Mechanism of beta-arrestin 1 mediated Src activation via Src SH3 domain revealed by cryo-electron microscopy

Domain: chemistry
Published: 2026-02-20
Editorial outcome: accept

Abstract
Beta-arrestins (βarrs) are key regulators and transducers of G-protein coupled receptor signaling; however, little is known of how βarrs communicate with their downstream effectors. Here, we delineate structural mechanisms underlying βarr-mediated signal transduction. Using cryo-electron microscopy, we elucidate how βarr1 recruits and activates the non-receptor tyrosine kinase Src, a well-established signaling partner of βarrs. βarr1 engages Src SH3 through two distinct sites, each employing a different recognition mechanism: a polyproline motif in the N-domain and a non-proline-based interaction in the central crest region. At both sites βarr1 interacts with the aromatic surface of SH3, disrupting the autoinhibited conformation of Src and directly triggering its allosteric activation. This structural evidence establishes βarr1 as an active regulatory protein rather than a passive scaffold and suggests a potentially general mechanism for βarr-mediated signaling across diverse effectors.

Introduction
Beta-arrestins 1 and 2 (βarr1 and βarr2) are ubiquitously expressed scaffold proteins that interact with most, if not all, G-protein coupled receptors (GPCRs) 1. βarrs mediate receptor desensitization and intracellular trafficking, as well as initiate downstream signaling cascades, independently or in concert with G-proteins 2. In their role as signal transducers, βarrs link GPCRs with numerous downstream effectors, including components of several mitogen-activated protein kinase cascades and Src family kinases, amongst many others 2.

Early studies proposed that βarrs function primarily as passive signaling scaffolds; however, more recent work has demonstrated that βarrs can directly and allosterically activate specific effector enzymes, including the non-receptor tyrosine kinase Src 3,4, the mitogen-activated protein kinases c-Raf 5 and extracellular signal-regulated kinase 6. Despite these advances, the molecular mechanisms underlying βarr-mediated signaling remain poorly understood, largely due to the absence of high-resolution structural information for complexes formed between βarrs and their diverse effector enzymes. Structural characterization of these interactions has been limited by their transient nature and mid-micromolar binding affinities, necessitating tailored strategies for complex stabilization.

Src homology 3 (SH3) domains are one of the most ubiquitous binding modules with nearly 300 members found in the human genome 7. SH3-containing proteins are involved in various signaling pathways, such as cell growth and proliferation, endocytosis, and cytoskeleton remodeling 8. Notably, βarrs have been shown to interact with the SH3 domains of several proteins 3, including the proto-oncogene kinase Src 9. Intriguingly, βarrs lack the canonical left-handed type II polyproline motifs required for SH3 binding; therefore, how βarrs recruit SH3 remains unknown.

Here, we report the structural insights into the fundamental mechanisms underlying βarr-mediated signal transduction. Using Src as a prototypical effector of βarr, we elucidate the molecular basis for Src SH3 recruitment and Src allosteric activation by cryo-electron microscopy (cryo-EM).

Results
βarr1 uses two distinct sites to bind SH3
To map the binding interface between the SH3 domain of Src and βarr1, we utilized a disulfide trapping strategy 10. The βarr1 and SH3 Src sequences were derived from rat (Rattus norvegicus) and chicken (Gallus gallus), respectively. Both proteins exhibit high inter-species sequence similarity—93.54% for βarr1 and 93.66% for Src (100% for Src SH3 domain), with the most notable sequence variations present in the unique domain of Src and at the C-terminus of βarr1 (Supplementary Fig. 1). We introduced cysteine substitutions at 18 different positions in βarr1 in proximity to P88-P91 and P120-P124, previously reported to be critical for βarr1–Src interaction (Fig. 1a) 3,9. Guided by the available structures of SH3 domains in complex with polyproline-rich peptides, we designed 13 cysteine mutants of SH3 (Fig. 1b). We comprehensively tested 234 combinations of purified βarr1 and SH3 cysteine mutant pairs by inducing disulfide bond formation with hydrogen peroxide in vitro. Active βarr1 was shown to bind more strongly to SH3 4; therefore, prior to disulfide trapping reactions βarr1 was activated using a synthetic phosphopeptide mimicking the C-tail of vasopressin 2 receptor (V2Rpp) and the stabilizing antibody fragment Fab30. Intriguingly, we observed the formation of disulfide cross-linked SH3–βarr1–V2Rpp–Fab30 complexes by two βarr1 mutants, E92C in the distal part of the N-domain (hereafter referred to as βarr1-N site) and P120C in the central crest region (hereafter referred to as βarr1-CC site) (Fig. 1c, d). In contrast, little to no SH3–βarr1 cross-linked complexes were formed by other βarr1 mutants, such as T116C (Fig. 1d). The SH3 mutant R95C in the RT-loop gave the strongest disulfide cross-linked band in both sites. As βarr1-N and βarr1-CC sites are more than 20 Å apart, we hypothesized that the SH3 domain might independently bind at two sites of βarr1. Complex formation at the βarr1-CC site was more efficient in the presence of V2Rpp and Fab30, whereas the βarr1-N site forms the complex equally well regardless of activation (Supplementary Fig. 2a).

SH3 E115C also formed a cross-linked complex with βarr1-CC (Fig. 1c). Analytical size-exclusion chromatography revealed that the SH3_R95C–βarr1_P120C complex displays a more prominent and symmetric peak compared to SH3_E115C–βarr1_P120C, suggesting greater homogeneity and stability (Supplementary Fig. 2b). βarr1 mutants P88C and P91C mutants also formed the cross-linked complex with SH3_R95C, but to a lesser degree than E92C suggesting the dynamic nature of the SH3–βarr1 interfaces (Fig. 1c).

To validate whether V2Rpp-activated βarr1 can bind SH3 at two distinct sites, we used isothermal titration calorimetry (Supplementary Fig. 2c). The integrated heat curve revealed two binding events: one with a Kd of ∼6 μM and the other with a significantly lower affinity of ∼50 μM. The favorable entropy (-TΔS) suggests that in both sites, SH3–hydrophobic interactions drive βarr1 binding. Furthermore, the favorable enthalpy (ΔH) at the higher affinity site indicates some contribution from hydrogen bonding and van der Waals forces.

βarr1 binds to the aromatic surface of SH3
To elucidate the mechanism of SH3 recruitment by βarr1, we formed cross-linked complexes with the best mutant pairs SH3_95C–βarr1_120C (SH3–βarr1-CC) and SH3_95C–βarr1_92C (SH3–βarr1-N) and determined the structures by cryo-EM (Fig. 2a–c, Supplementary Fig. 3, Supplementary Fig. 4, Supplementary Table 1). The map obtained from the SH3–βarr1-CC complex has a global resolution of 3.47 Å and showed a rigid interaction with SH3 in the central crest area of βarr1 (Fig. 2a, Supplementary Fig. 4e). The complex was activated by V2Rpp and stabilized by Fab30 and nanobody 32 (Nb32) 11. βarr1 adopts an active conformation 12, characterized by C-tail displacement and inter-domain rotation, and binds to the aromatic surface of SH3 containing residues from the RT loop, nSrc loop, and the 310 helix (Fig. 2b, Supplementary Video 1). βarr1 interacts with SH3 using β-strand V (residues 75-80) in the N domain and a part of the lariat loop in the C domain. Y90 and Y136 in SH3 interact with F75 in βarr1. The side chains of Y136 and N135 in SH3 are well-placed to make hydrogen bonds with the main chain oxygen of R76 in βarr1. Additional hydrogen bonds are possible between the main chain oxygen of E93 in SH3 and the side chain of N122 in βarr1, as well as between W118 in SH3 and the main chain oxygen of D78 in βarr1.

The SH3–βarr1-N complex was formed with V2Rpp and Fab30 but without Nb32, as Nb32 hinders complex formation due to the steric clashes with SH3. The cryo-EM map showed SH3 bound to the distal part of the N-terminal domain of βarr1 (Fig. 2c). Whereas SH3–βarr1-CC has a clearly resolved density for SH3, in SH3–βarr1-N the density for SH3 is weaker (local resolution is 4.2–4.6 Å) since SH3 binds to a flexible loop of βarr1, the 87FPPAPEDK94 motif (Fig. 2c, Supplementary Fig. 4f). We used molecular dynamics flexible fitting (MDFF) constrained by the position of the disulfide bond between βarr1 and SH3 to place the main chain of the SH3 domain. The proline region of the βarr1-N site (P88, P89, P91) appears to interact with the aromatic surface of SH3, similarly to the βarr1-CC site (Fig. 2c, Supplementary Video 2). However, the weak density for SH3 in the SH3–βarr1-N complex prevents confident identification of the specific SH3 residues involved in βarr1 binding. To confirm the interaction interfaces, we introduced mutations in SH3 and βarr1 and tested the binding by pull-down assay. Mutations in the RT loop (Y90A, Y92C) and the 310 helix (Y136A) of SH3 drastically reduced its ability to bind βarr1 (Fig. 2d). Likewise, the substitutions of prolines in βarr1 impair SH3 binding (Fig. 2e). These findings, together with the disulfide trapping data indicating that the RT loop faces the SH3–βarr1 interaction interface, confirm that residues within both the RT loop and 310 helix are critical for the SH3–βarr1 interaction at both sites of βarr1.

The structural and disulfide trapping data indicate a highly dynamic mode of SH3 binding to the βarr1-N site (Fig. 1c), with disulfide trapping capturing only a single state of this inherently dynamic interaction. Cross-linked complex formation by several SH3–βarr1 cysteine pairs (Fig. 1c) suggests that SH3 might adopt alternative orientations relative to the βarr1-N site, highlighting the complexity of low-affinity interactions in signaling cascades.

Notably, βarr1 uses a distinct recognition mechanism for SH3 binding at each site. The βarr1-N site binds SH3 exclusively via proline residues. In contrast, the recruitment of SH3 via the βarr1-CC site is driven by non-proline residues in β-strand V and the lariat loop, as well as two proline residues after β-strand VI.

Since the SH3–βarr1 complexes were stabilized using cysteine substitutions and disulfide trapping, we sought to determine whether these interactions also occur under physiological conditions, in the absence of engineered mutations. To address this, we mapped the interaction interfaces and dynamics of free βarr1, V2Rpp-activated βarr1, and SH3 using hydrogen–deuterium exchange mass spectrometry (HDX-MS) (Fig. 3; Supplementary Table 2; Supplementary Data 1–6). HDX-MS provides insights into protein interaction interfaces by measuring deuterium uptake in peptides typically 5–15 residues in length. Thus, this approach enables us to verify whether SH3 binds to two distinct regions of βarr1 in vitro without disulfide trapping. Approximately 5% of the analyzed peptides displayed a statistically significant decrease in deuterium uptake in the presence of SH3, defined as a statistically significant change greater than 0.3 Da for free βarr1 and 0.36 Da for βarr1–V2rpp (Supplementary Tables 2–3, Supplementary Data 1–2, 5). Consistent with the structures and the pull-down data, βarr1 showed a significant decrease in HDX rate in both the βarr1-CC and βarr1-N sites in the presence of SH3 (Fig. 3a, b; Supplementary Table 3). Although the differences in deuterium uptake for peptides corresponding to the βarr1-CC and βarr1-N sites were modest (0.2–0.8 Da), consistent with low-affinity interactions and partial complex occupancy arising from dilution during HDX labeling, the changes were reproducible across multiple time points and supported by several overlapping peptides in both free and V2Rpp-activated βarr1 (Supplementary Table 3; Fig. 3a, b; Supplementary Fig. 5a). Within these regions, longer peptides spanning both βarr1-CC and -N sites showed more pronounced changes, while shorter internal peptides displayed attenuated effects due to key interacting residue being positioned at the peptide N-terminus and increased back exchange (Supplementary Fig. 5a). Notably, a decrease in deuterium uptake was also observed for residues 34–47, which do not directly interact with SH3 in the structure but are located near the βarr1-N site, suggesting that SH3 binding may indirectly stabilize this region. Interestingly, reduced deuterium uptake was also detected in the hinge region of βarr1, in the C-domain spanning residues 257–310, and within the C-terminal tail (352–377) in V2Rpp-activated βarr1 (Fig. 3a). Since Src does not directly interact with the C-domain of βarr1 9, the reduced deuterium uptake in this region likely arises from long-range allosteric stabilization propagated through SH3 binding.

We next performed a qualitative HDX-MS experiment to assess changes in the deuterium uptake of SH3 in the presence of either free or V2Rpp-activated βarr1 (Supplementary Tables 2 and 4; Supplementary Fig. 5b–d; Supplementary Data 3–4, 6). Consistent with the structural data, peptides encompassing the nSrc loop and the 310 helix of SH3 exhibited decreased deuterium uptake, with more pronounced differences observed in the presence of V2Rpp-activated βarr1 (Supplementary Table 4; Supplementary Fig. 5b, c). In contrast, peptides corresponding to the RT loop predominantly showed increased deuterium uptake, likely reflecting local destabilization and enhanced conformational flexibility of the RT loop upon βarr1 binding. Taken together, the HDX-MS data reveal consistent qualitative trends that corroborate the disulfide trapping and cryo-EM results and support the presence of two SH3-binding regions on βarr1 and the interaction modes captured in the SH3–βarr1-CC and SH3–βarr1-N complexes.

