferrous heme affinity for O₂ binding. Thus, at higher temperatures (i.e., 37 °C) the contribution of k_r in determining the overall NO synthesis activity is increased, while the contribution of k_ox is diminished, with accompanying changes in nNOS enzyme behavior and O₂ concentration response.

Experimental Procedures

General Methods and Materials

All reagents and materials were obtained from Sigma-Aldrich, Amersham Biosciences or other sources as previously reported [20;70;76]. Absorption spectra and steady-state kinetic data were obtained using a Shimadzu UV-2401PC spectrophotometer. Stopped flow experiments were performed using a Hi-Tech Scientific KinetAsyst SF-61DX2 stopped-flow system equipped with anaerobic set-up and rapid scanning photodiode array detector. Data from multiple identical stopped-flow experiments were averaged to improve the signal-to-noise ratio. We didn’t see much difference in values when individual rates are averaged or the rate is calculated from averaged trace. We calculated SD values from the individual observed rates. The spectral traces were fit according to single or multiple exponential equations using software provided by the instrument manufacturer. The best fit was determined when adding further exponentials did not produce a significant improvement of the fit as judged from the residuals. All plots and some additional curve-fitting were done using Origin® 8.0 (OriginLab, Northampton, MA). Anaerobic samples were prepared in an air-tight cuvette using repeated cycles of vacuum followed by a positive pressure of catalyst-deoxygenated nitrogen gas.

Expression and Purification of Wild-type Protein

All proteins were purified in the presence of H₄B and L-Arg as described previously [20;26;70;76]. The ferrous heme-CO adduct absorbing at 444 nm was used to measure heme protein content with an extinction coefficient of ₄₄₄ = 74 mM⁻¹ cm⁻¹ (A₄₄₄–A₅₀₀) [76]. Purity of each protein was assessed by SDS-PAGE and spectral analysis.

NO Synthesis measurement

Steady-state NO synthesis was determined at 5, 10, 15, 20, 25, 30 and 37 °C using the spectrophotometric oxyhemoglobin assay as described previously [76].

Anaerobic Heme Reduction Measurements

The kinetics of heme reduction were analyzed at 5, 10, 15, 20, 25, 30 and 37 °C as described previously [20;26;70;76;77] using a stopped-flow apparatus and diode array detector (Hi-Tech Scientific KinetAsyst SF-61DX2) equipped for anaerobic analysis.

Reaction of Ferrous Heme-NO Complexes with Oxygen

The kinetics of Ferrous heme-NO oxidation were analyzed at 5, 10, 15, 20, 25, 30 and 37 °C as described previously in detail [24;70] using a stopped-flow apparatus and diode array detector (Hi-Tech Scientific KinetAsyst SF-61DX2) equipped for anaerobic analysis.

Single turnover reactions

Arginine hydroxylation experiments were carried out in a Hi-Tech SF61-DX2 stopped-flow instrument (Hi-Tech Scientific, Salisbury, UK) coupled to a diode array detector, as previously described [14]. The reactions were studied at 5, 10, 15, 20, 25, 30 and 37 °C in the presence of 50 μM H₄B, 2.5 mM L-Arginine, 150 mM NaCl, 10 % Glycerol, 1 mM DTT in 40 mM EPPS buffer, pH 7.6. 10 μM nNOSoxy was titrated with sodium dithionite and mixed with air-saturated buffer at respective temperatures ([O₂] = ~360 μM, 270 μM and ~215 μM at 10, 25 and 37 °C, respectively). NOHA oxidation was studied in the same conditions replacing L-Arginine by 1 mM NOHA. Sequential spectral data were fitted to A→ B→ C (Arg hydroxylation; Fe^II→ Fe^IIO₂→Fe^III) and A→ B→ C→ D (NOHA oxidation; Fe^II→Fe^IIO₂→Fe^IIINO→Fe^III) kinetic models using the Specfit/32 global analysis software, Version 3.0 (Spectrum Software Associates, Marlborough, MA) to obtain the spectra of the different species and the reaction rates. From the Specfit analysis software we calculated rates for k_cat1 (during Arg hydroxylation) and k_cat2 (during NOHA oxidation).

Temperature-Dependence and Thermodynamic Analysis

The natural log of the observed rates or rate constants for each averaged set of experimental data were plotted against the reciprocal of the absolute temperature. To calculate the activation energy, the data were then fit to the Arrhenius equation (Equation 3) using the linear fitting function using Origin® software (OriginLab, Northampton, MA). In this equation, A is the Arrhenius pre-exponential factor and R is the gas constant and E_a is activation energy. Activation enthalpy (ΔH^‡) and activation entropy (ΔS^‡) were calculated on the basis of Eyring equation (Equation 4) and Eyring plot (ln (k/T) vs. 1/T). Because the Gibbs free energy of activation (ΔG^‡) depends on temperature, it can be calculated for each step at a given temperature and we calculated ΔG^‡ at 25 °C using the Equation 5, whereas, ΔS^‡ and ΔH^‡ are virtually independent of temperature.

$$\text{ln}k=\text{ln}A-E_{\text{a}}/RT$$ (Equation 3)

$$ln(k/T)=-\mathrm{\Delta }{H}^{‡}/RT+\mathrm{\Delta }{S}^{‡}/R+ln(k_{\text{B}}/h)$$ (Equation 4)

$$\mathrm{\Delta }{G}^{‡}=RT[ln(k_{\text{B}}T/h)-\text{ln}k]$$ (Equation 5)

