Small-molecule ligands bind to a distinctpocket in Ras and inhibit SOS-mediatednucleotide exchange activityTill Maurera,1, Lindsay S. Garrentonb,1, Angela Oha, Keith Pittsc, Daniel J. Andersonb, Nicholas J. Skeltond,Benjamin P. Fauberd, Borlan Pana, Shiva Malekc, David Stokoeb, Mary J. C. Ludlamb, Krista K. Bowmana,Jiansheng Wue, Anthony M. Giannettic, Melissa A. Starovasnika, Ira Mellmanb, Peter K. Jacksonb,Joachim Rudolphd, Weiru Wanga,2, and Guowei Fangb,2

aStructural Biology, bResearch Oncology, cBiochemical and Cellular Pharmacology, dDiscovery Chemistry, and eProtein Chemistry, Genentech, Inc.,One DNA Way, South San Francisco, CA 94080

Edited by* Sung-Hou Kim, University of California, Berkeley, CA, and approved February 2, 2012 (received for review October 6, 2011)

The Ras gene is frequently mutated in cancer, and mutant Rasdrives tumorigenesis. Although Ras is a central oncogene, smallmolecules that bind to Ras in a well-defined manner and exertinhibitory effects have not been uncovered to date. Through anNMR-based fragment screen, we identified a group of small mole-cules that all bind to a common site on Ras. High-resolution cocrys-tal structures delineated a unique ligand-binding pocket on theRas protein that is adjacent to the switch I/II regions and can beexpanded upon compound binding. Structure analysis predictsthat compound-binding interferes with the Ras/SOS interactions.Indeed, selected compounds inhibit SOS-mediated nucleotideexchange and prevent Ras activation by blocking the formationof intermediates of the exchange reaction. The discovery of asmall-molecule binding pocket on Ras with functional significanceprovides a new direction in the search of therapeutically effectiveinhibitors of the Ras oncoprotein.

small G protein ∣ guanine nucleotide exchange ∣ nuclear magneticresonance ∣ crystal structure ∣ small-molecule inhibitors

Ras is a small GTP-binding protein that functions as a nucleo-tide-dependent switch for central growth signaling pathways(1, 2). In response to extracellular signals, Ras is converted from aGDP-bound (RasGDP) to a GTP-bound (RasGTP) state, as cata-lyzed by guanine nucleotide exchange factors (GEFs), notably theSOS1 protein. Active RasGTP mediates its diverse growth-stimu-lating functions through its direct interactions with effectorsincluding Raf, PI3K, and Ral guanine nucleotide dissociationstimulator. The intrinsic GTPase activity of Ras then hydrolyzesGTP to GDP to terminate Ras signaling. The Ras GTPase activitycan be further accelerated by its interactions with GTPase-acti-vating proteins (GAPs), including the neurofibromin 1 tumorsuppressor (2).

Ras, a human oncogene identified and characterized over 30 yago, is mutated in more than 20% of human cancers. Among thethree Ras isoforms (K, N, and H), KRas is most frequentlymutated (2). Mutant Ras has a reduced GTPase activity, whichprolongs its activated conformation, thereby promoting Ras-dependent signaling and cancer cell survival or growth (1, 2).

Mutations of Ras in cancer are associated with poor prognosis(2). Inactivation of oncogenic Ras in mice results in tumor shrink-age. Thus, Ras is widely considered an oncology target of excep-tional importance. However, development of small-moleculeinhibitors against Ras has thus far proven unsuccessful. Giventhe picomolar affinity between guanine nucleotides and Ras andthe high cytosolic concentration of guanine nucleotides, it is verychallenging to develop a conventional inhibitor competitiveagainst nucleotide binding (1, 2). Outside of the nucleotide-bind-ing pocket, the Ras protein does not contain obvious cavities forsmall-molecule binding. A number of small molecules have been

reported to bind to Ras (3–7), but their mechanisms of action andthe structural basis to achieve Ras inhibition remain elusive.

