Skip to main content
NCBI home page
ACS Medicinal Chemistry Letters logoLink to ACS Medicinal Chemistry Letters
. 2016 Nov 30;8(1):61–66. doi: 10.1021/acsmedchemlett.6b00373

Covalent Guanosine Mimetic Inhibitors of G12C KRAS

Yuan Xiong †,‡, Jia Lu §, John Hunter §, Lianbo Li §, David Scott †,‡, Hwan Geun Choi ∥, Sang Min Lim ⊥, Anuj Manandhar §, Sudershan Gondi §, Taebo Sim #,, Kenneth D Westover §,*, Nathanael S Gray †,‡,*
PMCID: PMC5238463  PMID: 28105276

Abstract

graphic file with name ml-2016-00373a_0008.jpg

Ras proteins are members of a large family of GTPase enzymes that are commonly mutated in cancer where they act as dominant oncogenes. We previously developed an irreversible guanosine-derived inhibitor, SML-8-73-1, of mutant G12C RAS that forms a covalent bond with cysteine 12. Here we report exploration of the structure–activity relationships (SAR) of hydrolytically stable analogues of SML-8-73-1 as covalent G12C KRAS inhibitors. We report the discovery of difluoromethylene bisphosphonate analogues such as compound 11, which, despite exhibiting reduced efficiency as covalent G12C KRAS inhibitors, remove the liability of the hydrolytic instability of the diphosphate moiety present in SML-8-73-1 and provide the foundation for development of prodrugs to facilitate cellular uptake. The SAR and crystallographic results reaffirm the exquisite molecular recognition that exists in the diphosphate region of RAS for guanosine nucleotides which must be considered in the design of nucleotide-competitive inhibitors.

Keywords: KRAS G12C, drug design, covalent inhibitor, GDP mimetic, bisphosphonate, bioisostere, CPM, ActivX

RAS mutations are frequently observed in malignant tumors and support diverse hallmarks of cancer including genomic instability,1,2 cell proliferation,3 suppression of apoptosis,4 reprogramming of metabolism,7,8 alteration of the microenvironment,9,10 evasion of immune responses,11,12 and promotion of metastases.13,14 Consistent with its pervasive influence on cancer cell function, extinction of oncogenic KRAS in many established tumor models results in tumor regression.8,15,16 Targeting RAS as a therapeutic measure is therefore a high priority.17

RAS mutations appear to fall into functional categories and will likely require tailored strategies to target each functional class.18−21 KRAS G12C is the most commonly found RAS mutation in non-small cell lung cancer, the leading cause of cancer death in the United States.22 Previously we hypothesized that the activating cysteine mutation in KRAS G12C might allow development of covalent GTP-competitive inhibitors. We reasoned that targeting the active site of KRAS would likely perturb KRAS G12C-mediated signaling based on the observation that RAS signaling is predicated on the identity of its bound ligand.23,24

We developed a GDP mimetic inhibitor SML-8-73-1, which contains an alpha-chloroacetamide electrophile and reacts with Cys12 upon inhibitor binding to KRAS G12C (Scheme 1). SML-8-73-1 is also able to compete in vitro with high concentration of GDP and GTP and decreases the affinity of KRAS G12C for the RAS binding domain (RBD) of BRAF.25 Furthermore, unbiased proteomics based profiling showed that SML-8-73-1 is highly selective for KRAS G12C among other GTP binding proteins.26 Nevertheless, SML-8-73-1 contains multiple charged phosphate groups and cannot pass through the cell membrane. Caging strategies to shield the charged phosphates were hampered by compound instability.25

Scheme 1. Analogue Design Based on Diphosphate Bioisosteres.

Scheme 1

Open in a new tab

To enable caging strategies and to explore the possibility of other chemical moieties that might improve properties of the diphosphate pharmacophore for covalent targeting of KRAS G12C, we investigated structure–activity relationships (SAR) on a series of analogues of SML-8-73-1 that varied the diphosphate group as well as the linker moiety. Here we report biochemical characterization of these compounds including an illustrative X-ray crystal structure demonstrating key difficulties inherent to this approach.

A diphosphate compound, SML-8-73-1 suffers from chemical and enzymatic instability, given that the phosphate anhydride bond is prone to hydrolysis.27 This is problematic from two perspectives: first, unstable compounds are inherently disadvantageous from a pharmacokinetic perspective; and second, a caging strategy will likely be required to shield charged atoms in the phosphate pharmacophore to enhance cell permeability, and the resulting steric bulk may further destabilize the phosphate anhydride bond. To address these potential issues, the central oxygen was substituted with a methylene group. The corresponding bisphosphonate is considered to be resistant to acidic and enzymatic hydrolysis at the P–C–P bond,29,30 and has been broadly used as an isostere of the diphosphate.