We then compared the mechanisms of SH3 recruitment by various SH3-binding proteins and βarr1-N (Supplementary Fig. 6a-c) and βarr1-CC sites (Supplementary Fig. 6d, e) 13–17. In all structures, SH3 utilizes its aromatic surface to engage with its binding partners (Supplementary Fig. 6). The βarr1-N site resembles the canonical polyproline-rich motif observed in Nef and ELMO1, although the polyproline sequence in βarr1 is considerably shorter (Supplementary Fig. 6a, c). In contrast, the βarr1-CC site employs non-proline residues to recruit SH3 (Fig. 2b, Supplementary Fig. 6d, e). Non-canonical binding was previously reported for several SH3-binding proteins 18,19. For example, endophilin A1 SH3 interacts with histidine, proline and two arginine residues in parkin Ubl (Supplementary Fig. 6b); SH3 of Sla1 endocytic protein engages with histidine and hydrophobic residues of ubiquitin (Supplementary Fig. 6e) 15,16. βarr1 is distinguished from these examples by engaging SH3 through two mechanistically distinct binding sites, revealing an additional layer of complexity in βarr-mediated interactions.

βarr1 drives autoinhibition relief of Src
To understand how βarr1 activates Src, we aimed to determine the structure of βarr1 in complex with three-domain Src (SH3-SH2-SH1, residues 83-533). We performed disulfide trapping experiments using the most promising βarr1 and Src mutants, based on the results of the βarr1–SH3 screening (Fig. 1c, d). Consistent with the findings for the isolated SH3 domain, we observed the formation of Src–βarr1 complexes at both βarr1-N and βarr1-CC sites, with βarr1 P120C (βarr1-CC) forming the highest proportion of the cross-linked complex with Src R95C (Supplementary Fig. 7a, b). Therefore, we formed the Src_R95C–βarr1_P120C complex (hereafter referred to as Src–βarr1-CC) and determined its structure by cryo-EM (Fig. 4, Supplementary Fig. 7c, Supplementary Table 1). The final cryo-EM map has a resolution of 3.32 Å with a clear density for the SH3 domain (Fig. 4a, Supplementary Fig. 8a-c). The superposition of SH3–βarr1-CC and the Src–βarr1-CC revealed a translational shift of 2.0-2.2 Å of the SH3 domain towards the central crest of βarr1 in the Src–βarr1-CC complex (Supplementary Fig. 8d). The orientation of SH3 in Src–βarr1-CC and the interaction network with βarr1 are similar to the SH3–βarr1-CC complex (Figs. 2b, 4b, Supplementary Fig. 8d). The 2D classes of the complex show fuzzy density around SH3 suggesting the high flexibility of the SH2 and SH1 domains (Supplementary Fig. 8e). Consistent with this observation, SH2 and SH1 were not resolved in the final map. Filtering the map to a lower resolution (15 Å) revealed the density attributable to SH2 but not to SH1 (Fig. 4c), suggesting that βarr1-CC binding to SH3 induces displacement of the SH1 domain. To determine whether a similar mechanism operates at the βarr1-N site, we formed a complex between Src R95C and βarr1 E92C (Src–βarr1-N) and collected a cryo-EM dataset. This dataset had too few usable particles to obtain a reliable 3D reconstruction. Still, using the SH3–βarr1-N map as a reference, we aligned 2D class averages of the Src–βarr1-N complex with projections of the SH3–βarr1-N map (Supplementary Fig. 8f). This comparison revealed that the 2D projections of Src–βarr1-N closely resemble those of the SH3–βarr1-N complex, with the addition of fuzzy densities likely corresponding to the liberated SH2 and SH1 domains, mirroring the SH1 displacement observed in the Src–βarr1-CC complex.

Src activity is tightly regulated by three mechanisms of intramolecular interactions named the latch, clamp, and switch that conformationally restrict SH1, the catalytic domain of Src, and maintain its autoinhibited conformation (Supplementary Fig. 9a-c) 20,21. Unlatching, unclamping, and switching are required for full activation of Src. The latch involves phosphorylation of C-terminal tail at Y527 and its binding to SH2 (Supplementary Fig. 9b); the clamp mediates binding of SH3 RT loop, the 310 helix and the nSrc loop to the SH2–SH1 linker and SH1 (Supplementary Fig. 9c); the switch involves the autophosphorylation of Src activation loop Y416 by another Src molecule 21. Interestingly, superimposing the SH3–βarr1 complexes onto the crystal structure of autoinhibited Src phosphorylated at Y527 reveals steric clashes of βarr1 with the SH2 and SH1 domains, suggesting that βarr1 cannot access the SH3 domain in this conformation (Supplementary Fig. 9d). Under physiological conditions, however, the interactions between SH3, SH2, and SH1 domains are likely to be in a dynamic equilibrium between bound and unbound states, allowing the SH3 domain engaged with the SH1-SH2 linker to transiently expose its binding interface to βarr1. The affinities of these interactions are evolutionarily fine-tuned: strong enough to maintain the kinase in an autoinhibited conformation, yet sufficiently weak to respond to diverse cellular stimuli 22. A conformation of Src lacking C-terminal phosphorylation at Y527 may further favor βarr1 accessibility, as the unphosphorylated tail forms fewer stabilizing interactions with the SH2 domain (Supplementary Fig. 9b). Supporting this, the crystal structure of unphosphorylated Src (PDB: 1Y57) shows the C-tail in a liberated state, with the C-tail-binding pocket on SH2 occupied by a sulfate ion from the crystallization buffer containing 1.3 M ammonium sulfate (Supplementary Fig. 9e). In vivo, however, unphosphorylated Src likely exists in a dynamic equilibrium between C-tail-bound and C-tail-liberated states. To determine which Src conformation is favored by V2Rpp-activated βarr1, we compared the binding of βarr1 to Src with and without C-terminal phosphorylation by pull-down assay. Phosphorylation at Y527 was performed in vitro using purified Csk, a known negative regulator of Src activity, and confirmed by western blotting (Fig. 4d). Although βarr1 is still capable of binding Y527-phosphorylated Src, it shows stronger binding to the unphosphorylated form, indicating a preference for the more dynamic, flexible conformation of Src (Fig. 4d). Previous studies have shown that the absence of C-tail phosphorylation increases the flexibility of the linker between the SH3 and SH2 domains, known as an “inducible snap lock”, that may further enable the SH3 domain to accommodate not only extended polyproline sequences but also structurally diverse interaction partners 23. Therefore, in the unphosphorylated state, enhanced flexibility in the SH3–SH2 linker appears to promote βarr1 accessibility by transiently exposing SH3 binding interface to βarr1 (Fig. 4e). As evident from the structures, βarr1 binding to SH3 RT loop and the 310 helix disrupts the clamp mechanism of Src autoinhibition and releases the SH2–SH1 linker and the SH1 domain (Fig. 2a, b, Fig. 4b, Supplementary Fig. 9a). This is further attested by the Src–βarr1-CC structure, in which high flexibility of SH1 is consistent with its unrestricted movement and the disruption of the autoinhibited state. Therefore, βarr1 causes the autoinhibition relief of Src by interacting with SH3 and disrupting the intramolecular clamp (Supplementary Video 3).

In addition to SH3, βarr1 is also known to bind the SH1 domain of Src; however, this interaction does not lead to Src activation 4. Furthermore, in contrast to SH3, which preferentially interacts with active βarr1, SH1 shows stronger binding to the inactive form 4. Consistent with this, our structural data show that SH1 is displaced from its position in inactive Src and does not appear to associate with V2Rpp-activated βarr1 in the conditions of our cryo-EM experiments. To gain insight into how inactive βarr1 engages SH1, we performed lysine-specific cross-linking mass spectrometry (CXMS) using purified βarr1 and SH1. CXMS provides proximity restraints between intermolecular lysine residues and between lysines and proteins N-termini (Fig. 5a, b). Two cross-linked peptides were identified between the N-terminal region of βarr1 and K356 within the C-lobe of the SH1 domain, positioned on the face opposite to the active site (Fig. 5c). To validate the CXMS data, we tested SH1 binding to different βarr1 constructs (full-length, N-domain, and C-domain) by pull-down assay. Whereas the full-length and N-domain of βarr1 bound to SH1, the βarr1 C-domain (residues 176-418) showed no detectable binding to SH1, indicating that the interaction is mediated by the N-domain, in agreement with the cross-linking results (Fig. 5d). The structural data and the proximity restraints from cross-linking analysis suggest that βarr1 engages either SH3 or SH1, but not both simultaneously (Fig. 5e). We propose that inactive βarr1 interacts with Src through SH1; however, upon activation, this interaction appears weakened or lost, and binding shifts in favor of SH3. Further work will be needed to confirm this hypothesis.

SH3 binding induces conformational changes in βarr1
Interestingly, βarr1 not only induces conformational changes in Src but also undergoes structural rearrangements upon Src SH3 binding (Fig. 6). The most dramatic changes are observed in the central crest region of βarr1 in the Src–βarr1-CC and SH3–βarr1-CC complexes (Fig. 6a). SH3 binding to βarr1-CC drastically changes the position of β-strand V: it shows a downward and inward movement by ~9 Å and displays a two-residue offset in its N-terminal part and a three-residue offset in its C-terminal part as compared to active βarr1 without SH3 (Fig. 6a–b). Furthermore, the middle loop in SH3–βarr1-CC adopts an intermediate conformation between the fully active and fully inactive βarr1 crystal structures (Fig. 6c). While we cannot rule out that the differences in the conformation of the middle loop may come from crystal packing constraints or the allosteric effects from binding to GPCRs 24–27, it is tempting to speculate that the larger conformational changes in β-strand V upon Src SH3 binding might affect the coupling of βarr1 to the receptor and the downstream signaling. We also observed conformational changes in βarr1 within the SH3–βarr1-N complex in proximity to the interaction interface. Specifically, the 45PEYLKER51 loop shifts by ~6-8 Å toward SH3 and adopts a distinct conformation compared to that seen in the SH3–βarr1-CC complex and other active βarr1 structures (Fig. 6d). Furthermore, the 87FPPAPEDK94 motif is shifted by 4 Å toward SH3, although it remains unclear whether this change is due to disulfide cross-linking or SH3 binding (Fig. 6e). Notably, the interaction of βarr1 with Src also introduces local conformational changes in SH3 by affecting the position of the nSrc loop and converting the 310 helix into a loop (Fig. 6f).

Physiological roles of SH3-binding sites of βarr1
We next sought to elucidate the physiological consequences of SH3 binding to βarr1-N and βarr1-CC. We generated four βarr1 mutants targeting these regions and assessed Src activation in HEK-293 βarr1/βarr2 double-knockout (CRISPR-Cas9-based, dKO) cells 28 by monitoring phosphorylation of the activation loop tyrosine Y416 following GPCR stimulation. Upon stimulation of the chimeric β2-adrenergic receptor containing the V2R tail (β2V2R) with BI-167107, βarr1 mutants exhibited a moderate reduction in βarr-dependent Src activation (Fig. 7a), indicating partial redundancy between the two Src-binding sites in this context. Notably, mutant 3, harboring combined mutations in both the βarr1-N and βarr1-CC sites (P88G_P91G_P121E_P124G), did not further decrease Src activation compared with mutant 1 (P88G_P91G) or mutant 2 (P121E_P124G). This finding suggests that βarr1–Src interactions are broadly distributed, and that the mutations tested, while disruptive, are not sufficient to fully abolish βarr-mediated Src activation.

We also tested the activation of endogenous Src in HEK-293 βarr1/βarr2 dKO cells downstream of dopamine 1 (D1R) receptor by βarr1 mutants (Fig. 7b). We chose D1R because Src is one of the key effectors of D1R signaling 29, and its activation was shown to exclusively depend on βarr 30. While basal levels of phospho-Src were detected in all non-stimulated cells, dopamine stimulation produced a significant increase in phospho-Src in cells transfected with βarr1 WT. In contrast, most βarr1 mutants failed to respond to dopamine, indicating a reduced ability to activate Src, except for the βarr1-CC mutant P121E_P124G, which exhibited modest activation following stimulation (Fig. 7b). Taken together, both the βarr1-CC and βarr1-N sites contribute to Src activation in HEK293 cells downstream of two receptors, β2V2R and D1R.

To dissect the contribution of individual residues, we generated six βarr1 mutants and assessed their ability to mediate Src activation in vitro by monitoring phosphorylation of a Src peptide substrate, using V2Rpp and Fab30 to activate βarr1. Whereas βarr1 WT increased Src activity approximately two-fold, the βarr1-N mutant (P88G_P91G), βarr1-CC triple/quadruple mutants (P121E_P124G_F75A_N122A and P121E_P124G_F75A_I314A), as well as the double-site mutant (P88G_P91G_P121E_P124G), failed to activate Src (Fig. 7c). In contrast, CC-site mutants P121E_P124G and F80A only modestly reduced Src activation relative to WT. These results support a model in which residues P88 and P91 mediate activation via the βarr1-N site, while F75, N122, and I314 are essential for activation via the βarr1-CC site.

Interestingly, in the in vitro assay, mutation of either the βarr1-N site or selected CC-site residues completely abolished Src activation despite the presence of the other intact site. This finding indicates that in the minimal in vitro reconstitution system the two sites act cooperatively rather than independently: loss of one site compromises the ability of the other to promote activation. In cells, however, single-site mutants (all mutants downstream of β2V2R; P121E_P124G downstream of D1R) still retained partial ability to activate Src (Fig. 7a, b), suggesting that the relative contribution of each site and the extent of cooperativity between them are context-dependent, with additional scaffolding interactions and membrane confinement likely providing partial functional compensation.

