Significance Despite decades of research, no approved drugs have been discovered for KRAS. Recently, a pocket occurring on the surface of the active and inactive form of KRAS was found, but, due to its comparatively shallow, polar nature, this pocket has been assumed to be “undruggable.” Starting from very weakly binding fragments and using structure-based drug design, we discovered BI-2852 (1), a nanomolar inhibitor to this pocket which is mechanistically distinct to covalent KRASG12C inhibitors; 1 modulates pERK and pAKT and has an antiproliferative effect in KRAS mutant cells. This work demonstrates the druggability of this so-called switch I/II pocket and provides the scientific community with a chemical probe that directly inhibits the active and inactive forms of KRAS.

Abstract The 3 human RAS genes, KRAS, NRAS, and HRAS, encode 4 different RAS proteins which belong to the protein family of small GTPases that function as binary molecular switches involved in cell signaling. Activating mutations in RAS are among the most common oncogenic drivers in human cancers, with KRAS being the most frequently mutated oncogene. Although KRAS is an excellent drug discovery target for many cancers, and despite decades of research, no therapeutic agent directly targeting RAS has been clinically approved. Using structure-based drug design, we have discovered BI-2852 (1), a KRAS inhibitor that binds with nanomolar affinity to a pocket, thus far perceived to be “undruggable,” between switch I and II on RAS; 1 is mechanistically distinct from covalent KRASG12C inhibitors because it binds to a different pocket present in both the active and inactive forms of KRAS. In doing so, it blocks all GEF, GAP, and effector interactions with KRAS, leading to inhibition of downstream signaling and an antiproliferative effect in the low micromolar range in KRAS mutant cells. These findings clearly demonstrate that this so-called switch I/II pocket is indeed druggable and provide the scientific community with a chemical probe that simultaneously targets the active and inactive forms of KRAS.

The 3 human RAS genes, KRAS, NRAS, and HRAS, encode 4 different RAS proteins (KRAS-4A, KRAS-4B, NRAS, and HRAS) which belong to the protein family of small GTPases that function as binary molecular switches involved in cell signaling (1). Activating mutations in RAS like the glycine 12 mutations are among the most common oncogenic drivers in human cancers. KRAS is the most frequently mutated oncogene, with mutation rates of 86 to 96% in pancreatic cancers (2), 40 to 54% in colorectal cancers (3), and 27 to 39% in lung adenocarcinomas (4). NRAS is predominantly mutated in melanoma and hematological malignancies (5, 6), while HRAS mutations are found in salivary gland and urinary tract cancers (7, 8).

The RAS family is known to cycle through 2 different conformational states that are defined by differential binding to nucleotides. In the “off” state, RAS proteins are bound to the nucleotide guanosine diphosphate (GDP), while in the “on” state they are bound to the nucleotide guanosine triphosphate (GTP). The γ-phosphate of GTP holds 2 regions, switch I and switch II (9), in a compact conformation that allows interaction with downstream effectors, such as CRAF, PI3Kα, and RALGDS, as well as with the allosteric site of SOS1 and SOS2. Hydrolysis of the γ-phosphate to produce GDP-RAS causes a conformational change in the switch regions, leading to the formation of an inactive state which is unable to bind effector molecules (10, 11). RAS itself has an intrinsic, but weak, GTPase activity that is enhanced by GTPase-activating proteins (GAPs) catalyzing RAS inactivation. The exchange of the bound nucleotide GDP into GTP is facilitated by guanine nucleotide exchange factors (GEFs) which, in the case of KRAS, is performed by SOS1 and SOS2 (12). GEFs catalyze the release of GDP from RAS in the cytoplasm and replace it with the more abundant intracellular GTP. Oncogenic mutations in RAS impair GTP hydrolysis, leading to stabilization of the activated GTP-RAS form and enhanced RAS signaling. The most common mutations occur as single-point mutations at codons 12, 13, and 61 (13).

Although KRAS could serve as an excellent drug target for many cancers, direct inhibition of oncogenic RAS has proven to be challenging. Despite decades of research, no therapeutic agent directly targeting RAS has been clinically approved. The main reason for this is the lack of druggable pockets on the surface of RAS. However, in recent years, there has been a resurgence of research around RAS, driven by the growing belief that RAS might be able to be drugged with low molecular weight organic molecules. This belief was sparked by the discovery of 2 pockets on the surface of RAS that could potentially be amenable to small-molecule drug discovery. The S.W.F. group at Vanderbilt (14), researchers at Genentech (15), and, more recently, the Rabbitts group (16, 17) discovered small molecules that bind to a shallow pocket between the switch I and II regions of KRAS. This pocket will be referred to as the switch I/II pocket (SI/II-pocket). In addition, the Shokat group discovered covalently linked small molecules which bind to a second pocket on RAS positioned above the switch II loop in GDP-KRASG12C, called the switch II pocket (SII-pocket) (11).

