Mutation-Specific RAS Oncogenicity Explains N-RAS Codon 61 Selection in Melanoma

Abstract

N-RAS mutation at codon 12, 13 or 61 is associated with transformation; yet, in melanoma, such alterations are nearly exclusive to codon 61. Here, we compared the melanoma susceptibility of an N-Ras^Q61R knock-in allele to similarly designed K-Ras^G12D and N-Ras^G12D alleles. With concomitant p16^INK4a inactivation, K-Ras^G12D or N-Ras^Q61R expression efficiently promoted melanoma in vivo, whereas N-Ras^G12D did not. Additionally, N-Ras^Q61R mutation potently cooperated with Lkb1/Stk11 loss to induce highly metastatic disease. Functional comparisons of N-Ras^Q61R and N-Ras^G12D revealed little difference in the ability of these proteins to engage PI3K or RAF. Instead, N-Ras^Q61R showed enhanced nucleotide binding, decreased intrinsic GTPase activity and increased stability when compared to N-Ras^G12D. This work identifies a faithful model of human N-RAS mutant melanoma, and suggests that the increased melanomagenecity of N-Ras^Q61R over N-Ras^G12D is due to heightened abundance of the active, GTP-bound form rather than differences in the engagement of downstream effector pathways.

Introduction

One-third of all human cancers harbor activating K-, H- or N-RAS mutations, which localize predominantly to codons 12, 13 or 61 (1, 2). RAS proteins function as canonical GTPase switches, binding to effectors in the presence of GTP and activating downstream signaling pathways to influence cellular proliferation, differentiation and survival. Return of RAS to an inactive, GDP-bound state is catalyzed by GTPase activating proteins (GAPs), which stimulate the weak, intrinsic GTPase activity of these proteins. Mutations at codons 12 or 13 render RAS proteins insensitive to GAP activity, resulting in constitutive, oncogenic signaling (3). Similarly, mutation of Q61, a catalytic residue required for efficient GTP hydrolysis, impedes the return of RAS to an inactive GDP-bound state (4).

Historically, RAS proteins with codon 12, 13 or 61 alterations have been considered oncogenic equivalents; however recent clinical observations suggest functional differences for each RAS mutation. For example, in colorectal cancer, K-RAS mutational status is used as a prognostic indicator of resistance to therapy with EGFR antibodies (e.g. cetuximab) (5–8). Retrospective analyses, however, of ‘all-comer’ trials suggests mutational specificity in this regard: patients harboring K-RAS codon 13 mutations appear to benefit from cetuximab therapy, while those with codon 12 mutations were unresponsive (9–13). Moreover, progression-free survival on targeted therapies may also be codon-specific in non-small cell lung cancer (NSCLC) (14). Here, molecular modeling and reverse phase protein analysis pinpointed differential effector engagement and downstream signaling as potential mediators of mutation-specific therapeutic response (14). Together, these results suggest that distinct, codon-specific properties of RAS mutations have important clinical and biological implications.

Cancers display tissue-specific preferences for mutation of the RAS homologs (Table S1). In melanoma, N-RAS is by far the most frequently mutated RAS isoform and notably, 84% of these mutations localize to codon 61 versus only 7% to glutamine 12 (Table S1). A similar preference for codon 61 mutations is noted in thyroid cancer, but is not observed in other cancer types. Codon 12 and 13 mutations constitute more than 90% of K-RAS mutations observed in human colon, pancreatic, lung and ovarian cancers (Table S1 and (1)). Likewise, glycine 12 is the most common site of N-RAS mutation in acute myeloid leukemia (Table S1). The mechanistic basis for codon 61 selection in melanoma and thyroid cancer is unclear. Some have suggested that cytosine to thymidine transversions caused by ultraviolet (UV) light may explain the preference for certain mutations in melanoma, but the majority of codon 61 mutations do not exhibit a characteristic UV-damage signature (15). Alternatively, it is possible that codon mutation preferences reflect differences in oncogenic signaling.

Comparing the oncogenic potential of various RAS mutants is challenging for several reasons. RAS gene dosage clearly influences downstream signaling, and artifacts of RAS overexpression are well-described. Likewise, endomembrane localization is critical for physiologic RAS signal transduction (16), and may not be adequately recapitulated using exogenous protein expression. In addition, genetic alterations private to a given cell line or tumor sample could obscure distinct functions of individual RAS mutants. To circumvent these issues, we generated a knock-in allele (LSL-N-Ras^Q61R) allowing for the conditional, tissue-specific and somatic expression of N-Ras^Q61R under control of the endogenous promoter. We used this allele in combination with similarly designed knock-in N-Ras^G12D (17) and K-Ras^G12D (18) models to compare the transforming potential of each mutant when expressed at physiological levels in melanocytes.

Results

Generation and characterization of LSL-N-Ras alleles

To compare the ability of N-Ras mutants to promote melanoma formation, we employed three conditional knock-in alleles: LSL-N-Ras^G12D (17), LSL-K-Ras^G12D (18) and LSL-N-Ras^Q61R. Each allele contains a floxed transcriptional stop sequence followed by a single missense mutation in the endogenous Ras gene (G12D or Q61R, respectively; Fig. 1A). The codon 12 LSL-K-Ras and N-Ras alleles have been previously described (17, 18). We generated and confirmed a related LSL-N-Ras^Q61R allele using standard homologous recombination followed by Southern blot, PCR and genomic sequencing (Fig. S1A–D). To minimize strain-specific effects, all alleles were backcrossed more than 7 generations to C57Bl/6J in the presence of a conditional p16^INK4a knockout allele (p16^L; (19)) and a melanocyte-specific, 4-hydroxytamoxifen inducible CRE recombinase (Tyr-CRE-ER^T2; (20)). Cohorts of all three alleles were born at normal Mendelian ratios (data not shown) and showed no defects in development or fertility.

Figure 1. Activation of LSL-N-Ras^Q61R or LSL-N-RAS^G12D slows melanocyte proliferation.

