Ca²⁺ is a key signal that regulates multiple biological phenomena ranging from neurotransmission to gene expression. Changes in the concentration of intracellular free Ca²⁺, its locus of action, and the amplitude and duration of Ca²⁺ influx are essential to transmit information through the nervous system. The mechanisms by which these changes can bring about such diverse neural responses rely on the ability of Ca²⁺ sensors to decode Ca²⁺ signals (McCue et al., 2010; Burgoyne et al., 2019). The neuronal calcium sensor (NCS) family of proteins has evolved to participate in specialized neuronal functions separate from calmodulin, due to their 10-fold higher affinity for Ca²⁺. The most abundant protein of the NCS family is the neuronal calcium sensor 1 (NCS-1), which was first discovered in Drosophila and named frequenin (Frq), is N-terminally myristoylated (Myr) and highly conserved from yeast to humans (Burgoyne et al., 2019; Pongs et al., 1993; Ames and Lim, 2012; Burgoyne and Haynes, 2012).

Unlike other NCSs, NCS-1 is found outside the nervous system. It does not contain a Ca²⁺/Myr switch, thus being constantly bound to the membrane, and has multiple binding partners (Burgoyne et al., 2019; Ames and Lim, 2012; Burgoyne and Haynes, 2012; Mansilla et al., 2017). NCS-1 participates in a wide range of important neuronal functions: it is a regulator of Ca²⁺ channels, exocytosis, synaptogenesis, and axonal growth, affecting higher functions such as learning and memory, neuroprotection, and axonal regeneration (Burgoyne et al., 2019; Burgoyne and Haynes, 2012; McFerran et al., 1999; Hui and Feng, 2008; Weiss et al., 2010; Dason et al., 2012). Furthermore, NCS-1 has been implicated in several pathological processes such as X-linked mental retardation and autism, schizophrenia, and bipolar disorder (Piton et al., 2008; Koh et al., 2003; Torres et al., 2009; Bahi et al., 2003). The multifunctionality of NCS-1 relies on its ability to recognize and regulate different and unrelated target proteins: G-protein-coupled receptors (GPCRs) and some of their regulators, Ca²⁺ channels, guanine nucleotide exchange factors (GEF), and kinases, both in a Ca²⁺-dependent or -independent manner (Burgoyne et al., 2019).

The structure of NCS-1 consists of two pairs of EF-hand motifs, of which only three are functional: EF-2, EF-3, and EF-4 (Bourne et al., 2001). EF-2 and EF-3 can recognize Mg²⁺ as well. It is known that the two Ca²⁺/Mg²⁺ binding sites are structural sites that allow the protein to adopt its tertiary fold. EF-4 has been suggested to be a regulatory Ca²⁺ binding site, able to sense changes in cytosolic calcium levels in neurons (Burgoyne et al., 2019; Aravind et al., 2008; Mikhaylova et al., 2009; Chandra et al., 2011; Tsvetkov et al., 2018; da Silva et al., 1995; Gifford et al., 2007).

The structures of several NCS proteins bound to their corresponding targets have shown that these Ca²⁺ sensors use a surface-exposed hydrophobic crevice to recognize their targets, which generally present short helical motifs that bind to the N- or C-terminal part of this large cavity (Figure 1A). It has been proposed that the structural determinants of target specificity are based on the shape and size of the hydrophobic crevice. NCS proteins contain a dynamic C-terminal helix (the so-called helix H10) that can insert into the crevice, thus contributing to its shape (Figure 1A). Since Ca²⁺ binding promotes structural rearrangements (Figure 1A), the occupancy of the three Ca²⁺ binding sites also determines affinity for protein partners. Also, the presence of hydrophilic residues at the border of the crevice contributes to target specificity and they constitute hot spots for interactions with the different targets (Burgoyne et al., 2019; Mansilla et al., 2017).

