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Structural insights into the mechanism of the human soluble guanylate cyclase

Soluble guanylate cyclase (sGC) is the primary nitric oxide (NO) sensor. It plays a central role in NO signaling and is implicated in many essential physiological processes and disease conditions. The binding of NO leads to a significant boost in sGC enzymatic activity. However, the mechanism of NO activation remains incompletely understood. Here, we report the cryo-electron microscopy structures of the human sGC α1β1 heterodimer in different functional states. These structures revealed that the transducer module bridges the NO sensor module and the catalytic module. NO binding to the β1 H-NOX domain triggers the structural rearrangement of the sensor module and the bending-straightening conformational switch of the transducer module. The resulting movement of the N-termini of the catalytic domains drives the structural changes within the catalytic module, which in turn boost sGC enzymatic activity. These observations indicate the structural framework for the mechanism of sGC activation induced by NO binding.

Introduction maps showed blurry features, which were indicative of continuous conformational heterogeneities.
Therefore, we divided the whole molecule into two bodies-the larger "N lobe" and the smaller "C lobe"-for further multibody refinement (Figures S1-S4) , and the subsequent local map qualities were dramatically improved . Analysis of the multibody refinements revealed monomodal particle distributions along the major eigenvectors given by the principal component analysis, indicating that the C lobe moved in a continuous fashion relative to the N lobe, probably due to the endogenous structural flexibility of the CC domains that connect the two lobes (Videos S1 and S2, Figures S2C-E and S4A-C). The two individual multibody maps were further aligned to the full consensus map and summed to generate the composite maps for model building and interpretation. The composite map qualities were sufficient to trace the main chain of most residues with the aid of available high-resolution homologous structures ( Figures S2B, S2F, S3E, S4D, S5, S6, and Table S1). For the β1 H105C mutant sample, we added Mg 2+ ions and the noncyclizable substrate analogue GMPCPP to the protein. The dataset of the β1 H105C mutant was processed as described above to obtain a reconstruction at 6.8 Å ( Figures S4F-G). At this resolution, the overall shape and domain organization of β1 H105C mutant were found to be similar to that of the NO-activated state, which was low-passed to the same 6.8 Å resolution, with a real space correlation of 0.96. However, the heme density is missing in the β1 H-NOX domain of the β1 H105C mutant, as expected ( Figure S4H).

Structure of sGC in the inactive state
Both the heme-unliganded and the heme-oxidized sGC were in a contracted conformation, which is akin to the low resolution "bent" conformation observed previously in negative stain EM (Campbell et al., 2014) (Figures 1C-E, Video S3). We found that the overall structure of sGC in the heme-unliganded state (4 Å) is essentially the same as that in the heme-oxidized state (3.9 Å), with a RMSD of only 0.28 Å ( Figure S7A). This was in accordance with the functional data, which showed that heme-unliganded state and heme-oxidized state have low activity ( Figure 1B) . Therefore, both of the structures were considered as the inactive state, and the 3.9 Å heme-oxidized state is used in further discussion of the inactive state due to its higher resolution and better map quality compared to the 4.0 Å heme-unliganded state. The structure of the inactive sGC occupies the 3D space of 140 Å × 75 Å × 75 Å ( Figures 1C-E). The large N lobe is composed of α1 H-NOX, α1 PAS, β1 PAS, and β1 H-NOX domains.
These domains are arranged in a pseudo two-fold symmetric manner, with the scaffolding PAS domains at the center and the H-NOX domains at the periphery ( Figure 1C). These domains are essential for NO sensing and form the N-terminal "sensor" module of sGC ( Figure 1C). The CC domains of both subunits form the "transducer" module that bridges the N-terminal sensor module and the C-terminal catalytic module ( Figure 1D).

