Positive cooperativity between RAS-binding and cysteine-rich domains regulates RAF membrane binding kinetics via lateral rebinding

Domain: neuroscience
Published: 2026-02-10
Editorial outcome: minor_revision

Abstract
RAF activation requires coordinated interactions with both RAS and membrane lipids, yet the molecular basis of this process remains unclear. Using a bottom-up reconstitution approach, we show how coordinated protein–protein and protein–lipid interactions regulate membrane binding dynamics of RAF to drive its multistep activation. Within membrane environments, the RAS-binding domain (RBD) and cysteine-rich domain (CRD) exhibit cooperativity, with CRD-mediated phosphatidylserine binding stabilizing the RBD:RAS complex. Importantly, RAF remains membrane-bound through lateral rebinding to RAS, where a weak CRD–lipid interaction plays an essential role. The lateral rebinding extends RAF’s membrane dwell time under high RAS density conditions. This prolonged membrane residence may facilitate completion of RAF’s multistep activation. Given the high abundance of weak multivalent membrane interactions, lateral rebinding may be a common mechanism for regulating the activity of signaling proteins through sustained membrane retention.

Introduction
The RAF kinases are critical regulators of the mitogen-activated protein kinase (MAPK) pathway, controlling diverse cellular processes1. RAF isoforms (A-, B-, and CRAF) share a conserved domain organization: an N-terminal regulatory region composed of the RAS-binding domain (RBD) and the cysteine-rich domain (CRD), and a C-terminal kinase domain (Fig. 1a). In their basal state, RAF kinases reside in the cytosol as autoinhibited monomers, and they are activated when the RBD binds to RAS⋅GTP at the membrane surface. Once activated, RAF initiates the three-tiered RAF-MEK-ERK kinase cascade, serving as a central node that modulates pathway dynamics. Mutations in RAF can lead to constitutive signaling in cancers and developmental disorders2,3. Paradoxical activation has been observed when inhibitors targeting oncogenic BRAFV600E are used under high RAS·GTP conditions, inadvertently promoting RAF dimerization and unintended ERK pathway activation4–11.

RAF activation proceeds through multiple intermediates governed by complex molecular interactions on membrane surfaces1,12–14. However, the precise mechanisms underlying the transition from inactive monomer to active dimer remain to be fully determined. In the active state, RAF associates with the plasma membrane via interaction of its RBD with active RAS⋅GTP15,16. The CRD of RAF further stabilizes its membrane association by engaging both RAS and negatively charged lipids, such as phosphatidylserine (PS)17–19. Thus, anchored at the membrane, RAF becomes dephosphorylated by the MRAS-SHOC2-PP1C phosphatase complex and dimerizes via its kinase domain—an essential step for allosteric activation20–24. The dimeric 14-3-3 protein reinforces an active dimeric assembly by binding a phosphoserine-containing motif on each RAF kinase domain25,26.

Recent cryo-electron microscopy (EM) structures of RAF provide important insight into how RAF activity is regulated27–30. The inactive monomeric CRAF:MEK1:14-3-3 complex can adopt multiple conformations, including a fully autoinhibited confirmation (Fig. 1b) that closely resembles that of BRAF, as well as more open conformations in which the CRD is structurally liberated30. In the fully autoinhibited conformation of both BRAF and CRAF, the 14-3-3 dimer binds two phosphoserine residues within the CR2 and CR3 regions of the same RAF molecule, forming a cradle-like structure that sequesters the CRD27,28. The CRD is a critical component that stabilizes the overall autoinhibited complex by making multiple contacts with the kinase domain and 14-3-3. For BRAF, RBD–RAS binding initiates the disassembly of the autoinhibited state, and CRD–lipid interaction is required for full activity31. Interestingly, in CRAF, the open monomer states expose the kinase dimerization interface, yet they do not spontaneously progress to form active dimers in solution30. These observations highlight the engagement of active RAS at the membrane surface as a critical step in RAF activation.

A key aspect of RAF activation is that RAS nanoclusters serve as primary sites for RAF recruitment and activation by presenting high RAS density and locally enriched lipids that facilitate activation32,33. The RBD and CRD—responsible for binding protein and lipids, respectively—are arranged in tandem and separated by a short, six-amino-acid-long linker. X-ray crystal structures show that the RBD and CRD together form an extended structural unit when bound to RAS34,35. Prior studies reported cooperativity between RBD and CRD in which RAS–RBD binding induces a transient interaction between RAS and CRD, and mapped CRD–lipid interfaces16. However, how these interactions control membrane kinetics has remained unclear. A quantitative understanding of membrane-binding kinetics and the underlying mechanisms is crucial because RAF is exclusively activated on cellular membranes14. RAF is also among the least abundant signaling proteins in many cancer cell types36. Upon receptor activation, only a small number of RAF molecules—estimated to be as few as 100–200 per cell—are recruited to the membrane, where they serve as a stoichiometric bottleneck that determines the overall signaling outcome37. Therefore, understanding RAF membrane dynamics is fundamental for elucidating how normal and oncogenic RAS activities are transduced into ERK signaling38.

Here, we employ a bottom-up reconstitution approach with supported lipid bilayers (SLBs) to systematically investigate how RAS proteins and lipids cooperatively regulate RAF membrane binding dynamics. By quantifying the kinetics, affinities, and diffusion of various CRAF regulatory domain constructs on RAS-functionalized membranes, we demonstrate membrane-specific positive cooperativity in which the RBD and CRD mutually modify their respective binding interactions. Specifically, RBD binding to RAS facilitates engagement of the RAF CRD with negatively charged PS lipids, resulting in two kinetically distinct steps during the binding of protein and lipid partners. This sequential association enables the CRD to bind abundant PS lipids in a RAS·GTP-dependent manner, ensuring that RAF membrane binding is strictly initiated by RAS activation. Moreover, CRD–lipid engagement reinforces RBD–RAS and CRD–RAS association, effectively reducing the membrane dissociation rate. Importantly, we show that RAF maintains its membrane-bound state through lateral rebinding to RAS, with a weak CRD–lipid interaction playing an essential role. This lateral rebinding mechanism significantly extends RAF’s membrane dwell time under conditions of high active RAS density and elevated anionic lipid content. This prolonged membrane residence may facilitate kinetic proofreading of RAF activation by increasing the likelihood of completing the multistep activation sequence. Overall, our study offers a mechanistic insight into how kinetic regulation by coordinated protein and lipid inputs modulate RAF signaling, advancing our understanding of signaling activation and specificity.

Results
RBD binding to RAS promotes lipid engagement of CRD
To investigate the mechanisms underlying RAF membrane binding, we reconstituted CRAF and RAS interaction on a SLB (Fig. 1a, c). A 20% PS composition was used to closely approximate the physiological fraction (~26%) of PS in the inner leaflet of plasma membranes while maintaining high-quality bilayers39. HRAS (1-184, C118S, hereafter referred to as RAS), which retains its native cysteine palmitoylation sites (181 and 184) in the hypervariable region, was linked to lipids via maleimide chemistry. This widely used lipidation method preserves the biological function and structure of RAS and has been applied to all three RAS isoforms (H-, N-, and KRAS) and other membrane-anchored small GTPases40–47. Maleimide-conjugated HRAS is expected to undergo rapid free rotational diffusion on membranes, allowing ready access for effector protein binding, as previously observed in fully processed NRAS and KRAS48,49. RAS was then activated by exchanging nucleotides with GTP using the catalytic domain of Son of Sevenless (SOS) (Supplementary Fig. 1)41. The membrane binding kinetics of CRAF constructs fluorescently labeled with mNeonGreen (mNG) or AZ647 were measured using total internal reflection fluorescence (TIRF) microscopy (Fig. 1c). The surface density of membrane-bound CRAF was determined from TIRF intensity, calibrated by fluorescence correlation spectroscopy (FCS) (Supplementary Fig. 2)50. Data demonstrating protein quality—SEC and SDS–PAGE, and single-molecule intensity analyses—are provided in Supplementary Figs. 4–6. The single-molecule intensity distribution suggests the presence of a minor dimeric fraction of the CRD-mNG construct, with no evidence of larger oligomers, at 150 mM NaCl, while all other proteins were purely monomeric.

RAF is a low-abundance protein, existing in the cytoplasm at a concentration ranging from high picomolar to low nanomolar51. Accordingly, we characterized membrane binding kinetics of truncated CRAF regulatory domains within this physiologically relevant range. At 20 nM, the RBD exhibited low-level, fast equilibrium binding (~30 molecules per µm2), independent of PS content (2% or 20%) (blue, Fig. 1d). Despite the known affinity of the CRD for PS, 20 nM of CRD displayed insignificant binding (approximately a few molecules per µm2) on both 2% and 20% PS membranes functionalized with RAS (gray, Fig. 1d). Notably, the tandem RBD-CRD exhibited a marked increase in membrane binding (~600 molecules per µm2) that depended on PS and had slower kinetics (red, Fig. 1d). This finding is intriguing because the CRD alone shows negligible binding, yet it effectively engages PS lipids when arranged in tandem with RBD. The enhanced lipid sensing exhibited by CRD requires a specific interaction between RAS·GTP and the RBD, as negligible binding was observed on RAS·GDP-functionalized membranes (Supplementary Fig. 7). Collectively, these results demonstrate that the RBD and CRD are functionally coupled, such that RBD binding to RAS promotes the CRD for lipid engagement.

To better understand how RAF integrates protein and lipid inputs for membrane binding, we sought to disentangle the contributions of protein interaction and lipid interaction to the enhanced affinity of RBD-CRD on RAS-functionalized membranes. To quantify the affinity arising solely from protein–protein interactions, the apparent dissociation constants (Kd) of RBD and RBD-CRD for RAS were determined on PEGylated (non-membrane) surfaces. An additional protein contact between RAS and the CRD increases the overall affinity of RBD-CRD by approximately fivefold—220 ± 80 nM for RBD and 40 ± 10 nM for RBD-CRD (Supplementary Fig. 8)—a trend similar to the twofold enhancement previously observed with KRAS35. For membrane-tethered RAS, the RBD exhibited Kd that depended little upon PS content, measuring 270 ± 90 nM and 240 ± 20 nM at 2% and 20% PS, respectively (Fig. 1e and Supplementary Fig. 9a, b). These values are consistent with our membrane-free measurements and previously reported affinities of the CRAF RBD for HRAS35,52. In contrast, RBD-CRD exhibited an affinity for membrane-tethered RAS more than an order of magnitude higher than that of the RBD alone (Kd = 25 ± 10 nM and 6 ± 3 nM at 2% and 20% PS, respectively; Fig. 1f and Supplementary Fig. 9c, d). Removing RAS essentially abolished this high-affinity binding (Fig. 1f). The progressive increase in affinity to RAS—from Kd = 220 nM for RBD (PEG surface), to 40 nM for RBD-CRD (PEG surface), to 6 nM for RBD-CRD (20% PS membrane)—demonstrates that low-nanomolar membrane affinity arises from the cooperative contributions of RAS–RBD binding, the additional RAS–CRD contact, and CRD–PS engagement. The CRD binds RAS with micromolar affinity via an interface distinct from the RAS:RBD contact17. However, this weak interaction between RAS and the isolated CRD does not independently promote lipid engagement, as the isolated CRD showed similarly low binding on both RAS-free and RAS-functionalized membranes (Fig. 1g and Supplementary Fig. 10). Together, these results demonstrate that both the protein (RAS) and lipid (PS) binding modalities of the CRAF CRD depend on the RBD being bound to RAS, particularly under the low nanomolar physiological RAF concentration.

CRD and lipid interaction selectively modulates the membrane dissociation rate
Characterizing the kinetic parameters of membrane binding is important for understanding the mechanisms that govern RAF activation. Given that the RBD and CRD have distinct primary binding partners—RAS and lipids, respectively—we first examined how these interactions interplay to modulate the association (kon) and dissociation (koff) rates. The kon values were quantified by counting the single-molecule recruitment events of AZ647-labeled CRAF to the membrane (Supplementary Fig. 11). In line with our ensemble binding measurements, clearly spatially resolved membrane recruitment events required the interaction with RAS (Fig. 2a). Interestingly, we observed highly transient interactions of RBD-CRD (appearing as faint, diffusive traces) on RAS-free 20% PS membranes. Although those particles could not be localized, manual intensity analysis estimated their dwell times on the order of several milliseconds (Supplementary Fig. 12). Importantly, these short-lived interactions between RBD-CRD and lipids do not result in productive RAS engagement, as kon values for both RBD and RBD-CRD were unaffected by PS concentration (Fig. 2b). This suggests that the initial recruitment is primarily driven by the protein–protein interaction between RBD and RAS, without involving CRD and lipid interactions. The CRD exerts a modest inhibitory effect on RBD–RAS binding, reducing the kon values for RBD-CRD (see statistical test summary in Supplementary Table 1). Because the RBD-CRD exhibits a lower kon than the RBD, its higher binding affinity must stem from a reduced membrane dissociation rate. We quantified the apparent koff on RAS-functionalized 20% PS membranes using an ensemble free-dissociation assay under continuous buffer flow. The RBD exhibited highly dynamic and transient interactions with a half-life of ~1 s, consistent with a previous study using proteins pulled down from cell lysates53. Indeed, mNG-fused RBD-CRD displayed an extended membrane residence time, with a half-life of several tens of seconds (Fig. 2c). Photobleaching had a minimal impact on the estimation of koff (Supplementary Fig. 14). Such a long dwell time cannot be achieved solely from protein–protein interactions. The RBD-CRD dissociates from RAS on a lipid-free PEGylated surface with a half-life of a few seconds (Supplementary Fig. 15a). This kinetic parameter analysis reveals that a crucial role for the CRD–PS interaction in selectively regulating membrane dissociation kinetics without affecting the association rate.

These results suggest that CRAF localizes to membranes through two kinetically distinct steps (Fig. 2d). In the first step, the RBD binds active RAS. In the second step, lipid engagement by the CRD—particularly a productive interaction—occurs only after the RBD has bound to RAS. This two-step mechanism ensures RAS·GTP-specific membrane localization of RAF while leveraging abundant PS lipids to prolong membrane dwell time and achieve high-affinity binding.

CRD persistently engages PS lipids upon RBD and RAS binding
To investigate how the RAS:RAF complex interacts with lipids, we performed diffusion analysis using single-particle tracking. Membranes containing 20% DOPS was functionalized with RAS at a density ranging from 200–900 per µm2 (see detailed values in Supplementary Fig. 17). For single particle tracking of RAS, a small fraction of RAS was labeled with Alexa647-GppNp (a fluorescent, nonhydrolyzable GTP analog), ensuring well-dispersed single molecules (Supplementary Fig. 16). RAS alone exhibited two distinct diffusion states (gray, Fig. 2e and Supplementary Fig. 17a, b). The majority (~90% occupancy, α) had a fast diffusion coefficient of 1.05 µm2/s (D1). The secondary, slower state exhibited a diffusion coefficient of 0.29 µm2/s (D2). This minor slow state may result from microscale heterogeneity of PS membranes or strong coupling with the solid support, a behavior also observed in fluorescent lipids (Supplementary Fig. 18). RBD-AZ647 bound to unlabeled RAS-functionalized membranes mirrored RAS diffusion, indicating minimal lipid interaction by RBD (blue, Fig. 2 and Supplementary Fig. 17c, d). By contrast, the RBD-CRD-AZ647 diffused significantly more slowly due to lipid engagement by CRD (red, Fig. 2e and Supplementary Fig. 17e, f). Two diffusional states were identified at 0.39 µm2/s (85% occupancy) and 0.14 µm2/s. Even the faster state is substantially slower than that of RBD with minimal lipid interaction, suggesting that once recruited by RAS, the RBD-CRD persistently engages PS lipids. Because RAS-RAF complexes were tracked at the single-molecule level and separated by several micrometers, the observed decrease in diffusion coefficient reflects lipid interactions with individual RAS:RBD-CRD complexes, rather than collective behaviors such as clustering. In support of this, the RBD-CRD at saturation density displayed diffusion behavior identical to that at single-molecule density, confirming the absence of clustering in our system (Supplementary Fig. 19). Previous studies have shown that RAS:RAF clustering requires more complex lipid compositions54,55. The profound reduction in diffusion coefficient in our study could be explained by multivalent lipid interactions, facilitated by the membrane insertion of interdigitated hydrophobic and positively charged residues within the CRD loop 1 (residues 143–149) and 2 (residues 157–164)2,55.

