The crystal structure of the catalytic domain of a eukaryotic guanylate cyclase
© Winger et al; licensee BioMed Central Ltd. 2008
Received: 17 September 2008
Accepted: 07 October 2008
Published: 07 October 2008
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© Winger et al; licensee BioMed Central Ltd. 2008
Received: 17 September 2008
Accepted: 07 October 2008
Published: 07 October 2008
Soluble guanylate cyclases generate cyclic GMP when bound to nitric oxide, thereby linking nitric oxide levels to the control of processes such as vascular homeostasis and neurotransmission. The guanylate cyclase catalytic module, for which no structure has been determined at present, is a class III nucleotide cyclase domain that is also found in mammalian membrane-bound guanylate and adenylate cyclases.
We have determined the crystal structure of the catalytic domain of a soluble guanylate cyclase from the green algae Chlamydomonas reinhardtii at 2.55 Å resolution, and show that it is a dimeric molecule.
Comparison of the structure of the guanylate cyclase domain with the known structures of adenylate cyclases confirms the close similarity in architecture between these two enzymes, as expected from their sequence similarity. The comparison also suggests that the crystallized guanylate cyclase is in an inactive conformation, and the structure provides indications as to how activation might occur. We demonstrate that the two active sites in the dimer exhibit positive cooperativity, with a Hill coefficient of ~1.5. Positive cooperativity has also been observed in the homodimeric mammalian membrane-bound guanylate cyclases. The structure described here provides a reliable model for functional analysis of mammalian guanylate cyclases, which are closely related in sequence.
The second messenger 3',5'-cyclic guanosine monophosphate (cGMP) is central to many signal transduction pathways, primarily eliciting effects by modulating the activities of phosphodiesterases, protein kinases, and ion channels [1–3]. In mammals, cGMP is synthesized by two distinct classes of guanylate cyclases, which are either cytoplasmic or membrane-bound . Both classes of guanylate cyclase share a catalytic module that is closely related in sequence to that of mammalian adenylate cyclases. The catalytic domain is a class III nucleotide cyclase domain , which is distributed widely from bacteria to humans. The class III nucleotide cyclase domain is often found fused to diverse regulatory domains, but is also found as an isolated protein [6–8]. The mammalian membrane-bound guanylate cyclases, which respond to extracellular peptide binding or to the levels of intracellular Ca2+, function in maintenance of fluid homeostasis, inhibition of myocyte hypertrophy, skeletal development, and visual and olfactory signal transduction . The mammalian soluble guanylate cyclases are regulated primarily by binding of nitric oxide (NO), and they modulate a wide range of physiological functions, such as maintenance of vascular tone, platelet aggregation, and neurotransmission . Dysfunction of guanylate cyclase signaling underlies many pathophysiological conditions, ranging from stroke and hypertension to gastrointestinal disease and neurodegeneration [11–13].
Mammalian soluble guanylate cyclases are heme-containing heterodimers of homologous α and β subunits . The N-terminal regulatory domain of each subunit contains a heme-NO and/or oxygen-binding (H-NOX) domain [14, 15], and the H-NOX domains of the β subunits have been shown to bind the heme cofactor [16, 17]. The homologous regions of the α subunits do not bind heme, but are predicted to possess a similar fold. The α and β subunits each contain a central region, shown to be involved in heterodimerization, that consists of an H-NOXA (H-NOX associated) domain and an amphipathic helical extension predicted to form a coiled-coil [18, 19]. The catalytic domain is located in the C-terminal segment of the protein, and it associates with the catalytic domain of the partner subunit to form a heterodimeric catalytic unit [20, 21]. The mechanism of soluble guanylate cyclase activation by NO involves the binding of NO to the heme cofactor , but the details of this activation mechanism are unknown. The response of soluble guanylate cyclase to NO is regulated allosterically by nucleotides [23, 24], but how this happens is also not understood.
