Native alanine substitution in the glycine hinge modulates

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Native alanine substitution in the glycine hinge modulates conformational flexibility of Heme Nitric oxide/Oxygen (H-NOX) sensing proteins Charles W Hespen, Joel J Bruegger, Yirui Guo, and Michael A Marletta ACS Chem. Biol., Just Accepted Manuscript • DOI: 10.1021/acschembio.8b00248 • Publication Date (Web): 14 May 2018 Downloaded from http://pubs.acs.org on May 15, 2018

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ACS Chemical Biology

Native alanine substitution in the glycine hinge modulates conformational flexibility of Heme Nitric oxide/Oxygen (H-NOX) sensing proteins Authors Charles W. Hespen1*, Joel J. Bruegger1*, Yirui Guo1, Michael A. Marletta1,2 1. QB3 Institute, University of California–Berkeley, 356 Stanley Hall, Berkeley, CA, 94720-3220, United States 2. Department of Chemistry, Department of Molecular and Cell Biology, QB3 Institute, 374B Stanley Hall, University of California–Berkeley, CA, 94720-3220, United States *Authors contributed equally to this work. Correspondence: Michael A. Marletta: [email protected] Abstract Heme nitric oxide/oxygen sensing (H-NOX) domains are direct NO sensors that regulate a variety of biological functions in both bacteria and eukaryotes. Previous work on H-NOX proteins has shown that upon NO binding, a conformational change occurs along two glycine residues on adjacent helices (termed the glycine hinge). Despite the apparent importance of the glycine hinge, it is not fully conserved in all H-NOX domains. Several H-NOX sensors from the family Flavobacteriaceae contain a native alanine substitution in one of the hinge residues. In this work the effect of the increased steric bulk within the Ala-Gly hinge on H-NOX function was investigated. The hinge in Kordia algicida OT-1 (Ka H-NOX) is composed of A71 and G145. Ligand-binding properties and signaling function for this HNOX were characterized. The variant A71G was designed to convert the hinge region of Ka H-NOX to the typical Gly-Gly motif. In activity assays with its cognate histidine kinase (HnoK), wild type displayed increased signal specificity compared to A71G. Increasing titrations of unliganded A71G gradually inhibits HnoK autophosphorylation, while increasing titrations of unliganded wild type H-NOX does not inhibit HnoK. Crystal structures of both wild type and A71G Ka H-NOX were solved to 1.9 and 1.6 Å, respectively. Regions of H-NOX domains previously identified as involved in protein-protein interactions with HnoK display significantly higher b-factors in A71G compared to wild-type H-NOX. Both biochemical and structural data indicate that the hinge region controls overall conformational flexibility of the H-NOX, affecting NO complex formation and regulation of its HnoK.

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Introduction Nitric oxide (NO) is a signaling molecule common throughout all domains of life. Soluble guanylate cyclase (sGC) is the primary NO sensor in mammals 1. The N-terminal domain of sGC belongs to a family of Heme Nitric oxide/Oxygen sensing (H-NOX) domains. Upon formation of a Fe2+–NO HNOX complex, the C-terminal cyclase domain catalyzes the formation of cyclic GMP, a second messenger that controls several processes, including vasodilation and neurotransmission 2,3. In addition to acting as the sensor domain of sGC in eukaryotes, H-NOX homologs have been identified in numerous bacterial species 4. Bacterial H-NOX sensors are divided into two subfamilies found either in facultative or obligate anaerobes. Those found in facultative anaerobes, e.g. gammaproteobacteria, are trans-acting NO sensors that are found within the same operon as the enzyme that they regulate, which is commonly a histidine kinase or diguanylate cyclase. Depending on the bacterial species, these H-NOX domains regulate phenotypes such as biofilm formation or bioluminescence

5-8

. H-NOX domains from obligate anaerobes,

e.g. Clostridia, are fused to methyl-accepting chemotaxis proteins. These cis-acting H-NOX domains contain conserved hydrogen-bonding residues that allow formation of a stable Fe2+–O2 complex in addition to a Fe2+–NO complex 9-12. H-NOX proteins bind gaseous ligands with a Kd in the picomolar-nanomolar range. H-NOX proteins from both facultative anaerobes and mammals can form stable complexes with NO and CO. These H-NOX domains specifically sense NO in vivo; no in vivo role as a CO sensor has been unambiguously reported. The orientation of a distal subdomain at the N-terminus and a proximal subdomain at the C-terminus regulate the activity of a cognate effector protein by a conformational change that occurs upon NO complex formation 13. The conformational shift is initiated by dissociation of the Fe–His bond of the heme, leading to the formation of a 5-coordinate NO complex

