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Mutations in the α4-α5 allosteric lobe of RAS do not significantly impair RAS signaling or self-association

  • Author Footnotes
    + These authors contributed equally to this work.
    Michael Whaby
    Footnotes
    + These authors contributed equally to this work.
    Affiliations
    Department of Cell and Molecular Pharmacology & Experimental Therapeutics, Medical University of South Carolina, Charleston, SC 29425, USA

    Hollings Cancer Center, Medical University of South Carolina, Charleston, SC 29425, USA
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  • Author Footnotes
    + These authors contributed equally to this work.
    Lauren Wallon
    Footnotes
    + These authors contributed equally to this work.
    Affiliations
    Department of Biochemistry and Molecular Pharmacology, New York University Langone Health, New York University School of Medicine, New York, NY 10016, USA
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  • Megan Mazzei
    Affiliations
    Department of Cell and Molecular Pharmacology & Experimental Therapeutics, Medical University of South Carolina, Charleston, SC 29425, USA

    Hollings Cancer Center, Medical University of South Carolina, Charleston, SC 29425, USA
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  • Imran Khan
    Affiliations
    Department of Cell and Molecular Pharmacology & Experimental Therapeutics, Medical University of South Carolina, Charleston, SC 29425, USA

    Hollings Cancer Center, Medical University of South Carolina, Charleston, SC 29425, USA

    Ralph H. Johnson VA Medical Center, Charleston, SC. 29401, USA
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  • Kai Wen Teng
    Affiliations
    Perlmutter Cancer Center, New York University Langone Health, New York University School of Medicine, New York, NY 10016, USA
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  • Shohei Koide
    Correspondence
    Corresponding authors: Shohei Koide.
    Affiliations
    Department of Biochemistry and Molecular Pharmacology, New York University Langone Health, New York University School of Medicine, New York, NY 10016, USA

    Perlmutter Cancer Center, New York University Langone Health, New York University School of Medicine, New York, NY 10016, USA
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  • John P. O’Bryan
    Correspondence
    Corresponding authors: John P. O’Bryan.
    Affiliations
    Department of Cell and Molecular Pharmacology & Experimental Therapeutics, Medical University of South Carolina, Charleston, SC 29425, USA

    Hollings Cancer Center, Medical University of South Carolina, Charleston, SC 29425, USA

    Ralph H. Johnson VA Medical Center, Charleston, SC. 29401, USA
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  • Author Footnotes
    + These authors contributed equally to this work.
Open AccessPublished:November 02, 2022DOI:https://doi.org/10.1016/j.jbc.2022.102661

      Abstract

      Mutations in one of the three RAS genes (HRAS, KRAS, and NRAS) are present in nearly 20% of all human cancers. These mutations shift RAS to the GTP-loaded active state due to impairment in the intrinsic GTPase activity and disruption of GAP-mediated GTP hydrolysis, resulting in constitutive activation of effectors such as RAF. Because activation of RAF involves dimerization, RAS dimerization has been proposed as an important step in RAS-mediated activation of effectors. The α4-α5 allosteric lobe of RAS has been proposed as a RAS dimerization interface. Indeed, the NS1 monobody, which binds the α4-α5 region within the RAS G domain, inhibits RAS-dependent signaling and transformation as well as RAS nanoclustering at the plasma membrane. Although these results are consistent with a model in which the G domain dimerizes through the α4-α5 region, the isolated G domain of RAS lacks intrinsic dimerization capacity. Furthermore, prior studies analyzing α4-α5 point mutations have reported mixed effects on RAS function. Here, we evaluated the activity of a panel of single amino acid substitutions in the α4-α5 region implicated in RAS dimerization. We found that these proposed “dimerization-disrupting” mutations do not significantly impair self-association, signaling, or transformation of oncogenic RAS. These results are consistent with a model in which activated RAS protomers cluster in close proximity to promote the dimerization of their associated effector proteins (e.g., RAF) without physically associating into dimers mediated by specific molecular interactions. Our findings suggest the need for a nonconventional approach to developing therapeutics targeting the α4-α5 region.

      Introduction

      RAS GTPases are important mediators of intracellular signaling cascades that regulate cell proliferation and survival (
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      ). Additionally, the K13 and K19 DARPins inhibit KRAS via binding of the α3-α4 allosteric lobe and disrupting RAS dimer/nanoclusters (
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      ). The importance of RAS dimers/nanoclusters was further highlighted by the endogenous RAS antagonist, DIRAS3, which inhibits RAS nanoclusters through binding of α5 region of RAS (
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      • Spencer-Smith R.
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      ,
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      • Marwood R.
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      • Khan I.
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      • Khan I.
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      ).
      Although these examples implicate RAS dimerization as an on-pathway event in RAS-mediated signaling and provide a rationale for drug discovery efforts aimed at development of RAS dimerization inhibitors, the importance of RAS dimerization remains a point of much debate. While active RAS stimulates the dimerization and activation of RAF kinases in the mitogen-activated protein kinase (MAPK) pathway (
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      ). Furthermore, the isolated HVR of KRAS, devoid of the G domain, is sufficient to drive dimerization of a fluorescent protein–HVR fusion protein in cells suggesting that KRAS G-domain is dispensable for dimerization (
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      • Pluckthun A.
      • Janne P.A.
      • Westover K.D.
      • Shan Y.
      • Shaw D.E.
      A structural model of a Ras-Raf signalosome.
      ,
      • Lee K.Y.
      • Fang Z.
      • Enomoto M.
      • Gasmi-Seabrook G.
      • Zheng L.
      • Koide S.
      • Ikura M.
      • Marshall C.B.
      Two Distinct Structures of Membrane-Associated Homodimers of GTP- and GDP-Bound KRAS4B Revealed by Paramagnetic Relaxation Enhancement.
      ,
      • Lee K.Y.
      • Enomoto M.
      • Gebregiworgis T.
      • Gasmi-Seabrook G.M.C.
      • Ikura M.
      • Marshall C.B.
      Oncogenic KRAS G12D mutation promotes dimerization through a second, phosphatidylserine-dependent interface: a model for KRAS oligomerization.
      ). Here, we show that mutations in the α4-α5 allosteric lobe do not affect RAS association in cells. In addition, these mutants did not impair MAPK activation compared with the parental oncogenic RAS. Although downstream signaling was unaffected by these mutations, we observed isoform-specific differences in the transforming activity of selected mutants. Overall, our results are consistent with a model in which RAS protomers associate in close proximity to promote effector dimerization (e.g., RAF) without formation of molecularly defined RAS dimers. Further, we propose that the ability of selected biologics such as NS1 monobody and K13/K19 DARPins to perturb RAS function is due to steric hinderance of RAS rather than disruption of bona fide dimers. This study provides further insights into RAS oligomerization and should inform efforts in developing therapeutics directed at the α4-α5 allosteric lobe of RAS.