Figure 1 with 1 supplement see all

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Interestingly, NCS-1 is an important regulator of G-protein signaling since it binds to proteins such as GPCRs including the dopamine D2, adenosine 2A, and cannabinoid CB1 receptors (Kabbani et al., 2002; Navarro et al., 2012; Pandalaneni et al., 2015; Angelats et al., 2018); and the molecular chaperone and GEF Ric-8A (Romero-Pozuelo et al., 2014). Although little is known of the regulatory activity of NCS-1 on GPCR function and the cellular and physiological consequences, Romero-Pozuelo et al., 2014, showed that the NCS-1/Ric-8A complex is implicated in the regulation of synapse number and probability of neurotransmitter release. In fact, the regulation of this protein-protein interaction (PPI) using small-molecule modulators allows synapse function control under pathological conditions. In neurodevelopmental disorders, where synapse number is abnormally high, the inhibition of the NCS-1/Ric-8A complex formation reduces synapse number and improves learning in Fragile X syndrome animal models (Cogram et al., 2022; Mansilla et al., 2017). In contrast, the stabilization of the PPI prevents synapse loss and the consequent impairment in locomotion in a Drosophila model of Alzheimer’s disease neurodegeneration at the motor neurons (Canal-Martín et al., 2019).

Ric-8A is an ubiquitously expressed cytosolic protein with two main functions: it constitutes a molecular chaperone that allows heterotrimeric Gα subunit biogenesis (Gabay et al., 2011) and additionally, works as a guanine exchange factor for G_i, G_q, and G_12/13 families (Tall et al., 2003; Chan et al., 2011; Van Eps et al., 2015). Both activities are stimulated by casein kinase II (CK2) phosphorylation (Papasergi-Scott et al., 2018). Ric-8A has been shown to regulate asymmetric cell division and is essential for embryonic development (Miller and Rand, 2000; Afshar et al., 2004; Couwenbergs et al., 2004; Tõnissoo et al., 2010; Woodard et al., 2010). Work in Drosophila has shown the relevance of Ric-8A in activating G_s for in vivo synaptogenesis and that this activity is downregulated by NCS-1 (Romero-Pozuelo et al., 2014). Recently, the structure of several Ric-8A/Gα or Gα fragment complexes have been solved at atomic level (Srivastava and Artemyev, 2019; McClelland et al., 2020; Seven et al., 2020). These works revealed the structural basis of Ric-8A as a Gα chaperone and GEF and showed how phosphorylation of Ric-8A residues S435 and T440 stabilizes a conformation that is competent for Gα recognition. However, there is scarce information on the molecular function of NCS-1 on Ric-8A activity. Based on genetic studies, it has been proposed that NCS-1 interacts with Ric-8A and prevents the Ric-8A/Gα interaction (Romero-Pozuelo et al., 2014). Here, we have combined biochemical, biophysical, and crystallographic studies to reveal the structural determinants of NCS-1/Ric-8A recognition and the mechanism of NCS-1-mediated downregulation of Ric-8A activity. This work shows how NCS-1 and Ric-8A constitute a hub that integrates Ca²⁺, phosphorylation, and G-protein signaling. The emergent picture indicates that Ric-8A activity is under NCS-1 control and that a Ca²⁺ signal triggers the disassembly of the NCS-1/Ric-8A complex, which in turn allows phosphorylation of Ric-8A, which stabilizes the Ric-8A/Gα complex.

Several studies have shown the structural determinants of Ric-8A chaperone and GEF activity for Gα protein subunits (Srivastava and Artemyev, 2019; McClelland et al., 2020; Seven et al., 2020; Zeng et al., 2019; Papasergi-Scott et al., 2023). Ric-8A phosphorylation promotes Gα recognition since it stabilizes a conformation that allows Ric-8A to trap the Ras-like domain of Gα (Figure 1B). This facilitates the folding of the Gα subunit and stabilizes a nucleotide-free state, which in turn would prepare the protein for GTP loading (Srivastava and Artemyev, 2019; McClelland et al., 2020; Seven et al., 2020). However, the mechanism by which the Gα chaperone and GEF activity of Ric-8A could be downregulated by NCS-1 remained unclear. Here, we have combined biochemical, biophysical, and PPI assays with X-ray crystallography to shed light on the mechanism that keeps Ric-8A inactive.