Structure of the sGC sensor module in the inactive state
The atomic model of sGC in the inactive state allowed us to characterize the domain-domain interfaces in detail (Figures 2A-I). The N-terminal α1 H-NOX domain was previously proposed to share a common H-NOX fold, but the critical histidine residue for heme coordination is not conserved (Figures S5 and S6).
Here, we show that the structure of the α1 H-NOX domain is similar to that of the β1 H-NOX domain.
However, the N-terminal αA helix of the α1 H-NOX occupies a large portion of the pseudo-heme binding pocket and contributes to the formation of the H-NOX hydrophobic core ( Figure S7B). Therefore, the α1 H-NOX domain does not retain the ability to bind heme under physiological conditions. The α1 H-NOX domain interacts with the adjacent PAS and β1 CC domains mainly through hydrophobic interactions ( Figure 2B). βA, the αH-βD loop, and αF of the α1 H-NOX domain make contacts with the αK-βH loop of the α1 PAS domain. αA, αE, and αF of the α1 H-NOX domain interact with αL, the αL-αM loop, and αM of the β1 subunit ( Figure 2B). Notably, there is no direct interaction observed between the α1 H-NOX and the β1 H-NOX domains, likely due to their distal spatial arrangement (Figure 2A).
The structure of each PAS domain resembles that of the Manduca sexta sGC α subunit (PDB ID: 4GJ4) (Purohit et al., 2013) (Figures S7C-D). The α1 and β1 PAS domains are arranged into a pseudosymmetric heterodimer ( Figure 2C). Hydrophobic residues on αI, βI and βJ of the α1 PAS domain and αH, βI and βJ of the β1 PAS domain are packed into the stable hydrophobic core of the α1β1 PAS heterodimer ( Figure 2C). In addition, αL of the PAS domain interacts with αM of the CC domain of the same subunit ( Figure 2C). Interestingly, the β1 PAS domain interacts extensively with αN of the β1 CC domain. S326 on βJ hydrophilically interacts with Q367 of αN, and R305 on βI and R258 on βG interact electrostatically with E370 and D374 on αN, respectively ( Figure 2E).
The H-NOX domain of the β1 subunit is the NO sensor of sGC (Derbyshire and Marletta, 2012). It shares common structural features with prokaryotic H-NOX domains (Plate and Marletta, 2013), which are composed of both N-terminal and C-terminal subdomains ( Figure S7B). A heme molecule binds inside the β1 H-NOX domain, and its five coordinated Fe ion is tightly bound to H105 of αF, as evidenced by the strong connecting density between them ( Figure 2D). β1 H-NOX extensively interacts with both the neighboring PAS heterodimers and the transducer module. On one side, I86 and V89 of αE hydrophobically interact with L390, F393, and L400 of the α1 PAS domain and L414 of the α1 CC domain, while G82, H107, and T110 of the β1 H-NOX domain form a complex hydrogen bonding network with R420, E417, and R428 of the α1 CC domain ( Figure 2F). On the other side, E96, Q99, and N100 on αF of the β1 H-NOX domain form a polar interaction nexus with S401 of the α1 PAS domain, N273 of the β1 PAS domain, and R410 of the α1 CC domain ( Figure 2E). In addition, E138 and D106 of the β1 H-NOX domain electrostatically interact with R381 of the β1 CC domain and R258 of the β1 PAS domain, respectively ( Figure 2E).

Structure of the sGC transducer module and the catalytic module in the inactive state
The polar interactions between the sensor module and the CC domains stabilize the transducer module in the "bent" conformation, in which both the α1 and β1 CC domains are broken into two short helixes (αM and αN) connected by a near 90° turn ( Figure 2G). The α1 CC helix breaks at the forth turn (Figures 2G and S5) and the β1 CC helix breaks at the third turn ( Figures 2G and S6). The αM and αN helixes of the α1 and β1 subunits still maintain two coiled-coil structures that are stabilized by the hydrophobic interactions of residues from both subunits ( Figure 2G). The majority of these residues are leucines, which pack into structures reminiscent of leucine zippers ( Figure 2G).
The C-terminal ends of the transducer module not only directly connect but also tightly interact with the catalytic module. I457 and F458 of the α1 CC insert into the hydrophobic pocket formed by A463, V462, L466, and Y594 of the α1 catalytic domain and M537 of the β1 catalytic domain ( Figures 2H-I). In a similar fashion, L394, Y395, V397, and L398 of the β1 CC insert into the hydrophobic pocket formed by L406, A403, V402, Y540, and Y470 of the β1 catalytic domain and M591 of the α1 catalytic domain ( Figures 2H-I). D391 of the β1 CC domain also electrostatically interacts with R407 of the β1 catalytic domain ( Figure 2H). In the catalytic module, the two subunits are bound in a pseudo-symmetric manner ( Figure S7E), consistent with previous observations of the isolated heterodimer (Seeger et al., 2014).
However, the structure of the catalytic module in the inactive state of the full-length sGC is different from that of the isolated α1β1 catalytic domain heterodimer (Seeger et al., 2014). By aligning their β1 subunits, we found that the αQ helix of the α1 subunit rotates for about 13º ( Figure S7F), indicating that the context of the full-length enzymeespecially the interactions between the transducer module and the catalytic modulewas important for the structure of the catalytic module. When we compared the structure of the sGC catalytic module in the inactive state with that of adenylate cyclase in the active state (PDB ID: 1CJU) (Tesmer et al., 1999), we found that there are steric clashes between the adenosine group of the substrate and the α1 subunit of sGC and between the tri-phosphate group of the substrate and the β1 subunit of sGC ( Figure S7G). This suggests that the structure of sGC in the inactive state is incompatible with substrate binding, which is in accordance with previous studies that showed that inactive sGC has a high K m (Denninger et al., 2000).
To characterize and validate the domain-domain interactions observed in the inactive state, we generated a sGC mutant (named cys-less sGC, sGC CL ) that has characteristic sGC enzymatic activity ( Figure 2J) but fewer cysteines, and thus reduces background cross-linking (see Methods section). Cysteine mutations were introduced at spatially neighboring positions on both subunits. One pair (α1 L275C and β1 A316C) was located between the PAS domains, and the other pair (α1 L425C and β1 L365C) was located between the CC domains. We found that for each cysteine mutation pair, oxidative cross-linking only happened when the cysteine mutants were present in both subunit simultaneously ( Figures 2K-L), which was consistent with our structural model of sGC in the inactive state.

