Journal of Inorganic Biochemistry 103 (2009) 1102–1112

Contents lists available at ScienceDirect

Journal of Inorganic Biochemistry

journal homepage: www.elsevier .com/locate / j inorgbio

Trp180 of endothelial NOS and Trp56 of bacterial saNOS modulate sigma bondingof the axial cysteine to the heme

Jérôme Lang a, Danelle Driscoll b, Stéphanie Gélinas a,1, Steven P. Rafferty b, Manon Couture a,*

a Département de biochimie et de microbiologie and PROTEO, Pavillon Charles-Eugène Marchand, Room 4165, Université Laval, Québec, Canada G1V 0A6b Department of Chemistry, Trent University, Peterborough, Ontario, Canada K9J 7B8

a r t i c l e i n f o

Article history:Received 27 November 2008Received in revised form 11 May 2009Accepted 15 May 2009Available online 24 May 2009

Keywords:Nitric oxide synthaseNOSResonance Raman spectroscopyHeme

0162-0134/$ - see front matter � 2009 Elsevier Inc. Adoi:10.1016/j.jinorgbio.2009.05.011

* Corresponding author. Tel.: +1 418 656 2131x135E-mail address: manon.couture@bcm.ulaval.ca (M

1 Present address: Centre de recherche sur les maladCHUL, université Laval, Québec, Canada G1K 7P4.

a b s t r a c t

The proximal ligand of thiolate-coordinated heme proteins is crucial for the activation of the oxygen mol-ecule and hydroxylation of substrates. In nitric oxide synthases (NOSs), the heme axial cysteine ligandforms a hydrogen bond to the side chain indole nitrogen of a tryptophan residue. Resonance Raman spec-troscopy was used to probe W56F and W56Y variants of the NOS of Staphylococcus aureus (saNOS) and theanalogous W180 variants of the endothelial NOS oxygenase domain (eNOSox). We show that the variantsdisplayed lower mFe–NO and mFe–CO frequencies indicating that these mutations increased the electron den-sity on the axial cysteine in their FeIIINO and FeIICO complexes. We also show by UV-visible spectroscopythat the FeIICO complexes of the variants displayed a red-shifted Soret optical transition in addition to thelower mFe–CO thus establishing that these properties are sensitive indicators of the modulation of the basi-city of the axial cysteine. We infer, based on its spectroscopic properties, that ferrous eNOSox W180Y sat-urated with L-arginine and tetrahydrobiopterin forms a tyrosine–cysteine hydrogen bond when bound toCO. Evidence for such a hydrogen bond was not obtained for the FeIIINO protein nor for the analogoussaNOS variant. These mutations reveal interesting differences in the response of NOS isotypes to analo-gous mutations at conserved residues and clearly show that the heme-Fe to cysteine r bond is modulatedby the Cys–Trp hydrogen bond in NOSs. These studies serve as a basis to gain information on the roleplayed by this hydrogen bond in oxygen activation in this class of enzymes.

� 2009 Elsevier Inc. All rights reserved.

1. Introduction

Nitric oxide synthases (NOSs) are heme proteins that catalyzethe oxidation of L-arginine to citrulline and nitric oxide. NOSs arefound across the eukaryotic kingdom; of these, the three mamma-lian NOS isoforms, inducible (iNOS), neuronal (nNOS), and endo-thelial (eNOS) are the best characterized. The nitric oxideproduced by these enzymes is utilized in immune response, signaltransduction, and vasodilation, respectively [1–4]. All three iso-forms share a common homodimeric structure. Each subunit con-sists of a carboxy-terminal reductase domain, which binds to andtransfers electrons from NADPH through the cofactors FAD andFMN to the amino-terminal oxygenase domain, which binds heme,(6R) 5,6,7,8-tetrahydro-L-biopterin (H4B), and the substrates oxy-gen and L-arginine [5,6]. These two domains are separated via acalmodulin (CaM)-binding region. Dimerization of the holo-en-

ll rights reserved.

15; fax: +1 418 656 7176.. Couture).ies lipidiques (CRML), Pavillon

zyme occurs primarily through the oxygenase domains; however,the dimer is strengthened through additional interactions betweenthe oxygenase and reductase domains [7]. Dimerization of iNOS re-quires the presence of H4B; this cofactor stabilizes the formed di-mer in nNOS and eNOS [7,8]. It is therefore observed that iNOSforms the weakest dimer and eNOS forms the strongest dimer ofthe mammalian isoforms.

Less well studied are the bacterial NOSs, such as those fromStaphylococcus aureus (saNOS) [9,10], Bacillus subtilis (bsNOS)[11], Deinococcus radiodurans (drNOS) [12] and Geobacillus stearo-thermophilus (gsNOS) [13]. In contrast to the multidomain struc-ture of the eukaryotic NOSs, those from bacteria contain only theoxygenase domain, thus electrons required for catalysis are likelysupplied from a separate reductase. Evidence that NO generatedby bacterial NOSs is physiologically important is scarce but mount-ing. NO produced by plant-pathogenic Streptomyces species is in-volved in the synthesis of a peptide phytotoxin and theproduction of NO was recently detected in vivo [14,15]. In B. subtil-is, bsNOS seems important in the protection against oxidativestress [16]. Interestingly, B. anthracis NOS (baNOS) is essential forspore virulence. This activity may also be related to the protection

J. Lang et al. / Journal of Inorganic Biochemistry 103 (2009) 1102–1112 1103

of the bacterium against oxidative stress [17] While the bacterialenzymes are capable of binding H4B and this molecule supportsthe in vitro synthesis of NO [11,18], this pterin is not the cofactorof all bacterial NOSs as many bacteria that possess a NOS are notable to synthesize this cofactor. Another cofactor that is more com-mon in bacteria, tetrahydrofolate, binds to the pterin site of bacte-rial NOSs and allows the synthesis of NO [11,12].

NOSs carry out the synthesis of NO at the active site of the oxy-genase domain in two consecutive enzymatic steps. The first reac-tion consumes one molecule of oxygen, two electrons and onemolecule of L-arginine and forms xN-hydroxy-L-arginine (NOHA)and water. The second step converts NOHA to citrulline and NOand requires a second oxygen molecule and one electron [19,20].In both of these reactions, the oxygen molecule must bind to theheme and be activated [21–24]. In the first reaction, it was pro-posed that two electrons and protons are used to activate the oxy-gen molecule and form a Compound I species defined as an ferryl(FeIV = O) centre with an associated radical that could reside onthe porphyrin, the pterin or an amino acid side chain. This mecha-nism was however recently called into question [19]. In the secondstep, a compound I species may not be formed and different sce-narios have been put forward [25–30]. As in P450 enzymes, thethiolate ligand of NOS is thought to be important for oxygen acti-vation at the active site and hydroxylation of substrates but directevidence of this is scarce.

Active site residues are well conserved between bacterial andeukaryotic NOSs, particularly those that constitute the arginineand heme-binding sites as determined from the crystal structuresof the oxygenase domain of all three mammalian NOSs[29,31,32] and of several bacterial NOSs [10,13,33]. All NOSs pos-sess a conserved overall three-dimensional structure and all are di-meric. Conserved residues include the proximal cysteine ligand to

Fig. 1. Superposition of eNOSox (cyan) and saNOS structures (green). Hydrogenbonds are drawn between the indole nitrogen of Trp180 and the sulphur atom ofCys186 of eNOSox (3.33 Å) and between the indole nitrogen of Trp56 and thesulphur atom of Cys62 of saNOS (3.29 Å). Weak hydrogen bonds may form betweenthe backbone amide nitrogen of the Ile/Val, Gly, and Arg residues and the sulphuratom of the axial cysteine in both proteins. However these hydrogen bonds wouldbe rather long in eNOSox (3.51–3.78 Å except for the hydrogen bond involving thenitrogen amide of Gly at 3.24 Å in subunit A) and saNOS (3.41–3.68 Å except for thehydrogen bond involving the nitrogen amide of Gly at 3.21 in subunit B) (notdrawn). The structures were superposed with the Swiss-PDB viewer software(GlaxoSmithKline, v3.7) from the 3NOS (eNOSox/Arg/H4B) and 1MJT (saNOS/NAD/S-ethylisothiourea) PDB file coordinates. The image was prepared from the Asubunits with the PYMOL software (Delano Scientific LLC, USA, v0.99). Nitrogen(blue), oxygen (red), sulphur (yellow) and carbon atoms (green, cyan) are identified.(For interpretation of the references to colour in this figure legend, the reader isreferred to the web version of this article.)