Synthesis of the bisphosphonates utilizes a one-pot reaction sequence, starting with nucleophilic addition–elimination reaction between methylenebis(phosphonic dichloride) and acetonide-protected guanosine 3 affording phosphonic chloride 5, which was subsequently reacted with ethanolamine 6 to give phosphoric ester 7 after aqueous quench (Scheme 2). Deprotection under acidic conditions followed by selective acylation using activated NHS-ester 9 gave rise to desired bisphosphonate diester 10 (XY-01-103). This synthetic sequence allows facile and quick access to bisphosphonate analogues 10 to 20 with various linkers and different guanine analogues, by utilizing different alcohol reaction partners.

Scheme 2. Synthesis of Bisphosphonate Analogues.

Scheme 2

Open in a new tab

(a) PO(OEt)3, 0 °C; (b) 6, NEt3, PO(OEt)3, 0 °C to rt; (c) TFA/DCM, rt; (d) 9, DIEA, DMF, 0 °C.

Different routes were employed to synthesize various bisphosphonate isosteres. Synthesis of phosphonate intermediate 25 commenced with DCC coupling of diethylphosphonoacetic acid 21 and ethanolamine 22, and the resulting phosphoryl acetate 23 was deprotected and coupled to guanosine 3 to yield phosphoryl acetate 25 (Scheme 3). Similarly, phosphoryl acetate 29 was obtained by an altered sequence. Deprotection and selective acylation of intermediates 25 and 29 gave rise to phosphoryl acetate compounds 30 and 31, respectively. The synthesis of 47 started from sulfonylation of ethylenediamine 32. The resulting sulfonamide 34 was deprotonated and reacted with diethyl chlorophosphate 35. Following deprotection, the collidine salt 37 was coupled with guanosine 3 to give sulfamoyl phosphonate 44, which was deprotected and acylated to provide 47 (XY-02-075). For 48, sequential addition of ethanolamine 22 and guanosine 3 to chlorosulfonyl isocyanate 38 gave rise to sulfamate 45, which after deprotection was acylated to yield 48. Alternatively, the synthesis of 49 started from reaction of ethylenediamine 40 with chlorosulfonyl acetate 41. Subsequent saponification and DCC coupling with guanosine 3 yielded the requisite sulfamoylacetate 46, which was deprotected and acylated to give 49.

Scheme 3. Synthesis of Bisphosphonate Isosteres.

Scheme 3

Open in a new tab

(a) 22, DCC, cat. DMAP, DCM, rt; (b) TMSBr, MeCN, 0 °C to rt; (c) 3, DCC, cat. DMAP, DMF, 60 °C; (d) 3 or 6, DCC, cat. DMAP, DMF, rt; (e) H2, Pd/C, MeOH, rt; then PTSA, DCM, rt; (f) 9, DIEA, DMF, 0 °C; (g) TFA/DCM, rt; (h) 33, NEt3, DCM, 0 °C; (i) 35, nBuLi, THF, −78 °C; (j) TMSBr, collidine, MeCN, rt; (k) 22, NEt3, DCM, 0 °C; (l) 41, NEt3, DCM, 0 °C to rt; (m) NaOH aq, EtOH, rt; (n) 3, MeCN/pyridine, rt; (o) H2, Pd/C, MeOH, rt; then 4 N HCl in dioxane, DCM, rt.

To measure GDP displacement we utilized a competitive “chemosensor assay”, wherein purified recombinant GDP-loaded KRAS G12C is incubated with compound, then probed at various time points with a probe compound that detects the presence of free (unreacted) G12C thiol. This assay provides a composite measurement of kinetic displacement of GDP and covalent inactivation of KRAS G12C (Figure S1).26

To determine the relative affinities of compounds we measured kinact/Ki using GMP-stabilized KRAS G12C (Figure S2). To obtain these values we used the strategy advocated by Copeland31 wherein reaction rates are plotted vs compound concentrations and fit to the curve described by the equation kobs = kinact[I]/(Ki + [I]). Using the fit, estimates for kinact and Ki can be extracted (Figure S2C,D). For this assay purified nucleotide free KRAS G12C was prepared and stabilized with an excess of GMP, which has a low affinity of 3.5 × 104 M–1 for RAS.32 GMP-stabilized RAS is incubated with compound then probed with a GTP-desthiobiotin probe which, similarly to widely used ATP-biotin (ActivX) probes,33,34 contains a reactive acyl phosphate anhydride that reacts with lysine 16 of KRAS. Desthiobiotinylated protein is detected using AlphaScreen (PerkinElmer) reagents (Figure S2A). Because the ActivX and AlphaScreen reagents are used in combination, we call this assay ActivAlpha (Figure S2D).