The combination of the crystallographic work using Ric-8A peptides of different lengths, together with cell-based PPI assays with different Ric-8A and NCS-1 mutants, has enabled the discovery of the necessary and sufficient region of Ric-8A that is required for NCS-1 recognition, a region that is conserved between human and rat (Figures 2 and 3). This region corresponds to the two-helix bundle that constitutes the HEAT repeat 9 of the ARM-HEAT repeat domain, and a disordered region that, in the presence of Gα, forms an extended coil to permit phosphorylated S435 and T440 (in human S436 and T441) to attach to positively charged patches of Ric-8A (Figures 1B and 7; Srivastava and Artemyev, 2019; McClelland et al., 2020; Seven et al., 2020; Zeng et al., 2019). Binding of phosphorylated rRic-8A (pRic-8A) to Gα does not substantially alter the conformation of the ARM-HEAT repeat domain, since the overall structure and orientation of repeats of uncomplexed pRic-8A and pRic-8A bound to Gα are very similar, such that any major differences are confined to the polypeptide loop regions. As shown by the crystal structures presented here, the HEAT repeat 9 undergoes a substantial conformational change: helix a9 unfolds while helix b9 refolds, resulting in the named regions R1 and R2 (Figure 3B and Figure 7B). The relative orientation of R1 and R2 change with respect to a9 and b9. If helices b9 and R2 are superposed, helix a9 is rotated ~180 degrees with respect to R1 (Figure 7B). To illustrate the structural rearrangement, a morph movie has been produced starting at the Gα-bound and ending at the NCS-1-bound conformations (Video 1). The structural reorganization affects the distribution of hydrophobic and positively charged amino acids. In the Ric-8A/Gα complex, hydrophobic residues in the HEAT repeat 9 are facing the ARM repeat 8 to create contacts between the two repeats (Figure 7A and C and Video 1). In the NCS-1ΔH10/Ric-8A-P complex, the same hydrophobic residues are exposed to the solvent, affording their interaction with the hydrophobic crevice of NCS-1, while positively charged residues are exposed on the opposite face, generating an amphipathic structure (Figure 3, Figure 3—figure supplement 2, Figure 7C and Video 1). This would create electrostatic repulsions with repeat 8 and would force the detachment of repeat 9 from repeat 8, so that NCS-1 properly recognizes Ric-8A. The unfolding of helix a9 could act as a hinge to assist in the rearrangement that occurs when uRic-8A binds NCS-1.

Figure 7

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Video 1

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The dramatic increase in thermal stability experienced by unphosphorylated Ric-8A upon binding to NCS-1 (>20°C in Ti; Figure 2B) is indicative of the strong interactions established in the complex between the two proteins. It also means that the free energy of binding would be high enough as to pay the energy penalty derived from the structural changes experienced by Ric-8A to give rise to the complex with NCS-1. Currently, it is unknown if the impact of NCS-1 binding to Ric-8A extends beyond the structural rearrangement that takes place at repeat 9 and its detachment from repeat 8. Nonetheless, it is worth noting that global changes within Ric-8A ARM-HEAT repeat domain have been observed when comparing the phosphorylated and unphosphorylated forms of Ric-8A-452 (Zeng et al., 2019). The combination of the crystallographic data with low-resolution SAXS data suggested that there were quasi-rigid body angular displacements of three subdomains (constituted by repeats 1–4, 5–6, and 7–9) of the ARM-HEAT repeat domain (Figure 7C). Therefore, interactions between the phosphorylated residues and the C-terminal repeat 9 (Figure 7A) are translated into different contacts between the subdomains, along with a different global shape and curvature of the ARM-HEAT repeat domain (Zeng et al., 2019). The structure solution of the complete Ric-8A ARM-HEAT repeat domain bound to NCS-1 will allow to understand the extent of the rearrangement that Ric-8A suffers upon NCS-1 binding.