the heme, and a tryptophan residue packed against the proximalface of the heme that hydrogen bonds to the cysteine ligand fromits indole ring nitrogen (Fig. 1). Mutations at this site, which re-move this hydrogen bond, may alter the electron density of the ax-ial cysteine in functionally-important ways. Such mutations thusoffer a means to probe the electron push provided by the axial cys-teine in relation to the electronic structure of heme–ligand com-plexes and to catalysis. Conserved mutations at this position innNOS (W409F, W409Y) that weaken or abolish the Trp–Cys hydro-gen bond have increased electron density on this axial ligand,resulting in a stronger Fe–cysteine bond and a weaker Fe–NO bondsince both NO and cysteine compete for Fe orbitals for r bond for-mation [34]. Surprisingly, the W409F and W409Y variants of nNOSwere able to synthesize NO and consume NADPH at faster ratesthan wild type nNOS under steady-state conditions [35,36]. A low-er rate of heme reduction, combined with a faster rate of oxidationof the FeIINO complex by O2 prevent the build-up of an inhibitoryFeIINO complex during catalysis and results in an increased rate ofNO synthesis [35–37]. In contrast, the W409H mutant of the sameprotein had reduced rates of NO synthesis and NADPH consump-tion which seems to result from less efficient electron transfer tothe heme; this suggests that W409 is important for mediating elec-tron transfer between the reductase and the oxygenase domains[38]. In contrast to these effects on nNOS, the corresponding muta-tions in iNOS (W188F, W188Y) result in heme-deficient, mono-meric, inactive protein, indicating that this tryptophan residuehas structural as well as catalytic roles [39]. Interestingly, the re-cent characterization of the W188H mutant of iNOSox indicatesthe formation of a heme-repleted enzyme with a strong His–Cyshydrogen bond that confers increased stability to the oxygenatedcomplex [40].

The effect of analogous mutations on the properties of themammalian endothelial NOS isotype has not been reported, norhave any such studies been done on the bacterial NOSs. It is impor-tant to characterize eNOS since it is less susceptible to NO inhibi-tion than nNOS and iNOS [41], and it may behave differentlyfrom these other isotypes when bound to NO. We also character-ized NOS of S. aureus (saNOS) since several liganded states havebeen reported [9,42], including the oxygenated complex [18,43],which can be used as a basis for comparison. The W56F andW56Y mutants of saNOS and the W180F and W180Y mutants ofeNOSox were thus investigated by resonance Raman spectroscopywith NO and CO as probes to evaluate how the extra electron den-sity introduced on the axial cysteine is shared with heme ligands.These results serve as a basis for future studies on the catalyticproperties of these mutants including the characterization of theoxygenated complex formed during catalysis.

2. Experimental procedures

2.1. Chemicals

L-arginine was obtained from Alexis Biochemicals (QBiogene,Carlsbad, CA, USA), H4B and NJYF were obtained from Sigma–Al-drich (Oakville, Canada), Sodium dithionite was purchased fromFluka (Sigma–Aldrich, Oakville, Canada), Argon and 12C16O werepurchased from Praxair (Mississauga, Canada), 14N16O was fromMEGS (Montreal, QC, Canada), 13C18O and 15N16O isotopes werefrom Icon Isotopes (Summit, NJ).

2.2. Mutagenesis

Site-directed mutagenesis of the saNOS coding sequence clonedin the pET30b expression plasmid (with a C-terminal hexahistidinetag) (Novagen, Madison, WI, USA) was performed using a protocol

1104 J. Lang et al. / Journal of Inorganic Biochemistry 103 (2009) 1102–1112

based on the QuikChange site-directed mutagenesis kit from Strat-agene [44]. Site-directed mutagenesis of the eNOSox coding se-quence cloned into the pET23 expression vector was performedwith the same protocol. The mutations were confirmed fromsequencing and no mutations other than that at codon 180 ofeNOSox protein sequence and at codon 56 of saNOS protein se-quence were found.

2.3. Protein expression and purification

The W56F and W56Y variants of saNOS were expressed andpurified in Escherichia coli cells exactly as described for wild typesaNOS [9]. In the absence of L-arginine, eNOSox variants W180Fand W180Y are prone to aggregation. Their purification was thusperformed with L-arginine present, and under these conditionsthe proteins were dimers as judged by size exclusion chromatogra-phy (not shown). The eNOSox wild-type, W180Y and W180F mu-tant proteins were cloned into a pET 23 expression vector(Novagen, Madison, WI) possessing a C-terminal His6 tag. The re-combinant proteins were expressed in E. coli BL21(DE3) cells. Thetransformed cells were grown in Terrific Broth (BioShop, Burling-ton, ON) supplemented with 100 lg/mL of ampicillin for 40 h. Cellpellets were resuspended in purification buffer (50 mM Tris buffer,pH 7.5, containing 0.5 M NaCl, 5 mM L-arginine, and 10% glycerol).Resuspended cells were lysed by mechanical grinding with 0.1 mmglass beads using the Bead Beater (Biospec, Bartlesville, OK) andthe cell lysate was centrifuged at 10,000g for 10 min at 4 �C to re-move the beads, and then clarified by centrifugation at 18,000g for20 min at 4 �C. Affinity chromatography was then carried out usinga column containing 2 mL of Ni-NTA Superflow resin (Qiagen,Mississauga, ON). The column was washed with four column vol-umes of purification buffer containing 15 mM imidazole. eNOSoxwas eluted in purification buffer containing 150 mM imidazole.The purified protein was desalted into purification buffer andstored at �80 �C. Protein concentration was determined from theUV-visible absorbance of the Soret band and an extinction coeffi-cient of 74,000 M�1cm�1 per heme for eNOSox [45]. SaNOS hemeconcentration was obtained by the pyridine hemochromogenmethod [46].

2.4. Sample preparation for spectroscopy

Proteins were put into a 40 mM HEPES pH 7.6 buffer containing40 mM NaCl using buffer exchange columns packed with P-6 DGresin (Bio-Rad Laboratories, CA). L-arginine (1 mM), H4B (1 mM)and NOHA (0.5 mM) were added to the protein samples asindicated.

2.5. Optical spectroscopy

Optical spectra were recorded with a Cary 3 spectrophotometerwith samples placed in a 1 cm-path-length anaerobic cuvette(Hellma, Müllheim, Germany). The FeII samples were prepared byequilibrating the sample solution with argon followed by reductionwith a small volume of a freshly prepared anaerobic sodium dithio-nite solution. The FeIICO complexes were obtained by equilibratingthe protein solution with argon, then adding a known volume ofgaseous CO into the cuvette, and reducing the heme with a smallvolume of a freshly prepared anaerobic solution of sodium dithio-nite. FeIIINO complexes were prepared by equilibrating samplesolutions with argon and then adding a known volume of gaseousNO that had been anaerobically bubbled through a solution ofNaOH and then water. FeIINO complexes were obtained by theaddition of a small amount of anaerobic dithionite to the FeIIINOcomplexes. The protein concentration used was 3–5 lM.

2.6. Resonance Raman spectroscopy

The equipment used to acquire resonance Raman spectra hasbeen previously described by Chartier et al. [9]. Briefly, the413.1 nm line of a Kr-ion laser (Innova 302 Kr laser, Coherent, San-ta Clara, CA) was used to probe the ferric and ferrous states. Theresonance Raman spectra of the FeIIINO and FeIICO complexes wereacquired with the 442 nm line of a He/Cd laser (Liconix laser,Melles-Griot, Canada). Several 1 min spectra were recorded atroom temperature with an excitation power ranging from 1.5–6 mW. The data collected over 30 min was averaged with Gramssoftware (ThermoGalactic, Salem, NH, USA). The spectrophotome-ter was calibrated with indene lines in the 200–1900 cm�1 region.Optical spectra of the samples were recorded prior to and after theresonance Raman experiment to verify their stability. The proteinconcentration in each sample was about 50 lM.

3. Results and discussion

3.1. Ferric forms of the saNOS W56 and eNOSox W180 mutants

In the presence of L-Arg, the optical spectra of the ferric state ofeNOSox and the W180 variants all possess a Soret optical transi-tion centered at 395 nm and a charge-transfer band (650 nm foreNOSox and 645 nm for the position-180 variants, not shown).These features are consistent with a 5-coordinate and high-spinheme as expected for the L-arginine-bound state of a NOS. Thiswas confirmed by obtaining the high-frequency region of the reso-nance Raman spectra. In the presence of L-Arg, W180Y and W180Fdisplayed frequencies of the m4 (1370 cm�1), m3 (1487 cm�1), m2

(1564 cm�1) and of the m10/mC–C(vinyl) (1624 cm�1) lines that arewithin 1–2 cm�1 of the values determined for wild type eNOSox/L-arginine (Fig. 2) and are consistent with a 5-coordinate high-spinheme [47–49]. Similar results were obtained with saNOS. With L-Arg, H4B, or both, the W56F and W56Y mutants of saNOS adopteda 5-coordinate high-spin state characterized by a Soret absorptionmaximum centered at 397 nm (not shown) and a single m3 line at1487 cm�1 in the high-frequency region of the resonance Ramanspectra (Fig. 3).