      Results

      Mutations in the α4-α5 region of KRAS do not impair oncogenic activity

      Given the contradictory reports on the importance of dimerization in RAS function, we tested whether mutations in the proposed dimerization interface (Fig. 1A and 1B; termed α4-α5 mutants hereafter) impaired MAPK pathway activation relative to the parental oncogenic KRAS mutant. To reduce overexpression artifacts, we determined the appropriate amounts of K/H/NRAS DNA that yielded protein expression and ERK phosphorylation below the point of saturation for the signaling assays (Fig. S1–S4). Surprisingly, there were no significant differences in MAPK activation between the KRASG12V α4-α5 mutants and the parental oncogenic KRASG12V with the exception of KRASG12V/K147D, a mutation that has been shown to decrease RAS-GTP levels (Fig.2A, 2B, and S5) (
      • Baker R.
      • Wilkerson E.M.
      • Sumita K.
      • Isom D.G.
      • Sasaki A.T.
      • Dohlman H.G.
      • Campbell S.L.
      Differences in the regulation of K-Ras and H-Ras isoforms by monoubiquitination.
      ,
      • Sasaki A.T.
      • Carracedo A.
      • Locasale J.W.
      • Anastasiou D.
      • Takeuchi K.
      • Kahoud E.R.
      • Haviv S.
      • Asara J.M.
      • Pandolfi P.P.
      • Cantley L.C.
      Ubiquitination of K-Ras enhances activation and facilitates binding to select downstream effectors.
      ).
      Figure thumbnail gr1
      Figure 1A model of an α4-α5 dimer of KRAS and the locations of residues tested in this study. A) The model was built by fitting the structure of KRAS4B•GMPPNP (residues 1-169; PDB ID 6VC8; (
      • Ingólfsson H.I.
      • Neale C.
      • Carpenter T.S.
      • Shrestha R.
      • López C.A.
      • Tran T.H.
      • Oppelstrup T.
      • Bhatia H.
      • Stanton L.G.
      • Zhang X.
      • Sundram S.
      • Di Natale F.
      • Agarwal A.
      • Dharuman G.
      • Kokkila Schumacher S.I.L.
      • Turbyville T.
      • Gulten G.
      • Van Q.N.
      • Goswami D.
      • Jean-Francois F.
      • Agamasu C.
      • Chen
      • Hettige J.J.
      • Travers T.
      • Sarkar S.
      • Surh M.P.
      • Yang Y.
      • Moody A.
      • Liu S.
      • Van Essen B.C.
      • Voter A.F.
      • Ramanathan A.
      • Hengartner N.W.
      • Simanshu D.K.
      • Stephen A.G.
      • Bremer P.T.
      • Gnanakaran S.
      • Glosli J.N.
      • Lightstone F.C.
      • McCormick F.
      • Nissley D.V.
      • Streitz F.H.
      Machine learning-driven multiscale modeling reveals lipid-dependent dynamics of RAS signaling proteins.
      )) onto the crystallographic dimer of HRAS•GMPPNP (residues 1-166; PDB ID 5P21; (
      • Pai E.F.
      • Krengel U.
      • Petsko G.A.
      • Goody R.S.
      • Kabsch W.
      • Wittinghofer A.
      Refined crystal structure of the triphosphate conformation of H-ras p21 at 1.35 A resolution: implications for the mechanism of GTP hydrolysis.
      )). The Switch I and II regions are colored in tan. The α4 and α5 helices in protomer 1 are labeled, and those in protomer 2, i.e., the symmetry-related copy, are labeled as α4’ and α5’. The residues subjected to mutational studies are shown in cyan and labeled for protomer 1 and shown in blue for protomer 2. The side chains of E49, K128 and R135 are disordered in the 6VC8 model and thus not depicted. B) RAS α4-α5 mutants included in this study.
      Next, we analyzed CRAF–BRAF association in cells expressing KRASG12V versus KRASG12V harboring α4-α5 mutants, given the well-established role of RAF dimerization in MAPK signaling (
      • Rajakulendran T.
      • Sahmi M.
      • Lefrancois M.
      • Sicheri F.
      • Therrien M.
      A dimerization-dependent mechanism drives RAF catalytic activation.
      ,
      • Freeman A.K.
      • Ritt D.A.
      • Morrison D.K.
      Effects of Raf dimerization and its inhibition on normal and disease-associated Raf signaling.
      ). In addition, a previous study demonstrated that D154Q and R161D RAS mutants decreased CRAF–BRAF heterodimers (
      • Ambrogio C.
      • Köhler J.
      • Zhou Z.W.
      • Wang H.
      • Paranal R.
      • Li J.
      • Capelletti M.
      • Caffarra C.
      • Li S.
      • Lv Q.
      • Gondi S.
      • Hunter J.C.
      • Lu J.
      • Chiarle R.
      • Santamaría D.
      • Westover K.D.
      • Jänne P.A.
      KRAS Dimerization Impacts MEK Inhibitor Sensitivity and Oncogenic Activity of Mutant KRAS.
      ). In agreement with the results from the MAPK signaling assays described above, there was no significant impairment in CRAF–BRAF interaction in cells expressing the KRAS α4-α5 mutants compared with parental KRASG12V (Fig. 2C and 2D). These data indicate that the α4-α5 mutations do not impair the activation of the canonical RAS/MAPK pathway mediated by KRASG12V.
      Figure thumbnail gr2
      Figure 2Mutations in the of α4-α5 allosteric lobe of KRAS do not impair oncogenic activity. A) ERK/MAPK activity assay in HEK 293 cells co-transfected with HA-tagged KRASG12V mutants and MYC-tagged ERK. B) Normalized pERK signal from panel (A). C) Immunoprecipitation (IP) of endogenous CRAF in HEK 293 cells transfected with KRASG12V mutants. D) Normalized BRAF signal from the CRAF IP in panel (C). E) NIH/3T3 transformation assay in cells transfected with KRASG12V mutants. F) Normalized foci number from the NIH/3T3 transformation assays in panel (E). All experiments were repeated at least three times (n=3) and results quantified using Welch’s t-test; error bars representing SEM (***p<0.0005, **p<0.005, and *p<0.05).
      Lastly, we performed transformation assays in NIH/3T3 cells transfected with KRASG12V or the KRASG12V α4-α5 mutants (Fig. 2E and 2F). Consistent with the signaling data, the KRASG12V α4-α5 mutants retained the ability to transform cells as well as parental KRASG12V. Taken together, these results suggest that mutations of amino acid residues proposed to be critical for KRAS–KRAS self-association do not significantly impair the signaling or transforming properties of oncogenic KRAS.