Structure of sGC in the NO-activated state
The NO-activated sGC has a dumbbell-shape structure in which the sensor module moves away from the catalytic module (Figure 3, Video S4). This is akin to the "extended" conformation previously observed by negative-stain EM (Campbell et al., 2014) and is dramatically different from the "bent" conformation of the inactive state. In addition, the overall structure of the constitutively active β1 H105C mutant shows a similar "extended" conformation in the absence of NO donor ( Figure S4F). These structures suggest that this large overall conformational change is associated with the enhanced enzymatic activity of sGC but probably does not result from the covalent S-nitrosylation of sGC by NO, which can lead to sGC desensitization (Mayer et al., 2009;Sayed et al., 2007).
Despite the large overall conformational change, the general domain arrangement within each module in the NO-activated state is similar to that in the inactive state (Figures 1 and 3). In the electron density map of the NO-activated state, the H105-Fe bond of β1 H-NOX is cleaved, as evidenced by the clear separation between each density ( Figure 4A). This suggests the current conformation corresponds to a NO-bound state, likely because excess NO donor DEA NONOate was added to the sample. However, we could not explicitly model the NO molecules or the heme deformation due to the limited resolution. The binding of NO induces a conformational change in β1 H-NOX in which the C-terminal subdomain rotates relative to the N-terminal subdomain ( Figure S7H). As described in the inactive state structure, αE and αF of β1 H-NOX contribute to a large portion of the inter-domain interactions in the inactive state (Figures 2E -F). When αF (residues 96-107) of the β1 subunit was used as the reference to superimpose the structure of the NO-bound β1 H-NOX domain onto the structure of the inactive state, the Cα atom of N62 in the N-terminal subdomain has 4.6 Å displacement ( Figure 4B). Furthermore, the NO-bound β1 H-NOX domain sterically clashes with the adjacent domains of the inactive state. Specifically, R40, I111, R116, and R121 of the NO-bound β1 H-NOX domain clash with Q422 and Q418 of the α1 CC domain, L380 of the β1 CC, and S270 of the β1 PAS domain of the inactive state, respectively ( Figure 4B). This indicates that the inactive state structure is incompatible with the NO-bound β1 H-NOX domain and, therefore, a structural rearrangement is required to accommodate the conformational change of the β1 H-NOX domain upon NO binding. Indeed, we observed structural changes within the sensor module in which α1 H-NOX has a small downward movement while β1 H-NOX has a large rotational and translational movement ( Figure 4C).  Figure 4E). To determine the function of these interfaces, alanine mutations were introduced to disrupt the interactions between them. We found that the D106A mutation in αF and the R258A mutation in βG of the β1 subunit both impaired NO activation ( Figure 4F), suggesting that this interface plays an essential role in sGC activity in the NO-activated state.