(A) Diagrammatic representation of the LSL-N-RAS^G12D (17) and LSL-N-Ras^Q61R alleles in the presence or absence of CRE recombinase. Green triangles denote lox P sites and unnumbered black boxes represent FRT sites. Blue boxes indicate the location of a puromycin (PURO) resistance cassette. SA-splice acceptor; SD-splice donor; 3×STOP- transcription and translational stop sequence. (B) PCR genotyping of melanocytes treated for 6 days with ethanol vehicle (E; lane 7) or 1.0 µM 4-OHT (lane 7). DNA from mice of the indicated genotypes is included as a control (lanes 3–5). (C) Melanocytes cultured as in ‘B’ were incubated with EdU, a thymidine analog, for 16 hours and then analyzed by flow cytometry. EdU incorporation in vehicle treated cells was set to 100%. Each dot represents a biological replicate with the mean and standard error of the mean indicated by black lines.

Using primary melanocytes derived from syngeneic Tyr-CRE-ER^T2 p16^L/L LSL-N-Ras^G12D/G12D (TpN^12D/12D) or LSL-N-Ras^Q61R/Q61R (TpN^61R/61R) neonates, we verified the functionality of the LSL alleles. Melanocyte purities of >99% were confirmed using Tyrosinase-Related Protein 1 (TRP-1) as a marker for flow cytometry ((21); Fig. S2A). In culture, melanocytes were treated with ethanol vehicle or 4-hydroxytamoxifen (4-OHT) to induce CRE activity. CRE-dependent excision of the transcriptional stop element was verified by PCR (Fig. 1B), and resulted in the production of mutant N-Ras mRNA (Fig. S2B). Activation of either allele did not induce changes in melanocyte morphology or pigmentation (Fig. S2C), but induced a decrease in melanocyte proliferation as measured by EdU incorporation (Fig. 1C). Despite co-deletion of p16^INK4a, physiological expression of either N-Ras mutant caused comparable anti-proliferative effects (Fig. 1C). Additionally, both TpN^12D/12D and TpN^61R/61R melanocytes failed to bypass senescence in the presence of 4-OHT, and the cells invariably ceased proliferating after 2–3 passages in culture (data not shown). Therefore, when expressed under the control of an endogenous promoter, neither N-Ras^G12D nor N-Ras^Q61R was capable of immortalizing p16^INK4a-deficient melanocytes.

N-RAS-driven melanomagenesis is codon-specific

We next sought to examine the phenotypic effects of melanocyte-specific Ras mutations in vivo. Toward that end, we generated contemporaneous colonies of p16^INK4a-deficient K-Ras or N-Ras mutant mice in a common genetic background. We elected to study N-Ras mutations in the homozygous state for two reasons: i) we noted no melanoma formation despite 80 weeks of monitoring in a large cohort of Tyr-CRE-ER^T2 p16^L/L LSL-N-ras^G12D/WT (TpN^12D/WT) animals (Fig. S3A), and ii) deep sequencing of nine N-RAS mutant human melanoma cell lines failed to detect the presence of a wild type allele, demonstrating consistent N-RAS loss of heterozygosity in vivo (22). Prior work from our group has demonstrated that melanocytic expression of K-Ras^G12D efficiently promotes melanoma in vivo in the setting of p16^INK4a inactivation (19, 23). Herein, syngeneic Tyr-CRE-ER^T2 p16^L/L LSL-K-Ras^G12D/WT (TpK^12D/WT), were maintained in a heterozygous state due to the homozygous lethality of K-Ras deletion (24). All mice in these cohorts were treated neonatally with 4-OHT (19) to induce melanocyte-specific expression of the desired Ras mutant and delete p16^INK4a.

Initial examination of the skin, paws and tails of these mice revealed the presence of nevi and hyperpigmented regions on the paws and tails (Fig. S4A). The penetrance and severity of these phenotypes was allele-specific with TpK^12D/WT animals having the most pronounced effect and the N-Ras codon 12 mutation producing the least (TpK^12D/WT > TpN^61R/61R > TpN^12D/12D). To quantify nevi frequency, nevi presence was scored weekly for 10 weeks. As expected, nevi were rarely observed on Tyr-CRE-ER^T2 p16^L/L (Tp) mice, but consistently found on TpK^12D/WT animals (Fig. S4B). Members of both the TpN^12D/12D and TpN^61R/61R cohorts had more nevi than control, Tp mice (p<0.001). However, the frequency with which N-Ras^Q61R triggered nevus formation was significantly higher than that observed in TpN^12D/12D animals (p = 0.03). These data suggest that in p16^INK4a-deficient melanocytes, physiological N-Ras^Q61R expression more efficiently promotes nevus formation than N-Ras^G12D mutation.

These established colonies of syngeneic TpN^12D/12D, TpN^61R/61R and TpK^12D/WT mice were aged and serially assessed for melanoma formation. In accordance with previous data, TpK^12D/WT animals developed tumors with high penetrance ((19); Fig. 2A, Table 1). Tumors were very rare in TpN^12D/12D mice (1 tumor found in 29 mice observed for 80 weeks), whereas TpN^61R/61R mice readily developed melanoma with high penetrance and a median latency of 26.3 weeks (Fig. 2A, B and Table 1). To explain this result, we considered the possibility that the LSL-N-Ras^Q61R and LSL-N-Ras^G12D alleles had different recombination efficiencies. Toward that end, we examined allelic recombination in primary melanocyte cultures and noted that if anything, the codon 61 allele recombined with lower efficiency than the codon 12 allele (Fig. S5A). We also considered the possibility that the recombined LSL-N-Ras^G12D allele was poorly expressed. Sequencing of cDNA from treated melanocytes confirmed 4-OHT-dependent expression of N-Ras^G12D mRNA (Fig. S2B). Lastly, functional validation of the LSL-N-Ras^G12D allele was accomplished by inducing allelic recombination in the hematopoietic lineage using an interferon-inducible Mx1-CRE driver. Expression of N-Ras^G12D in the hematopoietic compartment efficiently induced a myeloproliferative syndrome in accord with prior findings ((25) and data not shown). These data establish that when expressed at physiological levels in melanocytes, N-Ras^Q61R and K-Ras^G12D are inherently more transforming than N-Ras^G12D.

Figure 2. In vivo melanomagenesis is isoform and mutation-specific.

