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CCR Focus

Targeting RAS Membrane Association: Back to the Future for Anti-RAS Drug Discovery?

Adrienne D. Cox, Channing J. Der and Mark R. Philips
DOI: 10.1158/1078-0432.CCR-14-3214 Published April 2015

Abstract

RAS proteins require membrane association for their biologic activity, making this association a logical target for anti-RAS therapeutics. Lipid modification of RAS proteins by a farnesyl isoprenoid is an obligate step in that association, and is an enzymatic process. Accordingly, farnesyltransferase inhibitors (FTI) were developed as potential anti-RAS drugs. The lack of efficacy of FTIs as anticancer drugs was widely seen as indicating that blocking RAS membrane association was a flawed approach to cancer treatment. However, a deeper understanding of RAS modification and trafficking has revealed that this was an erroneous conclusion. In the presence of FTIs, KRAS and NRAS, which are the RAS isoforms most frequently mutated in cancer, become substrates for alternative modification, can still associate with membranes, and can still function. Thus, FTIs failed not because blocking RAS membrane association is an ineffective approach, but because FTIs failed to accomplish that task. Recent findings regarding RAS isoform trafficking and the regulation of RAS subcellular localization have rekindled interest in efforts to target these processes. In particular, improved understanding of the palmitoylation/depalmitoylation cycle that regulates RAS interaction with the plasma membrane, endomembranes, and cytosol, and of the potential importance of RAS chaperones, have led to new approaches. Efforts to validate and target other enzymatically regulated posttranslational modifications are also ongoing. In this review, we revisit lessons learned, describe the current state of the art, and highlight challenging but promising directions to achieve the goal of disrupting RAS membrane association and subcellular localization for anti-RAS drug development. Clin Cancer Res; 21(8); 1819–27. ©2015 AACR.

See all articles in this CCR Focus section, “Targeting RAS-Driven Cancers.”

Disclosure of Potential Conflicts of Interest

No potential conflicts of interest were disclosed.

Editor's Disclosures

The following editor(s) reported relevant financial relationships: S.E. Bates reports receiving a commercial research grant from Celgene via CRADA with NCI.

CME Staff Planners' Disclosures

The members of the planning committee have no real or apparent conflicts of interest to disclose.

Learning Objectives

Upon completion of this activity, the participant should have a better understanding of the strategies currently under investigation to improve treatment of RAS-driven cancers, and of the biologic rationale underlying novel therapeutic strategies that disrupt RAS membrane association.

Acknowledgment of Financial or Other Support

This activity does not receive commercial support.

Introduction

The three RAS genes (HRAS, NRAS, KRAS) are the most commonly mutated oncogenes in human cancers (1, 2). The role of oncogenic RAS proteins as key drivers in both common and uncommon cancers has led to intensive efforts over more than three decades to develop therapeutics that target RAS, encompassing both direct and indirect approaches. Oncogenically mutated RAS proteins fail to cycle “off” from the active GTP-bound state to the resting GDP-bound state, and thereby accumulate in the “on” configuration. Early direct approaches sought to attack this impaired molecular switch. Attempts to identify antagonists of GTP binding or to identify drug-like mimics of the negative regulatory GAP proteins have been unsuccessful, although new strategies of stabilizing conformational states may yet bear fruit, as discussed elsewhere in this CCR Focus section (3). More recent efforts to target specific RAS mutations (e.g., KRAS G12C), to interfere with RAS binding to its activator SOS1, and to block association with effectors such as RAF1 have been reviewed recently (1, 4). The consensus at present is that the most fruitful direction for anti-RAS therapeutics in the near future is indirect targeting of RAS signaling via inhibiting its downstream effectors, particularly the RAF–MEK–ERK and PI3K–AKT–MTOR kinase cascades that have been shown to be critical for RAS driver functions in specific cancers. These efforts are discussed elsewhere (1, 4, 5). Other approaches, such as attempts to identify additional targets for coinhibition with RAS, through synthetic lethality screens or metabolic dependencies, are also discussed elsewhere in this CCR Focus section (6, 7). Here, our focus is on direct targeting of RAS by interfering with its membrane association and trafficking. We argue that this approach, while challenging, remains both logical and potentially tractable, given information that has emerged over the past few years. Because the association of RAS proteins with membranes is absolutely required for their function, targeting this requirement can be viewed as the functional equivalent not of turning off the defective switch that is oncogenic RAS, but of removing it and thereby destroying the circuit.

CAAX Processing and RAS Membrane Association

The critical need for RAS protein association with cellular membranes has been appreciated for decades (8, 9). RAS association with the plasma membrane and with other membrane compartments upon which signaling occurs (10, 11) is promoted by a well-described series of posttranslational modifications at RAS C-terminal CAAX motifs (Fig. 1), where C = cysteine, A = (usually) aliphatic amino acids and X = a variable amino acid; in RAS, X = S or M (12, 13). In the initial and obligate step, a 15-carbon farnesyl polyisoprene lipid is added by farnesyltransferase (FTase) to the cysteine of the CAAX motif through a stable thioether linkage. Subsequently, the AAX amino acids are cleaved off by the farnesylcysteine-directed endoprotease, RAS converting CAAX endopeptidase 1, also known as RAS converting enzyme 1 (RCE1). The carboxyl group of the now C-terminal farnesylcysteine is next methylesterified by isoprenylcysteine carboxylmethyltransferase (LCMT) to produce RAS proteins with hydrophobic tails that have affinity for membranes. Both RCE1 and ICMT are restricted to the endoplasmic reticulum (14, 15), indicating that RAS must traffic to the plasma membrane via this compartment, and suggesting multiple layers of location-based regulation (Fig. 2). Each of the enzymes involved in these CAAX processing steps has been a target for drug discovery.

Figure 1.
Figure 1.

Membrane targeting sequences of RAS proteins. Top, the RAS on/off switch that is broken in oncogenically mutated RAS and fails to turn off from the active, GTP-bound state that interacts with effectors (E) to transmit downstream signals. Because membrane association is required for proper effector interaction, interfering with membrane targeting can impair signal transmission, like unwiring an electrical switch to prevent it from carrying current. Bottom, ribbon diagram of the four RAS proteins, which are 90% similar throughout their G domains that bind the guanine nucleotides, regulators, and effectors (including switch regions SI, SII), but differ greatly at their C-terminal membrane targeting domains. The latter consist of a carboxyterminal CAAX tetrapeptide motif (pink boxes) with an invariant cysteine that is the site of farnesylation, and an upstream hypervariable region (yellow boxes) that includes the “second signals” of one (NRAS) or two (HRAS, KRAS4A) palmitoylatable cysteines or clusters of positively charged (polybasic) residues (PBR), as well as “third signals” comprised of the surrounding residues. KRAS4B has a stretch of six contiguous lysines and no palmitoylatable cysteine, whereas KRAS4A has a hybrid motif of both a bipartite PBR and a palmitoylatable cysteine. Numbers refer to amino acid residues. Asterisks indicate sites of mutational hotspots at G12, G13, and Q61. Dots above the ribbon mark each 10 amino acid stretch. Brown bars in the ribbon mark sites of sequence variation. P, phosphorylation of KRAS4B at Serine181.

Figure 2.
Figure 2.