Structure of the sGC transducer module in the NO-activated state
The dramatically altered interactions between the CC domain and the sensor module lead to the conformational change of the transducer module ( Figure 5). Strikingly, the linkers between αM and αN fold into α helical structures, which fuse the αM with αN into the 71 Å long αMN helixes ( Figure 5A).
Specifically, R420-K426 of the α1 CC domain and L355-Q358 of the β1 CC domain fold into α-helical structures (Figures 5B, S5, and S6). As a result, the transducer module switches from a highly bent conformation in the inactive state to a long, continuous coiled-coil structure in the NO-activated state ( Figure 5B). The folding of the αM-αN loops results in a decrease in the exchangeability of the main chain hydrogens due to their hydrogen bonding in α helixes. This is consistent with previous hydrogendeuterium exchange mass spectrum results that showed that the αM-αN loops had a significantly slower exchange rate upon NO activation . To determine the functional importance of this bending-straightening conformational change, we mutated residues in the αM-αN linker to either prolines or alanines. Prolines generate kinks in helical structures due to their incapability to form hydrogen bonds on the main chain. Therefore, proline mutations should destabilize the helical structures of αMNs in the NO-activated state, and these proline mutants may favor the inactive conformation.
Indeed, proline mutations of D423 in the α1 CC domain or G356 in the β1 CC domain rendered sGC unresponsive for NO activation (Figures 5B-C). By contrast, mutation of the same set of residues into alanines had no such effect, indicating that the continuous helical structures of the αMNs are essential for NO activation of sGC ( Figure 5C).
In the NO-activated conformation, the interface between the α1 and β1 CC domain is drastically different from that observed in the inactive state. At the N-terminal part of the transducer module, A408, L409, V412, and I415 of α1 CC interact with A348, L352, and L355 of β1 CC ( Figure 5A). At the middle of the transducer module, L429 and L436 of α1 CC interact with L369 and L376 of β1 CC ( Figure 5A). At the C-terminal part of the transducer module, L443, L453, L454, and I457 of α1 CC make hydrophobic interactions with L383, L393, L394, and V397 of β1 CC ( Figure 5A). In addition, E446 and K447 of α1 CC electrostatically interact with K387 and E386 of β1 CC ( Figure 5A). Besides the overall "bendingstraightening" movements of each CC domain, the αN helix of the α1 subunit rotates approximately 70° around the αN helix of the β1 subunit ( Figure 5D). The separation of the C-termini of the transducer module also decreases. The distance of the Cα atoms between P459 of the α1 subunit and P399 of the β1 subunit shrinks from 26 Å to 20 Å ( Figure 5E).

Structure of the sGC catalytic module in the NO-activated state
As a consequence of the conformational change in the transducer module, the N-termini of the catalytic domains of each subunit move closer towards each other ( Figure 6A). This drives the structural reorganization of the entire catalytic module. The catalytic domain of the α1 subunit rotates 17º relative to the β1 catalytic domain ( Figures 6A-B). These movements reorganize the catalytic center to facilitate the binding of the substrate GTP and the cofactor Mg 2+ ions. In the map of the NO-activated state, we observe a strong density corresponding to the substrate analogue GMPCPP ( Figure 6C). The catalytic domains of both subunits contribute to GMPCPP binding. αS, βK, βL, and βM of the β1 subunit form the guanosine binding pocket and R552 of the β1 subunit binds the α phosphate of GMPCPP. D530 and D486 of the α1 subunit coordinate the Mg 2+ ions, which are essential for catalysis. Positively charged R574 of the α1 subunit and K593 of the β1 subunit bind the γ phosphate of GMPCPP ( Figure 6C). By comparing the current structure with the active adenylate cyclase structure (PDB ID: 1CJU) (Tesmer et al., 1999) ( Figure 6D), we found that the residues responsible for substrate binding and catalysis are at similar positions, indicating that the current sGC structure represents a catalytically competent conformation.
Interestingly, we found that the C-terminus of the α1 catalytic domain folds back to bind between αQ of the α1 subunit and αO-βK of the β1 subunit in the NO-activated state ( Figure S4E), although the exact Cterminal residues could not be explicitly modeled, probably due to their high flexibility.