      Mutations in the α4-α5 allosteric lobe do not affect KRAS association in cells

      Next, we addressed whether these α4-α5 mutations affected RAS–RAS association in cells. We employed Live-Cell NanoLuc® Binary Technology (NanoBiT), a protein–protein interaction (PPI) detection system where one protein partner is tagged with an 11-amino acid peptide (SmBiT) while the other protein partner is tagged with a 17.6 kDa NanoLuc fragment (LgBiT). When expressed in cells, PPIs between the two protein partners allows for complementation of the SmBiT and LgBiT tags to generate a luminescent signal upon substrate addition.
      To avoid potential interference of endogenous RAS proteins with the SmBiT/LgBiT-tagged RAS protein partners, we performed these assays in RAS-less MEFs transformed with BRAFV600E, a cell line that lacks all the RAS isoforms (
      • Drosten M.
      • Dhawahir A.
      • Sum E.Y.
      • Urosevic J.
      • Lechuga C.G.
      • Esteban L.M.
      • Castellano E.
      • Guerra C.
      • Santos E.
      • Barbacid M.
      Genetic analysis of Ras signalling pathways in cell proliferation, migration and survival.
      ). To account for variations between transfections, cells were lysed after measuring the live-cell luminescence (Fig S8A), and LgBiT-tagged proteins were quantified using HiBiT, an 11 amino acid peptide with high affinity (KD = 0.7 nM) for the LgBiT peptide. The HiBiT peptide out-competes the SmBiT peptide for LgBiT binding while still generating luminescent signal, allowing for a fast and sensitive method to quantify total LgBiT peptide levels in cells (Fig. 3A and S8B). When co-expressed in cells, SmBiT-KRASG12V and LgBiT-KRASG12V reconstituted luciferase activity (Fig. 3A and 3B). In contrast, co-expression of SmBiT-KRASG12V with EGFR-LgBiT, also a membrane localized protein which served as a negative control, generated a weak luminescent signal (Fig. 3B). The introduction of the α4-α5 mutations to LgBiT-KRASG12V resulted in no significant decreases in luciferase activity (Fig. 3B). These results are consistent with the results from signaling assays (Fig. 2), indicating that not only do the α4-α5 mutations have little to no effect on KRAS signaling and biology, they also do not impair RAS–RAS interactions in cells.
      Figure thumbnail gr3
      Figure 3Mutations in the α4-α5 allosteric lobe of RAS do not affect RAS–RAS association in cells. A) Illustration of the workflow for the NanoBiT assay used to measure PPIs in live cells. B) NanoBiT assay in RAS-less MEFs (BRAFV600E) co-expressing SmBiT-KRASG12V and LgBiT-KRASG12V, LgBiT-KRASG12V α4-α5 mutants, or EGFR-LgBiT. All experiments were repeated at least three times (n=3) and results quantified using Welch’s t-test; error bars representing SEM (***p<0.0005, **p<0.005, and *p<0.05).