(A) Kaplan-Meier curve of melanoma-free survival. Animals were followed for a total of 80 weeks. Survival of each strain was compared pairwise to that of TpN^61R/61R animals using a log-rank (Mantel-Cox) test. The following significant p-values were determined: Tp and TpN^12D/12D - p<0.0001; TpK^12D/WT- p<0.0027. (B) Representative images of the amelanotic tumors found throughout the skin of TpN^12D/12D, TpN^61R/61R and TpK^12D/WT animals. White arrowheads indicate the tumor location. (C) Hematoxylin and eosin staining of representative tumors of the indicated genotypes. Black bars represent a length of 20µm. White arrowheads indicate areas of melanin deposition.

Table 1.

Comparative summary of melanomagenesis and macrometastasis in LSL-Ras-driven models.

Genotype	Tumors/Treated Mice (%)	Median Tumor Latency (wks)	Macrometastasis
Tp	1/24 (4.1%)	> 80	None
TpN12D/12D	1/29 (3.4%)	> 80	None
TpN61R/61R	14/20 (70%)	26.3	None
TpK12D/WT	16/21 (76%)	36.3	None
TpLN61R/61R	17/20 (85%)	22.1	5/14 (36%)

We performed additional analyses to determine if there were phenotypic differences in the tumors of TpN^12D/12D, TpN^61R/61R and TpK^12D/WT mice. Similar growth rates were observed in tumors from TpN^12D/12D, TpN^61R/61R and TpK^12D/WT mice, albeit only one tumor was found in the TpN^12D/12D cohort (Fig. S5B). Moreover, melanomas from all three groups were histologically similar; containing both spindle-cell and desmoplastic cell types with no overt signs of macrometastatic spread (Fig. 2C, Table 1). In the TpN^61R/61R tumors, we confirmed the presence of the recombined LSL-N-ras^Q61R allele (Fig. S5C) and used quantitative real-time PCR to look for potential gene amplifications. Compared to TpK^12D/WT tumors, in which endogenous N-Ras expression is unaltered, TpN^61R/61R melanomas displayed no significant change in N-Ras mRNA levels, suggesting that the allele was not amplified during tumorigenesis (Fig. S5D). These data show that N-Ras^Q61R, N-Ras^G12D and K-Ras^G12D tumors, once established, are phenotypically similar, and suggests that oncogenic differences between the alleles is most pronounced during tumor initiation.

Lkb1 loss promotes melanoma metastasis in TpN^61R/61R mice

Melanoma mortality is predominantly attributed to metastatic disease, however very few RAS-driven genetically engineered mouse models, including the TpN^61R/61R model, show evidence of macrometastases (Tables 1 and (26)). We recently demonstrated that the loss of Liver Kinase 1 (Lkb1/Stk11) promotes widespread macrometastases in a K-Ras^12D-driven mouse melanoma model (23). LKB1 is mutated in ~10% of malignant human melanomas (15, 27, 28) and functionally inactivated by oncogenic BRAF (22, 29, 30). To test whether Lkb1 loss promotes metastasis in TpN^61R/61R tumors, we crossed these animals to a previously described conditional Lkb1 knockout allele (Lkb1^f; (31)). TpLN^61R/61R mice were treated neonatally or at adulthood with 4-OHT to stimulate melanocyte-specific deletion of both Lkb1 and p16^INK4a and induce N-Ras^61R expression. The onset of tumor formation in TpN^61R/61R neonates was not affected by Lkb1 loss (Fig. 3A; Table 1); however, the propensity for distant metastasis was markedly affected. TpLN^61R/61R animals, treated as adults or neonates, exhibited enhanced nevus formation relative to Lkb1-proficient counterparts (compare Fig. S4A to Fig. 3B). Upon sacrifice due to increasing tumor burden, many of TpLN^61R/61R showed signs of metastasis. We performed thorough autopsies on 14 TpLN^61R/61R mice, noting noting lymph node enlargement and other signs of macroscopic disease (Fig. 3C) in the majority of animals. The presence of visceral metastatic disease to the lung, spleen and/or liver was confirmed in 5 of these animals by histologic examination (36%; Fig. 3D, Table 1). Metastastases appeared similar in morphology and incidence to those observed in a prior K-Ras-driven, Lkb1^−/− melanoma model (23). Flow cytometric analyses of TpLN^61R/61R mice with primary melanomas showed the presence of infiltrating tumor cells expressing the melanocyte marker, Trp1 (Fig. 3E; Note that CD45+ cells were excluded from splenic analyses). Unlike cells from TpN^61R/61R tumors, we were able to culture TpLN^61R/61R melanoma cells in vitro. These cells lines exhibited varied morphologies, but were invariably Trp-1 positive (Fig. S6). Together, these findings establish TpLN^61R/61R as a faithful murine model of metastatic, cutaneous melanoma driven by an endogenous, oncogenic N-RAS allele.

Figure 3. Lkb1 deficiency promotes metastasis in TpN^61R/61R tumors.

(A) Kaplan-Meier curve comparing the melanoma-free survival of TpN^61R/61R and TpLN^61R/61Rmice treated neonatally with 4-OHT. A log-rank (Mantel-Cox) test revealed no significant difference in disease onset in these models (p>0.05). (B) Representative photographs showing severe melanocytic hyperproliferation in the ears, tails and paws of TpLN^61R/61Rmice. (C) Images of macrometastases in the lung, spleen and lymph nodes (LN) of TpLN^61R/61Rmice. (D) Representative hematoxylin and eosin stained TpLN^61R/61Rtissues showing tumor invasion into the liver, spleen and lymph node. (E) Expression of TRP1 in the spleen and liver of TpLN^61R mice with primary melanomas. The aqua line designates tissue from a wild type animal and the red line shows TRP1 expression in a TpLN^61R/61Rmouse with observed macrometastases. For splenic tissues, CD45+ cells were excluded prior to analysis.