We hypothesized that binding of the Src SH3 domain to the central crest region of β-arrestin1 (βarr1-CC), along with the resulting conformational changes in βarr1, could impair its engagement with the transmembrane core of GPCRs (Fig. 8a). We, therefore, evaluated whether excess of SH3 would interfere with the core coupling of βarr1 to the phosphorylated V2R in membranes. We labeled the finger loop of βarr1 V70C with an environment-sensitive bimane fluorophore (mBr) and measured the changes in fluorescence emission spectra with and without SH3 (Fig. 8b). Upon stimulation of the phosphorylated V2R with arginine-vasopressin peptide (AVP), the finger loop of βarr1 V70C binds to the receptor core, leading to a ~50% increase in bimane fluorescence. The presence of SH3 significantly reduces this effect, suggesting that SH3 binding to βarr1 sterically hinders the core coupling of βarr1 to V2R. Similar results were obtained with the purified phosphorylated chimeric M2 muscarinic receptor with V2R tail (M2V2R) reconstituted in lipid nanodiscs (Fig. 8c). These data suggest that high local concentration of Src can directly influence the conformational equilibrium of GPCR–βarr1 complexes by disrupting the fully engaged “core” conformation (Fig. 8d), which, in turn, may impair receptor desensitization and hinder the termination of G protein-mediated signaling. To test this hypothesis in an cellular context, we monitored real-time cAMP dynamics in HEK293 cells stably expressing the β2-adrenergic receptor (β2AR) using the live-cell FRET-based ICUE2 31 biosensor, which consists of a fusion between cyan and yellow fluorescent proteins (CFP-Epac2-YFP) (Fig. 8e). Stimulation of Gαs-coupled β2AR with isoproterenol triggers a rapid peak in cAMP levels, followed by a gradual decline as a result of β2AR desensitization. Overexpression of Src led to increased agonist-induced cAMP accumulation whereas the decline of cAMP accumulation was delayed compared to control cells, indicating that β2AR desensitization is affected (Fig. 8f).

Methods
Bacterial strains
Escherichia coli strain DH5α (C2987, New England Biolabs) was used for plasmid propagation. Protein expression was done in E. coli BL21 derivative strain (C3010, New England Biolabs).

Cell culture
Human embryonic kidney CRISPR-Cas9-based βarr1/βarr2 double knock-out cell line (HEK-293 βarr1/βarr2 dKO) 28 was maintained in Gibco Minimum Essential Media (MEM) supplemented with 1% penicillin-streptomycin and 10% (v/v) fetal bovine serum at 37 °C and 5% CO2. Expi-293F (Invitrogen, A14527) and tetracycline-inducible Expi293F cells (Invitrogen, A39241) were grown in suspension in Expi293 Expression Medium (Gibco) at 37 °C and 8% CO2.

Molecular biology
Bacterial expression constructs for wild-type (residues 2-418) and minimal cysteine (C59A, C125S, C140I, C150V, C242V, C251V, and C269S) and truncated (βarr1–MC-393) variants of rat βarr1 50, Fab30 51, and Nb32 11 have been previously reported and functionally verified. Plasmid pET28a-3D-Src expressing SH3, SH2 and SH1 domains of chicken Src (residues 83-533), plasmid expressing YopH phosphatase and Csk were generous gifts from Prof. John Kuriyan. For structural studies, a Src-5Cys mutant (C185S, C238S, C245S, C277S, C400S) was generated. Bacterial expression constructs for Src SH1 domain (250-536) and for βarr1 C-domain (176-418) were synthesized by GenScript. Plasmid expressing the SH3 domain of Src (residues 83-141) was designed by inserting a C-terminal FLAG-tag and a stop codon in pET28a-3D-Src. Mammalian expression constructs for human FLAG-M2-muscarinic receptor (M2R) with C-terminal sortase ligation consensus sequence (LPETGGH) and 6×His-tag 50, chimeric FLAG-β2-adrenergic receptor with C-terminal tail of vasopressin 2 receptor (β2V2R) 52, SNAP-vasopressin 2 receptor (V2R) 43, HA-tagged βarr1 9, and human Src 9 have been previously reported and functionally verified. Mammalian expression construct for HA-dopamine 1 receptor (D1R) was a gift from Prof. Marc Caron. Mutations were introduced using QuickChange II site-directed mutagenesis kit (Agilent) and verified by Sanger sequencing. Oligonucleotides used for site-directed mutagenesis are listed in Supplementary Table 5.

Protein expression and purification
The Src construct (residues 83-533) without the N-terminal disordered region was used for the structural and in vitro studies 53. Src was co-expressed in E. coli with YopH phosphatase and purified by immobilized metal ion affinity chromatography and anion exchange chromatography. The SH3 domain of Src (residues 83-141) and its variants and the SH1 domain of Src (251-533) were expressed in E. coli and purified by immobilized metal ion affinity chromatography followed by size exclusion chromatography (SEC). Wild-type βarr1 54 and its variants were expressed in E. coli and purified using GST affinity chromatography followed by anion-exchange chromatography. Fab30 55 was expressed in E. coli and purified using protein A affinity chromatography. Nb32 11 was expressed in E. coli and purified using immobilized metal ion affinity chromatography followed by SEC. Csk 56 was expressed in E. coli and purified using GST affinity chromatography followed by anion-exchange chromatography. M2R containing a C-terminal sortase ligation consensus sequence (LPETGGH) 25,50 was stably expressed in tetracycline-inducible Expi293F cells (Invitrogen, A39241), solubilized using 1% n-dodecyl-β-D-maltopyranoside (Anatrace), 0.05% cholesterol hemisuccinate and purified by M1 FLAG affinity chromatography. The synthetic phosphopeptide GGG-V2Rpp (GGG-ARGRpTPPpSLGPQDEpSCpTpTApSpSpSLAKDTSS) was ligated to the C-terminus of M2R using sortase, and the monomeric chimeric M2V2R was collected by SEC 50. M2V2R was reconstituted in lipid nanodiscs using Membrane Scaffold Protein 1, variant D1 with E3 helices added (MSP1D1E3) and a 3:2 molar ratio of phosphatidylcholine with phosphatidylglycerol. Detergent was removed using Biobeads (BioRad) 25.

Preparation of crude membranes with V2R and G-protein coupled receptor kinase 2 (GRK2)
Expi-293 cells grown in suspension were co-transfected with plasmids expressing the human V2R and the membrane targeted form of GRK2 to promote receptor phosphorylation. Transfected cells were grown in suspension for 48 hours and thereafter were stimulated with AVP (100 nM) for 30 min at 37 °C followed by harvesting by centrifugation at 4 °C. Cell pellets were washed with cold Hanks’ Balanced Salt Solution, snap frozen in liquid nitrogen and stored at −80 °C prior to preparation of crude membranes 57. Frozen cells were dounce-homogenized in cold homogenization buffer (20 mm Tris-HCl pH 7.4), containing EDTA-free protease inhibitor cocktail (Roche) and a PhosSTOP phosphatase inhibitor cocktail (Roche). Following differential centrifugation, the P2 microsomal membrane fraction was resuspended in cold buffer (50 mm Tris-HCl, 150 mm NaCl, 12.5 mm MgCl2, 0.2% BSA, and 10% glycerol, pH 7.4), supplemented with protease inhibitor cocktail and PhosSTOP phosphatase inhibitor cocktail, aliquoted, and stored at −80 °C until use.

Disulfide trapping
βarr1–MC-393 mutant (30 μM) was incubated with 3-fold molar excess of V2Rpp (ARGRpTPPpSLGPQDEpSCpTpTApSpSpSLAKDTSS), 1.5 molar excess of Fab30, and 2-fold molar excess of SH3 mutant or Src mutant in 20 mM HEPES 7.5, 150 mM NaCl buffer for 30 minutes at room temperature. The disulfide trapping reactions were initiated by 1 mM H2O2. After 1-hour incubation at room temperature the reactions were terminated by addition of 4x Laemmli sample buffer (without β-mercaptoethanol) (BioRad), subjected to SDS-PAGE, visualized by ReadyBlue Coomassie stain (Sigma-Aldrich) and quantified by ImageJ v1.52a. The band of SH3–βarr1 complex (∼60 kDa) or Src–βarr1 (∼130 kDa) was normalized to the total density of all bands in each sample. To test the effect of V2Rpp and Fab30 on formation of disulfide trapped complexes, the reactions were performed as described above; in reactions without V2Rpp and Fab30 the equivalent amount of buffer was added. The samples were subjected to SDS-PAGE and Western blotting, detected by rabbit polyclonal A1CT antibody generated in Lefkowitz lab 58 (1:5000) and HRP-conjugated ECL Rabbit IgG (NA9340, Amersham, RRID:AB_772191) (1:5000) for βarr1 and HRP-conjugated Anti-6X His-tag antibody (1:2000) (ab1187, Abcam, RRID: AB_298652) for SH3.

Isothermal titration calorimetry (ITC)
ITC measurements were performed using the MicroCal PEAQ-ITC system (Malvern). Purified βarr1–V2Rpp and SH3 were dialyzed in 20 mM HEPES, pH 7.5, 100 mM NaCl. 75 μM of βarr1–V2Rpp was loaded into the sample cell and 750 μM of SH3 into the injection syringe. The system was equilibrated to 25 °C. Titration curves were initiated by a 0.4 μL injection from syringe, followed by 2.0 μL injections (at 180 s intervals) into the sample cell. During the experiment, the reference power was set to 7 μcal·s-1 and the sample cell was stirred continuously at 750 rpm. Raw data, excluding the peak from the first injection, were baseline corrected, peak area integrated, and normalized. Data were analyzed using MicroCal Origin software to obtain thermodynamic parameters of binding and association constant (Ka=1/Kd).

Formation of SH3–βarr1 and Src–βarr1 complexes
The βarr1–MC-393 mutant (100 μM) was incubated with 2-fold molar excess of the SH3 mutant or Src mutant for 30 minutes at room temperature, followed by disulfide trapping as described above. SH3–βarr1 was separated from unbound βarr1 by immobilized metal ion affinity chromatography using Talon resin (Takara Bio). Src–βarr1 was separated from unbound βarr1 and Src by SEC. SH3–βarr1 and Src–βarr1 complexes were then incubated with 3-fold molar excess of V2Rpp, 1.5-fold molar excess of Fab30, 2-fold molar excess of Nb32 (for SH3–βarr1-CC and Src–βarr1-CC) for 30 minutes at room temperature. The complexes were subjected to SEC on a Superdex 200 Increase column (Cytiva Life Sciences) in 20 mM HEPES 7.5, 150 mM NaCl buffer. Peak fractions were concentrated to 6-8 mg/ml using Vivaspin® 6 column with molecular weight cut-off of 30,000 kDa (Sartorius).

Cryo-EM grid preparation and data acquisition
In the preliminary cryo-EM experiments, we observed that the complexes exhibited strong preferred orientation in vitreous ice. To alleviate the preferred orientation problem, 8 mM CHAPSO was added to the sample immediately before vitrification 59. The sample was applied to glow-discharged 300-mesh holey-carbon grids (Quantifoil R1.2/1.3, Electron Microscopy Sciences) or gold grids (Ultrafoil R0.6/1.0, Electron Microscopy Sciences) and vitrified using a Vitrobot Mark IV (Thermo Fisher Scientific) at 4 °C and 100% humidity. The data were collected on a Titan Krios transmission electron microscope (Thermo Fisher) operating at 300 kV equipped with a K3 direct electron detector (Gatan) in counting mode with a BioQuantum GIF energy filter (slit width of 20 eV) at a magnification of ×81,000 corresponding to a pixel size of 1.08 Å at the specimen level. 60-frame Videos with a dose rate of ~15 electrons per pixel per second and a total accumulated dose of ~53-60 electrons per Å2 were collected using the Latitude-S (Gatan) single-particle data acquisition program. The nominal defocus values were set from −0.8 to −2.5 μm.