Engagement of the CRD with lipids appears to be a critical step involved in the structural rearrangement of the autoinhibited RAF complex. To evaluate the kinetics and efficiency of CRD–lipid interaction following RBD binding, we compared the step-size distributions of RBD-CRD at the initial moment of contact (first-step sizes) versus the entire observation period (all-step sizes). The majority of RBD-CRD rapidly engage lipids within the 20 ms time resolution of the experiment, as evidenced by the similarity between these two distributions (Supplementary Fig. 20). This rapid lipid engagement (forward step 2 in Fig. 2d) effectively renders the two-step association mechanism unidirectional, with the first step serving as the rate-determining step. However, in the full-length protein, particularly in BRAF, CRD–lipid binding may be significantly slower, because the lipid-interacting residues of the CRD are occluded by 14-3-3 dimers. The rate of this step likely plays an important role in RAF activation and can be modulated by the degree of autoinhibition, which varies among isoforms and is alleviated by pathogenic mutations and inhibitors2,30,56.

CRD–lipid engagement enhances the affinity of protein–protein interaction in RAS:RAF complex at the membrane
A previous computational study predicted that CRD–lipid interactions reduce fluctuations between RAS and the RBD due to the short linker, thereby potentially increasing their overall association57. To test this prediction experimentally, we developed a FRET-based competition assay to examine whether CRD–lipid interactions influence the protein–protein interaction between RAS and CRAF (Fig. 3a). In this assay, RAS is labeled with the Atto488-GppNp donor, and CRAF (either RBD or RBD-CRD) is tagged with the AZ647 acceptor. The assay begins by forming the CRAF:RAS complex on the membrane, which generates a FRET signal. A high concentration of unlabeled RBD-CRD (3 µM) is introduced to compete with the labeled CRAF (see Supplementary Fig. 21 for the concentration optimization). Upon dissociation of AZ647-CRAF from RAS, unlabeled RBD-CRD occupies RAS, preventing the binding of labeled CRAF. The time-dependent change in FRET signal thus specifically tracks protein–protein dissociation (CRAF dissociation from RAS), while excluding dissociation of membrane-bound CRAF that is not associated with RAS. Using this assay, we observed that RBD-CRD dissociates from RAS significantly more slowly than RBD alone (Fig. 3b). The slower dissociation for RBD-CRD at 2% PS is primarily due to an additional interaction between the CRD and RAS, as its FRET dissociation curve closely follows the ensemble dissociation curve of RBD-CRD measured on lipid-free PEG surfaces (Supplementary Fig. 15b). Importantly, 20% PS lipids, which promote a greater degree of CRD–lipid interaction, further accentuates this slowed dissociation for RBD-CRD (Fig. 3b). These findings indicate that CRD–lipid interactions enhance the binding strength between protein domains (RAS:RBD and RAS:CRD), thereby stabilizing the complex and slowing its dissociation. Consequently, the protein–protein and protein–lipid interactions within the RAF regulatory domains mutually reinforce each other: RBD binding to RAS facilitates CRD–lipid engagement, which in turn strengthens RAS:RBD and RAS:CRD association (Fig. 3c).

A linker connecting RBD and CRD is crucial for positive cooperativity
We aimed to identify key structural features that mediate the positive cooperativity between RBD and CRD on membranes. X-ray crystallography and solution NMR studies reported that the RBD and CRD simultaneously bind to KRAS through distinct interfaces, forming a trimeric complex16,35. In this arrangement, the six-amino-acid-long linker connecting the RBD and CRD plays an essential role in forming the extended structure for KRAS binding by positioning these two tandem domains within close proximity (Fig. 4a). The residues involved in trimer formation are highly conserved across all RAS isoforms35. Guided by the KRAS:RBD-CRD (CRAF) structure, we investigated how the trimer formation mediates the positive cooperativity between the two domains, focusing on regulating membrane binding kinetics. We introduced double alanine mutations at L136 and T178 in the RBD-CRD construct (RBD-CRDL136A/T178A) (Fig. 4a). L136, located within the linker region, is buried within a hydrophobic pocket formed by F130 in the RBD and M183 in the CRD, bringing the two domains into close proximity35. The CRD residue T178 mediates contact with RAS through hydrogen bonding35. The L136A and T178A mutations are expected to disrupt the linker–RBD and CRD–RAS interactions, respectively, which would destabilize the RAS:RBD-CRD trimer complex. A previous study reported that L136A reduced KRAS binding affinity fourfold and T178A decreased RAF activity in cells35. Yet, we observed only marginal binding differences between the wild-type and mutant constructs on RAS-functionalized membranes containing 20% PS (red and blue, Fig. 4b), indicating that the CRD may have sufficient mobility to change its binding interface with RAS, as previously observed in molecular dynamic (MD) simulations58. Next, we replaced the entire linker sequence with a flexible GGGGGS linker of the same length. This flexible linker is expected to eliminate residue-specific interactions while increasing overall flexibility, which disrupts the proper orientation of the CRD. The replacement caused a modest decrease in membrane binding (yellow, Fig. 4b). To introduce a greater perturbation by increasing the physical separation and flexibility between the RBD and CRD, we characterized a construct containing an extended linker of four GGGGGS repeats (4×GGGGGS; 24 amino acids total). The extended linker constructs showed a greater reduction in membrane binding (cyan, Fig. 4b). Protein quality analyses (Supplementary Fig. 3–5) and circular dichroism (Supplementary Fig. 24) confirmed that all RBD-CRD linker variants were properly purified and folded. The reduced membrane binding observed for these constructs is therefore attributed to perturbations in the relative positioning between the RBD and CRD caused by the increased linker flexibility and length. Notably, the association rate remained largely unchanged across all four constructs (Fig. 4c), while differences in membrane binding affinity were mainly due to changes in the membrane dissociation rate (Fig. 4d). Thus, the CRD-mediated interactions (CRD–RAS and CRD–lipid) primarily contribute to membrane stabilization rather than to the initial association between RBD and RAS, consistent with the kon and koff analyses shown in Fig. 2.

RAS density modulates RAF membrane residence through lateral rebinding
We found that the dissociation kinetics of RBD-CRD differ markedly between free dissociation assays and competitive FRET assays. In free dissociation assays (red, Fig. 5a), the half-life was several tens of seconds—an order of magnitude longer than the half-life observed in competitive FRET assays (blue, Fig. 5a). The key difference between these two assays resides in the allowance of RAS rebinding pathways. In FRET assays, rebinding from both solution and the membrane is blocked by excess unlabeled RBD-CRD competing with the labeled CRAF. In contrast, free dissociation assays only prevent solution-based rebinding via continuous buffer flow, leaving membrane-based rebinding possible. This observation suggests that prolonged RAF membrane residence involves a lateral rebinding mechanism, where RAF briefly remains on the membrane via its lipid interactions and can subsequently rebind to another RAS molecule. In cases where RAF fails to rebind to RAS, it rapidly dissociates from the membrane. Based on these results, we propose a kinetic model for RAF membrane dissociation (Fig. 5b). The process begins with RAS dissociation from the RAS:RBD:CRD complex, resulting in a transient RBD-CRD:PS intermediate. This intermediate then unbinds from lipids, leading to complete dissociation of RAF into solution. Alternatively, it can rebind laterally to RAS, restoring the RAS:RBD-CRD:PS complex and extending its membrane residence time. We exclude the scenario where RBD-CRD first loses lipid contact yet remains bound to RAS, as a RAS:RBD-CRD complex with the CRD disengaged from lipids was not observed in single-particle tracking experiments (Fig. 2e). The slow diffusion coefficient of RBD-CRD is comparable to that of PKC regulatory domains known to penetrate into the hydrocarbon core of the bilayer59, indicating persistent lipid engagement of the CRD through its basic and hydrophobic residues in loop 1 and loop 216,55. Even if the CRD momentarily dissociates from lipids, it is likely to re-engage rapidly while still bound to RAS, given the fast lipid engagement timescale observed in our single-molecule tracking analysis (Supplementary Fig. 20).

Kinetic simulations further illuminate how elevated RAS·GTP densities, which can be achieved under receptor-triggered signaling, regulate RAF membrane dissociation kinetics. The lateral rebinding of a transient RBD-CRD:PS intermediate to RAS is a second-order process whose rate is linearly proportional to RAS surface density. Consequently, higher RAS densities increase RAF dwell times on membranes (left, Fig. 5c). In contrast, without lateral rebinding, dissociation follows a two-step, irreversible first-order pathway, removing any dependence on RAS density (right, Fig. 5c). To validate the lateral rebinding mechanism, we measured RAF dissociation kinetics at various RAS densities, revealing a strong modulation of membrane dissociation by RAS density. In particular, RBD-CRD remained on the membrane significantly longer at higher RAS densities (Fig. 5d). Although more complex models could be constructed, this two-step dissociation model successfully and quantitatively accounts for TIRF desorption data. The calculations and estimated rate constants for RAS dissociation from RAS:RBD-CRD:PS (k3), RAS rebinding (k-3), and lipid unbinding (k4) are presented in Supplementary Table 3 and its associated text.

Positively charged and hydrophobic residues in loops 1 and 2 of the CRD penetrate the bilayer and mediate membrane binding55. To test how these specific CRD–lipid interactions contribute to lateral rebinding, we characterized an RBD-CRD 4A construct containing four alanine mutations (K148A, L149A, K157A, and F158A; hereafter referred to as 4A). The 4A construct exhibited a diffusion behavior similar to that of the isolated RBD, and its membrane binding was substantially reduced compared to WT RBD-CRD (Supplementary Fig. 25a, b). Notably, the extension of membrane dwell time at higher RAS densities was significantly attenuated, confirming the critical role of CRD–lipid interactions in lateral rebinding and in the formation of the RBD-CRD:PS intermediate (Supplementary Fig. 25c). The rebinding mechanism may play a pivotal role in RAF activation under conditions of elevated RAS·GTP density. Together, these findings provide a kinetic framework for understanding how coordinated RAS and lipid inputs regulate RAF membrane residence time and activity.

Methods
Plasmid cloning
CRAF constructs were cloned into modified 2CT pET plasmids containing an N-terminal TwinStep II tag, maltose-binding protein (MBP), and a TEV protease cleavage site, followed by the protein of interest (2CT pET-TwinStrep-MBP). For mNG labeling, three repeats of the GGGGS amino acid linker were appended to the C-terminus, followed by the mNG tag. For sortase-mediated labeling with Gly-Gly-Gly-AZ647 (Vector Laboratories), a C-terminal LPETGG sequence was introduced. CRAF domains (51-131 for RBD, 51-188 for RBD-CRD, and 138-188 for CRD) were PCR-amplified and assembled using Gibson Assembly to yield TwinStrep-MBP-TEV-CRAF-mNG. Additionally, HRAS(1-184, C118S) and the SOS1 catalytic domain (SOScat, 565-1049) were cloned into a pProEX HTB plasmid to construct His6-TEV-HRAS and His6-TEV-SOScat, respectively.

Protein purification
CRAF constructs and mNG were expressed in Rosetta (DE3) cells. An overnight preculture was grown, then diluted into 1 L, and further grown at 37 °C. Protein expression was induced with 30 µM IPTG at an O.D. of 0.6 and continued for an additional five hours at 32 °C. Cells were harvested by centrifugation after five hours of induction, flesh-frozen in liquid nitrogen, and stored at −80 °C until purification. For purification, cell pellets were resuspended in lysis buffer consisting of Buffer A (20 mM HEPES [pH 7.3], 500 mM NaCl, 1 mM tris(2-carboxyethyl) phosphine [TCEP], and 10% glycerol) supplemented with 15 µg⋅mL−1 DNase and 1 mM phenylmethylsulfonyl fluoride (PMSF). Cells were lysed, and the lysate was clarified by centrifugation before purification using fast protein liquid chromatography on an NGC system (Bio-Rad). An initial capture step for CRAF constructs was performed using a Strep-Tactin affinity column (Strep-Tactin XT 4Flow, IBA Lifesciences), followed by elution with buffer A supplemented with 50 mM biotin. HiTrap chelating HP column (Cytiva Life Sciences™ HiTrap™ Chelating HP Column) was used for the initial capturing of (His)10-tagged mNG and eluted with buffer A containing 500 mM imidazole. Selected fractions were dialyzed overnight with TEV protease at 4 °C. The cleaved tag and uncleaved protein were removed by reintroducing the cleaved sample onto a Strep-Tactin column for CRAF or HiTrap chelating HP column for mNG. Further purification was performed using size-exclusion chromatography (Superdex 75 Increase HiScale, Cytiva) with buffer A. Collected fractions were analyzed by SDS-PAGE, pooled as appropriate, aliquoted, snap-frozen in liquid nitrogen, and stored at −80 °C. HRAS and SOS were expressed and purified as described elsewhere41.

SLB preparation
Glass coverslips (D263 Schott glass, Ibidi) were first cleaned by sonication in a 1:1 isopropanol/water mixture for 15 minutes, followed by incubation in 2% warm Hellmanex III for 30 min. The substrates were then etched for 10 minutes in a piranha solution (3:1 H₂SO₄/H₂O₂). The coverslips were assembled on six-channel flow chambers (sticky-Slide VI 0.4, Ibidi). SLBs were formed by fusion of small unilamellar vesicles as described elsewhere41. For SLBs containing 2% PS, the lipids were mixed at 97 mol% 1,2-dioleoyl-sn-glycero-3-phosphocholine (DOPC), 2 mol% 1,2-dioleoyl-sn-glycero-3-phospho-L-serine (DOPS), and 1 mol% 1,2-dioleoyl-sn-glycero-3-phosphoethanolamine-N-[4-(p-maleimidomethyl) cyclohexane-carboxamide] (MCC-DOPE). For SLBs containing 20% PS, the lipids were mixed at 77 mol% DOPC, 20 mol% DOPS, and 3 mol% MCC-DOPE. All lipids were purchased from Avanti Research. RAS was functionalized via a covalent reaction between the terminal cysteine residues and the maleimide group of MCC lipids as described elsewhere41. Typically, a concentration of RAS in PBS (Corning, Cat no. 46-013-CM) was incubated for 2.5 h. The reaction was terminated by incubation of 10 mM beta-mercaptoethanol (BME) for 10 minutes. Nucleotides bound to RAS were stripped with 10 mM EDTA in HBS (40 mM HEPES, 150 mM NaCl, pH 7.4). Finally, RAS was loaded with the desired nucleotide by overnight incubation of the sample with either 1 µM Atto488-GDP (Jena Bioscience) or 10 µM GppNp in HBS (40 mM HEPES, 150 mM NaCl, pH 7.4) supplemented with 5 mM MgCl2. For mNG FCS calibration measurements, SLB containing 96 mol% DOPC and 4 mol% 1,2-dioleoyl-sn-glycero-3-[(N-(5-amino-1-carboxypentyl)iminodiacetic acid)succinyl] (DOGS-NTA) were prepared by vesicle fusion method identical to SLBs containing PS lipids. A concentration ranging from 1 to 10 nM of His10-mNG was incubated in PBS for 10 min to yield various surface densities. The unbound mNG was rinsed with PBS.

PEGylated surface preparation
Glass coverslips (D263 Schott glass, Ibidi) were first cleaned by sonication in a 1:1 isopropanol/water mixture for 15 min, followed by incubation in 2% warm Hellmanex III for 30 min. The substrates were then etched for 10 minutes in a piranha solution (3:1 H₂SO₄/H₂O₂). The coverslips were assembled on six-channel flow chambers (sticky-Slide VI 0.4, Ibidi). A 2:1 ratio of Poly-L-Lysine mPEG:Poly-L-Lysine Biotin (SuSoS Surface Technology) at a total concentration of 0.1 mg/mL in DIW incubated in each flow chamber for 30 min, then washed out with twice with 1 mL of 20 mM TBS (Corning, Cat no. 46-012-CM). 0.1 mg/mL Streptavidin in TBS (MilliPore Sigma, Cat no. 18-973-010MG) was then incubated for 30 min and washed similarly. HRas 181-AviTag was then conjugated to the bound Streptavidin at 45 nM for 30 min before another wash with TBS followed by buffer exchange to HBS (40 mM HEPES, 150 mM NaCl, pH 7.4). Any nucleotides bound to RAS were stripped with 10 mM EDTA in HBS for 15 min before addition of 1 μM Atto488-GDP or non-fluorescent GppNp (Jena Bioscience) in HBS (40 mM HEPES, 150 mM NaCl, pH 7.4) supplemented with 5 mM MgCl2 and overnight incubation at 4 °C. RAS labeled with Atto488-GDP and non-fluorescent GppNp were prepared in parallel. Fluorescent samples were used to determine RAS density and non fluorescent samples were used for binding assays. The samples were rinsed with HBS (40 mM HEPES, 150 mM NaCl, pH 7.4) supplemented with 5 mM MgCl2 immediately before use.