N-RAS^Q61R and N-RAS^G12D similarly bind melanogenic effector pathways

The distinct oncogenicity of H-, K- and N-RAS is often attributed to the isoform-specific, preferential engagement of downstream effector pathways (17, 32–35). We postulated that a similar mechanism might also drive codon-specific melanomagenesis. To this end, the interaction of purified K-RAS, N-RAS, N-RAS^G12D and N-RAS^Q61R with PI3K and RAF (i.e. the BRAF RBD) was examined in vitro. Each GTPase was first loaded with a fluorescent GTP analog (mant-GMPPNP). Next, purified PI3K or RAF-RBD was titrated into reactions containing a constant amount of GMPPNP-RAS protein at both 5 mM and 100 µM Mg²⁺ concentrations. Fluorescence quenching, caused by effector binding was monitored to determine the K_d for each protein-protein interaction. In these assays, N-RAS^Q61R bound both PI3K and the RAF-RBD with a lower affinity than either wild-type N-RAS or N-RAS^G12D (Table 2). However, the noted differences were small (<1-fold for PI3K; ~4-fold for the RAF-RBD) and unlikely to translate to an in vivo phenotype. Likewise, isothermal titration calorimetry experiments did not reveal significant differences in the RAF-RBD binding affinities of N-RAS^Q61R, N-RAS^G12D and wild-type N-RAS (Figure S8). These data indicate that distinct engagement of the oncogenic RAS effector pathways frequently targeted in melanoma (i.e. RAF and PI3K), is not the cause of codon-specific melanomagenesis.

Table 2.

Binding affinities of RAS-mant-GMPPNP to PI3Kγ and RAF-RBD.

RAS Mutant	mant-GMPPNP Binding
	KD to bRaf-RBD (µM)	KD to PI3K (µM)
NRASWT	0.07 ± 0.03	2.2 ± 0.4
NRASG12D	0.08 ± 0.04	1.8 ± 0.5
NRASQ61R	0.28 ± 0.04	2.7 ± 1.2
KRASG12D	0.06 ± 0.03	2.5 ± 0.8

Activation of MAPK and ERK is codon-independent in NRAS mutant melanomas

The distinct subcellular localizations of H-, K- and N-RAS are suggested to influence effector availability and contribute to isoform-specific RAS oncogenicity (17, 32–35). To determine whether the availability of specific effector pools in vivo contributes to codon-specific signaling, we examined MAPK and PI3K activation in a variety of human melanoma cell lines harboring a mutation in N-RAS codon 12, 13 or 61 (n=11). These cell lines exhibited variable levels of activated ERK and AKT that did not correlate with genotype (Fig. 4A and B). NRAS^G12/13-mutant cell lines exhibited a moderate increase (<2-fold) in total ERK and AKT expression; however, the significance of this observation is unclear (Fig. 4A and B, right) and this finding would not explain the increased transforming activity of codon 61 mutants. In addition, no difference was observed in the proliferation of N-RAS codon 12/13 and 61-mutant cell lines as measured by EdU incorporation (Fig. S9).

Figure 4. Activation of ERK and AKT in human melanoma cell lines is not codon-specific.

Shown are immunoblots for total and phosphorylated ERK (A), or AKT (B) in established human melanoma cell lines harboring the indicated NRAS mutations. Protein expression levels were quantified using LI-COR ImageStudio software. Each dot represents a single cell line. The mean is indicated by a line, and the standard error is shown as whiskers. * p<0.05

Due to the confounding potential of secondary mutations acquired in human melanoma cell lines, we also examined ERK and AKT activation in primary and immortalized melanocytes derived from the TpN^12D/12D and TpN^61R/61R models. Expression of each N-RAS allele was induced in vitro using CRE recombinase and verified by genomic PCR (Fig. S810A and B, top). Immortalized TpN^61R/61R melanocytes were particularly resistant to sustained allelic recombination (Fig. S10B, top). Therefore, we analyzed these cells at an early time point following CRE induction (2 days). In both cases, we found that phospho-ERK and AKT levels were variable and codon-independent (Fig. S10A and B, bottom). These data, along with observations in human melanoma cell lines, demonstrate that melanomageneic and non-melanomageneic N-RAS mutants similarly activate the MAPK and PI3K pathways.

NRAS^Q61R exhibits distinct biochemical properties

Surprised by the finding that NRAS codon 12, 13 and 61 mutants appear equally to engage the oncogenic PI3K and MAPK effector pathways, we further investigated the biochemical properties that differentiate these two mutants. We observed during mant-GMPPNP-based binding assays, that the nucleotide exchange rate of NRAS^Q61R was significantly retarded compared to WT or NRAS^G12D (data not shown). To follow up on this observation, we measured the intrinsic and SOS-mediated nucleotide disassociation rates of purified NRAS, NRAS^G12D and NRAS^Q61R. To monitor exchange, each NRAS variant was pre-loaded with either mant-GMPPNP (mimicking the GTP bound state) or mant-GDP and then incubated with 1000-fold excess unlabeled nucleotide. The resulting change in fluorescence over time was used to monitor exchange rates. NRAS, NRAS^G12D and NRAS^Q61R showed a similar ability to exchange GDP (Table 3). However, the GMPPNP exchange rate of NRAS^Q61R was significantly slower than NRAS and NRAS^G12D (Table 3). This distinction became more pronounced when SOS was added to each reaction (Table 3). Specifically, SOS was unable to stimulate either GDP or GMPPNP exchange in the codon 61 variant (Table 3). These results indicate that NRAS^Q61R possesses a clear affinity for GTP that cannot be overcome by GEF interaction at the concentrations used in our assays.

Table 3.

Nucleotide binding affinities and intrinsic GTP hydrolysis rates of relevant NRAS isoforms.