Cryo-EM data processing
Videos were subjected to beam-induced motion correction using Patch Motion Correction in CryoSPARC v4.0.1 60 followed by determination of Contrast transfer function (CTF) parameters in Patch CTF. Micrographs with CTF fit better than 3.5 Å were used for further analysis. Particles manually selected from 15 micrographs were used to train a model in the particle picking tool Topaz v0.2.5a 61. The trained model was used to pick particles in all micrographs generating 200,270 particle projections for the SH3–βarr1-CC complex and 760,233 particle projections for the SH3–βarr1-N complex. The particles were rescaled to the pixel size of 1.3824 Å for further processing. For the SH3–βarr1-CC complex, the particle stack was subjected to 2D classification, Ab Initio model generation, non-uniform refinement and local refinement with a mask excluding the constant (CL/CH1) region of Fab30 (FabCR) in CryoSPARC. Particles were then imported to RELION v3.1 62 and subjected to 3D classification without alignment with a mask excluding the constant (CL/CH1) region of Fab30 (FabCR). Classes showing a clearly defined density for the SH3 domain (118,020 particles) were re-imported to CryoSPARC and subjected to local refinement with a mask on FabCR and a fulcrum on the SH3–βarr1 interface (Supplementary Fig. 3a). The resulting map was then processed by spIsoNet to improve map isotropy 63. The final map has a global resolution of 3.47 Å at a Fourier shell correlation of 0.143 (Supplementary Fig. 4c). For the SH3–βarr1-N complex, the particles picked with Topaz were subjected to 2D classification and the best 2D classes were used in template picking (5,607,258 particles). A subset of particles (1,000,000) was used to generate six Ab Initio classes. All particles were then subjected to four rounds of heterogeneous refinement. The resulting particle stack (345,529) was used in non-uniform refinement and local refinement with a mask excluding FabCR (Supplementary Fig. 3b). The resulting map was then subjected to spIsoNet to improve map isotropy 63. The final map has a global resolution of 3.34 Å at a Fourier shell correlation of 0.143 (Supplementary Fig. 4d). For the Src–βarr1-CC complex, the particles picked with Topaz (94,196 particles) were subjected to 2D classification and the best 2D classes were used in template picking (9,770,378 particles). The particles were rescaled to the pixel size of 1.44 Å for further processing. A subset of particles (1,000,000) was used to generate three Ab Initio classes. All particles were then subjected to three rounds of heterogeneous refinement, followed by non-uniform refinement and local refinement with a mask excluding FabCR. The resulting particle stack (692,850) was subjected to 3D classification without alignment in CryoSPARC and then in RELION with a mask excluding FabCR. Finally, the best class with 140,156 particles was subjected to another round of local refinement in CryoSPARC, generating a map with a global resolution of 3.32 Å (Supplementary Fig. 7c, Supplementary Fig. 8b). For the Src–βarr1-N complex, the particles picked with Topaz (61,962 particles) were subjected to 2D classification and matched with the 2D projections of the SH3–βarr1-N map using cluster selection mode in Reference Based Auto Select 2D module in CryoSPARC.

Model building and refinement
Cryo-EM maps for SH3–βarr1 complexes were post-processed using DeepEMhancer 64 to improve the interpretability of maps, specifically in the βarr1 loops and at the SH3–βarr1 interface. Since the overall resolution of maps was better than 4 Å, highRes deep learning model was used during DeepEMhancer processing of maps. The initial models of the SH3–βarr1-CC complex and Src–βarr1-CC complex were built manually by fitting the crystal structures of SH3 (PDB: 2PTK), βarr1–V2Rpp–Fab30–Nb32 complex (PDB code: 6NI2) into the experimental electron densities using UCSF Chimera v1.15 65. The structures were refined by combining manual adjustments in Coot v0.9.8.3 66 and ISOLDE 67 in UCSF ChimeraX 1.6.1 68, followed by real-space refinement in PHENIX v1.20.1-4487 69 with Ramachandran, rotamer, torsion, and secondary structure restraints enforced. For the SH3–βarr1-N complex, the crystal structure of the βarr1–V2Rpp–Fab30 complex (PDB code: 4JQI) was first built manually into the density and refined similarly to the SH3–βarr1-CC complex. After the refinement, SH3 (PDB: 2PTK) was docked into the density guided by the position of the disulfide bond between βarr1_E92C and SH3_R95C. The rigid body fitted SH3–βarr1-N model generated above was then subjected to molecular dynamics flexible fitting (MDFF) using Cryofit 70, integrated in Phenix software suite and Namdinator 71 followed by further iterative rounds of model building and real space refinement in Coot v0.9.8.3 66 and PHENIX v1.20.1-4487 69. Due to weak density for the SH3 domain in the SH3–βarr1-N complex, only its main chain was modeled. The models were validated with MolProbity v4.5.1 72. Refinement statistics are given in Supplementary Table 1.

Hydrogen-deuterium exchange mass spectrometry
To map the βarr1 interface involved in SH3 binding, βarr1–MC-393 (20 μM) or βarr1–MC-393–V2Rpp (20 uM βarr1; 100 uM V2Rpp) were incubated with 5-fold molar excess of SH3 in the 20 mM HEPES 7.5, 100 mM NaCl buffer for 30 minutes on ice. Conversely, to identify the βarr1-interacting interface on SH3, SH3 (20 uM) was incubated with 5-fold molar excess of βarr1–MC-393 or βarr1–MC-393–V2Rpp in the 20 mM HEPES 7.5, 100 mM NaCl buffer for 30 minutes on ice. The protein stock solution was diluted 17-fold in the 20 mM HEPES 7.5, 100 mM NaCl buffer prepared in D2O. The protein stock solution was diluted 17-fold into D2O-based buffer (20 mM HEPES pH 7.5, 100 mM NaCl) to achieve a final D2O fraction of 94% and to maintain an optimal dynamic range of deuterium uptake, thereby improving the reproducibility of HDX-MS measurements. The deuterium labeling reactions were performed 15 °C. At a designated time point (300, 1000, and 5000 s), the exchange was quenched by adding an equal volume of 0.8% formic acid in H2O, yielding pH 2.6. The sample was digested on Enzymate BEH Pepsin column (Waters) at a flow rate of 0.15 ml/min using 0.15% formic acid/3% acetonitrile as the mobile phase. An inline 4-μl C8-Opti-lynx II trap cartridge (Optimize Technologies) was used for desalting of the digested peptides. The peptides were then eluted through a C-18 column (Thermo Fisher Scientific, 50 × 1 mm Hypersil Gold C-18) using a rapid gradient from 10 to 90% acetonitrile containing 0.15% formic acid and a flow rate of 0.04 ml/min, leading directly into a maXis-II ETD ESI-QqTOF mass spectrometer (Bruker Daltonics). Up to 30, 150, and 150 pmol of βarr1, SH3, and V2Rpp, respectively, were used in each experiment to analyze βarr1 dynamics and interactions. To assess SH3 dynamics and interactions, up to 30, 150, and 150 pmol of SH3, βarr1, and V2Rpp, respectively, were used. An excess of proteins was included to ensure saturation of complex formation. The total time for the digest and desalting was 3 min, and all peptides had eluted from the C-18 column by 15 min. Pepsin and C-18 column were thoroughly washed after each run. The peptide fragments were identified using Bruker Compass and Biotools software packages. The level of deuterium incorporation was calculated using HDExaminer-3 (Trajan Scientific) from triplicate measurements of each time point. Only regions that exhibited statistically significant differences in deuterium uptake greater than 0.30 Da for free βarr1 and 0.36 Da for βarr1–V2rpp were considered relevant. To improve our confidence in the observed deuterium uptake differences, we sought to increase peptide coverage by analyzing a non-deuterated βarr1 sample. This analysis was performed under the previously employed HDX-LC conditions using a timsTOF fleX MALDI-2 mass spectrometer with ion mobility engaged. The identified peptides were then mapped to the existing HDX dataset. Although peptide overlap in the HDX dataset prevented the analysis of all peptide sequences, this approach ultimately increased the number of analyzable peptides by approximately 30%. Additionally, we conducted further experiments to measure the fully deuterated exchange profiles of βarr1 to estimate back-exchange. To prepare the fully deuterated protein, we first created a 480 μL stock solution of 3 M deuterated GuHCl in D2O. Subsequently, 20 μL of a 20 μM βarr1 protein stock was added to this GuHCl/D2O solution. The sample was then incubated at 42 °C for three days to allow for deuterium exchange. The GuHCl was deuterated beforehand by dissolving it in D2O and lyophilizing the solution to dryness. The peptides exhibited back exchange values ranging from 10% to 53%, with an average of 31%.

M2-FLAG pull-down assay
To test the binding between βarr1 and the SH3 domain of Src, βarr1 (20 μM) was incubated with 3-fold molar excess of V2Rpp for 30 minutes at room temperature, then 10 μM of SH3-FLAG was added. 30 μl of anti-FLAG M2 affinity gel (Millipore Sigma) was added thereafter and the mixture was incubated for 1 hour at room temperature with rotation. After incubation the anti-FLAG M2 resin was collected by centrifugation and washed with 1 ml of 20 mM HEPES pH 7.5, 100 mM NaCl buffer three times. The proteins were eluted with 0.2 mg/ml FLAG-peptide in 20 mM HEPES pH 7.5, 100 mM NaCl buffer, then mixed with Laemmli sample buffer (BioRad), subjected to SDS-PAGE and Western blotting and detected by monoclonal ANTI-FLAG M2-peroxidase (HRP) antibody (1:2000) (A8592, Sigma-Aldrich, RRID: AB_439702) for SH3-FLAG and polyclonal A1CT antibody 58 (1:5000) and HRP-conjugated ECL Rabbit IgG (1:5000) (NA9340, Amersham, RRID: AB_772191) for βarr1.

In vitro Src C-tail phosphorylation followed by M2-FLAG pull-down assay
Purified Src (20 μM) was incubated with 4 μM Csk in the presence of 100 μM ATP and 5 mM MgCl2 for 2 hours at room temperature. A control reaction was prepared identically, excluding ATP and MgCl2. The C-tail phosphorylation of Src was assessed by Western blotting using Anti-Src (phospho Y529) antibody (ab32078, Abcam, RRID: AB2286707). The samples of C-tail phosphorylated and unphosphorylated Src (20 μM) were added to 7 μM of βarr1-FLAG pre-incubated with 35 μM of V2Rpp. 30 μl of anti-FLAG M2 affinity gel (Millipore Sigma) was added thereafter and the mixture was incubated for 1 hour at room temperature with rotation. After incubation the anti-FLAG M2 resin was collected by centrifugation and washed with 1 ml of 20 mM HEPES pH 7.5, 100 mM NaCl buffer three times. The proteins were eluted with 0.2 mg/ml FLAG-peptide in 20 mM HEPES pH 7.5, 100 mM NaCl buffer, then mixed with Laemmli sample buffer (BioRad), subjected to SDS-PAGE and Western blotting. The total Src was detected by anti-Src antibody (1:5000) (EMD Millipore 05-184, RRID: AB_2302631) and HRP-conjugated ECL Mouse IgG (1:5000) (NA9310, Amersham, RRID: AB_772193). βarr1-FLAG was detected by monoclonal ANTI-FLAG M2-peroxidase (HRP) antibody (1:2000) (A8592, Sigma-Aldrich, RRID: AB_439702). Western Blot images were taken using BioRad ChemiDoc system and the densitometry analysis was performed by Image Lab v6.1.

Cross-linking mass-spectrometry
Purified βarr1 (90 μM) was incubated the purified SH1 domain of Src (90 μM) for 1 hour at room temperature. Cross-linking reagents (disuccinimidyl suberate (DSS), Bis(sulfosuccinimidyl) suberate (BS3), dithiobis(succinimidyl propionate) (DSP), 3,3´-dithiobis(sulfosuccinimidyl propionate) (DTSSP), ethylene glycol bis(succinimidyl succinate) (EGS)) were added thereafter to a final concentration of 2 mM. The reactions were incubated for 30 min and quenched by 50 mM Tris pH 8.0 for 15 min. The proteins were subjected to SDS-PAGE and visualized by ReadyBlue Coomassie stain (Sigma-Aldrich). The bands of interest (80-150 kDa) were excised from the gel and subjected to destaining using 25 mM ammonium bicarbonate in 50% acetonitrile. Destained gel slices were dehydrated using 100% acetonitrile and dried using speed-vac for 5 min to remove remaining acetonitrile. 10 mM dithiothreitol and 55 mM iodoacetamide were used for reduction and alkylation of cysteine residues, respectively. The gel slices were washed again in 25 mM ammonium bicarbonate and dried using 100% acetonitrile following by 5-min speed-vac evaporation. The proteins were then subject to in-gel trypsin digestion with 10 ng/μl trypsin (Sequencing grade, Promega) in 25 mM ammonium bicarbonate and overnight at 37 °C. The following day, the tryptic peptides were extracted with 100% acetonitrile and lyophilized. The peptides were loaded onto EvoSep tips (EvoSep) following manufacturer’s instructions. The peptide samples were analyzed using a timsTOF Pro 2 mass spectrometer (Bruker) coupled with an Evosep One LC system (Evosep) connected to an 8-cm Evosep performance column (EV-1109) using a pre-defined 30-samples-per-day extended gradient. The following parameters were applied for data-dependent acquisition: intensity threshold: 2.5E3; charge states: +1 to +5; dynamic exclusion: 0.4 min; target intensity for fragmentation: 2E4; isolation window (linear): 2 m/z at 700 m/z and 3 m/z at 800 m/z; collision energy (linear): 20 eV at 1/k0 of 0.60 V·s/cm2 and 59 eV at 1/k0 of 1.60 V·s/cm2. The cross-linked peptides were identified using MaxQuant v.2.6.3.0 with following parameters: digestion mode Trypsin/P; maximum missed cleavages 3; minimum length for a paired-peptide sequence 6; minimum score for cross-linked peptides 100; minimum matches 3; first search peptide tolerance 20 ppm; main search peptide tolerance 10 ppm; isotope match tolerance 0.005 Da, centroid match tolerance 10 ppm; maximum charge 6; isobaric weight exponent 0.75.