14-16

. In the

conversion from the unliganded to the active NO complex, the porphyrin ring shifts position within the heme-binding pocket, adjusting interactions with nearby amino acid residues within Van der Waals contact and causing a conformational shift of the distal subdomain at the N-terminus 17. The importance of the Fe-His bond to activity has been shown through a histidine to glycine substitution that creates an H-NOX that is constitutively in an “active” conformation

18,19

. Alternatively, the NO complex of an H-

NOX protein that can only form a 6-coordinate NO complex (e.g. H-NOX with Mn2+ protoporphyrin IX) leads to a protein in an “inactive” conformation 17. In H-NOX proteins from obligate anaerobes, a similar conformational change occurs. However, this H-NOX activation is due to 6-coordinate formation of the Fe2+–O2 complex 20. Prior structural investigations of H-NOX proteins have illustrated the global conformational changes that occur during sensor activation 17-21. In previously reported structures, two conserved glycine


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residues are situated on the αD and αG helices (Fig. 1A). These glycine residues (termed the “glycine hinge”) form a flexible joint between the two helices and appear to be at the center of the conformational change that occurs after the H-NOX sensor binds its physiological gas ligand.

17,18

. Due to the strict

conservation and position in previously solved H-NOX structures, the “glycine hinge” was assumed to be a necessary pivot point in the activation of H-NOX sensors. However, recent sequence alignments of HNOX homologs in Flavobacteriaceae have revealed native alanine substitutions in one of the glycine hinge residues (Fig. 1B). The effect on H-NOX function with this alanine substitution into one of the glycine hinge residues has not previously been examined. Due to the proximity of the adjacent helices, substitution with a larger amino acid may disrupt protein structure or impede the sensor from shifting between conformations. The work presented here has characterized an H-NOX from Kordia algicida OT1, which contains an uncommon native alanine substitution at the glycine hinge residue G71.

Results and Discussion Identification of H-NOX sensors with a native alanine substitution in the glycine hinge Putative H-NOX sequences from the Flavobacteriaceae species Kordia algicida OT-1, Croceibacter atlanticus, and Zunongwangia mangrovi were aligned to previously characterized H-NOX proteins (Fig. 1B). In each H-NOX, there is a single native alanine substitution of one of the glycine hinge residues. The potential steric interference of the side chain methyl group may alter the signaling function of these H-NOX proteins. To investigate the function of these H-NOX domains with an alanine residue at αD and glycine at αG (A/G hinge), the ligand binding and signaling properties of K. algicida OT-1 (Ka) H-NOX were characterized.

Ligand binding properties of Ka H-NOX Ka H-NOX constructs of both wild type (WT) and a point mutant designed to reintroduce the canonical glycine hinge (A71G) were investigated to compare any effects of hinge region steric bulk on ligand binding characteristics. Both WT and A71G have absorbance spectra very similar to previously characterized H-NOX proteins

11,22

. Fe2+–unliganded Ka H-NOX has a Soret maximum at 429 nm. Both

WT and A71G form a 6-coordinate Fe2+–CO complex (Soret: 420 nm) and a 5-coordinate Fe2+–NO complex (Soret: 397 nm) (Fig. 2A and Fig. S1A). When the Ka H-NOX heme is oxidized, WT and A71G have different Soret peaks (WT, 408 nm; A71G, 395 nm) (Fig. S1B). It is possible that A71 subtly tunes the shape of the ligand-binding pocket and, by extension, the planarity of the heme cofactor. In previously characterized H-NOX proteins, Soret maxima of ferric heme at 408 and 395 nm are indicative of Fe3+– OH and Fe3+–unliganded, respectively, and mutations that directly tune the degree of porphyrin distortion affect ferric heme ligand affinity 23,24.