The three-dimensional structure of a guanylate cyclase catalytic domain has not been reported. Crystal structures have been obtained for an oxygen-bound H-NOX domain of a methyl-accepting chemotaxis protein from the obligate anaerobe Thermoanaerobacter tencongensis [25, 26] and for NO- and carbon monoxide-bound forms of an H-NOX domain from a histidine kinase operon in the cyanobacterium Nostoc sp. , yielding clues to the mechanism of soluble guanylate cyclase heme ligand recognition and discrimination. Additionally, the crystal structure of the H-NOXA domain of a signal-transduction histidine kinase from Nostoc punctiforme was reported recently , revealing that the dimeric H-NOXA domain adopts a Per/Arnt/Sim (PAS) fold and suggesting a mechanism for the preferential heterodimerization exhibited by mammalian soluble guanylate cyclase. Homology modeling based on crystal structures of the related mammalian and bacterial class III adenylate cyclase catalytic domain dimers has provided some information concerning the structure of the catalytic domain of the guanylate cyclases (reviewed in ).
We have determined the structure of a C. reinhardtii soluble guanylate cyclase catalytic domain dimer by molecular replacement, using the structure of the mammalian adenylate cyclase heterodimer  as a search model. The structure contains one catalytic domain dimer per asymmetric unit. During refinement, we observed unexplained peaks in electron density maps around five of the fourteen cysteine residues in the dimer (Cys 499 and Cys 592 of both monomers, and Cys 621 of monomer B). High concentrations of reductant [5–10 mM dithiothreitol and 10 mM tris(2-carboxyethyl)phosphine] were present during crystallization, making it unlikely that cysteine oxidation or disulfide bond formation are responsible for these features. We wondered whether the apparent modification of the cysteine sidechains might be due to the addition of dimethylarsenic groups via reaction of the cysteine thiol group with the sodium cacodylate [sodium dimethylarsenate, (CH3)2AsO2Na] buffer and dithiothreitol reductant present during crystallization . This chemical modification has been observed previously for several proteins crystallized from solutions containing this buffer [37, 38]; it has even been used to obtain experimental phases for protein structure solution . Accordingly, we confirmed the presence and location of the dimethylarsenic-modified cysteines by taking advantage of the arsenic anomalous signal to calculate an anomalous difference map, which showed unambiguous peaks of electron density around the dimethylarsenic-modified cysteines (see Additional File 1: Dimethylarsenic cysteine modifications).
Crystallographic data and refinement statistics
a = 123.7, b = 123.7, c = 62. 8
α = 90, β = 90, γ = 120
Free R test set (%)
Monomers per A. U.
r.m.s. deviation, bond lengths (Å)
r.m.s. deviation, bond angles (Å)
The structure of the guanylate cyclase catalytic domain differs from that of the adenylate cyclases primarily in the elements that connect strands and helices, and in the less-conserved C-terminal subdomains (Figure 1). Biochemical and structural studies have shown that these regions in the adenylate cyclases couple to regulatory proteins such as the heterotrimeric G-protein subunits Gsα, Giα, and protein kinase C (reviewed in ). Differences in sequence and structure between individual catalytic subunits in heterodimeric adenylate and guanylate cyclases also localize to the same regions (Figure 1).
Modeling based on the adenylate cyclase structures has indicated the mechanism of nucleotide base discrimination [47, 48]. A glutamic acid and a cysteine, conserved in guanylate cyclases, have been proposed to mediate recognition of the exocyclic amine and carbonyl group of the guanine base, respectively. The specificity-determining residues in mammalian adenylate cyclase, a lysine and an aspartic acid, are located at the same relative positions in the adenylate cyclase protein sequence. In fact, swapping those residues into a guanylate cyclase catalytic domain results in conversion into an adenylate cyclase [47, 48], underscoring the equivalence of the catalytic machinery between guanylate and adenylate cyclases. In our structure, the corresponding residues are Glu 523(B) and Cys 592(B), which are situated close to the locations of their adenylate cyclase counterparts (Figure 3b). Local distortions caused by the dimethylarsenic modifications in each monomer appear to prevent optimal sidechain orientation for base recognition. For Cys 592(B), the dimethylarsenic modification of the thiol side chain prevents potential hydrogen bonding interaction with the exocyclic carbonyl group of a substrate GTP molecule (Figure 3b). The modification also results in distortion of the β2–β3 loop that contains the metal-binding residue Asp 527(A) and the base-recognition residue Glu 523(A), causing it to adopt a conformation incompatible with binding a metal-nucleotide complex (Figure 3a). Together, these local distortions would likely preclude any nucleotide binding in the active site, providing a rationale for our failure to visualize nucleotides and metals that have been soaked into the crystals.