GST pull down assay
To test the binding between different constructs of GST-βarr1 (full-length βarr1, βarr1-N domain (1-176), βarr1-C domain (177-420)) and the SH1 domain of Src, GST-βarr1 (3 μM) was incubated with SH1 (9 μM) for 1 hour at room temperature. 50 μL of GST-beads (GoldBio) equilibrated in 20 mM HEPES pH 7.5, 100 mM NaCl buffer were added thereafter, and the mixture was incubated for 1 hour at room temperature with rotation. After incubation the GST beads were collected by centrifugation and washed with 20 mM HEPES pH 7.5, 150 mM NaCl buffer three times. The proteins were eluted from GST beads with 100 μL of 50 mM reduced glutathione in 20 mM HEPES pH 8.0, 150 mM NaCl buffer, then mixed with 4x Laemmli sample buffer (BioRad), subjected to SDS-PAGE and Western blotting. SH1 was detected by polyclonal HRP-conjugated Anti-6X His-tag antibody (1:2000) (ab1187, Abcam, RRID: AB_298652); βarr1 was detected by monoclonal HRP-conjugated GST-tag antibody (1:2000) (8-326) (MA4-004-HRP, Thermo Fisher, RRID: AB_2537634).

In vitro Src continuous colorimetric kinase assay
Continuous colorimetric kinase assay 73 was performed in 100 mM HEPES 7.5, 150 mM NaCl, 5 mM MgCl2, 0.005% Triton X-100, containing 0.25 mM optimal Src peptide (AEEEIYGEFEAKKKK), 1 mM phosphoenolpyruvate, 0.3 mM NADH, 4 units of pyruvate kinase and 6 units of lactic dehydrogenase. The concentration of Src was 20 nM, the concentrations of βarr1 and V2Rpp were 100 nM and 200 nM, respectively. Reactions were started by the addition of ATP to a final concentration of 0.1 mM, and the decrease in NADH absorbance was monitored over 40 min at 25 °C using a CLARIOstar microplate reader (BMG Labtech). The initial velocity of the reaction (V0) determined using a linear regression curve fit (GraphPad Prism v9) and was converted to the amount of product formed in the reaction volume per minute using the Beer-Lambert law. Statistical comparisons were determined by one-way ANOVA with Dunnett’s post hoc test.

Src activation assay in HEK-293 βarr1/βarr2 dKO cells
HEK-293 βarr1/βarr2 dKO cells 28 were co-transfected with receptor (FLAG-D1R or chimeric FLAG-β2V2R), Src (only for chimeric FLAG-β2V2R) and wild-type or mutant βarr1 with a C-terminal HA tag with a 1:5 DNA:FuGENE® 6 (Promega) ratio according to the manufacturer’s instructions. All experiments were conducted 48 hours after transfection. Cells were serum starved for 16 hours in MEM supplemented with 1% penicillin-streptomycin and 0.1% (w/v) bovine serum albumin. The assay was initiated with 5-minute stimulation (10 μM dopamine for D1R) or 10-minute stimulation (10 μM BI-167107 for β2V2R) at 37 °C. The medium was removed, cells were placed on ice and lysed with 2x Laemmli sample buffer (BioRad) supplemented with 4% β-mercaptoethanol. Samples of equal volume from cell lysates were separated with SDS-PAGE, transferred to nitrocellulose membranes and immunoblotted. Primary antibodies and dilution used areas follow: monoclonal ANTI-FLAG M2 peroxidase (HRP) antibody (1:2000) (A8592, Sigma-Aldrich, RRID: AB_439702) to detect β2V2R, polyclonal A1CT antibody generated in Lefkowitz lab 58 (1:5000) for wild-type and mutant βarr1, anti-Src monoclonal antibody (1:1000) (MA5-15214, Thermo Fisher Scientific, RRID: AB_10980540) for total Src, anti-Src polyclonal Y418 for overexpressed Src (1:5000) (ab4816, Abcam, RRID: AB_304652), and Phospho-Src family polyclonal Y416 for endogenous Src (1:10000) (2101, Cell Signaling, RRID: AB_331697). Secondary antibody included HRP-conjugated ECL Rabbit IgG (1:5000) (NA9340, Amersham, RRID: AB_772191) and HRP-conjugated ECL Mouse IgG (1:5000) (NA9310, Amersham, RRID: AB_772193). Western Blot images were taken using BioRad ChemiDoc system and the densitometry analysis was performed by ImageJ v1.52a and Image Lab v6.1.

Bimane fluorescence
Purified βarr1-MC-393 V70C was labeled overnight at 4 °C with a 3-fold molar excess of monobromobimane (mBr) (Sigma-Aldrich) and an additional 3-fold molar excess added for 1 h at room temperature the next day. Reactions were quenched with L-cysteine, and free mBr was removed by SEC. The experiments were performed using membranes with phosphorylated V2R and purified chimeric M2V2R reconstituted in lipid nanodiscs. Aliquots of phosphorylated V2R membranes were activated with AVP (10 μM) then incubated with βarr1 V70C-mBr (20 nM), and SH3 (250 nM) in the assay buffer (20 mM HEPES 7.4, 100 mM NaCl, 1 mg/ml bovine serum albumin) for 120 min at room temperature in black solid-bottom 96-well microplates (Corning) with gentle agitation. Purified M2V2R reconstituted in lipid nanodiscs (25 nM) was activated with 10 μM iperoxo and subsequently incubated for 60 min at room temperature with 1.5-fold molar access of bimane-labeled βarr1, 2-fold molar excess of Fab30 and SH3 (500 nM). Fluorescence emission spectra were collected in top-read mode, with excitation at 370 nm (16 nm bandpass) and emission scanning from 410 nm to 640 nm (10 nm bandpass) in 0.5 nm increments using a CLARIOstar microplate reader (BMG Labtech). Statistical comparisons were determined by comparing the area under the curves (GraphPad Prism v9) and peak fluorescence using one-way ANOVA with Dunnett’s post hoc test from n=3 (M2V2R) or n=6 (V2R membranes) independent experiments.

Fluorescence Resonance Energy Transfer (FRET)-based live-cell cAMP accumulation assay
cAMP production mediated by Gαs-coupled β2AR activation was measured using Exchange Protein Activated by cAMP 2 (Epac2) sensor containing a CFP and YFP FRET pair 31. HEK293 cells stably expressing Epac2 and β2AR were transfected with Src (2 ug) or the equivalent amount of control plasmids (pcDNA 3.1) with FuGENE® 6 (Promega) transfection reagent according to the manufacturer’s instructions. 20 hours after transfection the cells were plated in poly-D-lysine-coated, black, clear-bottom 96-well plates (Corning) at a density of 50,000 cells per well. 24 hours after plating, cells were washed with PBS and incubated in imaging buffer (10 mM HEPES, 150 mM NaCl, 5 mM KCl, 1.5 mM MgCl2, 1.5 mM CaCl2, 10 mM glucose, 0.2% BSA, pH 7.4) for one hour at 37 °C. The cells were stimulated with 10 μM isoproterenol (ISO), and FRET changes were measured in real-time using FlexStation 3 plate reader (Molecular Devices). The changes in the background-subtracted 480 nm/535 nm fluorescence emission ratio (CFP/YFP) are indicative of changes in cAMP levels. To quantify the overall cAMP response, the area under the curve (AUC) was calculated from the time-course data in GraphPad Prism v9.

Statistics and reproducibility
Data were analyzed using GraphPad Prism v9. Data represent the mean ± SEM of at least three independent experiments. Statistical significance for more than two groups was determined by one-way ANOVA with Dunnett’s post hoc test. Statistical significance for two groups was determined by unpaired two-tailed Student’s t-test or two-tailed Welch’s t-test. No samples or data points were excluded from analysis. Uncropped and unprocessed scans of the blots are provided in the Source Data file.

The materials presented are available upon request from Robert J. Lefkowitz (lefko001@receptor-biol.duke.edu).

