Conformational resolution of nucleotide cycling and effector interactions for multiple small GTPases in parallel

Small GTPase proteins alternatively bind GDP/GTP guanine nucleotides to gate signaling pathways that direct most cellular processes. Numerous GTPases are implicated in oncogenesis, particularly three RAS isoforms HRAS, KRAS and NRAS, and the RHO family GTPase RAC1. Signaling networks comprising small GTPases are highly connected, and there is evidence of direct biochemical crosstalk between the functional G-domains of these proteins. The activation potential of a given GTPase is contingent on a co-dependent interaction with nucleotide and a Mg2+ ion, which bind to individual variants via distinct affinities coordinated by residues in the nucleotide binding pocket. Here, we utilize a selective-labelling strategy coupled with real-time nuclear magnetic resonance (NMR) spectroscopy to monitor nucleotide exchange, GTP hydrolysis and effector interactions of multiple small GTPases in a single complex system. We provide new insight on nucleotide preference and the role of Mg2+ in activating both wild-type and oncogenic mutant enzymes. Multiplexing reveals GEF, GAP and effector binding specificity in mixtures of GTPases and establishes the complete biochemical equivalence of the three related RAS isoforms. This work establishes that direct quantitation of the nucleotide-bound conformation is required to accurately resolve GTPase activation potential, as GTPases such as RALA or the G12C mutant of KRAS demonstrate fast exchange kinetics but have a high affinity for GDP. Further, we propose that the G-domains of small GTPases behave autonomously in solution and nucleotide cycling proceeds independent of protein concentration but is highly impacted by Mg2+ abundance.


Introduction
Small GTPases are a class of critical hub proteins, responsible for controlling both the direction and intensity of cell signals by acting as 'molecular switches' (1)(2)(3). The RAS and RHO subfamilies constitute about one third of all RAS superfamily GTPases (4), and are key regulators of both normal and oncogenic cellular processes including proliferation and migration.
Archetypally, small GTPases interconvert between active GTP-bound and inactive GDP-bound states. Nucleotide exchange and GTP hydrolysis occur intrinsically, however, exchange and hydrolysis rates are catalyzed by guanine nucleotide exchange factors (GEFs) or GTPase activating proteins (GAPs), respectively. Contributions from intrinsic versus catalyzed exchange in vivo are not well understood. In the active state, GTPases bind directly to downstream effector proteins via specialized recognition domains, predominantly RAS-binding domains (RBDs) for RAS GTPases and CRIB motifs for RHO GTPases (5,6).
Within this enzyme superfamily, the most heavily studied are three highly related RAS isoforms: HRAS, KRAS and NRAS. These small GTPases are critical mediators of signaling networks that stimulate cell growth and proliferation, including the MAPK and PI3K/mTOR pathways.
Oncogenic mutations at codons 12, 13 and 61 of the HRAS, KRAS and NRAS genes are amongst the most frequent genetic mutations in human cancers (7). The three RAS proteins share 80% sequence identity and are co-expressed in most cell types. Structurally, they share a nearly identical tertiary fold. As such, these proteins were initially considered functionally redundant, yet multiple lines of evidence support functional specificity: RAS genes exhibit different transforming potential (8)(9)(10), are distinctly mutated in cancers (11,12), exhibit unique sensitivities to GEFs (13) and are localized to discrete subcellular locales (14). Whether genuine biochemical variations in the core G-domains of these proteins contribute to these observed biological differences remains an open question.
Existing approaches to measure small GTPase kinetics include time course-HPLC, release of 32 Pi from GTP- 32 Pi and, most commonly, release or uptake of fluorescent nucleotide analogs. These are crucial assays used to decipher how much activated GTPase subsists in vivo. Unfortunately, several important drawbacks exist with these methods, foremost that fluorescently-tagged analogs can impact reaction kinetics (15). Indeed, the indirect nature of these approaches are a shortcoming that has led to improper conclusions concerning the rate of wild-type and oncogenic RAS mutant nucleotide exchange and its overall on/off state (16,17). To accurately quantify the activation state of GTPases requires consideration of relative nucleotide affinity (i.e. preference of a given GTPase to bind GDP or GTP), the availability of Mg 2+ cofactor, and the potential impact of multimer formation or membrane interactions. This would take into consideration the growing evidence that RAS GTPases dimerize (18,19), which would be intensified at high protein concentrations such as those in membrane nanoclusters.
Recently, real-time nuclear magnetic resonance (RT-NMR) experiments have been adapted to quantitate small GTPase activity (20,21). As GTPases undergo major conformational change upon binding to GDP or GTP, successive collection of 1 H-15 N HSQC spectra allows for kinetic analyses of exchange or hydrolysis. Importantly, these assays do not require fluorescent nucleotide analogs nor any chemical modification of the GTPase. Further, as NMR assays are functional over a wide range of protein concentrations and even on membrane-tethered GTPase (22), they can be used to probe the functional impact of proposed RAS GTPase oligomerization. To strengthen the RT-NMR approach, it is now possible to multiplex these assays (23), allowing quantification of activation states for several GTPases monitored simultaneously in real-time.
We employ here a multiplexed RT-NMR approach to study the full GTPase nucleotide exchange and hydrolysis cycle, as well as specificity of effector RBD binding. Using a selective-labelling strategy we employ RT-NMR to concurrently measure kinetics and effector specificity of the three related RAS isoforms, across RAS and RHO subfamily members, and between cancer-associated mutations of RAS and RAC1 and wild-type counterparts. The data improve our understanding of the complexity and inter-connectedness of small GTPases in the context of cell signaling, particularly the impact of relative nucleotide affinity, Mg 2+ availability and cross-talk mechanisms.