      RAS α4-α5 mutants are still susceptible to inhibition by the NS1 monobody

      NS1 inhibits oncogenic RAS-mediated signaling, biological transformation, and tumor formation (
      • Spencer-Smith R.
      • Koide A.
      • Zhou Y.
      • Eguchi R.R.
      • Sha F.
      • Gajwani P.
      • Santana D.
      • Gupta A.
      • Jacobs M.
      • Herrero-Garcia E.
      • Cobbert J.
      • Lavoie H.
      • Smith M.
      • Rajakulendran T.
      • Dowdell E.
      • Okur M.N.
      • Dementieva I.
      • Sicheri F.
      • Therrien M.
      • Hancock J.F.
      • Ikura M.
      • Koide S.
      • O'Bryan J.P.
      Inhibition of RAS function through targeting an allosteric regulatory site.
      ,
      • Khan I.
      • Spencer-Smith R.
      • O'Bryan J.P.
      Targeting the alpha4-alpha5 dimerization interface of K-RAS inhibits tumor formation in vivo.
      ,
      • Khan I.
      • MarElia-Bennet C.
      • Lefler J.
      • Zuberi M.
      • Denbaum E.
      • Koide A.
      • Connor D.M.
      • Broome A.M.
      • Pecot T.
      • Timmers C.
      • Ostrowski M.C.
      • Koide S.
      • O'Bryan J.P.
      Targeting the KRAS alpha4-alpha5 allosteric interface inhibits pancreatic cancer tumorigenesis.
      ) and disrupts higher order RAS associations at the plasma membrane (
      • Spencer-Smith R.
      • Koide A.
      • Zhou Y.
      • Eguchi R.R.
      • Sha F.
      • Gajwani P.
      • Santana D.
      • Gupta A.
      • Jacobs M.
      • Herrero-Garcia E.
      • Cobbert J.
      • Lavoie H.
      • Smith M.
      • Rajakulendran T.
      • Dowdell E.
      • Okur M.N.
      • Dementieva I.
      • Sicheri F.
      • Therrien M.
      • Hancock J.F.
      • Ikura M.
      • Koide S.
      • O'Bryan J.P.
      Inhibition of RAS function through targeting an allosteric regulatory site.
      ). Through binding of the α4-α5 region of RAS, NS1 allosterically inhibits RAS function irrespective of its nucleotide state (
      • Spencer-Smith R.
      • Koide A.
      • Zhou Y.
      • Eguchi R.R.
      • Sha F.
      • Gajwani P.
      • Santana D.
      • Gupta A.
      • Jacobs M.
      • Herrero-Garcia E.
      • Cobbert J.
      • Lavoie H.
      • Smith M.
      • Rajakulendran T.
      • Dowdell E.
      • Okur M.N.
      • Dementieva I.
      • Sicheri F.
      • Therrien M.
      • Hancock J.F.
      • Ikura M.
      • Koide S.
      • O'Bryan J.P.
      Inhibition of RAS function through targeting an allosteric regulatory site.
      ,
      • Spencer-Smith R.
      • Li L.
      • Prasad S.
      • Koide A.
      • Koide S.
      • O'Bryan J.P.
      Targeting the alpha4-alpha5 interface of RAS results in multiple levels of inhibition.
      ). We hypothesized that if the α4-α5 mutations decreased RAS–RAS interaction in cells, then NS1 would have less of an inhibitory effect on these mutants compared with the parental RAS oncogenic mutants. As illustrated in Fig. S6 and S7, α4-α5 mutations in KRASG12D did not impact ERK activation. Furthermore, co-expression of NS1 effectively inhibited downstream ERK phosphorylation in cells expressing KRASG12D and KRASG12D α4-α5 mutants (Fig. 4A and S9). The inhibitory effect of NS1 was highly specific as KRASG12D/R135E, which does not bind NS1 (
      • Spencer-Smith R.
      • Koide A.
      • Zhou Y.
      • Eguchi R.R.
      • Sha F.
      • Gajwani P.
      • Santana D.
      • Gupta A.
      • Jacobs M.
      • Herrero-Garcia E.
      • Cobbert J.
      • Lavoie H.
      • Smith M.
      • Rajakulendran T.
      • Dowdell E.
      • Okur M.N.
      • Dementieva I.
      • Sicheri F.
      • Therrien M.
      • Hancock J.F.
      • Ikura M.
      • Koide S.
      • O'Bryan J.P.
      Inhibition of RAS function through targeting an allosteric regulatory site.
      ), was refractory to inhibition by NS1 (Fig. 4A and S9). These results further support the notion that these α4-α5 mutations are not sufficient to impair RAS association or oncogenic RAS-mediated signaling in cells.
      Figure thumbnail gr4
      Figure 4RAS α4-α5 mutants are still susceptible to inhibition by the NS1 monobody. A) ERK/MAPK signaling assay in HEK 293 cells co-transfected with indicated HA-tagged KRASG12D mutants and FLAG-tagged NS1 or a negative control monobody (Mb (neg)). Results are representative of one of three biological replicates. Graph represents the relative pERK levels. The mean and sd (n=3) for the normalized pERK/ERK in the NS1 compared to Mb (neg) sample is shown. Dotted line at 1 represent pERK levels in Mb (neg) samples. All P values were generated using an unpaired Student’s t-test (***p<0.0005, **p<0.005, and *p<0.05).

      HRAS and NRAS α4-α5 mutants exhibit isoform-specific biological properties

      We then examined whether α4-α5 mutations of the other RAS isoforms had similar effects on their signaling and biological properties. In contrast to previously reported results with KRASD154Q (
      • Ambrogio C.
      • Köhler J.
      • Zhou Z.W.
      • Wang H.
      • Paranal R.
      • Li J.
      • Capelletti M.
      • Caffarra C.
      • Li S.
      • Lv Q.
      • Gondi S.
      • Hunter J.C.
      • Lu J.
      • Chiarle R.
      • Santamaría D.
      • Westover K.D.
      • Jänne P.A.
      KRAS Dimerization Impacts MEK Inhibitor Sensitivity and Oncogenic Activity of Mutant KRAS.
      ), we reported that mutations at D154 or R161 of HRASG12V had no effect on ERK-MAPK activation (
      • Spencer-Smith R.
      • Koide A.
      • Zhou Y.
      • Eguchi R.R.
      • Sha F.
      • Gajwani P.
      • Santana D.
      • Gupta A.
      • Jacobs M.
      • Herrero-Garcia E.
      • Cobbert J.
      • Lavoie H.
      • Smith M.
      • Rajakulendran T.
      • Dowdell E.
      • Okur M.N.
      • Dementieva I.
      • Sicheri F.
      • Therrien M.
      • Hancock J.F.
      • Ikura M.
      • Koide S.
      • O'Bryan J.P.
      Inhibition of RAS function through targeting an allosteric regulatory site.
      ), which might suggest isoform-specific effects of these mutations. Consistent with prior findings (
      • Spencer-Smith R.
      • Koide A.
      • Zhou Y.
      • Eguchi R.R.
      • Sha F.
      • Gajwani P.
      • Santana D.
      • Gupta A.
      • Jacobs M.
      • Herrero-Garcia E.
      • Cobbert J.
      • Lavoie H.
      • Smith M.
      • Rajakulendran T.
      • Dowdell E.
      • Okur M.N.
      • Dementieva I.
      • Sicheri F.
      • Therrien M.
      • Hancock J.F.
      • Ikura M.
      • Koide S.
      • O'Bryan J.P.
      Inhibition of RAS function through targeting an allosteric regulatory site.
      ,
      • Grozavu I.
      • Stuart S.
      • Lyakisheva A.
      • Yao Z.
      • Pathmanathan S.
      • Ohh M.
      • Stagljar I.
      D154Q Mutation does not Alter KRAS Dimerization.
      ) and the results with KRASG12V (Fig. 2), mutations at D154 or R161 had no effect on either HRASG12V or NRASG12V signaling (Fig. 5A–D and S10).
      Next, we assessed the impact of these mutations on the biological activity of H/NRASG12V. Whereas the KRASG12V mutants showed no significant differences in foci formation (Fig. 2E and 2F), all of the HRASG12V α4-α5 mutants (Fig. 5E and 5F) and one of the NRASG12V α4-α5 mutants (R161D) (Figure 5G and 5H) resulted in significantly fewer foci compared with the parental control. Together, these data show that mutations in the α4-α5 allosteric lobe of all three RAS isoforms have no effect on ERK-MAPK signaling but impair the transforming activity of HRASG12V and to a lesser extent NRASG12V.
      Figure thumbnail gr5
      Figure 5HRAS and NRAS α4-α5 mutants exhibit isoform-specific biological properties. A) ERK/MAPK activity assay in HEK 293 cells transfected with HA-tagged HRASG12V mutants. B) Normalized pERK signal from panel (A). C) ERK/MAPK activity assay in HEK 293 cells transfected with HA-tagged NRASG12V mutants. D) Normalized pERK signal from panel (C). E) NIH/3T3 transformation assay in cells transfected with HRASG12V mutants. F) Normalized foci number from the NIH/3T3 transformation assays in panel (E). G) NIH/3T3 transformation assay in cells transfected with NRASG12V mutants. H) Normalized foci number from the NIH/3T3 transformation assays in panel (E). All experiments were repeated at least three times (n=3) and results quantified using Welch’s t-test; error bars representing SEM ((***p<0.0005, **p<0.005, and *p<0.05).