In both isotypes, an increased back-bonding from the Fe to theporphyrin as a result of tryptophan mutation is not expected toperturb Raman modes in the high-frequency region of the FeIII

complexes. Spiro and collaborators suggested that the resistanceof these modes to perturbation arises from the higher effective nu-clear charge of FeIII that can absorb the electron donation from thethiolate ligand without significant back-bonding to the porphyrin[50]. Therefore the similar m4 frequencies of the Trp mutants of sa-NOS and eNOSox relative to their corresponding wild type proteinsin the ferric state were not unexpected if the mutation altered onlythe basicity of the axial ligand.

3.2. Ferrous forms of the saNOS W56 and eNOSox W180 mutants

With eNOSox and position-180 variants, the resonance Ramanspectra of the reduced, L-arginine-bound proteins display a m3 lineat 1466 cm�1 and m4 frequencies of 1347 cm�1 (eNOSox andW180Y, Fig. 2) and 1345 cm�1 (W180F) indicating that a 5-coordi-nate and high-spin heme population was present in all of theseproteins. A second 6-coordinate low-spin heme fraction, likely cor-responding to the ferrous form of the P420 state detected with CO(vide infra), was revealed by the m4 line at 1360 cm�1 in bothW180F and W180Y. As judged from the optical spectra, which re-vealed a Soret absorption band centered at 412 nm (5-coordinate)and no band near 420 nm expected for a 6-coordinate state [51], itappears that the 6-coordinate state originates from a minority

Fig. 3. High-frequency region of the resonance Raman spectra of the position-56variants of saNOS in the ferric and reduced states. All spectra are those ofL-arginine-bound proteins. The inset shows the optical spectra of W56F (dashed)and W56Y (solid line) saturated with L-arginine.

Fig. 2. High-frequency region of the resonance Raman spectra of wild type eNOSoxand the position-180 variants in the ferric and reduced states. All spectra are thoseof L-arginine-bound proteins. The inset shows the optical spectra of wild typeeNOSox (solid line) and of the W180Y variant (dot line). Although the resonanceRaman spectra of FeII W180Y and W180F display a m4 line at 1360 cm�1 indicatingthat a 6-coordinate and low-spin state is present, the optical spectra of W180Y andW180F (not shown) are very similar to that of wild type eNOSox. These resultsindicate that the 6-coordinate state is a minority species.

J. Lang et al. / Journal of Inorganic Biochemistry 103 (2009) 1102–1112 1105

population in each of these variants (Fig. 2, inset). Not surprisingly,the m3 line of this minor fraction was not detected.

Similarly, the position-56 variants of L-Arg-bound saNOS, in thepresence or absence of H4B, displayed a majority of heme in the 5-coordinate and high-spin state with a m4 line at 1346 cm�1 and a m3

line at 1467 cm�1 (Fig. 3). The corresponding frequencies of wildtype saNOS were 1347 and 1467 cm�1, respectively (not shown).A small population of 6-coordinate heme was present as indicatedby the minor m4 line at 1359 cm�1. The optical spectra of L-argi-nine-bound saNOS mutants displayed a Soret transition at411 nm with an additional band near 555 nm consistent with a5-coordinate ferrous heme (Fig. 3 inset). Thus in both isotypesthe replacement of the tryptophan led to the formation of a smallfraction of 6-coordinate state. As the removal of the tryptophan–cysteine hydrogen bond may facilitate protonation of the axialcysteine, this 6-coordinate state likely has heme coordinated to aprotonated cysteine thus forming a thiol bond.

The influence of thiolate ligation on the heme electronic struc-ture from a resonance Raman study perspective was thoroughlyanalyzed by Spiro’s group, which serves as a basis for understand-ing the properties of the mutants that we studied [50]. The m4 fre-quency of a ferrous, 5-coordinate high-spin hemeprotein with anaxial thiolate ligand (near 1345 cm�1) is lower than that the m4 fre-quency of an histidine-coordinated heme protein (near 1357 cm�1)[50]. This difference was proposed to arise from increased back-donation of the FeIIdp electrons to the porphyrin e�g orbitals in thepresence of a strong electron donor axial ligand such as cysteinethiolate. The m4 frequencies of saNOS position W56 and eNOSox po-sition W180 mutants at 1346 cm�1 and 1345–1347 cm�1, respec-tively, are fully consistent with the axial thiolate ligand stillbeing bound to the heme in these mutants. The m4 frequency ofthe W56 mutants was actually 1 cm�1 lower than wild type saNOSand 2 cm�1 lower for the W180F mutant of eNOSox saturated withL-arginine (1347 cm�1 for wild type eNOSox and W180Y) (Fig. 3).This downshift, although small, is consistent with an increased ba-sicity of the axial thiolate ligand in W56 variants and the W180Fvariant. For the W180Y variant, it is possible that its identical m4

frequency with respect to wild type eNOSox reflects the formationof a tyrosine–cysteine bond (vide infra).

3.3. FeIICO complexes of eNOSox, saNOS and mutants

To examine whether mutation of the conserved tryptophan res-idue to phenylalanine or tyrosine leads to increased electron den-sity on the axial cysteine, and how such mutations might modifythe interactions that a heme–ligand complex would experiencewith substrate L-arginine, we prepared and analyzed NO and COcomplexes of the position-180 variants of eNOSox (W180F,W180Y) and the homologous position-56 variants of saNOS.

First, the optical and resonance Raman spectra of the FeIICOform of the L-arginine-bound W180F and W180Y variants of eNOS-ox were obtained. Both variants displayed a Soret band at 449 nmwhen bound to CO indicating that they formed complexes in theP450 state as observed in wild type eNOSox, and consistent withaxial cysteinate ligation to the heme (Fig. 4A). The wavelengthmaxima of the Soret optical transition are red shifted by 3 nm com-pared to wild type eNOSox, which displays a Soret optical transi-tion centered at 446 nm. A second Soret transition was detectedat 422 nm that represents a population of proteins in the P420state that has lost the thiolate bond between the Fe and the axialcysteine [51]. This population increased in the W180Y mutantsindicating that the mutations had a destabilizing effect on the thio-late bond to the heme iron (Fig. 4A). However, the relative propor-tion of the P420 state of FeIICO complexes prepared from L-arginine- and H4B-bound proteins was not higher in the W180variants than in wild type eNOSox (Fig. 5A). Interestingly, in the

Fig. 4. Optical spectra (A) and resonance Raman spectra in the low-frequencyregion (B) of FeIICO complexes of eNOSox and the W180F and W180Y variants. Allspectra were obtained with L-arginine saturated proteins. (A) Wild type eNOSox(dot line), W180F (dashed line) and W180Y (solid line). The table inset lists thewavelength maxima of the Soret bands obtained by band fitting analysis as well asthe percentage of each fraction as estimated from the area under the curve. (B) Theresonance Raman spectra were obtained with 12C16O (as indicated) and with 13C18O(not shown). The 12C16O minus 13C18O difference spectrum is that of wild typeeNOSox.

Fig. 5. Optical spectra (A) and resonance Raman spectra in the low-frequencyregion (B) of FeIICO complexes of eNOSox and the W180F and W180Y variants. Allspectra were obtained with L-arginine and H4B saturated proteins. (A) Wild typeeNOSox (dot line), W180F (dashed line) and W180Y (solid line). The table inset liststhe wavelength maxima obtained by band fitting analysis as well as well as thepercentage of each fraction as estimated from the area under the curve. (B) Theresonance Raman spectra were obtained with 12C16O. The 12C16O minus 13C18Odifference spectrum is that of wild type eNOSox.

1106 J. Lang et al. / Journal of Inorganic Biochemistry 103 (2009) 1102–1112

presence of both L-arginine and H4B, the Soret transition of theP450 state of the W180Y variant was centered at the same wave-length as wild type eNOSox, at 445 nm, while no such change oc-curred for the W180F mutant (Fig. 5A). The wavelengthmaximum of the Soret optical transition of FeIICO complexes of ahemeprotein model was previously shown to be related to the ba-sicity of the proximal heme axial ligand [52]. The replacement ofthe tryptophan at position-180 by phenylalanine is expected toincrease the basicity of the axial cysteine, which may not be

observed when the tryptophan is replaced with tyrosine as it maystill lead to the formation of a hydrogen bond between the sidechain hydroxyl group and the sulphur atom. That the W180Y vari-ant containing both L-arginine and H4B displayed the same wave-length maximum of its Soret band as wild type eNOSox under thesame conditions indicates that the basicity of the axial cysteinewas not altered significantly in contrast to the W180F variant.