In this paper, we describe the discovery of nanomolar inhibitors that directly target the small, polar SI/II-pocket present on both the active and inactive form of KRAS. To discover small molecules that bind to KRAS, we conducted several fragment-based screens using uniformly 15N-labeled guanosine-5′-[(β,γ)-methyleno]triphosphate (GCP)-bound KRASG12D for validation. From these screens, we identified fragments that weakly bind to GCP-KRASG12D that were optimized using structure-based design. This was accomplished by developing a robust system for crystallizing small molecules bound to GTP-KRASG12D. The most potent KRAS inhibitor, BI-2852 (1), binds with nanomolar affinity to the active and inactive form of KRAS. Compound 1 blocks the interaction between GDP-KRAS and the catalytic site of SOS1, but, in contrast to covalent KRASG12C inhibitors, also inhibits the interactions between GTP-KRAS and the allosteric site of SOS1 as well as its effectors (CRAF and PI3Kα). In cells, 1 inhibits SOS1-catalyzed exchange of GDP to GTP as well as GAP-catalyzed exchange of GTP to GDP, which results in no net change in cellular GTP-RAS levels upon treatment. Compound 1 reduced pERK and pAKT levels in a dose-dependent manner, leading to an antiproliferative effect in NCI-H358 cells. The effects of 1 were confirmed to be KRAS-driven and not unspecific effects, through the consistent data generated for the 10-fold weaker distomer 44. Compound 1 demonstrates that the SI/II-pocket is indeed druggable and provides an ideal starting point for the design of more potent and selective RAS inhibitors. Compound 1 will also serve as a useful chemical probe for the scientific community in the study of RAS biology of simultaneous inhibition of active and inactive RAS in an in vitro setting.

Discussion Here, we describe the discovery of 1, an inhibitor of both the active and inactive form of KRAS, with nanomolar binding affinity to the SI/II-pocket, a small, shallow, and polar pocket deemed by many to be “undruggable.” Compound 1 is the first RAS inhibitor for the SI/II pocket with KRAS-driven cellular activity, displaying low micromolar pERK modulation and antiproliferative effects on a KRAS mutant cell line. Recent compounds (16, 17) claiming cellular activity do not provide negative control data and display antiproliferative effects under 2D cell culture conditions which should not be interpreted as KRAS-driven, given that KRAS antiproliferative effects are predominantly only observed under 3D, nonadherent conditions (35). Fragment screens delivered hits in the millimolar binding affinity range which were optimized using structure-based design. Although highly resource-intensive, the application of NMR to measure dissociation constants for newly designed compounds in the millimolar range served as an important method for establishing structure activity relationships. Guided by the X-ray structures of cocomplexes, we were able to optimize binding, as evidenced by the discovery of the isoindolinone 18, which bound to GCP-KRASG12D at 20 µM by ITC and also displayed inhibition of the key PPI with KRAS in a similar range. This allowed us to switch from NMR K D measurements to biochemical PPI assays to further optimize the potency of the compounds. Obtaining 3D crystallographic information on RAS proteins in the active form has been historically limited. The development of a reliable procedure to produce >10 mg/L of purified GCP-KRAS was instrumental in enabling crystallography, which, in turn, revealed critical information on the binding of the ligands to the SI/II-pocket of GCP-KRAS. Compound 1 maintained the polar interactions to D54 and E37 also addressed by 18 and, in addition, formed a H bond to S39 and displaced 3 water molecules, which are presumably responsible for the >100-fold improvement in potency. It should be noted that T74, which improves potency by 5- to 10-fold (SI Appendix, Tables S1 and S2), is not yet addressed by 1 and that 1 still contains 7 rotatable bonds. This highlights the potential for significant improvement beyond the current potency of 1 (e.g., IC 50 of 180 nM for the PPI between active KRASG12D and CRAF) and indicates that the SI/II-pocket is indeed druggable. Triple RAS knockout mice are not embryonically viable but can be rescued by reintroduction of an HRAS transgene, indicating functional redundancy among the RAS family (39) and suggesting that sparring at least one wild-type RAS isoform will be needed for a RAS drug. As the SI/II-pocket is conserved on both the inactive and active forms of all RAS isoforms, obtaining sufficient selectivity presents an additional significant challenge to drugging this pocket. Interestingly, 1 demonstrates a 10-fold selectivity of binding to GCP-KRASwt (SI Appendix, Table S5) which translates to a 4-fold selectivity of inhibition biochemically (inhibition of KRASwt versus KRASG12C binding to CRAF) (SI Appendix, Table S6). The relative lack of selectivity versus the KRASG12D::CRAF is expected due to the 5-fold weaker affinity of KRASG12D for CRAF, while KRASG12C and KRASwt maintain the same affinity for the RAS binding domain of CRAF (40). Also, a weaker affinity to GDP-NRAS was observed for 1. Together, this suggests that, despite the high conservation of the SI/II-pocket, it might be possible to design molecules with sufficient RAS isoform selectivity. The SI/II-pocket is involved in interactions with GEFs (41), GAPs (42), and downstream effectors (43, 44), and we provide evidence that compound 1 inhibits all of these PPIs. Functionally, 1 inhibits SOS1-catalyzed exchange of GDP-KRAS to GTP-KRAS as well as GAP-catalyzed exchange of GTP-KRAS to GDP-KRAS, which results in no net change in cellular GTP-RAS levels upon treatment. E37 on switch II, to which the phenolic oxygen of 1 H-bonds, is also an important residue for RAS binding to downstream effectors (45), GEFs (41, 46), and GAPs (47), explaining mechanistically how 1 inhibits the binding of multiple key RAS interactions partners. Compound 1 reduced pERK and pAKT levels in a dose-dependent manner in NCI-H358 cells, leading to an antiproliferative effect in NCI-H358 cells under nonadherent, low serum conditions. The effects of 1 were confirmed to be KRAS-driven and not off-target through the consistent data generated for the 10-fold less active distomer 44 and through the absence of any effects on BRAF(V600E) cell lines. We expect BI-2852 to serve as a useful chemical probe for the study of RAS biology in an in vitro setting, and it is available to the scientific community (https://opnme.com/molecules/kras-bi-2852). BI-2852 is also an ideal starting point for the design of more-potent and selective RAS inhibitors.