Activation of the mammalian adenylate cyclase has been proposed to proceed via two steps, based on structures of active and inactive forms of the protein [35, 41, 49]. In the first step, binding of Gsα between the α1–α2 and α3–β4 loops of the C2 subunit causes a 7° rotation of the core of the C1 subunit around an axis that runs parallel to the central cleft, priming the active site for catalysis by bringing the catalytic residues from one subunit ~2 Å closer to the catalytic residues of the other . The second step involves the closure of the active site around the bound nucleotide. This closure brings structural elements that bind the metal cofactors and the nucleotide triphosphate moiety, and residues in the opposite subunit that orient the ribose ring and stabilize the transition state, into optimal alignment for catalysis  (see Additional file 2: Activation mechanism of mammalian adenylate cyclase). In particular, helix α1 moves towards helix α4 of the opposite subunit, such that a triphosphate interaction site is formed from the helix α1 dipole and a P-loop-like structure between strand β1b and helix α1. The N-terminal end of strand β4, the C-terminal end of strand β1, and the N-termini of helices α2 and α3 also shift towards the active site, properly orienting metal-binding and triphosphate-binding residues for catalysis.
Comparison of our guanylate cyclase catalytic domain structure to structures of the mammalian adenylate cyclase suggests that the guanylate cyclase catalytic domain is in an inactive conformation. The dimethylarsenic modifications described above clearly distort several active site residues. But, in addition to these localized changes, the overall orientation of one subunit with respect to the other corresponds to an open state that must close considerably for catalysis to occur, as we discuss further below.
In the absence of nucleotide, it is not clear in our structure how occupation of one active site by nucleotide is detected by the other. The analysis is also complicated by the local distortions near the active site caused by the dimethylarsenic-modified cysteines. However, inspection of our structure provides two possible mechanisms for communication between the active sites. The first mechanism involves a direct connection between residues in the two active sites. The β2–β3 loop of each monomer carries both the invariant catalytic residue Asp 527 for one active site and the conserved guanine-binding residue Glu 523 for the opposite active site. We propose that the interaction of either Asp 527 or Glu 523 with nucleotide in one active site could alter the conformation of the loop in which they both reside, resulting in a change in nucleotide affinity or enhanced catalysis in the other active site. The second mechanism involves propagation of local changes in one active site, through changes in elements of secondary structure, to the other active site. As noted above, upon substrate binding, helix α1 and strands β1 and β4 in monomer A move inward towards crucial residues in helix α4 of monomer B, bringing residues from both monomers into correct alignment for catalysis in one active site (Figure 4). This movement also brings helix α4 of monomer A, which lies directly above, and packs against, strands β1 and β4 of monomer A, towards monomer B. As helix α4 of monomer A carries residues necessary for catalysis at the other active site, this movement might allow nucleotide binding at one active site to effectively begin pre-organizing the other active site for nucleotide binding.
We report the first structure of a eukaryotic guanylate cyclase catalytic domain. The resemblance of the domain to that of the mammalian adenylate cyclase is unsurprising, given the sequence and functional similarity between them. Nevertheless, more than ten years have elapsed between the first reports of the structures of the adenylate cyclases [35, 41] and our results, presented here. The difficulty in crystallizing a guanylate cyclase domain may reflect an increased intrinsic flexibility in the guanylate cyclase domain relative to the adenylate cyclase domain, and it is possible that we succeeded in part because of the fortuitous cysteine modifications that may have increased the rigidity of the domain, facilitating crystallization. We have been unable as yet to crystallize the catalytic domain in the absence of these modifications.
The high degree of sequence conservation between the soluble guanylate cyclase catalytic domain described here and the catalytic domains of mammalian soluble and membrane-bound guanylate cyclases (40 to 50% identity) suggests that our structure will serve as a superior model for functional studies, compared to the mammalian adenylate cyclase catalytic domains (25 to 30% sequence identity). Our structure indicates that the differences between the adenylate and guanylate cyclase are generally localized to flexible regions, some of which are proposed to mediate coupling with regulatory domains and other control elements. While specific differences in regulatory interactions are likely determined by the sequence and local structure of these variable elements, the overall activation mechanism, involving conformational switching by helix α1 and attendant changes in the adjacent β sheet, is expected to be conserved.
