Design, synthesis and biological evaluation of new 3,4-dihydroquinoxalin-2(1H)-one derivatives as soluble guanylyl cyclase (sGC) activators

Herein, we present the structure-based design, synthesis and biological evaluation of novel mono- and di-carboxylic 3,4-dihydroquinoxalin-2(1H)-one derivatives as potential heme-independent activators of soluble guanylate cyclase (sGC). Docking calculations of several known sGC agonists by utilizing both a homology model of human sGC β1 Η-ΝΟΧ domain and a recent cryo-EM structure of the same domain guided the structural optimization of various designed compounds. Among these, mono- and di-carboxylic 3,4-dihydroquinoxalin-2(1H)-one derivatives arose as promising candidate sGC activators. A series of such compounds was synthesized and assessed for their effect on sGC activity. None of them was able to trigger any detectable activation of native sGC in prostate cancer (LnCaP) or rat aortic smooth muscle (A7r5) cells, even after loss of heme by treatment with the heme oxidant ODQ. Furthermore, selected derivatives did not exhibit any antagonistic effect against the known heme-independent sGC activator BAY 60-2770 nor any additive or synergistic effect with the heme-dependent NO donor sodium nitroprusside (SNP) on heme-associated sGC in A7r5 cells. However, when tested in vitro using purified recombinant sGC enzyme, the dicarboxylic 3,4-dihydroquinoxalin-2(1H)-one derivative 30d was able to increase the enzymatic activity of both the wild-type α1/β1 sGC dimer (by 4.4-fold, EC50 = 0.77 μΜ) as well as the heme-free α1/β1 His105Ala mutant sGC (by 4.8-fold, EC50 = 1.8 μΜ). Notably, the activity of compound 30d towards the mutant α1/β1 Η105A enzyme was comparable with that previously reported by us for the bona fide activator BAY 60-2770, using the functionally equivalent wild-type sGC preparation treated with ODQ. These results indicate that compound 30d can indeed act as a promising sGC activator and may serve as a basic structure in the design of novel, optimized analogues with enhanced sGC agonistic activity and improved efficiency in cell-based and in vivo systems.


Novel
3,4-dihydroquinoxalin-2(1H)ones were designed and synthesized. Compounds were evaluated for activation of sGC in enzyme-and cell-based assays. Analogue 30d increased the activity of both the wild-type and mutant His105Ala sGC. 30d induces maximum enzyme activity comparable with that of activator BAY 60-2770.

Keywords:
Nitric oxide Soluble guanylyl cyclase sGC activators 3,4-Dihydroquinoxalin-2(1H)-one A B S T R A C T Herein, we present the structure-based design, synthesis and biological evaluation of novel mono-and dicarboxylic 3,4-dihydroquinoxalin-2(1H)-one derivatives as potential heme-independent activators of soluble guanylate cyclase (sGC). Docking calculations of several known sGC agonists by utilizing both a homology model of human sGC β1 Н-NΟХ domain and a recent cryo-EM structure of the same domain guided the structural optimization of various designed compounds. Among these, mono-and di-carboxylic 3,4-dihydroquinoxalin-2(1H)-one derivatives arose as promising candidate sGC activators. A series of such compounds was synthesized and assessed for their effect on sGC activity. None of them was able to trigger any detectable activation of native sGC in prostate cancer (LnCaP) or rat aortic smooth muscle (A7r5) cells, even after loss of heme by treatment with the heme oxidant ODQ. Furthermore, selected derivatives did not exhibit any antagonistic effect against the known heme-independent sGC activator BAY 60-2770 nor any additive or synergistic effect with the hemedependent NO donor sodium nitroprusside (SNP) on heme-associated sGC in A7r5 cells. However, when tested in vitro using purified recombinant sGC enzyme, the dicarboxylic 3,4-dihydroquinoxalin-2(1H)-one derivative 30d was able to increase the enzymatic activity of both the wild-type α1/β1 sGC dimer (by 4.4-fold, EC 50 ¼ 0.77 μМ) as well as the heme-free α1/β1 His105Ala mutant sGC (by 4.8-fold, EC 50 ¼ 1.8 μМ). Notably, the activity of compound 30d towards the mutant α1/β1 Н105A enzyme was comparable with that previously reported by us for