CO is widely used to probe the environment of the active site ofhemeproteins by resonance Raman spectroscopy as it forms stable

Table 1Frequencies of the mFe–CO and dFe–C–O modes of wild type eNOSox and saNOS and theirrespective position 180 and 56 variants.

mFe–CO (13C18O), width(cm�1)

dFe–C–O (13C18O), width(cm�1)

Reference

eNOSox/Arg 503 (D13), 18.9513 (D14), 13.8

568 (D20), 10.3 This work

W180F/Arg 500 (D�10), 15.4 568 (D20), 10.8 This work508 (D�17), 14.7

W180Y/Arg 501 (D�7), 26.8 568 (D19), 10.6 This work508 (D�13), 16.8

eNOSox/Arg/H4B

504 (D�8), 18.9 567 (D18), 10.4 This work514 (D�18), 11.6

W180F/Arg/H4B

505, 17.7 568, 10.8 This work512, 11.7

W180Y/Arg/H4B

504, 16.8 568, 10.1 This work514, 12.0

saNOS/Arg/H4B

502 (D12), 16.1 565 (D18), 11.0 [9], thiswork

W56F/Arg/H4B

500, 18.6 565, 10.9 This work

W56Y/Arg/H4B

500, 18.1 565, 10.2 This work

Width refers to the width at half height of the Raman lines.

J. Lang et al. / Journal of Inorganic Biochemistry 103 (2009) 1102–1112 1107

complexes and it is sensitive to polar and steric interactions withsurrounding groups [53,54]. For FeIICO complexes, it is known thatp-back donation from the heme-Fe to the CO p* orbitals leads todecreased mC–O frequencies and increased mFe–CO frequencies [53].Modulation of p-back bonding arises principally from electrostaticinteractions involving the heme-bound CO and distal groups, butcould also be modulated by the basicity of the proximal ligand aswell as steric interactions. The proximal ligand influences mFe–CO

frequencies by virtue of r bonding with the heme-Fe as both thecysteine and CO compete for the formation of r bonds with themetal. It is for this the reason that thiolate-coordinated hemepro-teins fall on different a correlation line than histidine-coordinatedhemeproteins and 5-coordinate heme–CO complexes. Evidencethat the electron density of the axial cysteine was increased inthe reduced heme state was obtained with CO as the ligand ofthe W180F mutant of eNOSox.

With wild type L-arginine-bound eNOSox, isotopic substitutionallowed the identification of an isotope-sensitive mode at�510 cm�1 (496 cm�1 with 13C18O) in the mFe–CO region (Fig. 4B).A second isotope-sensitive mode corresponding to the Fe–C–Obending mode was localized at 568 cm�1 (548 cm�1 with 13C18O).These wavelengths are close to those determined for full-lengtheNOS [55]. Regression analysis of the �510 cm�1 line revealed thatit contains two mFe–CO modes centered at 503 cm�1 and 513 cm�1,respectively (Supplementary Fig. 1). For the W180 variants, the513 cm�1 mFe–CO mode was downshifted by 5 cm�1 relative to thewild type protein while the frequency of the dFe–C–O mode was un-changed (Fig. 4B and Supplementary Fig. 1). The frequencies of thesecond mFe–CO mode were determined at 500 cm�1 (W180F) and501 cm�1 (W180Y), thus 3–4 cm�1 lower than wild type eNOSox(Supplementary Fig. 1). In this study, eNOSox and its variants werepurified with L-arginine present to avoid the formation of high-molecular weight multimers, which precluded the determinationof the mFe–CO and dFe–C–O frequencies of L-arginine-free proteins.Nevertheless, it is well known that both mFe–CO and dFe–C–O of NOSare sensitive to L-arg binding [55]. That the observed downshiftof mFe–CO in eNOSox W180F and W180Y results from altered inter-actions with L-arginine can be ruled out based on the detection oftwo mFe–CO modes in the W180F mutant as in wild type eNOSox,and with similar respective proportions (the mode near 510 cm�1

is dominant W180F and W180Y as in wild type eNOSox). In addi-tion, the frequency and widths of the Fe–C–O bending mode is un-changed in all three proteins (Table 1). Instead, we propose that thedownshift in frequency arises from the electronic effects on theproximal cysteine caused by the tryptophan mutation. A lowermFe–CO frequency is expected if r competition by the proximal cys-teine is increased due to the loss of the tryptophan–cysteinehydrogen bond [56]. In contrast, a higher mFe–CO frequency couldarise if p back-bonding increased as a result of the mutation[56]. Our results indicate that r competition by the axial cysteineincreased, as the W180F and W180Y mutants of eNOSox display3–5 cm�1 lower mFe–CO frequencies than wild type eNOSox whenprepared with L-arginine alone. These shifts are of the same mag-nitude as that observed in Q360P/L mutants of P450cam that alsodisturb hydrogen bonding to the axial cysteine [56]. Nonethelessit is still possible that back-bonding was affected as well, but to aminor extent. If some of the electron density of the axial cysteinewas transferred to CO though back-bonding interactions, thisshould lead to a decrease of mC–O and an increase of mFe–CO, partiallycancelling the effect produced by r bonding on mFe–CO. As we couldnot detect the mC–O mode in the very high-frequency region of theresonance Raman spectra of eNOSox and the position-180 variants,we cannot directly probe modulation to p-back-bonding (notshown). However, the 3 nm red shift in the wavelength of the Soretband of W180F and W180Y relative to wild type eNOSox is consis-tent with an increased basicity of the axial ligand (Fig. 4A) [52].

In the presence of both L-arginine and H4B, two mFe–CO modeswere detected with eNOSox at frequencies similar to those ob-served with L-arginine alone (Fig. 5B and Supplementary Fig. 2).A 2 cm�1 downshift of the 514 cm�1 mFe–CO relative to wild typeeNOSox was observed with the W180F mutant but not theW180Y mutant (Fig. 5B and Supplementary Fig. 2). Band fittinganalysis of the W56F spectrum was performed with two lines inthe mFe–CO region for the purpose of comparison with the other pro-teins. Such analysis revealed that the second mFe–CO was at504 cm�1 as in wild type eNOSox; however the fit in the mFe–CO re-gion with only one line centered at 508 cm�1 was equally good,casting doubts as to the exact frequency of the mFe–CO mode near504 cm�1. Although eNOSox W180F displays a downshift in fre-quency of the mFe–CO mode at 514 cm�1 we lack conclusive evidencethat the mode at 504 cm�1 is also sensitive to this mutation. Incontrast, the W180Y mutant had a spectrum comparable to thatof wild type eNOSox with frequencies of both mFe–CO modes identi-cal to those of wild type eNOSox (Fig. 5B and SupplementaryFig. 2). In addition, the wavelength maximum of the Soret transi-tion band was identical to that of wild type eNOSox (Fig. 5A). Theseobservations demonstrate that in the presence of both compounds,the electron density on the proximal cysteine in eNOSox W180Ywas similar to that of wild type which suggests the formation ofa hydrogen bond between tyrosine 180 and the proximal ligand.H4B is crucial for the formation of the tyrosine–cysteine hydrogenbond as the W180Y prepared with L-arginine alone did not displaythis behavior. As the H4B binding site is remote from the tyrosine-180 (over 12 Å) and cysteine-184 (over 9 Å), a direct interactioninvolving H4B, tyrosine 180 and the proximal cysteine is not ex-pected. However, H4B may stabilize the dimeric structure andtighten the active site [21], which could be critical to the formationof a tyrosine–cysteine hydrogen bond.

Resonance Raman spectra revealed that heme distortion isinduced by H4B binding as previously observed with iNOSox [57].Indeed, the lines at 692, 750 and 800 cm�1 gained intensity inthe L-arg and H4B-bound proteins with respect to the proteins thatcontained only L-arg (Figs. 4B and 5B). The mode at 692 cm�1 is as-signed to c15 (B2u symmetry) based on the analysis of Li et al. [57]and indicates that heme saddling increased upon H4B binding toeNOSox and the W180 variants.

Similar studies of the bacterial saNOS protein were performed(Fig. 6). Regression analysis of the mFe–CO region revealed that the

1108 J. Lang et al. / Journal of Inorganic Biochemistry 103 (2009) 1102–1112

prominent isotope-sensitive line centered at 503 cm�1, with ashoulder (Fig. 6B), contained two lines centered at 503 cm�1 (ma-jor) and 514 cm�1 (minor) (Supplementary Fig. 3). The weak line at514 cm�1 could correspond either to a minor Fe–CO stretchingmode or another mode that is enhanced through Fermi resonancecoupling [58]. Isotope analysis indicates suggests the latter, as theminor shift observed with 13C18O (Supplementary Fig. 3) is toosmall to be accounted for by a mFe–CO mode and the difference spec-trum does not show any feature in this region consistent with iso-

Fig. 6. Optical spectra (A) and resonance Raman spectra in the low-frequencyregion (B) of FeIICO complexes of the W56F and W56Y variants of bacterial saNOS.All spectra were obtained with L-arginine and H4B saturated proteins. (A) Wild typesaNOS (solid line), W56F (dot line) and W56Y (dashed line). The table inset lists thewavelength maxima obtained by band fitting analysis as well as the percentage ofeach fraction as estimated from the area under the curve. (B) The resonance Ramanspectra were obtained with 12C16O. The 12C16O minus 13C18O difference spectrum isthat of wild type saNOS.

tope substitution (Fig. 6B). Thus saNOS displays a single mFe–CO

mode at 503 cm�1 that has a similar frequency to that of nNOSox[58], gsNOS [59], bsNOS [60] and one of the mFe–CO modes of eNOS-ox described above. Unlike the W180Y mutant of eNOSox, the fre-quency of the single mFe–CO mode of the W56Y mutant of saNOSwas identical to that of the W56F mutant (500 cm�1) and 2 cm�1

lower than wild type saNOS in the presence of L-arginine andH4B (Fig. 6B). The identical mFe–CO frequencies of W56F and W56Yindicate that their respective proximal cysteines have similar bas-icities and no hydrogen bond involving Y56 has formed on theproximal cysteine of this variant. This conclusion is corroboratedby the similar wavelengths of the Soret optical transitions of theW56F and W56Y mutants which are 1.5–2 nm higher than wildtype saNOS for the P450-type population (Fig. 6A). These resultsindicate that the heme environment differed slightly between sa-NOS with respect to eNOSox as the tyrosine of W56Y could notform a hydrogen bond to the axial cysteine as was observed witheNOSox.