It is worth noting that most of the residues of Ric-8A implicated in the NCS-1/Ric-8A PPI are occluded in the absence of NCS-1 (Figures 3 and 7). The analysis of co-immunoprecipitation assays conducted with different NCS-1 and Ric-8A mutants point to the relevance of van der Waals interactions between the upper part of the NCS-1 crevice and the C-terminal half of Ric-8A helix R2 (Figure 3D and Table 2; see binding loss in NCS-1 W30A and Ric-8A L424A, M425A). In the complex with Gα, this region adopts an extended coiled structure and is attached to the ARM-HEAT repeat domain, due to the interactions with the phosphorylated Ric-8A residues (Figure 7A). In the absence of phosphorylation, this region would be exposed to the solvent and could acquire the helical structure needed to initiate the recognition of NCS-1. Initial contacts of the C-terminal part of helix R2 with NCS-1 could constitute the first steps in the recognition process and trigger the subsequent reorganization, which would imply the detachment of repeat 9 and complete the formation of helix R2 and, finally, the formation of helix R1 and the change in the relative orientation of R1 and R2 with respect to a9 and b9.

The dynamic C-terminal helix H10 of NCS-1 regulates the binding of Ric-8A by inserting into the hydrophobic crevice (Figure 1A; Romero-Pozuelo et al., 2014). Comparison of the structure of Ric-8A-P in complex with NCS-1ΔH10 with that of NCS-1 in which its helix H10 is inserted in the hydrophobic crevice explains why this helix constitutes a built-in competitive inhibitor of the NCS-1/Ric-8A interaction (Figure 1—figure supplement 1). The position of helix H10 completely overlaps with the Ric-8A region that is recognized by NCS-1 and displaces most of the H-bonds and contacts reported for the complex, including the C-terminus of helix R1, the N-terminus of helix R2, and the loop connecting them (Figure 3—figure supplement 2B, C and Figure 1—figure supplement 1A). Interestingly, the binding of Ric-8A does not cause a significant structural reorganization of NCS-1. Superimposition of the hNCS-1ΔH10/Ric-8A-P structure with that of free human NCS-1 (Canal-Martín et al., 2019) shows subtle changes: the NCS-1 helix H3 and the loop connecting the helices H3 and H4 rearrange to open up the cavity to accommodate helix R2 (Figure 1—figure supplement 1C). In addition, the loop connecting helices H7 and H8 helices, which interacts with the Ric-8A R1 region, undergoes a conformational change to permit the contacts between NCS-1 T135 and Ric-8A residues I407 and, to a lesser extent, V403 (Figure 3—figure supplement 2A, D and Figure 1—figure supplement 1C).

The data presented here show how Ca²⁺ recognition and Ric-8A phosphorylation serve as determinants of NCS-1/Ric-8A recognition. Binding of Ca²⁺ to the structural EF-hands EF-2 and EF-3 is necessary for protein recognition, while EF-4, the regulatory Ca²⁺ binding site (Aravind et al., 2008), is free of Ca²⁺, as shown in the crystal structure of the NCS-1ΔH10/Ric-8A-P complex (Figure 4A) and the in vitro reconstruction of the complex (Figure 2A). We show that a fully Ca²⁺-loaded protein does not properly recognize Ric-8A (assembly assays, Figure 2A, and BLI, Figure 4E and Table 3), suggesting that in the cellular context, an increase in Ca²⁺ concentration and the subsequent loading of EF-4 may trigger the disruption of the NCS-1/Ric-8A complex. This agrees with the previous cell-based PPI assays that showed a weakened affinity of NCS-1 for Ric-8A in Ca²⁺ saturating conditions (Romero-Pozuelo et al., 2014). Moreover, our in vitro nucleotide exchange assays demonstrate a loss of Ric-8A GEF function when in complex with NCS-1, while nucleotide exchange rates are partially rescued following addition of Ca²⁺ (Figure 6). With respect to the phosphorylation state of Ric-8A, NCS-1 binds to unphosphorylated Ric-8A protein (Figures 2A and 5A) and more importantly, in the context of the protein complex, NCS-1 protects Ric-8A from CK2-mediated phosphorylation (Figure 5B). In this situation, an increase of Ca²⁺ levels would permit the disassembly of the NCS-1/Ric-8A complex, allowing release of Ric-8A from the membrane, and subsequent phosphorylation (Figure 8). This would promote Ric-8A activation for Gα subunit binding and GEF activity. When inactivation of Ric-8A is required in the cellular context, Ric-8A dephosphorylation must precede NCS-1 binding, since the phosphorylated protein does not recognize NCS-1 properly (Figures 5C and 8). This can be explained as well in the context of the structure of phosphorylated Ric-8A bound to Gα, since the strong ionic interactions of phosphorylated S335 and T440 with the ARM-HEAT repeat domain would preclude the structural rearrangement of repeat 9 (Figure 7A and C). Cellular mechanisms of Ric-8A dephosphorylation are not understood. It has been proposed that in the cell, Ric-8A is constitutively phosphorylated (Papasergi-Scott et al., 2018). However, and in the light of the results presented here, in neurons and other tissues where NCS-1 is expressed (Burgoyne et al., 2019), Ric-8A phosphorylation may be under NCS-1 control and regulated by Ca²⁺ signals. Since NCS-1 is a Ca²⁺ sensor that is constantly bound to the membrane and close to Ca²⁺ channels, changes of cytosolic Ca²⁺ would promote the disassembly of the protein complex and the subsequent phosphorylation of Ric-8A by CK2 (Figure 8). Given that Ric-8A is a ubiquitously expressed protein, it is unknown if, in tissues where NCS-1 is not expressed, there are other Ca²⁺ sensors in charge of the regulation of its activity. Furthermore, it is likely that NCS-1 interacts with Ric-8B, a chaperone for other Gα (Tall et al., 2003; Chan et al., 2011), since the sequence and structure of the NCS-1 interacting region is conserved (Papasergi-Scott et al., 2023).