Introduction
Soluble guanylyl cyclase (sGC), a α/β-heterodimeric enzyme, is the primary receptor of endogenously produced nitric oxide (NO) or of NO delivered by donor molecules and catalyzes the conversion of guanosine triphosphate (GTP) to the second messenger cyclic guanosine monophosphate (cGMP) [1]. The prosthetic heme moiety, located in the Heme-NO OXygen domain (H-NOX) of the regulatory β 1 subunit, binds NO with high affinity, triggering an increase of catalytic activity [2]. Through this mechanism, the NO/sGC/cGMP pathway promotes vasodilation and inhibits smooth muscle proliferation, platelet aggregation and negative vascular remodeling by modulating the activity of a number of downstream targets, such as protein kinases, cGMP-gated ion channels and phosphodiesterases [3]. There is significant preclinical and clinical evidence that dysregulation of this signaling pathway, either by impaired NO bioavailability or by heme oxidation and conversion of sGC to its NO-unresponsive apo-enzyme state in oxidative stress conditions, contributes to various disorders, such as pulmonary hypertension, heart failure, glaucoma and chronic kidney disease [4]. During the last two decades, novel treatments of the aforementioned disorders have targeted the sGC, aiming to restore or augment its enzymatic activity.
Through these efforts, two new classes of compounds that are capable of activating sGC in an NO-independent manner have been discovered, distinct from NO donors; heme-dependent sGC "stimulators" and hemeindependent sGC "activators" (Figure 1) [5]. Because of their broad clinical potential, sGC stimulators attract increasing interest from research groups and pharmaceutical companies [6,7], exemplified by two approved per os drugs, riociguat and vericiguat [8,9]. Riociguat (Adempas ® ) was approved in 2013 for the treatment of two forms of pulmonary hypertension, pulmonary arterial hypertension (PAH) and chronic thromboembolic pulmonary hypertension (CTEPH) [10,11,12,13]. More recently, FDA approved vericiguat (Verquvo ® ) for the treatment of heart failure with reduced ejection fraction (HFrEF) [14,15]. Besides these two compounds, a rising number of sGC stimulators have been introduced in clinical studies for the treatment of various diseases, e.g., praliciguat is in clinical phase II for the treatment of heart failure with preserved ejection fraction (HFpEF) and diabetic nephropathy, whereas olinciguat is in clinical phase II trials for the treatment of achalasia and sickle cell disease [16].
Despite the progress in the clinic, there is still insufficient information regarding the mechanism of action of stimulators. Recent studies, involving Nuclear Resonance Spectroscopy (NMR) and cryogenic Electron Microscopy (cryo-EM) methods, applied to complexes of known sGC stimulators with either bacterial H-NOX domains or with human sGC, pointed to two main hypotheses regarding their binding mode to the enzyme. The first postulates that the stimulators' binding pocket resides in the β1 Н-NΟХ domain, whereas the second suggests that the binding pocket is formed by the interface of three different domains: the β1 Н-NΟХ, the α1 subunit and the β1 Coiled-coil (CC) domain [17,18].
In comparison, sGC activators' mode of action has been better characterized in numerous publications over the years [19,20]. All of these studies are in agreement with the hypothesis that these compounds activate the NO-insensitive forms of sGC through interaction with residues that bind heme in a pocket located in the β1 Н-NΟХ domain. More specifically, the development of crucial hydrogen-bonding interactions with Tyr 135 , Ser 137 and Arg 139 , also known as the Y-S-R motif, mimics those formed by the two propionic groups of the heme moiety.
In contrast to the stimulators, the clinical development of sGC activators has encountered several challenges. A typical example is Cinaciguat, the most clinically advanced sGC activator. Phase IIb clinical trials of this compound, administered intravenously for the treatment of acute heart failure, were terminated due to a marked hypotensive adverse effect observed even at low doses [21]. Another interesting sGC activator, ataciguat, which reached Phase II clinical trials for aortic valve stenosis, was also discontinued.
In an effort to reduce such systemic effects, the development of current sGC activators focuses, among others, on other pathophysiological indications, where local application is a possibility. Recent examples are ocular instillation of MGV354 and inhalation of BAY 1211163, which were introduced in Phase II clinical studies for the treatment of openangle glaucoma and acute respiratory distress syndrome, respectively [22,23].
Another interesting approach to deal with the poor clinical profile of sGC activators was presented by Elgert et al. [24]. Based on this, second generation of sGC activators, mainly consisting of monocarboxylic derivatives, act as partial agonists of sGC through the formation of a non-stabilizing dead-end complex with sGC. Due to their binding and activation profile, monocarboxylic sGC activators may not cause hypotension, as dicarboxylic derivatives do. This hypothesis was further supported by point mutation studies of the Y-S-R motif residues, which indicated differences in the binding mode between a full (BAY 60-2770) and a partial sGC agonist (BAY-543). The first formed hydrogen-bonding interactions with all three Y-S-R motif residues, whereas the second selectively developed hydrogen-bonding interactions with Tyr 135 only [25]. Based on the aforementioned pharmacological studies and focusing on a sGC activator with improved pharmacokinetic properties, Bayer recently identified the orally administered monocarboxylic acid runcaciguat via high-throughput screening. This second generation sGC activator is currently investigated in a clinical phase II trials in patients with chronic kidney disease and nonproliferative diabetic retinopathy [26].
Motivated by these observations, we decided to focus on the discovery of novel sGC activators through a structure-based design approach. Initially, we performed in silico analysis of the binding mode of known full and partial sGC agonists in the heme binding pocket. The absence of PDB-submitted complexes of these compounds with the human sGC β1 Н-NΟХ domain led us to the utilization of computational methods. Thus, a homology model of human sGC β1 Н-NΟХ domain was constructed and used docking calculations of known sGC activators. Detailed analysis indicated crucial structural features of these molecules that were involved into residue-specific interactions with the β1 Н-NΟХ domain. Based on these findings, further docking calculations of several designed compounds were performed, among which monocarboxylic 3,4-dihydroquinoxalin-2(1H)-one derivatives exhibited promising binding modes. Further attempts to increase the binding affinity of these derivatives through enhancement of hydrogen-bonding and van der Waals interactions with the Y-S-R motif and a previously unoccupied hydrophobic pocket, led to the design of potentially more effective dicarboxylic 3,4-dihydroquinoxalin-2(1H)-ones. Herein, we describe the results of our computational studies, the synthesis of selected designed compounds and the evaluation of their ability to stimulate sGC activity.