In the low-frequency regions of the resonance Raman spectra,saNOS and its W56 mutants displayed heme deformation lines at690, 751 and 800/808 cm�1 (Fig. 6B) similar to those describedfor eNOSox and the W180 variants. The relative intensity of theselines that is identical in all three proteins indicates that the hemeadopts a similar structure in the W56 variants as in saNOS. Thevery high-frequency region of the resonance Raman spectrum ofW56 variants was obtained with 12C16O and 13C18O but the poorsignal to noise ratio did not allow the detection of the mC–O mode.

Regression analysis of the optical spectra (Fig. 6A) revealedthat the protein fraction in the P420 state was increased slightlywith the saNOS W56 variants indicating that the mutations per-turbed the stability of the thiolate bond. SaNOS W56 variants werecharacterized with substrate and H4B present. In fact, it is only inthese conditions that the P450 state of the saNOS variants at posi-tion 56 could be formed. When the substrate was absent, or withsamples containing only L-arginine, nearly all the heme complexwith CO was in the P420 state (not shown) in contrast to wild typesaNOS that is stable in these conditions [9].

3.4. Heme–NO interactions with distal molecules

NO is a sensitive probe of the electronic structure of the hemecomplexes with relation to the hydrogen bonding interactionsinvolving the proximal cysteine in the ferric state although resultsare less straightforward to interpret than those with CO due to thefrequent mixing of the mFe–NO and dFe–N–O mode of FeIIINO com-plexes [57,60,61]. The FeIIINO complexes of eNOSox and saNOSvariants display optical transitions expected for 6-coordinate andlow-spin heme complexes with a Soret transition at 437 nm(W180 variants) and 435–437 nm (W56 variants) and two addi-tional bands near 545 and 578 nm (insets of Figs. 7–9).

mFe–NO modes of thiolate-bound proteins typically display small15N isotopic shifts (64 cm�1) while dFe–N–O modes display muchlarger isotopic shifts (>13 cm�1). Although the frequencies of themFe–NO and dFe–N–O modes are close and the resonance Raman linesmay overlap [61], in substrate-free and pterin-free wild type sa-NOS these modes are well separated at 539 cm�1 (D4 cm�1 with15N16O) for the mFe–NO mode and at 550 cm�1 (�D16 cm�1 with15N16O) for the dFe–N–O mode [42]. Using isotope substitution, themFe–NO mode of substrate-free saNOS W56F and W56Y was de-tected at 531 cm�1, 8 cm�1 lower in frequency than the mFe–NO

mode of wild type saNOS (Fig. 7). This mode is characterized bya small isotope shift with 15N16O (3–4 cm�1) as expected for aFe–NO stretching mode (Supplementary Fig. 4 and Table 2). ThedFe–N–O mode was detected at 551 cm�1 in the difference spectraand by regression analysis (�533 cm�1 with 15N16O) which is2.5 cm�1 lower than wild type saNOS indicating that the dFe–N–O

Fig. 7. Low-frequency region of the resonance Raman spectra of the position-56variants of saNOS in the FeIIINO state. The spectra were obtained from substrate-free and pterin-free proteins with 14N16O (as indicated) and 15N16O (not shown).The difference spectra are those of the 14N16O minus 15N16O spectra. The insetshows the optical spectra of the FeIIINO complex of W56F (dashed line) and W56Y(solid line). The absorption maxima are centered at 435, 542 and 576 nm for W56Fand W56Y and 438, 548 and 579 nm for wild type saNOS (not shown).

Fig. 8. Low-frequency region of the resonance Raman spectra of the L-arginine-bound, FeIIINO complexes of the position-56 variants of saNOS and wild type saNOS.The spectra were obtained from L-arginine-bound proteins with 14N16O (asindicated) and 15N16O (not shown). The difference spectra are those of the 14N16Ominus 15N16O spectra. The inset shows the optical spectra of the L-arginine-bound,FeIIINO complexes of W56F (solid line) and W56Y (dashed line). The absorptionmaxima are centered at 437, 545 and 578 nm for W56F and W56Y and 441, 549 and586 nm for wild type saNOS (not shown).

Fig. 9. Low-frequency region of the resonance Raman spectra of the FeIIINOcomplexes of eNOSox and the position-180 variants. The spectra were obtainedfrom L-arginine-bound proteins and from L-arginine and H4B bound proteins with14N16O (as indicated) and 15N16O (not shown). The difference spectra are those ofthe 14N16O minus 15N16O spectra. The inset shows the optical spectra of the L-arginine- and H4B-bound, FeIIINO complexes of eNOsox (solid line), W180F (dashedline) and W180Y (dot line). The absorption maxima are centered at 441, 548 and580 nm for eNOSox and 437, 545 and 576 nm for W180F and W180Y.

J. Lang et al. / Journal of Inorganic Biochemistry 103 (2009) 1102–1112 1109

mode was less sensitive to the W56 mutation than the mFe–NO mode(Supplementary Fig. 4 and Table 2). The strength of the r bond

between the proximal cysteine and iron is expected to increaseupon the removal of the hydrogen bond to this ligand, while thecapacity of NO to form a strong r bond with the Fe should decreaseand thus lower the mFe–NO frequencies [62]. The unambiguousdetection of the mFe–NO mode of substrate-free W56F and W56Y,well separated from the dFe–N–O mode, reveals that the position-56 mutations weakened the Fe–NO bond which is reflected inthe lower mFe–NO frequency (Table 2).

The FeIIINO spectra were next obtained with substrate presentto determine if L-arginine–NO interactions were maintained inthe W56 variants. With L-arginine present, wild type saNOS dis-plays an isotope-sensitive band at 541 cm�1 with contributionsfrom the Fe–N–O bending mode and the Fe–NO stretching mode,which is characterized by an isotope shift intermediate to thoseof the mFe–NO and dFe–N–O modes (Fig. 8, Supplementary Fig. 5 andTable 2) [42]. Regression analysis revealed that a weak line near524 cm�1 also displays isotope sensitivity. Although its frequencyis in the region observed for the mFe–NO mode of substrate-boundP450cam [61], the magnitude of the isotopic shift is too large to as-sign the line to a Fe–NO stretching mode. Mode mixing is sus-pected based on changes in the shape and width of the band asits frequency changes due to isotope substitution or W56 mutation(Supplementary Figs. 5–7), and formal assignment of this band re-quires further studies.

An isotope-sensitive line was detected at 537 cm�1 for L-argi-nine-bound and for L-arginine and H4B-bound W56F and W56Y(Fig. 8, Supplementary Figs. 5 and 6 and Table 2). The magnitudeof the isotope shift was intermediate to those of the mFe–NO anddFe–N–O modes and was similar to that of the mFe–NO/dFe–N–O modeof wild type saNOS (8–9 cm�1); consequently this line is assignedto a mixed mFe–NO/dFe–N–O mode for W56F and W56Y (Table 2). Asecond weak isotope-sensitive line detected in wild type saNOS(near 524 cm�1) is also present in the spectra of the W56 variants

Table 2Frequencies of the mFe–NO and dFe–N–O modes of saNOS and eNOSox mutants.

mFe–NO (15N16O), width(cm�1)

mFe–NO/dFe–N–O (15N16O),width (cm�1)

Reference

saNOS 539 (D3), 18.8 553 (�D16), 8.5a [42], thiswork

W56F 531 (D3), 18.4 551 (�D18), 8.3a This workW56Y 531 (D4), 19.5 551 (�D18), 11.5a This worksaNOS/Arg – 541 (D7), 16.0 [42]W56F/Arg – 537 (D8), 15.4 This workW56Y/Arg – 537 (D9), 15.1 This worksaNOS/Arg/

H4B– 542 (D8), 14.6 [42]

W56F/Arg/H4B

– 538 (ND), 15.0 This work

W56Y/Arg/H4B

– 537 (ND), 15.0 This work

eNOSox/Arg 530 (D8), 12.8 549 (D 11), 13.5a This workW180F/Arg – 539 (D6), 19.1 This workW180Y/Arg – 539 (D6), 18.5 This workeNOSox/

Arg/H4B529 (D6), 12.4 548 (D8), 12.6a This work

W180F/Arg/H4B

– 540 (D7), 17.4 This work

W180Y/Arg/H4B

– 540 (D6), 17.9 This work

ND: not determined.–: Absent or not detected.Width refers to the width at half height of the Raman lines.

a Substrate-free and pterin-free saNOS and W56 variants display two, well sep-arated isotope-sensitive lines. The line near 550 cm�1 which displays a very largeisotope shift correspond to a dFe–N–O mode with no or minimal contributions fromthe mFe–NO mode. The isotopic shift was estimated from the 14N16O minus 15N16Odifference spectrum. Similarly, L-arginine bound and L-arginine and H4B bound wildtype eNOSox display well separated mFe–NO and dFe–N–O modes.