Structural mechanism of sGC activation
By analyzing the structures of individual sGC domains in both inactive and NO-activated states, we found that the structures of the scaffolding PAS dimer remain relatively unchanged among the different states, with a RMSD of 0.91 Å ( Figure S7I). Therefore, we used the structures of the PAS dimer as a reference point to align and to compare the two full-length structures ( Figures 7A-B). During activation, the α1 H-NOX domain has a small concomitant downward movement, while the interfaces between α1 H-NOX and its adjacent domains are largely maintained ( Figure 4C). This suggested that the α1 H-NOX domain may play a role that is mainly structural instead of one involved in NO signal transduction (Video S5). This is in agreement with the activity assay result that the H-NOX domain of the α1 subunit is dispensable for NO activation ( Figure 7C) (Koglin and Behrends, 2003). In contrast, the local conformational change of the β1 H-NOX domain upon NO binding drives the structural rearrangement of the sensor module ( Figure 4C), which, along with previous functional data, suggested that the H-NOX domain of the β1 subunit plays an essential role in NO sensing (Derbyshire and Marletta, 2012). Indeed, complete removal of the β1 H-NOX domain rendered the sGC enzyme trapped in a relatively low activity state and unresponsive to NO activation ( Figure 7C). This suggests that the β1 H-NOX domain in the NO-bound state is necessary to stabilize the sGC enzyme in an active conformation. Further supporting this conclusion, disruption of the interactions between β1 H-NOX and adjacent domains by mutation also diminished the NO activation ( Figure 4F).
The structural changes in the sensor module upon NO binding trigger the bending-straightening conformational switch of the transducer module. As a result, the distal catalytic module rotates 86º in a swing-like manner and is spatially displaced by 101 Å (Figures 7A-B). Inhibition of the straightening by proline mutations abolishes the NO activation ( Figure 5C), which suggests that the conformational change in the transducer module is essential for the activation of the catalytic module. Furthermore, we found that the catalytic module changes from a substrate binding incompatible conformation to a catalytically competent conformation ( Figures 6A-B), which explained how the binding of NO decreases the K m (GTP) and increases the k cat of sGC (Denninger et al., 2000). However, the information flow in reverse direction, from the catalytic module to the sensor module, was implicated by previous functional data described in the literatures. Previous studies have reported that GTP binding to the catalytic module can stabilize the initial five-coordinate sGC-NO complex (Tsai et al., 2011). Moreover, ATP and GTP binding to the catalytic module slows NO dissociation (Cary et al., 2005). As a result, the catalytic module may act as a sensor for intracellular nucleotides and affect the sensitivity of sGC to NO (Ruiz-Stewart et al., 2004). We speculate that the nucleotide binding events in the catalytic module can alter its local conformation and send the signal via the middle transducer module to prime the sensor module and enhance NO sensitivity. Therefore, the transducer module acts as an allosteric structural coupler between the sensor module and the catalytic module to allow the bi-directional flow of information within the sGC molecule.
The structures of human α1β1 sGC presented here provide the first high-resolution view of the architecture and domain organization of this enzyme. Moreover, visualizing the sGC structures of multiple states, together with accompanying biochemical analysis, allowed us to characterize its allosteric activation mechanism ( Figure 7D),. Our work not only paves the way towards a better mechanistic understanding of sGC activity, but also provides a template for structure-based drug design targeting related human diseases.

Acknowledgments
We thank all of Chen Lab members for kindly help. We also thank Jiahuai Han for sharing the human

Declaration of Interests
The authors declare no competing interests.  (B) End-point activity assay of the purified non-tagged wild type sGC protein sample for cryo-EM, compared with that of NS2028 treated non-tagged wild type sGC, wild type sGC with CGFP tag (WT) and β1 H105C mutant with CGFP tag (Data are shown as means ± standard error, n = 3).         human α1 and non-tagged β1 yielded a stable heterodimer. The two coding sequences were transformed into the pFastbac dual vector and expression was driven by p10 or polyhedrin promoters. The corresponding baculovirus was generated using the Bac-to-Bac system.