Selective Isotopic GTPase Labelling and Intrinsic Nucleotide Cycling
To juxtapose structure-function data for multiple small GTPases in parallel, we selected six enzymes from the RAS and RHO subfamilies (Fig. 1A). Structures of these proteins demonstrate the exceptional similarity of their tertiary folds (Fig. 1B). Identical sizes, shapes and enzymatic functions makes in vitro profiling of numerous GTPase activities together a massive challenge.
For NMR-based analyses, uniformly labeled samples of multiple GTPases results in excessive crowding of resonance peaks, as demonstrated in Fig. 1C using HRAS, KRAS and NRAS. To observe multiple GTPases concurrently required a selective isotopic labelling approach with single 15 N amino acids. Fig. 1D and 1E show isotopically labeled RAS isoforms ( 15 N-Ile HRAS, 15 N-Thr KRAS and 15 N-Leu NRAS) in both the GDP-bound and GTP-bound conformation (using nonhydrolyzable GMPPNP). The reduction in spectral complexity allowed us to monitor nucleotide exchange, GTP hydrolysis or effector binding in mixtures of even the most highly related GTPases.
We performed exchange and hydrolysis assays on individual selectively-labeled GTPases to profile their kinetic activity in isolation. NMR spectra and nucleotide exchange plots for 15 Table S1 (for all individually measured GTPase kinetics). Exchange was initiated by the addition of a 10:1 molar ratio of the GTP analog GTPγS, used to reflect the cellular GTP/GDP ratio. Measured exchange rates for HRAS match what has been calculated by RT-NMR using uniformly labeled samples (21). As individual GTPases reach an activation plateau (i.e. nucleotide exchange no longer proceeds), they exhibit differential GTPγS-loading that reflects their activation state at equilibrium. We report these data in Supplemental Table S1 as "% Activated". Further, as we control the GTPγS:GDP ratio at 10:1 we can also calculate the preference of these GTPases to each nucleotide, reported as "GDP Preference". We observed the three RAS isoforms and RHEB exhibited a much higher ratio of GTPγS-bound (~65-75%) in these conditions compared to RALA (38%) or RHOG (49%), a direct reflection of differential nucleotide affinities. We next measured intrinsic GTP hydrolysis for each RAS isoform ( 15 N-Tyr HRAS, 15 N-Thr KRAS and 15 N-Leu NRAS) (Supplemental Fig. S2H-J). The measured rate for HRAS matches what was previously determined on uniformly labeled protein (21), and NRAS and KRAS exhibited near identical rates. Overall, the selective-labeling approach provides an opportunity to precisely measure GTPase activity while reducing spectral complexity.