A third assay consists of incubating recombinant G12C KRAS with new inhibitors and then performing electrospray ionization mass spectrometry as reported previously.25 The percent labeling of the protein can be detected as shown in Figure S3.

To meet our overarching goal of developing GTP-competitive inhibitors that have anticancer effects in cells, compounds will likely need to achieve adequate RAS inhibition well within the typical time scale of the RAS protein turnover; the half-life has been estimated at 12–24 h.35,36 We previously performed rudimentary kinetics simulations37 showing that G12C KRAS inactivators with a kinact of 0.6 min–1 and Ki of 10 nM will yield 50% inhibition of KRAS G12C in 5 h. We therefore adopt these as preliminary standards for inhibitor development. Nevertheless it should be noted that the ActivAlpha assay utilizes an excess of GMP to stabilize the KRAS protein and, unlike the in vivo situation, does not require displacement of GDP or GTP.

In the ActivAlpha assay diphosphate 1 (SML-8-73-1) shows an excellent Ki of 9 nM and a relatively fast kinact of 0.86 min–1, which is consistent with the reactive chloroacetamide electrophile of SML-8-73-1. However, the corresponding methylenebisphosphonate analogue 10 loses 300-fold in binding affinity (Ki), and labels KRAS at a rate that is 11-fold slower (Table 1). This highlights the importance of interactions between the oxygen atom of the diphosphate and the various residues in the P-loop of KRAS. By substituting the central oxygen with a methylene group in 10, both the pKa of the resulting phosphoric acid and bond lengths and bond angles of the P–X–P linkage (X = O, CH2) are altered, which may result in lower affinity.38 To regain the electronic and conformational properties as in SML-8-73-1, difluoromethylene and monofluoromethylene groups were incorporated in compounds 11 and 12, respectively. Fluorinated phosphonates are widely used in the pharmaceutical industry as phosphate isosteres and are found to be better phosphate mimetics than phosphonates.39,40 In this case, difluoromethylene bisphosphonate analogue 11 improved the affinity for KRAS by 7-fold, compared to bisphosphonate analogue 10, while the monofluoromethylene bisphosphonate analogue 12 showed a lower affinity and slower labeling rate.41

Table 1. SAR of Bisphosphonate Analogues and Diphosphate Isosteres.

graphic file with name ml-2016-00373a_0006.jpg

graphic file with name ml-2016-00373a_0007.jpg

Open in a new tab

The acrylamide analogue of SML-8-73-1 (2) exhibits a higher rate constant kinact than α-chloroacetamide-containing SML-8-73-1, although it is generally believed that the α-chloroacetamide warhead is more reactive toward cysteine group than acrylamide.42 We postulated that longer length between the β-phosphate and the reactive site of the acrylamide in 2 may be optimal for the requisite covalent bond formation. Analogues with a propyl linker (13, 14) improved the affinity to KRAS G12C, possibly due to a combinatory effect of optimal spacing, and better trajectory of the electrophile warhead toward nucleophilic substitution of Cys12. On the contrary, analogue 20, with an internal electrophile and shorter distance between the β-phosphate and the reactive site for Cys12, showed complete loss of activity.

We hypothesized that a rigid linker with a preferred trajectory may further improve labeling efficiency, and a series of analogues with cyclic linkers were synthesized. In the case of 16 with a pyrrolidine linker, a larger kinact of 2.2 min–1 was observed, along with fast labeling of G12C KRAS in the chemosensor assay (t1/2 of 1.1h). Cyclopentane-containing analogues 18 and 19 which place the nitrogen in the exocyclic position also showed larger kinact, however the affinities for these two compounds are significantly lower. A piperidine linker in 17 did not improve affinity or labeling efficiency.

We also examined substitution of one or both phosphates with carbonyl or sulfonyl groups. The results showed that both phosphates are crucial in achieving high binding affinity to KRAS; substituting one or both phosphates significantly lowered affinity and resulted in longer labeling time. Among these, phosphonyl sulfonamide analogue 47 (XY-02-075) had the lowest Ki of 1.6 μM and a t1/2 of 2.3 h. A cocrystal structure of XY-02-075 with KRAS G12C showed an intramolecular hydrogen bonding between the sulfonamide hydrogen and the phosphonate oxygen (see below). Phosphoryl acetate analogue 31, in which the α-phosphate was replaced with an acetate group, showed a 7-fold lower Ki than the corresponding reversed phosphoryl acetate 30. This observation is in line with previous reports that interactions between the β-phosphate in GDP and Ras or magnesium are stronger than that of the α-phosphate.32 Further replacement of the remaining phosphonate with sulfonamide in compounds 48 and 49 did not significantly improve affinity. Incorporation of a squaryldiamide group, a common isostere for diphosphates and bisphosphonates,43 in 50 resulted in poor affinity.