2 Evidence that some L-arginine was bound to wild type eNOSox in the FeIIINOate was obtained from a sample passed through a small desalting column to remove

xcess L-arginine. This sample displayed the same Raman lines at 539 cm�1 and49 cm�1 as an eNOSox sample prepared with 1 mM L-arginine except that the line at39 cm�1 (mFe-NO) was more intense and the line at 549 cm�1 (dFe-N-O) was lesstense (Supplementary Fig. 9).

1110 J. Lang et al. / Journal of Inorganic Biochemistry 103 (2009) 1102–1112

but at a frequency 5 cm�1 lower, indicating that this line is sensi-tive to the W56 mutation in addition to isotope substitution (Sup-plementary Figs. 5 and 6). Overall, we observed that L-arginineinduces a shift from the detection of well separated mFe–NO anddFe–N–O modes to the detection of a mixed mFe–NO/dFe–N–O mode inwild type saNOS and in W56 variants of saNOS. In addition to theseL-arginine-induced changes, we observed that the magnitude ofthe isotope shift of mFe–NO/dFe–NO (7–9 cm�1) is similar in wild typesaNOS and the two variants (Table 2). However, the frequency shiftin going from the mFe–NO mode to the mFeNO/dFe–N–O mode did not oc-cur to the same extent. While L-arginine binding induces a 2 cm�1

upshift in frequency in wild type saNOS, a 6 cm�1 upshift occurswith both of the position-56 variants (Table 2). The relatively lar-ger change in frequency of the mFe–NO (D8 cm�1) compared to thedFe–N–O mode (D2.5 cm�1) of the substrate-free proteins likely ex-plains the larger shift upon L-arginine binding with the W56 vari-ants, which involves a change in mode composition from a mFe–NO

to a mFe–NO/dFe–N–O mode. We therefore conclude that the sub-strate–NO interactions in W56 variants of saNOS are similar tothose of wild type saNOS. As both W56F and W56Y exhibit thesame behavior, there is no evidence that tyrosine-56 of theW56Y mutant of saNOS formed a hydrogen bond with the proxi-mal cysteine. This is consistent with the results with FeIICO com-plexes that also showed that saNOS W56Y did not form such ahydrogen bond.

L-arginine binding also induces several other modifications oflow-frequency modes which reflect increased heme distortion pre-viously observed with wild type saNOS [42]. The assignment of theheme deformation modes is based on the analysis by Li et al. withiNOSox [42]. L-arginine-bound W56F and W56Y saNOS variantsdisplay strong lines at 684 cm�1 (c15), 708 cm�1 (c11), 728 cm�1

(c5), 743 cm�1 (c1) (Fig. 8). These lines, which are present but withmuch lower intensity in substrate-free and pterin-free proteins(Fig. 7), belong to various symmetries: c15 has B2u symmetry

and indicates that the heme is saddled; c5 has A2u symmetry andindicates that the heme is domed; c11 has B1u symmetry whichindicates heme ruffling and c1 has A1u symmetry which indicatespropeller deformation. In addition, L-arginine binding increasedthe intensity of a heme propionate mode at 394 cm�1. Finally, aweak line of unknown origin at 717 cm�1 disappeared upon L-argi-nine binding. All these changes were observed upon L-arginine-binding in wild type saNOS and W56F and W56Y which indicatedsimilar structural changes to the heme in all three cases. The onlydifference observed was that the 684 cm�1 line was more intensein L-arginine-bound W56 variants than in wild type saNOS(Fig. 9) indicating that the heme was slightly more saddled in theW56 variants.

The FeIIINO complexes of eNOSox and position-180 variantswere analyzed by resonance Raman spectroscopy to look at L-argi-nine and NO interactions at the active site. L-arginine was includedas the variants formed high molecular weight complexes in its ab-sence. In the low-frequency region, isotopic substitution was usedwith L-arginine-saturated wild type eNOSox to determine the fre-quencies of Fe–N–O modes. As shown in Fig. 9, an isotope-sensitiveline was detected near 542 cm�1 (536 cm�1 with 15N16O). Regres-sion analysis revealed two lines centered at 549 cm�1 and539 cm�1 in the 14N16O spectrum (Supplementary Fig. 7). Theselines, which are merged in the 15N16O spectrum at 537 cm�1, wereused to calculate isotopic shifts of 11.4 and 1.3 cm�1, respectively.The magnitude of the isotopic shift of the 549 cm�1 line (11 cm�1)is high, and close to that displayed by the dFe–N–O mode of sub-strate-free saNOS [42]. We therefore attribute the 549 cm�1 lineof eNOSox to the Fe–N–O bending mode (Table 2). The line at539 cm�1, which displays a very small isotope shift of 1–2 cm�1,is assigned to the Fe–NO stretching mode. Thus eNOSox displayswell separated mFe–NO and dFe–N–O modes when bound to L-argi-nine2. Similarly, L-arginine and H4B-bound eNOSox displays wellseparated mFe–NO and dFe–N–O modes at similar frequencies (Fig. 9,Supplementary Fig. 8 and Table 2). In this respect wild type eNOSoxis distinct from saNOS, iNOSox and nNOSox which all display amixed mFe–NO/dFe–N–O mode when bound to L-arginine and H4B[34,42,57]. In contrast, in the L-arginine bound states (with and with-out H4B) of the position-180 variants of eNOSox, a single isotope-sensitive line was observed which corresponds to a mixed mFe–NO/dFe–N–O mode, based on the amplitude of the isotope shift (6 cm�1)(Fig. 9, Supplementary Figs. 6 and 7 and Table 2). Thus the W180Fand W180Y eNOSox variants more closely resemble other NOSs iso-types, in this respect, rather than wild type eNOSox. These resultsclearly show that the substrate–NO interactions are altered in theW180 variants.

Heme deformation modes were observed in the low-frequencyregion of W180F, W180Y and wild type eNOSox as described abovefor saNOS and position-56 variants. Lines were observed at684 cm�1 (assigned to c15, B2u symmetry, indicates heme saddling),708 cm�1 (assigned to c11, B1u symmetry, indicates heme ruffling),729 cm�1 (assigned to c5, A2u symmetry, indicates heme doming)and 743 cm�1 (assigned to c1, A1u symmetry, indicates propellerdeformation) in eNOSox and position-180 variants. The intensityof these lines was similar in all three proteins with the exceptionof the line at 684 cm�1; this was more intense in the W180F andW180Y variants and it indicates that heme saddling is increasedin these variants as was observed for the W56 variants of saNOS.The altered L-arginine–NO interactions detected in the W180 vari-

ste55in

J. Lang et al. / Journal of Inorganic Biochemistry 103 (2009) 1102–1112 1111

ants with respect to eNOSox and the lack of a tyrosine–cysteinehydrogen bond in FeIIINO complexes of W180Y contrasts to obser-vations made on the FeIICO complex and reveal that this mutationhas a more pronounced effect when the heme is bound to NOrather than CO.

It has been suggested that the tryptophan–cysteine hydrogenbond may be important in the process of auto-inhibition by NOas loss of this interaction weakens the mFe–NO bond of nNOS, andW409 mutants of nNOS were less susceptible to NO inhibition[35–37]. On the other hand, it is also known that eNOS is less proneto autoinhibition by NO than nNOS and iNOS. We note that eNOSoxis distinct from nNOS and iNOS in that well separated mFe–NO anddFe–N–O modes are present for eNOSox saturated with L-arginineand H4B rather than a mixed mFe–NO/dFe–N–O mode in nNOSox andiNOSox [34], which suggests that the conformation of the Fe–NOcomplex, or the mode of L-arginine binding, or both, differ in eNOSrelative to the other two mammalian isoforms. However, wenote that the W180F and W180Y mutants display a singlemFe–NO/dFe–N–O mode at the same frequency as that of the W409Fmutant of nNOSox, which suggests that the W180 hydrogen bondto the proximal cysteine confers unique properties to the FeIIINOcomplex of wild type eNOSox. It is possible that the unique prop-erties conferred by the tryptophan–cysteine hydrogen bond ofeNOSox may help modulate the extent of autoinhibition of theenzymatic activity by NO, in addition to its contributions to therate of heme reduction, the affinity for NO and the reactivity ofthe FeIINO complex toward oxygen [35–37]. All our results indicatethat the analogous mutants of saNOS and eNOSox behave similarlythus indicating that tyrosine is not a functional replacement oftryptophan in these proteins when bound to NO, as observed alsowith nNOSox [34].