RAS Mutant	GDP Off	GDP off + 1:2 SOScat	GMPPNP Off	GMPPNP Off + 1:2 SOScat	Intrinsic Hydrolysis
	Rate (s−1 × 10−4)	Rate (s−1 × 10−4)	Rate (s−1 × 10−4)	Rate (s−1 × 10−4)	Rate (s−1 × 10−4)
NRASWT	1.5 ± 0.2	8.0 ± 0.7	1.7 ± 0.4	9.8 ± 0.9	4.6 ± 0.4
NRASG12D	1.3 ± 0.3	3.3 ± 0.4	1.6 ± 0.5	3.9 ± 0.6	2.3 ± 0.5
NRASQ61R	1.1 ± 0.2	1.3 ± 0.3	0.4 ± 0.2	0.5 ± 0.2	0.002 ± 0.03

These data, along with previous observations in other RAS isoforms (36), suggest a decreased intrinsic GTPase activity of NRAS^Q61R. Indeed, when directly assessed, the intrinsic GTP hydrolysis rate of NRAS^Q61R was much slower than NRAS^G12D or NRAS (1,150 and 2,300 times slower, respectively; See Table 3). These results, taken together, suggest that NRAS^Q61R has higher affinity for GTP relative to NRAS and NRAS^G12D. Therefore, we examined the relative stability of NRAS, NRAS^G12D and NRAS^Q61R using thermal unfolding measurements for both GDP- and GMPPCP-bound proteins. NRAS^Q61R remained stable until reaching 80 ± 5°C, whereas the wild type and NRAS^G12D proteins were destabilized at much lower temperatures (67 ± 3°C and 70 ± 3°C, respectfully; See Table S2). The thermostability of NRAS^Q61R was lower in the GDP-bound versus GMPPCP-bound state (74 ± 4°C versus 80 ± 5°C, See Table S2), suggesting that NRAS^Q61R adopts a conformation which stabilizes nucleotide binding, especially in the GTP-bound state. Together, our work reveals that NRAS^Q61R exhibits distinct nucleotide binding capacity, stability and GTPase resistance likely responsible for its exceptional melanomagenic properties.

Discussion

Our data explain a long-standing mystery in the field, demonstrating through biochemical and genetic analyses that the predominance of codon 61 mutants in human melanoma can be attributed to their distinct oncogenic properties. In our novel suite of mouse models, endogenous levels of NRas^Q61R, but not NRas^G12D were able to efficiently drive in vivo melanomagenesis (Fig. 2). Although prior work has linked isoform-specific RAS oncogenicity to differential effector binding (17, 32–35), we found that NRAS^Q61R and NRAS^G12D similarly engaged the PI3K and MAPK pathways (Table 2, Fig. 4, Fig. S8, Fig. S10). Our data suggest an alternative model for codon-specific oncogenicity, demonstrating that the melanomagenic NRAS^Q61R mutant possesses distinct biochemical properties not found in NRAS^G12D (Table 3).

Prior to this study, it was unclear whether the prevalence of NRAS codon 61 mutations in melanoma reflected a preferential pattern of mutagenesis or codon-specific differences in the biological function of each allele. We now show that NRAS^G12D does not efficiently drive cutaneous melanomagenesis (Fig. 2), and therefore, codon-specific RAS biology influences tumorigenic potential. Tissue- and temporal-specificity are also likely to affect oncogenic RAS activity. For instance, activation of the LSL-NRAS^G12D allele in hematopoietic stem cells and early melanocyte progenitors drives leukemogenesis and melanoma of the central nervous system ((25, 37, 38), data not shown). Moreover, it is possible that our experimental system, which relies upon p16^INK4a loss to facilitate melanomagenesis, is biased to favor N-Ras codon 61 mutations. Patient studies have suggested exquisite cooperation between N-RAS codon 61 mutations and p16^INK4a inactivation (39, 40), and N-Ras^Q61K transgenic mice appear to be ‘addicted’ to the activity of CDK4/6, which is enhanced by p16^INK4a loss (41). Therefore, perhaps other cooperating oncogenic events commonly found in melanoma would more efficiently synergize with N-Ras^G12D in tumor formation. Nonetheless, p16^INK4a loss is found in the majority of human melanomas, occurring through a variety of mechanisms (i.e. epigenetic silencing, mutation and deletion). Given the significance of this event in the clinical setting, we believe the choice of the p16^INK4a-deficient genetic context is highly relevant to the human disease.

Widely efficacious and durable treatment options are not available for the 17–30% of patients with NRAS mutant, metastatic melanomas (1, 15, 42). With mounting data to suggest that genetically engineered mouse models of cancer more faithfully report therapeutic efficacy (43–45), a number of RAS-mutant melanoma models have been developed (See Table S3). Most of these models employ transgenic technologies that can alter the expression, subcellular compartmentalization, intracellular signaling and transforming potential of RAS (16). To this end, the activation of endogenous RAS oncogenes triggers minimal ERK and AKT activation in our system and others (Fig. S10 and (17, 18, 25)); yet, transgenic RAS alleles robustly stimulate these pathways (46, 47). Other murine melanoma models employ oncogenic RAS isoforms rarely observed in human melanomas (i.e. H-RAS and K-RAS mutants). As each RAS isoform can initiate a unique set of downstream signals, it is unclear to what degree these models faithfully recapitulate the oncogenic mechanisms found in human melanomas. For these reasons, we believe that the TpN^61R and TpLN^61R models will be extremely valuable for pre-clinical drug testing, especially given the metastatic nature of the Lkb1-deficient tumors.

NRAS^Q61R exhibits unique biochemical properties

In contrast to prior studies showing that preferential effector usage drives isoform-specific RAS oncogenesis (17, 32–35), we found that distinct biochemical properties inherent to each RAS mutant likely drives codon-specific melanomagenesis (Tables 2 and S2). Nearly 3 decades ago, Der and colleagues reported that HRAS codon 61 mutants, regardless of their transforming potential, exhibited a ~10-fold decrease in intrinsic GTP hydrolysis (48). Later, Donovan, Shannon and Bollag used predicted RAS-GTP levels to form the hypothesis that defects in intrinsic GTPase activity and nucleotide exchange dictate the transforming potential of individual RAS mutants (49). Our experimental results are consistent with this model, demonstrating that the melanomagenic NRAS^Q61R variant exhibits high affinity GTP binding, increased stability and reduced intrinsic GTPase activity when compared to NRAS^G12D (Tables 2 and S2).