Figure 8

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The transmission of the nerve impulse between neurons begins with the generation of an action potential in the axon initial segment. An action potential is a depolarization of the plasma membrane that depends on Na⁺ entry through specific voltage-gated channels and that propagates along the axon to the terminal. When the depolarization Na⁺ wave reaches the axon terminal, voltage-dependent Ca²⁺ channels open allowing the massive entrance of Ca²⁺ which in turn triggers neurotransmitter release (Llinás et al., 1981; Oheim et al., 2006). Therefore, Na⁺ precedes Ca²⁺ waves. Here, we demonstrate that Na⁺ decreases the affinity of NCS-1 for Ca²⁺ (Figure 4D and Table 4). The crystal structure of the hNCS-1ΔH10/Ric-8A-P complex shows that NCS-1 binds Na⁺ at EF-4, the regulatory Ca²⁺ binding site. From a physiological point of view this finding is relevant since NCS-1/Ric-8A disassembly and Ric-8A activation would occur at a Ca²⁺ signal that is higher than initially expected, ensuring the return to the Ric-8A inactive and NCS-1-complexed state when Ca²⁺ levels decrease, due to sequestering by several high capacity, low specificity, Ca²⁺ binding proteins.

This work also sheds light into the mechanism of specificity that allows NCS-1 to recognize different targets. NCS-1 interacts with several proteins related to G-protein signaling: Ric-8A and GPCRs such as the adenosine A2A, cannabinoid CB1, or the dopamine D2 receptors (Kabbani et al., 2002; Navarro et al., 2012; Angelats et al., 2018). The structure and Ca²⁺ determinants of dopamine D2 receptor recognition by NCS-1 are known (Figure 1; Pandalaneni et al., 2015). It is relevant that the Ca²⁺ signals that trigger protein-protein recognition are opposite: while the binding of NCS-1 to Ric-8A takes place at low Ca²⁺ concentrations at which the functional site EF-4 is free of Ca²⁺ (Figures 3A and 4A), binding to dopamine D2 receptor takes place at high Ca²⁺ levels (Figure 1A). Therefore, while Ric-8A is being blocked by NCS-1, dopamine D2 receptor is free of NCS-1 and vice versa. Also, the role of the NCS-1 dynamic C-terminal helix H10 is different in the two molecular recognition processes, while NCS-1 uses this helical element to properly recognize dopamine D2 receptor (Pandalaneni et al., 2015), the helix H10 negatively regulates the binding to Ric-8A. Finally, if the crystal structure of the NCS-1ΔH10/Ric-8A-P complex is compared with other NCS proteins bound to their corresponding targets, the complex that is structurally most similar is the KChIP1/Kv4.3 pair, with an elongated helix resembling the R2 bound to Ric-8A (Figure 1—figure supplement 1D). Interestingly, the N- to C-orientation of the helix is opposite, and no extra helical segment (R1) is recognized. In addition, the Ca ²⁺ content is different (Figure 1A; Pioletti et al., 2006). The number of structures that have been determined of NCS proteins bound to different targets continues to grow. Comparison of these structures show that, while all of the ligands use the same NCS hydrophobic crevice, differences in Ca²⁺ site ligation, role of the helix H10, and the loop that connects EF-3 and EF-4 in NCS-1 shape the crevice to recognize, very specifically, different target protein motifs. In this sense polar interactions between NCS-1 and the target protein play also an important role to ensure specificity and affinity (Figure 1A and Figure 1—figure supplement 1).