Fragment-based lead discovery (FBLD) has provided a start-ing point to develop a number of chemical entities for targetspreviously considered “undruggable,” notably Bcl2 (reviewed inref. 8). This approach has benefited in particular from the use ofNMR and surface plasmon resonance for ultrasensitive detectionof interactions between ligand and protein, even at millimolaraffinities. Aside from the identification of ligands for a knownbinding site, a fragment-based screen can also reveal unique bind-ing sites in proteins. Central to this discovery process is the abilityto obtain high-resolution structures of the ligand-protein com-plex. Characterization of the site through a combination of struc-tural studies and biophysical and biochemical examination formthe basis for the hit-to-lead optimization process. We have ap-plied FBLD to probe oncogenic KRas, and report here the dis-covery of a unique small-molecule binding pocket on Ras and thecharacterization of the effect of the ligands on Ras activation.

ResultsA Fragment Screen Identifies Ras-Binding Small Molecules.An NMR-based saturation transfer difference (STD) assay (9) was used todetect ligand binding to the recombinant KRas4B-G12D, KRasm(Fig. S1A). A screen of a 3,300-compound library was performed(using pools of up to six fragments each) by measuring compoundbinding to a 1∶1 mixture of KRasmGDP and KRasm bound toGDP or guanosine-5′-[(β, γ)-methyleno]triphosphate (GMPPCP)proteins. Representative screening data and their deconvolutionare shown in Fig. 1A and Fig. S1B. Binders were identified thatshowed an STD signal/noise ratio greater than five. Candidatefragments from positive pools were retested as single compoundsfor their binding to KRasmGDP and KRasmGMPPCP separately,yielding 240 primary hits. These hits were further validated bycomparing 2D 1H15N heteronuclear single quantum coherence(HSQC) spectra of isotopically labeled KRasmGDP in the pre-sence or absence of 2 mM compounds. Of these, 25 compoundsproduced chemical shift perturbations that can be mapped to a

Author contributions: S.M., D.S., M.A.S., I.M., P.K.J., J.R., W.W., and G.F. designed research;T.M., L.S.G., A.O., K.P., D.J.A., N.J.S., B.P., A.M.G., andW.W. performed research; A.O., B.P.F.,M.J.C.L., K.K.B., J.W., and J.R. contributed new reagents/analytic tools; T.M., L.S.G., N.J.S.,J.R., W.W., and G.F. analyzed data; and T.M., L.S.G., J.R., W.W., and G.F. wrote the paper.

Conflict of interest statement: All authors are employees of Genentech, Inc.

*This Direct Submission article had a prearranged editor.

Data deposition: The KRas structure factors have been deposited in the Protein DataBank, www.pdb.org (PDB ID codes 4DSO, 4DST, and 4DSU).1T.M. and L.S.G. contributed equally to this work.2To whom correspondence may be addressed. E-mail: fang.guowei@gene.com or wang.weiru@gene.com.

This article contains supporting information online at www.pnas.org/lookup/suppl/doi:10.1073/pnas.1116510109/-/DCSupplemental.

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contiguous site on the KRasm structure and were thus classifiedas confirmed hits (Fig. S1A). Intriguingly, each of the 25 con-firmed hits showed chemical shift perturbations for residues V8,T74/G75 and L56/D57 (Fig. 1B and Fig. S1C). Some compoundsalso perturbed additional resonances corresponding to residuesclose to the region of D57.

High-Resolution Cocrystal Structures Reveal a Unique Binding Pocketon Ras. To understand the molecular basis of Ras interaction withthese compounds, we determined the first full-length KRasm crys-tal structure in the presence of GDP or nonhydrolyzable GTPanalogs (GMPPCP) at 1.75- and 2.05-Å resolution, respectively.Within the GTPase domain (amino acids 1–169), the KRasm

structure (Fig. 2A) is similar to the structure of truncatedKRas4B-Q61H [amino acids 1–169 with residue 61 mutated fromglutamine to histidine; Protein Data Bank (PDB) code 3GFT;Structural Genomics Consortium]. Excluding the switch regions,the root-mean-square deviation in Cα positions between thesetwo structures is 0.5 Å. In addition, they share a high degreeof similarity to the truncated HRas (10) and NRas (PDB code3CON; Structural Genomics Consortium) structures. The switchI and II regions display conformational heterogeneity amongdifferent Ras structures because they are sensitive to the nucleo-tide state and, to some extent, influenced by crystal packinginteractions.