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Although WT and A71G form a 5-coordinate NO complex, there is a significant difference in the apparent rate of proximal histidine (H104) dissociation. In stopped-flow experiments performed on WT Ka H-NOX, the 6-coordinate Fe2+–NO complex (416 nm) is observable and decays to the 5-coordinate complex over a time course of 0.75 s at 20 °C (Fig. 2B). However, in the same time course, A71G forms a nearly complete 5-coordinate NO complex within the dead time of the stopped-flow (Fig. 2C). The rate of formation of the 5-coordinate complex is dependent on the concentration of NO in solution 16. At varying concentrations of NO (50-200 µM), WT Ka H-NOX consistently displays slower formation of the 5coordinate complex compared to A71G. However, a rate for 5-coordinate formation could not be calculated because of variability of the apparent rate values at low NO concentrations, likely due to limitations of the instrument. Even at low NO concentrations, no distinct Soret peak denoting the 6coordinate intermediate could be observed in A71G at 20 °C. Given the unusual effect on H104 dissociation, ligand affinity was investigated. The association rate for NO could not be calculated as it occurs within the dead time of the stopped-flow. Instead, the dissociation rate of NO was determined to probe any differences in NO dissociation between WT and A71G. A CO/dithionite trap was used to measure koff for NO, which resulted in a decrease of the Soret peak at 397 nm and an increase in the 6-coordinate CO complex at 420 nm

25,26

. As has been observed

with other H-NOX proteins, two dissociation rates were observed for both WT and A71G (Fig. S2), and values for k1 and k2 were averaged over 3 replicates. There is not a substantial difference in the dissociation rate constants of WT and A71G, indicating that A71 does not influence NO dissociation. Although A71 is on a helix near the ligand-binding pocket, the residue is predicted to face away from the pocket and would likely not interact directly with any potential ligand. There is a significant difference in the percent of k1 between wild type and A71G, which may be caused by differences in protein conformation. The two NO dissociation rates observed in H-NOX proteins are due to kinetically distinct protein conformations 27. In addition to the observation that A71G forms the 5-coordinate NO complex more rapidly, A71 may affect the conformational flexibility of the H-NOX, slowing dissociation of H104. Table 1. NO dissociation rate constants of Ka H-NOX Ka H-NOX

k1 (s-1)

k2 (s-1)

∆A1 (%)

Wild type

0.0063 ±0.0008

0.0007 ±0.0001

32.62 ±1.22

A71G

0.0051 ±0.0003

0.0008 ±0.0002

51.06 ±3.76

Ka H-NOX regulation of Ka HnoK The additional steric bulk in the hinge region of Ka H-NOX affects formation of the 5-coordinate NO complex, and so the effect on signal transduction was investigated next. The hnoX operon of K. algicida includes a predicted histidine kinase immediately downstream of the hnoX open reading frame.


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As with a previously characterized H-NOX and HnoK pair from Shewanella oneidensis, Ka HnoK has a helical domain at the N-terminus predicted to directly interact with the H-NOX

28

. Three regions

homologous to Per-Arnt-Sim (PAS) domains are located between the N-terminal helical domain and the histidine kinase domain (Fig. 3A). The function of these predicted PAS domains is unknown, but all putative HnoK proteins in Flavobacteriaceae share similar domain architecture. Autophosphorylation assays with [32P]-γ-ATP were performed to measure H-NOX regulation of HnoK activity. Wild type Ka H-NOX displays kinase inhibition only in the Fe2+–NO ligation state (Fig. 3B). Interestingly, maximal kinase inhibition (~20% total autophosphorylation) occurs with only 2:1 molar equivalents of NO-coordinated H-NOX to HnoK monomer (4 µM H-NOX, 2 µM HnoK). In previous HnoK characterizations with kinases from different organisms, much higher molar ratios of HNOX:HnoK (10:1-20:1 equivalents) were required to observe significant inhibition 7,8,28-30. Ferric or Fe2+– CO H-NOX do not inhibit the kinase (Fig. 3C). The presence of the NO donor DEA-NONOate in a kinase only control also does not affect kinase autophosphorylation. Lastly, concentrations of DEANONOate were added to yield final concentrations of NO in either excess (5 molar equivalents) or substoichiometric (sub, 0.8 molar equivalents) amounts relative to Ka H-NOX concentration. In the presence of sub [NO], the 5-coordinate complex forms, albeit more slowly compared to excess NO concentrations. Ka H-NOX exposed to either sub or excess [NO] was found to completely inhibit kinase activity. Regulation of Ka HnoK autophosphorylation was also tested with the mutant, A71G. As observed with the wild type, NO-coordinated A71G inhibits Ka HnoK (Fig. 4A and Fig. S3). Modest inhibition of HnoK autophosphorylation with Fe2+–unliganded A71G was also observed at 10:1 molar equivalents of H-NOX:HnoK. In order to compare potential Ka HnoK inhibition by Fe2+–unliganded A71G, kinase autophosphorylation was measured with increasing concentrations of unliganded H-NOX up to 50:1 molar equivalents. Higher inhibition of autophosphorylation is observed with increasing concentrations of Ka H-NOX A71G (Fig. 4B). Less than 40% of HnoK is phosphorylated with 50:1 molar equivalents of unliganded A71G. In contrast, unliganded WT H-NOX does not inhibit HnoK activity at any of the tested concentrations. Compared to the HnoK control without H-NOX, all assays with unliganded WT display ≥ 80% 32P–HnoK. The substantial degree of HnoK inhibition with high concentrations of A71G suggests that substitution of the bulkier alanine with glycine increases conformational flexibility, allowing the HNOX to sample an inhibitory conformation resembling that in the Fe2+–NO complex. A decrease in signaling specificity at increasing H-NOX:HnoK ratios has also been observed in the H-NOX/HnoK pair from Shewanella oneidensis 29. This H-NOX has a typical Gly-Gly hinge and inhibits its cognate HnoK with increasing concentrations of both Fe2+–CO and Fe3+ heme ligation states.