Small-molecule PPI modulators with therapeutic potential have been discovered in the past few years (Mansilla et al., 2017; Canal-Martín et al., 2019; Roca et al., 2018). They are able to inhibit (e.g. the phenothiazine FD-44) or stabilize (e.g. the acylhydrazone 3b) the formation of the NCS-1/Ric-8A complex and in doing so, they regulate synapse number and function and show a promising prospect in the treatment of neurodevelopmental disorders (Mansilla et al., 2017; Cogram et al., 2022) and neurodegenerative diseases (Canal-Martín et al., 2019). It has been suggested that these small molecules, which bind to the same NCS-1 site despite their opposite activity, are not Ric-8A competitors. Instead, they have been suggested to be allosteric modulators that contribute to the stabilization of the dynamic helix H10 inside (inhibitors) or outside (stabilizers) the crevice, favoring NCS-1 conformations that hinder or allow the entrance of Ric-8A and the subsequent formation of the protein complex. Supporting this, the modulator FD-44 does not inhibit the formation of the NCS-1/Ric-8A complex when the dynamic helix H10 is deleted. The structure of the hNCS-1ΔH10/Ric-8A-P complex explains the molecular mechanism of action of the regulatory compounds. Comparison of the different complexes shows that the binding sites for these molecules overlap with the C-terminal half of helix R2 (Figure 1—figure supplement 1A, B). With respect to the inhibitors, hydrophobic interactions of FD-44 with the helix H10 stabilize the crevice-inserted helix H10 conformation of NCS-1. The combination of the compound plus helix H10 makes the Ric-8A interacting region of NCS-1 completely unavailable (Figure 1—figure supplement 1A). On the contrary, for the PPI stabilizer, the polar characteristics of the molecule hinders the approach of the helix H10 to the crevice, making the C-terminal part of the cavity, which is more relevant in terms of the PPI, available for Ric-8A recognition (Figure 1—figure supplement 1B). The structure of hNCS-1ΔH10/Ric-8A-P does not support the existence of a ternary NCS-1/Ric-8A/stabilizer complex, since there would be no room for the repositioning of the compound. Therefore, the binding of Ric-8A to NCS-1 would displace modulator 3b from the crevice, which would be likely given the moderate affinity of the compound (Canal-Martín et al., 2019). Finally, we believe that the high-resolution data on the NCS-1/Ric-8A PPI interface presented here will be essential to rationally develop improved compounds with better PPI regulatory properties and selectivity, since the relevant interactions between NCS-1 and Ric-8A have been finally revealed. The fact that the NCS-1 Ca²⁺ and structural determinants are different in recognizing Ric-8A and the dopamine D2 receptor is conceptually relevant, since opens the path to the design of selective drugs that target specifically these neuronal pathways.

The atomic coordinates and structure factors have been deposited in the Protein Data Bank (https://www.pdb.org/) with codes: Structure 1 (8ALH), Structure 2 (8AHY), Structure 3 (8ALM). All data generated or analysed during this study are included in the manuscript and supporting files. Source data files have been provided for Figures 2–5.