Dependence of Nucleotide Exchange on Mg 2+
To begin assay of multiplexed GTPases, we first focused on the three highly related RAS isoforms.
There has been little attempt to study differences in nucleotide exchange and/or GTP hydrolysis rates for the RAS isoforms in their individual contexts, however, one study found that the intrinsic rates of GTP catalysis differed across isoforms using 32 Pi-GTP single turnover (24). Due to the inefficiency of selective-labeling, monitoring HSQC chemical shifts of the three GTPases required each at a concentration of 300 μM. Interestingly, this meant we could assay GTPase activity at concentrations that should drive RAS dimerization. The existence of KRAS homodimers remains contentious, with proposed dissociation binding constants ranging from low µM (25) to mM (26).
Recent biophysical data dispute the existence of RAS dimers even at high concentrations (27,28).
Performing these assays at total GTPase concentrations approaching 1 mM would resolve whether G-domain oligomers can influence RAS activation in solution. Initial multiplexed nucleotide exchange assays (GDP-to-GTPγS) at a 10:1 molar excess of GTPγS showed exchange rates substantially faster than those measured for GTPases individually at comparatively lower concentrations. To resolve this, we considered that exchange assays with each RAS isoform alone at various concentrations (150, 250 or 350 μM), showed rates increase with protein concentration ( Fig. 2A-C). Mg 2+ cofactor is absolutely required for nucleotide binding to RAS GTPases, and both RAS (29) and RHO (30) nucleotide exchange is highly dependent on Mg 2+ concentration.
The majority of GTPase kinetic studies use MgCl2 at a steady concentration of 5 mM, and we postulated that increasing protein concentrations lower the [Mg 2+ ]:[GTPase] ratio, leading to Mg 2+ scarcity and faster exchange rates. To test this, we measured nucleotide exchange of NRAS at 200 µM or 350 µM in 5 mM MgCl2, again showing that the rate increases at a higher concentration ( Fig. 2D). When we repeated the assay using 15 mM MgCl2, exchange rates were identical. Thus, Mg 2+ availability is a key determinant of GTPase exchange, which otherwise proceeds independent of protein concentration even approaching 1 mM.
Interestingly, there are several recurrent oncogenic mutants of RAS that have been determined to function by via rapid intrinsic exchange (21). As these mutations lie proximal to the nucleotide/Mg 2+ binding pocket we were curious if increasing [Mg 2+ ] may slow their intrinsic exchange rate. We purified isotopically labeled KRAS proteins of two fast exchange mutants, G13D (found within the P-loop) and Q61L (in the switch II region) (21, 31) (Fig. 2E). We performed nucleotide exchange assays at either 5 or 50 mM MgCl2 (Fig. 2F). There was no Mg 2+ dependence on the exchange rate of either mutant, indicating these amino acid mutations likely alter nucleotide affinity rather than disrupt coordination of the Mg 2+ ion.
Finally, we tested whether increasing concentrations of RAS with a steady concentration of Mg 2+ would lead to differences in GTP hydrolysis. Neither KRAS, HRAS nor NRAS exhibited altered GTP hydrolysis rates with increasing protein concentration at a constant [Mg 2+ ] of 5 mM ( Fig.   2G-I). This is consistent with the absence of competing nucleotide in these assays, and with data suggesting GDP-binding is more dependent on Mg 2+ than is GTP binding (32).

Multiplexed Nucleotide Exchange Assays
With a strategy for selective amino acid labelling and conditions optimized for simultaneously monitoring multiple GTPase activities, we performed a series of multiplexed nucleotide exchange assays. concentrations did not lead to obvious binding or higher order complexes, as no significant chemical shift perturbations or peak broadening was observed. Intrinsic exchange rates for each of the three isoforms were not changed from those measured for HRAS, KRAS and NRAS alone at lower concentrations. Kinetic analyses and parameters for all multiplexed assays are detailed in Supplemental Table S2. All three GTPases were 80-90% activated at equilibrium and demonstrate a minor preference for GDP over GTPγS. We can conclude that neither the presence of alternative RAS isoforms nor high concentrations of these GTPases significantly impact their activation state.
We next examined the NRAS GTPase in parallel with a related GTPase, RALA, and a RHO subfamily GTPase, RHOG. Fig. 3F-J depicts the labelling strategy and multiplexed exchange.
RALA and RHOG exhibited slightly faster exchange rates than NRAS (1.4-fold and 1.3-fold, respectively), but their % activation at equilibrium is significantly lower than that of NRAS (84% GTPγS-bound for NRAS, 46% RALA, and 49% RHOG). This reveals that RALA and RHOG have a greater preference for GDP over GTPγS compared to NRAS. These data highlight the need to directly monitor the nucleotide-bound state when quantifying GTPase activation, as kinetic rates alone can significantly misrepresent the total activated GTPase in a given system.