Microscopy
Imaging was performed using a Nikon Ti-2 microscope equipped with an Apo TIRF 100× oil immersion objective and an EMCCD camera (iXon Life 897, Oxford Instruments). For RAS density and RAF-mNG measurements, a 488 nm laser (Coherent), filtered through a laser cleanup filter (ZET488/10x, Chroma Inc.), was directed onto the sample using a dichroic mirror (zt488/640rpc, Chroma Inc.). The fluorescence was collected through two stacked emission filters (ZET488/640m and ET535/70m, Chroma Inc.). RAF-AZ647 was excited with a 640 nm laser (Coherent), filtered through a laser cleanup filter (LD01-640/8, Semrock) and directed by the same dichroic mirror (zt488/640rpc). The fluorescence was collected through two stacked emission filters (ZET488/640m and ET706/95m, Chroma Inc.).

Ensemble adsorption and desorption
In ensemble experiments, SLBs were initially prepared with RAS loaded with the fluorescent nucleotide Atto488-GDP to determine surface density. Nucleotide exchange to GTP was facilitated by incubation of 100 nM catalytic domain of SOS and 100 µM GTP in HBS (40 mM HEPES, 150 mM NaCl, pH 7.4) supplemented with 5 mM MgCl2 and 0.1 mg·mL−1 casein at 23 °C for 6 min. Active RAS·GTP-functionalized membranes are nonfluorescent and suitable for subsequent binding assays with RAF fused with mNG. RAF-mNG was diluted as specified in the figure captions according to the protein concentration determined by absorbance at 280 nm. The binding measurements were performed in HBS (40 mM HEPES, 150 mM NaCl, pH 7.4) supplemented with 5 mM MgCl2, 0.1 mg·mL−1 casein, 1 µM GTP, and 1 mM BME at 23 °C. Upon reaching a plateau in the binding curve (typically in 5 min), desorption measurements were conducted in the same complete buffer including all supplemented components. Desorption was initiated by injecting 200 µL of the buffer, followed by continuous buffer flow at a rate of 1 mL·min−1 using a syringe pump. Kd measurements and desorption assays on PEGylated surfaces were performed at 23 °C identically to SLB samples, except that nucleotide exchange by SOS was omitted; RAS was incubated overnight with GppNp. For Kd measurements, RBD-mNG and RBD-CRD-mNG were incubated at concentrations ranging from 5 to 750 nM. For desorption measurements, 200 nM was used. Background signals, Ibg, were obtained from the average intensities of the frames prior to RAF injection and were subtracted from the raw binding intensities. The intensity values were then divided by the maturation efficiency, MEff, of RAF-mNG proteins. The solution contribution, ISoln, of mNG was then subtracted to calculate the binding/desorption net intensities, I, for analysis.

I = (Iraw - Ibg) / MEff - ISoln (1)

The net intensity was converted to the surface density using the TIRF-FCS calibration curve.

Competitive FRET dissociation
Competitive desorption assays were performed using Förster Resonance Energy Transfer (FRET) to quantitatively assess the stability of the RAS:RAF complex. The donor Atto488-GppNp (Jena Bioscience) was loaded on RAS (RAS-Atto488-GppNp) and the acceptor AZ647 (Vector Laboratories) was conjugated to either RBD or RBD-CRD of CRAF (RAF-AZ647). The assays were performed in HBS (40 mM HEPES, 150 mM NaCl, pH 7.4) supplemented with 5 mM MgCl2, 0.1 mg·mL−1 casein, 1 µM GTP, and 1 mM BME at 23 °C. To mitigate photobleaching, the buffer was further supplemented with enzymatic oxygen scavenging reagents: 0.32 mg·mL−1 glucose oxidase from Aspergillus niger (Serva), 0.05 mg·mL−1 catalase from bovine (Sigma-Aldrich), 2 mM Trolox (Sigma-Aldrich), and 20 mM glucose. The RAS:RAF complex was formed by incubating 200 nM of RBD-AZ647 or 20 nM RBD-CRD-AZ647 (according to the protein concentration determined by absorbance at 280 nm) with RAS tethered on SLBs for 5 min. 3 µM unlabeled RBD-CRD was subsequently introduced. FRET measurements were acquired and analyzed as described elsewhere68. Three fluorescence signals were recorded: donor/acceptor (DA), acceptor/acceptor (AA), and donor/donor (DD). DD and AA are regular fluorescence signal acquired with the optical configurations described in the “Microscopy” section. The DA signal was acquired with a 488 nm laser excitation and emission configured for 640 nm channel. The measured DA signal, representing total acceptor fluorescence under donor excitation, is described by the following equation:

FLDA = EmFRET + α FLDD + β FLAA (2)

Here, FLDA is the total acceptor emission under donor excitation, EmFRET is the sensitized emission due to FRET, α FLDD accounts for donor emission bleed-through into the acceptor channel, and β FLAA accounts for direct acceptor excitation at the donor excitation wavelength. The FRET signal was calculated by subtracting the contribution of α FLDD and β FLAA from FLDA.

Single-molecule recruitment
Single-particle recruitment measurements of RAF-AZ647 were conducted on RAS·GTP-functionalized membranes in HBS (40 mM HEPES, 150 mM NaCl, pH 7.4) supplemented with 5 mM MgCl2, 0.1 mg·mL−1 casein, 1 µM GTP, and 1 mM BME at 23 °C RAF-AZ647 was diluted as specified in figure captions according to the AZ647 concentration. TIRF measurements were acquired using a 20 ms exposure time in stream mode, with the 640 nm laser power set to 5 mW. Single-molecule trajectories were detected and built using the TrackMate plugin in ImageJ69. Cumulative binding events were then counted and plotted over the duration of the measurement using custom MATLAB scripts. The slope of this plot, normalized by RAS density, was used to determine kon41.

Single-molecule tracking
Single-molecule diffusion experiments were conducted at 23 °C in HBS (40 mM HEPES, 150 mM NaCl, pH 7.4) supplemented with 5 mM MgCl2, 0.1 mg·mL−1 casein, 1 µM GTP, and 1 mM BME, as well as oxygen scavenging reagents including 0.32 mg·mL−1 glucose oxidase from Aspergillus niger (Serva), 0.05 mg·mL-1 catalase from bovine (Sigma-Aldrich), 2 mM Trolox (Sigma-Aldrich), and 20 mM glucose. RAF-AZ647 was diluted as specified in figure captions according to the AZ647 concentration. RAF-AZ647 and RAS-Alexa647-GppNp diffusion measurements were performed on a RAS·GTP-functionalized SLBs with 10 ms time resolution. Single-molecule trajectories were detected and built using the TrackMate plugin in ImageJ69. Trajectories with a mean speed of 0.5 pixels per 10 ms were accepted to exclude a minor fraction of immobile particles. The step size distribution was calculated using custom MATLAB scripts70. A Brownian diffusion model, described below, was used to fit the step size distribution of each measurement:

p(r, t, D) = α r/2D1t exp(-r2/4D1t) + (1-α) r/2D2t exp(-r2/4D2t) (3)

Diffusion coefficients, D1 and D2, and relative population, for the fast species, α were calculated from the corresponding fitting. Step size is represented by r, at an experimental time interval, t.

Fluorescent correlation spectroscopy
Measurements were performed on a custom-built FCS setup integrated with a Nikon Ti-2 microscope. A SuperK Evo HP white-light laser source (NKT Photonics) was reflected by a dichroic mirror (zt488rdc, Chroma Inc.) and filtered through a cleanup filter (LL01-488, Semrock) to select a 488 nm wavelength. The beam was coupled into a polarization-maintaining single-mode fiber (PM-S405-XP, Thorlabs) via a fiber coupler (PAF2-A4A, Thorlabs). The beam collimated via a reflective collimator (RC08FC-P01, Thorlabs), directed to the back port of the microscope and then to the objective by a quad-band dichroic filter cube (ZT405/488/561/640rpc, Chroma Inc.). The excitation beam was focused by Apo TIRF 100× oil immersion objective. Fluorescence emission from the sample was spatially filtered through a 50 µm pinhole, collimated by an achromatic doublet lens (AC254-125-A, Thorlabs), reflected by a dichroic mirror (550lpxr, Chroma Inc.), and further filtered through a band pass filter (ET520/40m, Chroma Inc.). It was then refocused by another achromatic doublet lens (AC254-75-A, Thorlabs) onto an avalanche photodiode (SPCM-AQRH-16, Excelitas Technologies) connected to a time-correlated single-photon counting module (PicoHarp 300, PicoQuant). Data acquisition was performed using PicoHarp software (PicoQuant) and analyzed in Prism (GraphPad). The autocorrelation function, G(τ), was fitted to a two-dimensional Gaussian diffusion equation to calculate surface density of RAF and RAS.

G(τ) = 1/N * 1/(1+τ/τd) (4)

where N is the number of particles within the focal area, τ is the time delay, and τd is the correlation time. The focal radius was determined to be 0.169 µm and was used to calculate the protein density.