Where,

• k = reaction rate constant

• T = absolute temperature

• R = gas constant (8.314472 J mol⁻¹ K⁻¹)

• k_B = Boltzmann constant(1.3806504 × 10⁻²³ J K⁻¹)

• h = Planck’s constant (6.626 × 10⁻³⁴J s)

• ΔH^‡ = Enthalpy of activation

• ΔS^‡ = Entropy of activation

• ΔG^‡ = Gibbs free energy of activation

• E_a = Energy of activation

Simulations of nNOS Distribution During Steady-state NO Synthesis

The NO synthesis kinetics were simulated using the global model as implemented in MathCAD 7.0 [18;21] or Gepasi 3.30 [78]. Slight changes in the model were introduced to account for the pterin reduction step as shown in Fig. 1. Both methods yielded similar results. Simulations assume constant values for [O₂] = ~360 μM, 270 μM and ~215 μM at 10, 25 and 37 °C, respectively, and [NADPH] = 40 μM. This approximation allows to treat all the processes as unimolecular reactions and assimilate the observed rates to apparent rate constants. Thus, for bimolecular reactions k₂, k₆ and k₁₀ (Table 3) the observed rate is the product of the real rate constant multiplied by the concentration of oxygen. We base our simulations on the observed rates using a correction factor to estimate the value in air-saturated buffer instead of the half air-saturated experimental conditions. Experimentally observed oxidation rates (k_ox), which we derived at half air-saturated condition, were multiplied by a factor of 1.5 to approximate a full air-saturated condition according to the dependence observed previously [24]. We also multiplied the observed oxygen binding rates k₂ and k₆ values by a factor of 2.0 to get a full air-saturated condition assuming that the rate is proportional to [O₂]. The values for heme reduction (k_r), NO dissociation (k_d), ferrous heme-NO oxidation (k_ox) and all other rates at different temperatures are shown in Table 3. Percentage of species in the steady state is calculated after 50 s of simulation, although a steady state distribution is reached within 5–10 seconds in all cases.

Table 3.

Calculated rates used for computer simulations

			Temperature
			5 °C	10 °C	15 °C	20 °C	25 °C	30 °C	37 °C
k1	FeIII(a) → FeII(a)	kr	4.16	5.26	6.54	8.11	10.4	12.7	17.2
k2	FeII(a) (+ O2) → FeIIO2(a)		56.8	115	228	438	826	1526	3488
k3	FeIIO2(a) → FeIII(b*)	kcat1	9.2	13.5	20.6	31	46.1	67.7	113
k4	FeIII(b*) → FeIII(b)	kr′	4.16	5.26	6.54	8.11	10.4	12.7	17.2
k5	FeIII(b) → FeII(b)	kr″	4.16	5.26	6.54	8.11	10.4	12.7	17.2
k6	FeII(b) (+ O2) → FeIIO2(b)		56.8	115	228	438	826	1526	3488
k7	FeIIO2(b) → FeIIINO	kcat2	16.4	28.7	52.2	92.9	162	278	572
k8	FeIIINO → FeIII(a) + NO	kd	1.58	3.68	5.89	10.7	20.1	31.1	58.2
k9	FeIIINO → FeIINO	kr‴	4.16	5.26	6.54	8.11	10.4	12.7	17.2
k10	FeIINO (+ O2) → FeIII(a) + NOx	kox	0.0948	0.121	0.153	0.191	0.233	0.31	0.381
k11	FeIINO → FeII(a) + NO		0.000052	0.000096	0.00018	0.00031	0.00054	0.00092	0.0019

^(a)denotes L-Arg-bound enzyme;

^(b)denotes NOHA-bound enzyme;

Fe^III(b*) indicates the ferric enzyme with NOHA and H₄B radical bound, see scheme in Fig. 1 for details. All values are in s-1. For bimolecular reactions (k2, k6, k10) the value used in the calculations is the product of the “actual” value and the concentration of oxygen, which is considered to remain constant during the reaction. For k2 and k6 the values used are the experimental values multiplied by 2 (assumes the rate is proportional to [O₂], experimental values are determined in half air-saturated conditions). For k10 the values used are the experimentally determined values x 1.5 (based on observed oxygen dependence seen in Tejero et al [24]). k11 values are interpolated from Salerno et al paper [17].

Supplementary Material

Supp Figure S1-S2

Supplemental Fig. S1: Global kinetic model for NOS.

Supplemental Fig. S2: Analysis of the temperature dependence data of observed heme reduction rates for nNOS using the modified Marcus equation with distance dependence.

Abbreviations

EPPS

4-(2-hydroxyethyl)-1-piperazinepropanesulfonic acid

Fe^IIINO

ferric heme–NO complex

Fe^IINO

ferrous heme–NO complex

H₄B

(6R)-5,6,7,8-tetrahydro-L-biopterin

k_d

dissociation rate of NO from the ferric heme–NO complex of NOS

k_ox

oxidation rate of the ferrous heme–NO complex of NOS

k_r

reduction rate of heme by the reductase (flavoprotein) domain of NOS

nNOS

neuronal nitric oxide synthase, nNOSoxy, oxygenase domain of the nNOS

NOS

nitric oxide synthase

CaM

Calmodulin

FNR

ferredoxin NADP-reductase