These results suggest that NRAS^Q61R may more efficiently activate downstream effectors in vivo. However, the activation of ERK and AKT were similar in both primary melanocytes and human melanoma cultures harboring an NRAS codon 12 or 61 mutation (Figs. 4 and S10). Moreover, even through unbiased probes of the kinome using PhosphoScan technology (Cell Signaling), we were unable to detect consistent differences between NRas^G12D and NRas^Q61R signaling in primary melanocytes (data not shown). Due to feedback mechanisms within many pathways downstream of RAS, changes in signaling flux may not be readily apparent. Alternatively, enhanced cooperation with secondary oncogenic events during the initial stages of tumor development may be responsible for the initial selection of NRAS^Q61R mutants in melanoma. In line with this observation, RAF activation is a common event in both thyroid carcinomas and melanomas, the two major tumor types wherein NRAS codon 61 mutations predominate (1, 2). Mounting evidence supports the idea that the intensity of oncogenic RAS signaling can influence transformation potential (38). In addition, RAF binding in conjunction with unidentified allosteric regulators is speculated to further reduce the intrinsic GTPase activity of HRAS codon 61 mutants (36). These findings, along with our work, suggest that NRAS codon 61 mutants exhibit unique biochemical properties which promote the transformation of RAF-responsive tissues. Parsing the mechanism of this specific dependency will require the structural analyses of multiple NRAS mutants as well as comprehensive functional screens using endogenous RAS expression systems and relevant cell types. However, our data suggest that this work would be extremely valuable, identifying codon-specific tumor vulnerabilities for therapeutic targeting.

Methods

Murine Alleles and Husbandry

Animal work was conducted in accordance with protocols approved by the Institutional Care and Use Committee for animal research at the University of North Carolina and the Ohio State University. The Tyr-CRE-ER^T2, p16^L, Lkb1^f, LSL-K-ras^12D and LSL-N-Ras^12D alleles have been previously described (17–20, 31). All animals in this study were backcrossed more than 7 generations to C57BL/6. The Lkb1^f allele was initially established on an albino C57BL/6J background and then crossed into the TpN^61R colony. Therefore, some of the mice in this cohort are albino. Analysis of tumor-free survival was conducted using GraphPad Prism software. To determine statistical significance, a Logrank (Mantel-Cox) test was performed for each experimental pairing.

Generation of the LSL-N-Ras^Q61R Allele

Standard homologous recombination procedures were used to insert a conditionally excisable transcriptional stop codon and point mutation into the endogenous N-Ras locus (See Fig. S1A). Founding mice were first crossed to C57Bl/6 FlpE mice (Jackson Labs #005703) to remove the neomycin resistance cassette. Southern blotting was performed on ES cells and founder DNA using standard procedures. PCR primers and cycling conditions used for genotyping LSL-N-Ras^Q61R were as follows: Q61R GENO2- 5'-GCAAGAGGCCCGGCAGTACCTA-3' (0.15 µM); Primer 1 – 5'-AGACGCGGAGACTTGGCGAGC-3' (0.15 µM); Primer 2 – 5'-GCTGGATCGTCAAGGCGCTTTTCC-3' (0.15 µM); Cycling - 95°C 15 min, 35 × [94°C 30s, 62°C 30s, 72°C 45s], 72°C 5 min. The resulting PCR products were 487 (wild type), 371 (LSL-N-ras^Q16R), and 562 (LSL-N-ras^Q16R+ CRE) base pairs in size.

Primary Melanocyte Culture

Skin was isolated from newborn pups and placed dermis side down in 0.25% trypsin, 0.1% EDTA for 3 hours at 37°C and 5% CO₂. Using forceps, the epidermal layer was separated from the dermis. Using surgical scissors, epidermal cells were minced finely in phosphate buffered saline supplemented with 0.02% EDTA. To further dissociate the cell suspension, each sample was subjected to 2 rounds of program A on the GentleMACs dissociator (Miltenyi). The resulting samples were spun down and plated on rat tail collegen-coated dishes in melanocyte growth medium (Ham’s F10, 10 µg/mL insulin, 0.5 ng/mL bovine serum albumin, 5% fetal bovine serum, 1 µM ethanolamine, 1 µM phophoethanolamine, 10 nM sodium selenite, 20 nM TPA, 50 pM cholera toxin, 1× penicillin/streptomycin, 100 nM melanocyte stimulating hormone, and 0.05 mM di-butyril cyclicAMP). Media was changed every other day.

Induction of CRE Recombinase

For neonatal induction, pups were painted dorsally (postnatal days 2–4) with 25mg/mL 4-hydroxytamoxifen (4-OHT) dissolved in DMSO. Adult induction of CRE was performed as previously described (50). For cell culture studies in primary melanocytes, 4-OHT was dissolved in ethanol and added to the growth media. Cells were treated for 6 consecutive days replacing the media and 4-OHT every other day. In the immortalized TpN^61R cultures, which were resistant to recombination, an adenoviral CRE recombinase was employed (Ad5-MCV-Cre-GFP, Baylor College of Medicine Vector Development Lab). Prior to proliferation or morphological assessment, melanocyte cultures were returned to untreated media for 3 days. Allelic recombination in tumors and primary cultures was confirmed by genomic PCR using the genotyping primer sets and conditions described for each allele.

Scoring of Murine Nevi

Nevi on a 4 cm² dorsal area were counted by staff, blinded of the animal’s genotype, every week for 10 weeks. So as not to interfere with melanomagenesis, we chose not to depilate or shave these animals during the study. The presence of a single nevus at any time point was scored as a ‘1’. If no nevus was observed a ‘0’ was entered for that time point. The percentage of time nevus positive was calculated as: (sum of all 10 measurements/10)* 100. Each dot represents 1 animal with the mean indicated by a line. Comparisons between sample pairs were conducted using a student’s t-test.

Culture of Murine and Human Melanoma Cell Lines

The NZM24 and NZM63 cell lines were developed in the laboratory of Dr. Bruce C. Baguley (51) and obtained from the Cell Line Collection maintained by the Auckland Cancer Society Research Centre, Faculty of Medical and Health Sciences, The University of Auckland. The MaMel27II cell line was generated by Drs. G. Finlay, C. Posch and S. Ortiz (Mount Zion Cancer Research Center, San Francisco; (52)). WM3670, WM3629 and WM1366 cells were created in the Heryln lab and obtained from the Wistar Institute (53). VMM39 cells were the kind gift of C. Slingluff (U. Virginia)(54). The SK-Mel-119, SK-Mel-103 and SK-Mel-147 cell lines were generously provided by Dr. A. Houghton (Memorial Sloan-Kettering)(55). The Mel224 cell line was produced and supplied by J. Hansson (Karolinska Institutet). Tumor-derived murine cell lines form the TpLN^61R/61R and TpK^12D/WT models were generated, genotyped and maintained as previously described (23).

