RICTOR Amplification Promotes NSCLC Cell Proliferation through Formation and Activation of mTORC2 at the Expense of mTORC1

Lung cancer is the leading cause of cancer-related deaths worldwide, despite significant advances in therapeutic options for these patients. Lung cancer is divided into two subtypes, non–small cell lung cancer (NSCLC) and small cell lung cancer (SCLC), which account for approximately 85% and 15% of cases, respectively. NSCLC is a disease typically characterized by its genomic alterations, although nearly half of all patients lack a known targetable alteration (1). Recent advances in tumor sequencing technologies have identified amplification of mTORC2-specific component RICTOR as potential targetable alteration in several types of cancer including NSCLC, SCLC, breast cancer, and gastric cancer (2–5).

The mechanistic target of rapamycin (mTOR) is a serine/threonine kinase that acts in two distinct complexes, rapamycin-sensitive mTORC1 and rapamycin-insensitive mTORC2. Both complexes share the mTOR kinase and scaffolding protein mLST8, whereas mTORC1 contains scaffolding protein Raptor and mTORC2 contains Raptor-analogous scaffolding protein Rictor and regulatory component Sin1 (6, 7). Growth factors, amino acids, and cellular energy activate mTORC1, which has well-characterized functions including regulation of cell growth, protein translation, metabolism, and autophagy (6, 7). Although less well understood, mTORC2 is activated by growth factors and the PI3K pathway, primarily through binding of PIP3 in the plasma membrane via the PH domain of cofactor Sin1 (8). Once localized to the membrane, mTORC2 can phosphorylate (S473) and activate its downstream effector AKT (9), also situated at the membrane. In addition to AKT, mTORC2 targets include PKCα and SGK (10, 11). mTORC2 and its downstream effectors act together to regulate cell proliferation, metabolism, and cytoskeletal organization (6, 7).

Aberrant signaling of both mTOR complexes has been implicated in many types of cancers, although most studies have focused on mTORC1. mTORC2′s role in cancer has also been reported; for example, mTORC2 is required for tumor formation in PTEN-null prostate cancer (12, 13) and drives tumor progression and resistance in HER2-positive breast cancer (3, 14). Additionally, the recent identification of RICTOR amplification as a potential actionable target in several cancer types suggests an additional subtype of cancer may rely on mTORC2 signaling for tumorigenesis (2–5). However, the mechanism by which amplification of a single scaffolding component within mTORC2 drives its oncogenic function remains unknown.

In this report, we investigate the roles of RICTOR amplification as a driver of mTORC2 formation and promoter of NSCLC tumor growth. Using the Cas9 Synergistic Activation Mediator (SAM) system (15) to increase expression of Rictor or the CRISPR-Cas9 system to knock out Rictor, we show that alterations in Rictor lead to corresponding changes in not only mTORC2 formation, but also mTORC1. Further, overexpression of Rictor in NSCLC cells leads to increased growth of 3D cultures and in vivo tumor xenografts. We also found that RICTOR-amplified NSCLC is sensitive to loss of mLST8, an mTOR cofactor specifically required for mTORC2 function (13), both in vitro and in vivo. Collectively, these data show that Rictor promotes NSCLC through mTORC2 formation and could potentially be targeted with an mTORC2-specific inhibitor.

H1975, H358, H23, and H1703 cells were obtained from ATCC and maintained in RPMI-1640 media containing 10% fetal bovine serum (FBS) and 1% penicillin/streptomycin. BEAS-2B and H460 cells were obtained from the Vanderbilt-Ingram Cancer Center Core Facility and maintained in the same conditions. 293T cells were obtained from ATCC and maintained in DMEM containing 10% FBS. All cells were cultured in a humidified incubator with 5% CO₂ at 37°C. Cell lines were used between passages 1 and 50 after thaw. Cell lines from ATCC were authenticated using short tandem repeat profiling. Mycoplasma testing was performed every 6 months, most recently in November 2019, using the PlasmoTest kit (Invitrogen).

CRISPR-Cas9 backbone vectors were obtained from Addgene, and guide RNA sequences were cloned into the vector according to depositor's instructions. Plasmids and guide RNA primers are listed in Supplementary Table S1.