Crystal structures of Ka H-NOX wild type & A71G The ability for the A71G single point mutation to both affect the rate of 5-coordinate NO complex formation and regulate HnoK activity demonstrates that the hinge region forms the crux of the H-NOX conformational shift. To understand how an alanine or glycine residue at position 71 affects the overall HNOX structure, the crystal structures of both Ka H-NOX Fe2+–unliganded wild type and A71G (PDB 6BDD and 6BDE) were solved at 1.90 and 1.64 Å, respectively (Table S1). When aligning each structure to the C-terminal subdomain, both wild type and A71G share the same overall conformation (Fig. 5A). Alanine 71 on the wild type structure is positioned within the αD helix directly across from G145 on the αG helix. The distance between the methyl side chain of A71 and the α-carbon of G145 is approximately 3.7 Å (Fig. 5B and Fig. S4A and B). There are no major conformational changes of the N-terminal subdomain or flattening of the heme cofactor, which have been observed in previous structures comparing active and inactive H-NOX proteins 17,18,21,31. Instead of generating a recognizable conformational shift, it is possible that substitution of the A-G hinge with a canonical G-G hinge increases conformational flexibility of the H-NOX. Reviewing b-factors of specific regions of Ka H-NOX will provide some evidence for increased flexibility in the A71G variant. A slight increase in the average b-factor of the whole protein between A71G and wild type is shown in Table S1. However, specific regions of the H-NOX structure important for signaling and kinase interaction display higher b-factor changes relative to the rest of the protein. Bfactor values for Cα atoms were normalized to the average b-factor of each structure (Fig. S5). The loop between the αB and αC helices (residues 31-40) displays a modest increase in b-factor values between A71G and wild type. This loop has previously been characterized in other H-NOX proteins (from both S. oneidensis and Vibrio cholerae) as important for protein-protein interactions with the cognate HnoK 28,32. The largest differences in b-factors are observed at the end of the αD helix (Asp80) and near the edge of the αF helix (residues 109-111) (Fig. 5C and D, Fig. S4C and D). The normalized fold-difference of bfactors between A71G and WT was also mapped to the model, with the largest increases in b-factor values of Cα atoms highlighted in red (Fig. 5E and F). The proximal histidine ligand (H104) is located on the αF helix, and the higher flexibility of this helix may explain the apparent increase in the rate of formation of the 5-coordinate NO complex observed in A71G H-NOX. In a recent HDX-MS study of V. cholerae (Vc) H-NOX and HnoK interactions the αF helix exhibited the highest conformational changes after NO binding

32

. Additionally, there is evidence in both S. oneidensis (So) and Vc H-NOX that

conformational changes upon NO binding of the αF helix expands the binding interface with HnoK, regulating kinase activity

28,32

. To determine whether increased flexibility of Ka H-NOX affects affinity

for Ka HnoK, pull-down assays were performed with HisGST-tagged Ka HnoK and either untagged wild type or A71G H-NOX. A71G has a higher affinity for both full-length Ka HnoK and the truncation of the


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N-terminal helical domain (Fig. S6). The greater conformational flexibility of A71G Ka H-NOX increases the affinity for its cognate HnoK, which explains why the A71G H-NOX inhibits kinase activity without formation of the 5-coordinate NO complex.