Peer review rounds
[{"round": 1, "reviewer_comments": {"reviewer_1": "In this manuscript, Jimenez Salinas et al. investigate how the tandem RAS-binding domain (RBD) and cysteine-rich domain (CRD) of RAF cooperatively regulate membrane binding kinetics using a supported lipid bilayer-based reconstitution system. They show that RBD binding to RAS facilitates CRD engagement with phosphatidylserine (PS) lipids, thereby extending RAF membrane dwell time via a proposed lateral rebinding mechanism. The authors further suggest that this prolonged residence could enable kinetic proofreading to enhance signaling specificity in RAS nanoclusters.\nWhile the study utilizes multiple biophysical approaches and presents kinetic measurements, it lacks critical experimental information that compromises both reproducibility and interpretation. Key parameters—such as buffer composition, salt concentration, protein concentration, temperature, and protein solubility/stability—are either absent or inadequately described. In addition, no methodological detail is provided for the generation, expression, or biochemical validation of mutant and linker-variant constructs, making it impossible to assess whether these proteins are comparably expressed, folded, or stable relative to wild-type controls.\nBeyond these technical limitations, the conceptual contribution of the study is modest. The major conclusions—positive cooperativity between RBD and CRD, and the role of CRD–lipid interactions have been established previously through structural, computational, and biochemical studies. This work largely validates existing models in a minimal in vitro system and provides limited advancement in our mechanistic understanding of RAF activation.\nSpecific concerns:\nThe study does not use full-length BRAF constructs, including autoinhibited and active forms resolved by cryo-EM, which are necessary to explore physiologically relevant conformational transitions and regulation. The study uses BRAF cryoEM structures and utilizes isolated CRAF RBD, CRD, and RBD-CRD constructs.\nThe intrinsic instability of the isolated CRD domain relative to RBD or RBD-CRD is not acknowledged or controlled for, despite likely influencing the observed behaviors. Previous studies have suggested utilizing high salt concentration to keep it soluble/stable.\nThe authors use an HRAS(1–184) construct, even though mature HRAS is 185 residues long. They do not specify which cysteine is modified by maleimide or address the implications of this for membrane anchoring and orientation.\nMaleimide conjugation is a non-physiological means of membrane tethering that does not replicate the native prenylation of KRAS, which has been extensively utilized in the recent past. This limitation is not discussed, despite its potential impact on membrane interaction kinetics.\nThe manuscript omits critical methodological details: the final composition of protein storage buffers, assay buffers, salt concentrations, assay temperatures, and precise protein concentrations used in kinetic and diffusion experiments are not reported.\nThere is no description or validation of the linker mutant or extended linker constructs. The authors provide no data on their expression levels, solubility, or folding.\nThe rationale for the L136A/T178A double mutant is unclear. T178 is not part of the linker, and its selection is not justified. Similarly, the purpose/rationale of introducing a 24-residue flexible linker is not explained.\nThe claim that the T178A mutation stabilizes the extended linker construct is not supported by any biochemical data.\nThe known ~2-fold higher affinity of RBD-CRD for RAS compared to RBD alone is not factored into the interpretation of results. Many observations attributed to RBD-CRD vs RBD or CRD could reflect this baseline affinity difference.\nThe proposed enhancement of RBD–RAS association by CRD–lipid interaction is not clearly demonstrated or mechanistically explained.\nThe claim that RAS·GTP mediates membrane recruitment of RBD-CRD species not bound to RAS is conceptually inconsistent, especially under conditions where all RAS binding sites are presumably occupied.\nThe term “RAS unbinding” is used ambiguously throughout and should be clarified to distinguish between RAS–RAF dissociation and membrane detachment.\nThe central conclusion regarding nanocluster-specific signaling is not experimentally supported. The lipid compositions used do not promote RAS nanocluster formation, and the system lacks the complexity to model this behavior. Therefore, extrapolating these findings to nanocluster biology is not justified.", "reviewer_2": "The manuscript by Salinas et al. presents a well-thought-out and comprehensive kinetic analysis of the RAF1 RBD-CRD domain interactions with HRAS and a simple supported lipid bilayer (SLB). The SLB is composed of DOPS, DOPC, and a functionalized DOPE that links HRAS to the bilayer via maleimide chemistry through an available cysteine in HRAS’s hypervariable region. I believe this study proposes a model that is both intriguing and valuable to the field, and that the supporting data and analysis are sound.\nIn brief, the authors propose that, following initial binding of RAF to RAS on the membrane, subsequent lipid interactions involving the RAF CRD domain are sufficient to extend RAF’s residence time. Nanoclustering of RAS and a CRD–lipid intermediate increases the likelihood of subsequent rebinding to other RAS monomers, enabling the full RAF activation sequence—including RAF dimerization—to proceed. This model has important implications for understanding the RAF activation cycle, particularly the possibility of a kinetic proofreading function.\nTherefore, I recommend the manuscript for publication, provided the following points are addressed:\n• The binding and dissociation rates of RAS/RAF complexes were calculated in a previous single-molecule pull-down study: Lee, H.W., Kyung, T., Yoo, J., et al. Nat Commun 4, 1505 (2013). In this study, RAS was immobilized, and RAF molecules were pulled down from a lysate onto coverslips and imaged using TIRF microscopy. Residence times and kinetic parameters were measured. The authors should explicitly compare their data to this study. Moreover, as in Lee et al., they should account for potential photobleaching effects to ensure that their measured rates are not significantly influenced by the photophysics of the fluorophores.\n• Lee et al. describe the formation of “pre-equilibrating encounter complexes.” The authors should clarify whether these results are consistent with or differ from those in the current manuscript, especially with respect to the proposed lateral rebinding model. Although Lee et al. used immobilized RAS (i.e., non-diffusing and in a lipid-free system), it is possible that their findings contradict the current interpretation. Are there alternative explanations that support the proposed lateral rebinding and kinetic proofreading model?\n• The authors cite a recent study using KRAS4b on SLBs, which analyzes KRAS4b/RBD-CRD complexes on simple and complex lipid bilayers: Shrestha R., Carpenter T.S., Van Q.N., et al. Commun Biol. 2024; 7(1):242. The authors should comment on the specific CRD-lipid contact points identified by the simulations and NMR in Shrestha et al. and evaluate experimentally whether mutations of these lipid-penetrating CRD residues affect lateral rebinding interactions in the RBD-CRD complex or the proposed RBD-CRD–PS intermediate.\n• In the final sentence of paragraph 1 in the section titled “RAS density modulates RAF membrane residence through lateral rebinding,” the authors state: “we exclude the scenario where RBD-CRD first loses lipid contact yet remains bound to RAS, as a lipid-free RAS:RBD-CRD complex was not observed in single-particle tracking experiments (Fig. 2e).” To more robustly support this claim, the authors should measure the diffusion of fluorescently labeled RAS in the presence of RBD-CRD on the same SLB and compare those diffusion rates to the measurements where the fluorescent tag is on RBD-CRD. Comparing the distributions and fractional occupancies of the different diffusion states in both labeling conditions would provide a stronger test of the assertion that lipid-free RAS:RBD-CRD complexes do not form or persist. I would also encourage a different term than “lipid-free.” We know that there are extensive and dynamic contacts between RAS and lipids. There is probably more precise language to talk about the conformational state of the complex and contact with the lipids in the bilayer.", "reviewer_3": "This is an interesting and potentially impactful article that would be suitable for publication in a major journal such as Nature Communications, following adequate attention to significant shortcomings. The article summarizes well-reasoned biophysical studies of the multi-pronged interactions between the important signaling proteins CRaf, HRas and anionic lipids (phosphatidylserine or PS) during CRaf membrane binding, 2D-diffusion and dissociation on a lipid bilayer possessing anchored HRas molecules. The quite promising results suggest that, as often observed for signaling proteins that target the plasma membrane, the CRaf molecule possesses cooperative, multi-component binding contacts involving both protein-lipid and protein protein interactions that together provide adequate specificity and affinity for the correct membrane surface. In CRaf this multi-component interaction is provided by its distinct cysteine-rich domain (CRD) and Ras binding domain (RBD) that provide protein-lipid (CRD-PS) and protein-protein (RBD-HRas), respectively. While these results are not surprising, this manuscript is the first to address the cooperative nature of these essential Ras:CRaf:PS interactions in a quantitative, biophysical manner. The following major and minor points should be addressed prior to publication.\n1) The article places a major emphasis on Raf targeting, retention, and activation in Ras nanoclusters with language about relevance to nanoclusters on 10 pages of the text including the Abstract, Introduction, Results, Discussion, and Figure Legends. However, as the article itself notes, the system employed observes individual Ras:RBD-CRD complexes interacting with PS lipids on the target bilayer with no detectable clusters (pg 8, bottom half of paragraph). Moreover, it appears that Ras densities and clustering are not measured in most or all experiments. Thus the article should be more careful about its claims of relevance to clustered systems, and should limit those claims to the description of the plausible, but as yet unproven, model in the Discussion.\n2) The experiments are well chosen and the data is promising, but there is inadequate attention to reproducibility which could lead to questions about the rigor of the work. In general, statistical tests of significance are absent and there is information on replicates for only two of the many experiments (in both cases N=3, presumably representing technical rather than biological replicates). Perhaps I missed it, but I could find no indication that experiments were carried out more than once, and were repeated on multiple days. Such lack of information about replicates and statistical tests makes it impossible to judge the reproducibility and statistical significance of the findings.\n3) The slow diffusion observed for Ras:RBD-CRD:PS complexes on the DO-lipid bilayer raises the possibility that, in addition to the frictional drag of the Ras anchor lipid and the bound PS molecules against the bilayer, there may well be additional frictional drag provided by protein insertion into the bilayer. See PMID 23701821.\n4) The article is missing key references to directly relevant single molecule studies of Ras-effector interactions, and of the effects of membrane environment on the affinities and kinetics of protein-protein and protein-membrane interactions. See, for example, PMID 29211993, 39217418, 22263647.", "reviewer_4": "This manuscript provides interesting results and confirms several observations from previous experimental and computational studies. I recommend that authors stress the novelty of their findings in the context of broader Ras biology during the revision of this manuscript.\nThe manuscript reports that RBD binds to RAS, allowing CRD to bind to negatively charged PS lipids (as determined by TIRF microscopy). They propose two kinetically distinct binding steps. Also, CRD binds to PS in a RAS-GTP-dependent manner. It ensures RAF membrane binding (by CRD) is strictly driven by RAS activation (by RBD). Using FRET, they demonstrate that CRD-lipid binding enhances the association between RBD and RAS (as measured by FRET). RBD and CRD binding to RAS and PS lipids exhibit cooperativity. RAF undergoes lateral rebinding with RAS (using kinetic simulations in SimBiology and TIRF microscopy). Weak CRD-lipid interactions play a role in lateral rebinding. This extends the RAF's membrane dwell time under high RAS density (e.g., nanoclusters). This prolonged membrane residence increases the likelihood of completing RAF's multistep activation sequence (i.e., facilitates kinetic proofreading of RAF activation).\nComments:\nThis work would have had a significant impact if the authors had investigated the kinetics in the presence of PIP2 lipid and distinguished the binding mechanisms.\nAdditionally, DOPC is not a particularly abundant component of the plasma membrane. Any reason for its selection? (CRD is more likely to bind to POPC:POPS membranes than DOPC:DOPS)\nAlso, is a 20% PS concentration sufficient to mimic biomembranes?\nWhat would be the effect of having the kinase domain? Past experiments were based on the same RBD-CRD construct. Engaging the kinase domain and seeing the effect of homodimerization can be quite novel.\nI find it challenging to convince myself that RAF unbinding occurs primarily by detaching from RAS first, then from the membrane, rather than the other way around (end of first paragraph on p. 12). Strong arguments to support their claim can be useful.\nSeveral xray structures clearly show that CRD creates contacts with RAS. I agree that these contacts may not be formed in the absence of RBD. Therefore, lipids are not the only driving force for membrane interaction. Claiming that CRD is driven only by lipids is not completely correct.\nIt is interesting that RBD alone has a higher k_on to RAS-associated membranes than RBD-CRD; however, the presence of CRD lowers the off rate, resulting in RBD-CRD having a stronger binding affinity than RBD alone. Additionally, the rate constants for RBD alone and RBD-CRD are not affected by PS concentration, so only k_off is influenced by PS levels. I don’t quite understand that easily. Some clarity would be helpful.\nAdditionally, I find it puzzling that the RBD-CRD linker does not affect the initial RBD-RAS association but is relevant for the membrane dissociation rate, and thus for the cooperative binding of RBD-CRD to RAS and PS lipids.", "reviewer_5": "In this study, the authors use quantitative TIRF microscopy to analyze the binding kinetics and equilibrium behavior of two RAF domains (RBD and CRD). Particularly, they looked at RBD/CRD interactions with phosphatidylserine lipids, RAS, and cooperative binding interactions with each other. The purpose of this study was to develop a model for membrane-dependent RAF activation, which is dependent on recruitment to the membrane by GTP-RAS, and retention on the membrane through weak PS interactions which cooperatively strengthen binding to RAS. The carefully designed experiments elucidate KD values, koff rates, and kon rates to elucidate the impact of each component in the system. The RAF-MEK-ERK signaling cascade is a classical biochemistry interaction that is fundamentally important to numerous cell signaling events. Understanding activation of this pathway is critical to being able to regulate cell signaling in cancer cells. This work will be of high interest to a number of fields because of the extreme importance of RAF activation in cancer.\nOverall the conclusions of the study are supported by the results. The experiments are well-designed, and the interpretations are logical. There are some minor points where the language and reporting in the manuscript could be clarified, as noted below. The manuscript is well written and the figures are generally of good production quality. The biggest concern to this reviewer is there may be an over-extrapolation of these results to connections concerning RAS nanoclusters, RAF auto-inhibition, and involvement of the SHOC2-MRAS-PP1C complex. I feel that these results provide a convincing argument about RBD/CRD cooperativity, interactions with PS lipids, and importance of RAS concentration. However, the attempt to explain a full mechanism of RAF activation seems to be speculation and extrapolation based on other published works. I recommend the manuscript undergo minor revisions to address the points listed below, where the issue of over-extrapolation is elaborated on in the last point.\nPoints to address:\n• Some additional clarifications could be added to the introduction to aid readers who are not in this field:\no It was not immediately obvious to me that “14-3-3” is the name of a protein. Please add a (very) brief explanation. Even just add the word “protein” after it the first time it appears.\no Please briefly explain what are BRAF, KRAS, and CRAF, compared to just RAF and RAS.\n• Evaluating how the RBD-CRD had measured KD values of 14 nM vs 5 nM for binding 2% and 20% PS membranes as a “evident dependence on PS” seemed strange in contrast to the previous comparison of 222 nM vs 163 nM as “depended little on upon PS content”. Please adjust the language or clarify.\n• Was there a 0% PS control performed for any of these samples? Why were 2% and 20% PS concentrations chosen for the experiments? Please explain the lipid choice briefly.\n• The number of reported significant figures should be reduced for reported KD values in caption for Figure 1. Example: 222 +/- 88 should be changed to 220 +/- 90\n• Figure 2a, middle panel: I think I’m seeing some low-intensity spots indicating protein binding in the -RAS condition. However, I can’t rationalize how you can have less than one molecule showing up, unless this is a product of partial occupancy over the duration of a given exposure time? Please explain.\n• It is odd that the experiment wasn’t successful for the 4XGGGGGS linker construct, but it was successful for the 4XGGGGGS/T178A construct. Can the authors please provide their thoughts on this? What part of the experiment was unsuccessful? Could the protein not be expressed? Perhaps this doesn’t need to be explained in the manuscript, but I think it is appropriate for discussion during the review.\n• The statement in the discussion “Consistent with this, we observed stronger binding of RBD-CRD on RAS-free membranes” is unclear. After a few minutes of consideration, I am now thinking that perhaps you mean that RBD-CRD has stronger binding on RAS-free membranes compared to RBD alone or CRD alone? When I originally read this sentence, I was interpreting it as RBD-CRD has stronger binding on RAS-free membranes, compared to membranes containing RAS. Please revise for clarity.\n• To improve clarity, it would be helpful to reference figure numbers throughout the discussion.\n• This point is in regards to the statement in the discussion: “… whereby molecules that remain on the membrane for extended periods – such as those within RAS nanoclusters – are disproportionately more likely to become activated.” Based on the evidence presented here, I don’t see how a RAS nanocluster would retain RAF any more than a high concentration of monomeric RAS. I’m seeing how this is related to concentration, but not necessarily how it’s related to nanoclusters. This needs to be made clearer. Perhaps making it explicit that clustering is creating a high local concentration, and RAF will be retained due to this high local concentration. This is distinct, for example, from requiring a patch of RAS that is a certain diameter or particular shape."}, "author_response": "Response to the reviewers’ comments\nWe are grateful to reviewers for their constructive comments, all of which have been addressed\nin the revised manuscript. Below, we respond to each comment point-by point. The inclusion of\nadditional replicates and standardizing significant figures led to small changes in some reported\nvalues; these revisions do not alter any interpretation and conclusions. We also made minor edits\nto maintain the logical flow and the manuscript length after incorporating the reviewers’\nsuggestions and the new data.