Protein Expression and Purification
Sf9 insect cells at a density of 4 × 10 6 /mL in SIM SF medium were infected with the baculovirus and cultured at 27°C in a shaker for 72 h before harvesting and storage at -80°C . Cells corresponding to 500 ml culture were thawed and resuspended with 20 mL lysis buffer (50 mM Tris pH 8.0 at 4º C, 150 mM NaCl) containing 1 μg/mL aprotinin, 1 μg/mL pepstatin, 1 μg/mL leupeptin, 1 mM phenylmethanesulfonylfluoride, 2 mM dithiothreitol (DTT), and 1 mM EDTA. Cells were broken by sonication in 5 s intervals followed by a 5 s pause at 50% output for 20 min. Unbroken cells, cell debris, and membranes were removed by ultracentrifugation at 40,000 rpm for 1 h at 4°C using a Ti45 rotor (Beckman). Excess amount of purified glutathione S transferase-tagged GFP-nanobody (Tang et al., 2018) was added to the supernatant and incubated at 4°C for 10 min with rotation. Samples were then loaded onto 4 mL Glutathione Sepharose 4B columns (GE Healthcare) and washed with TBS buffer (20 mM Tris, pH 8.0, 150 mM NaCl) containing 1 mM dithiothreitol at 4°C. Protein was eluted with elution buffer (50mM Tris, pH 8.0, 10 mM reduced glutathione, 1 mM dithiothreitol) at room temperature. The eluate was diluted with buffer A (20 mM Tris, pH 8.0 at 4°C ) to a conductance lower than 5000 Ω -1 and loaded onto a 1 mL HiTrap Q HP (GE Healthcare). The protein was eluted with buffer B (20 mM Tris, pH 8.0, 500 mM NaCl) at 4°C in a linear gradient using the AKTA pure system (GE Healthcare). The peak fractions containing sGC were pooled and incubated with prescission protease overnight to cleave the tag off from the protein. The digested protein was further purified by Superdex 200 increase (GE Healthcare) running in buffer containing 20 mM HEPES, 50 mM NaCl and 2 mM Tris (2-carboxyethyl) phosphine (TCEP). The peak fractions containing the sGC protein were pooled and concentrated. UV-Vis spectrum is measured using a spectrometer (Pultton) in the cuvette mode.

Activity Assay
The protein used for cryo-EM sample preparation was diluted with 20 mM triethanolamine (pH 7.6), 300 mM NaCl, 1 mM dithiothreitol and subjected to activity assay as described below. For the heme-oxidized sample, the protein was diluted with 20 mM triethanolamine (pH 7.6), 300 mM NaCl and preincubated with 20 μM NS2028 at 25°C for 30 min and then added dithiothreitol to the final concentration of 1mM for activity assay. To generate the sGC mutant protein for activity assay, the coding sequences of the α1 subunit with a C-terminal GFP-strep tag and the β1 subunit were cloned into pFastBac1 expression vectors, respectively. To generate α1-ΔNOX and β1-ΔNOX constructs, the N-terminal 273 amino acids of the α1 subunit and 200 amino acids of the β1 subunit were removed, respectively. The constructs carrying desired point mutations were generated by Quick Change method and corresponding baculoviruses were generated by Bac-to-Bac system. Sf9 insect cells at the density of 4 × 10 6 /ml in SIM SF medium were infected with baculovirus and cultured in a shaker at 27°C for additional 72 h before harvest. Cells were resuspended with buffer containing 50 mM Tris (pH 8.0 at 4°C), 150 mM NaCl, 1 μg/ml aprotinin, 1 μg/ml pepstatin, 1 μg/ml leupeptin, 1 mM phenylmethanesulfonylfluoride and 1mM dithiothreitol, and broken by passing through the syringe needle with 0.45 μm inner diameter for 6 times. Cell debris and membrane were removed by ultracentrifugation at 40,000 rpm for 30min at 4°C using TLA55 rotor (Beckman). The supernatants were loaded onto Streptactin Beads 4 FF (Smart-Lifesciences) and washed with buffer containing 20 mM Tris (pH 8.0 at 4°C), 150 mM NaCl and 1 mM dithiothreitol. The protein was eluted with buffer containing 50 mM Tris (pH 8.5 at 4°C), 50 mM NaCl, 10 mM D-desthiobiotin and 1 mM dithiothreitol. The eluates were diluted with equal volume of 20 mM triethanolamine (pH 7.6) and loaded onto Q Beads 6 FF (Smart-Lifesciences) by gravity and washed with buffer containing 20 mM triethanolamine (pH 7.6), 150 mM NaCl and 1 mM dithiothreitol. The protein was eluted by 20 mM triethanolamine (pH 7.6), 300 mM NaCl and 1 mM dithiothreitol. The protein concentrations of various GFP-tagged sGC mutants were determined by comparing their GFP fluorescence signal to that of a purified GFP-tagged sGC standard on fluorescence-detection size-exclusion chromatography (FSEC) (Kawate and Gouaux, 2006). The activity assay mixture contained 10 nM sGC, 60 mM triethanolamine (pH 7.6), 150 mM NaCl, 0.5 mM dithiothreitol, 5 mM MgCl 2 , 200 µM GTP with or without 200 µM DEA NONOate (Cayman Chemical) in a final volume of 20 μl. The assay mixture was incubated at 25°C for 10 min and stopped by adding 80 μl 125 mM Zn(OAc) 2 and 100 μl 125 mM Na 2 CO 3 . The GTP-ZnCO 3 precipitation was removed by centrifugation at 17,000 × g for 5min and the supernatant was used for cGMP quantification, which was done by using the Cyclic GMP ELISA Kit (Cayman Chemical) according to the instruction. Each assay was independently repeated for at least 3 times.