Ferrous NO complexes of the position-180 variants of eNOSoxand position-56 variants of saNOS were unstable and displayedSoret transitions in the 400–408 nm range, indicating that theydid not form the 6-coordinate low-spin heme complex like wildtype eNOSox (Soret centered at 440 nm) and wild type saNOS (Sor-et centered at 440 nm) when prepared in the presence of L-arginine(not shown). This behavior is similar to that of the W409F andW409Y variants of nNOSox [34]. However, none of the W180 vari-ants of eNOSox and of the W56 variants of saNOs formed the 6-coordinate low-spin state when prepared at pH 9.5 in contrast tothe W409Y variant of nNOSox that formed a 6-coordinate FeIINOcomplex at high pH similar to that of the wild type protein, whichsuggested that at neutral pH, the proximal cysteine was protonatedand became deprotonated with the help of Y409 at pH 9.5 [34]. Ascomparison between the mutants and their respective wild typeproteins was not possible, the FeIINO complexes of the W56 andW180 variants were not characterized further by resonance Ramanspectroscopy.

3.5. Structural comparison between the mammalian and bacterialNOSs

Our results show that the hydrogen bond formed betweenW180 and the proximal cysteine of eNOSox and between W56and the proximal cysteine of saNOS, as previously shown withnNOSox [34] is important for the stability of the Fe–S bond inthe ferrous, ferrous CO and ferrous–NO bound states and is impor-tant for the fine tuning of substrate–NO interactions in FeIIINOcomplexes of eNOSox and the electronic structure of these com-plexes in saNOS. Comparison of Trp variants of other NOSs givesus a better idea of specific structural roles played by the Trp–cys-teine hydrogen bond of mammalian and bacterial NOSs isoforms.First, the iNOS W188F variant is unable to incorporate heme whenexpressed in E. coli and is predominantly isolated in a heme-freestate [39]. This is not seen in the Trp to Phe variants of other mam-

malian or bacterial NOSs, but it was observed with a W409L mu-tant of nNOSox, possibly as a result of decreased hemestabilization by p–p interactions [38]. This suggests that the inabil-ity to incorporate heme in the Trp variants is not specific to iNOS,but rather that iNOS is more sensitive to the mutation than otherNOSs. As iNOS forms the weakest dimer of the mammalian NOSs,mutation to this active site residue may have greater repercussionon this isoform.

Second, a comparison of the Trp–Cys hydrogen bond distancefrom crystallographic data shows similar bond lengths for all NOSs,between 3.3 Å and 3.6 Å for mammalian NOSs [29,32,63,64] andbetween 3.3 Å and 3.8 Å for bacterial NOSs [10,13], which suggestsa similar hydrogen bond strength across all NOSs. This is in agree-ment with the similar downshift in frequency caused by the Trpmutation on the mFe–NO/dFe–N–O mode of W56 variants of saNOS(7–8 cm�1, with L-arg and H4B) and of W409 variants of nNOSox(6–8 cm�1, with L-arg and H4B) [34]. The similar shifts of themFe–NO mode of substrate-free and pterin-free variants of saNOS(8 cm�1) and nNOSox (6–8 cm�1) [34] with respect to their wildtype counterparts are also in agreement with a tryptophan–cys-teine hydrogen bond of similar strength in these NOSs.

On the other hand, the W56 and W180 variants revealed someinteresting structural differences between the eNOS and nNOS iso-forms. While the eNOSox W180Y variant is able to form a Tyr–Cyshydrogen bond when saturated with L-arginine and H4B in the FeII-

CO state, it is not able to do so in the FeIINO state as the analogousnNOS W409Y variant studied by Couture et al. [34]. Although thestrength of the Trp–cysteine hydrogen bond may be similar in allNOS isoforms, small structural features unique to each isoform ap-pear to confer properties specific to some of them when tyrosinereplaces tryptophan.

3.6. Functional relevance

Our results indicate that the Trp–cysteine hydrogen bond ofeNOSox and saNOS has an important structural role, as unstableFeIICO complexes were formed in W56 variants of saNOS whensubstrate and H4B were absent, and FeIIINO complexes of W180variants of eNOSox displayed altered substrate–NO interactions.However, when FeIICO and FeIIINO complexes structurally similarto those of the wild type proteins were formed, our results are con-sistent with modulation of r bonding in FeIICO complexes ofW180F and W56F variants of eNOSox and saNOS, respectively,and in FeIIINO complexes of W56 variants of saNOS. Indeed, theFe–NO and Fe–CO bonds were weakened as the basicity of the axialligand increased as a result of the mutations. In this sense, theW180 and W56 mutants resemble the Q360P mutant of cyto-chrome P450cam, designed to disrupt the proximal hydrogen bondnetwork, which displays lower mFe–CO frequencies as a result of in-creased r donation by the thiolate [56]. The catalytic activity of theQ360P mutant was similar to that of the wild type enzyme whichled to the conclusion that increased r donation by the axial thio-late does not promote heterolysis of the O–O bond during catalysis.In contrast the L358P mutant of cytochrome P450cam, also de-signed to disrupt the proximal hydrogen bond network, had in-creased p-back-bonding from the thiolate as a result of themutation and displayed enhanced rate of heterolysis [65]. Themechanism of NO synthesis differs from oxygenation reactions cat-alysed by P450 in a number of ways and may or may not implicateheterolysis of the O–O bond to catalyze the oxygenation of L-argi-nine and NOHA. It thus remains to be determined if the Trp–Cyshydrogen bond, which clearly modulates r bonding, is importantfor oxygen activation in NOSs. We have recently begun the enzy-matic characterization of our NOS variants to understand howcatalysis is affected by the Trp mutations, and preliminary results

1112 J. Lang et al. / Journal of Inorganic Biochemistry 103 (2009) 1102–1112

indicate that the saNOS variants are catalytically active. These vari-ants thus offer the possibility of determining how the increased ba-sicity of the axial cysteine, which affects r bonding, affects theelectronic properties of the oxygenated complex and impactscatalysis in NOS.

4. Conclusions

We have shown that thiolate-coordination is maintained in theferric and ferrous states of W56 and W180 variants of saNOS andeNOSox, respectively. Furthermore, we demonstrated that interac-tions with substrates are not affected by these mutations (with theexception of the FeIIINO complexes of eNOSox W180 variants),while modulation of heme–ligand complexes occurs and involveschanges in r bonding with the axial cysteine in NO and CO com-plexes. Additionally, we revealed that a direct correlation exists be-tween the wavelength of the Soret optical transition and thefrequency of the Fe–CO stretching mode as a result of modulationof the basicity of the axial cysteine. These studies serve as a basis todetermine how mutations to the tryptophan–cysteine interactionsimpact the catalytic steps in NOSs thus offering clues regardingdriving forces for the strict conservation of this tryptophan inNOSs.

5. Abbreviations

NOS nitric oxide synthaseeNOSox, nNOSox and iNOSoxm oxygenase domain of endothelial,

neuronal ad inducible nitric oxide synthases, respectivelysaNOS nitric oxide synthase-like protein of S. aureusNOHA Nx-hydroxy-L-arginineH4B (6R) 5,6,7,8 tetrahydro-L-biopterinWT wild type

Acknowledgements

This work was supported by NSERC discovery Grant 250073 andCFI Grant 7128 to M.C. and NSERC Discovery Grant 216849-03 andCFI grant 2318 to S.P. Rafferty.

Appendix A. Supplementary material

Supplementary data associated with this article can be found, inthe online version, at doi:10.1016/j.jinorgbio.2009.05.011.

References

[1] R. Bruckdorfer, Mol. Aspects Med. 26 (2005) 3–31.[2] F.X. Guix, I. Uribesalgo, M. Coma, F.J. Munoz, Prog. Neurobiol. 76 (2005) 126–

152.[3] S. Moncada, A. Higgs, New Engl. J. Med. 329 (1993) 2002–2012.[4] S. Moncada, E.A. Higgs, Br. J. Pharmacol. 147 (Suppl. 1) (2006) S193–S201.[5] C.S. Raman, P. Martasek, B.S.S. Masters, in: K.M. Kadish, K.M. Smith, R. Guilard

(Eds.), Structural Themes Determining Function in Nitric Oxide Synthases,Academic Press, New York, 2000.

[6] H. Li, T.L. Poulos, J. Inorg. Biochem. 99 (2005) 293–305.[7] R.C. Venema, H. Ju, R. Zou, J.W. Ryan, V.J. Venema, J. Biol. Chem. 272 (1997)

1276–1282.[8] B. Hemmens, A.C. Gorren, K. Schmidt, E.R. Werner, B. Mayer, Biochem. J. 332

(1998) 337–342.[9] F.J. Chartier, M. Couture, Biophys. J. 87 (2004) 1939–1950.