Acknowledgments We thank Andreas Bergner, Helmut Berger, Matthias Klemencic, Norbert Kraut, Erik Patzelt, Jens Quant, Michaela Streicher, Diane Thompson, Ingrid Vorwahlner, Anika Weiss, and Piro Lito. Funding to support this work came from US National Institutes of Health (NIH) Grants P50A095103 (National Cancer Institute Specialized Programs of Research Excellence in Gastrointestinal Cancer) and RC2CA148375 (NIH American Recovery and Reinvestment Act Stimulus Grant), and through grants from the Lustgarten Foundation for Pancreatic Cancer Research to S.W.F. Additional Austrian governmental funding was provided by the Austrian Forschungsförderungsgesellschaft through Grants 854341 and 861507 (Basisprogramme). Use of the Advanced Photon Source, an Office of Science User Facility operated for the US Department of Energy Office of Science by the Argonne National Laboratory, was supported by US Department of Energy Contract DE-AC02-06CH11357. In addition, the X06SA beamline at the Swiss Light Source, Paul Scherrer Institut, Villigen, Switzerland, was used for crystallographic measurements, with special thanks to Dirk Reinert and Expose GmbH for data measurement. We thank the Vanderbilt High-Throughput Screening (HTS) Core, in which some of these experiments were performed. The HTS Core receives support from the Vanderbilt Institute of Chemical Biology and the Vanderbilt-Ingram Center (NIH Grant P30CA68485).

Footnotes Author contributions: D.K., M.G., A.M., L.J.M., A.Z., M.M., A.G., M.K., J.P., J.R., R.S., A.W., M.Z., M.P., S.W.F., and D.B.M. designed research; D.K., M.G., A.M., L.J.M., A.Z., M.M., A.G., D.C., S.F., T. Gerstberger, T. Gmaschitz, C.G., P.G., D.H., W.H., J.H., J.K.-O., P.K., S.K., M.K., R.K., L.L., F.M., S.M.-M., C.P., J.P., J.S., C.S., Y.S., K.S., R.S., A.S., B.S., G.S., Q.S., B.W., and M.Z. performed research; D.C., S.F., T. Gerstberger, T. Gmaschitz, C.G., P.G., D.H., W.H., J.H., J.K.-O., P.K., S.K., R.K., L.L., F.M., S.M.-M., C.P., J.P., J.S., C.S., Y.S., K.S., A.S., B.S., G.S., Q.S., and B.W. contributed new reagents/analytic tools; D.K., M.G., A.M., L.J.M., A.Z., M.M., A.G., D.C., S.F., T. Gerstberger, T. Gmaschitz, C.G., P.G., D.H., W.H., J.H., J.K.-O., P.K., S.K., M.K., R.K., L.L., F.M., S.M.-M., C.P., J.P., J.R., J.S., C.S., Y.S., K.S., R.S., A.S., B.S., G.S., Q.S., A.W., B.W., M.Z., M.P., S.W.F., and D.B.M. analyzed data; and D.K., M.G., A.M., L.J.M., A.Z., M.M., M.P., and D.B.M. wrote the paper.

Conflict of interest statement: D.K., M.G., A.M., L.J.M., A.Z., M.M., A.G., D.C., S.F., T. Gerstberger, T. Gmashitz, P.G., D.H., W.H., J.H., J.K.-O., P.K., S.K., M.K., R.K., L.L., F.M., S.M.-M., C.P., J.R., C.S., Y.S., K.S., R.S., A.S., B.S., G.S., B.W., M.Z., M.P., and D.B.M. were employees of Boehringer Ingelheim at the time of this work.

This article is a PNAS Direct Submission.

Data deposition: The atomic coordinates and structure factors have been deposited in the Protein Data Bank, www.wwpdb.org (PDB ID codes 6GJ5, 6GJ6, 6GJ7, and 6GJ8).

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