CRISPR SAM, knockout, and inducible Cas9 cell lines were established using the lentiviral delivery system. Briefly, lentiviruses were packaged in HEK293T cells by transfecting cells with CRISPR or expression plasmids together with psPAX2 (lentiviral packaging) and pCMV-VSV-G (envelope) plasmids at a 1:1:1 molar ratio using the Lipofectamine 2000 Reagent. Media were changed after 16 hours of transfection, and virus was collected after 24 to 48 hours. Indicated cells were transduced with 1:1 virus and complete growth media with polybrene (8 μg/mL) for 24 hours and selected with puromycin (1–2 μg/mL), blasticidin (10 μg/mL), or hygromycin (40–50 μg/mL) for at least 48 hours to establish stable cell lines before being used for experiments.

2.5 × 10⁶ cells suspended in 100 μL of Matrigel and PBS (1:1) were injected into the hind flanks of 6-week-old athymic nude (Foxn1^nu; Envigo) or Rag1^−/− C57BL/6J mice. For xenograft experiments using an inducible Cas9, doxycycline feed (Envigo #TD.00426) was given for 10 days. Tumor measurements were started 10 days after injection and measured for 30 to 60 days after injection. Tumors were measured every 2 to 3 days with digital calipers, and tumor volume was calculated according to the formula (V = 4/3π(l/2)(h/2)(w/2)). Data are presented as mean ± SEM. Two-way ANOVA with Bonferroni correction was used for statistical analysis. Experiments with mice were pre-approved by the Vanderbilt Institutional Animal Care and Use Committee and followed all state and federal rules and regulations.

Cell lysates for Western blotting only were collected in RIPA buffer. Tumor lysates were collected in Triton X lysis buffer (1% Triton X-100, 0.5 mmol/L EDTA, 50 mmol/L Tris-Cl). All lysis buffers were supplemented with protease inhibitors and phosphatase inhibitors (Complete Mini and PhosStop inhibitor cocktail, Roche). Protein concentration was determined by Pierce BCA Protein Assay kit, and equal amounts of protein extracts were mixed with 4× Laemmli sample buffer and separated by electrophoresis on an SDS-PAGE gel, and then transferred onto nitrocellulose membranes. Membranes were blocked with 5% milk in TBST buffer and incubated with corresponding primary antibodies and IRdye-conjugated or HRP-conjugated secondary antibodies. Immunoreactivity was detected using the Odyssey scanner (Li-cor Biosciences) or enhanced chemiluminescence. To perform immunoprecipitation, equal amounts of input lysates (500 μg) collected in CHAPS buffer (40 mmol/L Tris, pH 7.5, 120 mmol/L NaCl, 1 mmol/L EDTA, 0.3% CHAPS) were incubated with the primary antibodies (1–2 μg) for 2 hours to overnight at 4°C. Protein G Dynabeads (Thermo Fisher Scientific #10-003-D) were added and lysates were incubated for 1 hour and washed four times with CHAPS lysis buffer. Immunoprecipitate and whole-cell lysates were then subjected to Western blot analysis. All antibodies are listed in Supplementary Table S2. Quantification of Western blots was performed using ImageJ software.

Cells cultured on glass coverslips were fixed with 4% PFA, permeabilized with 1% Triton X-100 in PBS, and stained with the Duolink (Sigma-Aldrich #92102) proximity ligation assay according to the manufacturer's protocol using antibodies listed in Supplementary Table S2 and counterstained and mounted with DAPI (Thermo Scientific #P36941). Proximity ligation assay (PLA) puncta and DAPI-stained nuclei were enumerated using ImageJ software.

Cell growth was measured by MTT, colony formation, and BrdUrd assays. For MTT assays, 2,000 cells were plated into each well of a 96-well plate in 100 μL of complete growth medium. Cell viability was measured by incubating cells with 20 μL of 5 μg/mL Tetrazolium salt 3-(4,5-dimethylthiazol-2-yL)-2,5-diphenyltetrazolium bromide (MTT, Sigma-Aldrich) and quantified by reading absorbance at 590 nm after resuspending in MTT solvent. For colony formation assay, 500 cells were plated in complete growth medium in each well of a 12-well plate. Cells were grown for 10 to 14 days, and the media were changed every 3 days. Colonies were stained with 0.5% crystal violet in methanol and colony area was quantified. For the BrdUrd assay, cells were grown on glass coverslips for 24 hours, then incubated with BrdUrd (10 ng/mL) for 30 minutes. Cells were fixed in 4% paraformaldehyde for 15 minutes and then permeabilized with 1% Triton X-100 in PBS for 5 minutes. DNA was denatured by incubation with 2N HCl for 30 minutes at 37°C, followed by rinsing in PBS. Cells were blocked for 1 hour using 2.5% goat serum in PBS and then probed with Alexa 647-conjugated anti-BrdUrd antibody (1:50, Invitrogen) overnight at 4°C. Coverslips were mounted onto slides using ProLong Gold antifade reagent with DAPI (Thermo Scientific #P36941). 40× images were quantified by counting BrdUrd-positive nuclei/total nuclei using ImageJ software.