New GDP analogues did not achieve the comparable labeling efficiency observed for SML-8-73-1. To gain insight into why our analogues were inferior to SML-8-73-1, we determined the X-ray crystal structure of XY-02-075 bound to KRAS G12C. Complete labeling of KRAS G12C with XY-02-075 was confirmed prior to crystallization using mass spectrometry (Figure S3). Crystals were in the monoclinic space group C2 with a unit cell similar to other KRAS structures obtained previously.26 Molecular replacement using WT KRAS GDP-bound structure as a search model (PDB 4OBE) was used to obtain phase information, and the final model was refined to a resolution of 2.70 Å with R-work of 28.0%, R-free of 33.5%, and average B-factor of 89.0 Å2 (Table S2).

The structure of the G domain is similar to previously solved RAS family protein structures including the previously reported SML-8-73-1-bound structure (PDB 4NMM) (RMSD = 0.43 Å, 166 atoms aligned). Continuous positive density connecting the terminal carbon atom of XY-02-075 to Cys12 confirmed a covalent link between the compound and protein (Figure S4). The conformation of residues surrounding the guanosine binding site, including P-loop (residue 10–17), 57DXXG, 116NKXD, and 146SAK motifs, were similar to the SML-8-73-1-bound structure with several differences. Importantly there is a lack of density where a magnesium ion and coordinated water molecules have been observed in nearly all previous structures of HRAS and KRAS (Figure 1B,C). Also, in the XY-02-075-bound structure we observed a hydrogen bond between the amide carbonyl oxygen atom and the backbone nitrogen atom of Gly13, whereas the linker amide group formed a hydrogen bond with the ε-nitrogen group of Lys16 in the SML-8-73-1 structure. We also noted an outward displacement of switch I residues Tyr32 and Asp33 (Figure 1A). Finally, due to poor electron density and a high B-factor, switch II residues 63 and 64 could not be unambiguously assigned in our model.

Figure 1.

Figure 1

Open in a new tab

Comparison between G12C in complex with XY-02-075 (cyan/blue) and SML 8-73-1 (gray/orange). (A) Structural superposition of nucleotide binding pocket: switch I is highlighted in XY-02-075-bound structure in yellow and in SML-8-73-1 in salmon. Tyr32 and Asp33 are in sticks as labeled. (B, C) Schematic representation comparing interactions between SML-8-73-1 (B) or XY-02-075 (C) and Kras G12C. Dashed lines represent the hydrogen bonding interactions.

Replacement of β-phosphate with sulfonamide apparently leads to dissociation of the magnesium ion and its coordinated water network. Compared to the SML-8-73-1 structure, the magnesium-bridged interaction between the β-phosphate and Tyr 32 and Asp 33 is lost when bound to XY-02-075. Additionally, an intramolecular hydrogen bond forms within XY-02-075 between the amide hydrogen of the sulfonamide and the oxygen atom of the α-phosphonate resulting in a kink in the inhibitor conformation relative to SML-8-73-1 (Figure 1B,C). This kink contributes to displacement of switch I residues Tyr32 and Asp33 by 3.3 Å relative to the position observed in the SML-8-73-1-bound structure. The folded conformation of XY-02-075 also does not allow the sulfonamide oxygens to form hydrogen bonds with the backbone nitrogen atoms of P-loop residues Gly15 and Lys16 as observed with SML-8-73-1. Consequently, the orientation of the linker amide group of XY-02-075 is shifted in comparison to SML-8-73-1.

To confirm that our compounds do not coordinate efficiently with Mg2+, we performed a 31P NMR MgCl2 titration study44 against fixed concentrations of GDP, GTP, and several of our compounds. These showed a Mg2+-dependent shift in the 31PNMR signal with the half-maximal effect at ∼0.5 equiv for GDP and GTP and a plateau by 2 equiv. However, with the same concentration of our phosphonate compounds, up to 20 equiv of Mg2+ was required before the shift in signal began to plateau, demonstrating that our phosphonate analogues bind Mg2+ with much lower affinity (Figures S5, S6).

Mono-, di-, and triphosphate esters are ubiquitously found in biological molecules including small molecules such as nucleotides, acetyl CoA, inositides, and phospholipids and as post-translational modifications to serine, threonine, and tyrosine on many proteins. Therefore, it is not surprising that a host of protein catalytic and binding pockets have evolved to use these molecules as substrates, cofactors, and recognition motifs. While phosphates are powerful binding elements and vastly improve the water solubility of compounds to which they are attached, they typically impair passive diffusion across a lipid bilayer and therefore are typically not found in small molecule drugs. Medicinal chemists have found a number of phosphate bioisosteres that are recognized by these pockets, but typically there is a considerable loss of binding affinity.40 An alternative approach is to create phosphate prodrugs in which the phosphate is unveiled following intracellular metabolism by a suitable enzyme.