Computational studies and design
Homology modeling of human β1 H-NOX domain (hH-NOX) was carried out using the X-ray crystal structure of the corresponding domain from the cyanobacterium Nostoc sp. as template (NsH-NOX, PDB ID: 2O09) [27]. Based on their sequence identity of 35.1% ( Figure S1) an acceptable homology model of hH-NOX domain was obtained using Modeller [28]. Subsequently, the homology model was subjected to energy minimization using UCSF Chimera [29]. Superimposition of the 3D structures obtained before and after the energy minimization indicated subtle changes, mostly at the helices forming the heme binding pocket ( Figure S4). Before docking of the designed compounds, we assessed the performance of AutoDock (v.4.2.6) [30] to identify the binding pose of the heme prosthetic group. Indeed, AutoDock predicted the crystallographic pose of NsH-NOX heme within a mean root-mean-square deviation of 0.7 Å in 99/100 docking rounds and with a mean estimated free energy of binding (ΔG est ) of -17.9 kcal/mol. Inspection of the complex revealed hydrogen-bonding interactions between heme's propionate groups and residues Tyr 2 , Tyr 135 , Ser 137 , Arg 139 , as well as van der Waals interactions with residues of helix aF (Leu 101 , Leu 104 , His 105 , Leu 108 ) and helix aG (Leu 142 , Ile 145 , Val 146 , Ile 149 ).
Subsequently, we carried out molecular docking of two known activators with hH-NOX, ataciguat (HMR-1766) and cinaciguat. The topranked cluster of ataciguat (54% population, mean ΔG est ¼ -10.23 kcal/mol) displayed hydrogen-bonding interactions between the amide -NH of the ligand and the imidazole side chain of His 105 , and between the morpholine ring of ataciguat and the side chain of Arg 139 . A number of van der Waals interactions were also formed with residues of the aF and aG helices, similar to those formed by heme ( Figure S5). In contrast to ataciguat, the high flexibility of cinaciguat (7 and 17 rotatable bonds respectively) gave rise to a large number of different bound conformations (17 clusters with >1 pose). Visual investigation of the 2 top-ranked clusters (12% and 6% populations with mean ΔG est of -13.97 kcal/mol and -13.24 kcal/mol, respectively) revealed hydrogen-bonding interactions between the carboxylate groups of cinaciguat and the Y-S-R motif ( Figure S6), in accordance with previously published data. 19 Hydrophobic-aromatic interactions between the phenoxy or terminal phenethyl moiety of cinaciguat and residues of helices αF and αG were also formed.
The increased flexibility of known sGC activators within the heme binding pocket, combined with their moderate mean binding energies, led us to design new compounds with rigid heterocyclic skeletons that can be functionalized appropriately. First, considering the relatively short distance between the amide and sulfonamide -NH groups of ataciguat (equivalent to 2-3 C-C bonds) we decided to incorporate them into a fused bicyclic heterocycle. Then, we incorporated carboxylic substituents in specific positions of the bicyclic scaffold in order to enhance hydrogen-bonding interactions with the Y-S-R motif residues ( Figure S7A). In an attempt to exploit the available hydrophobic cavities of hH-NOX ( Figure S7B), we introduced suitably substituted heteroaromatic rings at either NH positions of the skeleton.
Furthermore, the spatial arrangement of the -NH groups in the bicyclic skeleton was probed aiming at the optimal orientation of their substituents within the heme binding pocket. On this basis we designed 4 compounds and performed docking calculations (C1-C4, Table S1). Comparison of their top-ranked poses indicated the 3,4-dihydroquinoxalin-2(1H)-one derivative C4 as the most promising ligand (28% population, mean ΔG est ¼ -10.60 kcal/mol). As predicted for cinaciguat, C4 displayed hydrogen-bonding interactions between the amide-attached carboxylate and residues Tyr 2 , Arg 116 and Arg 139 (Figure 2A, B). Additionally, the pyrrole substituent, attached to the secondary amide through amide linkage, displayed similar van der Waals interactions with Leu 101 (aF helix) and Val 146 , Ile 149 (aG helix) as those observed for heme.
We also investigated the potential replacement of the pyrrole moiety of C4 by other heteroaromatic rings with the designed compounds C5-C9. However, their docking results did not display any significant improvement of the estimated binding affinities (Table S1). Therefore, we maintained the pyrrole substituent as a structural feature of our compounds during the optimization process and we planned to synthesize monocarboxylic 3,4-dihydroquinoxalin-2(1H)-one derivatives decorated with selected heteroaromatic substituents at the N4 position of the skeleton to gain insights into structure activity relationships.
In order to enhance binding affinity of C4, a thorough investigation of the heme binding site was undertaken. During this process, two previously unoccupied cavities by C4 were revealed. These involve, a hydrophilic cavity, that incorporates residues Tyr 135 , Ser 137 of the Y-S-R motif, as well as a hydrophobic one, formed mostly by alkyl and aromatic side chains of residues, situated in helices αA and αF ( Figure 2C). In an attempt to exploit the hydrophilic cavity, the decoration of the 3,4dihydroquinoxalin-2(1H)-one skeleton involved the introduction of several phenyl-or phenoxy-substituents with an additional carboxylic group, at position C7 of the basic skeleton. The newly designed compounds C10-C13 were introduced to docking studies (Table S1). Among  them, dicarboxylic 3,4-dihydroquinoxalin-2(1H)-one derivative C13 exhibited enhanced binding affinity, as the 4-oxy-benzoic acid substituent at C7 presented optimal proximity to the aforementioned residues. Furthermore, the addition of a phenyl group at position C6 of the basic skeleton contributed to the efficient exploitation of the hydrophobic cavity and led to the optimized dicarboxylic derivative C14, which showed increased stability (population C4 ¼ 29% vs C14 ¼ 70%) and binding affinity (mean ΔG est C4 ¼ À10.60 kcal/mol vs C14 ¼ À15.65 kcal/mol) within the heme binding pocket. The improved binding mode of C14 were basically attributed to the increased amount of hydrogenbonding and van der Waals interactions with residues of heme binding pocket ( Figure 3A). Notably, superimposition of the docked conformations of heme and C14 revealed a similar orientation, especially regarding the two carboxyl groups of C14, which are crucial for the establishment of a hydrogen-bonding network with the Y-S-R motif ( Figure 3B, C).
Interestingly, during the course of this project, various cryo-electron microscopy (cryo-EM) structures of human sGC were resolved, in active and inactive state [31,32]. We decided to verify our initial design rationale by utilizing this available information and conducted docking calculations of the known sGC activators and designed compounds C1-C14 with β1 H-NOX domain, obtained from a cryo-EM structure of human sGC in the heme-oxidized state (PDB ID: 6JT1) [31], which is targeted by sGC activators. Analysis of the results revealed a similar classification of the known and designed compounds based on their mean estimated binding energy and population of their top-ranked clusters (Table S1), supporting our scaffold optimization strategy. Notably, the interaction analysis of selected designed compounds (e.g. C14) bound to the heme binding pocket revealed their similar orientation in the heme cavity as well as the formation of hydrogen-bonding interactions with the Y-S-R motif, especially with Arg 139 , and other residues nearby, such as Tyr 2 , Arg 116 and Glu 138 ( Figure S8). Based on these results, we proceeded to the synthesis of a series of mono-and di-carboxylic 3,4-dihydroquinoxalin-2(1H)-one derivatives which incorporated structural features, capable of promoting binding properties and activation of sGC, as it was indicated by our molecular design studies. Thus, various substituents were introduced at position N4, C6 and C7 of the bicyclic quinoxalin-2(1H)-one skeleton of the target molecules, depending on the synthetic challenges might emerged from their introduction, in order to elucidate the relationships between the ligand estimated binding affinities and the sGC agonistic activity.
Another route, outlined in Scheme 2, was employed for the appropriate decoration of the 3,4-dihydroquinoxalin-1(2H)-one core that eventually would furnish the target dicarboxylic compounds. Thus, commercially available 3-bromoaniline 10 was introduced in a series of previously reported transformations, including p-bromination (11), amine acetylation (12), o-nitration (13) and acetamide acidic hydrolysis, to furnish the 4,5-dibromo-2-nitroaniline 14 with a total yield of 51% [35]. Subsequently, selective etherification of 14 at C5 position was achieved by treatment with freshly prepared benzyl 4-hydroxybenzoate (23) affording the diphenyl ether 15 [36]. Next, chloroacetylation of 15 to the anilide 16, followed by reduction of the nitro group using metallic iron in acidic conditions, provided the substituted aniline 17. The latter was introduced to a halogen-exchange Finkelstein reaction using sodium iodide and the generated intermediate alkyl iodide was cyclized in situ via intramolecular nucleophilic attack by the amine group to the resulting substituted quinoxalin-2(1H)-one derivative 18. Upon treatment with 9 and NaH, N-benzylated lactam 19 was obtained in 68% yield. Finally, imine reduction with sodium cyanoborohydride afforded the 3,4-dihydroquinoxalin-1(2H)-one derivative 20, while Suzuki cross-coupling with phenylboronic acid followed by imine reduction furnished the corresponding C7-phenylated derivative 22.
Further diversification of the dihydroquinoxalinone cores 4, 8, 20 and 22 was achieved by amide coupling of secondary amine with several heteroaromatic acyl chlorides (24)(25)(26) in the presence of triethylamine as base. Following this methodology, amide derivatives 27(a-e) and 29(a-e) were obtained with overall good yields (23-99 %). Finally, cleavage of the benzyl ester protective group by applying catalytic hydrogenolysis conditions using ammonium formate and palladium on activated carbon as catalyst furnished the target mono-and dicarboxylic 3,4-dihydroquinoxalin-1(2H)-one derivatives, 28(a-e) and 30(a-e), respectively. Notably, the deprotection of di-benzyl ester derivatives 29(a-b) was accompanied by unavoidable debromination of the skeleton (Scheme 3).