Our full-length KRasm structure reveals additional featurespertaining to the C-terminal domain (CTD) (Fig. 2A). Aside fromthe C-terminal CAAX farnesylation motif, the CTD sequence ishypervariable across K, N, and HRas. The KRasm CTD containsa unique hexalysine stretch for direct association with cytoplasmicmembrane. In the crystal structure, helix α5 from the GTPase do-main is extended by one turn after which the CTDmakes a π-turnfollowed by an extended loop containing the hexalysine stretch.This extended conformation is likely to promote interaction ofthe lysine stretch with negatively charged lipid head groups fromthe membrane (Fig. 2A).

Cocrystals of ligands and KRasm were obtained by soaking theconfirmed fragment hits into the KRasm crystals. Fig. S2 A–Cdepicts the complexes with benzamidine (BZDN), benzimidazole(BZIM), and 4,6-dichloro-2-methyl-3-aminoethyl-indole (DCAI),respectively. In all three cases, compounds bind to a similar siteon KRasm.

This small-molecule binding site (Fig. 2A and Fig. S2 A–D)resides between helix α2 and the core β-sheet, β1–β3. The com-pounds show amphipathic interactions centering on a hydrophobicpocket of approximately 7 × 7 Å at the opening and 5-Å deep(Fig. 2B). This binding pocket has a size sufficiently large to accom-modate a benzyl and a chloro group. For comparison, a typicalATP-binding site found at kinase active sites is about 20 × 10 ×10 Å [as an example, see the p38MAPkinase (PDB code 1WFC)].

Residues surrounding the binding pocket in Ras include K5,L6, V7, I55, L56, and T74 (Fig. S2D). This observation is largelyconsistent with the mapping from the NMR spectrum perturba-tions because all of the perturbed residues (Figs. 1B and 2 A–C,

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Fig. 1. NMR-based fragment screen identifies Ras-binding compounds. (A)One-dimensional proton spectra showing representative screening results.(i) The 1D reference spectrum of a compound pool containing DCAI. Arrowspoint to the peaks of DCAI. (ii) The STD spectrum of the DCAI-containingpool. (B) These panels show the 1H15N HSQC of KRasmGDP at a concentrationof 100 μM titrated with 0, 250, 500, 1,000, 1,500, and 4,000 μM DCAI (fromblack to purple). Residues V8, T74, G75, L56, and D57 make up one contig-uous site in the Ras protein structure. Side-chain resonances are labeled “sc.”

Fig. 2. Ras-ligand cocrystal structures reveal anexpandable small-molecule binding pocket in KRas.(A) Structure of KRasm bound to GTPγS-Mg2þ. Theswitch I/II regions are colored in green and orange,respectively. The spheres indicate residues withchemical shift perturbations upon DCAI binding inthe NMR experiments. The bound GTPγS is shownas sticks and Mg2þ as a blue dot. The disordered re-gion of KRasm is drawn as a dotted line. Hypotheticcytoplasmic membrane is shown to illustrate itspotential interactions with the KRasm C-terminal do-main. (B) The small-molecule binding pocket onKRas. Surface of KRasm in complex with BZDN(orange) and BZIM (cyan) is colored by electrostaticpotential with red being negative and blue beingpositive. (C) The DCAI-binding pocket on KRasm.D54 and R41 side chains are shown in rotamer con-formation with (cyan) and without (blue) DCAIbound. Arrows indicate the direction of their rota-tion upon DCAI binding. Residues with chemical shiftperturbations in the NMR spectra are colored in gray.(D) A close-up surface rendering of the DCAI 4-chlorine atom (in green) and its interacting surfaceon KRasm (in blue) in comparison with the KRasmsurface with no ligand bound (in brown).