In stark contrast to the relative success in finding monophosphate mimetics, there has been much more limited effort and success in developing diphosphate mimetics. For example, squaryldiamides have been studied as diphosphate surrogates in mannosyltransferase targeting sugar-nucleotide mimics;43 phosphinylformate has been used as diphosphate isostere in squalene synthetase inhibitors;45 furthermore, sulfonylbenzoyl-nitrostyrenes were explored as bisubstrate type inhibitors of EGFR, in which the sulfonylbenzoyl moiety served as a diphosphate mimic.46 However, these bioisosteres suffer from lowered bioactivity when compared with their diphosphate counterparts. We faced this challenge when considering how to generate a cell penetrant version of our prototype G12C KRAS labeling compound SML-8-73-1. We envisioned two possible approaches: to find noncharged phosphate mimetics or to create diphosphate prodrugs. Here we described the synthesis and biological characterization of 18 analogues where we explored phosphate replacements and “linker” variation. We discovered that relatively subtle changes to the diphosphate group resulted in dramatic reduction in affinity and labeling efficiency.

Based on our structural analysis the loss of affinity appears to be, at least in some cases, driven by disruption of the coordination between our compounds and magnesium resulting in loss of numerous interactions seen in native ligand-bound structures. Future efforts to optimize the phosphate pharmacophore will likely benefit from analysis of coordination between compounds and Mg2+. This structure also adds evidence to the already extensive data supporting the highly dynamic nature of the RAS switches and their dependence on bound ligand for sampling certain conformations. In the structure reported here, switch I (especially residues 32 and 33) is shifted outward to accommodate the folded XY-02-075, which does not coordinate magnesium and water molecules. This suggests that even small alterations to a ligand bound to KRAS will result in conformational changes that will likely disrupt interactions with RAS effectors that use the switch I or switch II interfaces. The list of such proteins includes important proteins that transduce major signaling cascades from RAS including PI3K and B-RAF. This suggests that development of GTP-competitive inhibitors, if possible, will be highly likely to perturb RAS-dependent cellular processes such as oncogenesis in the case of activated RAS.

A promising compound obtained in this study, 11, possesses a difluoromethylene bisphosphonate moiety and exhibits a 40-fold reduced affinity relative to SML-8-73-1 but has the potential advantage of more facile prodrug creation due to not having the chemical instability that results from the phosphate anhydride bond. Current efforts focus on development of prodrugs of difluoromethylene bisphosphonate analogues as well as replacement of the guanine heterocycle with moieties that possess better binding affinity in order to regain affinity lost in the phosphate binding pocket.

Acknowledgments

Results shown in this article are derived from work performed at Argonne National Laboratory, Structural Biology Center at the Advanced Photon Source. Argonne is operated by U Chicago Argonne, LLC, for the U.S. Department of Energy, Office of Biological and Environmental Research, under Contract DEAC02-06CH11357. Mass spectrometry work shown in this article was performed at the UT Southwestern proteomics core facility.

Glossary

Abbreviations

DMAP

4-dimethylaminopyridine

DCC

N,N′-dicyclohexylcarbodiimide

PTSA

p-toluenesulfonic acid

DCM

dichloromethane

TFA

trifluoroacetic acid

DIEA

N,N-diisopropylethylamine

TMSBr

bromotrimethylsilane

Supporting Information Available

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acsmedchemlett.6b00373.

  • Experimental methods, X-ray data, and Figures S1–S6 (PDF)

Y.X., N.S.G., and K.D.W. were supported by Astellas Pharma Inc. K.D.W. was also supported by the Welch Foundation I1829, the V Foundation for Cancer Research, and DOD W81XWH-16-1-0106.

The authors declare no competing financial interest.