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Author details

1. Daniel Muñoz-Reyes

   Department of Crystallography and Structural Biology, Institute of Physical-Chemistry 'Blas Cabrera', CSIC, Madrid, Spain

   Contribution

   Investigation, Writing – review and editing, D M-R performed protein purification, protein complex assemblies, biochemical and biophysical experiments, protein crystallization, X-ray diffraction experiments and structure resolution

   Competing interests

   No competing interests declared

2. Levi J McClelland

   Center for Biomolecular Structure and Dynamics, and Division of Biological Sciences, University of Montana, Missoula, United States

   Contribution

   Funding acquisition, Investigation, Writing – review and editing, L.J.M. purified the proteins needed for functional assays and performed the experiments. Also designed and analyzed functional assays

   Competing interests

   No competing interests declared

3. Sandra Arroyo-Urea

   Institute for Biocomputation and Physics of Complex Systems (BIFI) and Laboratorio de Microscopías Avanzadas (LMA), University of Zaragoza, Zaragoza, Spain

   Contribution

   Investigation, Writing – review and editing

   Competing interests

   No competing interests declared

4. Sonia Sánchez-Yepes

   Department of Neurobiology, Instituto Ramón y Cajal de Investigación Sanitaria, Hospital Universitario Ramón y Cajal, Madrid, Spain

   Contribution

   Investigation, S S-Y performed co-IPs

   Competing interests

   No competing interests declared

5. Juan Sabín

   1. AFFINImeter Scientific & Development team, Software 4 Science Developments, Santiago de Compostela, Spain
   2. Departamento de Física Aplicada, Universidad de Santiago de Compostela, Santiago de Compostela, Spain

   Contribution

   Formal analysis, J S analyzed and interpreted ITC data

   Competing interests

   No competing interests declared

6. Sara Pérez-Suárez

   Department of Crystallography and Structural Biology, Institute of Physical-Chemistry 'Blas Cabrera', CSIC, Madrid, Spain

   Contribution

   Investigation, S P-S performed cloning, protein expression and purification

   Competing interests

   No competing interests declared

7. Margarita Menendez

   1. Department of Biological Physical-Chemisty, Institute of Physical-Chemistry 'Blas Cabrera', CSIC, Madrid, Spain
   2. Ciber of Respiratory Diseases, ISCIII, Madrid, Spain

   Contribution

   Conceptualization, Formal analysis, Funding acquisition, Investigation, Writing – review and editing, M M performed ITC experiments, analyzed and interpreted ITC data. M M performed the joined analysis of biophysical and functional assays

   Competing interests

   No competing interests declared

8. Alicia Mansilla

   1. Department of Neurobiology, Instituto Ramón y Cajal de Investigación Sanitaria, Hospital Universitario Ramón y Cajal, Madrid, Spain
   2. Department of Systems Biology, Universidad de Alcala, Madrid, Spain

   Contribution

   Formal analysis, Funding acquisition, Investigation, Writing – review and editing, A M designed the co-IPs

   Competing interests

   No competing interests declared

9. Javier García-Nafría

   Institute for Biocomputation and Physics of Complex Systems (BIFI) and Laboratorio de Microscopías Avanzadas (LMA), University of Zaragoza, Zaragoza, Spain

   Contribution

   Formal analysis, Funding acquisition, Investigation, Writing – review and editing

   Competing interests

   No competing interests declared

10. Stephen Sprang

    Center for Biomolecular Structure and Dynamics, and Division of Biological Sciences, University of Montana, Missoula, United States

    Contribution

    Formal analysis, Funding acquisition, Investigation, Writing – review and editing, S R S designed and analyzed functional assays

    Competing interests

    No competing interests declared

11. Maria Jose Sanchez-Barrena

    Department of Crystallography and Structural Biology, Institute of Physical-Chemistry 'Blas Cabrera', CSIC, Madrid, Spain

    Contribution

    Conceptualization, Formal analysis, Supervision, Funding acquisition, Investigation, Writing – original draft, Writing – review and editing, M J S -B contributed with the conception, design of the study, data analysis and interpretation. Performed the joined analysis of biophysical, functional assays and structural data, M J S-B wrote the manuscript with contributions from all authors

    For correspondence

    xmjose@iqf.csic.es

    Competing interests

    No competing interests declared

    "This ORCID iD identifies the author of this article:" 0000-0002-5986-1804

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