      Discussion

      The link between RAS dimerization and RAS-mediated signaling was first observed by Santos and colleagues in 1998 (
      • Santos E.
      • Nebreda A.R.
      • Bryan T.
      • Kempner E.S.
      Oligomeric structure of p21 ras proteins as determined by radiation inactivation.
      ) and was revisited at the turn of the 21st century (
      • Inouye K.
      • Mizutani S.
      • Koide H.
      • Kaziro Y.
      Formation of the Ras dimer is essential for Raf-1 activation.
      ). Indeed, artificial dimerization of RAS at the plasma membrane activated the MAPK pathway (
      • Nan X.
      • Tamguney T.M.
      • Collisson E.A.
      • Lin L.J.
      • Pitt C.
      • Galeas J.
      • Lewis S.
      • Gray J.W.
      • McCormick F.
      • Chu S.
      Ras-GTP dimers activate the Mitogen-Activated Protein Kinase (MAPK) pathway.
      ) while inhibition of RAS clusters at the plasma membrane is associated with inhibition of the MAPK pathway (
      • Spencer-Smith R.
      • Koide A.
      • Zhou Y.
      • Eguchi R.R.
      • Sha F.
      • Gajwani P.
      • Santana D.
      • Gupta A.
      • Jacobs M.
      • Herrero-Garcia E.
      • Cobbert J.
      • Lavoie H.
      • Smith M.
      • Rajakulendran T.
      • Dowdell E.
      • Okur M.N.
      • Dementieva I.
      • Sicheri F.
      • Therrien M.
      • Hancock J.F.
      • Ikura M.
      • Koide S.
      • O'Bryan J.P.
      Inhibition of RAS function through targeting an allosteric regulatory site.
      ,
      • Sutton M.N.
      • Lu Z.
      • Li Y.C.
      • Zhou Y.
      • Huang T.
      • Reger A.S.
      • Hurwitz A.M.
      • Palzkill T.
      • Logsdon C.
      • Liang X.
      • Gray J.W.
      • Nan X.
      • Hancock J.
      • Wahl G.M.
      • Bast R.C.
      DIRAS3 (ARHI) Blocks RAS/MAPK Signaling by Binding Directly to RAS and Disrupting RAS Clusters.
      ). Although there is still debate surrounding the exact mechanism of RAS nanoclustering at the plasma membrane, the α4-α5 region of RAS has been proposed in several studies to be an important interface contributing several stabilizing interactions to facilitate formation of RAS dimers (
      • Spencer-Smith R.
      • Koide A.
      • Zhou Y.
      • Eguchi R.R.
      • Sha F.
      • Gajwani P.
      • Santana D.
      • Gupta A.
      • Jacobs M.
      • Herrero-Garcia E.
      • Cobbert J.
      • Lavoie H.
      • Smith M.
      • Rajakulendran T.
      • Dowdell E.
      • Okur M.N.
      • Dementieva I.
      • Sicheri F.
      • Therrien M.
      • Hancock J.F.
      • Ikura M.
      • Koide S.
      • O'Bryan J.P.
      Inhibition of RAS function through targeting an allosteric regulatory site.
      ,
      • Guldenhaupt J.
      • Rudack T.
      • Bachler P.
      • Mann D.
      • Triola G.
      • Waldmann H.
      • Kotting C.
      • Gerwert K.
      N-Ras forms dimers at POPC membranes.
      ,
      • Muratcioglu S.
      • Chavan T.S.
      • Freed B.C.
      • Jang H.
      • Khavrutskii L.
      • Freed R.N.
      • Dyba M.A.
      • Stefanisko K.
      • Tarasov S.G.
      • Gursoy A.
      • Keskin O.
      • Tarasova N.I.
      • Gaponenko V.
      • Nussinov R.
      GTP-Dependent K-Ras Dimerization.
      ,
      • Mysore V.P.
      • Zhou Z.W.
      • Ambrogio C.
      • Li L.
      • Kapp J.N.
      • Lu C.
      • Wang Q.
      • Tucker M.R.
      • Okoro J.J.
      • Nagy-Davidescu G.
      • Bai X.
      • Pluckthun A.
      • Janne P.A.
      • Westover K.D.
      • Shan Y.
      • Shaw D.E.
      A structural model of a Ras-Raf signalosome.
      ,
      • Lee K.Y.
      • Fang Z.
      • Enomoto M.
      • Gasmi-Seabrook G.
      • Zheng L.
      • Koide S.
      • Ikura M.
      • Marshall C.B.
      Two Distinct Structures of Membrane-Associated Homodimers of GTP- and GDP-Bound KRAS4B Revealed by Paramagnetic Relaxation Enhancement.
      ,
      • Lee K.Y.
      • Enomoto M.
      • Gebregiworgis T.
      • Gasmi-Seabrook G.M.C.
      • Ikura M.
      • Marshall C.B.
      Oncogenic KRAS G12D mutation promotes dimerization through a second, phosphatidylserine-dependent interface: a model for KRAS oligomerization.
      ,
      • Herrero A.
      • Crespo P.
      RAS Dimers: The Novice Couple at the RAS-ERK Pathway Ball.
      ). Based on these studies, mutations within the α4-α5 allosteric lobe (e.g., D154Q) have been proposed to disrupt the interactions necessary to form higher-order RAS assemblies. However, there has been conflicting data surrounding the ability of these RAS mutants to impact signaling and biology.
      Consistent with our previous results (
      • Spencer-Smith R.
      • Koide A.
      • Zhou Y.
      • Eguchi R.R.
      • Sha F.
      • Gajwani P.
      • Santana D.
      • Gupta A.
      • Jacobs M.
      • Herrero-Garcia E.
      • Cobbert J.
      • Lavoie H.
      • Smith M.
      • Rajakulendran T.
      • Dowdell E.
      • Okur M.N.
      • Dementieva I.
      • Sicheri F.
      • Therrien M.
      • Hancock J.F.
      • Ikura M.
      • Koide S.
      • O'Bryan J.P.
      Inhibition of RAS function through targeting an allosteric regulatory site.
      ), we found that single point mutations within the α4-α5 allosteric lobe of RAS predicted to disrupt dimer formation (Fig. 1) did not decrease downstream MAPK pathway activation compared to parental oncogenic RAS, with the exception of the KRASG12V/K147D mutant. K147 is an important site for post-translational modifications (PTMs) that regulate RAS activity. Monoubiquitylation of K147 impedes GAP interaction with RAS, thereby favoring the RAS-GTP state by decreasing GAP-stimulated GTP hydrolysis (
      • Baker R.
      • Wilkerson E.M.
      • Sumita K.
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      ,
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      ,
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      Site-specific monoubiquitination activates Ras by impeding GTPase-activating protein function.
      ). Furthermore, K147 acetylation regulates nucleotide binding and is associated with increased KRAS activity and tumor growth in vivo (
      • Song H.Y.
      • Biancucci M.
      • Kang H.J.
      • O'Callaghan C.
      • Park S.H.
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      • Jiang H.
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      SIRT2 deletion enhances KRAS-induced tumorigenesis in vivo by regulating K147 acetylation status.