Supplementary Material

ml6b00373_si_001.pdf (1.6MB, pdf)

References

  1. Denko N. C.; Giaccia A. J.; Stringer J. R.; Stambrook P. J. The human Ha-ras oncogene induces genomic instability in murine fibroblasts within one cell cycle. Proc. Natl. Acad. Sci. U. S. A. 1994, 91, 5124. 10.1073/pnas.91.11.5124. [DOI] [PMC free article] [PubMed] [Google Scholar]
  2. Saavedra H. I.; Knauf J. A.; Shirokawa J. M.; Wang J.; Ouyang B.; Elisei R.; Stambrook P. J.; Fagin J. A. The RAS oncogene induces genomic instability in thyroid PCCL3 cells via the MAPK pathway. Oncogene 2000, 19, 3948. 10.1038/sj.onc.1203723. [DOI] [PubMed] [Google Scholar]
  3. Pylayeva-Gupta Y.; Grabocka E.; Bar-Sagi D. RAS oncogenes: weaving a tumorigenic web. Nat. Rev. Cancer 2011, 11, 761. 10.1038/nrc3106. [DOI] [PMC free article] [PubMed] [Google Scholar]
  4. Cox A. D.; Der C. J. The dark side of Ras: regulation of apoptosis. Oncogene 2003, 22, 8999. 10.1038/sj.onc.1207111. [DOI] [PubMed] [Google Scholar]
  5. Flier J. S.; Mueckler M. M.; Usher P.; Lodish H. F. Elevated levels of glucose transport and transporter messenger RNA are induced by ras or src oncogenes. Science 1987, 235, 1492. 10.1126/science.3103217. [DOI] [PubMed] [Google Scholar]
  6. Ying H.; Kimmelman A. C.; Lyssiotis C. A.; Hua S.; Chu G. C.; Fletcher-Sananikone E.; Locasale J. W.; Son J.; Zhang H.; Coloff J. L. Oncogenic Kras maintains pancreatic tumors through regulation of anabolic glucose metabolism. Cell 2012, 149, 656. 10.1016/j.cell.2012.01.058. [DOI] [PMC free article] [PubMed] [Google Scholar]
  7. Rak J.; Mitsuhashi Y.; Bayko L.; Filmus J.; Shirasawa S.; Sasazuki T.; Kerbel R. S. Mutant ras oncogenes upregulate VEGF/VPF expression: implications for induction and inhibition of tumor angiogenesis. Cancer Res. 1995, 55, 4575. [PubMed] [Google Scholar]
  8. Ancrile B. B.; O’Hayer K. M.; Counter C. M. Oncogenic ras-induced expression of cytokines: a new target of anti-cancer therapeutics. Mol. Interventions 2008, 8, 22. 10.1124/mi.8.1.6. [DOI] [PMC free article] [PubMed] [Google Scholar]
  9. Weijzen S.; Velders M.; Kast W. Modulation of the immune response and tumor growth by activated Ras. Leukemia 1999, 13, 502. 10.1038/sj.leu.2401367. [DOI] [PubMed] [Google Scholar]
  10. Seliger B.; Harders C.; Lohmann S.; Momburg F.; Urlinger S.; Tampe R.; Huber C. Down-regulation of the MHC class I antigen-processing machinery after oncogenic transformation of murine fibroblasts. Eur. J. Immunol. 1998, 28, 122.. [DOI] [PubMed] [Google Scholar]
  11. Lundy J.; Grimson R.; Mishriki Y.; Chao S.; Oravez S.; Fromowitz F.; Viola M. Elevated ras oncogene expression correlates with lymph node metastases in breast cancer patients. J. Clin. Oncol. 1986, 4, 1321. [DOI] [PubMed] [Google Scholar]
  12. Campbell P. M.; Der C. J. Oncogenic Ras and its role in tumor cell invasion and metastasis. Semin. Cancer Biol. 2004, 14, 105. 10.1016/j.semcancer.2003.09.015. [DOI] [PubMed] [Google Scholar]
  13. Millis A. J.; Hoyle M.; McCue H. M.; Martini H. Differential expression of metalloproteinase and tissue inhibitor of metalloproteinase genes in aged human fibroblasts. Exp. Cell Res. 1992, 201, 373. 10.1016/0014-4827(92)90286-H. [DOI] [PubMed] [Google Scholar]
  14. Fisher G. H.; Wellen S. L.; Klimstra D.; Lenczowski J. M.; Tichelaar J. W.; Lizak M. J.; Whitsett J. A.; Koretsky A.; Varmus H. E. Induction and apoptotic regression of lung adenocarcinomas by regulation of a K-Ras transgene in the presence and absence of tumor suppressor genes. Genes Dev. 2001, 15, 3249. 10.1101/gad.947701. [DOI] [PMC free article] [PubMed] [Google Scholar]