EM sample preparation
Purified sGC was concentrated and supplemented with the ligands before freezing. Protein samples were loaded onto glow-discharged Quantifoil 0.6/1 holey carbon gold grids and plunged into liquid ethane by Vitrobot Mark IV (Thermo Fisher Scientific).

Disulfide bond cross-linking
To generate the cys-less sGC construct (sGC CL ), the cys-rich N-terminal 63 amino acids of α1 subunit were removed. Additional mutations of C176A, C239A, C669S, C455Y, and C460G were done on α1 subunit and C292N were done on β1 subunit. The coding sequences of α1 CL with C-terminal GFP-strep tag and β1 CL subunit without tags were cloned into modified BacMam expression vectors (Goehring et al., 2014;Li et al., 2017) respectively. Then specific amino acids were mutated into cysteines by Quick Change method. Cysteine mutants were transfected into HEK293F cells with 3 μg/ml polyethylenimines

Cryo-EM Data Acquisition
Cryo-grids were screened on a Talos Arctica electron microscope (Thermo Fisher Scientific) operated at 200 kV using a Ceta 16M camera (Thermo Fisher Scientific). The screened grids were transferred to a Titan Krios electron microscope (Thermo Fisher Scientific) operated at 300 kV with an energy filter set to a slit width of 20 eV. Images were recorded using a K2 Summit direct electron camera (Thermo Fisher Scientific) in super-resolution mode at a nominal magnification of 130,000 ×, corresponding to a calibrated super-resolution pixel size of 0.5225 Å. The defocus range was set from -1.5 μm to -2 μm.
Each image was acquired as a 7.68 s movie stack (32 frames) with a dose rate of 6.25 e -Å -2 s -1 , resulting in a total dose of about 48 e -Å -2 . All data acquisition was done using SerialEM.
Micrographs with ice or ethane contamination, empty carbon, and poor CTF fit (> 5 Å) were manually removed. All classification and reconstruction was performed with Relion 3.0 (Zivanov et al., 2018) unless otherwise stated. Particles were picked using Gautomatch (developed by Kai Zhang) and subjected to reference-free 2D classification to remove bad particles. Initial models were generated by cryoSPARC (Punjani et al., 2017) using the selected particles from 2D classification. The particles were further subjected to 3D classification to remove bad particles using the initial model, which was low-passed filtered to 30 Å as the reference. The particles selected from good 3D classes were re-centered and reextracted, and their local CTF parameters were individually determined using Gctf v1.06 (Zhang, 2016).
These particles were imported into cisTEM (Grant et al., 2018) and subjected to 3D classification with auto-masking. The particles from the best 3D classes calculated by cisTEM were exported into Relion 3.0 and subjected to 3D auto-refinement to generate the consensus map. Two soft masks that cover the N lobe and C lobe were generated from the consensus map, which was edited manually in UCSF Chimera using the volume eraser tool (Pettersen et al., 2004). 3D multi-body refinements  were performed using the two soft masks of the lobes and the parameters determined from previous 3D autorefinement. The motions of the bodies were analyzed by relion_flex_analyse in Relion 3.0. The two halfmaps of each lobe generated by 3D multi-body refinement were subjected to post-processing in Relion 3.0. The masked and sharpened maps of each lobe were aligned to the consensus map using UCSF Chimera and summed to generate the composite map for visualization and interpretation. All of the resolution estimations were based on the Fourier shell correlation (FSC) of 0.143 cutoff after correction of the masking effect (Chen et al., 2013). B-factors used for map sharpening were automatically estimated by the post-processing procedure in Relion 3.0.