Summary and Conclusions The results reported here show that the native alanine substitution found in K. algicida decreases the conformational flexibility of the H-NOX protein by acting as a molecular wedge at the crux of the HNOX conformational shift. The same would be expected for the other Flavobacteriaceae species that contain an H-NOX with an alanine substitution in the glycine hinge. Further evidence supporting this conclusion includes the results of the A71G variant that re-introduced the native glycine hinge. First, comparison of the crystal structures of both WT and A71G Ka H-NOX exhibit higher b-factor values in regions of the A71G structure previously implicated in protein-protein interactions and H-NOX sensor activation

14,28,32,33

. Second, stopped-flow spectroscopy has shown that A71G forms the 5-coordinate NO

complex more rapidly than wild type, which is direct biochemical evidence for increased conformational flexibility. However, the most striking evidence of the Ala-Gly hinge interface decreased the conformational flexibility is that increasing titrations of unliganded Ka H-NOX A71G inhibit HnoK autophosphorylation, while HnoK activity is unaffected by unliganded wild type H-NOX, even at H-NOX to HnoK ratios of 50:1. These results lead to a model of the glycine hinge mechanism (Fig. 6). The methyl side chain of A71 provides sufficient steric bulk in the hinge region to lock the conformation of Ka H-NOX (Fig. 6A). This effectively prevents ligand-independent inhibition of Ka HnoK by stabilizing the distal subdomain, and also slows the formation of the 5-coordinate NO complex in the proximal subdomain. In the A71G variant that represents a return to the canonical glycine hinge, conformational flexibility increases to that typically seen in Gly-Gly hinge H-NOXs (Fig. 6B). Both the formation of the 5-coordinate NO complex and the conformational shift of the distal subdomain are inextricably connected by the hinge region at the αD-αG helical interface. H-NOX proteins regulate unique signaling functions in both bacteria and eukaryotes. Elucidating the underlying mechanism of H-NOX signaling activation may expand biological understanding of mammalian sGC. Based on the structure of full-length sGC solved by electron microscopy, the enzyme exhibits a high degree of conformational flexibility

34

. Recombinant H-NOX proteins have also been

modified through incorporation of artificial porphyrins for a variety of uses, ranging from fluorescent reporters to MRI contrasting agents 35-37. Modifying conformational flexibility of H-NOX sensors can also be utilized to tune reporter output for future design of H-NOX-based probes.



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The in vivo function of H-NOX signaling in K. algicida is currently unknown. K. algicida was originally isolated from an algal bloom of the phytoplankton Skeletoma costatum and, as the name suggests, can kill and lyse phytoplankton 38,39. Although K. algicida does not encode a homolog for nitric oxide synthase, there is evidence of NO production and signaling in several phytoplankton species including S. costatum

40-42

. For K. algicida, NO may signal the presence of its diatom prey. Another

unanswered question is whether there is a signaling effect in vivo on K. algicida and other members of Flavobacteriaceae that encode hnoX genes with the Ala-Gly hinge. It is evident that the alanine residue in the H-NOX hinge region increases conformational stability, but how this affects a phenotypic response to NO is not yet known. Materials and Methods Cloning and expression: H-NOX sequences containing the native alanine substitution of G71 were identified using NCBI Protein BLAST of H-NOX proteins in Flavobacteriaceae. Gene fragments (gBlocks) of the hnoX gene and the H-NOX associated histidine kinase (hnoK) from Kordia algicida OT1 were ordered through Integrated DNA Technologies (IDT). Each gBlock was codon optimized for expression in Escherichia coli using a codon optimization tool from IDT. The gBlock of the hnoX gene contained a region encoding a C-terminal tobacco etch virus (TEV) protease cleavage site and a 6histidine tag. Golden Gate cloning was used to insert the hnoX gene fragment into a pET-28b expression vector (kindly provided by the Tullman-Ercek lab, Northwestern) 43. The gBlock of Ka hnoK was cloned into pHisMBP vector using restriction digestion of EcoRI and NotI on the 5’ and 3’ ends of the gBlock, respectively, and ligation with T4 DNA ligase (New England Biosciences)

44,45

. Plasmids were

transformed into E. coli DH5α and insertion of the correct gene was confirmed by sequencing through the UC Berkeley sequencing facility. Expression plasmid containing hnoX was transformed and expressed in E. coli RP523(DE3) as reported previously 46,47. The expression plasmid of Ka hnoK was transformed into E. coli BL21(DE3) and expressed using an identical protocol for the previously reported expression of the hnoK from Vibrio cholerae