  15. Stephen A. G.; Esposito D.; Bagni R. K.; McCormick F. Dragging ras back in the ring. Cancer Cell 2014, 25, 272. 10.1016/j.ccr.2014.02.017. [DOI] [PubMed] [Google Scholar]
  16. Hunter J. C.; Manandhar A.; Carrasco M. A.; Gurbani D.; Gondi S.; Westover K. D. Biochemical and structural analysis of common cancer-associated KRAS mutations. Mol. Cancer Res. 2015, 13, 1325. 10.1158/1541-7786.MCR-15-0203. [DOI] [PubMed] [Google Scholar]
  17. Al-Mulla F.; Milner-White E. J.; Going J. J.; Birnie G. D. Structural differences between valine-12 and aspartate-12 Ras proteins may modify carcinoma aggression. J. Pathol. 1999, 187, 433.. [DOI] [PubMed] [Google Scholar]
  18. De Roock W.; Jonker D. J.; Di Nicolantonio F.; Sartore-Bianchi A.; Tu D.; Siena S.; Lamba S.; Arena S.; Frattini M.; Piessevaux H. Association of KRAS p. G13D mutation with outcome in patients with chemotherapy-refractory metastatic colorectal cancer treated with cetuximab. JAMA 2010, 304, 1812. 10.1001/jama.2010.1535. [DOI] [PubMed] [Google Scholar]
  19. Siegel R. L.; Miller K. D.; Jemal A. Cancer statistics. Ca-Cancer J. Clin. 2015, 65, 5. 10.3322/caac.21254. [DOI] [PubMed] [Google Scholar]
  20. Milburn M. V.; Tong L.; Brunger A.; Yamaizumi Z.; Nishimura S.; Kim S. Molecular switch for signal transduction: structural differences between active and inactive forms of protooncogenic ras proteins. Science 1990, 247, 939. 10.1126/science.2406906. [DOI] [PubMed] [Google Scholar]
  21. Satoh T.; Nakafuku M.; Kaziro Y. Function of Ras as a molecular switch in signal transduction. J. Biol. Chem. 1992, 267, 24149. [PubMed] [Google Scholar]
  22. Lim S. M.; Westover K. D.; Ficarro S. B.; Harrison R. A.; Choi H. G.; Pacold M. E.; Carrasco M.; Hunter J.; Kim N. D.; Xie T.; Sim T.; Janne P. A.; Meyerson M.; Marto J. A.; Engen J. R.; Gray N. S. Therapeutic targeting of oncogenic K-Ras by a covalent catalytic site inhibitor. Angew. Chem., Int. Ed. 2014, 53, 199. 10.1002/anie.201307387. [DOI] [PMC free article] [PubMed] [Google Scholar]
  23. Hunter J. C.; Gurbani D.; Ficarro S. B.; Carrasco M. A.; Lim S. M.; Choi H. G.; Xie T.; Marto J. A.; Chen Z.; Gray N. S.; Westover K. D. In situ selectivity profiling and crystal structure of SML-8-73-1, an active site inhibitor of oncogenic K-Ras G12C. Proc. Natl. Acad. Sci. U. S. A. 2014, 111, 8895. 10.1073/pnas.1404639111. [DOI] [PMC free article] [PubMed] [Google Scholar]
  24. Budovskii E. I.; Shibaev V. N. Kinetics of the acid hydrolysis of uridine diphosphate glucose and 3-N-methyluridine diphosphate glucose. Chem. Nat. Compd. 1968, 4, 199. 10.1007/BF00571136. [DOI] [Google Scholar]
  25. Harris M. R.; Usher D. A.; Albrecht H. P.; Jones G. H.; Moffatt J. G. The hydrolysis of uridine cyclic phosphonate catalyzed by ribonuclease-A: implications for the mechanism of action of the enzyme. Proc. Natl. Acad. Sci. U. S. A. 1969, 63, 246. 10.1073/pnas.63.2.246. [DOI] [PMC free article] [PubMed] [Google Scholar]
  26. Widler L.; Jahnke W.; Green J. R. The chemistry of bisphosphonates: from antiscaling agents to clinical therapeutics. Anti-Cancer Agents Med. Chem. 2012, 12, 95. 10.2174/187152012799014959. [DOI] [PubMed] [Google Scholar]
  27. Copeland R. A.Evaluation of enzyme inhibitors in drug discovery a guide for medicinal chemists and pharmacologists; John Wiley & Sons: 2013. [PubMed] [Google Scholar]
  28. John J.; Sohmen R.; Feuerstein J.; Linke R.; Wittinghofer A.; Goody R. S. Kinetics of interaction of nucleotides with nucleotide-free H-ras p21. Biochemistry 1990, 29, 6058. 10.1021/bi00477a025. [DOI] [PubMed] [Google Scholar]
  29. Patricelli M. P.; Szardenings A. K.; Liyanage M.; Nomanbhoy T. K.; Wu M.; Weissig H.; Aban A.; Chun D.; Tanner S.; Kozarich J. W. Functional interrogation of the kinome using nucleotide acyl phosphates. Biochemistry 2007, 46, 350. 10.1021/bi062142x. [DOI] [PubMed] [Google Scholar]