PCR was used to amplify the gene encoding a 991-residue soluble guanylate cyclase homolog CYG12 (GenBank: XM_001700795) from a C. reinhardtii cDNA library obtained from the Chlamydomonas Center . Forward and reverse PCR primers were 5'-ATGCTGGGCTGGTATGACCGT-3' and 5'-TTACTCCAAACACGGGTTGTCA-3', respectively. PCR products were phosphorylated, blunt-cloned into the vector pGEM, and verified by sequencing (UC Berkeley DNA Sequencing Facility). The guanylate cyclase catalytic domain was expressed and purified using a SUMO-based system (LifeSensors) as follows: residues 468–655, which comprise the catalytic domain of CYG12, were subcloned into a vector containing the yeast SUMO homolog SMT3 with an N-terminal His-tag, and the fusion protein was expressed in Escherichia coli Tuner(DE3) (Invitrogen) for 18 h at 20°C. The fusion protein was purified from supernatant by passage over a HisTrap Ni Sepharose affinity column (GE Healthcare). A His-tagged version of the SMT3-specific protease was used to cleave the N-terminal SMT3 fusion partner from the guanylate cyclase domain, which was separated from the protease and SMT3 by a second Ni affinity step. The guanylate cyclase domain was further purified by Q Sepharose anion-exchange chromatography, followed by gel filtration into a final buffer of 25 mM triethanolamine, pH 7.5, 25 mM NaCl, and 10 mM dithiothreitol. Purified protein was concentrated and stored at -20°C until use. Protein concentrations were determined by absorbance using the calculated extinction coefficient ε280 = 7680 M-1 cm-1.
Crystals were grown using the sitting-drop vapor diffusion method. Equal volumes (200 nl) of protein [40–60 mg/ml in 25 mM triethanolamine, pH 7.5, 25 mM NaCl, 5–10 mM dithiothreitol, 10 mM tris(2-carboxyethyl)phosphine] were mixed with crystallization solution [0.1 M sodium cacodylate, pH 5.0–6.4, 42–62% saturated (NH4)2HPO4] and then equilibrated with a 100-μl reservoir of the same crystallization buffer at 20°C. Crystals grew in the trigonal space group and appeared within 1–2 days. Crystals were transferred to a solution of mother liquor containing 28% glycerol as a cryoprotectant, and cryo-cooled and stored in liquid nitrogen. Diffraction data were collected at 100 K using synchrotron radiation at beam line 8.2.2 at the Advanced Light Source, Lawrence Berkeley National Laboratory. Reflections were integrated and scaled with the programs MOSFLM  and SCALA . The structure was solved by molecular replacement with the program PHASER  using the mammalian adenylate cyclase catalytic domain (PDB code 1AZS)  as the search model, and the space group was identified as P3221. Map improvement was carried out using ARP/wARP  and RESOLVE . The model was built using COOT  and refined using PHENIX . Six TLS domains were used during refinement: residues 467–475 and 578–595, monomer A/B; residues 476–560, 569–577, and 596–608, monomer A/B; and residues 613–651, monomer A/B. Analysis of model quality was carried out using MOLPROBITY . Figures were prepared using PYMOL . The atomic coordinates and structure factors have been deposited in the Protein Data Bank (3ET6).
Guanylate cyclase assays were performed in duplicate at 24°C as described previously . Assays contained 5 μg of guanylate cyclase in 25 mM triethanolamine, pH 7.5, 25 mM NaCl, 4 mM MnCl2 or MgCl2 and 5 mM dithiothreitol. Assays were initiated by addition of indicated amounts of GTP, and were quenched after 2 minutes by addition of 400 μl of 125 mM Zn(CH3CO2)2 and 500 μl of 125 mM Na2CO3. cGMP was quantified using a cGMP enzyme immunoassay kit, format B (Biomol), per the manufacturer's instructions. Experiments were repeated three times to ensure reproducibility.
We would like to thank Carmen Schwechheimer, Caleb Cassidy-Amstutz, and Bianca Lee for technical assistance, and Nicholas Levinson and Markus Seeliger for critical reading of the manuscript. This work was supported in part by NIH grant GM077365 (M.A.M.). J.A.W. was supported by an American Heart Association Postdoctoral Fellowship. The Advanced Light Source is supported by the U.S. Department of Energy under Contract DE-AC03-76SF00098 at the Lawrence Berkeley National Laboratory.
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