Evaluation of bioactivity
The biological activity of the synthesized derivatives was evaluated via a cGMP ELISA assay in LNCaP and A7r5 RAoSM cells, which naturally express the soluble guanylate cyclase (sGC) α 1 β 1 isoenzyme [37,38]. The  assay was performed both in cells pretreated or not with the heme oxidant 1H-[1,2,4]oxadiazolo[4,3-a]quinoxalin-1-one (ODQ), which has been previously described to potentiate the effect of heme-and NO-independent activators on the function of the sGC, while it reduces those of heme-dependent activators, such as the NO donor sodium nitroprusside (SNP) [39]. Results were determined as cGMP levels normalized for total protein (pmol/mg) in the respective well and are represented as means AE SD of fold-induction relatively to untreated (Control, CTL) cells.

Bioactivity of monocarboxylic 3,4-dihydroquinoxalin-1(2H)-one derivatives in LnCaP cells
In LNCaP cells, the oxidant ODQ had no significant effect by itself on basal cGMP levels. Both the NO donor SNP and the heme-independent sGC activator BAY 60-2770 raised cGMP levels similarly in ODQuntreated (naïve) cells. As expected, the effect of SNP was abrogated by pretreatment with ODQ, while the effect of BAY 60-2770 was enhanced ( Figure 4). None of the monocarboxylic derivatives tested elicited a significant response, either in the absence or presence of ODQ. Therefore, at the concentrations and cellular background we conducted our tests, the synthesized molecules are unable to trigger by themselves detectable activation of sGC. Indicatively, we present the results of the compounds 28a, 28c and 28d in Figure 4.

Bioactivity of dicarboxylic 3,4-dihydroquinoxalin-1(2H)-one derivatives in A7r5 RAoSM cells
The oxidant ODQ did not significantly affect basal cGMP levels in A7r5 RAoSM cells, either. The control sGC agonists, SNP and BAY 60-2770, exhibited an identical pattern of biological activity and sensitivity to ODQ to that seen in LNCaP cells. None of the dicarboxylic derivatives tested elicited a raise in cGMP levels, either on naïve or ODQtreated cells. Indicatively, we present the results of the compounds 30b, 30d and 30e in Figure 5.