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and Figs. S1C and S2 A–D) are located in or around the bindingpocket. The surface area outside the pocket is more hydrophilicand engages in specific interactions with the polar portion of thecompounds. For example, KRasm D54 at the rim of the bindingpocket forms hydrogen bonds with the NH group in BZDN andBZIM (Fig. 2B). H, N, and KRas have identical amino acid se-quences in the binding site, which suggests that the compoundswould likely bind to all three isoforms. We confirmed that DCAIand BZIM bind to HRas in the STD assay (Fig. S2 E and F).

The Small-Molecule Binding Pocket on Ras Can Be Expanded uponDCAI Binding. The crystal structure of DCAI-KRasmGMPPCP re-veals a high degree of plasticity in the binding pocket (Fig. 2 Cand D). The chloro-substituent at the six-position of DCAI an-chors the ligand into the hydrophobic pocket. DCAI is larger thanBZDN and BZIM, and the 4-chloro group of DCAI is accommo-dated in the binding pocket through reorienting its indole coreby 65° relative to BZIM. Additionally, the side chain of D54of KRasm moves away from the binding site by 2.5 Å (Fig. 2C).D54 normally forms a salt bridge with R41 in the absence of com-pound binding. This compound-induced rotamer conformationof D54 effects an outward flipping (7.5 Å) of R41 and preservesthe salt bridge (Fig. 2C). Overall, the pocket surface retreats byabout 3.5 Å upon DCAI binding (Fig. 2D), rendering the ligandbinding site opening to a size of 7 × 10.5 Å, although the ex-tended region is relatively shallow.

DCAI Inhibits SOS-Mediated Nucleotide Exchange for Ras. Compari-son of the Ras-DCAI structure with the structure of the Ras-SOS1 complex (11) indicates that the small-molecule bindingpocket described above is located adjacent to the Ras–SOS inter-action surface (Fig. 3A), suggesting that ligand binding may affectSOS-mediated nucleotide exchange for KRas. To investigatethe nucleotide exchange reaction, a nucleotide exchange and anucleotide release assay were employed using fluorescentN-methylanthraniloyl (MANT) derivatives of guanine nucleo-tides, recombinant wild-type KRas, and a catalytically active formof SOS1 (SOScat) consisting of its Ras exchanger motif and Cdc25domains (11, 12) (Fig. S3). The kinetics of these reactions weremonitored by changes in fluorescence intensity of MANT-nucleo-tide in its Ras-bound versus free state. Our nucleotide exchangeassay detects inhibitors against any step in the reaction mechan-ism, whereas the nucleotide release assay specifically identifiesinhibitors against the first phase of the exchange reaction, nucleo-tide release (Fig. S3).

We analyzed representative hits from the fragment screen inthese two assays and found that DCAI effectively inhibited bothreactions (Fig. S4A). Titration experiments indicated that DCAIblocked both nucleotide exchange and release reactions with anIC50 of 342� 22 μM and 155� 36 μM, respectively, whereasBZIM and indole (INDL) had no or minimal effects (Fig. 3Band Fig. S4B). We conclude that DCAI blocks the first phaseof the exchange reaction by preventing nucleotide release fromKRas. This inhibitory effect is not restricted to DCAI. We foundthat 2-(4,6-dichloro-2,3-dimethyl-1H-indol-1-yl)ethanamine, aclose structural analog of DCAI, also binds to KRas and inhibitsSOS-mediated nucleotide release from KRas (Fig. S4C).

In addition towild-typeKRas,DCAIalso inhibits SOS-catalyzednucleotide release from mutant KRas (KRasm) (Fig. 3C). In fact,DCAI exhibits a lower IC50 against KRasm than wild-type KRas,consistent with the different kinetics of the SOS-catalyzed releasereaction for KRas vs. KRasm. At saturating SOS concentrations,the nucleotide release rate is fivefold slower for KRasm than forwild-type KRas (Fig. S4D), indicating that SOScat is less effectiveat releasing nucleotide from KRasm compared to wild-type KRas.