      ). Thus, the reduction in downstream MAPK signaling from KRASG12V/K147D, especially since it did not impair self-association of KRAS (Fig. 3), could be a consequence of the K147D mutation preventing these tumor-promoting PTMs. Consistent with the MAPK signaling activity, there was no significant impairment in foci formation from the KRASG12V α4-α5 mutants compared to KRASG12V. Together, these results demonstrate that mutations at residues in the α4-α5 region that have been proposed to mediate RAS dimerization do not impair the oncogenic signaling or biological activity of KRAS. This suggests that if these mutations truly disrupted dimerization, then RAS dimerization, per se, may not be a necessary, on-pathway step for oncogenic RAS signaling. Conversely, if RAS dimerization is necessary for its activity, then our data suggest that these mutations do not affect RAS–RAS interactions to the extent necessary to inhibit oncogenic signaling.
      This study provides evidence that not only do the KRAS α4-α5 mutants retain the ability to activate MAPK signaling, but they also do not have impaired PPIs with other RAS monomers in cells. While NS1 does not disrupt the association of RAF with HRAS, it decreases CRAF–BRAF association in cells, reflecting the ability of NS1 to sterically interfere with RAS clustering at the plasma membrane (
      • Spencer-Smith R.
      • Koide A.
      • Zhou Y.
      • Eguchi R.R.
      • Sha F.
      • Gajwani P.
      • Santana D.
      • Gupta A.
      • Jacobs M.
      • Herrero-Garcia E.
      • Cobbert J.
      • Lavoie H.
      • Smith M.
      • Rajakulendran T.
      • Dowdell E.
      • Okur M.N.
      • Dementieva I.
      • Sicheri F.
      • Therrien M.
      • Hancock J.F.
      • Ikura M.
      • Koide S.
      • O'Bryan J.P.
      Inhibition of RAS function through targeting an allosteric regulatory site.
      ). In contrast, the KRASG12V α4-α5 mutants were not significantly impaired in their ability to induce CRAF–BRAF interaction compared to KRASG12V, which was also reflected in the MAPK signaling assays. These results were further corroborated by the NanoBiT assays showing that the α4-α5 mutations did not affect RAS–RAS association in cells. Furthermore, these mutants remained sensitive to inhibition by NS1. Overall, we have shown that oncogenic KRAS is essentially unimpaired by mutations in the α4-α5 allosteric lobe.
      The isoform-specific differences in transformation observed with the D154 and R161 mutations in HRAS and the R161 mutation in NRAS were unexpected but raise important questions. First, could these mutations disrupt differential effector activation from the RAS isoforms? For instance, Yan et al. (1998) reported that KRAS potently activated RAF but was less efficient at activating PI3K when compared with HRAS (
      • Yan J.
      • Roy S.
      • Apolloni A.
      • Lane A.
      • Hancock J.F.
      Ras isoforms vary in their ability to activate Raf-1 and phosphoinositide 3-kinase.
      ). While there is no evidence for PI3K dimerization, targeting RAS with NS1 monobody decreased downstream phospho-AKT levels (
      • Spencer-Smith R.
      • Koide A.
      • Zhou Y.
      • Eguchi R.R.
      • Sha F.
      • Gajwani P.
      • Santana D.
      • Gupta A.
      • Jacobs M.
      • Herrero-Garcia E.
      • Cobbert J.
      • Lavoie H.
      • Smith M.
      • Rajakulendran T.
      • Dowdell E.
      • Okur M.N.
      • Dementieva I.
      • Sicheri F.
      • Therrien M.
      • Hancock J.F.
      • Ikura M.
      • Koide S.
      • O'Bryan J.P.
      Inhibition of RAS function through targeting an allosteric regulatory site.
      ) suggesting that disrupting RAS nanoclusters may affect multiple pathways. Second, it is possible that each isoform may utilize distinct mechanisms of nanoclustering (
      • Herrero A.
      • Crespo P.
      RAS Dimers: The Novice Couple at the RAS-ERK Pathway Ball.
      ). Nevertheless, given the inability of these α4-α5 mutations to impair oncogenic RAS signaling (i.e., RAF-MAPK activation), we conclude that either these mutations do not affect RAS dimerization or that RAS dimerization is not needed for RAS-induced MAPK activation.
      Our results are consistent with a model in which RAS protomers rely on proximity, but not direct association with one another to form a signaling-competent complex (Fig. 6). The required proximity may be in the form of loosely associated nanoclusters where RAS protomers are close enough to promote RAF dimerization but do not require well-defined interactions between amino acid side chains of residues within the α4-α5 allosteric lobe. Disruption of these nanoclusters may require larger molecules, such as NS1 or DIRAS3, which may reduce the density of RAS on the membrane surface and/or distort the RAS–RAF complex into an inactive conformation (
      • Fang Z.
      • Marshall C.B.
      • Nishikawa T.
      • Gossert A.D.
      • Jansen J.M.
      • Jahnke W.
      • Ikura M.
      Inhibition of K-RAS4B by a Unique Mechanism of Action: Stabilizing Membrane-Dependent Occlusion of the Effector-Binding Site.
      ). In contrast, point mutations of specific amino acid residues on the α4-α5 lobe of RAS do not appear sufficient to disrupt the interactions necessary for downstream pathway activation. The implications of these results for drug design are that smaller molecules targeting the α4-α5 allosteric lobe may be insufficient to impair downstream pathway activation. Instead, an approach to bring a large molecule to the α4-α5 region utilizing “glue” compounds (
      • Shigdel U.K.
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      • Stiles D.T.
      • Blodgett J.A.V.
      • Udwary D.W.
      • Rajczewski A.T.
      • Mann A.S.
      • Mostafavi S.
      • Hardy T.
      • Arya S.
      • Weng Z.
      • Stewart M.
      • Kenyon K.
      • Morgenstern J.P.
      • Pan E.
      • Gray D.C.
      • Pollock R.M.
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      • Klausner R.D.
      • Townson S.A.
      • Verdine G.L.
      Genomic discovery of an evolutionarily programmed modality for small-molecule targeting of an intractable protein surface.
      ) may be required to exploit this vulnerability of RAS.
      Figure thumbnail gr6
      Figure 6Model for the role of the α4-α5 region of oncogenic RAS higher-order assembly. GTP loading of RAS results in recruitment of RAF through binding of the CRD-RBD region of RAF resulting in unmasking of the dimerization interface on RAF. This results in RAF-mediated clustering of activated RAS. NS1 disrupts RAS clusters and RAS signaling through steric hinderance which prevents RAF from forming productive dimers. Mutations within the α4-α5 region are insufficient to disrupt this clustering due to a lack of electrostatic interactions between requisite amino acid side chains. Created with BioRender.com