60 μL of Matrigel (Corning #354230) was added to each well of an 8-well chamber slide (Thermo Scientific) and incubated at 37°C to solidify. Cells (5,000) were suspended in 400 μL of complete media with 2.5% Matrigel and 5 ng/mL of EGF then added to the chamber slides. Cultures were monitored for 20 days, and media were replaced every 4 days. Area of cultures was quantified using ImageJ software.

Xenograft tumor sections were paraffin-embedded and sectioned. Rehydrated paraffin sections were subjected to antigen retrieval (Retrievagen A, BD Pharmingen), and endogenous peroxidases were blocked by 3% H₂O₂ for 30 minutes. Sections were blocked in 2.5% goat serum in PBS and stained with primary antibody and biotinylated secondary antibody, followed by avidin–peroxidase reagent and DAB. Antibodies are listed in Supplementary Table S2. Sections were then counterstained with hematoxylin and mounted with Cytoseal-XYL (Thermo Scientific Richard-Allan Scientific).

Alteration frequency and copy-number segmentation plots and analysis were generated using the online platform cbioportal.org. Proportion of copy-number gain/loss was analyzed using the GenVisR package (16).

Amplification of RICTOR has been identified in several different types of cancers, including breast cancer, gastric cancer, SCLC, and NSCLC (2–5). Analysis of the TCGA PanCancer Atlas identified RICTOR amplification in 12% of total NSCLC cases, and 15% of squamous cell carcinoma and 11% of adenocarcinoma cases, with amplification being the most frequent alteration of RICTOR (Fig. 1A). RICTOR is located on the 5p chromosome, a site of frequent copy-number gain in NSCLC (17). In addition to the overall 5p gain, copy-number segment and GISTIC analysis reveals a focal amplification of the chromosomal locus surrounding the RICTOR gene (Supplementary Fig. S1A and S1B). Interestingly, no other components of the mTOR complexes were coaltered with RICTOR amplification (Fig. 1B). Further analysis of adenocarcinoma cases, which accounts for the majority of NSCLC cases, showed a positive correlation between copy-number and mRNA expression of RICTOR, suggesting amplification does indeed lead to an increase in Rictor expression in NSCLC cases (Fig. 1C).

In order to examine how amplification of a single mTOR complex component might drive tumorigenesis, we used the Cas9 Synergistic Activation Mediator system (SAM) to upregulate expression of RICTOR (15). This system utilizes an enzymatically inactive Cas9 (dCas9) fused to VP64, coexpressed with MS2-P65-HSF1 activation helper proteins. A small guide RNA (sgRNA) with two MS2 aptamers targeting the promoter region of the gene of interest is also expressed, such that all components of the system colocalize and activate transcription of the targeted gene. Multiple sgRNAs targeting the promoter region of RICTOR (Fig. 1D) were tested in a panel of nontumorigenic, immortalized lung epithelial (BEAS-2B) and NSCLC (H1975, H358, H460, and H2030) cell lines, all of which have lower copy-number values compared with the H23 cell line (Fig. 1E), an NSCLC cell with a verified amplification of RICTOR (2). Western blot analysis of these cell lines at baseline demonstrated Rictor protein levels that largely correlate with copy-number values (Fig. 1F). In all lines, SAM Rictor cells exhibited increased mRNA (Fig. 1G) and protein (Fig. 1H) levels of Rictor compared with SAM empty vector (EV) control cells. Across all cell lines analyzed, RICTOR mRNA was upregulated approximately 2- to 4-fold with the CRISPR SAM system (Fig. 1G). Analysis of mRNA expression in RICTOR-amplified samples compared with diploid samples of TCGA lung adenocarcinoma samples (Fig. 1C) showed an approximately ∼2.25-fold increase in Rictor expression. This correlates well with the expression changes in Rictor when using the CRISPR SAM system in Fig. 1G, suggesting that SAM-induced Rictor expression is comparable to endogenous Rictor levels in human lung cancer patients.