\nThe original comments from the reviewers are in bold italic; our responses are in regular font.\nChanges in the manuscript are colored in blue.\nTable of Contents\nReviewer #1 (Remarks to the Author): ...................................................................... 2\nReviewer #2 (Remarks to the Author): .................................................................... 13\nReviewer #3 (Remarks to the Author): .................................................................... 17\nReviewer #4 (Remarks to the Author): .................................................................... 20\nReviewer #5 (Remarks to the Author): .................................................................... 26\nIn this manuscript, Jimenez Salinas et al. investigate how the tandem RAS-binding domain\n(RBD) and cysteine-rich domain (CRD) of RAF cooperatively regulate membrane binding\nkinetics using a supported lipid bilayer-based reconstitution system. They show that RBD\nbinding to RAS facilitates CRD engagement with phosphatidylserine (PS) lipids, thereby\nextending RAF membrane dwell time via a proposed lateral rebinding mechanism. The\nauthors further suggest that this prolonged residence could enable kinetic proofreading to\nenhance signaling specificity in RAS nanoclusters.\nWhile the study utilizes multiple biophysical approaches and presents kinetic measurements, it\nlacks critical experimental information that compromises both reproducibility and\ninterpretation. Key parameters—such as buffer composition, salt concentration, protein\nconcentration, temperature, and protein solubility/stability—are either absent or inadequately\ndescribed. In addition, no methodological detail is provided for the generation, expression, or\nbiochemical validation of mutant and linker-variant constructs, making it impossible to assess\nwhether these proteins are comparably expressed, folded, or stable relative to wild-type\ncontrols.\nWe have addressed all issues in our point-by-point responses below.\nBeyond these technical limitations, the conceptual contribution of the study is modest. The\nmajor conclusions—positive cooperativity between RBD and CRD, and the role of CRD–lipid\ninteractions have been established previously through structural, computational, and\nbiochemical studies. This work largely validates existing models in a minimal in vitro system\nand provides limited advancement in our mechanistic understanding of RAF activation.\nWe respectfully propose that our study is the first quantitative biophysical analysis providing a\nkinetic model for RAF–membrane interaction, with novel mechanistic insights at the molecular\nlevel. Reviewer #3 also acknowledged that our manuscript is “the first to address the cooperative\nnature of these essential Ras:CRaf:PS interactions in a quantitative, biophysical manner.”\nWe agree that positive cooperativity between RBD and CRD has been extensively studied;\nhowever, many questions remain regarding their functional and structural communication,\nparticularly within the membrane context. This was one of the central questions in this work:\nhow do RBD and CRD cooperate to integrate protein and lipid inputs in membrane\nenvironments? We sought to address these questions through quantitative measurements in\nsimplified model membrane systems, moving beyond qualitative description.\nSpecifically, previous structural and NMR studies have reported positive cooperativity between\nthe RBD and CRD, in which nucleotide-dependent KRAS–RBD binding promotes transient\ninteractions between KRAS and the CRD. In contrast, our study provides a comprehensive\nkinetic analysis of protein–protein and protein–lipid interactions within membrane environments.\nOur findings reveal membrane-specific positive cooperativity and resulting kinetics in which\nCRD–lipid engagement enhances the protein–protein interaction within the RAS:RAF complex,\nand vice versa. We also identified the distinct mechanistic roles of RAS–RBD, RAS–CRD, and\ndiscovered that RAF membrane residence is modulated by RAS density through a lateral\nrebinding mechanism, in which a weak CRD–lipid interaction plays a key role. This lateral\nrebinding mechanism in RAF has not been previously observed or proposed. Because RAF\nactivation involves a series of multistep processes on the membrane, these findings provide\ncritical mechanistic insight into how coordinated RAS and lipid interactions collectively drive\nfull RAF activation.\nTo better reflect this point, we modified the introduction as follows (page 3, line 64):\n“Prior studies reported cooperativity between RBD and CRD in which RAS–RBD binding\ninduces a transient interaction between RAS and CRD, and mapped CRD–lipid interfaces16.\nHowever, how these interactions control membrane kinetics has remained unclear. A quantitative\nunderstanding of membrane-binding kinetics and the underlying mechanisms is crucial because\nRAF is exclusively activated on cellular membranes14.”\nSpecific concerns:\n1. The study does not use full-length BRAF constructs, including autoinhibited and active\nforms resolved by cryo-EM, which are necessary to explore physiologically relevant\nconformational transitions and regulation. The study uses BRAF cryoEM structures and\nutilizes isolated CRAF RBD, CRD, and RBD-CRD constructs.\nThe reviewer raises two concerns: (1) our use of truncated CRAF constructs rather than full-\nlength BRAF, and (2) our use of BRAF cryo-EM structure while utilizing CRAF constructs. We\nwill address each point below. :\n(1) Why use truncated CRAF constructs rather than full-length BRAF constructs?\nRAF activation is a complex process that involves multiple protein–protein and protein–\nlipid interactions within membrane environments. We strategically planned to dissect this\ncomplex process with the conserved regulatory module (RBD-CRD) in a minimal, well-\ncontrolled setting. This approach enabled high-resolution kinetic analysis and revealed a\npreviously unrecognized membrane-interaction mechanism—lateral rebinding—by\nwhich RAF integrates protein and lipid inputs to achieve prolonged membrane residence.\nCRAF was chosen because its regulatory domains are extensively characterized\nbiochemically and structurally, enabling structure-driven hypotheses and direct\ncomparison between solution and membrane contexts. Given the modular domain\narchitecture and conservation across RAF, we expect that the positive cooperativity and\nlateral rebinding observed for the RBD-CRD module will also be present in the full-\nlength proteins. We agree that full-length protein is essential to understand the complete\nactivation process, but full-length RAF study is beyond the scope of this work. Our\ncurrent work will provide a mechanistic foundation for future work with full-length RAF.\nWe now make this limitation explicit in our planned extensions to full-length RAF\nreconstitutions (page 17, line 418):\n3\n“To investigate how membrane-binding kinetics influence activation outcomes under\nmore physiologically relevant conditions, it will be necessary to use the autoinhibited\nfull-length RAF:14-3-3:MEK complex.”\n(2) Why show BRAF cryo-EM structures while using CRAF constructs experimentally?\nThe cryo-EM structure of autoinhibited BRAF was used as the current structural\narchetype for RAF because autoinhibited CRAF structures were not yet officially\navailable at the time of submission; autoinhibitory mechanisms observed in BRAF are\nwidely accepted to be conserved in CRAF (Ref. 27; Part et al., Nature 2019, 575, 545).\nRecently, autoinhibited CRAF structures—including a conserved autoinhibited structure\nsimilar to BRAF—have been released (Ref. 30; Jang et al. Nat Commun 2025, 16, 8150).\nWe have replaced the BRAF cryo-EM structure in Figure 1b with the fully autoinhibited\nCRAF structure.\nTo reflect this change, we have modified the main text as follows (page 2, line 47):\n“The inactive monomeric CRAF:MEK1:14-3-3 complex can adopt multiple\nconformations, including a fully autoinhibited confirmation (Fig. 1b) that is closely\nresembles that of BRAF, as well as more open conformations in which the CRD is\nstructurally liberated30.”\nPlease note that all mutational designs and linker variants in our experiments were guided\nby the CRAF RBD-CRD structure (Ref. 35; Tran et al., Nat. Commun. 2021, 12:1176),\nnot BRAF.\nRegarding the BRAF cryo-EM structure, we originally stated that RAS binding to the\nRBD alone is insufficient to release RAF from autoinhibition, based on the reported\nstructure of the RAS:BRAF recruitment complex (Ref. 29; Park et al. Nat Commun 2023,\n14, 4580). However, more recent structural and biochemical studies by Ritt et al. (Ref.\n31; PNAS 2025, 122, 38.) have shown that RAS binding to the RBD initiates the\ndisassembly of the autoinhibited complex. We have updated the manuscript to reflect this\nmost up-to-date understanding by deleting the previous statement and adding the\nfollowing text (page2, line 54):\n“For BRAF, RBD–RAS binding initiates the disassembly of the autoinhibited state, and\nCRD–lipid interaction is required for full activity31.”\n2. The intrinsic instability of the isolated CRD domain relative to RBD or RBD-CRD is not\nacknowledged or controlled for, despite likely influencing the observed behaviors. Previous\nstudies have suggested utilizing high salt concentration to keep it soluble/stable.\nWe appreciate the reviewer’s thoughtful comment regarding this important control. During the\nfinal size-exclusion chromatography (SEC) purification step of purification (buffer: 20 mM\nHEPES [pH 7.3], 500 mM NaCl, 1 mM TCEP, and 10% glycerol), the CRD eluted at the\n4\nexpected volume based on protein standards, and SDS–PAGE analysis confirmed high purity.\nThese results indicate that the CRD was robustly expressed and properly purified.\nIn our membrane assays, we used a buffer containing 40 mM HEPES (pH 7.4), 150 mM NaCl,\nand 5 mM MgCl₂. We did not observe any detectable aggregation on membranes in either\nensemble or single-molecule binding assays. To examine this more thoroughly, we\nnonspecifically immobilized CRD-mNG on a glass surface in our assay buffer and analyzed its\nfluorescence intensity distribution at the single-molecule level. The CRD exhibited a slightly\nbroadened distribution with a mean intensity approximately 25-30% higher than other constructs,\nand the distribution tail extended only modestly (Supplementary Fig. 5). These results suggest\nthat the CRD may contain a small fraction of dimers but no evidence of larger oligomeric\nspecies.\nTo reflect this discussion in Results, the following text has been added to the Results (page 5,\nline 112):\n“Data demonstrating protein quality—SEC and SDS–PAGE, and single-molecule intensity\nanalyses—are provided in Supplementary Figure 4–6. The single-molecule intensity distribution\nsuggests the presence of a minor dimeric fraction of the CRD-mNG construct, with no evidence\nof larger oligomers, at 150 mM NaCl, while all other proteins were purely monomeric”\nAdditionally, a previous study has shown that the CRD is sufficiently stable at a physiological\nsalt concentration to allow characterization of its membrane binding. Spencer-Smith et al. (Ref.\n2; Spencer-Smith et al. Mol Cell 2022, 82, 4262-4272) quantified CRD binding to nanodiscs\ncontaining 20% PS at 150 mM NaCl using SPR. They observed minimal binding at 0.3 M, and\nmicromolar concentrations were required to detect substantial SPR signals. This weak binding of\nthe CRD at nanomolar concentrations is consistent with our observations (Fig. 1g). Similarly,\nTravers et al. (Ref. 60; Travers et al. Biophys J 2020, 119, 1-14) also reported comparably low\nlevels of CRD binding at 1 M on 30% PS liposomes at 250 mM NaCl.\n3. The authors use an HRAS(1–184) construct, even though mature HRAS is 185 residues\nlong. They do not specify which cysteine is modified by maleimide or address the implications\nof this for membrane anchoring and orientation.\nDetails of the specific residues used for maleimide-conjugation of HRAS and the validity of this\nmethod are described in the Result (page 4, line 100) with relevant references. We used its native\ncysteine palmitoylation sites (C181 and C184) to covalently link to lipids via maleimide\nchemistry. Maleimide conjugation has been extensively used in membrane reconstitution studies\nand has been validated to preserve the biological function and membrane orientation of RAS at\nlevels comparable to those of the fully processed protein. The relevant references are provided\n(Ref. 40-47, including three newly added references).\nAlthough HRAS orientation is less studied than other RAS isoforms, prior work shows that the\nNRAS G-domain remains largely free of lipid interaction, exhibiting rapid free rotational\nmobility (Ref. 48; A. Werküller et al. ChemPhyChem 2013, 14, 3698-3705). Furthermore, a\nrecent NMR study revealed that ~90% of KRAS adopts the membrane-distal state in which the\nG-domain does not directly contact the membrane, but it is tethered via its C-terminal HVR (Ref.\n5\n49; Van et al. PNAS 2020, 117, 24258-24268), facilitating the capture of effector proteins.\nConsistent with these observations, we expect the HRAS G-domain to undergo rapid free\nrotational diffusion on membranes, remaining largely distal from the membrane surface and\nthereby facilitating efficient RBD binding.\nOur experimental results also indicate that G-domain of maleimide-conjugated HRAS is freely\navailable for RBD binding. The measured K for RBD–RAS binding on the membrane surface\nd\n(Fig. 1e and Supplementary Fig. 9a,b) agrees well with K determined in solution (Ref. 52;\nd\nZheng et al. PNAS 2022, 119, e2119990119) (page 6, line 141).\nTo reflect this discussion, we have added the following sentence (page 5, line 104):\n“Maleimide-conjugated HRAS is expected to undergo rapid free rotational diffusion on\nmembranes, allowing ready access for effector protein binding, as previously observed in fully\nprocessed NRAS and KRAS48,49.”\n4. Maleimide conjugation is a non-physiological means of membrane tethering that does not\nreplicate the native prenylation of KRAS, which has been extensively utilized in the recent\npast. This limitation is not discussed, despite its potential impact on membrane interaction\nkinetics.\nWe chose maleimide conjugation here because the study aims to determine membrane kinetic\nmechanisms in a minimal system with precise control of HRAS surface density. Covalent\nattachment to maleimide lipids provides advantages that are difficult to achieve with prenylated\nKRAS under our flow-based assays: (i) stable tethering that withstands multiple wash/flow steps;\nand (ii) avoidance of variability from uncontrolled reversible farnesyl insertion/extraction, which\nwould otherwise couple RAS association/dissociation to the measured RAF kinetics and\nconfound rate estimates. This robustness enables unambiguous interpretation of relative kinetic\neffects, because all measurements were performed at a stable, well-defined RAS density.\nMoreover, recent SPR studies using KRAS robustly demonstrated that farnesylation (F) and\nmethylation (Me) do not measurably alter its affinity for CRAF RBD-CRD, and that the CRD\ninteracts with neither fully processed KRAS (KRAS-FMe) nor the fully processed hypervariable\nregion (HVR-FMe) (Ref. 35; Tran et al., Nat. Commun. 2021, 12:1176). Therefore, the absence\nof prenylation in our tethering strategy is unlikely to meaningfully affect RBD-CRD or CRD\nbinding to RAS, or to alter our conclusions.\n5. The manuscript omits critical methodological details: the final composition of protein\nstorage buffers, assay buffers, salt concentrations, assay temperatures, and precise protein\nconcentrations used in kinetic and diffusion experiments are not reported.\nWe appreciate the reviewer’s attention to these details and apologize for the omission of the final\nprotein storage buffer composition. Buffer A (20 mM HEPES, pH 7.3, 500 mM NaCl, 1 mM\nTCEP, and 10% glycerol) was used throughout the protein purification process, including the\n6\nfinal SEC step. The composition is described under the section Protein Purification in the\nMethods.\nWe identified that the assay buffer composition for the competitive FRET assay was missing and\napologize for this omission. The complete buffer composition has now been added. The\ncompositions of all other assay buffers were originally provided as abbreviations in the Methods,\nwith full definition given at their first mention. To improve clarity, we have now revised the\nmanuscript to present full buffer compositions—including salt concentrations—throughout,\nwithout using abbreviations.\nIn addition, the assay temperature is now specified alongside the full buffer compositions in the\nMethods.\nWe also found the omission of some protein concentrations. All protein concentrations have been\nspecified in figure captions. The availability of this information has been indicated in the\nMethods.\n6. There is no description or validation of the linker mutant or extended linker constructs. The\nauthors provide no data on their expression levels, solubility, or folding.\nGlycine-rich flexible linkers are widely used to connect two protein domains without interfering\nwith their individual structures and functions. (Chen et al., Adv Drug Deliv Rev 2013, 65, 1357-\n1369). The L136A/T178A mutant and short linker (GGGGGGS) and extended linker\n(4xGGGGGGS) variants were expressed at levels comparable to wild-type RBD-CRD and were\nstable under our assay conditions. Each protein eluted at the expected volume on SEC (calibrated\nwith protein standards, Supplementary Fig. 3) and was monomeric in the assay buffer (40 mM\nHEPES, pH 7.4, 150 mM NaCl, 5 mM MgCl ), as indicated by single-molecule fluorescence\n2\nintensity distributions (Supplementary Fig. 5). Additionally, we acquired CD spectra\n(Supplementary Fig. 24) for these constructs and found no evidence of folding variation among\nthem.\nTo include protein quality analysis and to reflect this discussion, the manuscript has been\nmodified as follows (page 12, line 289):\n“Protein quality analyses (Supplementary Fig. 3–5) and circular dichroism (Supplementary Fig.\n24) confirmed that all RBD-CRD linker variants were properly purified and folded. The\nreduced membrane binding observed for these constructs is therefore attributed to perturbations\nin the relative positioning between the RBD and CRD caused by the increased linker flexibility\nand length.”\n7. The rationale for the L136A/T178A double mutant is unclear. T178 is not part of the linker,\nand its selection is not justified. Similarly, the purpose/rationale of introducing a 24-residue\nflexible linker is not explained.\n7\n(1) Rationale for the L136A/T178A double mutant: This set of measurements (Figure 4) was\nperformed to identify structural elements that mediate positive cooperativity between the\nRBD and CRD. The mutations and linker variants were selected based on the X-ray structure\nof KRAS:RBD-CRD(CRAF) complex reported by Tran et al. (Ref. 35; Tran et al., Nat.\nCommun. 2021, 12:1176). In this structure, the RBD and CRD bind RAS through distinct\ninterfaces, forming a trimer (Figure 4a). The L136A/T178A double mutant was selected to\ndisrupt this assembly. To clarify the rationale for the selection of the L136A/T178A double\nmutant, we have revised the corresponding text as follows (page 11, line 273):\n“L136, located within the linker region, is buried within a hydrophobic pocket formed by\nF130 in the RBD and M183 in the CRD, bringing the two domains into close proximity35. The\nCRD residue T178 mediates contact with RAS through hydrogen bonding35. The L136A and\nT178A mutations are expected to disrupt the linker–RBD and CRD–RAS interactions,\nrespectively, which would destabilize the RAS:RBD-CRD trimer complex. A previous study\nreported that L136A reduced KRAS binding affinity fourfold and T178A decreased RAF\nactivity in cells35.”