Multiplexed GEF and GAP Assays
Our multiplexed NMR strategy presents an opportunity to observe the effects and specificity of exchange-promoting GEFs and hydrolysis-activating GAPs on mixtures of small GTPases (Fig.   4A). We performed a GEF assay in a mixture of the three isoforms of RAS using a catalytic domain from the major RAS regulator SOS1 (SOScat (21,33)), added at a molar ratio of 1:15000 (versus [total GTPase]) (Fig. 4B). We calculated similar increases in the SOS-catalyzed exchange rate for all three isoforms, (4.5-fold increase for HRAS, 3.3-fold for KRAS and 3-fold for NRAS) indicating that they have comparable sensitivities to this GEF.
To observe SOS specificity, we performed the same assay using multiplexed NRAS, RALA and RHOG (Fig. 4C). In the presence of SOScat, we observed a significant increase in the exchange rate of NRAS relative to intrinsic and no effect on the exchange rates of RALA or RHOG. We next performed a multiplexed assay using KRAS and RAC1, as SOS is reported to have GEF activity towards both. The activating GEF domains differ for each GTPase: the REM-CDC25 domains (SOScat) mediate activity towards RAS (33,34), and the DH-PH domains activate RAC1 (35,36). The intrinsic exchange of KRAS proceeds 1.3-fold faster than that of RAC1, with % activated at equilibrium measured at 83% and 65%, respectively (Fig. 4D). Upon addition of SOScat, KRAS nucleotide exchange increases 2.5-fold while RAC1 rates remained unchanged (Fig. 4E). Notably, the nucleotide bound ratio of KRAS at equilibrium is not altered by the presence of SOS (83%), supporting a model whereby GEFs do not actively exchange nucleotide but function by a passive mechanism.
We next employed our approach to measure GAP activation of GTPase hydrolysis by performing multiplexed assays with the three RAS isoforms. Calculated intrinsic hydrolysis rates for HRAS, KRAS and NRAS were indistinguishable when measured in tandem (Fig. 4F/G). In the presence of recombinant GAP-334 domain from the major RAS regulator p120GAP (added at a 1:5500), hydrolysis rates of each isoform were uniformly increased (2.3-fold HRAS, 2.6-fold KRAS and 2.9-fold NRAS (Fig. 4H)). These results indicate that the three isoforms share sensitivity to GAP activation, in addition to having comparable rates of intrinsic hydrolysis.

Multiplexed GTPase Assays with Oncoproteins
There are conflicting reports on wild-type RAS isoforms influencing the transformation potential of oncogenic mutants, and little is known about cross-talk between wild-type and mutant GTPases from a biophysical standpoint. We used multiplexed RT-NMR to concurrently monitor nucleotide exchange and GTP hydrolysis of wild-type RAS isoforms and several oncogene-derived mutants nucleotide exchange compared to their wild-type counterparts, while KRAS Q61L exhibits rapid intrinsic exchange. This was also observed when these oncoproteins were multiplexed with the matched wild-type, whereby KRAS G12C intrinsically exchanges at a rate 2.9-fold slower than wild-type, HRAS G12V at a rate 1.6-fold slower, and KRAS Q61L 1.6-fold faster (Supplemental Fig. S4D-F). Significantly, the KRAS G12C variant reaches equilibrium at only 67% activated and HRAS G12V at only 36%. In contrast, KRAS Q61L reaches 87% activation under these conditions. We extended these analyses to measure GEF activity by adding SOScat. The KRAS G12C mutant had a 1.5-fold slower exchange rate in the presence of SOScat than wild-type KRAS, similar to the intrinsic difference (Fig. 5B). A multiplexed GEF assay with wild-type KRAS and the Q61L variant again demonstrated that Q61L exchanges at a faster rate than wild-type (1.3fold) and reaches equilibrium at nearly 90% GTPγS-bound (Fig. 5C). These data suggest that there is no biophysical interplay between wild-type and oncogenic RAS GTPases at even high concentrations of G-domain. They suggest that large pools of the RAS G12V or G12C variants likely remain GDP-bound even in the presence of activating GEF. Impaired GTP hydrolysis is a key biochemical defect in oncogenic RAS GTPases. We determined that KRAS G12C has a 2.6-fold slower intrinsic rate of hydrolysis than wild-type KRAS when measured individually (Supplemental Fig. S4G) or in a multiplexed assay (Supplemental Fig.   S4H). The KRAS Q61L variant showed effectively no intrinsic GTP hydrolysis over a 10 hr time course (Supplemental Fig. S4I). To monitor rates in the presence of GAP, we added GAP-334 to multiplexed samples. The addition of GAP at 1:5000 did not affect the GTP hydrolysis rate of either G12C or Q61L KRAS (Fig. 5D/E), while the hydrolysis rate of wild-type KRAS increased 2.5-fold. Thus, the presence of neither wild-type KRAS nor GAP significantly alters the ability of these oncogenic mutants to hydrolyze GTP.
Genetic defects impacting small GTPase function in cancer are not limited to RAS proteins, so we sought to examine biophysical interplay between RAC1 and a RAC1 P29S mutant recurrently found in melanoma (Fig. 5F). A nucleotide exchange assay on the P29S variant alone demonstrated this oncoprotein intrinsically exchanges 7-fold faster than wild-type RAC1 (Supplemental Fig. S4J). A multiplexed approach with wild-type and P29S RAC1 provided the same result (Fig. 5G). Importantly, P29S RAC1 reaches almost 100% activation in these conditions, demonstrating a clear preference for binding GTPγS over GDP, while wild-type RAC1 shows a 6-fold preference for GDP. The utility of an approach that directly monitors GTPase conformation is fully demonstrated in Fig. 5H and Supplemental Fig. S4K. These NMR resonances demonstrate how poorly the HRAS G12V and KRAS G12C oncoproteins exchange over time in a 10-fold excess of GTPγS, as compared to their wild-type counterparts, KRAS Q61L or RAC1 P29S.