Model building
The position of β1 H-NOX domain was firstly identified according to its distinguishable heme group density. Other domains were assigned by the domain-domain linkers that are visible in the post-processed map ( Figure S2G). The homology models of individual H-NOX, PAS and catalytic domains were generated by Phyre2 server (Kelley et al., 2015) based on the structures of human β1 H-NOX (PDB:5MNW), Manduca sexta α PAS domain (PDB: 4GJ4) (Purohit et al., 2013) and human α1, β1 catalytic domain heterodimer (PDB:3UVJ) (Allerston et al., 2013). The models were placed into the corresponding composite maps using UCSF chimera (Pettersen et al., 2004) and manually rebuilt in Coot (Emsley et al., 2010). The composite maps were then converted into mtz files and the models were further refined by Phenix in reciprocal space (Adams et al., 2010) and Coot in real space. During model building, we found the structure of catalytic module of heme-oxidized state and the heme-unliganded state are essentially the same, but we observed a positive difference density around the βK-αP loop of α1 catalytic domain in the heme-oxidized state sample ( Figure S2H). During the sample preparation of the hemeoxidized state, we supplemented oxidizing reagent NS2028, substrate GTPγS and cofactor Mg 2+ ions into the sGC protein. Therefore, based on the local chemical environment, this positive density might represent Mg 2+ ions together with highly negatively charged phosphate groups that possibly came from the decomposition of the GTPγS molecule. However, this putative phosphate groups were not modeled.

Quantification and statistical analysis
Global resolution estimations of cryo-EM density maps are based on the 0.143 Fourier Shell Correlation criterion (Rosenthal and Henderson, 2003). The local resolution was estimated using relion_postprocess in Relion 3.0 (Zivanov et al., 2018). The number of technical replicates (N) and the relevant statistical parameters for each experiment (such as mean or standard error) are described in the figure legends. No statistical methods were used to pre-determine sample sizes.

Data availability
Cryo-EM maps and atomic coordinates of the heme-unliganded, heme-oxidized, NO-activated and β1

Consensus map
After multi-body refinement

Consensus refinement
Multi-body refinement of N-lobe Multi-body refinement of C-lobe 7.6Å 6.8Å 6.8Å Catalytic module of the NO-activated state   Figure S5. Sequence alignment of the sGC α subunit. Related to Figure 1. The sequences of the Homo sapiens α1 subunit, Homo sapiens α2 subunit, Danio rerio α1 subunit, Danio rerio α2 subunit, and Manduca sexta α1 subunit were aligned. Conserved residues are colored in gray. Residues that are mutated to cysteines for oxidative cross-linking are indicated with black box. Mutations for activity assay are indicated with red box. The residues corresponding to H105 in the β1 subunit are indicated with yellow box. Secondary structural elements are indicated as follows: β sheets by arrows, α helixes by cylinders, and loops by lines. Unmodeled residues are shown as dashed lines. The color of arrows and cylinders are the same as in Figure 1A.  The sequences of the Homo sapiens β1 subunit, Danio rerio β1 subunit, and Manduca sexta β1 subunit were aligned. Conserved residues are colored in gray.
Residues that are mutated to cysteines for oxidative cross-linking are indicated with black box. Mutations for activity assay are indicated with red box. Secondary structural elements are indicated as follows: β sheets by arrows, α helixes by cylinders, and loops by lines. Unmodeled residues are shown as dashed lines. The color of arrows and cylinders are the same as in Figure 1A. (E) Structural comparison between the catalytic module of the full-length sGC in the inactive state (colored) and the isolated catalytic domain heterodimer (gray, PDB ID: 4NI2). The β1 subunit was used for structural alignment. (F) 90º rotated view compared to (E). (G) Structural comparison between the catalytic module of the full-length sGC in the inactive state (colored) and the catalytic domain of the active adenylate cyclase (gray, PDB ID: 1CJU, chain A&B). The β1 subunit was used for structural alignment. The residues of sGC that are within 2.2 Å of the substrate are shown as red spheres. (H) Structural comparison between the NO-activated state (purple) and the inactive state (gray) of the human β1 H-NOX domain. The N-terminal subdomain was used for alignment and the movements are indicated as red arrows. (I) Structural comparison between the NO-activated state (colored) and the inactive state (gray) of the human α1 and β1 PAS heterodimer.)