\n(2) Rationale for the linker variants: The flexible linker variants were designed to induce greater\nperturbations in the RBD-CRD arrangement, particularly in how the CRD can be positioned\n(or oriented) relative to the RBD. To clarify the rationale for introducing the short GGGGGS\nlinker, we have revised the corresponding text as follows (page 12, line 283):\n“This flexible linker [GGGGGS] is expected to eliminate residue-specific interactions while\nincreasing overall flexibility, which disrupts the proper orientation of the CRD.”\nTo clarify the rationale for introducing the extended 4×GGGGGS linker, we have revised the\ncorresponding text as follows (page 12, line 286):\n“To introduce a greater perturbation by increasing the physical separation and flexibility\nbetween the RBD and CRD, we characterized a construct containing an extended linker of\nfour GGGGGS repeats (4×GGGGGS; 24 amino acids total).”\n8. The claim that the T178A mutation stabilizes the extended linker construct is not supported\nby any biochemical data.\nIn the original manuscript, we were unable to express the RBD-CRD(4×GGGGGS) construct\ndespite multiple attempts. The open reading frame of the plasmid had been verified by Sanger\nsequencing, so the lack of expression was puzzling. During the revision, we performed whole-\nplasmid sequencing and found an unexpected insertion that had not been detected by earlier\nSanger sequencing. After re-cloning the plasmid, we were able to purify RBD-\nCRD(4×GGGGGS). The RBD-CRD(4×GGGGGS/T178A) construct was used out of necessity\nbecause we were initially unable to express the RBD-CRD(4×GGGGGS) construct. Now that the\nRBD-CRD(4×GGGGGS) construct has been successfully expressed, we have replaced the RBD-\nCRD(4×GGGGGS/T178A) data with RBD-CRD(4×GGGGGS) in Figure 4. The manuscript has\nbeen revised to reflect this change and provide adequate description as follows (page 12, line\n288):\n8\n“The extended linker constructs showed a greater reduction in membrane binding (cyan, Fig.\n4b). Protein quality analyses (Supplementary Fig. 3–5) and circular dichroism (Supplementary\nFig. 24) confirmed that all RBD-CRD linker variants were properly purified and folded. The\nreduced membrane binding observed for these constructs is therefore attributed to perturbations\nin the relative positioning between the RBD and CRD caused by the increased linker flexibility\nand length.”\nThe 4×GGGGGS construct occasionally exhibited a minor residual signal, about 5% higher than\nothers, in the normalized desorption curve (Fig. 4d). The absolute residual intensities were\ncomparable across constructs, suggesting that its lower equilibrium binding intensity may have\ncaused the apparent residual fraction after normalization. The dissociation rate of the\n4×GGGGGS construct was nonetheless faster than that of the other constructs.\nAdditionally, we found that RBD-CRD(4×GGGGGS) exhibits relatively fast equilibrium\nkinetics, making it difficult to reliably compare relative association rates using ensemble\nassociation assays. Therefore, we quantified k using single-molecule recruitment assays for the\non\nWT RBD-CRD, the L136A/T178A mutant, and the two linker variants. All k values remained\non\nunchanged, consistent with our previous conclusions. The manuscript has been revised to include\nthis new data in Figure 4c.\n9. The known ~2-fold higher affinity of RBD-CRD for RAS compared to RBD alone is not\nfactored into the interpretation of results. Many observations attributed to RBD-CRD vs RBD\nor CRD could reflect this baseline affinity difference.\nWe appreciate the Reviewer’s critical feedback. We agree that it is important to disentangle the\ncontributions of protein–protein and protein–lipid interactions to the high affinity of RBD-CRD\non RAS-functionalized membranes. To account for the intrinsically higher protein–protein\naffinity between RBD-CRD and RAS, we performed additional experiments measuring the K\nd\nvalues of RBD and RBD-CRD for RAS in the absence of membranes (PEGylated surfaces) and\ncompared them with the corresponding K values for membrane-tethered RAS.\nd\nBriefly, we observed that an additional protein contact between RAS and the CRD increases the\noverall affinity of RBD-CRD for RAS by approximately fivefold. The CRD–PS interaction\nfurther enhances the overall affinity of RBD-CRD for membrane-tethered RAS by approximately\nsevenfold. To reflect this new result, we have revised the main text describing the K\nd\nmeasurements and have added K values determined on lipid-free PEG surfaces as\nd\nSupplementary Fig 8 as follows (page 6, line 131):\n“To better understand how RAF integrates protein and lipid inputs for membrane binding, we\nsought to disentangle the contributions of protein interaction and lipid interaction to the\nenhanced affinity of RBD-CRD on RAS-functionalized membranes. To quantify the affinity\narising from solely from protein–protein interactions, the apparent dissociation constants (K ) of\nd\nRBD and RBD-CRD for RAS were determined on PEGylated (non-membrane) surfaces. An\nadditional protein contact between RAS and the CRD increases the overall affinity of RBD-CRD\nby approximately fivefold—220 ± 80 nM for RBD and 40 ± 10 nM for RBD-CRD (Supplementary\nFig. 8)—a trend similar to the twofold enhancement previously observed with KRAS35. For\nmembrane-tethered RAS, the RBD exhibited K that depended little upon PS content, measuring\nd\n270 ± 90 nM and 240 ± 20 nM at 2% and 20% PS, respectively (Fig. 1e and Supplementary Fig.\n9a,b). These values are consistent with our membrane-free measurements and previously\nreported affinities of the CRAF RBD for HRAS35,52. In contrast, RBD-CRD exhibited an affinity\nfor membrane-tethered RAS more than an order of magnitude higher than that of the RBD alone\n(K = 25 ± 10 nM and 6 ± 3 nM at 2% and 20% PS, respectively; Fig. 1f and Supplementary Fig.\nd\n9c,d). Removing RAS essentially abolished this high-affinity binding (Fig. 1f). The progressive\nincrease in affinity to RAS—from K = 220 nM for RBD (PEG surface), to 40 nM for RBD-CRD\nd\n(PEG surface), to 6 nM for RBD-CRD (20% PS membrane)—demonstrates that low-nanomolar\nmembrane affinity arises from the cooperative contributions of RAS–RBD binding, the additional\nRAS–CRD contact, and CRD–PS engagement.”\nAdditionally, we quantified the dissociation rate of RBD-CRD from RAS on lipid-free\nPEGylated surfaces and compared it with that measured for RAS tethered to 20% PS\nmembranes. We found that the lipid interaction is essential to achieve the prolonged membrane\nresidence observed in Figure 2c. To reflect this new result, we have revised the main text\ndescribing the dissociation kinetics (Fig. 2c) as follows (page 8, line 182):\n“Such a long dwell time cannot be achieved solely from protein–protein interactions. The RBD-\nCRD dissociates from RAS on a lipid-free PEGylated surface with a half-life of a few seconds\n(Supplementary Fig. 15a). This kinetic parameter analysis reveals that a crucial role for the\nCRD–PS interaction in selectively regulating membrane dissociation kinetics without affecting\nthe association rate.”\n10. The proposed enhancement of RBD–RAS association by CRD–lipid interaction is not\nclearly demonstrated or mechanistically explained.\nThe conclusion is derived from the competitive FRET assay shown in Figure 3. This assay\nmeasures energy transfer between fluorescently labeled RAS and RAF (RBD or RBD-CRD),\nthereby selectively reporting the dissociation kinetics of RAF from membrane-tethered RAS.\nAs such, the FRET decay curve reflects only the kinetics of the protein–protein dissociation.\nUsing this setup, we examined how this protein–protein (RAS–RAF) interaction is influenced by\nCRD–lipid engagement. A key conclusion arises from comparing the FRET dissociation curves\nat 2% and 20% PS membranes. The dissociation at 20% PS was slower than that at 2% PS,\nindicating that CRD–lipid interactions strengthen the protein–protein interaction. If the\nprotein-protein interaction were unaffected by CRD–lipid interaction, the FRET dissociation\nkinetics would be identical between 2% and 20% PS.\nThanks to the additional experiments measuring the dissociation kinetics of RBD-CRD from\nRAS on lipid-free PEGylated surfaces (related to Reviewer 1, Comment #9), we were able to\nidentify the contribution of the RAS-CRD interaction to the observed slow FRET dissociation\n10\nkinetics. The manuscript has been revised to clarify the role of the RAS-CRD interaction in the\nslower dissociation kinetics, as follows (page 10, line 250):\n“The slower dissociation for RBD-CRD at 2% PS is primarily due to an additional interaction\nbetween the CRD and RAS, as its FRET dissociation curve closely follows the ensemble\ndissociation curve of RBD-CRD measured on lipid-free PEG surfaces (Supplementary Fig. 15b).\nImportantly, 20% PS lipids, which promote a greater degree of CRD–lipid interaction, further\naccentuates this slowed dissociation for RBD-CRD (Fig. 3b). These findings indicate that CRD–\nlipid interactions enhance the binding strength between protein domains (RAS:RBD and\nRAS:CRD), thereby stabilizing the complex and slowing its dissociation.”\nThe potential mechanism underlying this behavior may involve reduced fluctuations of the\nRAS:RBD complex due to additional anchoring by CRD–lipid interactions, as suggested by MD\nsimulations (ref. 57; Structure 2018, 26, 513–525). This mechanism is described in the Result\n(page 10, line 237–239) and the Discussion (page 15, line 373–375).\n11. The claim that RAS·GTP mediates membrane recruitment of RBD-CRD species not bound\nto RAS is conceptually inconsistent, especially under conditions where all RAS binding sites\nare presumably occupied.\nWe found that the major population of lipid-engaged RBD-CRD species not bound to RAS (the\nRBD-CRD:PS complex) is primarily generated through the dissociation of RAS from the\nRAS:RBD-CRD:PS complex. Therefore, RAS·GTP indirectly mediates the membrane\nrecruitment of the RBD-CRD:PS species. This process is described in detail in the Results (page\n13, line 313–315; Fig. 5b).\nWe believe the confusion may have arisen because this observation was introduced too early—\nbefore the full explanation of the lateral rebinding mechanism that accounts for the formation of\nthe RAS-free RBD-CRD:PS species. To improve clarity, we have removed this earlier\ndescription on the formation of the RBD-CRD:PS species, this process is described in detail in\nthe later section RAS density modulates RAF membrane residence through lateral rebinding.\n12. The term “RAS unbinding” is used ambiguously throughout and should be clarified to\ndistinguish between RAS–RAF dissociation and membrane detachment.\nWe appreciate for the suggestion. The term “RAS unbinding” has been revised to indicate “RAS\ndissociation from the RAS:RBD:PS complex.” (page 13, line 313; page 14, line 336; page 15,\nline 376)\n13. The central conclusion regarding nanocluster-specific signaling is not experimentally\nsupported. The lipid compositions used do not promote RAS nanocluster formation, and the\nsystem lacks the complexity to model this behavior. Therefore, extrapolating these findings to\nnanocluster biology is not justified.\nremoved from the Abstract and Results. Additionally, we have substantially revised the\nDiscussion section to present this concept briefly as an unproven hypothesis, as follows (page 16,\nline 409):\n“It is worth mentioning that the average density of RAS on the plasma membrane is in the range\nof 30–150 RAS molecules per μm2 68. RAS is not uniformly distributed; instead, it forms\ndynamically exchanging nanoclusters that serve as the primary sites for recruitment and\nactivation of RAF32. The radius of a nanocluster is estimated to be 6–12 nm, with a local surface\ndensity of ~4000 to ~16,000 RAS per μm2 32,33,68. Such high local RAS densities within\nnanoclusters may facilitate RAF activation by extending its membrane residence time through\nlateral rebinding, although this hypothesis remains to be experimentally validated.”\nThe manuscript by Salinas et al. presents a well-thought-out and comprehensive kinetic\nanalysis of the RAF1 RBD-CRD domain interactions with HRAS and a simple supported lipid\nbilayer (SLB). The SLB is composed of DOPS, DOPC, and a functionalized DOPE that links\nHRAS to the bilayer via maleimide chemistry through an available cysteine in HRAS’s\nhypervariable region. I believe this study proposes a model that is both intriguing and valuable\nto the field, and that the supporting data and analysis are sound.\nIn brief, the authors propose that, following initial binding of RAF to RAS on the membrane,\nsubsequent lipid interactions involving the RAF CRD domain are sufficient to extend RAF’s\nresidence time. Nanoclustering of RAS and a CRD–lipid intermediate increases the likelihood\nof subsequent rebinding to other RAS monomers, enabling the full RAF activation\nsequence—including RAF dimerization—to proceed. This model has important implications\nfor understanding the RAF activation cycle, particularly the possibility of a kinetic\nproofreading function.\nTherefore, I recommend the manuscript for publication, provided the following points are\naddressed:\n1. The binding and dissociation rates of RAS/RAF complexes were calculated in a previous\nsingle-molecule pull-down study: Lee, H.W., Kyung, T., Yoo, J., et al. Nat Commun 4, 1505\n(2013). In this study, RAS was immobilized, and RAF molecules were pulled down from a\nlysate onto coverslips and imaged using TIRF microscopy. Residence times and kinetic\nparameters were measured. The authors should explicitly compare their data to this study.\nMoreover, as in Lee et al., they should account for potential photobleaching effects to ensure\nthat their measured rates are not significantly influenced by the photophysics of the\nfluorophores.\nWe appreciate the reviewer’s thoughtful suggestions. First, we would like to clarify that, unlike\nthe previous study by Lee et al. (Nat Commun 2013), pre-equilibrating encounter complexes\nwere not observed in our system (see also our response to Reviewer 2, Comment #2). Therefore,\nthe reported k values in the Nat Commun 2013 study and in the present work describe different\non\nprocesses and cannot be directly compared. In our measurements, the k for the RBD binding to\non\nRAS was estimated to be ~0.006 nM-1s-1 (Fig. 2b) and the k was ~1.4 s-1 (converted from a\noff\nhalf-life of ~1 s, Fig. 2c). From these results, the corresponding K = k /k was estimated to be\nd off on\n~230 nM, which agrees well with independently determined K values (270 nM on 2% PS and\nd\n240 nM on 20% PS membranes) obtained from titration experiments (Fig. 1e and Supplementary\nFig. 9a,b). This close agreement supports the validity of our kinetic rate estimates. The transient\nnature of RBD–RAS interaction observed in our study is also consistent with that reported by\nLee et al. (~300 ms), indicating that both systems exhibit dynamic, fast exchange kinetics.\nBecause the k values in the previous study and in our present study represent different\non\nprocesses, we chose not to compare them directly. To highlight the dynamic nature of RBD–RAS\ninteraction observed in both studies, we have revised the manuscript and provide the reference as\nfollows (Page 7, line 178, related to Fig. 2c):\n13\n“The RBD exhibited highly dynamic and transient interactions with a half-life of ~1 s, consistent\nwith a previous study using proteins pulled down from cell lysates53.”\nRegarding the potential effect of photobleaching on k we observed minimal impact. In our\noff\nstudy, k was determined using an ensemble desorption assay, in which fluorescence intensity\noff\ndecay was measured at specified time intervals rather than by continuous streaming acquisition.\nEach measurement involved the acquisition of fewer than 20 frames. To quantify the contribution\nof photobleaching to the observed intensity decay, we measured the photobleaching kinetics of\nmNG stably tethered to supported lipid bilayers. The laser was illuminated every 0.5 seconds\nusing the same laser power and exposure time as in the desorption assays. Only a minor\nphotobleaching effect was observed over 100 frames, with a 3% decrease in fluorescence at the\n20th frame, clearly demonstrating that photobleaching had little to no effect on our\nmeasurements.\nSupplementary Fig. 14: Photobleaching kinetics has a minimal impact on determining k .\noff\nTo reflect this discussion, the manuscript has been revised to include the photobleaching data as\nSupplementary Figure 14 and the following statement for describing Figure 2c (page 8, line\n181):\n“Photobleaching had a minimal impact on the estimation of k (Supplementary Fig. 14).”\noff\n2. Lee et al. describe the formation of “pre-equilibrating encounter complexes.” The authors\nshould clarify whether these results are consistent with or differ from those in the current\nmanuscript, especially with respect to the proposed lateral rebinding model. Although Lee et\nal. used immobilized RAS (i.e., non-diffusing and in a lipid-free system), it is possible that\ntheir findings contradict the current interpretation. Are there alternative explanations that\nsupport the proposed lateral rebinding and kinetic proofreading model?\nWe examined the possibility of pre-equilibrating encounter complex formation in our system by\nconducting an additional experiment based on a similar approach in Figure 2f of Lee et al. (Nat\nCommun 2013). In this experiment, we measured background fluorescence intensity as a\nfunction of membrane-bound RAS surface density, in the presence of 3 nM RBD-mNG.\nAccording to Lee et al., an increase in background signal at higher RAS densities would indicate\nformation of pre-equilibrating encounter complexes. We calculated background intensity by\n14\naveraging fluorescence signals from membrane regions excluding detectable single-molecule\nRBD spots. The 3 nM concentration was chosen to maintain low enough particle density for\nrobust particle exclusion during background analysis. Across RAS densities titrated up to ~20\nmolecules/μm2, we observed little to no change in background intensity (Response Fig. 1). The\ntrend closely followed control measurements using eGFP reported by Lee et al., which does not\ninteract with RAS. This result suggests the absence—or minimal presence—of pre-equilibrating\nencounter complexes in our system. Thus, we conclude that such complexes, if they exist, do not\nsignificantly contribute to RAF membrane binding dynamics under our experimental conditions.\nThus, the observed RAS density-dependent modulation of RAF dwell time is best explained by\nthe lateral rebinding mechanism in which RAF forms a transient RBD-CRD:lipid intermediate\non membrane surface (Fig. 5b). This is supported by additional data (see also our response to\nReviewer 2, Comment #3), showing that disruption of CRD–lipid interactions by mutations\nreduces the RAS density dependence of RAF dissociation kinetics (Supplementary Fig. 25c).\nResponse Figure 1: Normalized background fluorescence in the presence of fluorescent RBD.\nData from RAS-functionalized lipid membranes (open red circles, this study) are compared with\nthose from RAS-functionalized PEGylated surfaces (solid symbols, Lee et al. Nat. Commun.\n2013). No change in background fluorescence was detected on RAS-functionalized lipid\nmembranes, indicating the absence of pre-equilibrating encounter complex formation under our\nexperimental conditions.\n3. The authors cite a recent study using KRAS4b on SLBs, which analyzes KRAS4b/RBD-\nCRD complexes on simple and complex lipid bilayers: Shrestha R., Carpenter T.S., Van Q.N.,\net al. Commun Biol. 2024; 7(1):242. The authors should comment on the specific CRD-lipid\ncontact points identified by the simulations and NMR in Shrestha et al. and evaluate\nexperimentally whether mutations of these lipid-penetrating CRD residues affect lateral\nrebinding interactions in the RBD-CRD complex or the proposed RBD-CRD–PS intermediate.\nWe thank the reviewer for the thoughtful suggestion. Based on the study by Shrestha et al.