20

. The mutant, A71G, was generated in Ka hnoX by QuikChange

mutagenesis (Agilent) using primers synthesized by IDT. The A71G mutant was transformed into E. coli DH5α and gene inserts were sequenced to confirm the point mutation. The hnoX mutant was expressed identically to wild type. Protein purification: All purification steps for Ka H-NOX wild type and A71G were performed using an identical protocol. Post expression cell pellets were thawed on ice and resuspended in Buffer A (50 mM Tris pH 8.0, 300 mM NaCl, 20 mM imidazole, 5 mM TCEP, 5 % glycerol). Cells were lysed using a C5 homogenizer (Emulsiflex), and cell debris was pelleted by ultracentrifugation at 40000 rpm with a Type 45ti rotor (Beckman Coulter). Clarified lysate was poured over a column containing His60 resin (Clontech). Resin was washed with 10 column volumes of Buffer A and 10 column volumes of


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Buffer A containing 40 mM imidazole. H-NOX was eluted with Buffer A containing 300 mM imidazole. After elution, 0.25 mg TEV protease was mixed with the H-NOX and the solution was dialyzed overnight into Buffer B (50 mM HEPES pH 7.5, 150 mM NaCl, 2 mM TCEP, 5 % glycerol). H-NOX was passed through clean His60 resin to remove the cleaved 6-His tag and TEV protease. The H-NOX was then run through a gel filtration column (Superdex 200 16/60 pg, GE Healthcare) equilibrated in Buffer B as a final purification step. H-NOX fractions were pooled, and purity was confirmed by SDS-PAGE and UVVis spectroscopy. H-NOX was flash frozen with liquid N2 and stored at -80 °C. Ka HnoK was purified using an identical protocol as previously reported on the HnoK from V. cholerae 20. UV-Vis spectroscopy of Ka H-NOX: Ligation and oxidation states of Ka H-NOX were generated similarly to previously reported H-NOX proteins 11. Briefly, H-NOX was brought into an anaerobic glove bag (Coy). All buffers and reagents used were purged with argon gas. Both wild type and A71G H-NOX were purified as Fe2+–unliganded. H-NOX concentration was determined by measuring the Soret absorbance of Fe2+–H-NOX (429 nm) and using the extinction coefficient from V. cholerae H-NOX (137 mM-1cm-1)

11

. The Fe2+–NO complex was formed by adding 500 mM (excess NO) or 120 mM

(substoichiometric NO) of DEA-NONOate to 200 mM H-NOX. DEA-NONOate (Cayman) releases 1.5 equivalents of NO. H-NOX was incubated with DEA-NONOate for 30 min before spectra were measured to ensure complete release of NO. The Fe2+–CO complex was formed by filling the headspace of a reactivial containing 200 mM H-NOX with CO gas. H-NOX was oxidized with 10 mM K3[Fe(CN)3] and desalted in Buffer B to form the Fe3+ oxidation state. Spectra were taken with either a Cary UV-Vis spectrophotometer or Nanodrop 2000. Stopped-flow spectroscopy: Stopped-flow spectroscopy was used to observe the conversion of 6coordinate Fe2+–NO to 5-coordinate Fe2+–NO. Fe2+–unliganded H-NOX was desalted into anaerobic Buffer B without TCEP and removed from the anaerobic chamber in a sealed glass tonometer. NO solution was generated by bubbling NO gas into a sealed anaerobic reactivial containing Buffer B without TCEP. H-NOX and NO buffer were added to separate sample syringes of the stopped-flow spectrometer (TgK Scientific). Stopped-flow spectra were taken over a time course of 0.75 s at 20 °C. NO dissociation rates: NO dissociation rates were measured similarly to previously described methods 27,48. NO dissociation was measured at 10 °C over a time course of 120 min. A CO/dithionite trap was used to prevent rebinding of NO during the experiment. Dissociation rates were calculated by measuring the difference spectra over time of CO complex formation and fit to a two-phase decay curve using the graphing software Prism (GraphPad). To eliminate the possibility of trap dependence, three concentrations of dithionite were used (100 mM, 30 mM, and 10 mM).



Page 10 of 21

Ka HnoK autophosphorylation assays: Kinase activity was determined using similar methods to previously reported HnoK enzymes

20,29

. Briefly, 2 µM Ka HnoK in Buffer C (50 mM TEA pH 8.0, 150

mM NaCl, 10 mM MgCl2) was mixed with 10 mM ATP and 2.5 µCi [32P]-γ-ATP in 100 mM Tris pH 8.0 to begin the assay. Assays were performed in 20 µL total volume. Reactions were quenched at 30 s by adding 5 µL 6x SDS-PAGE loading dye with 100 mM EDTA. Quenched reactions were resolved by SDS-PAGE. A storage phosphor plate (GE) was then exposed to the gel and phosphorylated HnoK was visualized using a Typhoon imager (GE). Band intensity of