We hypothesized that amplification of RICTOR could promote tumorigenesis by driving formation of mTORC2. To test this hypothesis, we used the PLA to quantitate the interactions between mTOR and either Rictor (mTORC2) or Raptor (mTORC1). The same rabbit antibody against mTOR was used in all PLA experiments, whereas mouse anti-Rictor or anti-Raptor were used to differentiate between the two complexes (Supplementary Table S2). Compared with SAM EV cells, SAM Rictor cells exhibited increased fluorescent foci indicating an increase in mTOR–Rictor interactions (Fig. 2A), thereby suggesting an increase in the formation of mTORC2. In contrast, mTOR–Raptor interactions were decreased in Rictor-overexpressing cells, consistent with a previous report showing an increase in mTOR–Rictor precipitation when mTORC1 was inhibited (18). To complement the PLA studies, we also immunoprecipitated for mTOR and found that Rictor and Sin1 binding to mTOR was increased in SAM Rictor cells compared with control, whereas binding of Raptor to mTOR was reduced (Fig. 2B). Furthermore, this Rictor-driven increase in mTORC2 formation increased downstream phosphorylation of AKT (S473) in Rictor-overexpressing cells compared with control (Fig. 2C), suggesting the increased mTORC2 was indeed functional.

In order to assess the effect of Rictor overexpression on mTORC1 activity while excluding the effects of cross-talk between the mTOR complexes, we stimulated SAM Rictor or EV cells with glutamine, a known activator of mTORC1, but not mTORC2 (19). SAM Rictor cells showed a reduction in phosphorylation of S6K1, a direct substrate of mTORC1, compared with SAM EV cells in response to glutamine stimulation (Fig. 2D), confirming a reduction in mTORC1 activity caused by a loss of mTORC1 upon Rictor overexpression. These data suggest amplification of Rictor in NSCLC promotes the formation and activity of mTORC2 at the expense of mTORC1.

To complement overexpression experiments, we used CRISPR-Cas9–mediated genome editing (20–22) to target RICTOR for loss of function in H23 cells, a cell line with verified RICTOR amplification (Fig. 1E; ref. 2). Rictor-deficient H23 cells showed a reduction in mTOR–Rictor interactions and an increase in mTOR–Raptor interactions compared with sgLacZ control cells when assessed by PLA (Fig. 2E). Additionally, immunoprecipitation of mTOR in sgRictor cells showed a loss of Rictor and Sin1 binding to mTOR, whereas Raptor binding was increased (Fig. 2F). As expected with a loss of mTORC2, phosphorylation of AKT (S473) was reduced in H23 sgRictor compared with sgLacZ control cells in response to serum stimulation (Fig. 2G). Glutamine stimulation, however, increased the levels of p-S6RP (S235/236) in Rictor-deficient cells (Fig. 2H), suggesting mTORC1 activity was increased. Together, these results demonstrate that changes in cellular Rictor alter the amount of mTORC2, consequently increasing or decreasing the amount of mTORC1 in the opposite direction.

mTORC2 is a known regulator of cell proliferation and promoter of tumorigenesis (6, 7). However, we found that Rictor overexpression did not alter cell number when grown in 2D (data not shown). To determine if Rictor-driven mTORC2 provides a growth advantage to NSCLC cells, control or Rictor-overexpressing H1975 or H358 cells were grown as Matrigel-embedded 3D cultures for 20 days. Imaging of cultures on the final day showed the area of SAM Rictor cultures was significantly larger than SAM EV cultures (Fig. 3A), suggesting that Rictor overexpression promotes growth of NSCLC.

Conversely, loss of Rictor in H23 RICTOR-amplified cells significantly reduced the number of viable cells compared with control when measured by MTT assay (Fig. 3B). The reduction in the cell number of Rictor-deficient cells was due to an inhibition of cell proliferation, as judged by BrdUrd incorporation (Fig. 3C). These results are consistent with the possibility that Rictor-driven mTORC2 formation promotes the cell proliferation of NSCLC tumor cells.