RAS trafficking pathway. Nascent RAS proteins leaving the polysome are rapidly modified by farnesyltransferase (FTase), which attaches a C15 farnesyl isoprenoid lipid to the cysteine of the CAAX motif. This provides them sufficient affinity for the endoplasmic reticulum (ER), where they are further modified by RAS-converting CAAX endopeptidase 1 (RCE1)-catalyzed proteolytic removal of the AAX residues, and by reversible isoprenylcysteine carboxylmethyltransferase (ICMT)-catalyzed carboxylmethylation of the now terminal farnesylated cysteine residue. By preventing the first and obligate step, FTIs prevent all three of these modifications. In the presence of FTIs, KRAS and NRAS are alternatively prenylated by geranylgeranyltransferase I (GGTase I), which attaches a C20 geranylgeranyl isoprenoid that allows the same subsequent processing steps. RAS trafficking to the inner leaflet of the plasma membrane (PM) requires a second membrane-targeting element that dictates the pathway it will take to the plasma membrane. NRAS and HRAS have one or two cysteine residues, respectively, that undergo reversible acylation by a Golgi-resident protein acyltransferase (PAT) to promote their trafficking to the plasma membrane. Rapid deacylation by an acyl protein thioesterase (APT1/2) frees them up to be re-acylated and trafficked back to the plasma membrane. The nonpalmitoylated pool of APT in the cytosol is the active form, and is in dynamic equilibrium with a palmitoylated pool on the Golgi. KRAS4B, which has no palmitoylatable cysteine but a stretch of 6 lysines (polybasic region) does not go to the Golgi but trafficks the more directly to the plasma membrane, where it binds by virtue of its electrostatic charge. KRAS4A, which has a hybrid motif of a palmitoylated cysteine and a bifurcated polybasic region, undergoes an intermediate form of trafficking. PDE6δ recognizes the farnesyl isoprenoid and solubilizes nonpalmitoylated RAS proteins from any compartment, thereby promoting their availability for restoration to the plasma membrane; deltarasin blocks this interaction. Not pictured: other chaperone proteins that guide the lipidated RAS proteins between and within membrane regions. Each enzyme depicted has been a target for drug discovery.

Targeting CAAX Prenylation: FTase and GGTase, Statins

RAS farnesylation by FTase is the first, irreversible, and rate-limiting step of CAAX processing. As it was quickly determined to be both an obligate modification for oncogenic RAS biologic activity and a process governed by an enzyme that recognized a simple short tetrapeptide CAAX motif, this step was rapidly exploited for inhibition. Both rational drug design and library screening were employed in numerous intensive and successful efforts to identify FTase inhibitors (FTI; refs. 16, 17). Two of these, lonafarnib and tipifarnib, progressed to advanced clinical trials but failed to show efficacy against KRAS-driven cancers (16, 17). The failure of FTIs, once anticipated to be magic bullets for RAS-driven cancers, to serve as broadly effective anti-RAS drugs led to a widespread misperception that even RAS itself is not a good target. However, it is critical to recognize that, in the presence of FTIs, NRAS and KRAS, but not HRAS, become substrates for geranylgeranyltransferase I (GGTase I) through a process known as alternative prenylation (18, 19). This phenomenon was revealed only when FTIs became available. Because geranylgeranylated RAS proteins still associate with membranes and are still biologically active, FTIs were ineffective despite hitting their FTase target. Thus, FTIs failed as anti-RAS drugs not because blocking RAS association with membranes is a flawed approach, but rather because FTIs failed to achieve this goal. Accordingly, we and others have continued to pursue the RAS modification and trafficking pathways as logical targets for potential therapeutics. One potential solution to the problem of alternative prenylation might be dual inhibition of FTase and GGTase I (20), either by combining individual inhibitors of each enzyme, or by dual specificity inhibitors. Such inhibitors have been developed and some have reached the clinic (16, 17), but have been limited by toxicity (21–23). There are still hopes that a therapeutic window can be found, possibly by targeting their delivery to RAS-driven cancer cells (17), where oncogenic RAS rendered cytosolic by these agents could sequester effectors and act as dominant negatives (24, 25).

Because farnesyl pyrophosphate (FPP) is both an intermediate in the cholesterol biosynthetic pathway and is also used for modification of CAAX proteins, it has long been speculated that statins may limit cancer by inhibiting prenylation of RAS or related small GTPases. However, due to the differential KMs for squalene synthase versus FTase, cholesterol synthesis is 1,000-fold more easily inhibited by loss of FPP than is FTase-mediated modification of RAS (26). Consistent with this, only suprapharmacologic levels of statins block protein prenylation in cell culture (27) and cause mislocalization of RAS (28), whereas therapeutic levels do not (28). Moreover, the effects of statins on cell growth are RAS-independent (27); nor is there evidence for these effects on RAS in animals administered pharmacologically relevant doses. Thus, statins show no promise for use as anti-RAS drugs.

Targeting Postprenylation CAAX Processing: RCE1 and ICMT

Although it is clear that the postprenylation CAAX-processing enzymes also contribute to RAS membrane association, making them potentially attractive targets for drug development, there are numerous challenges to overcome before inhibition of either RCE1 or ICMT can be translated to the clinic. These include uncertainty regarding the requirements for RCE1 and ICMT in oncogenic RAS functions, perplexing results from genetic validation studies, lack of understanding of the mechanism of RCE1 CAAX protease activity, and difficulties in developing inhibitors of these enzymes that can achieve sufficient potency and selectivity to become clinically efficacious drugs. Nevertheless, resolving these issues is likely to reveal important facets of RAS biology that may in turn reveal additional targets, warranting some discussion here.

RCE1 generates the substrate for ICMT: a prenylcysteine with an α carboxyl group. Thus, ICMT can act only after RCE1. The sequential nature of these modifications implies that RCE1 deficiency should produce cell biologic consequences at least as strong as ICMT deficiency, if one assumes that the effects of partial processing are similar for each substrate. At present, due to the lack of drug-like potent and selective inhibitors, this can be tested genetically but not pharmacologically. These studies reveal significant context dependence of deficiencies in these enzymes. Contrary to initial suppositions, Icmt-null mice die earlier in gestation than do Rce1-null mice (29, 30), and Icmt deficiency completely blocked the transformed growth of RAS-transformed rodent fibroblasts, whereas Rce1 deficiency had modest effects (31). Furthermore, the consequences of genetic loss of Rce1 and Icmt can be both opposing and context dependent. In a mouse model, the same myeloproliferative disease driven by oncogenic Kras G12D that was ameliorated by Icmt deficiency (32) was enhanced by Rce1 deficiency (33). However, Icmt deficiency accelerated the disease in a mouse model of pancreatic cancer driven by Kras G12D (34), due to inhibition of signaling from Notch-1, which acts as a tumor suppressor in this model (35). Importantly, Icmt deficiency ameliorated Kras-driven disease in other mouse tumor models (M.G. Dalin and M.O. Bergö; unpublished results), supporting the context dependency of ICMT impairment. These results suggest that a better understanding of the biology of CAAX processing will be critical to further validation of RCE1 and ICMT as potential drug targets, particularly with respect to the indications and populations in which they may be best applied. The complexity inherent in the context-dependent roles of RCE1 and ICMT is likely explained by the myriad substrates of these enzymes other than RAS, many of which are signaling molecules.

Many drugs fail in development due to cardiac toxicity, which is a demonstrated consequence of Rce1 depletion in the heart (36). Similarly, targeted depletion of Rce1 in the retina resulted in rapid degeneration of specific photoreceptor cells (37). On the other hand, the cleavage of RHOA by the bacterial toxin and cysteine protease YopT requires RCE1 but not ICMT (38). Thus, even if they can be developed, RCE1 inhibitors may be more toxic overall than ICMT inhibitors. Nevertheless, attempts have been made to develop inhibitors of both RCE1 (39) and ICMT (40–42), although each will necessarily also affect alternate substrates in addition to RAS. Disappointingly, even the most potent and selective of the RCE1 inhibitors have recently been shown to lack mechanism-based activity (43). The antiproliferative effects of ICMT inhibitors on RAS-transformed cells may be more compelling (40); whether they can be converted into pharmacologic leads is currently unclear.