\n(Commun Biol 2024), we generated an RBD-CRD construct containing tetra-alanine mutations\n(K148A, L149A, K157A, and F158A; hereafter referred to as 4A). We characterized membrane\nbinding, diffusion, and desorption kinetics for the 4A construct. To reflect this new result, the\nmanuscript has now been revised to include the data as Supplementary Fig. 25 and the following\ntext (Page 14, line 339):\n15\n“Positively charged and hydrophobic residues in loops 1 and 2 of the CRD penetrate the bilayer\nand mediate membrane binding55. To test how these specific CRD–lipid interactions contribute to\nlateral rebinding, we characterized an RBD-CRD4ᴬ construct containing four alanine mutations\n(K148A, L149A, K157A, and F158A; hereafter referred to as 4A). The 4A construct exhibited a\ndiffusion behavior similar to that of the isolated RBD, and its membrane binding was\nsubstantially reduced compared to WT RBD-CRD (Supplementary Fig. 25a,b). Notably, the\nextension of membrane dwell time at higher RAS densities was significantly attenuated,\nconfirming the critical role of CRD–lipid interactions in lateral rebinding and in the formation of\nthe RBD-CRD:PS intermediate (Supplementary Fig. 25c).”\n4. In the final sentence of paragraph 1 in the section titled “RAS density modulates RAF\nmembrane residence through lateral rebinding,” the authors state: “we exclude the scenario\nwhere RBD-CRD first loses lipid contact yet remains bound to RAS, as a lipid-free RAS:RBD-\nCRD complex was not observed in single-particle tracking experiments (Fig. 2e).” To more\nrobustly support this claim, the authors should measure the diffusion of fluorescently labeled\nRAS in the presence of RBD-CRD on the same SLB and compare those diffusion rates to the\nmeasurements where the fluorescent tag is on RBD-CRD. Comparing the distributions and\nfractional occupancies of the different diffusion states in both labeling conditions would\nprovide a stronger test of the assertion that lipid-free RAS:RBD-CRD complexes do not form\nor persist. I would also encourage a different term than “lipid-free.” We know that there are\nextensive and dynamic contacts between RAS and lipids. There is probably more precise\nlanguage to talk about the conformational state of the complex and contact with the lipids in\nthe bilayer.\nWe appreciate the reviewer’s suggestion. However, we believe the proposed experiment would\nnot directly address the presence or absence of RAS:RBD-CRD complexes that lack CRD–lipid\nengagement (hereafter referred to as CRD-disengaged RAS:RBD-CRD complexes). The main\ncomplication originates from interpretating the diffusion behavior of fluorescently labeled RAS*\n(the asterisk denotes a fluorescent label). Based on our findings, CRD-disengaged RAS*:RBD-\nCRD complexes are expected to diffuse similarly to free RAS*, as the diffusion of free RAS*\nand RAS:RBD* are nearly identical (Figure 2e). Therefore, it would be difficult to distinguish\nwhether fast-diffusing RAS* corresponds to free RAS or CRD-disengaged RAS:RBD-CRD\ncomplexes. Diffusion analysis of labeled RBD-CRD* on membranes provides a more direct\nmean for detecting CRD-lipid engagement. In our experiment, we did not observe a fast-\ndiffusing population of RBD-CRD* (Figure 2e), suggesting the majority of membrane-bound\nRBD-CRD remains engaged with lipids.\nTo revise a term “lipid-free”, the corresponding sentence has been modified as follow (page 13,\nline 317):\n“We exclude the scenario where RBD-CRD first loses lipid contact yet remains bound to RAS, as\na RAS:RBD-CRD complex with the CRD disengaged from lipids was not observed in single-\nparticle tracking experiments (Fig. 2e).”\nThis is an interesting and potentially impactful article that would be suitable for publication in\na major journal such as Nature Communications, following adequate attention to significant\nshortcomings. The article summarizes well-reasoned biophysical studies of the multi-pronged\ninteractions between the important signaling proteins CRaf, HRas and anionic lipids\n(phosphatidylserine or PS) during CRaf membrane binding, 2D-diffusion and dissociation on\na lipid bilayer possessing anchored HRas molecules. The quite promising results suggest that,\nas often observed for signaling proteins that target the plasma membrane, the CRaf molecule\npossesses cooperative, multi-component binding contacts involving both protein-lipid and\nprotein protein interactions that together provide adequate specificity and affinity for the\ncorrect membrane surface. In CRaf this multi-component interaction is provided by its distinct\ncysteine-rich domain (CRD) and Ras binding domain (RBD) that provide protein-lipid (CRD-\nPS) and protein-protein (RBD-HRas), respectively. While these results are not surprising,\nthis manuscript is the first to address the cooperative nature of these essential Ras:CRaf:PS\ninteractions in a quantitative, biophysical manner. The following major and minor points\nshould be addressed prior to publication.\n1. The article places a major emphasis on Raf targeting, retention, and activation in Ras\nnanoclusters with language about relevance to nanoclusters on 10 pages of the text including\nthe Abstract, Introduction, Results, Discussion, and Figure Legends. However, as the article\nitself notes, the system employed observes individual Ras:RBD-CRD complexes interacting\nwith PS lipids on the target bilayer with no detectable clusters (pg 8, bottom half of\nparagraph). Moreover, it appears that Ras densities and clustering are not measured in most\nor all experiments. Thus the article should be more careful about its claims of relevance to\nclustered systems, and should limit those claims to the description of the plausible, but as yet\nunproven, model in the Discussion.\nWe appreciate the reviewer’s critical feedback. The description of RAS nanoclusters has been\nremoved from the Abstract and Results. Additionally, we have substantially revised the\nDiscussion section to present this concept briefly as an unproven hypothesis, as follows (page 16,\nline 409):\n“It is worth mentioning that the average density of RAS on the plasma membrane is in the range\nof 30–150 RAS molecules per μm2 68. RAS is not uniformly distributed; instead, it forms\ndynamically exchanging nanoclusters that serve as the primary sites for recruitment and\nactivation of RAF32. The radius of a nanocluster is estimated to be 6–12 nm, with a local surface\ndensity of ~4000 to ~16,000 RAS per μm2 32,33,68. Such high local RAS densities within\nnanoclusters may facilitate RAF activation by extending its membrane residence time through\nlateral rebinding, although this hypothesis remains to be experimentally validated.”\nRAS densities were quantified for all experiments. They are presented in either figures or figure\ncaptions.\n17\n2. The experiments are well chosen and the data is promising, but there is inadequate\nattention to reproducibility which could lead to questions about the rigor of the work. In\ngeneral, statistical tests of significance are absent and there is information on replicates for\nonly two of the many experiments (in both cases N=3, presumably representing technical\nrather than biological replicates). Perhaps I missed it, but I could find no indication that\nexperiments were carried out more than once, and were repeated on multiple days. Such lack\nof information about replicates and statistical tests makes it impossible to judge the\nreproducibility and statistical significance of the findings.\nWe thank the reviewer for directing our attention to the significance of statistical rigor and\nreproducibility. We apologies the omission of this information. We have now revised manuscript\nto report the number of total technical replicates (n = SLBs in total) and independent experiments\n(N) in figure captions, with the addition of replicate experiments. In some cases—particularly\nthose dependent on RAS surface density—performing identical replications on different days\nwas challenging because the resulting RAS densities often varied. However, all RAS densities\nwere quantified, and in such cases we clearly indicated in the figure captions that similar results\nwere obtained in independent experiments, with references to the corresponding data in the\nSupplementary Information.\nThe experiments are highly reproducible, largely due to our ability to directly measure the\nabsolute surface density of proteins—including RAS—using a TIRF-FCS calibration standard.\nThis allows quantitative reconstitution of well-defined protein-functionalized membranes.\nAdditionally, statistical tests have been performed for k measurements in Figure 2b (p values in\non\nSupplementary Table 1) and, as well as ensemble binding measurements in Supplementary Fig.\n10.\n3. The slow diffusion observed for Ras:RBD-CRD:PS complexes on the DO-lipid bilayer\nraises the possibility that, in addition to the frictional drag of the Ras anchor lipid and the\nbound PS molecules against the bilayer, there may well be additional frictional drag provided\nby protein insertion into the bilayer. See PMID 23701821.\nWe fully agree with this interpretation. The possibility of membrane insertion by CRD loop 1\nand 2 is described in the Results (page 9, line 217–220).\nTo further highlight this point, we have now included the suggested reference (added to Ref. 59;\nZiemba et al. Chem Phys Lipids 2013) and the following text (page13, line 319):\n“The slow diffusion coefficient of RBD-CRD is comparable to that of PKC regulatory domains\nknown to penetrate into the hydrocarbon core of the bilayer59, indicating persistent lipid\nengagement of the CRD through its basic and hydrophobic residues in loop1 and loop 216,55.”\n4. The article is missing key references to directly relevant single molecule studies of Ras-\n18\neffector interactions, and of the effects of membrane environment on the affinities and kinetics\nof protein-protein and protein-membrane interactions. See, for example, PMID 29211993,\n39217418, 22263647.\nWe appreciate for providing important references relevant to our work. We have now added the\nfollowing three references in the main text (page 5, line 102):\nRef. 45: Buckles et al., Biophys J 2017, 113, 2396-2405\nRef. 46: McCombs et al., Biophys J 2024, 123, 3277-3280\nRef. 47: Buckles et al., Biophys J 2020, 229, 2305-2218\n“This widely used lipidation method preserves the biological function and structure of RAS and\nhas been applied to all three RAS isoforms (H-, N-, and KRAS) and other membrane-anchored\nsmall GTPases40–47.”\nThis manuscript provides interesting results and confirms several observations from previous\nexperimental and computational studies. I recommend that authors stress the novelty of their\nfindings in the context of broader Ras biology during the revision of this manuscript.\nWe appreciate the reviewer’s constructive feedback. To emphasize the novelty and significance\nof our study in the context of broader RAS biology, we have revised our manuscript to include\nthe following text in the Introduction section (page 3, line 64):\n“Prior studies reported cooperativity between RBD and CRD in which RAS–RBD binding\ninduces a transient interaction between RAS and CRD, and mapped CRD–lipid interfaces16.\nHowever, how these interactions control membrane kinetics has remained unclear. A quantitative\nunderstanding of membrane-binding kinetics and the underlying mechanisms is crucial because\nRAF is exclusively activated on cellular membranes14. RAF is also among the least abundant\nsignaling proteins in many cancer cell types36. Upon receptor activation, only a small number of\nRAF molecules—estimated to be as few as 100–200 per cell—are recruited to the membrane,\nwhere they serve as a stoichiometric bottleneck that determines the overall signaling outcome37.\nTherefore, understanding RAF membrane dynamics is fundamental for elucidating how normal\nand oncogenic RAS activities are transduced into ERK signaling38.”\nThe manuscript reports that RBD binds to RAS, allowing CRD to bind to negatively charged\nPS lipids (as determined by TIRF microscopy). They propose two kinetically distinct binding\nsteps. Also, CRD binds to PS in a RAS-GTP-dependent manner. It ensures RAF membrane\nbinding (by CRD) is strictly driven by RAS activation (by RBD). Using FRET, they\ndemonstrate that CRD-lipid binding enhances the association between RBD and RAS (as\nmeasured by FRET). RBD and CRD binding to RAS and PS lipids exhibit cooperativity. RAF\nundergoes lateral rebinding with RAS (using kinetic simulations in SimBiology and TIRF\nmicroscopy). Weak CRD-lipid interactions play a role in lateral rebinding. This extends the\nRAF's membrane dwell time under high RAS density (e.g., nanoclusters). This prolonged\nmembrane residence increases the likelihood of completing RAF's multistep activation\nsequence (i.e., facilitates kinetic proofreading of RAF activation).\nComments:\n1. This work would have had a significant impact if the authors had investigated the kinetics\nin the presence of PIP2 lipid and distinguished the binding mechanisms.\nLipid specificity of RAF kinases is an important question. PIP2 is involved in upstream signaling\nevents such as RAS activation by SOS. Several molecular dynamics simulations and\nexperimental studies suggest that the CRD engages with PIP2 (Johnson et al., Biochemistry\n2005, 44(9), 3432–3440; Carpenter et al., Biophys. J. 2026, PMID: 40849684). Our lab is\nactively investigating this topic. Preliminary experiments comparing RBD-CRD binding to\nsupported lipid bilayers containing 5% PIP2 versus 20% PS (both carry the same –20 net charge)\nin the absence of RAS show largely similar association behavior (Revision Fig. 2). While we\nagree that this is a significant question, we believe it requires a more detailed investigation in the\npresence of RAS, which we plan to focus on in a separate study.\nmembranes containing either 20% DOPS or 5% PIP2. Both conditions show similar levels of\nmembrane binding. Data represent means ± standard deviations from three technical replicates (n\n= 3 SLBs).\n2. Additionally, DOPC is not a particularly abundant component of the plasma membrane.\nAny reason for its selection? (CRD is more likely to bind to POPC:POPS membranes than\nDOPC:DOPS)\nWe thank the reviewer for this comment. We selected DO-lipids based on prior success in\ngenerating high-quality supported lipid bilayers. We have recently begun using PO-lipids in our\nexperiments. Our latest measurements indicate that RBD-CRD binds to POPC:POPS membranes\nwith comparable (or slightly lower) affinity with respect to DOPC:DOPS membranes (Revision\nFig. 3). However, this small to modest difference does not affect the main conclusions of this\nstudy.\nRevision Figure 3. Equilibrium surface densities of 100 nM RBD-CRD on RAS-free\nmembranes composed of either DOPC:DOPS or POPC:POPS. Both membranes contain 20% PS.\nData represent means ± standard deviations from three technical replicates (n = 3 SLBs).\n21\n3. Also, is a 20% PS concentration sufficient to mimic biomembranes?\nWe believe that 20% PS is sufficiently close to physiological levels for studying RAF–PS\ninteractions. According to recent lipidomics data from Lorent et al. (Nat. Chem. Biol. 2020, 16,\n644–652), PS constitutes approximately 26% of the inner leaflet lipids in erythrocyte plasma\nmembranes. Currently, 20% PS is a technical upper limit we can reliably incorporate while\nmaintaining high-quality supported lipid bilayers and efficient protein coupling, although further\noptimization might be possible.\nTo provide this information, we have revised the manuscript to include the following sentence\nand cite the study by Lorent et al. (page 4, line 98):\n“A 20% PS composition was used to closely approximate the physiological fraction (~26%) of\nPS in the inner leaflet of plasma membranes while maintaining high-quality bilayers39.”\n4. What would be the effect of having the kinase domain? Past experiments were based on the\nsame RBD-CRD construct. Engaging the kinase domain and seeing the effect of\nhomodimerization can be quite novel.\nWe strategically designed this study to dissect complex interactions between RAF and\nmembranes with the conserved regulatory module (RBD-CRD) in a minimal, well-controlled\nsetting. We plan to extend this work to investigate the membrane dynamics of full-length RAF.\nBased on our observations, we expect that kinase domain dimerization would further extend\nmembrane dwell time, as increased multivalency (two RBD-CRD modules) could enhance the\nlateral rebinding process, which in turn increases sensitivity to RAS density. This hypothesis\nrequires experimental validation with full-length proteins.\nWe provided our planned future studies in the final paragraph of the Discussion section as\nfollows (page 17, line 418):\n“To investigate how membrane-binding kinetics influence activation outcomes under more\nphysiologically relevant conditions, it will be necessary to use the autoinhibited full-length\nRAF:14-3-3:MEK complex. These advancements will enable us to rigorously recapitulate the\nsequential steps of RAF activation and elucidate the underlying molecular mechanisms and\nkinetics with high resolution.”\n5. I find it challenging to convince myself that RAF unbinding occurs primarily by detaching\nfrom RAS first, then from the membrane, rather than the other way around (end of first\nparagraph on p. 12). Strong arguments to support their claim can be useful.\nWe identified the primary membrane detachment pathway of RAF as sequential dissociation\nfrom RAS followed by unbinding from lipids (hereafter referred to as Scenario A). This\nconclusion is supported by RAF diffusion analysis. The RBD, which is lacking the lipid-\ninteracting CRD, diffuses at ~1 m2/s when bound to RAS. However, this fast-diffusing species\n22\nwas not observed for RBD-CRD:RAS complexes. Instead, these complexes exhibit much slower\ndiffusion (~0.3 m2/s), indicating persistent lipid engagement. This slow diffusion coefficient is\ncomparable to that of PKC regulatory domains known to penetrate into the hydrocarbon core of\nthe bilayer (Ref. 59). Indeed, NMR and computational studies have shown that loop 1 and 2 of\nthe CRD insert into the membrane (Ref. 16,55). The absence of the fast-diffusing population\nsuggests that RBD-CRD persistently engages lipids via the CRD as long as it remains\nmembrane-bound, making Scenario A favorable.\nFor detachment from RAS to be the final step in RAF membrane dissociation (hereafter referred\nto as Scenario B), all RBD-CRD:PS intermediates would need to laterally rebind to RAS, which\nis unlikely to occur. Scenario B also requires the CRD to disengage from lipids before RAS\nunbinding. However, our single-molecule tracking data (Supplementary Fig. 20) show that when\nRBD binds to RAS, the CRD engages lipids rapidly, within the 20 ms temporal resolution of our\nassay; which is much shorter than the dwell time of RBD:RAS (0.3 to 1 second, according to the\npresent study and previous study by Lee et al. reported in Ref. 55 Nat Commun 2013, 4, 1505).\nThis result suggests that if the CRD disengages from lipids, it is likely to reengage immediately\nas long as RBD-CRD is bound to RAS, making Scenario B unfavorable.\nTo provide a clearer rationale for selecting the primary RAF membrane dissociation pathway, we\nhave now included the above discussion in the main text as follows (page 13, line 319):\n“The slow diffusion coefficient of RBD-CRD is comparable to that of PKC regulatory domains\nknown to penetrate into the hydrocarbon core of the bilayer59, indicating persistent lipid\nengagement of the CRD through its basic and hydrophobic residues in loop1 and loop 216,55.\nEven if the CRD momentarily dissociates from lipids, it is likely to re-engage rapidly while still\nbound to RAS, given the fast lipid engagement timescale observed in our single-molecule\ntracking analysis (Supplementary Fig. 20).”\n6. Several xray structures clearly show that CRD creates contacts with RAS. I agree that these\ncontacts may not be formed in the absence of RBD. Therefore, lipids are not the only driving\nforce for membrane interaction. Claiming that CRD is driven only by lipids is not completely\ncorrect.\nWe appreciate the reviewer’s constructive feedback. We performed additional experiments to\nmeasure the K values for RBD and RBD-CRD for RAS immobilized on lipid-free PEGylated\nd\nsurfaces. These experiments allowed us to quantitatively assess how the affinity of RBD-CRD\nincreases progressively with the addition of each binding contact—RAS–RBD, RAS–CRD, and\nCRD–lipid interactions.