Authentication of Cell Lines

All cell lines were obtained from their original sources or authorized distributors and maintained as previously described (51–56). Cell lines from the Wistar institute (WM3670, WM3629 and WM1366) are routinely validated by short tandem repeat (STR) profiling using the AmpFISTR Identifiler PCR Amplification kit (Life Technologies) and were used for the experiments shown within 3 months of receipt. For primary cell lines where original STR data is not currently available, N-RAS mutations were validated by PCR followed by Sanger sequencing.

Flow Cytometry

Cell cycle analysis of primary melanocytes: After 10 days in culture, 4.4×10⁵ primary melanocytes were seeded onto collagen coated 60mm dishes. A day later, the media was changed and cells were allowed to recover overnight. EdU was added to each culture at a concentration of 2.5ug/mL for a period of 16 hours. Cells were harvested and processed as described in the Click-iT EdU Flow Cytometry Assay Kit (Invitrogen) using saponin for permeabilization. Cell cycle analysis of human melanoma cell lines: Human melanoma cell lines grown to 50–70% confluence were treated with 10 µM EdU for a period of 6 hours. Cells were harvested, processed, stained for EdU incorporation and analyzed on a Flowsight cytometer (Amnis). Trp1 Staining: To validate that our cultures contained a pure population of melanocytes and verify the identity of TpLN^61R metastases, PEP1 antibody (αTrp1; (21)) was added at a 1:100 dilution to saponin permeabilized cells followed by incubation with an AlexaFluor488 conjugated anti-rabbit secondary (Molecular Probes, 1:1000 dilution). In the analysis of splenic tissues (Fig. 3E), co-staining with CD45 was first used to gate out any CD45+ cells.

In vitro protein purification

A vector encoding human N-RAS (1-172) was acquired from Addgene (vector #25256, NRAS-A). The G12D and Q61R mutations were each introduced into this vector using standard quick change mutagenesis. All proteins were expressed and purified from Rosetta 2 BL21(DE3) cells (Novagen, Madison, WI, USA). Briefly, cells were grown at 37°C in a shaking culture of Terrific Broth media supplemented with kanamycin. Once the cells reached an OD600 between 0.6–1.0, the culture was chilled in an ice bath (to ~20°C) and 0.1mM IPTG added. The cultures were then shaken for an additional 12–15 hours at 18°C. Cells were harvested by centrifugation and pellets stored at −80°C. N-RAS proteins were purified on Ni Sepharose6 columns following the manufacturer’s protocol (GE Life Sciences, Piscataway, NJ, USA). To further purify the proteins from contaminants, size-exclusion chromatography was performed (Superdex 75 10/300 GL; GE Life Sciences, Piscataway, NJ, USA). Purity exceeding 95% was confirmed by SDS-PAGE analysis.

The minimum catalytic domain of the human protein Son of Sevenless (SOS^cat) was expressed in the pQlinkH vector (Addgene) and purified as previously described (57). The E.coli codon-optimized RAS-binding domain of BRAF (amino acids 149–232) containing an N-terminal purification tag (MGHHHHHHSSGVDLGTENLYFQS) was synthesized by Genewiz (South Plainfield, NJ), sub-cloned into pET28a and expressed in BL21 (DE3) cells. The resulting BRAF-RBD was purified on Ni Sepharose6, cleaved overnight with TEV protease and then subjected to subtractive Ni column purification. The tag-less BRAF-RBD was further purified using size exclusion chromatography and verified to be > 95% pure by SDS-PAGE analysis. Purified PI3Kγ (amino acids 144-1102) was kindly provided by Genentech.

Loading of N-RAS with nucleotide derivatives

To observe nucleotide dissociation activity in the GTP-bound state, N-RAS was loaded with 2’−/−3’-O-(N’methylanthraniloyl) -guanosine-5'-[(β,γ)-imido]triphosphate (^mantGMPPNP). N-RAS was exchanged out of excess MgCl₂ into a buffer containing 20 mM HEPES, 50 mM NaCl, 125 mM (NH₄)₂SO₄, 1 mM MgCl₂ at pH 7.4. The concentration of N-RAS was adjusted to 100 µM and a 5-molar excess of ^mantGMPPNP was added. Alkaline phosphatase conjugated to sepharose beads was added to 1:10 volume and EDTA was added to 1 mM to increase the nucleotide exchange rate. The protein mixture was incubated at 4°C until all unlabeled nucleotide was converted to guanosine. The alkaline phosphatase beads were removed by centrifugation and 10 mM MgCl₂ was added to induce nucleotide binding. The concentration of ^mantGMPPNP-bound RAS was determined by measuring the absorbance of the mant fluorophore (ε₃₅₀ = 5700 M⁻¹cm⁻¹). For assays requiring unlabeled N-RAS in the GTP-bound state, N-RAS·GMPPCP was prepared in an identical manner.

For assays observing N-RAS nucleotide dissociation activity in the GDP-bound state, the protein was loaded with ^mantGDP (2’−/−3’-O-(N’methylanthraniloyl) –guanosine diphosphate) as previously described (58). N-RAS was exchanged into a buffer containing 20 mM HEPES, 125 mM (NH₄)₂SO₄, 100 µM EDTA, 200 µM DTPA at pH 8.0 to remove excess MgCl₂ and nucleotide. A 5-molar excess of ^mantGDP was added to the protein and the mixture was incubated for 30 min at 37°C to increase the nucleotide exchange rate. MgCl₂ was then added to 20 mM and the protein was placed at 4°C for 2 hours to induce binding, after which N-RAS was exchanged into a buffer containing 20 mM HEPES, 50 mM NaCl, 5 mM MgCl₂, 100 µM DTPA, pH 7.4 for storage.

Quantitative Real-time PCR

RNA from melanocytes or melanomas was isolated using the Qiagen All-Prep Kit. Equal amounts of RNA were subjected to reverse transcription using a PrimeScript First Strand cDNA Synthesis kit (Takara). Real-time PCR was run in triplicate and a standard curve of known plasmid concentration was included on every plate (ABI Taqman primer set #Mm00477878). Molecules of N-RAS/µg total RNA were calculated assuming a cDNA conversion efficiency of 100%.