      Experimental Procedures

      Cell culture and cloning

      Freshly thawed HEK 293 (MUSC Tissue Culture Facility) and NIH/3T3 (National Institutes of Health) cells were maintained in Dulbecco’s modified Eagle’s medium (DMEM) (Corning) supplemented with 10% fetal bovine serum (FBS) or 10% calf serum, respectively. RAS-less MEFs (BRAFV600E) were obtained from the National Cancer Institute and maintained in DMEM supplemented with 10% FBS and 4 ug/mL blasticidin. RAS α4-α5 mutants were generated via site-directed mutagenesis using pCGN-HA-RASG12V or pCGN-HA-RASG12D as templates for each isoform. Primers used to generate RAS α4-α5 mutants are listed in Table 1. All monobodies were subcloned into CMV-driven expression vectors containing an mCherry-tag followed by a FLAG-tag on the N-terminus. KRASG12V was subcloned into a high-expression, CMV-driven vector downstream of the SmBiT cDNA sequence. Similarly, KRASG12V and the α4-α5 mutants were cloned downstream of LgBiT-containing vectors; however, these vectors were low-expression, HSV-driven vectors. Lastly, EGFR-LgBiT clone was in a high-expression, CMV-driven vector. All NanoBiT® vectors were provided by Matt Robers and Dr. Jim Vasta (Promega).
      Table 1Primers used to generate the RASG12V α4-α5 mutants analyzed in this study
      α4–α5 mutationForward Primer (5' to 3')Reverse Primer (5' to 3')
      KRASG12VK128ECCAGAACAGTAGACACAGAACAGGCTCAGGACTTAGCATGCTAAGTCCTGAGCCTGTTCTGTGTCTACTGTTCTGG
      Q131ECAGTAGACACAAAACAGGCTGAGGACTTAGCAAGAAGTACTTCTTGCTAAGTCCTCAGCCTGTTTTGTGTCTACTG
      R135ECAGGCTCAGGACTTAGCAGAAAGTTATGGAATTCCTTTAAAGGAATTCCATAACTTTCTGCTAAGTCCTGAGCCTG
      K147DTTTATTGAAACATCAGCAGACACAAGACAGGGTGTTGATCAACACCCTGTCTTGTGTCTGCTGATGTTTCAATAAA
      D154QACAAGACAGGGTGTTGATCAAGCCTTCTATACATTAGTTAACTAATGTATAGAAGGCTTGATCAACACCCTGTCTTGT
      D154RACAAGACAGGGTGTTGATAGAGCCTTCTATACATTAGTTAACTAATGTATAGAAGGCTCTATCAACACCCTGTCTTGT
      R161DGCCTTCTATACATTAGTTGATGAAATTCGAAAACATAAATTTATGTTTTCGAATTTCATCAACTAATGTATAGAAGGC
      HRASG12VD154QACCCGGCAGGGAGTGGAGCAGGCCTTCTACACGTTGGTGCACCAACGTGTAGAAGGCCTGCTCCACTCCCTGCCGGGT
      D154RACCCGGCAGGGAGTGGAGCGGGCCTTCTACACGTTGGTGCACCAACGTGTAGAAGGCCCGCTCCACTCCCTGCCGGGT
      R161DGATGCCTTCTACACGTTGGTGGACGAGATCCGGCAGCACGTGCTGCCGGATCTCGTCCACCAACGTGTAGAAGGCATC
      NRASG12VD154QACCAGACAGGGTGTTGAACAAGCTTTTTACACACTGGTATACCAGTGTGTAAAAAGCTTGTTCAACACCCTGTCTGGT
      D154RACCAGACAGGGTGTTGAAAGAGCTTTTTACACACTGGTATACCAGTGTGTAAAAAGCTCTTTCAACACCCTGTCTGGT
      R161DGCTTTTTACACACTGGTAGATGAAATACGCCAGTACCGATCGGTACTGGCGTATTTCATCTACCAGTGTGTAAAAAGC