\nThe main text describing Figure 2 has been revised to incorporate these new results. The\ncontribution of the CRD–RAS interaction to the overall membrane binding affinity is now\nexplicitly stated as follows (page 6, line 146):\n“The progressive increase in affinity to RAS—from K = 220 nM for RBD (PEG surface), to 40\nd\nnM for RBD-CRD (PEG surface), to 6 nM for RBD-CRD (20% PS membrane)—demonstrates\n23\nthat low-nanomolar membrane affinity arises from the cooperative contributions of RAS–RBD\nbinding, the additional RAS–CRD contact, and CRD–PS engagement”\n7. It is interesting that RBD alone has a higher k to RAS-associated membranes than RBD-\non\nCRD; however, the presence of CRD lowers the off rate, resulting in RBD-CRD having a\nstronger binding affinity than RBD alone. Additionally, the rate constants for RBD alone and\nRBD-CRD are not affected by PS concentration, so only k is influenced by PS levels. I don’t\noff\nquite understand that easily. Some clarity would be helpful.\nWe also found these results are interesting and unexpected. The observed trend is reproducible\nacross multiple independent experiments (Fig. 2). First, we would like to clarify that the k\non\nvalues were measured by a single-molecule recruitment assay. In this assay, the time resolution is\ninherently set by the camera frame rate (20 ms) as described in the Methods section; this\ninformation has now been added to the Figure 2 caption as well. CRD–lipid interactions alone\nare highly transient and rarely result in detectable binding events at our current time resolution.\nWe occasionally observed faint, short-lived diffusive traces of RBD-CRD on 20% PS membrane\nin the absence of RAS (see figure 2a for RBD-CRD, (-) RAS). However, these events are too\ntransient to be reliably detected by the particle localization algorithm. Based on rough estimates,\nthe dwell time of these interactions is likely on the order of a few to several milliseconds; even\nshorter-lived species may exist but are not visible under our experimental conditions\n(Supplementary Fig. 12).\nThe key observation is that these transient interactions between RBD-CRD and lipids do not lead\nto productive RAS engagement (clearly spatially resolved binding events). If they did, we would\nexpect k to increase at a higher PS concentration, which was not observed. The highly transient\non\nnature of PS-recruited RBD-CRD and its inability to engage RAS prevent it from accumulating\non the membrane. Therefore, PS does not contribute to the initial membrane recruitment of RAF.\nTo reflect this discussion, the manuscript has been modified to the following text in the Results\n(page 7, line 164):\n“In line with our ensemble binding measurements, clearly spatially resolved membrane\nrecruitment events required the interaction with RAS (Fig. 2a). Interestingly, we observed highly\ntransient interactions of RBD-CRD (appearing as faint, diffusive traces) on RAS-free 20% PS\nmembranes. Although those particles could not be localized, manual intensity analysis estimated\ntheir dwell times on the order of several milliseconds (Supplementary Fig. 12). Importantly,\nthese short-lived interactions between RBD-CRD and lipids do not result in productive RAS\nengagement, as k values for both RBD and RBD-CRD were unaffected by PS concentration\non\n(Fig. 2b).”\nWe were also surprised that such a weak CRD–lipid interaction has a large impact on the\ndissociation rate (Fig. 2b). This arises from the combined effect of multiple factors: (1)\npromotion of CRD–PS engagement upon RAS–RBD binding (Fig. 1d,f), (2) stabilization of the\nRAS:RBD-CRD complex by PS engagement (Fig. 3), and (3) lateral rebinding (Fig. 5). A more\ndetailed discussion of this point is provided in the Discussion (page 15, line 365-380).\non\nimpose partial steric hindrance on RBD access to RAS. Although it is a plausible explanation,\nwe do not have direct evidence.\n8. I find it puzzling that the RBD-CRD linker does not affect the initial RBD-RAS association\nbut is relevant for the membrane dissociation rate, and thus for the cooperative binding of\nRBD-CRD to RAS and PS lipids.\nThis point is closely related to our response to Reviewer 4, Comment #7. RAF membrane\ninteractions involve three different molecular contacts: (1) RBD–RAS interaction, (2) CRD–\nRAS interaction, and (3) CRD–lipid (PS) interaction. The CRD-mediated interactions (2) and (3)\nare dependent of RBD–RAS interaction (1) (Fig. 1g). Therefore, interactions (2) and (3) play no\nrole in the initial membrane recruitment event. Instead, k is primarily governed by the RBD–\non\nRAS interaction (1) (Fig 2a,b).\nIncreasing the flexibility or length of the RBD-CRD linker likely alters the orientation and\nposition of the CRD relative to RAS and the membrane surface (Ref. 35 and 58), thereby\nreducing the efficiency of both CRD–RAS and CRD–lipid interactions. Because these CRD-\nmediated interactions (2) and (3) primarily contribute to membrane stabilization and lateral\nrebinding rather than initial recruitment, their disruption selectively affects the membrane\ndissociation rate (𝑘 ), without altering the initial RBD–RAS association.\noff\nTo clarify the role of the linker, the manuscript has been revised to include the following text in\nthe Result section (page 12, line 296):\n“Thus, the CRD-mediated interactions (CRD–RAS and CRD–lipid) primarily contribute to\nmembrane stabilization rather than to the initial association between RBD and RAS, consistent\nwith the K and K analyses shown in Figure 2.”\non off\nIn this study, the authors use quantitative TIRF microscopy to analyze the binding kinetics\nand equilibrium behavior of two RAF domains (RBD and CRD). Particularly, they looked at\nRBD/CRD interactions with phosphatidylserine lipids, RAS, and cooperative binding\ninteractions with each other. The purpose of this study was to develop a model for membrane-\ndependent RAF activation, which is dependent on recruitment to the membrane by GTP-RAS,\nand retention on the membrane through weak PS interactions which cooperatively strengthen\nbinding to RAS. The carefully designed experiments elucidate KD values, koff rates, and kon\nrates to elucidate the impact of each component in the system. The RAF-MEK-ERK signaling\ncascade is a classical biochemistry interaction that is fundamentally important to numerous\ncell signaling events. Understanding activation of this pathway is critical to being able to\nregulate cell signaling in cancer cells. This work will be of high interest to a number of fields\nbecause of the extreme importance of RAF activation in cancer.\nOverall the conclusions of the study are supported by the results. The experiments are well-\ndesigned, and the interpretations are logical. There are some minor points where the language\nand reporting in the manuscript could be clarified, as noted below. The manuscript is well\nwritten and the figures are generally of good production quality. The biggest concern to this\nreviewer is there may be an over-extrapolation of these results to connections concerning RAS\nnanoclusters, RAF auto-inhibition, and involvement of the SHOC2-MRAS-PP1C complex. I\nfeel that these results provide a convincing argument about RBD/CRD cooperativity,\ninteractions with PS lipids, and importance of RAS concentration. However, the attempt to\nexplain a full mechanism of RAF activation seems to be speculation and extrapolation based\non other published works. I recommend the manuscript undergo minor revisions to address\nthe points listed below, where the issue of over-extrapolation is elaborated on in the last point.\nPoints to address:\n1. Some additional clarifications could be added to the introduction to aid readers who are not\nin this field:\n(1) It was not immediately obvious to me that “14-3-3” is the name of a protein. Please add a\n(very) brief explanation. Even just add the word “protein” after it the first time it\nappears.\nWe have changed “14-3-3 dimer” to “dimeric 14-3-3 protein” when it first appears in the\nIntroduction (page 2, line 44).\n(2) Please briefly explain what are BRAF, KRAS, and CRAF, compared to just RAF and\nRAS.\nWe have now listed all three RAF isoforms in the Introduction as follows (page 2, line 26):\n“RAF isoforms (A-, B-, and CRAF) share a conserved domain organization: an N-terminal\nregulatory region composed of the RAS-binding domain (RBD) and the cysteine-rich\ndomain (CRD), and a C-terminal kinase domain (Fig. 1a).”\n“This widely used lipidation method preserves the biological function and structure of RAS\nand has been applied to all three RAS isoforms (H-, N-, and KRAS) and other membrane-\nanchored small GTPases40–47.”\n2. Evaluating how the RBD-CRD had measured KD values of 14 nM vs 5 nM for binding 2%\nand 20% PS membranes as a “evident dependence on PS” seemed strange in contrast to the\nprevious comparison of 222 nM vs 163 nM as “depended little on upon PS content”. Please\nadjust the language or clarify.\nWe appreciate the reviewer’s detailed attention to our work. In the original submission, we\nevaluated the differences based on the mean values and 95% confidence intervals. The K values\nd\nof RBD binding to RAS on 2% and 20% PS membranes largely overlapped due to wide\nconfidence intervals, whereas those for RBD-CRD did not. During the revision, we performed\nadditional experiments and now report mean values and standard deviations from three\nindependent replicates. Incorporating these new data, the K of RBD was determined to be 270 ±\nd\n90 nM and 240 ± 20 nM on 2% and 20% PS membranes, respectively, showing little dependence\non PS. The K for RBD-CRD was 25 ± 10 nM and 6 ± 3 nM on RAS-functionalized membranes\nd\ncontaining 2% and 20% PS, respectively. The revised manuscript no longer directly compares the\n2% and 20% PS data; instead, it focuses on comparing K values between lipid-free surfaces and\nd\n20% PS membranes. Therefore, we believe this issue has been fully addressed.\n3. Was there a 0% PS control performed for any of these samples? Why were 2% and 20% PS\nconcentrations chosen for the experiments? Please explain the lipid choice briefly.\nWe selected 20% PS as a physiologically relevant level, based on lipidomics showing ~26% PS\nin the erythrocyte inner leaflet plasma membrane (Lorent et al. Nat. Chem. Biol. 2020, 16, 644–\n652). 20% is the current technical upper limit for incorporating PS while maintaining high\nbilayer quality and efficient protein coupling; further optimization may increase this threshold.\nFor a lower PS membrane, we selected 2% because complete removal of PS (0% PS) resulted in\na higher frequency of membrane defects.\nTo provide this information, we have revised the manuscript to include the following sentence\nand cite the study by Lorent et al. (page 4, line 98):\n“A 20% PS composition was used to closely approximate the physiological fraction (~26%) of\nPS in the inner leaflet of plasma membranes while maintaining high-quality bilayers39. ”\n4. The number of reported significant figures should be reduced for reported KD values in\ncaption for Figure 1. Example: 222 +/- 88 should be changed to 220 +/- 90\nreduce significant figures for all reported K values.\nd\n5. Figure 2a, middle panel: I think I’m seeing some low-intensity spots indicating protein\nbinding in the -RAS condition. However, I can’t rationalize how you can have less than one\nmolecule showing up, unless this is a product of partial occupancy over the duration of a\ngiven exposure time? Please explain.\nWe agree with reviewer’s interpretation. In the (–) RAS condition for RBD-CRD, the faint,\nlow-intensity spots correspond to RBD-CRD-AZ647 recruited by PS interaction alone. These spots\ndwell much shorter than a single exposure (20 ms). The integrated signal is therefore\nproportionally reduced, which explains the appearance of “less than one molecule.”\nTo explain this observation with an appropriate context, the manuscript has been revised to\ninclude the following text in the Results (page 7, line 164):\n“In line with our ensemble binding measurements, clearly spatially resolved membrane\nrecruitment events required the interaction with RAS (Fig. 2a). Interestingly, we observed highly\ntransient interactions of RBD-CRD (appearing as faint, diffusive traces) on RAS-free 20% PS\nmembranes. Although those particles could not be localized, manual intensity analysis estimated\ntheir dwell times on the order of several milliseconds (Supplementary Fig. 12). Importantly,\nthese short-lived interactions between RBD-CRD and lipids do not result in productive RAS\nengagement, as k values for both RBD and RBD-CRD were unaffected by PS concentration\non\n(Fig. 2b).”\n6. It is odd that the experiment wasn’t successful for the 4XGGGGGS linker construct, but it\nwas successful for the 4XGGGGGS/T178A construct. Can the authors please provide their\nthoughts on this? What part of the experiment was unsuccessful? Could the protein not be\nexpressed? Perhaps this doesn’t need to be explained in the manuscript, but I think it is\nappropriate for discussion during the review.\nIn the original manuscript, we were unable to express the RBD-CRD(4×GGGGGS) construct\ndespite multiple attempts. The open reading frame of the plasmid had been verified by Sanger\nsequencing, so the lack of expression was puzzling. During the revision, we performed whole-\nplasmid sequencing and found an unexpected insertion that had not been detected by earlier\nSanger sequencing. After re-cloning the plasmid, we were able to purify RBD-\nCRD(4×GGGGGS). The RBD-CRD(4×GGGGGS/T178A) construct was used out of necessity\nbecause we were initially unable to express the RBD-CRD(4×GGGGGS) construct. Now that the\nRBD-CRD(4×GGGGGS) construct has been successfully expressed, we have replaced the RBD-\nCRD(4×GGGGGS/T178A) data with RBD-CRD(4×GGGGGS) in Figure 4. The manuscript has\nbeen revised to reflect this change and provide adequate description as follows (page 12, line\n288):\n28\n“The extended linker constructs showed a greater reduction in membrane binding (cyan, Fig.\n4b). Protein quality analyses (Supplementary Fig. 3–5) and circular dichroism (Supplementary\nFig. 24) confirmed that all RBD-CRD linker variants were properly purified and folded. The\nreduced membrane binding observed for these constructs is therefore attributed to perturbations\nin the relative positioning between the RBD and CRD caused by the increased linker flexibility\nand length.”\nThe 4×GGGGGS construct occasionally exhibited a minor residual signal, about 5% higher than\nothers, in the normalized desorption curve (Fig. 4d). The absolute residual intensities were\ncomparable across constructs, suggesting that its lower equilibrium binding intensity may have\ncaused the apparent residual fraction after normalization. The dissociation rate of the\n4×GGGGGS construct was nonetheless faster than that of the other constructs.\nAdditionally, we found that RBD-CRD(4×GGGGGS) exhibits relatively fast equilibrium\nkinetics, making it difficult to reliably compare relative association rates using ensemble\nassociation assays. Therefore, we quantified k using single-molecule recruitment assays for the\non\nWT RBD-CRD, the L136A/T178A mutant, and the two linker variants. All k values remained\non\nunchanged, consistent with our previous conclusions. The manuscript has been revised to include\nthis new data in Figure 4c.\n7. The statement in the discussion “Consistent with this, we observed stronger binding of\nRBD-CRD on RAS-free membranes” is unclear. After a few minutes of consideration, I am\nnow thinking that perhaps you mean that RBD-CRD has stronger binding on RAS-free\nmembranes compared to RBD alone or CRD alone? When I originally read this sentence, I\nwas interpreting it as RBD-CRD has stronger binding on RAS-free membranes, compared to\nmembranes containing RAS. Please revise for clarity.\nWe now recognize that the original sentence could be confusing. Our intended meaning was that\non RAS-free membranes, RBD-CRD exhibits stronger binding than CRD alone. We have\ndemonstrated this behavior in a separate study (BioRxiv:\nhttps://www.biorxiv.org/content/10.1101/2025.08.03.668294v1).\nRevision Figure 4. CRAF RBD-CRD exhibits higher binding to RAS-free membranes\ncontaining 20% PS compared to RBD.\nconclusion without referencing our unpublished work. Therefore, we have removed the\ncorresponding sentence. The revised text with the deletion now reads as follows (page 15, line\n380):\n“Previous computational and experimental studies have shown that RBD induces local anionic\nlipid enrichment independently of RAS, thereby enhancing the overall membrane affinity of RBD-\nCRD60. The RBD-induced local enrichment of anionic lipids may support lateral rebinding by\nslowing the dissociation of the transient RAS-free RBD-CRD:PS intermediate.”\n8. To improve clarity, it would be helpful to reference figure numbers throughout the\ndiscussion.\nWe have included figure numbers throughout the discussion.\n9. This point is in regards to the statement in the discussion: “… whereby molecules that\nremain on the membrane for extended periods – such as those within RAS nanoclusters – are\ndisproportionately more likely to become activated.” Based on the evidence presented here, I\ndon’t see how a RAS nanocluster would retain RAF any more than a high concentration of\nmonomeric RAS. I’m seeing how this is related to concentration, but not necessarily how it’s\nrelated to nanoclusters. This needs to be made clearer. Perhaps making it explicit that\nclustering is creating a high local concentration, and RAF will be retained due to this high\nlocal concentration. This is distinct, for example, from requiring a patch of RAS that is a\ncertain diameter or particular shape.\nThe original intent of the discussion on RAS nanoclusters was to propose a hypothesis that a\nhigh local concentration of RAS within nanoclusters may extend RAF membrane residence. To\nreflect the reviewer’s feedback, the description of RAS nanoclusters has been removed from the\nAbstract and Results. Additionally, we have substantially revised the Discussion section to\npresent this concept briefly as an unproven hypothesis, as follows (page 16, line 409):\n“It is worth mentioning that the average density of RAS on the plasma membrane is in the range\nof 30–150 RAS molecules per μm2 67. RAS is not uniformly distributed; instead, it forms\ndynamically exchanging nanoclusters that serve as the primary sites for recruitment and\nactivation of RAF32. The radius of a nanocluster is estimated to be 6–12 nm, with a local surface\ndensity of ~4000 to ~16,000 RAS per μm2 32,33,67. Such high local RAS densities within\nnanoclusters may facilitate RAF activation by extending its membrane residence time through\nlateral rebinding, although this hypothesis remains to be experimentally validated.”\nAdditionally, we have removed detailed descriptions of the RAF activation mechanism involving\nRAS nanoclusters. We also revised the manuscript to explicitly state the need to use the\nautoinhibited full-length RAF protein in future studies to enable a more comprehensive\nmechanistic investigation, as follows (page 17, line 418):\n30\n“To investigate how membrane-binding kinetics influence activation outcomes under more\nphysiologically relevant conditions, it will be necessary to use the autoinhibited full-length\nRAF:14-3-3:MEK complex. These advancements will enable us to rigorously recapitulate the\nsequential steps of RAF activation and elucidate the underlying molecular mechanisms and\nkinetics with high resolution.”\n31"}, {"round": 2, "reviewer_comments": {"reviewer_1": "I acknowledge that the authors have resolved the earlier technical issues and that the strengthened controls and lipid-free assays make the experimental framework sound. However, as I noted in my first-round review, the conceptual advance remains limited: the reported RBD–CRD cooperativity aligns with established structural expectations, and the proposed lateral rebinding mechanism appears to be an expected consequence of multivalent interactions on crowded membrane surfaces rather than a genuinely new insight. Although the manuscript offers careful quantitative confirmation of established ideas, it still falls short of delivering a substantive mechanistic advance.", "reviewer_2": "Thank you again for addressing my comments. I see no need for further revisions, and this manuscript should be accepted for publication.", "reviewer_3": "My previous concerns have all been adequately addressed, and I recommend publication.", "reviewer_4": "I have no further comments. Most of previous comments have been addressed in the revision", "reviewer_5": "The authors have satisfactorily addressed all of my comments."}, "author_response": ""}]

Decision letter