[10] L.E. Bird, J. Ren, J. Zhang, N. Foxwell, A.R. Hawkins, I.G. Charles, D.K. Stammers,Structure 10 (2002) 1687–1696.

[11] S. Adak, K.S. Aulak, D.J. Stuehr, J. Biol. Chem. 277 (2002) 16167–16171.[12] S. Adak, A.M. Bilwes, K. Panda, D. Hosfield, K.S. Aulak, J.F. McDonald, J.A. Tainer,

E.D. Getzoff, B.R. Crane, D.J. Stuehr, Proc. Natl. Acad. Sci. USA 99 (2002) 107–112.[13] J. Sudhamsu, B.R. Crane, J. Biol. Chem. 281 (2006) 9623–9632.

[14] J.A. Kers, M.J. Wach, S.B. Krasnoff, J. Widom, K.D. Cameron, R.A. Bukhalid, D.M.Gibson, B.R. Crane, R. Loria, Nature 429 (2004) 79–82.

[15] E.G. Johnson, J.P. Sparks, B. Dzikovski, B.R. Crane, D.M. Gibson, R. Loria, Chem.Biol. 15 (2008) 43–50.

[16] F. Hochgrafe, C. Wolf, S. Fuchs, M. Liebeke, M. Lalk, S. Engelmann, M. Hecker, J.Bacteriol. 190 (2008) 4997–5008.

[17] K. Shatalin, I. Gusarov, E. Avetissova, Y. Shatalina, L.E. McQuade, S.J. Lippard, E.Nudler, Proc. Natl. Acad. Sci. USA 105 (2008) 1009–1013.

[18] F.J. Chartier, S.P. Blais, M. Couture, J. Biol. Chem. 281 (2006) 9953–9962.[19] Y. Zhu, R.B. Silverman, Biochemistry 47 (2008) 2231–2243.[20] A.C. Gorren, B. Mayer, Biochim. Biophys. Acta 1770 (2007) 432–445.[21] W.K. Alderton, C.E. Cooper, R.G. Knowles, Biochem. J. 357 (2001) 593–615.[22] O.W. Griffith, D.J. Stuehr, Annu. Rev. Physiol. 57 (1995) 707–736.[23] J.T. Groves, C.C. Wang, Curr. Opin. Chem. Biol. 4 (2000) 687–695.[24] K.B. Cho, E. Derat, S. Shaik, J. Am. Chem. Soc. 129 (2007) 3182–3188.[25] B.R. Crane, A.S. Arvai, S. Ghosh, E.D. Getzoff, D.J. Stuehr, J.A. Tainer,

Biochemistry 39 (2000) 4608–4621.[26] H. Huang, J.M. Hah, R.B. Silverman, J. Am. Chem. Soc. 123 (2001) 2674–2676.[27] K.-B. Cho, J.W. Gauld, J. Am. Chem. Soc. 126 (2004) 10267–10270.[28] K.-B. Cho, J.W. Gauld, J. Phys. Chem. B 109 (2005) 23706–23714.[29] H. Li, J. Igarashi, J. Jamal, W. Yang, T.L. Poulos, J. Biol. Inorg. Chem. 11 (2006)

753–768.[30] D.L. Tierney, H. Huang, P. Martasek, B.S. Masters, R.B. Silverman, B.M. Hoffman,

Biochemistry 38 (1999) 3704–3710.[31] H. Li, C.S. Raman, C.B. Glaser, E. Blasko, T.A. Young, J.F. Parkinson, M. Whitlow,

T.L. Poulos, J. Biol. Chem. 274 (1999) 21276–21284.[32] B.R. Crane, A.S. Arvai, D.K. Ghosh, C. Wu, E.D. Getzoff, D.J. Stuehr, J.A. Tainer,

Science 279 (1998) 2121–2126.[33] K. Pant, A.M. Bilwes, S. Adak, D.J. Stuehr, B.R. Crane, Biochemistry 41 (2002)

11071–11079.[34] M. Couture, S. Adak, D.J. Stuehr, D.L. Rousseau, J. Biol. Chem. 276 (2001)

38280–38288.[35] S. Adak, D.J. Stuehr, J. Inorg. Biochem. 83 (2001) 301–308.[36] S. Adak, C. Crooks, Q. Wang, B.R. Crane, J.A. Tainer, E.D. Getzoff, D.J. Stuehr, J.

Biol. Chem. 274 (1999) 26907–26911.[37] S. Adak, Q. Wang, D.J. Stuehr, J. Biol. Chem. 275 (2000) 17434–17439.[38] T. Yumoto, I. Sagami, S. Daff, T. Shimizu, J. Inorg. Biochem. 82 (2000) 163–170.[39] D.J. Wilson, S.P. Rafferty, Biochem. Biophys. Res. Commun. 287 (2001) 126–

129.[40] J. Tejero, A. Biswas, Z.Q. Wang, R.C. Page, M.M. Haque, C. Hemann, J.L. Zweier, S.

Misra, D.J. Stuehr, J. Biol. Chem. 283 (2008) 33498–33507.[41] J. Santolini, A.L. Meade, D.J. Stuehr, J. Biol. Chem. 276 (2001) 48887–

48898.[42] F.J.M. Chartier, M. Couture, Biochem. J. 401 (2007) 235–245.[43] F.J. Chartier, M. Couture, J. Biol. Chem. 282 (2007) 20877–20886.[44] L. Zheng, U. Baumann, J.L. Reymond, Nucleic Acids Res. 32 (2004) e115.[45] D.J. Stuehr, M. Ikeda-Saito, J. Biol. Chem. 267 (1992) 20547–20550.[46] C.A. Appleby, Methods Enzymol. 52 (1978) 157–166.[47] T.G. Spiro, X.Y. Li, in: T.G. Spiro (Ed.), Resonance Raman Spectroscopy of

Metalloproteins, Wiley, New York, 1988.[48] J. Wang, W.S. Caughey, D.L. Rousseau, in: M. Feelish, J.S. Stamler (Eds.),

Resonance Raman Scattering: A Probe of Heme Protein-Bound Nitric Oxide,John Wiley & Sons, New York, 1996.

[49] S. Hu, K.M. Smith, T.G. Spiro, J. Am. Chem. Soc. 118 (1996) 12638–12646.[50] P. Anzenbacher, R. Evangelista-Kirkup, J. Schenkman, T.G. Spiro, Inorg. Chem.

28 (1989) 4491–4495.[51] J. Wang, D.J. Stuehr, D.L. Rousseau, Biochemistry 34 (1995) 7080–7087.[52] N. Suzuki, T. Higuchi, Y. Urano, K. Kikuchi, H. Uekusa, Y. Ohashi, T. Uchida, T.

Kitagawa, T. Nagano, J. Am. Chem. Soc. 121 (1999) 11571–11572.[53] T.G. Spiro, I.H. Wasbotten, J. Inorg. Biochem. 99 (2005) 34–44.[54] G.N.J. Phillips, M.L. Teodoro, T. Li, B. Smith, J.S. Olson, J. Phys. Chem. B 103

(1999) 8817–8829.[55] B. Fan, J. Wang, D.J. Stuehr, D.L. Rousseau, Biochemistry 36 (1997) 12660–

12665.[56] S. Yoshioka, T. Tosha, S. Takahashi, K. Ishimori, H. Hori, I. Morishima, J. Am.

Chem. Soc. 124 (2002) 14571–14579.[57] D. Li, D.J. Stuehr, S.R. Yeh, D.L. Rousseau, J. Biol. Chem. 279 (2004) 25489–

25499.[58] J. Wang, D.J. Stuehr, D.L. Rousseau, Biochemistry 36 (1997) 4595–4606.[59] M. Kabir, J. Sudhamsu, B.R. Crane, S.R. Yeh, D.L. Rousseau, Biochemistry 47

(2008) 12389–12397.[60] J. Santolini, M. Roman, D.J. Stuehr, T.A. Mattioli, Biochemistry 45 (2006) 1480–

1489.[61] S. Hu, J.R. Kincaid, J. Am. Chem. Soc. 113 (1991) 2843–2850.[62] M.L. Fernandez, M.A. Marti, A. Crespo, D.A. Estrin, J. Biol. Inorg. Chem. 10

(2005) 595–604.[63] T.O. Fischmann, A. Hruza, X.D. Niu, J.D. Fossetta, C.A. Lunn, E. Dolphin, A.J.

Prongay, P. Reichert, D.J. Lundell, S.K. Narula, P.C. Weber, Nat. Struct. Biol. 6(1999) 233–242.

[64] C.S. Raman, H. Li, P. Martasek, V. Kral, B.S. Masters, T.L. Poulos, Cell 95 (1998)939–950.

[65] S. Yoshioka, S. Takahashi, K. Ishimori, I. Morishima, J. Inorg. Biochem. 81 (2000)141–151.