Bioactivity of new 3,4-dihydroquinoxalin-1(2H)-one derivatives on purified recombinant sGC
We hypothesized that the inactivity of our 3,4-dihydroquinoxalin-1(2H)-one derivatives to act as sGC agonists in cell-based assays might be due in part to suboptimal physicochemical properties, such as impaired cellular permeability, and therefore we decided to test their activity in vitro using purified recombinant sGC enzyme [40]. To this end, we assayed selected derivatives which were substituted at position N1, N4, C6 and C7 of the 3,4-dihydroquinoxalin-1(2H)-one skeleton with moieties which may display enhanced sGC binding affinity, such as the monocarboxylic derivatives 28a and 28e, and the dicarboxylic compounds 30b and 30d. Experiments were performed in preparations of the wild-type α1/β1 as well as with the heme-free α1/β1 His105Ala mutant enzyme [40]. Results are shown as sGC activity expressed in nmol cGMP/(mg x min) generated, as means AE SD of three independent experiments (Table 1).
Among the four tested compounds, the dicarboxylic derivative 30d bearing a N4 furanoyl, a C6 phenyl and a C7 4-carboxyphenoxy moiety, exhibited potent sGC agonistic activity, followed by 30b, whereas the monocarboxylic derivatives displayed no activity. However, the latter molecule showed statistically-significant activity only on the mutant enzyme and only at the highest concentration (10 μМ). Specifically, compound 30d increased the activity of the wild-type sGC α1/β1 enzyme by as much as 4.4-fold (from 61.59 AE 15.45 nmol cGMP/mg x min to 271.0 AE 113.8 nmol cGMP/mg x min, at 10 μМ), exhibiting an apparent EC 50 value ¼ 0.77 μМ ( Figure 8A). Crucially, the ability of 30d to increase the activity of the heme-free sGC α1/β1 His105Ala mutant enzyme was undiminished, since it raised sGC activity by 4.8-fold (from 112.0 AE 63.38 nmol cGMP/mg x min to 541.5 AE 55.21 nmol cGMP/mg x min) at the same maximal concentration, with an apparent EC 50 value ¼ 1.8 μМ ( Figure 8B).
Comparing the activity of dicarboxylic derivatives 30b and 30d, it is clearly evident that not only the N1 and C7 carboxylic substituents, but also the presence of the C6 phenyl group significantly improved the sGC agonistic activity. These results are in agreement with the information obtained from docking simulations of designed analogues, where C6phenylated dicarboxylic derivative C14 presented better binding mode compared to the non-phenylated one, C13. This observation is also in line with our applied design approach which indicated that the introduction of an aromatic moiety at the C6 position of the 3,4-dihydroquinoxalin-1(2H)-one skeleton could not only enhance the binding affinity through efficient van der Waals interactions with specific residues of the heme binding pocket, but also improve the activation of sGC elicited by this compound.
It is noteworthy that the compound 30d induces maximum enzyme activity at the highest-tested concentration (10 μM) comparable with that of the bona fide activator BAY 60-2770, as it was determined previously by us using the functionally equivalent (i.e. heme-free) ODQ-treated wild-type enzyme [24].

Conclusions
In summary, guided by a structure-based design approach we synthesized appropriately substituted mono-and di-carboxylic 3,4-dihydroquinoxalin-2(1H)-one derivatives as potential sGC activators, i.e. molecules capable of stimulating the heme-free enzyme. Although the initial design strategy was based on a homology model of hH-NOX, we have verified our approach using a recent cryo-EM structure of the same domain. A series of synthesized compounds were subsequently evaluated in intact LnCaP prostate cancer or A7r5 vascular smooth muscle cells, however we were unable to detect enzyme activation in either the absence or presence of the prosthetic heme group, effected by the use of the heme-oxidant ODQ. Furthermore, the monocarboxylic derivative 28e and the dicarboxylic compound 30d, both at 3-fold excess concentrations, did not exhibit any antagonistic effect against the hemeindependent sGC activator BAY 60-2770 in the presence of ODQ, whereas derivative 30d did not display any additive or synergistic effect with the known NO donor SNP on the heme-associated enzyme in A7r5 RAoSM cells.
In contrast, among four selected mono-and dicarboxylic derivatives, the dicarboxylic molecules compounds 30b and 30d alone upregulated the activity of the purified recombinant sGC enzyme in vitro. Significantly, 30d increased the activity of both the wild-type α1/β1 (4.4-fold) as well as the mutant α1/β1 His105Ala (4.8-fold) sGC at the maximal concentration tested (10 μМ), with EC 50 values of 0.77 and 1.8 μМ, respectively. It is also noteworthy that the ability of 30d to stimulate the α1/β1 Н105A (heme-free) enzyme is comparable with that of the known activator BAY 60-2770, reported previously by us, against the functionally equivalent ODQ-treated wild-type purified enzyme (i.e. in the absence of the heme moiety), over the same range of concentrations [24]. These last bioactivity data indicate that the addition of appropriate   substituents at positions N1, N4, C6 and C7 of the 3,4-dihydroquinoxalin-2(1H)-one skeleton, as indicated by our structure-based design approach, may result to generation of new compounds with sGC agonistic activity characteristic of sGC "activators". The future incorporation of optimal structural features to the new 3,4-dihydroquinoxalin-2(1H)-one derivatives is expected not only to enhance their sGC binding and agonistic properties, but also to improve some of their suboptimal physicochemical characteristics, which may account for the lack of their activity in the cell-based assays. Undoubtedly, among all compounds presented herein, the dicarboxylic 3,4-dihydroquinoxalin-2(1H)-one 30d seems the most promising starting point for the design of additional, optimized analogues with enhanced sGC agonistic activity and improved efficiency in intact cells.