DCAI inhibits nucleotide release from Ras in an SOS-depen-dent manner only. EDTA chelates Mg2þ ions and promotes nu-cleotide release from KRas independent of SOS; DCAI had no

effect on the rate of EDTA-stimulated nucleotide release fromKRas or KRasm (Fig. S5A). In addition, we found that DCAIhas no effect on the intrinsic ability of KRas to release nucleotidein the absence of EDTA and SOS (Fig. S5 B–C).

DCAI specifically inhibits the SOS-mediated nucleotide ex-change on Ras, but not the Dbs-mediated nucleotide exchangeon RhoA and Cdc42 (Fig. 3D), consistent with the fact that bulkyresidues, Trp58 in RhoA and Phe56 in Cdc42, occupy the spaceequivalent to the DCAI-binding site in RhoA and Cdc42, therebypotentially preventing the binding of DCAI to RhoA and Cdc42(Fig. S6).

DCAI Inhibits Nucleotide Exchange by Blocking the InteractionsBetween Ras and SOS. Next, we investigated the mechanism of theDCAI-mediated inhibition of nucleotide release. Based on theoverlap of the small-molecule binding pocket with theRas-SOS in-teraction surface (Fig. 3A), we predicted that DCAI binds to KRasand competes with SOS association. Further structural analysisof the apoRas-SOS complex suggests two mechanisms by whichbinding of DCAI to KRas interferes with the formation of the ex-change reaction intermediates and thereby inhibits SOS-mediatedcatalysis (Fig. 4A and Fig. S7A). First, the 2-methyl and 3-ami-noethyl side chains of DCAI sterically hinder the binding ofSOScat to Ras. Second, binding of DCAI to KRas results in a rota-tion of both D54 and R41 to enlarge the binding pocket (Figs. 2 Cand D, 4A, and Fig. S7A); reorientation of these side chains willprevent them from participating in two salt bridges that stabilizethe Ras–SOS interaction (to SOS H911 and SOS D910, respec-tively; Fig. 4A and Fig. S7A) (11). Consistent with this model,the KRas R41S mutant reduces the rate of the SOS-mediatednucleotide release reaction by 2.5-fold (Fig. 4B), even though thismutation does not affect the binding of DCAI to KRas (Fig. S2G).We conclude that R41 is important for SOS-catalyzed nucleotideexchange on Ras, but dispensable for recognition by DCAI.

In contrast, structural analysis suggests that the DCAI-relatedcompound, BZIM, would not sterically hinder SOS binding(Fig. 4A and Fig. S7A) and would not alter the ability of D54 andR41 to participate in the interactions (Figs. 2B, 4A, and Fig. S7A)at the apoRas–SOS interface. Consistent with this structuralprediction, BZIM does not inhibit the nucleotide release or ex-change (Fig. 3B and Fig. S4 A–C). Interestingly, both DCAI andBZIM bind to KRas with similar affinity as measured by com-pound-induced chemical shift perturbations of Ras in NMR ex-periments (Kd of 1.1� 0.5 mM and 1.5� 0.3 mM for DCAI andBZIM, respectively), suggesting that binding to Ras itself is notsufficient to inhibit exchange. Our data indicate that the sterichindrance and/or disruption of salt bridges contribute to the in-hibition of the SOScat-catalyzed nucleotide exchange by DCAI.

To further elucidate the mechanism of DCAI function, wedeveloped biochemical assays to directly analyze the interactionbetween KRas and SOS. Biotinylated wild-type KRasGDP wasincubated with SOScat in the absence of free nucleotide with orwithout EDTA. The KRas-associated SOScat was purified onstreptavidin beads and assayed by Western blotting (Fig. 4C).In the absence of EDTA, KRasGDP and SOScat formed a complexthat represents the first intermediate of the nucleotide releasereaction, which is then converted into the apoRas–SOS complexupon nucleotide release (Fig. S3A). Addition of EDTA to the re-action chelates Mg2þ from the nucleotide-binding site in KRasand promotes the formation of the apoRas–SOS complex, a laterintermediate in the exchange reaction (Fig. S3A). We found thatDCAI, but not BZIM, compromised the formation of the Ras–SOS complex independent of the presence of EDTA (Fig. 4C andFig. S7 B and C). We conclude that DCAI inhibits the nucleotiderelease reaction by blocking the association of KRas with SOS.