RAS Trafficking: “Second Signals,” the Acylation/Deacylation Cycle, and RAS Chaperones

RAS membrane association and trafficking are regulated in a complex manner that has yet to be fully unraveled (Fig. 2). This complexity has revealed both additional challenges and additional targets for drug discovery. First, in addition to modification of the CAAX sequence, RAS plasma membrane association requires a proximal “second signal” that is either palmitoylation of one (NRAS) or two cysteines (HRAS) or a polybasic stretch of lysine residues (KRAS4B; refs. 44, 45) to confer additional hydrophobicity or an electrostatic interaction with the negatively charged headgroups of the phospholipids at the inner leaflet of the plasma membrane, respectively (Fig. 1). KRAS4A is unique among the four RAS proteins in possessing a dual membrane targeting motif that consists of both a palmitoylated cysteine and two short polybasic regions flanking that acylated cysteine (ref. 46; Fig. 1). Second, RAS association with membranes is not a one-way street. Instead, RAS proteins undergo a cycle of delivery to the plasma membrane followed by return to endomembranes for recycling, where they can interact with a Golgi-resident palmitoyl acyltransferase (47). Depalmitoylation of NRAS and HRAS at the plasma membrane initiates the recycling process (48, 49), whereas KRAS4B dissociation from the plasma membrane is mediated in part by PKC-catalyzed phosphorylation of serine 181 within the polybasic region (50). Third, RAS protein trafficking is also modulated by interactions with GDI-like chaperones (51, 52) that help to shepherd the lipidated RAS proteins among hydrophobic membranes through aqueous cytosol, and within specific plasma membrane regions (53). Among these chaperones are galectin-1 and-3 that escort HRAS and KRAS4B, respectively (54–56), and nucleolin that shepherds NRAS (57, 58). Phosphodiesterase-6δ (PDE6δ) has been variously proposed to chaperone NRAS (46, 59) or nonpalmitoylated HRAS (59), and KRAS4B (46, 60–62) but not KRAS4A (46). RAS proteins are released from PDE6δ upon binding of ARL2/3 to an allosteric site on PDE6δ (63). Each of these interactions is potentially a source of targets for anti-RAS drugs, and each has been pursued to varying degrees.

Targeting RAS Trafficking by Disrupting Palmitoylation: Protein Acyl (Palmitoyl) Transferases

Aside from CAAX-signaled modifications, enzymatic targets in the RAS processing pathway include those that regulate the palmitoylation/depalmitoylation cycle. KRAS, the predominant RAS gene mutated in human cancers, is expressed in two splice variants. The nonpalmitoylated KRAS4B has been generally accepted to be the major driver of cancer. However, recent work implicating the palmitoylated splice variant KRAS4A in colorectal adenocarcinoma (46) may force a re-evaluation of the notion that palmitoylation is not a good target in RAS-driven cancers. Moreover, palmitoylation inhibitors may help to limit the activity of oncogenic NRAS (64, 65), a key driver of melanoma and hematopoietic malignancies. As inhibitors of palmitoylation and depalmitoylation are developed, it may also be important to better understand the roles of palmitoylated wild-type RAS isoforms in supporting or impairing oncogenic RAS-driven cancers (1). Nonspecific inhibitors of protein palmitoylation such as 2-bromopalmitate (66) have been useful as tool compounds, but are unlikely to be developed into drugs given the vast number of palmitoylated proteins in the human genome (67). The recent identification of 23 DHHC (aspartic acid-histidine-histidine-cysteine tetrapeptide motif) proteins in the mammalian repertoire of protein acyltransferases (PAT) supports the feasibility of developing palmitoylation inhibitors that are specific for a subset of substrates such as RAS proteins (68). Consensus motifs to link specific palmitoylated substrates with their respective PATs have not yet been determined, but identification of the DHHC9/GPC16 complex as the PAT that modifies NRAS and HRAS (47) suggests the possibility of targeting this process selectively. Although progress in this direction has been limited to date (66, 68, 69), the recent advent of metabolic labeling with the bioorthogonal fatty acid 17-octadecynoic acid (17-ODYA) followed by click chemistry-mediated retrieval of labeled substrates promises to simplify transferase assays that can be applied to compound screens. In addition, because sequences around the palmitoylation sites stabilize membrane association and signaling of palmitoylated RAS proteins without affecting palmitoylation status itself (70, 71), these may also need to be taken into account for successful PAT inhibitor development.

Targeting RAS Trafficking by Disrupting Depalmitoylation: APT1/LYPLA1 Acyl Protein Thioesterases

There has been more progress in the area of inhibiting depalmitoylation, although translating the present tool compounds to drugs is also likely to be challenging. Counterintuitive although it may seem at first glance, the rationale for inhibiting depalmitoylation of palmitoylated RAS proteins is that depalmitoylation on all cellular membranes seems to be required for dynamic cycling of NRAS and HRAS among their membrane locations (72). This process is facilitated by acyl protein thioesterase 1 (APT1, also known as lysophospholipase 1, LYPLA1) and possibly also APT2, which are themselves reversibly S-acylated (73). The existence of APT in both palmitoylated Golgi-bound and nonpalmitoylated cytosolic pools (74) due to autodepalmitoylating activity of APTs (74) may explain how cytosolic (nonpalmitoylated) APT can promote cycling of palmitoylatable isoforms of RAS proteins to and from membranes, ultimately resulting in enrichment of the palmitoylated forms at the plasma membrane and Golgi (72, 74). These features provide the rationale for development of APT1/2 inhibitors as potential anti-RAS agents. The ability of a series of β-lactone–based inhibitors such as palmostatins B and M (derived from the over-the-counter weight loss drug tetrahydrolipstatin; refs. 75, 76) to mislocalize HRAS and NRAS proteins to internal membranes (75, 76) and to inhibit the proliferation of myeloid progenitor cells expressing oncogenic NRAS in treated mice (77) suggests that such inhibitors have the potential to disrupt oncogenic NRAS function, although questions remain regarding kinetics and target specificity (78). Additional tool compounds such as boron-based APT1/2 inhibitors have been identified (79). Other recent screens have identified LYPLA1/- and LYPLA2-selective inhibitors based on a triazole urea scaffold (80). Interestingly, APTs are not the only regulators of RAS depalmitoylation; for example, HRAS becomes depalmitoylated following the peptidyl-prolyl isomerase activity of FKBP12 on a proline near the palmitoylated cysteines (81). Understanding and optimizing the possible effects of such inhibitors may be complicated by the differential dynamics of wild-type and oncogenic RAS palmitoylation (82), and by the numerous non-RAS substrates of APT1/2/LYPLA1/2 that also undergo dynamic palmitoylation, including proteins as varied as heterotrimeric G protein alpha subunits, MAGUK scaffolds, Src family tyrosine kinases, nucleoporin, and BK ion channels (69). Nevertheless, the advent of new labeling technologies and new probes are making it possible to better study dynamic palmitoylation (83), which in turn is expected to reveal novel paths toward improved inhibitors of this key modification.

Targeting RAS Trafficking by Disrupting Other Modifications: PKC-Mediated Phosphorylation

In addition to CAAX processing and palmitoylation, other posttranslational modifications of RAS are potentially targetable (1, 84). Of these, phosphorylation of KRAS4B on serine 181, a process that is mutually exclusive with calmodulin binding (85), can alter subcellular localization dramatically, displacing the modified GTPase from the plasma membrane (50, 86), and converting KRAS4B from a growth-promoting to a growth-suppressing protein (87). Consistent with this, a PKC agonist, bryostatin, slowed the growth of mouse tumors driven by oncogenic KRAS4B but not those driven by a phosphorylation-deficient KRAS4B mutant (50). It is unclear why, in another study, rodent fibroblasts transformed with phosphorylation-deficient KRAS4B12V,181A failed to produce tumors in nude mice (88), leading these investigators to conclude that phosphorylation of KRAS4B is required for oncogenesis rather than leading to growth suppression. We have found that crossing p48-Cre mice with animals with a phosphorylation-deficient LSL-Kras4B-G12D/S181A double knock-in allele produced pancreatic tumors with equal frequency as crossing to mice with a phosphorylatable LSL-Kras4B-G12D/S181S allele (unpublished results), arguing strongly against a requirement for KRAS4B phosphorylation. Importantly, a recent study (89) showed that numerous cancer-associated mutations of PKC isoforms are loss-of-function mutations. This finding suggests that these PKCs act as tumor suppressors, consistent with the effect of phosphorylating KRAS4B on S181. Regardless of the explanation, although bryostatin and analogues have been under preclinical and clinical investigation as anticancer treatments, are reasonably well tolerated and have antitumor activity (90), enthusiasm for this approach to anti-KRAS therapy is diminished by the low probability of finding a drug that stimulates KRAS4B phosphorylation without affecting PKC-mediated activation of other signaling molecules that promote tumor growth and/or lead to toxicities.