Next, we investigated whether the growth advantage conferred by Rictor overexpression would result in faster growing tumors in vivo. SAM Rictor #5 or EV control H1975 cells were injected into the hind flanks of Rag1-null immunodeficient mice and tumor growth was monitored for approximately 1 month. Rictor overexpression significantly increased tumor volume compared with control in the xenograft model (Fig. 3D). Tumors were harvested and weighed upon removal. The mass of SAM Rictor tumors was significantly larger than control tumors (Fig. 3E). Western blot analysis of tumor lysates showed that SAM R5 tumors retained the increased expression of Rictor throughout the course of tumor development and exhibited increased levels of p-AKT, a readout of mTORC2 activity (Fig. 3F). Furthermore, SAM Rictor tumors had an increase in Ki-67 staining (Fig. 3G), consistent with studies showing mTORC2 activity drives cell proliferation.

Increased proliferation of Rictor-overexpressing tumors suggested that mTORC2 inhibition might be beneficial for treatment of RICTOR-amplified tumors. However, an mTORC2-specific small molecule is not currently available. A previous study from our lab identified mLST8, a cofactor associated with both complexes, to be selectively required for mTORC2 integrity and function, but not mTORC1 (13); thus, targeting of mLST8 could be used to specifically inhibit mTORC2. As shown in Fig. 4A, CRISPR-Cas9–mediated targeting of mLST8 in H23 cells inhibited coimmunoprecipitation of mTORC2-specific components Rictor and Sin1 with the mTOR kinase, whereas Raptor coimmunoprecipitation increased (Fig. 4A). Western blot analysis of downstream signaling also confirmed that p-AKT was decreased upon mLST8 knockout (Fig. 4A). Consistent with Rictor knockout in Fig. 3, mLST8 knockout also reduced the relative number of viable cells and inhibited cell proliferation as measured by MTT assays and BrdUrd uptake assays, respectively (Fig. 4B and C).

To achieve equal cell numbers between WT and mLST8 cells for implantation in vivo, we utilized a doxycycline-inducible Cas9 system coexpressed with an sgRNA targeting mLST8 (23). In vitro experiments showed that a single dose of doxycycline was enough to induce Cas9-mediated LOF of mLST8, although continuous treatment was required for sustained expression of Cas9 (Fig. 4D). Inducible knockout of mLST8 in H23 resulted in a reduction in colony formation after a single dose of doxycycline in sgMLST8 cells compared with sgLacZ (Fig. 4E).

For xenograft studies, H23 cells expressing a doxycycline-inducible Cas9 and either sgLacZ control or sgMLST8 were injected into the hind flanks of athymic nude mice. Doxycycline was given for 10 days to induce Cas9 expression and mLST8 LOF and tumor growth was measured every 2 to 4 days for 2 months. At the end of the study, tumors were removed, weighed, and subjected to further analysis by IHC. Tumor volume and weight of sgMLST8 tumors were significantly reduced compared with sgLacZ (Fig. 4F and G). Western blot analysis of tumor lysates showed sustained reduction of mLST8 to varying degrees in tumors expressing sgMLST8 (Fig. 4H). Additionally, IHC staining of Ki-67 was reduced in sgMLST8 tumors compared with sgLacZ tumors (Fig. 4I), recapitulating the in vitro results of loss of cell proliferation caused by inhibition of mTORC2 activity.

Amplification of RICTOR has been identified as an actionable target in NSCLC, yet no other mTORC2 component is coamplified with RICTOR in human lung cancer data sets (Fig. 1). Because Rictor was shown to have mTOR-independent functions (24), the mechanism by which amplification of RICTOR could drive oncogenesis has not been clearly defined. We hypothesized that overexpression of Rictor would suppress Raptor–mTOR interactions and increase Rictor–mTOR interactions, driving activity of mTORC2, a known activator of AKT and cell proliferation. Here we show that overexpression of Rictor in NSCLC increases the amount of mTORC2 at the expense of mTORC1, thereby increasing mTORC2 downstream signaling and facilitating tumor cell proliferation.