32

P-phosphorylated HnoK was measured

using ImageQuant. For assays that measured H-NOX regulation of HnoK, 1-50 molar equivalents (2-100 µM H-NOX) were incubated with 2 µM HnoK for 30 min before ATP was added. Band intensity of 32Pphosphorylated HnoK + H-NOX was normalized to a kinase only control. Pull-down assay: HisGST Ka HK full-length or Ka HK 1-70 (4 µM) were mixed with 18 and 36 µM Ka H-NOX (WT or A71G) and 15 µL Pierce glutathione agarose (Thermo Fisher Scientific) in pulldown buffer (50 mM HEPES pH 8.0, 50 mM NaCl, 10 mM MgCl2, 2 mM TCEP, 5 % v/v glycerol) at 4 °C. After overnight incubation, glutathione agarose were pelleted by centrifuge at 3500 x g for 30 sec and washed with pull-down buffer twice before elution. Protein was eluted by 10 µL 6x SDS loading dye and was resolved in SDS-PAGE. Band intensity was quantified by ImageJ 49. Crystallization of wild type and A71G Ka H-NOX: Fe2+–unliganded Ka H-NOX was desalted into Buffer D (50 mM HEPES pH 7.5, 50 mM NaCl) for crystallization screens. Aerobic screens of Ka HNOX did not yield any diffraction-quality crystals. Frozen samples of Fe2+–unliganded WT Ka H-NOX were shipped to the laboratory of Cathy Drennan (MIT) and screened using the lab’s Mosquito Crystallization Robot housed in an anaerobic chamber. Initial crystallization conditions were 0.5 M LiCl, 0.1 M Tris pH 8.5, 28 % (w/v) PEG 6000. A71G also crystallized in similar conditions in an anaerobic glove bag. All reagents were flushed with argon prior to being placed in an anaerobic glove bag. Final optimized crystallization conditions were 0.3 M LiCl, 0.1 M Tris pH 8.5, 34 % (w/v) PEG 6000 with 500 µL of well solution and 2 µL of protein plus 2 µL well solution in sitting drop trays. Crystals were harvested shortly after formation because they began to lose diffraction quality after one week. Crystals of wild type and A71G were looped and snap frozen with liquid N2 before being removed from the anaerobic glove bag. X-ray diffraction and structure elucidation: X-ray diffraction data were collected on ALS beamline 5.0.1. The data were indexed, integrated and scaled using HKL2000

50

A71G structure were obtained through molecular replacement (PHENIX Phaser)

. Initial phases of the

48

with monomer A of

the Shewanella oneidensis Fe(II)-unliganded H-NOX (PDB: 4U99; modified by deleting loop regions and using PHENIX Sculptor) 51, and an initial model was built using PHENIX Autobuild 52. Initial phases of Ka WT structure were obtained through molecular replacement (PHENIX Phaser) with the solved


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structure of A71G. Models were improved through multiple rounds of manual model building (COOT) 53 and refinement (PHENIX Refine) 54. Well-defined density for the unliganded heme was readily identified and placed into the model (PHENIX LigandFit) 55. Water molecules were added with PHENIX Refine, and refinement of the model continued until final refinement statistics were obtained (Table S1).

Associated Content Supporting Information: The Supporting Information is available free of charge on the ACS Publications website, http://pubs.acs.org. Figures S1-S6. Table S1

Accession codes Coordinates for the crystal structures included in this work were deposited in the Protein Data Bank. Ka H-NOX Fe(II)-unliganded WT is 6BDD, and Ka H-NOX Fe(II)-unliganded A71G is 6BDE.

Acknowledgments This work was funded in part by the George E. Hewitt Foundation for Medical Research. The authors thank 5.0.1. beamline scientists and Lawrence Berkeley National Lab for technical assistance. For assistance in setting up initial screens and use of anaerobic high-throughput crystallization equipment, the authors thank C.M. Drennan, C. Nguyen, and R.E. Bjork.



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Figures and Legends

Fig. 1. Comparison of glycine hinge regions in H-NOX proteins. (A) Crystal structures of Caldanaerobacter subterraneus Fe2+–O2 (orange) and Fe2+–unliganded (salmon) (1U55 & 5JRU) and Shewanella oneidensis Fe2+–unliganded (wheat) and Fe2+–NO (red) (4U99 & 4U9B) H-NOX. Conserved glycine hinge residues on the D and G helices are highlighted in blue. The crux of H-NOX conformational change occurs along the two helices with these conserved glycine residues. The distal (Nterminal) and proximal (C-terminal) subdomains are labeled. (B) Primary sequence alignment of H-NOX proteins from S. oneidensis, Kordia algicida, Croceibacter atlanticus, Zunongwangia mangrovi, C. subterraneus, and Nostoc sp. The conserved hinge residues are noted. Although most characterized HNOX proteins have two conserved glycine residues, three species from Flavobacteriaceae with a single native alanine substitution are included in the alignment.