Targeting RAS Trafficking by Disrupting RAS–Chaperone Interactions

In addition to blocking enzymatic activities regulating RAS membrane interactions, several distinct approaches have been taken to disrupting farnesylated RAS binding to chaperones, largely with the intent to block oncogenic KRAS4B specifically.

Inhibitors of the Ras–PDE6δ interaction have recently been identified. Deltarasin, at low micromolar levels, impaired the accumulation of KRAS4B on the plasma membrane and slowed the growth in vitro and in vivo of a tumor cell line harboring mutant KRAS (62, 91). However, the degree of dependence of KRAS on PDE6δ is not yet clear. For example, PDE6d knockout mice are viable and fertile (92), whereas knockout of Kras in mice is embryonic lethal (93), indicating that KRAS can still function in the absence of PDE6δ. On the other hand, the crystal structure of PDE6δ indicates little to no specificity for one farnesylated protein over another (63), which may be reflected in the discordant observations of RAS interactions mentioned above. It may also be that, similarly to effective multikinase inhibitors that were once disparaged as “dirty” drugs, inhibition of multiple farnesylated proteins contributes to the salutary effects of deltarasin. Future translation of current PDE6δ inhibitors to clinical leads will likely require a better understanding of its chaperone specificity (94) and trafficking patterns.

Salirasib, also known as farnesylthiosalicylic acid (FTS), resembles the S-farnesylated cysteine on Ras (95, 96) and is proposed to compete with it for binding to chaperones such as galectins (52, 56, 97). A small clinical trial showed that salirasib was well tolerated in pancreatic cancer patients (98), yet definitive answers as to whether it effectively perturbed KRAS function in patients, and can provide clinical benefit, remain to be determined. Like the other inhibitors of RAS membrane association, salirasib is not specific for RAS, but also inhibits other farnesylated proteins, including RHEB and MTOR (1). Larger trials are merited, but await a better understanding of salirasib mechanism of action and elucidation of tractable biomarkers.

Meanwhile, galectins are also known to modulate RAS nanoclustering and localization within defined membrane microdomains (53–56, 99), that in turn are regulated by phosphatidylserine (99). Thus, perhaps it was not surprising that an unbiased high-content screen to observe mislocalization of KRAS4B from plasma membrane to endomembranes revealed that staurosporine and analogues blocked endosomal recycling of phosphatidylserine and displaced KRAS4B from the plasma membrane to endosomes where it was degraded (100). Interestingly, fendiline, an L-type calcium channel blocker, was also identified as a compound that induced KRAS4B mislocalization (101); however, this activity was channel independent. The third class of compound identified in this visual screen was metformin, that also displaced KRAS4B from the plasma membrane (102). These findings are very exciting because they suggest that unbiased screens have the potential to uncover existing drugs that can be repurposed to block KRAS4B membrane association. To further identify new regulators of KRAS4B membrane association, the Philips lab has designed a genome-wide siRNA screen employing a dual luciferase assay that quantitatively reports displacement of KRAS4B from cellular membranes (unpublished data). We expect that additional roles for still other proteins in membrane targeting and/or stabilization have yet to be revealed. As this new information adds to the surge in understanding how nascent RAS proteins are delivered to the plasma membrane, we expect the RAS trafficking pathway to continue to provide a target-rich environment for drug discovery.

Authors' Contributions

Conception and design: A.D. Cox, C.J. Der, M.R. Philips

Writing, review, and/or revision of the manuscript: A.D. Cox, C.J. Der, M.R. Philips

Grant Support

A.D. Cox and C.J. Der were supported by the NIH under award numbers R01CA042978, R21CA179193, and R01CA175747; by a grant from the Lustgarten Foundation for Pancreatic Cancer Research; and the 2012 Pancreatic Cancer Action Network-AACR Innovative Grant, supported by Tempur-Pedic Retailers, grant number 12–60-25-DER. M.R. Philips was supported by the NIH under award numbers R01GM055279, R01CA116034, and R01CA163489.

Footnotes

  • Note: A.D. Cox, C.J. Der, and M.R. Philips share senior authorship.

  • Received January 26, 2015.
  • Revision received February 25, 2015.
  • Accepted February 25, 2015.