Computational methods
The coordinates of the NsH-NOX crystal structure in complex with heme were retrieved from the Protein Data Bank with accession code: 2O09 [27]. Residues 1-182 from chain A were used as template for the modeling of hH-NOX domain residues 1-184 (UniProt assessing code: Q02153) according to the sequence alignment obtained with Clustal Omega ( Figure S1). Modeller v9.14 was used to produce 200 models of hH-NOX and the model with the lowest DOPE score was selected [28]. Its quality was assessed using the Structural Analysis and Verification Server v6.0 (https://saves.mbi.ucla.edu/), which revealed a well-defined model ( Figure S2). Next, hH-NOX coordinates were superimposed with the template structure of NsH-NOX using the MatchMaker extension of UCSF Chimera 1.10.1 [29], for the preparation of hHNOX complex with the heme moiety at its crystallographic position. The complex was subjected to energy minimization using the AmberTools extension of UCSF Chimera 1.10.1 [29], which was carried out with 1,000 and 4,000 steps of the steepest descent and the conjugate gradient methods, respectively (quality assessment in Figure S3). The energy-minimized model of hH-NOXheme complex was used without heme for the molecular docking calculations of Ataciguat, Cinaciguat and of the designed compounds C1-C14. Structure coordinates of human sGC complex with heme moiety in oxidized state were retrieved from the Protein Data Bank with accession code: 6JT1 [31]. Residues 1-184 from chain B were used without heme for the molecular docking calculations of the designed compounds C1-C14.
The initial 3D coordinates of ligands were obtained from SMILES representations using the program OMEGA v4.0.0.4 [41]. Protein and ligands were assigned with united-atom types and Gasteiger partial charges using AutoDockTools v1.5.7. The search space was defined at the center of the heme-binding cavity as a grid box of 18.75 Å in each dimension (50 points Â 0.375 Å spacing). One hundred docking rounds were performed with AutoDock v.4.2.6 using the Lamarckian genetic algorithm with the default parameters [30,42]. The maximum number of energy evaluations was set to 2.5Â10 6 and the resulting conformations were clustered within a 2-Å cutoff. Hydrogen-bonding and van der Waals interactions were visually inspected using Discovery Studio Visualizer v20.1.0.19295 [43], whereas rendering of figures was performed with PyMOL 2.4.0 [44].

Chemistry
All reagents were obtained commercially from Alfa Aesar, Sigma-Aldrich or Merck and used without further purification. Reactions involving moisture-sensitive reactants were conducted in flame-dried glassware under an atmosphere of argon. Reagents and anhydrous solvents were transferred via syringe. Analytical TLC was performed on Merck Silica gel 60 F254 on precoated silica gel plates. Visualization was accomplished under UV light (254 and 365 nm), by exposure to iodine vapors and by use of Seebach staining solution. Flash column chromatography was performed on silica gel (SDS 60A, 40-63 lm). Melting points were determined on an Haake Buchler melting point apparatus in capillary tubes and are uncorrected. The IR spectra were recorded on a FT-IR Jasco spectrophotometer and a FT-IR Spectrum100 Perkin Elmer spectrophotometer equipped with KBr pellet accessory. NMR spectra ( 1 H and 13 13 C) Avance III HD Ascend TM spectrometer. The chemical shifts (δ) are reported in parts per million (ppm) and the residual solvent signal is used as an internal standard. The following abbreviations are used for the proton spectra multiplicities: singlet (s), doublet (d), triplet (t), quartet (q), multiplet (m), broad signal (br) and combinations thereof. Coupling constants (J) are given in Hertz (Hz). Mass spectra were obtained on a single quadrupole LC/MS system of Agilent LC/MSD1260 Infinity II equipped with an ESI ion source.

7-Bromoquinoxalin-2(1H)-one (5) [33]
To a solution of 2 (5 g, 34.21 mmol) in acetic acid (165 mL) was added a solution of bromine (1.75 mL, 34.21 mmol) in acetic acid (103 mL), and then the resulting mixture was stirred at rt for 21 h. The reaction mixture was poured into a mixture of ice and water, the resulting precipitate was collected by filtration, washed with H 2 O (50 mL) and dried in vacuo yielding 5 as white crystal solid (6.92 g, 30

Benzyl 4-(bromomethyl)benzoate (9) [34]
To a solution of 4-(bromomethyl)benzoic acid (2 g, 9.3 mmol) in dry toluene (23 mL) were added p-TsOH monohydrate (177 mg, 0.93 mmol) and benzyl alcohol (1.5 mL, 14 mmol) and the resulting mixture was stirred at 120 C for 22.5 h using Dean-Stark apparatus for collection of formed H 2 O. All volatiles were removed under reduced pressure and the resulting residue was diluted with EtOAc (100 mL) and sat. NaHCO 3 (45 mL). The layers were mixed and then separated. The combined organic layers were washed with brine, dried over Na 2 SO 4 and filtered. The crude was concentrated in vacuo and purified via flash column chromatography (SiO 2 , n-hexane/EtOAc 100:0 to 95:5, v/v) yielding 9 as light-brown solid (1.

N-(3,4-Dibromophenyl)acetamide (12) [35]
To a solution of 11 (3.7 g, 14.6 mmol) in dry pyridine (28 mL) was added dropwise acetyl chloride (2.1 mL, 29.2 mmol) at 10-15 C using ice bath and the resulting mixture was stirred at rt for 2 h. All volatiles were removed under reduced pressure and the resulting residue was diluted with DCM (100 mL) and H 2 O (50 mL). The layers were mixed and then separated. The combined organic layers were washed with brine, dried over Na 2 SO 4 and filtered. The crude was concentrated in vacuo and purified via flash column chromatography (SiO 2 , DCM) yielding 12 as light-yellow solid (

Вenzyl 4-(5-amino-2-bromo-4-nitrophenoxy)benzoate (15)
To a suspension of 23 (231 mg, 1.012 mmol) and K 2 CO 3 (420 mg, 3.038 mmol) in dry DMSO (5.8 mL) was added dropwise solution of 14 (250 mg, 0.845 mmol) in dry DMSO (5.8 mL) and the resulting mixture was stirred at 100 C for 2 h. The reaction mixture was poured into a mixture of ice and water. The mixture was basified to a pH of 13 by addition of 2 M NaOH aq. sol. and the resulting precipitate was collected by filtration. The crude was purified via flash column chromatography (SiO 2 , n-hexane/