Determination of N-RAS binding to BRAF-RBD and PI3Kγ

For quantitative binding to BRAF-RBD and PI3Kγ (amino acids 144-1102), N-RAS·^mantGMPPNP was incubated with different concentrations of the desired effectors in a buffer containing 50 mM HEPES, 50 mM NaCl, 5 mM MgCl₂ at pH 7.4. Nucleotide dissociation was initiated by the addition of 1000-molar excess of unlabeled nucleotide and the rate of dissociation was determined by monitoring the change in fluorescence of the ^mantGMPPNP loaded protein (excitation and emission wavelengths of 365 and 435 nm, respectively) using a Spectramax M2 plate reader (59). Each nucleotide dissociation curve was fit to a one-phase single exponential to determine k_obs. The dissociation rates were plotted against the effector concentrations and fit to the equation:

$$Y=A-(A-B)*\frac{{(([P1]+[P2]+Kd)-{((P1)+(P2)Kd)}^2-4*[P1]*[P2])}^{0.5}}{2*[P1]}$$

Y: Observed rate, A: Maximum rate, B: Minimum rate, P1: Concentration of protein 1 (cell), P2: Concentration of protein 2 (ligand/effector), which was used to determine the N-RAS·^mantGMPPNP : effector binding constant (K_D) (60).

Nucleotide exchange in the absence and presence of SOS^cat

Nucleotide dissociation rates were measured using a PerkinElmer LS50B fluorimeter as previously reported (58). N-RAS was loaded with mant-labeled nucleotide and added to a final concentration of 1 µM in 1 mL of exchange buffer (20 mM HEPES, 50 mM NaCl, 5 mM MgCl₂, 100 µM DTPA, pH 7.4). A 1000-fold excess of unlabeled nucleotide was added to initiate the dissociation reaction and the rate of nucleotide dissociation was measured by monitoring the change in fluorescence (λ_ex = 365 nm, λ_em = 435 nm). For reactions that showed dissociation rates too slow to reach completion within the time frame of the assay, 25 mM EDTA was added to induce complete nucleotide dissociation. For GEF-facilitated nucleotide exchange, the minimum catalytic domain of son of sevenless (SOS^cat) was added to a 1:2 RAS-to-SOS^cat ratio (chosen to limit RAS binding to the allosteric site in SOS^cat). The fluorescence data was normalized and fit to one-phase exponentials to determine rates.

Determination of intrinsic GTP hydrolysis activity of N-RAS

To measure GTP hydrolysis rates, the phosphate sensor FLIPPi (Addgene, Cambridge, MA, USA) was used for the reaction, which binds to free phosphate in solution (61). Due to the nature of the assay, all trace phosphate was removed from buffers prior to use to minimize background fluorescence. N-RAS was exchanged into a buffer containing 20 mM HEPES, 20 mM (NH₄)₂SO₄, 1 mM EDTA, 1 mM inosine, pH 8.0. N-RAS was incubated with GTP and passed through a PD-10 column to remove trace Mg²⁺ (62). N-RAS was diluted to 10 µM and combined with 10 µM FLIPPi, and 2 mM MgCl₂ was added to initiate the hydrolysis reaction. A phosphate standard curve was used to convert the raw fluorescence output to a measurement of [GTP] hydrolyzed.

Thermal Melting Analysis by Circular Dichroism (CD) Spectroscopy

A Jasco J-815 CD Spectrometer was used for CD measurements. N-RAS was exchanged into a buffer containing 10 mM KH₂PO₄/K₂HPO₄ at pH 7.45 and diluted to 15 µM. MgCl₂ and guanine nucleotide (GDP or GMPPCP) was added to a final concentration of 500 µM and 80 µM respectively, immediately before analysis. A thermal melt scan from 20° to 90°C was performed to determine the temperature (T_m) at which half of the protein is unfolded.

RAS Mutational Analysis in Human Cancers

Data were downloaded from COSMIC (release v64, March 26, 2013)(1). Cell lines and cultured cells were removed from the analysis to prevent inclusion of events secondary to culture. Remaining events were limited to observations of somatic missense mutations in codons 12, 13 or 61 of HRAS, NRAS or KRAS. Multiple subtypes and histologies were then summed to give information for the displayed tumor types.

Significance.

This work explains the curious predominance in human melanoma of mutations of codon 61 of N-RAS over other oncogenic N-RAS mutations. Using conditional ‘knock-in’ mouse models, we show that physiological expression of N-RAS^Q61R, but not N-RAS^G12D, drives melanoma formation.

Abbreviations List

4-OHT

4-hydroxytamoxifen

AKT

v-akt murine thymoma viral oncogene

BRAF

v-raf murine sarcoma viral oncogene homolog B

CDK4/6

cyclin dependent kinase 4/6

DMSO

dimethyl sulfoxide

DTPA

diethylene triamine pentaacetic acid

EDTA

Ethylenediaminetetraacetic acid

EdU

5-ethynyl-2’-deoxyuridine

EGFR

epidermal growth factor receptor

ERK

extracellular signal-regulated kinases

FLIPPi

fluorescence resonance energy transfer (FRET) sensors for phosphate (P(i))

GAP

GTPase-activating protein

GEF

guanine nucleotide exchange factor

GMPPNP

guanosine-5'-[(β,γ)-imido]triphosphate

GMPPCP

guanosine-5'-[(β,γ)-methyleno]triphosphate

HEPES

4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid

H-RAS

Harvey rat sarcoma viral oncogene homolog

K-RAS

Kirsten-rat sarcoma viral oncogene homolog

LKB1/STK11

liver kinase B1/serine threonine kinase 11

MAPK

mitogen-activated protein kinase

N-RAS

neuroblastoma RAS viral oncogene homolog

PI3K

Phosphatidylinositol-4,5-bisphosphate 3-kinase

RAF

rapidly accelerated fibrosarcoma

SOS

Son of Sevenless

TEV

tobacco etch virus protease cleavage site

TPA

12-O-Tetradecanoylphorbol-13-Acetate

TRP-1

Tyrosinase-Related Protein 1