Peer review rounds
[{"round": 1, "reviewer_comments": {"reviewer_1": "In this manuscript, Pakharukova et al showed that b-arr1 interacts with the Src SH3 domain via two distinct sites (N- and CC-) and activates the auto-inhibited Src, thereby plays as an active signal transducer. Authors have identified two distinct sites of the Src SH3 domain that bind to b-arr1, determined the complex structures using cryoEM, and performed various biochemical and mutational analysis to support the activation mechanism of Src by b-arr1. They properly addressed concerns raised by previous reviewers and the revised manuscript provides insights into understanding the molecular mechanism by which Src activated by b-arr1.\nAlthough most of the concerns by the previous reviewers have been addressed, I am bit confused on the mutational analysis that performed to understand the physiological meaning of the current study (Fig 7 and lines 275 ~ 288). Unlike the authors described in the discussion session “… this configuration enables βarr1 to ensure robust recruitment and activation of Src when one of the interaction sites is sterically hindered or occupied by a competing binding partner…”., I noticed that mutation of either site exhibited similar to that of the b-arr1 lacking effect in Fig 7.\nQ1. Fig 7a, mutation of both N- and CC sites (mut 3) exhibited higher normalized density for pSrc compared to mut 1 or mut 2. Perhaps, interaction sites are broadly distributed and multiple mutation may not provide decisive effects. Nevertheless, after reading the entire manuscript, the mutation studies are puzzling. Authors may comment on their mutational analysis.\nQ2. Mutation on one of the interaction sites should exhibit some degree of activation. In Fig 7c, mutation on N-site (mut 1) or CC-site (mut 5 or 6) exhibited an effect similar to reaction in the absence of b-arrestin. Authors should comment on this.\nRelated to above Q. Lines 285~290\n“ Whereas β-arr1 WT (activated by V2Rpp and Fab30) increases Src activity two-fold, βarr1-N mutant P88G_P91G, .. as well as P88G_P91G_P121E_P124G mutant failed to activate Src (Fig. 7c). βarr-CC mutants P121E_P124G and F80A only slightly reduced Src activation compared to βarr1 WT. Therefore, this supports a mechanism where P88 and P91 drive Src activation via the βarr1-N site and F75, N122 and I314 are critical for Src activation via the βarr1-CC site.”\nWhy N-mutant failed to increase the Src activation despite it has functional CC-site?\nQ3. Line 281-282\n“..In contrast, none of the βarr1 mutants showed a response to dopamine stimulation, suggesting their reduced ability to activate Src (Fig. 7b)…”\nThe band intensity of each line is difficult to quantify by eyes. But mutant 2 (cc-mutant) showed similar increase to that of the WT (and showed some response). Authors may consider to revise this part for clarity.", "reviewer_2": "Please find my technical review of the manuscript by Pakharukova et al. below. The current presentation and analysis of the HDX-MS data are insufficient to support the main conclusions of the study. The data as presented do not meet the current standards for reporting and validation in the field, which prevents a full and fair evaluation of the work. The suggestions below focus on areas relevant to HDX-MS experiments that should be clarified and expanded to enhance the paper's impact.\nFor this reason, I cannot recommend publication in its present form. However, the concerns are addressable. I would be willing to re-evaluate a revised manuscript that thoroughly addresses the major points listed below.\nMajor points:\nThe central issue with the manuscript is the lack of a comprehensive and transparent presentation of the HDX-MS data. Without this, the technical quality of the experiment and the statistical validity of the conclusions cannot be verified.\nThe manuscript currently presents deuterium uptake data for a small subset of peptides (three in Fig. 3, two in Supp. Fig. 5). While these examples are illustrative, this selective presentation makes it difficult to assess the overall quality and robustness of the entire dataset. No other peptides are presented that show no differences between protein states being compared. To build confidence in the findings, the data must be presented more comprehensively:\n● To establish the technical quality of the experiment, the manuscript must be expanded to include standard HDX-MS quality metrics. Specifically, please report the following for the final, quality-controlled dataset: The total number of identified peptides, protein sequence coverage (%), average peptide redundancy. These metrics are readily available in HDExaminer and are essential for a complete report.\n● The authors state that significance was defined as a deuterium uptake difference of >0.2 Da across at least two peptides or time points. This is a very small difference. To validate this, please clarify:\n○ How many peptides in total from the experiment met this significance criterion?\n○ Are the identified regions of differential uptake supported by multiple, overlapping peptides? This is a key part of validating HDX-MS findings.\n● To provide the necessary evidence for the claims made, a comprehensive view of the dataset is required. Two standard approaches would be acceptable:\n○ Include a comprehensive data table that displays the D-uptake values for all identified peptides, following established reporting guidelines (e.g., as shown in Masson et al., 2019: https://pubmed.ncbi.nlm.nih.gov/31249422/).\n○ A heatmap with differential D-uptake between states would prove useful. Alternatively, provide the individual deuterium uptake plots for all peptides identified as part of the Supplementary Information.\nAddressing the points above by providing a complete and transparent view of the dataset is essential to support findings.\nMinor points:\n● Statistical significance (p-values) is missing from Supplementary Fig. 5. Is this because the differences shown are not statistically significant?\n● Please include the amount of protein (in pmol) that was injected on-column for the analysis.\n● Please clarify the labeling temperature, as this is not clear. Was the deuterium labeling reaction performed on ice or at another controlled temperature?\n● A final but important point on technical accuracy: The manuscript repeatedly describes HDX-MS as a measure of 'solvent accessibility' (e.g., line 142). This phrasing should be revised throughout the text. HDX rates are primarily governed by the stability of backbone hydrogen bonds and local protein dynamics, not simply by solvent exposure. A more accurate description of the goal would be 'to map the interaction interface' or 'to probe changes in protein conformation'. Please ensure this correction is applied consistently for accuracy.", "reviewer_3": "I co-reviewed this manuscript with one of the reviewers who provided the listed reports. This is part of the Nature Communications initiative to facilitate training in peer review and to provide appropriate recognition for Early Career Researchers who co-review manuscripts."}, "author_response": "Point-by-point responses to the reviewers\nReviewer #1 (Remarks to the Author):\nIn this manuscript, Pakharukova et al showed that b-arr1 interacts with the Src SH3 domain\nvia two distinct sites (N- and CC-) and activates the auto-inhibited Src, thereby plays as an\nactive signal transducer. Authors have identified two distinct sites of the Src SH3 domain\nthat bind to b-arr1, determined the complex structures using cryoEM, and performed\nvarious biochemical and mutational analysis to support the activation mechanism of Src by\nb-arr1. They properly addressed concerns raised by previous reviewers and the revised\nmanuscript provides insights into understanding the molecular mechanism by which Src\nactivated by b-arr1.\nWe thank the reviewer for the thorough review of our manuscript and the valuable suggestions.\nAlthough most of the concerns by the previous reviewers have been addressed, I am bit\nconfused on the mutational analysis that performed to understand the physiological meaning\nof the current study (Fig 7 and lines 275 ~ 288). Unlike the authors described in the discussion\nsession “… this configuration enables βarr1 to ensure robust recruitment and activation of\nSrc when one of the interaction sites is sterically hindered or occupied by a competing\nbinding partner…”., I noticed that mutation of either site exhibited similar to that of the b-\narr1 lacking effect in Fig 7.\nWe thank the reviewer for the thoughtful analysis of the mutational studies. As correctly noted,\nmutations in either the N-site of βarr1 or selected residues in the CC-site abolished βarr-mediated\nSrc activation in the in vitro kinase assay (Fig. 7c), consistent with cooperative rather than\nredundant roles of the two sites. In HEK293 cells (Fig. 7a,b), particularly downstream of the\nβ2V2R receptor, some mutants nevertheless retained ability to activate Src despite impairment of\none site. These findings suggest that the contribution of each site, and the degree of cooperativity\nbetween them, is context-dependent, with additional scaffolding interactions and membrane\nconfinement likely providing partial functional compensation.\nTo clarify that, we have added the following sentences to the Results section (also in response to\nQ1 and Q2):\n“Upon stimulation of the chimeric β2-adrenergic receptor containing the V2R tail (β2V2R) with\nBI-167107, βarr1 mutants exhibited a moderate reduction in βarr-dependent Src activation (Fig.\n7a), indicating partial redundancy between the two Src-binding sites in this context.”\n“Interestingly, in the in vitro assay, mutation of either the βarr1-N site or selected CC-site residues\ncompletely abolished Src activation despite the presence of the other intact site. This finding\nindicates that in the minimal in vitro reconstitution system the two sites act cooperatively rather\nthan independently: loss of one site compromises the ability of the other to promote activation. In\ncells, however, single-site mutants (all mutants downstream of β2V2R; P121E_P124G\ndownstream of D1R) still retained partial ability to activate Src (Fig. 7a, b), suggesting that the\nrelative contribution of each site and the extent of cooperativity between them are context-\ndependent, with additional scaffolding interactions and membrane confinement likely providing\npartial functional compensation.”\nWe have also revised the Discussion as following:\n“Although Src contains a single SH3 domain and can engage only one βarr1 site at a time, the\npresence of two spatially proximal SH3-binding motifs within βarr1 is likely to enhance Src\nrecruitment through avidity-like effects. Rather than functioning as redundant, independent\ndocking sites, the two weak-affinity sites increase the effective local concentration of SH3 ligands\nand promote rapid rebinding following dissociation, thereby stabilizing what would otherwise be\ntransient interactions. This model explains why mutation of either site abolishes Src activation in\nthe minimal in vitro system, where cooperative rebinding is essential, while single-site mutants\nretain partial activity in cellular contexts, where additional scaffolding interactions and membrane\nconfinement can partially compensate. Thus, the dual-site architecture of βarr1 appears optimized\nto enhance the kinetics and stability of Src engagement rather than to provide simple redundancy.”\nQ1. Fig 7a, mutation of both N- and CC sites (mut 3) exhibited higher normalized density\nfor pSrc compared to mut 1 or mut 2. Perhaps, interaction sites are broadly distributed, and\nmultiple mutation may not provide decisive effects. Nevertheless, after reading the entire\nmanuscript, the mutation studies are puzzling. Authors may comment on their mutational\nanalysis.\nWe thank the reviewer for this comment. We agree with the reviewer’s suggestion that, in the case\nof mutant 3 (P88G_P91G_P121E_P124G), these mutations did not exert a decisive effect. This is\nlikely because residues F75, N122, and I314 are critical for Src activation via the CC-site (as shown\nin the in vitro experiment). Consequently, mutant 3 still retained some ability to activate Src.\nTo clarify that, we have revised the Results section as follows (with the added text underlined):\nWe generated four βarr1 mutants targeting these regions and assessed Src activation in HEK-293\nβarr1/βarr2 double-knockout (CRISPR-Cas9-based, dKO) cells28 by monitoring phosphorylation\nof the activation loop tyrosine Y416 following GPCR stimulation. Upon stimulation of the\nchimeric β2-adrenergic receptor containing the V2R tail (β2V2R) with BI-167107, βarr1 mutants\nexhibited a moderate reduction in βarr-dependent Src activation (Fig. 7a), indicating partial\nredundancy between the two Src-binding sites in this context. Notably, mutant 3, harboring\ncombined mutations in both the βarr1-N and βarr1-CC sites (P88G_P91G_P121E_P124G), did\nnot further decrease Src activation compared with mutant 1 (P88G_P91G) or mutant 2\n(P121E_P124G). This finding suggests that βarr1–Src interactions are broadly distributed, and that\nthe mutations tested, while disruptive, are not sufficient to fully abolish βarr-mediated Src\nactivation.\nQ2. Mutation on one of the interaction sites should exhibit some degree of activation. In Fig\n7c, mutation on N-site (mut 1) or CC-site (mut 5 or 6) exhibited an effect similar to reaction\nin the absence of b-arrestin. Authors should comment on this.\nWe thank the reviewer for this suggestion. These results point to cooperativity between the two\nsites in the minimal in vitro system, whereby loss of one site impairs activation mediated by the\nother.\nTo address this point, we have revised the Results section as follows (with the added text\nunderlined):\nTo dissect the contribution of individual residues, we generated six βarr1 mutants and assessed\ntheir ability to mediate Src activation in vitro by monitoring phosphorylation of a Src peptide\nsubstrate, using V2Rpp and Fab30 to activate βarr1. Whereas βarr1 WT increased Src activity\napproximately two-fold, the βarr1-N mutant (P88G_P91G), βarr1-CC triple/quadruple mutants\n(P121E_P124G_F75A_N122A and P121E_P124G_F75A_I314A), as well as the double-site\nmutant (P88G_P91G_P121E_P124G), failed to activate Src (Fig. 7c). In contrast, CC-site mutants\nP121E_P124G and F80A only modestly reduced Src activation relative to WT. These results\nsupport a model in which residues P88 and P91 mediate activation via the βarr1-N site, while F75,\nN122, and I314 are essential for activation via the βarr1-CC site. Interestingly, in the in vitro assay,\nmutation of either the βarr1-N site or selected CC-site residues completely abolished Src activation\ndespite the presence of the other intact site. This finding indicates that in the minimal in vitro\nreconstitution system the two sites act cooperatively rather than independently: loss of one site\ncompromises the ability of the other to promote activation. In cells, however, single-site mutants\n(all mutants downstream of β2V2R; P121E_P124G downstream of D1R) still retained partial\nability to activate Src (Fig. 7a, b), suggesting that the relative contribution of each site and the\nextent of cooperativity between them are context-dependent, with additional scaffolding\ninteractions and membrane confinement likely providing partial functional compensation.\nWe have also revised the Discussion as follows:\n“Rather than functioning as redundant, independent docking sites, the two weak-affinity sites\nincrease the effective local concentration of SH3 ligands and promote rapid rebinding following\ndissociation, thereby stabilizing what would otherwise be transient interactions. This model\nexplains why mutation of either site abolishes Src activation in the minimal in vitro system, where\ncooperative rebinding is essential, while single-site mutants retain partial activity in cellular\ncontexts, where additional scaffolding interactions and membrane confinement can partially\ncompensate. Thus, the dual-site architecture of βarr1 appears optimized to enhance the kinetics\nand stability of Src engagement rather than to provide simple redundancy.”\nRelated to above Q. Lines 285~290: “ Whereas β-arr1 WT (activated by V2Rpp and Fab30) increases Src activity two-fold, βarr1-N mutant P88G_P91G, .. as well as\nP88G_P91G_P121E_P124G mutant failed to activate Src (Fig. 7c). βarr-CC mutants\nP121E_P124G and F80A only slightly reduced Src activation compared to βarr1 WT.\nTherefore, this supports a mechanism where P88 and P91 drive Src activation via the βarr1-\nN site and F75, N122 and I314 are critical for Src activation via the βarr1-CC site.”\nWhy N-mutant failed to increase the Src activation despite it has functional CC-site?\nWe believe this reflects the cooperative nature of the two binding sites, particularly in the minimal\nin vitro reconstitution system, where disruption of one site impairs the ability of the other to drive\nactivation. We have clarified this point in the revised Results section (see answer to the question\nabove).