      Transfections and cell signaling assays

      Transfections and cell signaling assays were performed as previously described (
      • Khan I.
      • Koide A.
      • Zuberi M.
      • Ketavarapu G.
      • Denbaum E.
      • Teng K.W.
      • Rhett J.M.
      • Spencer-Smith R.
      • Hobbs G.A.
      • Camp E.R.
      • Koide S.
      • O'Bryan J.P.
      Identification of the nucleotide-free state as a therapeutic vulnerability for inhibition of selected oncogenic RAS mutants.
      ,
      • Khan I.
      • O'Bryan J.P.
      Probing RAS Function with Monobodies.
      ). Briefly, HEK 293 cells were transfected with HA-tagged RAS using polyethylenimine (PEI). Typically, we transfected cells using 3 ul of PEI for every 1 ug of DNA. When indicated, HA-tagged RAS was co-transfected with MYC-tagged ERK for signaling assays. Transfected cells were incubated for 30 hours in complete media (DMEM with 10% FBS) then serum-starved overnight. MYC-tagged ERK was immunoprecipitated from the cell lysates using α-MYC antibody (Millipore-Sigma), then ERK and pERK levels were analyzed via Western Blot using α-ERK (Cell Signaling Technology) and α-pERK (Cell Signaling Technology) antibodies. ERK and pERK protein levels were quantified using Image Studio Lite (Ver 5.2) software. pERK/ERK ratio was determined for each mutant and normalized to the parental oncogenic RAS mutant. Each experiment was performed three times (n=3).
      To analyze CRAF–BRAF association, HEK 293 cells were transfected with the indicated RAS mutants using the same conditions as described above. After cell lysates were collected, a co-immunoprecipitation was done by pulling down endogenous CRAF using α-CRAF (BD Biosciences) antibody and probing for CRAF and BRAF via Western Blot with α-CRAF (BD Biosciences) and α-BRAF (Santa Cruz) antibodies. BRAF/CRAF ratio was determined for each mutant and all values were normalized to the parental oncogenic RAS mutant. Each experiment was performed three times (n=3).
      For signaling assays performed with KRASG12D and the NS1 monobody, 1x106 HEK293T cells were cultured 24 hours before transfection on a 6-well plate using DMEM supplemented with 10% FBS. Cells that were between 70-90% confluent were then serum starved and transfected with either KRAS variants or in a 1:1 ratio with monobodies using lipofectamine according to the manufacturer’s protocol. Raw band intensity was evaluated using Image Studio Lite® Version 5.2. For calculating the normalized (pERK/ERK)/HA, each band was first normalized to vinculin. For calculating the NS1/ Mb (neg) ratio, pERK/ERK ratio was first calculated then used to generate the NS1/ Mb (neg) ratio. Statistical analysis was performed using GraphPad Prism 9.

      Immunoblotting and antibodies

      Following experimental endpoints as described above, cell lysates were made by washing cells with PBS followed by addition of PLC lysis buffer (50 mM HEPES, pH 7.5, 150 mM NaCl, 10% glycerol, 1% Triton X-100, 1 mM EGTA, 1.5 mM magnesium chloride, 100 mM sodium fluoride supplemented with 1 mM vanadate, 10 mg/ml leupeptin and 10mg/ml aprotinin). Lysates were nutated at 4° C for 1 hour, then centrifuged at 14K RPM for 10 minutes. Supernatant containing protein was transferred to new tubes and stored at -80° C. BCA method (ThermoFisher) was used to quantify protein in cell lysates before Western Blot analyses. The following antibodies were used: monoclonal HA (clone 16B12, Biolegend #90154), monoclonal FLAG (Clone M2, Sigma #F1804), phospho-ERK (Thr202/Tyr204, CST #9101), total ERK (CST #9102), Vinculin (SC #73614), Anti-MYC (Clone A46, Millipore-Sigma #05–724, CRAF (BD Biosciences # 610151), BRAF (Santa Cruz #sc-9002).

      NIH/3T3 transformation assays

      For NIH/3T3 transformation assays, 2.5x105 cells were split into 60 mm tissue culture plates and seeded overnight. The following day, cells were transfected with the indicated RAS mutants using PEI transfection method(
      • Khan I.
      • O'Bryan J.P.
      Probing RAS Function with Monobodies.
      ). Media on cells was changed every two days following transfections. Oncogenic RAS induced foci formation approximately 2-3 weeks following transfections, and cells were fixed and stained with 0.1% crystal violet before quantification of foci. Assays were performed three times each (n=3).

      NanoBiT® protein–protein interaction assays

      For NanoBiT® PPI assays, 3.0 x 104 cells per well (RAS-less MEFs) were plated in a white-wall, clear-bottom 96-well plate (Thermo Scientific™ 165306) and incubated at 37 oC overnight. The next day, all wells were transfected with SmBiT-KRASG12V, and selected wells were transfected (technical replicates per experiment = 6) with either EGFR-LgBiT, LgBiT-KRASG12V, or LgBiT-KRASG12V α4-α5 mutants using PEI transfection method. 24 hours after transfection, media was aspirated from wells, and luminescence was measure using NanoGlo® Live-Cell Substrate (Promega; Cat # N2012) suspended in Opti-MEM® reduced serum media (Gibco; cat # 31985070). After the live-cell luminescence measurement, cells were lysed with 1.0% Triton X-100 and incubated with HiBiT peptide (0.1 μM) for 10 minutes on orbital shaker. Then, luminescence was measured to quantify LgBiT peptide levels. Live-cell luminescence was normalized to luminescence after HiBiT peptide addition, and all samples were normalized to SmBiT-KRASG12V/LgBiT-KRASG12V. Assays were performed three times each (n=3).

      Statistical Analysis

      All statistical analyses were performed using GraphPad Prism 9 software.

      Data and materials availability

      All reagents in this manuscript are available upon request and completion of an MTA with the Medical University of South Carolina and/or New York University.

      Competing interests

      J.P.O, A.K. and S.K. are listed as inventors on a patent application on Monobodies targeting the nucleotide-free state of RAS files by the Medical University of South Carolina and New York University (No. 62/862,924). K.W.T., A.K., and S.K. are listed as inventors on a patent application on mutant RAS targeting Monobodies filed by New York University (Application No. 63/121,903). A.K. and S.K. are listed as inventors on issued and pending patents on Monobody technology filed by The University of Chicago (US Patent 9512199 B2 and related pending applications). S.K. was an SAB member and holds equity in and received consulting fees from Black Diamond Therapeutics; receives research funding from Black Diamond Therapeutics, Puretech Health and Argenx BVBA. The other authors declare no competing interests.

      Acknowledgments

      We wish to thank members of the O’Bryan and Koide laboratories for comments on the manuscript. We also thank Matt Robers and Dr. Jim Vasta (Promega Corporation) for providing the NanoBiT reagents along with helpful advice on establishing the NanoBiT assay in the laboratory.

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