DCAImayblock the formationof theKRasGDP-SOScat complex(step I) and/or inhibit the conversion of this complex to apoRas–SOS (step II) in the nucleotide release assay (Fig. S3A). If DCAI

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blocks step I in the reaction, we predict that the inhibitory effectof DCAI at a given compound concentration should be reducedas the SOS concentration increases. On the other hand, if DCAIblocks step II in the reaction, the inhibitory effect of DCAI shouldbe constant independent of the SOS concentration. In the nucleo-tide release assay, we titrated the SOS concentration in the pre-sence of fixed DCAI concentrations (either 125 or 250 μM) andobserved greater inhibition by DCAI at lower SOS concentrations(Fig. 4D), indicating that DCAI blocks the formation of theKRasGDP-SOScat complex. Thus, DCAI acts as a competitive inhi-bitor of nucleotide release via its association with RasGDP. Thismechanism of action forDCAI also predicts that the IC50 ofDCAIin the nucleotide release assay should be highly dependent on theconcentration of SOScat with lowerSOScat concentrations resultingin lower IC50 values. Titration of SOScat in the nucleotide releaseassay confirmed this prediction (Fig. S7D).

DCAI Inhibits Ras Activation in Cells. To measure the intracellularRasGTP level, we developed a biosensor system in HEK-293Tcells inducibly expressing Venus fluorescence protein fused to theRas-binding domain and cysteine-rich domain (RBD-CRD) do-main of cRaf (13). In live cell imaging experiments, we demon-strated that endogenous plasma membrane-associated RasGTP

recruited Venus-RBD-CRD to the membrane (13) (Fig. S8A;

time 0). DCAI disrupted this recruitment with EC50 values of15.8� 0.4 μM (Fig. S8 A–C). Neither BZIM nor INDL, whichare inactive in in vitro exchange assays, had any effect onRBD-CRD localization (Fig. S8 A and C).

This inhibition of membrane targeting of the RBD-CRD re-porter is not due to loss of Ras at the membrane, as DCAI hadno effect on the localization of fluorescently tagged Ras in HEK-293T cells (Fig. S8D). In addition, the inhibition by DCAI doesnot result from a general disruption of protein interactions at theplasma membrane, as the recruitment of Smad-7 by Smurf-1 atthe plasma membrane was not affected by DCAI (Fig. S8E).

We also examined the effect of DCAI on activation of Ras byEGF, a response critically dependent on SOS. Addition of EGFincreased the RasGTP level in HEK-293Tcells, whereas DCAI at-tenuated this EGF-stimulated activation of Ras (Fig. S8 FandG).

DiscussionThis paper describes our efforts to identify and characterizesmall molecules that inhibit Ras activity. Extensive efforts havebeen invested in the past two decades in the development of aneffective therapy against Ras-driven tumors. Initial work wasfocused on farnesyl transferase inhibitors (FTIs) and inhibitors ofRas converting enzme 1 and isoprenylcysteine methyltransferase,which disrupt the processing and localization of Ras. However,these strategies failed for the KRas mutant tumors because these

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enzymes have many substrates besides Ras and because KRas canbe modified by geranylgeranylation in the presence of FTIs.

More recent efforts have focused on directly targeting the Rasoncoprotein by (i) restoring the GTPase activity, which is greatlyreduced due to Ras mutations, (ii) blocking the formation ofRasGTP (i.e., inhibiting Ras-GEFs), or (iii) blocking Ras signalingto downstream effectors. The first approach failed despite exten-sive efforts; retrospective analysis of the mutant Ras-GAP struc-ture indicates a lack of any small-molecule binding space at theGTPase active site (14). Targeting Ras signaling to effectors andinhibition of Ras nucleotide exchange both require disruption ofprotein–protein interactions, which is challenging.