\nQ3. Line 281-282 “..In contrast, none of the βarr1 mutants showed a response to dopamine\nstimulation, suggesting their reduced ability to activate Src (Fig. 7b)…”\nThe band intensity of each line is difficult to quantify by eyes. But mutant 2 (cc-mutant)\nshowed similar increase to that of the WT (and showed some response). Authors may\nconsider to revise this part for clarity.\nWe thank the reviewer for pointing that out.\nWe have clarified the Results section as follows:\nWhile basal levels of phospho-Src were detected in all non-stimulated cells, dopamine stimulation\nproduced a significant increase in phospho-Src in cells transfected with βarr1 WT. In contrast,\nmost βarr1 mutants failed to respond to dopamine, indicating a reduced ability to activate Src,\nexcept for the βarr1-CC mutant P121E_P124G, which exhibited modest activation following\nstimulation (Fig. 7b). Taken together, both the βarr1-CC and βarr1-N sites contribute to Src\nactivation in HEK293 cells downstream of two receptors, β2V2R and D1R.\nReviewer #2 (Remarks to the Author):\nPlease find my technical review of the manuscript by Pakharukova et al. below. The current\npresentation and analysis of the CThe suggestions below focus on areas relevant to HDX-MS\nexperiments that should be clarified and expanded to enhance the paper's impact. For this\nreason, I cannot recommend publication in its present form. However, the concerns are\naddressable. I would be willing to re-evaluate a revised manuscript that thoroughly\naddresses the major points listed below.\nMajor points:\nThe central issue with the manuscript is the lack of a comprehensive and transparent\npresentation of the HDX-MS data. Without this, the technical quality of the experiment\nand the statistical validity of the conclusions cannot be verified.\nWe thank the reviewer for thorough assessment of the HDX data and for the helpful suggestions\nthat enhance the impact of the manuscript. We have addressed the reviewer’s concerns as detailed\nbelow. According to reviewer’s recommendation, we also provide Supporting Data Files 1–6: a\ncomprehensive view of the dataset with the D-uptake values for all identified peptides and the\nindividual uptake plots for all datasets, following established reporting guidelines (Masson et al.,\n2019).\nThe manuscript currently presents deuterium uptake data for a small subset of peptides\n(three in Fig. 3, two in Supp. Fig. 5). While these examples are illustrative, this selective\npresentation makes it difficult to assess the overall quality and robustness of the entire\ndataset. No other peptides are presented that show no differences between protein states\nbeing compared.\nWe thank the reviewer for this comment. We have now included the examples of the peptides that\nshow no differences between the protein states in Fig. 3c and Supplementary Fig. 5c).\nTo build confidence in the findings, the data must be presented more comprehensively:\n● To establish the technical quality of the experiment, the manuscript must be expanded to\ninclude standard HDX-MS quality metrics. Specifically, please report the following for the\nfinal, quality-controlled dataset: The total number of identified peptides, protein sequence\ncoverage (%), average peptide redundancy. These metrics are readily available in\nHDExaminer and are essential for a complete report.\nWe have now included HDX-MS quality metrics and the experiment details in Supplementary\nTable 2.\n● The authors state that significance was defined as a deuterium uptake difference of >0.2\nDa across at least two peptides or time points. This is a very small difference. To validate\nthis, please clarify:\n○ How many peptides in total from the experiment met this significance criterion?\n○ Are the identified regions of differential uptake supported by multiple, overlapping\npeptides? This is a key part of validating HDX-MS findings.\nWe thank the reviewer for these insightful comments. To improve our confidence in the observed\ndeuterium uptake differences, we sought to increase peptide coverage by analyzing a non-\ndeuterated βarr1 sample. This analysis was performed under the previously employed HDX-LC\nconditions using a timsTOF fleX MALDI-2 mass spectrometer with ion mobility engaged. The\nnewly identified peptides were then mapped to the existing HDX dataset. Although peptide overlap\nin the HDX dataset prevented the analysis of all new peptide sequences, this approach ultimately\nincreased the number of analyzable peptides by approximately 30%. Additionally, we conducted\nfurther experiments to measure the fully deuterated exchange profiles of βarr1 to estimate back-\nexchange. To prepare the fully deuterated protein, we first created a 480 µL stock solution of 3 M\ndeuterated GuHCl in D₂O. Subsequently, 20 µL of a 20 µM βarr1 protein stock was added to this\nGuHCl/D₂O solution. The sample was then incubated at 42 °C for three days to allow for deuterium\nexchange. The GuHCl was deuterated beforehand by dissolving it in D₂O and lyophilizing the\nsolution to dryness. The peptides exhibited back exchange values ranging from 10% to 53%, with\nan average of 31%.\n1) Number of peptides meeting the significance criterion: In the comparison between free βarr1\nand βarr1 in complex with SH3, 5% of the analyzed peptides exhibited significantly decreased\ndeuterium uptake in the presence of SH3, as defined by our established significance criterion (>0.2\nDa across at least two overlapping peptides or time points). In the comparison between activated\nβarr1–V2rpp and βarr1–V2rpp in the presence of SH3, 5% of peptides showed decreased uptake\nand 0.5% (1 peptide) showed increased uptake meeting the same criterion. These peptides,\ntogether with their average ΔD values and the cumulative ΔD over time, are listed in\nSupplementary Table 3. Although the differences in deuterium uptake for peptides corresponding\nto the βarr1-CC and βarr1-N sites were relatively small (0.2–0.6 Da), likely reflecting the low\naffinity of the interactions, the changes were reproducible across multiple time points and observed\nin several overlapping peptides in both free and V2Rpp-activated βarr1.\n2) Validation through overlapping peptides: The regions exhibiting differential deuterium\nuptake are supported by several overlapping peptides that display consistent patterns of change\nover several time points (Supplementary Table 3). Importantly, these overlapping peptides were\nidentified in both free inactive βarr1 and V2Rpp-activated βarr1 in the presence of SH3, supporting\nthe observed HDX-MS differences. Together, these results corroborate the disulfide trapping data\nand cryo-EM structures.\nWe have modified the section on the HDX experiments as follows (with the added or modified\ntext underlined):\nTo address this, we mapped the interaction interfaces of free βarr1, V2Rpp-activated βarr1, and\nSH3 using hydrogen–deuterium exchange mass spectrometry (HDX-MS) (Fig. 3; Supplementary\nTable 2). HDX-MS provides insights into protein interaction interfaces by measuring deuterium\nuptake in peptides typically 5–15 residues in length. Thus, this approach enables us to verify\nwhether SH3 binds to two distinct regions of βarr1 in vitro without disulfide trapping.\nApproximately 5% of the analyzed peptides displayed a statistically significant decrease in\ndeuterium uptake in the presence of SH3, defined as a change greater than 0.2 Da observed in at\nleast two overlapping peptides or time points (Supplementary Table 3). Consistent with the\nstructures and the pull-down data, βarr1 showed a significant decrease in HDX rate in both the\nβarr1-CC and βarr1-N sites in the presence of SH3 (Fig. 3a, b; Supplementary Table 3). Although\nthe differences in deuterium uptake for peptides corresponding to the βarr1-CC and βarr1-N sites\nwere relatively small (0.2–0.6 Da), likely reflecting the low affinity of the interactions, the changes\nwere reproducible across multiple time points and observed in several overlapping peptides in both\nfree and V2Rpp-activated βarr1 (Supplementary Table 3; Fig. 3b). Notably, a decrease in\ndeuterium uptake was also observed for residues 42–47, which do not directly interact with SH3\nin the structure but are located near the βarr1–N site, suggesting that SH3 binding may indirectly\nstabilize this region. Interestingly, reduced deuterium uptake was also detected in the hinge region\nof βarr1 and its C-domain within the C-terminal region (residues 351–388), as well as in residues\n257–300 and 328-344 in V2Rpp-activated βarr1 (Fig. 3a). Since Src does not directly interact with\nthe C-domain of βarr19, the reduced deuterium uptake in this region likely arises from long-range\nallosteric stabilization propagated through SH3 binding.\n● To provide the necessary evidence for the claims made, a comprehensive view of the dataset is\nrequired. Two standard approaches would be acceptable:\n○ Include a comprehensive data table that displays the D-uptake values for all identified peptides,\nfollowing established reporting guidelines (e.g., as shown in Masson et al.,\n2019: https://pubmed.ncbi.nlm.nih.gov/31249422/).\n○ A heatmap with differential D-uptake between states would prove useful. Alternatively, provide\nthe individual deuterium uptake plots for all peptides identified as part of the Supplementary\nInformation.\nAddressing the points above by providing a complete and transparent view of the dataset is\nessential to support findings.\nWe thank the reviewer for this suggestion.\nWe now include additional Supporting Data Files to provide the comprehensive view of the\ndataset:\n1) Tables with the D-uptake values for all identified peptides for each dataset (Supporting\nData Files 1–4), following established reporting guidelines as in Masson et al., 2019.\n2) The individual deuterium uptake plots for all peptides for each dataset (Supporting Data\nFiles 5-6).\nThese files are provided for both βarr1 and SH3 datasets.\nMinor points:\n● Statistical significance (p-values) is missing from Supplementary Fig. 5. Is this because the\ndifferences shown are not statistically significant?\nWe thank the reviewer for this insightful comment. Statistical significance (p-values) was not\nreported for Supplementary Fig. 5 because only a single HDX-MS experiment was performed for\nthe SH3 states; therefore, statistical testing was not applicable. Nevertheless, the observed\ndifferences are reproducible across multiple overlapping peptides and reach magnitudes of up to\n1.3 Da at individual time points, supporting the biological relevance of these changes\n(Supplementary Table 4). Accordingly, we present these data as qualitative yet informative\nobservations for readers. To enhance transparency, comprehensive experimental details have been\nincluded in Supplementary Table 2, and the legend for Supplementary Fig. 5 has been updated to\nclarify that the data are based on a single experiment. Furthermore, Supplementary Fig. 5 has been\nrevised to include peptides showing no consistent changes in deuterium uptake.\nWe added the following text to the manuscript:\nWe next performed a qualitative HDX-MS experiment to assess changes in the deuterium uptake\nof SH3 in the presence of either free or V2Rpp-activated βarr1 (Supplementary Tables 2 and 4;\nSupplementary Fig. 5). Consistent with the structural data, peptides encompassing the nSrc loop\nand the 3 helix of SH3 exhibited decreased deuterium uptake, with more pronounced differences\n10\nobserved in the presence of V2Rpp-activated βarr1 (Supplementary Table 4; Supplementary Fig.\n5a, b). In contrast, peptides corresponding to the RT loop predominantly showed increased\ndeuterium uptake, likely reflecting local destabilization and enhanced conformational flexibility\nof the RT loop upon βarr1 binding. Taken together, the HDX-MS data corroborate the disulfide\ntrapping and cryo-EM results, confirming the presence of two SH3-binding sites on βarr1 and\nvalidating the SH3–βarr1-CC and SH3–βarr1-N complexes as bona fide interaction structures.\n● Please include the amount of protein (in pmol) that was injected on-column for the analysis.\nUp to 30, 150, and 150 pmol of βarr1, SH3, and V2Rpp, respectively, were used in each experiment\nto analyze βarr1 dynamics and interactions. To assess SH3 dynamics and interactions, up to 30,\n150, and 150 pmol of SH3, βarr1, and V2Rpp, respectively, were used. An excess of proteins was\nincluded to ensure saturation of complex formation. This information is now included in Methods\nsection.\n● Please clarify the labeling temperature, as this is not clear. Was the deuterium labeling reaction\nperformed on ice or at another controlled temperature?\nThe deuterium labeling reaction was performed at 15°C (now included in the Methods section and\nSupplementary Table 2).\n● A final but important point on technical accuracy: The manuscript repeatedly describes HDX-\nMS as a measure of 'solvent accessibility' (e.g., line 142). This phrasing should be revised\nthroughout the text. HDX rates are primarily governed by the stability of backbone hydrogen bonds\nand local protein dynamics, not simply by solvent exposure. A more accurate description of the\ngoal would be 'to map the interaction interface' or 'to probe changes in protein conformation'.\nPlease ensure this correction is applied consistently for accuracy.\nWe thank the reviewer for this important point. We have now corrected the phrasing throughout\nthe manuscript.\nReviewer #3 (Remarks to the Author):\nI co-reviewed this manuscript with one of the reviewers who provided the listed reports. This is\npart of the Nature Communications initiative to facilitate training in peer review and to provide\nappropriate recognition for Early Career Researchers who co-review manuscripts.\nWe thank the reviewer for the assessment of the manuscript and valuable suggestions.\nPoint-by-point responses to the reviewers\nReviewer #2 (Remarks to the Author):\nI thank the authors for their detailed rebuttal and for providing the comprehensive datasets,\nincluding the D-uptake values and peptide plots. The inclusion of these data significantly\nimproves the transparency of the study.\nAfter reviewing the full dataset and experimental parameters, the authors need to further\nrefine the interpretation of the D-uptake differences. Specifically, the experimental\nconditions (high dilution) appear to have limited the target occupancy, resulting in small\nshifts that are close to the noise (see comments below). While the HDX-MS data provides\nvaluable validation, the current interpretation overstates the statistical certainty of the\nresults given these experimental limitations.\nI do not object to the publication of this work; however, I recommend adjusting the text to\nframe these results as \"consistent qualitative trends\" that corroborate the CryoEM model,\nrather than a quantitative definition of the interface. This shift in presentation, along with\nthe data reporting clarifications detailed below, will strengthen the manuscript.\nWe thank the Reviewer for the careful evaluation of the HDX datasets. In response to this\nsuggestion, we have adjusted the text in the manuscript to present the HDX results as consistent\nqualitative trends that corroborate the cryo-EM structures:\n“Taken together, the HDX-MS data reveal consistent qualitative trends that corroborate the\ndisulfide trapping and cryo-EM results and support the presence of two SH3-binding regions on\nβarr1 and the interaction modes captured in the SH3–βarr1-CC and SH3–βarr1-N complexes.”\nMajor technical concerns\n(cid:0) Target occupancy during HDX: The authors state in the Methods that SH3 was added in\n5-fold excess to ensure saturation. However, the dilution step into D2O significantly alters\nthe equilibrium. Based on the stated Kd of 6 uM and the stock concentrations of 20 uM and\n100 uM, the initial occupancy during equilibration is high (~93%). However, the protocol\ndescribes a 17-fold dilution with D2O. Under these final conditions, the calculated target\noccupancy of βarr1 (when saturated with SH3) or SH3 (when saturated with βarr1) drops to\n~46%. This low occupancy likely explains why the observed differences are so small and why\nmany peptides show no change. A smaller dilution (e.g., 5-fold) would maintain higher\noccupancy (~70%). Because the occupancy is low, the authors must acknowledge this\nlimitation in their text. The small magnitude of changes is likely an artifact of the\nexperimental design rather than interaction dynamics.\nWe thank the Reviewer for highlighting this important point. We agree that, for such weak\ninteractions, it is desirable to maximize both complex occupancy and the D₂O fraction during\nexchange, while minimizing back-exchange in solution and in the gas phase. In this study, we\nprioritized maintaining 94% D₂O fraction at 17-fold dilution to preserve optimal dynamic range\nand improve the reproducibility of HDX measurements. Although this results in a lower complex\noccupancy (~46%), the improved fidelity of deuterium uptake data provides greater confidence in\nidentifying bona fide interaction sites, particularly given the small ΔHDX values expected for\nweak complexes. We therefore assumed that the potential gain in occupancy at a 5-fold dilution\nwould be outweighed by the reduced HDX data quality associated with lowering the D₂O fraction\nto ~80%.