Aberrant mTORC2 signaling has been identified as an important mediator of tumorigenesis in the context of other cancer-causing mutations such as loss of PTEN or activation of PI3K (12, 25). Amplification of RICTOR in NSCLC also co-occurs with common oncogenic driver mutations, including KRAS and EGFR (2). In this study, we performed all experiments with both KRAS (H358 and H23) and EGFR (H1975)-mutant cell lines. In H358 or H1975 cells harboring these mutations, overexpression of Rictor increased 3D culture growth, yet 3D cultures of immortalized, nontumorigenic lung epithelial cells (BEAS2B) were unchanged in response to overexpression of Rictor (data not shown). Conversely, in H23 RICTOR-amplified and KRAS-mutant NSCLC, loss of mTORC2 inhibited tumor growth, suggesting that mTORC2 activity can promote tumor cell proliferation but may not be sufficient to cause cellular transformation on its own. In addition to proliferation, mTORC2 has known roles in regulation of metabolism and cytoskeletal organization. Additional studies focused on understanding the impact of RICTOR amplification on these other aspects of tumor biology are warranted.

Our study shows that increased mTORC2 formation occurs at the expense of mTORC1 formation, a finding that seems counter to the large body of work demonstrating the importance of mTORC1 in promoting tumor growth. It is important to note that the increased proliferative effect of Rictor overexpression was observed in 3D culture experiments, a condition that more closely mimics the nutrient gradients present in the in vivo tumor microenvironment. A recent study has suggested mTORC1 may inhibit tumor growth in nutrient-starved conditions, offering a potential explanation to how reduced mTORC1 could still promote tumor growth (26). Thus, further investigation of nutrient availability in 3D or in vivo tumor settings is required to better understand how mTORC2 upregulation impacts tumor cell proliferation.

RICTOR is located on the p arm of chromosome 5 (5p13.1), a common site of copy-number gain in lung cancer and other diseases (17). Recent studies, including our own analysis, show that in addition to this overall copy-number gain, the loci surrounding the RICTOR gene is a site of focal amplification (Supplementary Fig. S1; ref. 2). Amplification of RICTOR often co-occurs with amplification or copy-number gain of several other genes along the 5p chromosome, including SKP2 and GOLPH3, both of which have been implicated in cancer progression and modulating the PI3K–mTOR–AKT signaling axis (27–29). Other coamplified genes with known tumorigenic properties include OSMR and LIFR, also located at the 5p13.1 cytoband (30–32). Although we were able to show that overexpression of Rictor alone was able to increase the proliferative capacity of lung cancer cell lines, further studies exploring the interaction between RICTOR and its coaltered genes in cancer are necessary to fully understand the oncogenic effects of amplification occurring at the 5p chromosome.

The identification of Rictor overexpression as a driver of mTORC2 formation and tumor progression suggests that specific inhibition of mTORC2 would be a logical therapeutic strategy for patients with RICTOR-amplified cancers. However, only mTORC1-specific inhibitors (rapamycin analogues) or mTOR kinase inhibitors that block the activity of both complexes are available (33). Our data suggest that increased mTORC2 formation can reduce the levels of mTORC1, thus inhibition of both mTOR complexes may not be necessary for the treatment of RICTOR-amplified cancers. Additionally, inhibition of mTORC1 releases a negative feedback loop on PI3K/AKT signaling (34, 35), which could potentially lead to an unwanted upregulation of mTORC2 signaling. In our study, we target mLST8 to specifically inhibit mTORC2 and show that tumor growth of RICTOR-amplified H23 NSCLC cells is reduced, suggesting inhibition of the mTOR–mLST8 interaction (13) may be an efficacious way to specifically inhibit mTORC2 and treat RICTOR-amplified NSCLC.

L.C. Kim reports grants from National Cancer Institute during the conduct of the study. J. Chen reports grants from VA and grants from NCI during the conduct of the study. No potential conflicts of interest were disclosed by the other author.

L.C. Kim: Conceptualization, data curation, formal analysis, investigation, writing–original draft, writing–review, and editing. C.H. Rhee: Data curation. J. Chen: Conceptualization, resources, funding acquisition, project administration, writing–review, and editing.

We would like to acknowledge the Vanderbilt University Medical Center Translational Pathology Share Resource for their help with IHC experiments. This work was supported by a VA Merit Award 5101BX000134 and a VA Research Career Scientist Award (J. Chen), NIH grants R01 CA177681 (J. Chen), R01 CA95004 (J. Chen), T32 CA009592 (L.C. Kim), and F31 CA2220804-01 (L.C. Kim).

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