Benzyl 4-hydroxybenzoate (23) [36]
To a solution of 4-hydroxybenzoic acid (7 g, 50.7 mmol) in MeOH (85 mL) and H 2 O (9.5 mL) was added Cs 2 CO 3 (8.25 g, 25.3 mmol) and the resulting mixture was stirred at rt for 2 h. All volatiles were removed under reduced pressure. The obtained cesium salt of 4-hydroxybenzoic acid was suspended in dry DMF (46.5 mL), benzyl bromide (6 mL, 50.7 mmol) was added at 10-15 C using ice bath and the resulting mixture was stirred at rt for 16 h. All volatiles were removed under reduced pressure and the resulting residue was diluted with EtOAc (150 mL) and H 2 O (75 mL). The layers were mixed and then separated. The combined organic layers were washed with brine, dried over Na 2 SO 4 and filtered. To a solution of the appropriate amide (2, 5 or 18) (1.0 equiv) in dry DMF was added sodium hydride (80% w/w in paraffin oil) (1.7 equiv) and the resulting mixture was stirred at rt for 1 h. Then solution of 9 (1.1 equiv) in dry DMF was added and the reaction mixture was stirred at rt for 15 h. All volatiles were removed under reduced pressure and the resulting residue was diluted with EtOAc (70 mL) and saturated Na 2 CO 3 aq. sol. (35 mL). The layers were mixed and then separated. The combined organic layers were washed with brine, dried over Na 2 SO 4 and filtered. The crude was concentrated in vacuo and purified via flash column chromatography (SiO 2 , n-hexane/EtOAc 100:0 to 90:10, v/v). To a suspension of the appropriate imine (3, 7, 19 or 21) (1.0 equiv) in MeOH and acetic acid (90:10, v/v) was added NaCNBH 3 (5.0 equiv) at 10-15 C using ice bath and the resulting mixture was stirred at rt for 1 h. All volatiles were removed under reduced pressure and the resulting residue was neutralized (pH ¼ 7) using saturated NaHCO 3 aq. sol. The resulting aqueous phase was extracted with EtOAc (3 Â 50 mL) and the combined organic layers were washed with brine, dried over Na 2 SO 4 and filtered. The crude was concentrated in vacuo to furnish the corresponding amines without any further purification.  13 (8). Compound 8 (246 mg, 0.55 mmol, white solid) was prepared in 98% yield from 7 (250 mg, 0.56 mmol) following the general procedure except that the reaction mixture was poured into a mixture of ice and water and, after neutralization, the resulting precipitate was obtained through filtration in vacuo furnishing product 8; m.p.: 119-120  13 (20). Compound 20 (22 mg, 0.03 mmol, white solid) was prepared in 95% yield from 19 (23 mg, 0.03 mmol) following the general procedure except that 7 equivalents of NaCNBH 3 (15 mg, 0.24 mmol) were added and the resulting mixture was stirred for 4 h. Then it was poured into a mixture of ice and water and, after neutralization, the resulting precipitate was obtained through filtration in vacuo furnishing product 20; m.p.: 66-68 C; TLC (n-hexane/  (22). Compound 22 (66 mg, 0.1 mmol, white solid) was prepared in 94% yield from 21 (73 mg, 0.11 mmol) following the general procedure except that 10.5 equivalents of NaCNBH 3 (72 mg, 1.15 mmol) were added and the resulting mixture was stirred for 8.5 h. Then it was poured into a mixture of ice and water and, after neutralization, the resulting precipitate was obtained through filtration in vacuo furnishing product 22; m.p.: 73-75 C; TLC (n-hexane/   13   was diluted with saturated Na 2 CO 3 aq. sol. (25 mL To a solution of appropriate mono-or di-benzyl ester (27a-e or 29a-e) in dry THF and EtOH abs. (50:50, v/v) were added Pd, 10% on activated carbon (0.5 equiv) and HCOONH 4 (6 equiv) and the resulting mixture was stirred at rt for 4 h. The reaction mixture was filtered in vacuo and the obtained filtrate was concentrated under reduced pressure. The resulting residue was acidified to a pH of 1 by addition of conc. HCl and the aqueous phase was extracted with EtOAc (3 Â 15 mL). The combined organic layers were washed with brine, dried over Na 2 SO 4 and filtered. The crude was concentrated in vacuo and purified via recrystallization with EtOAc and MeOH. )-2-oxo-7-phenyl-3,4-dihydroquinoxalin-1(2H)-yl]methyl}benzoic acid (28e). Compound 28e (6.6 mg, 0.015 mmol, white solid) was prepared in 79% yield from 27e (10 mg, 0.018 mmol) following the general procedure except that the reaction mixture was stirred at rt for 2 h and the crude was purified via flash column chromatography (SiO 2 , n-hexane/EtOAc 50:50 to 0:100, v/v); m.p.: The extracts were collected by 0.1 M HCl and analyzed by a commercial ELISA kit (Cayman Chemicals) for total cell cGMP content, following the manufacturer's instructions. cGMP levels were normalized for total protein in the respective well, determined by a Micro BCA Protein assay kit (Thermo Scientific).