\nTo clarify this for readers, we have included the following statement in the Methods sections\n(added text shaded in yellow):\n“The protein stock solution was diluted 17-fold into D O-based buffer (20 mM HEPES pH 7.5,\n2\n100 mM NaCl) to achieve a final D₂O fraction of 94% and to maintain an optimal dynamic range\nof deuterium uptake, thereby improving the reproducibility of HDX-MS measurements.”\nAnd in the Results section:\n“Although the differences in deuterium uptake for peptides corresponding to the βarr1-CC and\nβarr1-N sites were modest (0.2–0.8 Da), consistent with low-affinity interactions and partial\ncomplex occupancy arising from dilution during HDX labeling, the changes were consistent across\nmultiple time points and supported by several overlapping peptides in both free and V2Rpp-\nactivated βarr1 (Supplementary Table 3; Fig. 3a, b; Supplementary Fig. 5a).”\n(cid:0) Overlapping peptides: There are inconsistencies in the data where longer peptides show\n\"significant\" differences (highlighted in yellow in table below), but shorter, internal peptides\ncovering the same region do not. For example, Supplementary Table 3 highlights the peptide\n76–109 as significantly different. However, multiple shorter peptides nested within this\nsequence (covering the interacting region FRKDLF, residues 80–88) show no difference.\nTheoretically, the shorter peptides (which are not \"diluted\" by non-interacting residues)\nshould show larger % D-uptake differences than the longer parent peptide. The fact that\nthey do not, suggests the difference in the longer peptide may be a technical artifact (e.g.,\nintegration boundary differences) rather than a biological signal. The authors should re-\nexamine these inconsistencies. If the \"significant\" hit is not supported by internal\noverlapping peptides, it should be treated with extreme caution.\nWe thank the Reviewer for this careful analysis. The apparent discrepancy in deuterium uptake\ndifferences between longer and shorter peptides can be explained by several technical and\nstructural factors.\nFirst, the first two N-terminal residues do not contribute to measurable deuterium uptake.\nStructural analysis indicates that interaction of βarr1 with SH3 in the CC-site relies on residues\nF75 (hydrophobic interactions) and R76 (two hydrogen bonds). These residues are located at or\nnear the peptide N-terminus in several short internal peptides, such as TFRKDLFVA, FRKDL,\nFRKDLF, and RKDLF, inherently limiting detectable changes.\nSecond, the longer peptide RKDLFVANVQSFPPAPEDK spans residues that contribute to both\nthe βarr1-CC site (RKDLF) and the -N site (FPPAP), effectively integrating interactions from two\nbinding regions. As a result, deuterium uptake differences are more readily detected in this peptide\nthan in shorter fragments that sample only a subset of the interacting surface.\nThird, chromatographic behavior further contributes to these differences. Most short peptides (e.g.\nGLTFRKDLF, FRKDLF, DVLGLTFRKDL, DVLGLTFRKDLF) elute substantially later (∼9–\n13 min) than the longer peptides RKDLFVANVQSFPPAPEDK and LTFRKDLFVANVQ\nexhibiting significant reduction in HDX uptake (∼3 min). Therefore, these shorter peptides are\nmore susceptible to back exchange which further attenuates small differences in deuterium uptake.\nTo address this issue systematically, we reanalyzed a curated set of peptides encompassing the\nβarr1-N and -CC interaction sites (residues 66–126) and performed a focused volcano plot analysis\n(Supplementary Fig. 5a; see below). The significance threshold for this group is 0.2112 Da. This\nanalysis reveals a consistent trend toward decreased deuterium uptake in the presence of SH3\nacross most peptides, with smaller differences for shorter peptides due to the technical\nconsiderations described above. We have now included this analysis as Supplementary Figure 5a\nand added the following sentence to the Results section:\n“Within these regions, longer peptides spanning both βarr1-CC and -N sites showed more\npronounced changes, while shorter internal peptides displayed attenuated effects due to key\ninteracting residue being positioned at the peptide N-terminus and increased back exchange\n(Supplementary Figure 5a).”\nTaken together, these observations indicate that the differences observed in the longer peptides\nreflect peptide length, residue positioning, and chromatographic behavior rather than technical\nartifacts. Importantly, as suggested by the Reviewer, the HDX data are now presented as\nqualitative, supportive trends that are fully consistent with the cryo-EM structures:\n“Taken together, the HDX-MS data reveal consistent qualitative trends that corroborate the\ndisulfide trapping and cryo-EM results and support the presence of two SH3-binding regions on\nβarr1 and the interaction modes captured in the SH3–βarr1-CC and SH3–βarr1-N complexes.”\n(cid:0) The threshold of 0.2 Da is set at the absolute limit of detection (approximately 2x\nexperimental noise), which increases the risk of false positives. HDExaminer offers multiple\nrepresentations to evaluate a dataset, such as Residual plot or Wood’s plot, where statistical\nsignificance can be automatically calculated based on replicate variance (especially when 3\nreplicates were performed). What is the automatic cutoff threshold on the Residual plot or\nWood’s plot for the βarr1 vs βarr1:SH3 dataset?\nWe thank the Reviewer for this helpful suggestion. For the curated set of peptides encompassing\nthe βarr1-N and βarr1-CC interaction sites (residues 66–126), the significance threshold in the\nvolcano plot is 0.21 Da, while the corresponding residual plot cutoff is 0.63 Da (calculated as 0.21\nDa multiplied by three data points). For the remainder of the dataset, slightly higher automatic\nthresholds were obtained, with cutoffs of 0.30 Da for datasets with free βarr1 and 0.36 Da for\ndatasets with V2Rpp-activated βarr1. Given the Reviewer’s concern, we applied a more stringent\ncutoff across the dataset and report only statistically significant differences exceeding 0.30 Da for\nβarr1 and 0.36 Da for βarr1–V2rpp at a single time point. Figures 3a and Supplementary Tables 3\nand 4 have been revised accordingly.\nThe following sentences were revised in the Results section:\n“Approximately 5% of the analyzed peptides displayed a statistically significant decrease in\ndeuterium uptake in the presence of SH3, defined as a statistically significant change greater than\n0.3 Da for free βarr1 and 0.36 Da for βarr1–V2rpp (Supplementary Table 2–3, Supplementary\nData 1–2, 5)”.\nand in the Methods section:\n“Only regions that exhibited statistically significant differences in deuterium uptake greater than\n0.30 Da for free βarr1 and 0.36 Da for βarr1–V2rpp were considered relevant”.\n(cid:0) Furthermore, given these small magnitudes and potential variance differences between\nstates, a standard Student’s t-test is prone to false positives. The field standard is the\nWelch’s t-test (Hageman & Weis, 2019, Anal. Chem.), which accounts for unequal variances and is\navailable in HDExaminer. Please verify if the peptides in Figure 3 (residues 76-94, 86-101,\n106-126) remain statistically significant using a Welch’s t-test (e.g. they should show as\nsignificant in the volcano plot or represented with a star in the D-uptake plot when compared\nto the control state). If these peptides do not pass the Welch’s test, or given the inconsistencies\nnoted in Point 2, I strongly recommend presenting the HDX-MS data as qualitative trends.\nThis allows the data to successfully serve its purpose (validation of CryoEM) without\noverstating the quantitative evidence.\nWe re-evaluated the statistical significance of peptides in Figure 3b using Welch’s t-test and found\nthat the changes in the presented peptides are statistically significant; they also show as significant\nin the volcano plot (see above). The exact significance values are now shown in Figure 3b.\nImportantly, as suggested by the Reviewer, we frame the HDX data as consistent qualitative trends\nsupporting the cryo-EM structures:\n“Taken together, the HDX-MS data reveal consistent qualitative trends that corroborate the\ndisulfide trapping and cryo-EM results and support the presence of two SH3-binding regions on\nβarr1 and the interaction modes captured in the SH3–βarr1-CC and SH3–βarr1-N complexes.”\nMinor points and data presentation\n(cid:0) The current tables are incomplete. The authors should re-export all data using the “Export\nUptake Summary Table” option in HDExaminer. To ensure the data is reviewable, the table\nmust explicitly contain the following fields: Protein State, Protein, Start, End, Sequence,\nPeptide Mass, RT, Deut Time, maxD, #D, %D, Conf Interval (#D), #Rep, Confidence, Stddev,\nand p-value.\nWe thank the Reviewer for this suggestion. We have now re-exported all data from HDExaminer\nto include all fields used in the analysis.\n(cid:0) It is known that different charge states can sometimes result in different D-uptake values\nfor the same peptide. It appears that for many peptides in this dataset, multiple charge states\nhave been detected (based on the exported plots). The authors must ensure that for direct\ncomparisons between protein states, only similar charge states are taken into account to\navoid technical bias.\nWe thank the Reviewer for the careful analysis of the data. Although multiple charge states were\ndetected, only identical charge states were used for direct comparisons between protein states.\n(cid:0) The data exports currently include short peptides that, due to sequence similarity, cannot\nbe distinguished based on MS/MS data. While HDExaminer warns of these peptides with\nsimilar masses, it does not automatically exclude them. The authors must manually curate\nthe dataset to identify such cases and include only one peptide representation. A couple of\nexamples found in this dataset include:\nβarr1: Residues 42-47 (VDPVG) is indistinguishable from 43-48 (DPVGV)\nSH3: Residues 27-30 (VTTF) is indistinguishable from 28-31 (TTFV)\nWe thank the Reviewer for this comment. We have curated the datasets and included only one\npeptide representation with the same mass.\n(cid:0) The D-uptake plots provided in the supplementary files display Standard Deviation (SD)\nbut lack confidence intervals. Please re-export these plots to include confidence intervals to\nallow for a proper visual assessment of significance.\nWe now present the D-uptake plots with confidence intervals.\n(cid:0) I strongly encourage the authors to upload their RAW HDX-MS (including identification\nruns) data to a public repository (e.g., PRIDE or MassIVE). Access to the RAW data is\ncritical for reviewers to evaluate the manuscript efficiently and fairly and allow for future\nmining.\nWe thank the Reviewer for this suggestion. The HDX-MS data have now been deposited to the\nProteomeXchange Consortium via the MassIVE repository with the dataset identifier PXD073493\n(MSV000100571).\nI thank the authors for their detailed rebuttal and for providing the comprehensive datasets,\nincluding the D-uptake values and peptide plots. The inclusion of these data significantly\nimproves the transparency of the study.\nAfter reviewing the full dataset and experimental parameters, the authors need to further refine\nthe interpretation of the D-uptake differences. Specifically, the experimental conditions (high\ndilution) appear to have limited the target occupancy, resulting in small shifts that are close to\nthe noise (see comments below). While the HDX-MS data provides valuable validation, the\ncurrent interpretation overstates the statistical certainty of the results given these experimental\nlimitations.\nI do not object to the publication of this work; however, I recommend adjusting the text to frame\nthese results as \"consistent qualitative trends\" that corroborate the CryoEM model, rather than a\nquantitative definition of the interface. This shift in presentation, along with the data reporting\nclarifications detailed below, will strengthen the manuscript.\nMajor technical concerns\n Target occupancy during HDX: The authors state in the Methods that SH3 was added in\n5-fold excess to ensure saturation. However, the dilution step into D2O significantly\nalters the equilibrium. Based on the stated Kd of 6 uM and the stock concentrations of\n20 uM and 100 uM, the initial occupancy during equilibration is high (~93%). However,\nthe protocol describes a 17-fold dilution with D2O. Under these final conditions, the\ncalculated target occupancy of βarr1 (when saturated with SH3) or SH3 (when saturated\nwith βarr1) drops to ~46%. This low occupancy likely explains why the observed\ndifferences are so small and why many peptides show no change. A smaller dilution\n(e.g., 5-fold) would maintain higher occupancy (~70%). Because the occupancy is low,\nthe authors must acknowledge this limitation in their text. The small magnitude of\nchanges is likely an artifact of the experimental design rather than interaction dynamics.\n Overlapping peptides: There are inconsistencies in the data where longer peptides show\n\"significant\" differences (highlighed in yellow in table below), but shorter, internal\npeptides covering the same region do not. For example, Supplementary Table 3\nhighlights the peptide 76–109 as significantly different. However, multiple shorter\npeptides nested within this sequence (covering the interacting region FRKDLF, residues\n80–88) show no difference. Theoretically, the shorter peptides (which are not \"diluted\" by\nnon-interacting residues) should show larger % D-uptake differences than the longer\nparent peptide. The fact that they do not, suggests the difference in the longer peptide\nmay be a technical artifact (e.g., integration boundary differences) rather than a\nbiological signal. The authors should re-examine these inconsistencies. If the\n\"significant\" hit is not supported by internal overlapping peptides, it should be treated\nwith extreme caution.\n The threshold of 0.2 Da is set at the absolute limit of detection (approximately 2x\nexperimental noise), which increases the risk of false positives. HDExaminer offers\nmultiple representations to evaluate a dataset, such as Residual plot or Wood’s plot,\nwhere statistical significance can be automatically calculated based on replicate\nvariance (especially when 3 replicates were performed). What is the automatic cutoff\nthreshold on the Residual plot or Wood’s plot for the βarr1 vs βarr1:SH3 dataset?\n Furthermore, given these small magnitudes and potential variance differences between\nstates, a standard Student’s t-test is prone to false positives. The field standard is the\nWelch’s t-test (Hageman & Weis, 2019, Anal. Chem.), which accounts for unequal variances and is\navailable in HDExaminer. Please verify if the peptides in Figure 3\n(residues 76-94, 86-101, 106-126) remain statistically significant using a Welch’s t-test\n(e.g. they should show as significant in the volcano plot or represented with a star in the\nD-uptake plot when compared to the control state). If these peptides do not pass the\nWelch’s test, or given the inconsistencies noted in Point 2, I strongly recommend\npresenting the HDX-MS data as qualitative trends. This allows the data to successfully\nserve its purpose (validation of CryoEM) without overstating the quantitative evidence.\nMinor points and data presentation\n The current tables are incomplete. The authors should re-export all data using the\n“Export Uptake Summary Table” option in HDExaminer. To ensure the data is\nreviewable, the table must explicitly contain the following fields: Protein State, Protein,\nStart, End, Sequence, Peptide Mass, RT, Deut Time, maxD, #D, %D, Conf Interval (#D),\n#Rep, Confidence, Stddev, and p-value.\n It is known that different charge states can sometimes result in different D-uptake values\nfor the same peptide. It appears that for many peptides in this dataset, multiple charge\nstates have been detected (based on the exported plots). The authors must ensure that\nfor direct comparisons between protein states, only similar charge states are taken into\naccount to avoid technical bias.\n The data exports currently include short peptides that, due to sequence similarity, cannot\nbe distinguished based on MS/MS data. While HDExaminer warns of these peptides\nwith similar masses, it does not automatically exclude them. The authors must manually\ncurate the dataset to identify such cases and include only one peptide representation. A\ncouple of examples found in this dataset include:\nβarr1: Residues 42-47 (VDPVG) is indistinguishable from 43-48 (DPVGV)\nSH3: Residues 27-30 (VTTF) is indistinguishable from 28-31 (TTFV)\n The D-uptake plots provided in the supplementary files display Standard Deviation (SD)\nbut lack confidence intervals. Please re-export these plots to include confidence intervals\nto allow for a proper visual assessment of significance.\n I strongly encourage the authors to upload their RAW HDX-MS (including identification\nruns) data to a public repository (e.g., PRIDE or MassIVE). Access to the RAW data is\ncritical for reviewers to evaluate the manuscript efficiently and fairly and allow for future\nmining."}]

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