FBLD represents an effective approach to probe protein sur-face using low molecular weight but highly soluble compounds.Through FBLD, we identified a group of compounds that bindto a site adjacent to the functionally important switch I/II regionsof KRas. This binding site exists on part of the Ras surface com-monly involved in Ras-effector, Ras-GEFs, and Ras-GAP inter-actions (11, 14, 15). The fact that all 25 confirmed hits from theNMR screen of 3,300 fragments bind to the same site suggeststhat this site is likely to be the primary compound-binding siteon Ras. High-resolution crystal structures elucidate the specificmode of molecular interactions and delineate a well-defined siteresponsible for binding.

Although several previous studies have reported potentialsmall-molecule association with Ras, none have characterizedthe binding site by a high-resolution structure analysis and nonereported a defined binding pocket (3–7). The best-characterizedRas-binding compounds reported so far are SCH54292 (16) andits more water-soluble derivative (7). In silico calculation basedon NMR mapping suggests that these compounds bind to aregion between switch II and helix 4 of RasGDP, although otherbinding areas cannot be ruled out. This hypothesized SCH54292-

binding site is distinct from the DCAI-binding pocket that wedetermined.

An intriguing feature of the DCAI-binding pocket is its abilityto expand upon compound binding and the functional impor-tance of such compound-induced conformational change. Alter-ing rotamer conformation of KRas R41 and D54 removes twohydrogen bonds at the interface between Ras and SOS, whichlikely contributes to the DCAI-mediated inhibitory activity. ADP-ribosylation on R41 has been reported to interfere with SOS-cat-alyzed nucleotide exchange on Ras (17), and we speculate thatADP-ribosylation on R41 also compromises the salt bridge withSOS. Interestingly, sequence and structure homology betweenSOS1 and RasGRF1 (18), another exchange factor, suggests thatthese hydrogen bonds are likely conserved in the complex of Rasand RasGRF1 as well, and therefore DCAI may inhibit otherRas-GEFs.

DCAI not only binds to Ras, but also blocks the interactionof Ras as a substrate with SOS (Fig. S7E). RasGTP also binds toan allosteric site on SOS1 and up-regulates its nucleotide ex-change activity (19). DCAI does not appear to affect the bindingof Ras to the allosteric site (Fig. S9A). Functionally, DCAI blocksthe SOS-mediated nucleotide release and inhibits the activationof Ras. Consistent with this specific mechanism of action, the in-hibitory effect of DCAI is only toward SOS-catalyzed nucleotideexchange and does not affect the intrinsic nucleotide exchangeby Ras. This mode of action is in sharp contrast to that of aRac exchange inhibitor, EHT 1864, which inhibits both intrinsicand GEF-mediated nucleotide exchange for Rac (20). Patgiriet al. recently described an SOS1-derived peptide that inhibitedRas signaling (21). Although this peptide and DCAI interact withdifferent regions of Ras, both inhibit Ras nucleotide release invitro and Ras activation in vivo.

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Fig. 4. DCAI is a competitive inhibitor that blocks the Ras-SOScat interaction. (A) Ras coordinates from the DCAI-KRasm and BZIM-KRasm complexes weresuperimposed onto those from the HRas–SOS complex (1NVV; HRas bound to the substrate site of SOS; ref. 19) to highlight the locations of the DCAIand BZIM binding sites. The carbon atoms of Ras R41 and D54 are colored in green in the Ras–SOS complex and in yellow in the Ras–DCAI complex. (B) Nucleo-tide release assays were performed with wild-type RasGDP (black) or Ras-R41SGDP (red) across a range of SOScat concentrations. SOScat is less effective in cat-alyzing nucleotide release from Ras-R41Smant-GDP. (C, Top) Nucleotide-free biotinylated KRas was generated by incubation with EDTA. Streptavidin beads wereincubated with SOScat alone (lane 1; negative control) or with SOScat plus nucleotide-free biotinylated KRas in the presence of increasing concentrations ofDCAI (lanes 3–7). Bead-associated SOScat was purified, washed, and detected by Western blotting. (Bottom) Identical reactions as above, with the exceptionthat EDTA was omitted in the reaction. Results shown are representative of three independent experiments. (D) Nucleotide release assays were performed, induplicates, with a dose titration of SOScat in the presence of DMSO or of either 125 or 250 μMDCAI. The relative reaction rates were determined by normalizingto those of DMSO control, and the percentage of inhibition by DCAI was plotted for each concentration of SOScat. Error bars in B and D are the SEM.