Cloning of constructs and mutagenesis.
For infection of Sf-9 cells human α 1 and β 1 were cloned into the pFASTBAC vector as it was described by Koglin und Behrends [49,50]. For the purification of sGC a C-terminally strep tagged or N-terminally twin-strep-TEV tagged α 1 subunit was used. For the C-terminally strep tagged α 1 subunit synthetic DNA encoding the C-terminal region of α 1 and a strep tag II (PHGE-PIKMRIGLHSGSVFAGVVGVKMPRYCLFGNNVTLANKFESCSVPRKINVSP TTYRLLKDCPGFVFTPRSREELPPNFPSEIPGICHFLDAYQQGTNSKPCFQ KKDVEDGNANFLGKASGIDWSHPQFEK) was purchased from GeneArt ® (Invitrogen ™ , Thermo Fisher Scientific; Waltham, USA) and cloned Ncol/Kpnl into α 1 human in pFASTBAC [50]. To purify the wildtype α 1 containing constructs by Nterminal fusion of a TEV protease-cleavable twin strep tag II an artificial gene encoding a twin strep tag II, a TEV protease-cleavage side and the N-terminus of the α 1 subunit (LGSATM-SAWSHPQFEKGGGSGGGSGGSAWSHPQFEKSGGGGGENLYFQGMFCTK LKDLKITGECPFSLLAPGG) was cloned BamHI/SexAI into the α 1 construct in pFASTBAC. This artificial gene was obtained from GeneArt ® (Invitrogen ™ , Thermo Fisher Scientific; Waltham, USA). The mutation of histidine at position 105 to alanine in the β 1 subunit was introduced into the human β 1 in pFASTBAC.

Protein purification.
For peptide purifications Sf-9 cells, cultivated at 27 C and 140rpm, were infected with recombinant baculoviruses encoding for a twin strep TEV tagged α 1 and β 1 wildtype or an Cterminally strep-tagged α 1 subunit and β 1 H105A. 300-400ml of cell suspension with a concentration of 3 Â 10 6 cells/ml were coinfected with virus. After 72h cells were harvested at 3020xg for 4min and resuspended in 9-12ml lysis buffer (50 Mm triethanolamine/HCl, 10 mM DTT, 1 mM ethylenediaminetetraacetic acid, pH 7.4) containing complete ™ protease inhibitor cocktail (Roche, Mannheim, Germany). The resuspended cells were sonified and centrifuged at 15,000Âg and 4 C for 90 min. The supernatant was clarified by filtration through 0.45 μM and 0.2 μM filters before purification via affinity chromatography.
The samples were treated 3 min with a temperature of 99 C before storage at À20 C. After resolution on a 10% SDS polyacrylamide electrophoresis gel the samples of the purified protein were stained with Coomassie Brilliant Blue as described in Kang et al [51]. 4.3.2.6. Densitometric analysis and determination of purified protein concentration. The protein concentration was determined by the Warburg-Christian method with a NanoPhotometer ™ (Implen, München, Germany) [52]. As there were clear differences in protein concentrations detected by Coomassie blue staining between the different α 1 S/β 1 H105A Proteins, we adjusted the concentration by densitometric analysis. The densitometric volume was quantified with ImageJ (National Institutes of Health, Bethesda, MD, USA). For determination of purified protein concentration from densitometric volume after Coomassie Brilliant Blue staining, a standard curve was generated in triplicate. BSA was loaded on a 10% SDS polyacrylamide electrophoresis gel in amounts ranging from 0.125 mg to 4 mg. A standard curve was created by linear regression with Prism5 (GraphPad, La Jolla, USA), relating the loaded protein amount to the densitometric volume after staining with Coomassie Brilliant Blue. To determine purified sGC concentration, the sum of the densitometric volume of both subunits was determined.
4.3.2.7. sGC activity assay. sGC activity assays were performed as described previously in Kraehling et al. [53]. In short, cGMP forming activity of purified protein was determined by incubation for 10 min at 37 C in the presence of 50 mM triethanolamine/HCl buffer pH 7.4, containing 3 mM DTT, 1 mM 3-isobutyl-1-methylxanthine, 1 mM cGMP, 5 mM creatine phosphate, 10units/tube creatine phosphokinase, 0.5 mM [α-32 P]GTP and 3 mM magnesium chloride. The reaction was stopped by adding 500 μL of 125 mM zinc acetate and 500 μL of 120 mM sodium carbonate which results in a co-precipitation of zinc carbonate and 5 0 -nucleotides. The mixture was centrifuged at 18.000g for 9 min. The supernatant was purified on short alumina columns before eluting the enzyme formed cGMP with natrium acetate (pH 6.5; 250 mM). CGMP was quantified by liquid scintillation counting in a Beckman Coulter Counter LS6500 (Beckmann GmbH, Krefeld, Germany) or a Hidex 600 SL Automatic TDCR Liquid Scintillation Counter (Hidex Oy, Turku, Finland) with a counting time of 2 min [54]. 4.3.2.8. Statistical analysis. All results are expressed as means AE standard deviation (SD) of three independent experiments. In sGC activity assay samples were analysed as duplicates and averaged for each assay. Analyses and curve fitting were performed with Prism 5 (GraphPad, La Jolla, USA). For comparison of sGC activity one-way ANOVA was performed followed by Tukey's multi comparison test with Prism 5. A value of P < 0.05 was considered statistically significant.

Author contribution statement
Dionysios-Panagiotis Kintos: Conceived and designed the experiments; Performed the experiments; Analyzed and interpreted the data; Wrote the paper.
Konstantinos Salagiannis, Theresa Wittrien: Performed the experiments; Analyzed and interpreted the data; Wrote the paper.
Soenke Behrends, Stavros Topouzis, Manolis A. Fousteris: Conceived and designed the experiments; Analyzed and interpreted the data; Contributed reagents, materials, analysis tools or data; Wrote the paper.

Funding statement
This work was supported by Operational Programme "Competitiveness, Entrepreneurship and Innovation" (NSRF, 2014-2020), co-financed by Greece and the European Union (European Regional Development Fund) ("INSPIRED-The National Research Infrastructures on Integrated Structural Biology, Drug Screening Efforts and Drug target functional characterization" (MIS 5002550), implemented under the Action "Reinforcement of the Research and Innovation Infrastructure"). The publication of this article has been financed by the Research Committee of the University of Patras.

Data availability statement
No data was used for the research described in the article.

Declaration of interest's statement
The authors declare no conflict of interest.

Additional information
Supplementary content related to this article has been published online at https://doi.org/10.1016/j.heliyon.2022.e11438.