References

  1. 1.↵
    1. Cox AD,
    2. Fesik SW,
    3. Kimmelman AC,
    4. Luo J,
    5. Der CJ
    . Drugging the undruggable RAS: Mission Possible? Nat Rev Drug Discov 2014;13:828–51.
    OpenUrlCrossRefPubMed
  2. 2.↵
    1. Prior IA,
    2. Lewis PD,
    3. Mattos C
    . A comprehensive survey of Ras mutations in cancer. Cancer Res 2012;72:2457–67.
    OpenUrlAbstract/FREE Full Text
  3. 3.↵
    1. Marcus K,
    2. Mattos C
    . Direct attack on RAS: intramolecular communication and mutation-specific effects. Clin Cancer Res 2015;21:1810–8.
    OpenUrlAbstract/FREE Full Text
  4. 4.↵
    1. Stephen AG,
    2. Esposito D,
    3. Bagni RK,
    4. McCormick F
    . Dragging ras back in the ring. Cancer Cell 2014;25:272–81.
    OpenUrlCrossRefPubMed
  5. 5.↵
    1. Pylayeva-Gupta Y,
    2. Grabocka E,
    3. Bar-Sagi D
    . RAS oncogenes: weaving a tumorigenic web. Nat Rev Cancer 2011;11:761–74.
    OpenUrlCrossRefPubMed
  6. 6.↵
    1. Downward J
    . RAS synthetic lethal screens revisited: still seeking the elusive prize? Clin Cancer Res 2015;21:1802–9.
    OpenUrlAbstract/FREE Full Text
  7. 7.↵
    1. Kimmelman AC
    . Metabolic dependencies in RAS-driven cancers. Clin Cancer Res 2015;21:1828–34.
    OpenUrlAbstract/FREE Full Text
  8. 8.↵
    1. Jackson JH,
    2. Cochrane CG,
    3. Bourne JR,
    4. Solski PA,
    5. Buss JE,
    6. Der CJ
    . Farnesol modification of Kirsten-ras exon 4B protein is essential for transformation. Proc Natl Acad Sci U S A 1990;87:3042–6.
    OpenUrlAbstract/FREE Full Text
  9. 9.↵
    1. Willumsen BM,
    2. Christensen A,
    3. Hubbert NL,
    4. Papageorge AG,
    5. Lowy DR
    . The p21 ras C-terminus is required for transformation and membrane association. Nature 1984;310:583–6.
    OpenUrlCrossRefPubMed
  10. 10.↵
    1. Bivona TG,
    2. Perez De Castro I,
    3. Ahearn IM,
    4. Grana TM,
    5. Chiu VK,
    6. Lockyer PJ,
    7. et al.
    Phospholipase Cgamma activates ras on the golgi apparatus by means of RasGRP1. Nature 2003;424:694–8.
    OpenUrlCrossRefPubMed
  11. 11.↵
    1. Chiu VK,
    2. Bivona T,
    3. Hach A,
    4. Sajous JB,
    5. Silletti J,
    6. Wiener H,
    7. et al.
    Ras signalling on the endoplasmic reticulum and the Golgi. Nat Cell Biol 2002;4:343–50.
    OpenUrlCrossRefPubMed
  12. 12.↵
    1. Cox AD,
    2. Der CJ
    . Ras history: the saga continues. Small GTPases 2010;1:2–27.
    OpenUrlCrossRefPubMed
  13. 13.↵
    1. Wright LP,
    2. Philips MR
    . Thematic review series: lipid posttranslational modifications. CAAX modification and membrane targeting of Ras. J Lipid Res 2006;47:883–91.
    OpenUrlAbstract/FREE Full Text
  14. 14.↵
    1. Dai Q,
    2. Choy E,
    3. Chiu V,
    4. Romano J,
    5. Slivka SR,
    6. Steitz SA,
    7. et al.
    Mammalian prenylcysteine carboxyl methyltransferase is in the endoplasmic reticulum. J Biol Chem 1998;273:15030–4.
    OpenUrlAbstract/FREE Full Text
  15. 15.↵
    1. Schmidt WK,
    2. Tam A,
    3. Fujimura-Kamada K,
    4. Michaelis S
    . Endoplasmic reticulum membrane localization of Rce1p and Ste24p, yeast proteases involved in carboxyl-terminal CAAX protein processing and amino-terminal a-factor cleavage. Proc Natl Acad Sci U S A 1998;95:11175–80.
    OpenUrlAbstract/FREE Full Text
  16. 16.↵
    1. Berndt N,
    2. Hamilton AD,
    3. Sebti SM
    . Targeting protein prenylation for cancer therapy. Nat Rev Cancer 2011;11:775–91.
    OpenUrlCrossRefPubMed
  17. 17.↵
    1. Tamanoi F,
    2. Lu J
    . Recent progress in developing small molecule inhibitors designed to interfere with ras membrane association: toward inhibiting K-Ras and N-Ras functions. Enzymes 2013;34 Pt. B:181–200.
    OpenUrlCrossRef
  18. 18.↵
    1. Rowell CA,
    2. Kowalczyk JJ,
    3. Lewis MD,
    4. Garcia AM
    . Direct demonstration of geranylgeranylation and farnesylation of Ki-Ras in vivo. J Biol Chem 1997;272:14093–7.
    OpenUrlAbstract/FREE Full Text
  19. 19.↵
    1. Whyte DB,
    2. Kirschmeier P,
    3. Hockenberry TN,
    4. Nunez-Oliva I,
    5. James L,
    6. Catino JJ,
    7. et al.
    K- and N-Ras are geranylgeranylated in cells treated with farnesyl protein transferase inhibitors. J Biol Chem 1997;272:14459–64.
    OpenUrlAbstract/FREE Full Text
  20. 20.↵
    1. Liu M,
    2. Sjogren AK,
    3. Karlsson C,
    4. Ibrahim MX,
    5. Andersson KM,
    6. Olofsson FJ,
    7. et al.
    Targeting the protein prenyltransferases efficiently reduces tumor development in mice with K-RAS-induced lung cancer. Proc Natl Acad Sci USA 2010;107:6471–6.
    OpenUrlAbstract/FREE Full Text
  21. 21.↵
    1. deSolms SJ,
    2. Ciccarone TM,
    3. MacTough SC,
    4. Shaw AW,
    5. Buser CA,
    6. Ellis-Hutchings M,
    7. et al.
    Dual protein farnesyltransferase-geranylgeranyltransferase-I inhibitors as potential cancer chemotherapeutic agents. J Med Chem 2003;46:2973–84.
    OpenUrlCrossRefPubMed
  22. 22.↵
    1. Lobell RB,
    2. Liu D,
    3. Buser CA,
    4. Davide JP,
    5. DePuy E,
    6. Hamilton K,
    7. et al.
    Preclinical and clinical pharmacodynamic assessment of L-778,123, a dual inhibitor of farnesyl:protein transferase and geranylgeranyl:protein transferase type-I. Mol Cancer Ther 2002;1:747–58.
    OpenUrlAbstract/FREE Full Text
  23. 23.↵
    1. Lobell RB,
    2. Omer CA,
    3. Abrams MT,
    4. Bhimnathwala HG,
    5. Brucker MJ,
    6. Buser CA,
    7. et al.
    Evaluation of farnesyl:protein transferase and geranylgeranyl:protein transferase inhibitor combinations in preclinical models. Cancer Res 2001;61:8758–68.
    OpenUrlAbstract/FREE Full Text
  24. 24.↵
    1. Fiordalisi JJ,
    2. Holly SP,
    3. Johnson RL 2nd.,
    4. Parise LV,
    5. Cox AD
    . A distinct class of dominant negative Ras mutants: cytosolic GTP-bound Ras effector domain mutants that inhibit Ras signaling and transformation and enhance cell adhesion. J Biol Chem 2002;277:10813–23.
    OpenUrlAbstract/FREE Full Text
  25. 25.↵
    1. Lerner EC,
    2. Qian Y,
    3. Blaskovich MA,
    4. Fossum RD,
    5. Vogt A,
    6. Sun J,
    7. et al.
    Ras CAAX peptidomimetic FTI-277 selectively blocks oncogenic Ras signaling by inducing cytoplasmic accumulation of inactive Ras-Raf complexes. J Biol Chem 1995;270:26802–6.
    OpenUrlAbstract/FREE Full Text
  26. 26.↵
    1. Sinensky M,
    2. Beck LA,
    3. Leonard S,
    4. Evans R
    . Differential inhibitory effects of lovastatin on protein isoprenylation and sterol synthesis. J Biol Chem 1990;265:19937–41.
    OpenUrlAbstract/FREE Full Text
  27. 27.↵
    1. DeClue JE,
    2. Vass WC,
    3. Papageorge AG,
    4. Lowy DR,
    5. Willumsen BM
    . Inhibition of cell growth by lovastatin is independent of ras function. Cancer Res 1991;51:712–7.
    OpenUrlAbstract/FREE Full Text
  28. 28.↵
    1. Cho KJ,
    2. Hill MM,
    3. Chigurupati S,
    4. Du G,
    5. Parton RG,
    6. Hancock JF