Multiplexed Effector Binding Assays
The specificity of effector binding domains for small GTPases is a question of huge interest, and one that has not been well explored. Effectors targeting RAS subfamily GTPases typically comprise an RBD domain, and there are >50 potential RBDs in the human proteome (37). To directly observe effector specificity and competition for GTPase binding partners in a complex system we used NMR and multiple selectively labeled GTPases. We purified RBDs from two isoforms of the effector RAF kinases (ARAF and BRAF) and an RBD from the RAL effector RLIP76 (Fig. 6A). The RBD of ARAF was added to GMPPNP-loaded HRAS, KRAS and NRAS, and displayed uniform binding to all three isoforms as determined by peak broadening and chemical shift perturbations (Fig. 6B). Next, we titrated the RBD of BRAF and it also induced complete broadening across all three isoforms of RAS (Fig. 6C). To observe specificity across GTPase subfamilies, the BRAF RBD was titrated into a mixture of GMPPNP-loaded KRAS, NRAS and RHOG. Once again, severe peak broadening was observed for the KRAS and NRAS resonances, while peaks derived from RHOG were left unperturbed (Fig. 6D). The structurally unrelated RBD from RLIP76 was then titrated into a mixture of HRAS, KRAS and RALA. Only RALA peaks displayed peak broadening while HRAS and KRAS peaks were unperturbed (Fig.   6E). Finally, we looked to titrate an engineered monobody against RAS, NS1 (38), into a multiplexed mixture of HRAS, KRAS and NRAS. This monobody was designed to interact with the distal site (from the nucleotide-binding pocket) of HRAS and KRAS to prevent its dimerization/clustering on the membrane. Upon titration of NS1 into the three isoforms of RAS, peak broadening was observed only in HRAS and KRAS (Fig. 6F), while NRAS-specific chemical shifts were unperturbed. Overall, the multiplexed NMR approach is a powerful technique to observe interactions and binding specificities of multiple, unmodified proteins simultaneously.

Discussion
Accurate data characterizing small GTPase nucleotide cycling are crucial to our understanding of both wild-type and mutant activation potential in cells. These are extremely challenging experiments that must consider co-dependent nucleotide and Mg 2+ affinity, competing nucleotides and metal ions, and the distinct biochemical properties of a given GTPase. We have investigated the complete nucleotide cycle and effector interactions for multiple GTPases in parallel using a selective labeling approach coupled with RT-NMR. These multiplexed assays reveal details of GEF and GAP specificities, protein binding preferences, and nucleotide-dependent activation states that cannot be accurately assessed using conventional techniques.