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DCAI exhibits cellular activity in HEK-293Tcells; it attenuatesthe EGF-stimulated activation of Ras and blocks the recruitmentof the cRaf RBD-CRD domain to the cytoplasmic membrane.Given that DCAI binds to Ras and acts as a competitive inhibitoragainst association of SOS with Ras, it is expected that its potencyin cellular assays is in part determined by the cellular concentra-tions of SOS and Ras. Quantitative analysis determined the cel-lular SOS1 and Ras concentrations at 5 and 350 nM in HEK-293Tcells, respectively. The EC50 of DCAI in the RBD-CRD redistri-bution assay is at 15.8� 0.4 μM (Fig. S8C), which is consistentwith the inhibitory potency of DCAI in the in vitro nucleotiderelease reaction (Fig. S7D). Although these studies suggest thatDCAI inhibits the nucleotide exchange via binding on Ras incells, we cannot exclude the possibility of off-target driven effects,given its low biochemical affinity to Ras. In addition, our cellularassays in Fig. S8 measure RasGTP level via interaction with RBDor RBD-CRD of cRaf, and it is possible that the cellular activitywe observed results from the inhibition of both Ras nucleotideexchange and Ras–Raf interaction. Superimposing the Ras-DCAI and the Ras-effector complexes suggests that DCAI bind-ing does not alter the protein–protein interface and that DCAIshould exert minimal effects on the Ras/effector interactions(Fig. S9 B and C).

Through an integrated approach of fragment-based small-molecule screening, high-resolution structural analyses andfunctional studies, we have identified a unique small-moleculebinding pocket and elucidated its functional role in inhibitingRas activation. The identified fragment hits have low affinity toRas, and our efforts represent only the very first step toward de-veloping a lead compound. Although further medicinal chemistryefforts are needed to enhance potency while controlling selectiv-ity, the current results may reinvigorate the search for therapeu-

tics of this challenging and yet important oncology target (seeSI Text for discussion). Interestingly, additional small G proteinsshare similar structural features. Thus, the FBLD approachrepresents a potential route to target these small G proteinswith small-molecule inhibitors, thereby opening up therapeuticopportunities for this class of biologically important regulatoryproteins.

Materials and MethodsFragment Screening and Hit Validation. STD experiments were performed at280 K in pools of up to six compounds each at a concentration of 250 μM percompound. The pools of compounds were incubated with a 1∶1 mixture ofKRasmGDP and KRasmGMPPCP (5 μM each) in 20 mM d11-tris (hydroxymethyl)aminomethane hydrochloride pH 8.0, 200 mM NaCl, 2 mM MgCl2, 99%2H2O, and 50 μM 4,4-dimethyl-4-silapentane-1-sulfonic acid. Compoundswith STD signal-to-noise ratio greater than five were remeasured as indivi-dual compounds for binding to KRasmGDP or KRasmGMPPCP separately.

Hits were further tested for specific binding to KRasm by measurement of1H15N HSQC protein spectra. Positive hits were defined as compounds thatexhibit cross-peak perturbation by more than 15 Hz (combined 15N and 1Hchemical shift change) when compared to the spectrum recorded in theabsence of compounds.

Crystallization, Structure Determination, and Biochemical and Cellular Assays.See SI Materials and Methods for details.

ACKNOWLEDGMENTS. We thank Dr. Sharon Campbell (University of NorthCarolina, NC) for helpful discussions on Ras biochemistry. We are gratefulto Ken Dong, Yvonne Franke, William Forrest, Imola Fodor, Janet Jin, JuliaKong, Hong Li, Maria Lorenzo, Kyle Mortara, Olivier René, Qinghua Song,and Christine Tam for technical contributions. We thank Reciprocal SpaceConsulting (Berkeley, CA) for collecting diffraction data and the StanfordSynchrotron Facility and the Advanced Light Source for radiation lightsources.

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