    . Therapeutic levels of the hydroxmethylglutaryl-coenzyme A reductase inhibitor lovastatin activate ras signaling via phospholipase D2. Mol Cell Biol 2011;31:1110–20.
    OpenUrlAbstract/FREE Full Text
  29. 29.↵
    1. Bergo MO,
    2. Leung GK,
    3. Ambroziak P,
    4. Otto JC,
    5. Casey PJ,
    6. Gomes AQ,
    7. et al.
    Isoprenylcysteine carboxyl methyltransferase deficiency in mice. J Biol Chem 2001;276:5841–5.
    OpenUrlAbstract/FREE Full Text
  30. 30.↵
    1. Lin X,
    2. Jung J,
    3. Kang D,
    4. Xu B,
    5. Zaret KS,
    6. Zoghbi H
    . Prenylcysteine carboxylmethyltransferase is essential for the earliest stages of liver development in mice. Gastroenterology 2002;123:345–51.
    OpenUrlCrossRefPubMed
  31. 31.↵
    1. Bergo MO,
    2. Ambroziak P,
    3. Gregory C,
    4. George A,
    5. Otto JC,
    6. Kim E,
    7. et al.
    Absence of the CAAX endoprotease Rce1: effects on cell growth and transformation. Mol Cell Biol 2002;22:171–81.
    OpenUrlAbstract/FREE Full Text
  32. 32.↵
    1. Wahlstrom AM,
    2. Cutts BA,
    3. Liu M,
    4. Lindskog A,
    5. Karlsson C,
    6. Sjogren AK,
    7. et al.
    Inactivating Icmt ameliorates K-RAS-induced myeloproliferative disease. Blood 2008;112:1357–65.
    OpenUrlAbstract/FREE Full Text
  33. 33.↵
    1. Wahlstrom AM,
    2. Cutts BA,
    3. Karlsson C,
    4. Andersson KM,
    5. Liu M,
    6. Sjogren AK,
    7. et al.
    Rce1 deficiency accelerates the development of K-RAS-induced myeloproliferative disease. Blood 2007;109:763–8.
    OpenUrlAbstract/FREE Full Text
  34. 34.↵
    1. Court H,
    2. Amoyel M,
    3. Hackman M,
    4. Lee KE,
    5. Xu R,
    6. Miller G,
    7. et al.
    Isoprenylcysteine carboxylmethyltransferase deficiency exacerbates KRAS-driven pancreatic neoplasia via Notch suppression. J Clin Invest 2013;123:4681–94.
    OpenUrlCrossRefPubMed
  35. 35.↵
    1. Hanlon L,
    2. Avila JL,
    3. Demarest RM,
    4. Troutman S,
    5. Allen M,
    6. Ratti F,
    7. et al.
    Notch1 functions as a tumor suppressor in a model of K-ras-induced pancreatic ductal adenocarcinoma. Cancer Res 2010;70:4280–6.
    OpenUrlAbstract/FREE Full Text
  36. 36.↵
    1. Bergo MO,
    2. Lieu HD,
    3. Gavino BJ,
    4. Ambroziak P,
    5. Otto JC,
    6. Casey PJ,
    7. et al.
    On the physiological importance of endoproteolysis of CAAX proteins: heart-specific RCE1 knockout mice develop a lethal cardiomyopathy. J Biol Chem 2004;279:4729–36.
    OpenUrlAbstract/FREE Full Text
  37. 37.↵
    1. Christiansen JR,
    2. Kolandaivelu S,
    3. Bergo MO,
    4. Ramamurthy V
    . RAS-converting enzyme 1-mediated endoproteolysis is required for trafficking of rod phosphodiesterase 6 to photoreceptor outer segments. Proc Natl Acad Sci U S A 2011;108:8862–6.
    OpenUrlAbstract/FREE Full Text
  38. 38.↵
    1. Fueller F,
    2. Bergo MO,
    3. Young SG,
    4. Aktories K,
    5. Schmidt G
    . Endoproteolytic processing of RhoA by Rce1 is required for the cleavage of RhoA by Yersinia enterocolitica outer protein T. Infect Immun 2006;74:1712–7.
    OpenUrlAbstract/FREE Full Text
  39. 39.↵
    1. Manandhar SP,
    2. Hildebrandt ER,
    3. Schmidt WK
    . Small-molecule inhibitors of the Rce1p CaaX protease. J Biomol Screen 2007;12:983–93.
    OpenUrlAbstract/FREE Full Text
  40. 40.↵
    1. Judd WR,
    2. Slattum PM,
    3. Hoang KC,
    4. Bhoite L,
    5. Valppu L,
    6. Alberts G,
    7. et al.
    Discovery and SAR of methylated tetrahydropyranyl derivatives as inhibitors of isoprenylcysteine carboxyl methyltransferase (ICMT). J Med Chem 2011;54:5031–47.
    OpenUrlCrossRefPubMed
  41. 41.↵
    1. Lau HY,
    2. Ramanujulu PM,
    3. Guo D,
    4. Yang T,
    5. Wirawan M,
    6. Casey PJ,
    7. et al.
    An improved isoprenylcysteine carboxylmethyltransferase inhibitor induces cancer cell death and attenuates tumor growth in vivo. Cancer Biol Ther 2014;15:1280–91.
    OpenUrlCrossRefPubMed
  42. 42.↵
    1. Winter-Vann AM,
    2. Baron RA,
    3. Wong W,
    4. delaCruz J,
    5. York JD,
    6. Gooden DM,
    7. et al.
    A small-molecule inhibitor of isoprenylcysteine carboxyl methyltransferase with antitumor activity in cancer cells. Proc Natl Acad Sci U S A 2005;102:4336–41.
    OpenUrlAbstract/FREE Full Text
  43. 43.↵
    1. Dechert AM,
    2. MacNamara JP,
    3. Breevoort SR,
    4. Hildebrandt ER,
    5. Hembree NW,
    6. Rea AC,
    7. et al.
    Modulation of the inhibitor properties of dipeptidyl (acyloxy)methyl ketones toward the CaaX proteases. Bioorg Med Chem 2010;18:6230–7.
    OpenUrlCrossRefPubMed
  44. 44.↵
    1. Hancock JF,
    2. Cadwallader K,
    3. Paterson H,
    4. Marshall CJ
    . A CAAX or a CAAL motif and a second signal are sufficient for plasma membrane targeting of ras proteins. EMBO J 1991;10:4033–9.
    OpenUrlPubMed
  45. 45.↵
    1. Hancock JF,
    2. Paterson H,
    3. Marshall CJ
    . A polybasic domain or palmitoylation is required in addition to the CAAX motif to localize p21ras to the plasma membrane. Cell 1990;63:133–9.
    OpenUrlCrossRefPubMed
  46. 46.↵
    1. Tsai FD,
    2. Lopes MS,
    3. Zhou M,
    4. Court H,
    5. Ponce O,
    6. Fiordalisi JJ,
    7. et al.
    K-Ras4A splice variant is widely expressed in cancer and uses a hybrid membrane-targeting motif. Proc Natl Acad Sci U S A 2015;112:779–94.
    OpenUrlAbstract/FREE Full Text
  47. 47.↵
    1. Swarthout JT,
    2. Lobo S,
    3. Farh L,
    4. Croke MR,
    5. Greentree WK,
    6. Deschenes RJ,
    7. et al.
    DHHC9 and GCP16 constitute a human protein fatty acyltransferase with specificity for H- and N-Ras. J Biol Chem 2005;280:31141–8.
    OpenUrlAbstract/FREE Full Text
  48. 48.↵
    1. Goodwin JS,
    2. Drake KR,
    3. Rogers C,
    4. Wright L,
    5. Lippincott-Schwartz J,
    6. Philips MR,
    7. et al.
    Depalmitoylated Ras traffics to and from the Golgi complex via a nonvesicular pathway. J Cell Biol 2005;170:261–72.
    OpenUrlAbstract/FREE Full Text
  49. 49.↵
    1. Rocks O,
    2. Peyker A,
    3. Kahms M,
    4. Verveer PJ,
    5. Koerner C,
    6. Lumbierres M,
    7. et al.
    An acylation cycle regulates localization and activity of palmitoylated Ras isoforms. Science 2005;307:1746–52.
    OpenUrlAbstract/FREE Full Text
  50. 50.↵
    1. Bivona TG,
    2. Quatela SE,
    3. Bodemann BO,
    4. Ahearn IM,
    5. Soskis MJ,
    6. Mor A,
    7. et al.
    PKC regulates a farnesyl-electrostatic switch on K-Ras that promotes its association with Bcl-XL on mitochondria and induces apoptosis. Mol Cell 2006;21:481–93.
    OpenUrlCrossRefPubMed
  51. 51.↵
    1. Nancy V,
    2. Callebaut I,
    3. El Marjou A,
    4. de Gunzburg J
    . The delta subunit of retinal rod cGMP phosphodiesterase regulates the membrane association of Ras and Rap GTPases. J Biol Chem 2002;277:15076–84.
    OpenUrlAbstract/FREE Full Text
  52. 52.↵
    1. Rotblat B,
    2. Niv H,