Biochemical diversity in the three isoforms of RAS is an open question with large implications for
developmental biology and cellular transformation. We find that intrinsic rates of exchange and hydrolysis are nearly identical across the three isoforms, as are their sensitivities to activation by SOS or inactivation by p120GAP. It appears that the core G-domains of HRAS, KRAS and NRAS are biochemically equivalent and that biological differences are likely determined by posttranslational modification, subcellular localization or distinct effector/protein interactions. It will be interesting to determine if these results hold true for other RASGEF proteins such RASGRFs or RASGRPs (39,40), and to examine GEF specificity against all 35 RAS subfamily GTPases (4).
Using high protein concentrations to profile GTPase kinetics revealed a strong dependence on Mg 2+ for nucleotide exchange. The Mg 2+ concentration in mammalian cells has been estimated at 17-20 mM, however, less than 5% of that is presumed free (41,42). The cytoplasm (where membrane-tethered small GTPase proteins are exposed) is expected to have only 0.5-1 mM free Mg 2+ , significantly less than used in most in vitro kinetic assays. Several thoughtful experiments from nearly three decades ago estimated the Mg 2+ affinity for wild-type HRAS at 2.8 µM (43), and resolved that high Mg 2+ concentrations in RAS exchange assays significantly slow GDP dissociation rates. Moreover, there are intriguing data that Mg 2+ -GTP affinity for HRAS is higher than that of Mg 2+ -GDP (32). This would be consistent with our observations of increased nucleotide exchange in conditions of Mg 2+ scarcity, but how this affects RAS activity in cells is unknown. We can speculate that high density RAS nanoclusters may be influenced by Mg 2+ availability, which could act to promote nucleotide exchange. Supplemental Table S3 complies the existing knowledge of RAS nucleotide binding and the little that is known about Mg 2+ affinity.
Most of these early data were generated by measuring retention of radiolabeled nucleotides on filters, but these detailed assessments of GTPase biochemistry and function should be reconsidered using modern approaches.
Multiple lines of evidence suggest functional interplay between wild-type and mutant RAS proteins, with claims that wild-type RAS can suppress oncogenic mutant activity (19,(44)(45)(46) and others that mutants promote wild-type RAS activation (47,48). Multiplexing GTPase mutants with their wild-type counterpart G-domains provided an opportunity to explore direct biophysical effects on hydrolysis or exchange. Our results corroborate observations that mutations at G12 (Cys or Val) or Q61 (Leu) have unique deficiencies. The G12 oncoproteins exhibit very slow intrinsic exchange and hydrolysis rates, minor sensitivity to GEF and complete insensitivity to GAPs, while Q61L is a fast exchange mutant that exhibits virtually no hydrolysis of GTP. RAC1 P29S, the third most frequently observed hotspot mutation in melanoma (after NRAS and BRAF) (49,50), is also a fast exchange mutant. We observed no significant cross-talk between wild-type and oncogenic mutant proteins, either in chemical shift perturbations upon incubation or by measuring nucleotide cycling kinetics.
Multiple selectively labeled proteins provide an excellent opportunity to directly measure binding competition in a complex in vitro system. Addition of the ARAF or BRAF RBDs to mixtures of the three RAS isoforms showed uniform binding to all three GTPases, while we could observe clear specificity differences using either distinct effector binding domains or distally related GTPases. While these NMR mixing experiments led to peak broadening in our system, improved approaches able to detect chemical shift perturbations will allow for calculation of affinities in a competitive setting. Future combination of multiplexed GTPases with multiple competing effectors (51) will facilitate study of ever-more complex in vitro systems that retain a capacity to deliver atomic-level, quantitative data for composite protein-protein interactions.
Precise and dynamic measurements of GTPase activation states are essential to elucidate their role in development and disease. Here, we have collectively demonstrated how selective isotopic labelling and RT-NMR offer improved measures of intrinsic, GEF and GAP enzymatic activities and effector binding specificities for multiple GTPases.

Protein Expression and Purification
For unlabeled GST-tagged proteins, proteins were expressed in Escherichia coli BL21 codon+ Spectra were processed with NMRPipe (53) and analyzed using NMRView (54). For intrinsic exchange and GEF assays, GTPγS was added at a 8-10 molar fold excess (GTPγS: Total Protein) and SOScat was added at molar ratios described in our results. To calculate the GDP-bound ratio [IGDP/(IGDP + IGTP)], peak intensities were extracted from each individual spectrum using NMRView . Exchange curves were plotted and fitted to a single phase exponential decay function using GraphPad Software. For intrinsic GTP hydrolysis and GAP assays, peak intensities were extracted and data were fit to a one phase exponential association function. For effector/monobody titrations, unlabeled RBD domain or NS1 monobody was titrated into selectively labeled mixtures of GTPases.