  30. Xiao Y. S.; Guo L.; Jiang X. N.; Wang Y. S. Proteome-wide discovery and characterizations of nucleotide-binding proteins with affinity-labeled chemical probes. Anal. Chem. 2013, 85, 3198. 10.1021/ac303383c. [DOI] [PMC free article] [PubMed] [Google Scholar]
  31. Ulsh L. S.; Shih T. Y. Metabolic turnover of human c-rasH p21 protein of EJ bladder carcinoma and its normal cellular and viral homologs. Mol. Cell. Biol. 1984, 4, 1647. 10.1128/MCB.4.8.1647. [DOI] [PMC free article] [PubMed] [Google Scholar]
  32. Shukla S.; Allam U. S.; Ahsan A.; Chen G.; Krishnamurthy P. M.; Marsh K.; Rumschlag M.; Shankar S.; Whitehead C.; Schipper M. KRAS protein stability is regulated through SMURF2: UBCH5 complex-mediated β-TrCP1 degradation. Neoplasia 2014, 16, 115. 10.1593/neo.14184. [DOI] [PMC free article] [PubMed] [Google Scholar]
  33. Westover K. D. Enzymology of GTP-competitive RAS inhibitors. http://www.cancer.gov/research/key-initiatives/ras/ras-central/blog/Enzymology-of-GTP-competitive-RAS-inhibitors.
  34. For a table of pKa, bond lengths, and bond angles for different diphosphate isosteres, see the Supporting Information.
  35. Romanenko V. D.; Kukhar V. P. Fluorinated phosphonates: synthesis and biomedical application. Chem. Rev. 2006, 106, 3868. 10.1021/cr051000q. [DOI] [PubMed] [Google Scholar]
  36. Elliott T. S.; Slowey A.; Ye Y.; Conway S. J. The use of phosphate bioisosteres in medicinal chemistry and chemical biology. MedChemComm 2012, 3, 735. 10.1039/c2md20079a. [DOI] [Google Scholar]
  37. Imidobisphosphate (P-NH-P linkage) is also a common isostere for diphosphate. Due to difficulties in the synthesis of the corresponding diester, this analogue was not obtained.
  38. Flanagan M. E.; Abramite J. A.; Anderson D. P.; Aulabaugh A.; Dahal U. P.; Gilbert A. M.; Li C.; Montgomery J.; Oppenheimer S. R.; Ryder T.; Schuff B. P.; Uccello D. P.; Walker G. S.; Wu Y.; Brown M. F.; Chen J. M.; Hayward M. M.; Noe M. C.; Obach R. S.; Philippe L.; Shanmugasundaram V.; Shapiro M. J.; Starr J.; Stroh J.; Che Y. Chemical and Computational Methods for the Characterization of Covalent Reactive Groups for the Prospective Design of Irreversible Inhibitors. J. Med. Chem. 2014, 57, 10072. 10.1021/jm501412a. [DOI] [PubMed] [Google Scholar]
  39. Niewiadomski S.; Beebeejaun Z.; Denton H.; Smith T. K.; Morris R. J.; Wagner G. K. Rationally designed squaryldiamides-a novel class of sugar-nucleotide mimics?. Org. Biomol. Chem. 2010, 8, 3488. 10.1039/c004165c. [DOI] [PubMed] [Google Scholar]
  40. Pecoraro V. L.; Hermes J. D.; Cleland W. W. Stability constants of Mg2+ and Cd2+ complexes of adenine nucleotides and thionucleotides and rate constants for formation and dissociation of MgATP and MgADP. Biochemistry 1984, 23, 5262. 10.1021/bi00317a026. [DOI] [PubMed] [Google Scholar]
  41. Biller S. A.; Forster C.; Gordon E. M.; Harrity T.; Rich L. C.; Marretta J.; Ciosek C. P. Jr. Isoprenyl phosphinylformates: new inhibitors of squalene synthetase. J. Med. Chem. 1991, 34, 1912. 10.1021/jm00110a024. [DOI] [PubMed] [Google Scholar]
  42. Traxler P. M.; Wacker O.; Bach H. L.; Geissler J. F.; Kump W.; Meyer T.; Regenass U.; Roesel J. L.; Lydon N. Sulfonylbenzoyl-nitrostyrenes: potential bisubstrate type inhibitors of the EGF-receptor tyrosine protein kinase. J. Med. Chem. 1991, 34, 2328. 10.1021/jm00112a003. [DOI] [PubMed] [Google Scholar]

Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

ml6b00373_si_001.pdf (1.6MB, pdf)

Articles from ACS Medicinal Chemistry Letters are provided here courtesy of American Chemical Society

RESOURCES