    3. Andre S,
    4. Kaltner H,
    5. Gabius HJ,
    6. Kloog Y
    . Galectin-1(L11A) predicted from a computed galectin-1 farnesyl-binding pocket selectively inhibits Ras-GTP. Cancer Res 2004;64:3112–8.
    OpenUrlAbstract/FREE Full Text
  53. 53.↵
    1. Abankwa D,
    2. Gorfe AA,
    3. Inder K,
    4. Hancock JF
    . Ras membrane orientation and nanodomain localization generate isoform diversity. Proc Natl Acad Sci U S A 2010;107:1130–5.
    OpenUrlAbstract/FREE Full Text
  54. 54.↵
    1. Ashery U,
    2. Yizhar O,
    3. Rotblat B,
    4. Elad-Sfadia G,
    5. Barkan B,
    6. Haklai R,
    7. et al.
    Spatiotemporal organization of Ras signaling: rasosomes and the galectin switch. Cell Mol Neurobiol 2006;26:471–95.
    OpenUrlPubMed
  55. 55.↵
    1. Belanis L,
    2. Plowman SJ,
    3. Rotblat B,
    4. Hancock JF,
    5. Kloog Y
    . Galectin-1 is a novel structural component and a major regulator of h-ras nanoclusters. Mol Biol Cell 2008;19:1404–14.
    OpenUrlAbstract/FREE Full Text
  56. 56.↵
    1. Shalom-Feuerstein R,
    2. Plowman SJ,
    3. Rotblat B,
    4. Ariotti N,
    5. Tian T,
    6. Hancock JF,
    7. et al.
    K-ras nanoclustering is subverted by overexpression of the scaffold protein galectin-3. Cancer Res 2008;68:6608–16.
    OpenUrlAbstract/FREE Full Text
  57. 57.↵
    1. Farin K,
    2. Schokoroy S,
    3. Haklai R,
    4. Cohen-Or I,
    5. Elad-Sfadia G,
    6. Reyes-Reyes ME,
    7. et al.
    Oncogenic synergism between ErbB1, nucleolin, and mutant Ras. Cancer Res 2011;71:2140–51.
    OpenUrlAbstract/FREE Full Text
  58. 58.↵
    1. Inder KL,
    2. Lau C,
    3. Loo D,
    4. Chaudhary N,
    5. Goodall A,
    6. Martin S,
    7. et al.
    Nucleophosmin and nucleolin regulate K-Ras plasma membrane interactions and MAPK signal transduction. J Biol Chem 2009;284:28410–9.
    OpenUrlAbstract/FREE Full Text
  59. 59.↵
    1. Chandra A,
    2. Grecco HE,
    3. Pisupati V,
    4. Perera D,
    5. Cassidy L,
    6. Skoulidis F,
    7. et al.
    The GDI-like solubilizing factor PDEdelta sustains the spatial organization and signalling of Ras family proteins. Nat Cell Biol 2012;14:148–58.
    OpenUrlCrossRefPubMed
  60. 60.↵
    1. Bhagatji P,
    2. Leventis R,
    3. Rich R,
    4. Lin CJ,
    5. Silvius JR
    . Multiple cellular proteins modulate the dynamics of K-ras association with the plasma membrane. Biophys J 2010;99:3327–35.
    OpenUrlCrossRefPubMed
  61. 61.↵
    1. Schmick M,
    2. Vartak N,
    3. Papke B,
    4. Kovacevic M,
    5. Truxius DC,
    6. Rossmannek L,
    7. et al.
    KRas localizes to the plasma membrane by spatial cycles of solubilization, trapping and vesicular transport. Cell 2014;157:459–71.
    OpenUrlCrossRefPubMed
  62. 62.↵
    1. Zimmermann G,
    2. Papke B,
    3. Ismail S,
    4. Vartak N,
    5. Chandra A,
    6. Hoffmann M,
    7. et al.
    Small molecule inhibition of the KRAS-PDEdelta interaction impairs oncogenic KRAS signalling. Nature 2013;497:638–42.
    OpenUrlCrossRefPubMed
  63. 63.↵
    1. Ismail SA,
    2. Chen YX,
    3. Rusinova A,
    4. Chandra A,
    5. Bierbaum M,
    6. Gremer L,
    7. et al.
    Arl2-GTP and Arl3-GTP regulate a GDI-like transport system for farnesylated cargo. Nat Chem Biol 2011;7:942–9.
    OpenUrlCrossRefPubMed
  64. 64.↵
    1. Song SP,
    2. Hennig A,
    3. Schubert K,
    4. Markwart R,
    5. Schmidt P,
    6. Prior IA,
    7. et al.
    Ras palmitoylation is necessary for N-Ras activation and signal propagation in growth factor signalling. Biochem J 2013;454:323–32.
    OpenUrlAbstract/FREE Full Text
  65. 65.↵
    1. Cuiffo B,
    2. Ren R
    . Palmitoylation of oncogenic NRAS is essential for leukemogenesis. Blood 2010;115:3598–605.
    OpenUrlAbstract/FREE Full Text
  66. 66.↵
    1. Resh MD
    . Targeting protein lipidation in disease. Trends Mol Med 2012;18:206–14.
    OpenUrlCrossRefPubMed
  67. 67.↵
    1. Yang W,
    2. Di Vizio D,
    3. Kirchner M,
    4. Steen H,
    5. Freeman MR
    . Proteome scale characterization of human S-acylated proteins in lipid raft-enriched and non-raft membranes. Mol Cell Proteomics 2010;9:54–70.
    OpenUrlAbstract/FREE Full Text
  68. 68.↵
    1. Chavda B,
    2. Arnott JA,
    3. Planey SL
    . Targeting protein palmitoylation: selective inhibitors and implications in disease. Expert Opin Drug Discov 2014;9:1005–19.
    OpenUrlCrossRefPubMed
  69. 69.↵
    1. Hernandez JL,
    2. Majmudar JD,
    3. Martin BR
    . Profiling and inhibiting reversible palmitoylation. Curr Opin Chem Biol 2013;17:20–6.
    OpenUrlCrossRefPubMed
  70. 70.↵
    1. Willumsen BM,
    2. Cox AD,
    3. Solski PA,
    4. Der CJ,
    5. Buss JE
    . Novel determinants of H-Ras plasma membrane localization and transformation. Oncogene 1996;13:1901–9.
    OpenUrlPubMed
  71. 71.↵
    1. Laude AJ,
    2. Prior IA
    . Palmitoylation and localisation of RAS isoforms are modulated by the hypervariable linker domain. J Cell Sci 2008;121:421–7.
    OpenUrlAbstract/FREE Full Text
  72. 72.↵
    1. Rocks O,
    2. Gerauer M,
    3. Vartak N,
    4. Koch S,
    5. Huang ZP,
    6. Pechlivanis M,
    7. et al.
    The palmitoylation machinery is a spatially organizing system for peripheral membrane proteins. Cell 2010;141:458–71.
    OpenUrlCrossRefPubMed
  73. 73.↵
    1. Kong E,
    2. Peng S,
    3. Chandra G,
    4. Sarkar C,
    5. Zhang Z,
    6. Bagh MB,
    7. et al.
    Dynamic palmitoylation links cytosol-membrane shuttling of acyl-protein thioesterase-1 and acyl-protein thioesterase-2 with that of proto-oncogene H-ras product and growth-associated protein-43. J Biol Chem 2013;288:9112–25.
    OpenUrlAbstract/FREE Full Text
  74. 74.↵
    1. Vartak N,
    2. Papke B,
    3. Grecco HE,
    4. Rossmannek L,
    5. Waldmann H,
    6. Hedberg C,
    7. et al.
    The autodepalmitoylating activity of APT maintains the spatial organization of palmitoylated membrane proteins. Biophys J 2014;106:93–105.
    OpenUrlCrossRefPubMed
  75. 75.↵
    1. Dekker FJ,
    2. Rocks O,
    3. Vartak N,
    4. Menninger S,
    5. Hedberg C,
    6. Balamurugan R,
    7. et al.
    Small-molecule inhibition of APT1 affects Ras localization and signaling. Nat Chem Biol 2010;6:449–56.
    OpenUrlCrossRefPubMed
  76. 76.↵
    1. Hedberg C,
    2. Dekker FJ,
    3. Rusch M,
    4. Renner S,
    5. Wetzel S,
    6. Vartak N,
    7. et al.
    Development of highly potent inhibitors of the Ras-targeting human acyl protein thioesterases based on substrate similarity design. Angew Chem Int Ed Engl 2011;50:9832–7.
    OpenUrlCrossRefPubMed
  77. 77.↵
    1. Xu J,
    2. Hedberg C,
    3. Dekker FJ,
    4. Li Q,
    5. Haigis KM,
    6. Hwang E,
    7. et al.
    Inhibiting the palmitoylation/depalmitoylation cycle selectively reduces the growth of hematopoietic cells expressing oncogenic Nras. Blood 2012;119:1032–5.
    OpenUrlAbstract/FREE Full Text