Fig. 2. Ligand binding characterization of K. algicida H-NOX. (A) UV-Vis spectroscopy of Ka H-NOX Fe2+–unliganded, NO, and CO. Wavelength of Soret maximum is included next to each peak. (B) Stopped-flow spectroscopy showing conversion of 6-coordinate (416 nm) to 5-coordinate (397 nm) Fe2+– NO for wild type Ka H-NOX. Arrows indicate the loss of the 6-coordinate complex (416 nm) and the formation of the 5-coordinate complex (397 nm). (C) Stopped-flow spectroscopy showing conversion of


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6-coordinate to 5-coordinate Fe2+–NO for Ka H-NOX A71G. The arrow indicates the formation of the 5coordinate complex.

Fig. 3. Ka HnoK regulation by wild type Ka H-NOX. (A) Ka H-NOX and HnoK domain architecture. (B) Kinase autophosphorylation assay. HnoK (2 µM) was mixed with increasing molar equivalents of Fe2+– unlig or NO-coordinated H-NOX. Autophosphorylation was measured using radiolabeled [32P]-γ-ATP. Quantified phospho-HnoK bands were normalized to a control sample that does not contain H-NOX (-) (C) Autophosphorylation of 2 µM HnoK with 10 µM H-NOX with different heme oxidation or ligation states. Notably, Fe2+–NO H-NOX inhibits kinase autophosphorylation. NO ex and NO sub indicate either excess or substoichiometric concentrations of DEA-NONOate were added relative to H-NOX concentration. The (-) NO control denotes that 500 µM DEA-NONOate was preincubated to a reaction mixture that only contains HnoK. All phospho-HnoK bands were normalized to the kinase only (-) control.



Fig. 4. Ka H-NOX A71G regulation of HnoK autophosphorylation. (A) Kinase autophosphorylation assay. HnoK (2 µM) was mixed with increasing molar equivalents of Fe2+–unlig or NO-coordinated A71G mutant. Autophosphorylation was measured using radiolabeled [32P]-γ-ATP. (B) A71G H-NOX inhibits Ka HnoK. Increasing molar equivalents of WT and A71G (2-50 eq.) were mixed with 2 µM HnoK. HnoK with up to 50 equivalents of Fe2+–unliganded WT H-NOX retains ≥ 80% autophosphorylation compared to kinase only control. Autophosphorylation is inhibited with increasing equivalents of Fe2+–unliganded H-NOX A71G.


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Fig. 5. Crystal structures of WT and A71G Ka H-NOX. (A) Structural alignment of WT (green) and A71G (cyan). Helices are labeled αA-αG, and the proximal and distal subdomains are annotated. (B) Rotated view of WT H-NOX displaying αD and αG helices. The methyl group side chain of A71 is shown on the αD helix, and G145 is highlighted in red. The distal and proximal subdomains are also annotated with respect to the current structural view. (C) B-factor sausage representation of WT H-NOX. The heme cofactor was included as marker to show that angle of image is identical to panel A. Areas of lower flexibility compared to A71G are noted by the blue arrow or box. (D) B-factor putty of A71G HNOX. Areas of increased flexibility compared to WT are highlighted by the orange box and arrow. Bfactor scale is by color with lower values closer to blue and higher values closer to red. (E and F) Normalized b-factor fold difference between A71G and WT Ka H-NOX. The b-factor difference after normalization plotted in Fig. S5 was mapped to the structure of Ka H-NOX. Red portions indicate significantly higher b-factor values of H-NOX residues on the A71G mutant structure compared to WT H-NOX. Blue portions denote lower b-factor values of A71G H-NOX residues compared to WT H-NOX.

Fig. 6. Model of hinge region steric locking. (A) Schematic of WT Ka H-NOX highlighting helices αD and αG. The methyl side chain of A71 provides conformational stability to the H-NOX until the sensor binds NO. (B) H-NOX proteins with a canonical glycine hinge have higher conformational flexibility and can sample an active state even in the absence of NO.


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Glycine hinge TOC 79x30mm (300 x 300 DPI)


Native alanine substitution in the glycine hinge modulates

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