  78. 78.↵
    1. Cox AD
    . Protein localization: Can too much lipid glue stop Ras? Nat Chem Biol 2010;6:483–5.
    OpenUrlCrossRefPubMed
  79. 79.↵
    1. Zimmermann TJ,
    2. Burger M,
    3. Tashiro E,
    4. Kondoh Y,
    5. Martinez NE,
    6. Gormer K,
    7. et al.
    Boron-based inhibitors of acyl protein thioesterases 1 and 2. Chembiochem 2013;14:115–22.
    OpenUrlCrossRefPubMed
  80. 80.↵
    1. Adibekian A,
    2. Martin BR,
    3. Chang JW,
    4. Hsu KL,
    5. Tsuboi K,
    6. Bachovchin DA,
    7. et al.
    Characterization of a selective, reversible inhibitor of lysophospholipase 1 (LYPLA1). Probe Reports from the NIH Molecular Libraries Program [monograph on the Internet]. Bethesda, MD: National Center for Biotechnology Information; 2010 [cited 2015 Jan 26]. Available from http://www.ncbi.nlm.nih.gov/books/NBK189924/.
  81. 81.↵
    1. Ahearn IM,
    2. Tsai FD,
    3. Court H,
    4. Zhou M,
    5. Jennings BC,
    6. Ahmed M,
    7. et al.
    FKBP12 binds to acylated H-ras and promotes depalmitoylation. Mol Cell 2011;41:173–85.
    OpenUrlCrossRefPubMed
  82. 82.↵
    1. Baker TL,
    2. Zheng H,
    3. Walker J,
    4. Coloff JL,
    5. Buss JE
    . Distinct rates of palmitate turnover on membrane-bound cellular and oncogenic H-ras. J Biol Chem 2003;278:19292–300.
    OpenUrlAbstract/FREE Full Text
  83. 83.↵
    1. Martin BR
    . Chemical approaches for profiling dynamic palmitoylation. Biochem Soc Trans 2013;41:43–9.
    OpenUrlAbstract/FREE Full Text
  84. 84.↵
    1. Ahearn IM,
    2. Haigis K,
    3. Bar-Sagi D,
    4. Philips MR
    . Regulating the regulator: post-translational modification of RAS. Nat Rev Mol Cell Biol 2012;13:39–51.
    OpenUrlCrossRefPubMed
  85. 85.↵
    1. Villalonga P,
    2. Lopez-Alcala C,
    3. Chiloeches A,
    4. Gil J,
    5. Marais R,
    6. Bachs O,
    7. et al.
    Calmodulin prevents activation of Ras by PKC in 3T3 fibroblasts. J Biol Chem 2002;277:37929–35.
    OpenUrlAbstract/FREE Full Text
  86. 86.↵
    1. Plowman SJ,
    2. Ariotti N,
    3. Goodall A,
    4. Parton RG,
    5. Hancock JF
    . Electrostatic interactions positively regulate K-Ras nanocluster formation and function. Mol Cell Biol 2008;28:4377–85.
    OpenUrlAbstract/FREE Full Text
  87. 87.↵
    1. Sung PJ,
    2. Tsai FD,
    3. Vais H,
    4. Court H,
    5. Yang J,
    6. Fehrenbacher N,
    7. et al.
    Phosphorylated K-Ras limits cell survival by blocking Bcl-xL sensitization of inositol trisphosphate receptors. Proc Natl Acad Sci U S A 2013;110:20593–8.
    OpenUrlAbstract/FREE Full Text
  88. 88.↵
    1. Barcelo C,
    2. Paco N,
    3. Morell M,
    4. Alvarez-Moya B,
    5. Bota-Rabassedas N,
    6. Jaumot M,
    7. et al.
    Phosphorylation at Ser-181 of oncogenic KRAS is required for tumor growth. Cancer Res 2014;74:1190–9.
    OpenUrlAbstract/FREE Full Text
  89. 89.↵
    1. Antal CE,
    2. Hudson AM,
    3. Kang E,
    4. Zanca C,
    5. Wirth C,
    6. Stephenson NL,
    7. et al.
    Cancer-associated protein kinase C mutations reveal kinase's role as tumor suppressor. Cell 2015;160:489–502.
    OpenUrlCrossRefPubMed
  90. 90.↵
    1. Kollar P,
    2. Rajchard J,
    3. Balounova Z,
    4. Pazourek J
    . Marine natural products: bryostatins in preclinical and clinical studies. Pharm Biol 2014;52:237–42.
    OpenUrlCrossRefPubMed
  91. 91.↵
    1. Zimmermann G,
    2. Schultz-Fademrecht C,
    3. Kuchler P,
    4. Murarka S,
    5. Ismail S,
    6. Triola G,
    7. et al.
    Structure guided design and kinetic analysis of highly potent benzimidazole inhibitors targeting the PDEdelta prenyl binding site. J Med Chem 2014;57:5435–48.
    OpenUrlCrossRefPubMed
  92. 92.↵
    1. Zhang H,
    2. Li S,
    3. Doan T,
    4. Rieke F,
    5. Detwiler PB,
    6. Frederick JM,
    7. et al.
    Deletion of PrBP/delta impedes transport of GRK1 and PDE6 catalytic subunits to photoreceptor outer segments. Proc Natl Acad Sci U S A 2007;104:8857–62.
    OpenUrlAbstract/FREE Full Text
  93. 93.↵
    1. Johnson L,
    2. Greenbaum D,
    3. Cichowski K,
    4. Mercer K,
    5. Murphy E,
    6. Schmitt E,
    7. et al.
    K-ras is an essential gene in the mouse with partial functional overlap with N-ras. Genes Dev 1997;11:2468–81.
    OpenUrlAbstract/FREE Full Text
  94. 94.↵
    1. Azoulay-Alfaguter I,
    2. Strazza M,
    3. Mor A
    . Chaperone-mediated specificity in Ras and Rap signaling. Crit Rev Biochem Mol Biol 2014:1–9.
  95. 95.↵
    1. Elad G,
    2. Paz A,
    3. Haklai R,
    4. Marciano D,
    5. Cox A,
    6. Kloog Y
    . Targeting of K-Ras 4B by S-trans, trans-farnesyl thiosalicylic acid. Biochim Biophys Acta 1999;1452:228–42.
    OpenUrlPubMed
  96. 96.↵
    1. Haklai R,
    2. Weisz MG,
    3. Elad G,
    4. Paz A,
    5. Marciano D,
    6. Egozi Y,
    7. et al.
    Dislodgment and accelerated degradation of Ras. Biochemistry 1998;37:1306–14.
    OpenUrlCrossRefPubMed
  97. 97.↵
    1. Kloog Y,
    2. Elad-Sfadia G,
    3. Haklai R,
    4. Mor A
    . Ras chaperones: new targets for cancer and immunotherapy. Enzymes 2013;33 Pt A:267–89.
    OpenUrlCrossRef
  98. 98.↵
    1. Laheru D,
    2. Shah P,
    3. Rajeshkumar NV,
    4. McAllister F,
    5. Taylor G,
    6. Goldsweig H,
    7. et al.
    Integrated preclinical and clinical development of S-trans, trans-Farnesylthiosalicylic Acid (FTS, Salirasib) in pancreatic cancer. Invest New Drugs 2012;30:2391–9.
    OpenUrlCrossRefPubMed
  99. 99.↵
    1. Zhou Y,
    2. Liang H,
    3. Rodkey T,
    4. Ariotti N,
    5. Parton RG,
    6. Hancock JF
    . Signal integration by lipid-mediated spatial cross talk between Ras nanoclusters. Mol Cell Biol 2014;34:862–76.
    OpenUrlAbstract/FREE Full Text
  100. 100.↵
    1. Cho KJ,
    2. Park JH,
    3. Piggott AM,
    4. Salim AA,
    5. Gorfe AA,
    6. Parton RG,
    7. et al.
    Staurosporines disrupt phosphatidylserine trafficking and mislocalize Ras proteins. J Biol Chem 2012;287:43573–84.
    OpenUrlAbstract/FREE Full Text
  101. 101.↵
    1. van der Hoeven D,
    2. Cho KJ,
    3. Ma X,
    4. Chigurupati S,
    5. Parton RG,
    6. Hancock JF
    . Fendiline inhibits K-Ras plasma membrane localization and blocks K-Ras signal transmission. Mol Cell Biol 2013;33:237–51.
    OpenUrlAbstract/FREE Full Text
  102. 102.↵
    1. Cho KJ,
    2. van der Hoeven D,
    3. Hancock JF
    . Inhibitors of K-Ras plasma membrane localization. Enzymes 2013;33 Pt A:249–65.
    OpenUrlCrossRef
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Clinical Cancer Research: 21 (8)
April 2015
Volume 21, Issue 8

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Targeting RAS Membrane Association: Back to the Future for Anti-RAS Drug Discovery?
Adrienne D. Cox, Channing J. Der and Mark R. Philips
Clin Cancer Res April 15 2015 (21) (8) 1819-1827; DOI: 10.1158/1078-0432.CCR-14-3214
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