The protein phosphatase 2A-B56α complex regulates N-Myc degradation in neuroblastoma

TL;DR

This study shows that the PP2A-B56α complex can regulate N-Myc protein levels in neuroblastoma, offering a new potential treatment approach for high-risk pediatric tumors.

Contribution

The study reveals that PP2A-B56α modulates N-Myc stability via proteasomal degradation, extending prior findings on c-Myc.

Findings

PP2A reactivation reduces N-Myc protein expression in neuroblastoma cells.

DT-061 treatment inhibits tumor growth and reduces N-Myc in a xenograft model.

N-Myc S62 mutation abrogates the effects of PP2A-B56α modulation on cell viability.

Abstract

High-risk neuroblastoma is one of the most common and deadliest pediatric solid tumors. Proliferation, differentiation, and treatment resistance have been linked to the amplification of MYCN. Although N-Myc has proven to be a difficult therapeutic target, our group and others have previously demonstrated that a small-molecule targeting PP2A, DT-061, drives c-Myc degradation in MYC-driven cancers. This results from its ability to bias PP2A toward heterotrimers that contain the B56α regulatory subunit, which dephosphorylates the S62 c-Myc residue, affecting protein stability and driving its proteasomal degradation. Interestingly, despite a high degree of sequence homology in the phosphodegron of c-Myc and N-Myc, the role of PP2A-B56α in regulating the analogous S62 residue on N-Myc is unknown. Here, we show how N-Myc protein expression is significantly reduced after PP2A reactivation in neuroblastoma cell lines. Treatment with DT-061 combined with its inactive competitive antagonist, DT-766, and the proteasome inhibitor, MG-132, reversed this effect on the loss of N-Myc protein expression, suggesting that PP2A-B56α modulation affects N-Myc stability via the proteasomal degradation pathway. A loss in cell viability and inhibition of the colony formation potential of neuroblastoma cells accompanied this loss in N-Myc expression. Conversely, these effects were abrogated when the N-Myc S62 phosphosite was mutated. In a xenograft model, we observed tumor growth inhibition upon DT-061 treatment, along with a reduction in N-Myc protein expression in vivo. Combined, these results highlight the importance of the PP2A tumor suppressor in regulating MYCN oncogenic signaling and open new potential treatment regimens for high-risk neuroblastoma patients.

Figures

Full text

Neuroblastoma (NB) is an extracranial pediatric cancer affecting immature neuroblast cells of the developing sympathetic nervous system. Even with aggressive therapy, children with high-risk NB have a 5-year survival rate of less than 50%, making NB one of the deadliest pediatric solid tumors. High-risk NB is defined by unfavorable histology and is strongly associated with the presence of genetic alterations (1). Amplification of the N-Myc protein, encoded by MYCN, is present in up to 50% of high-risk patients with NB and is a validated molecular driver of NB pathogenicity (2, 3). Proliferation, dedifferentiation, and treatment resistance have all been linked to MYCN amplification and its role in activating tumorigenic transcriptomic core regulatory circuitries, making it the most important prognostic marker for NB patients (1). However, N-Myc remains a challenging therapeutic target, and to date no direct targeted therapies are clinically available, making approaches that indirectly downregulate its activity an area of interest (4). N-Myc belongs to the Myc family of proto-oncogenes, which share a conserved transcriptional activation domain at the N-terminus known as Myc Box 1 (MB1). The MB1 phospho-degron contains the serine 62 (S62) and threonine 58 (T58) amino acid residues, which, when regulated by phosphorylation, determine protein stability through modulating the binding of specific E3 ubiquitin ligase complexes (5, 6). Therefore, understanding the precise mechanisms and effector proteins that regulate the phosphorylation status of this phosphodegron to modulate N-Myc stability could potentially lead to the discovery of new therapeutic approaches to downregulate N-Myc activity and function.

Protein phosphatase 2A (PP2A) is a family of serine/threonine phosphatase heterotrimeric holoenzymes comprised of a scaffolding (A) subunit, a substrate-directing regulatory (B) subunit, and a catalytic (C) subunit. In aggregate, PP2A is considered a tumor suppressor protein, although certain heterotrimers have been described to play tumor-promoting roles (7). In cancer, distinct genetic and non-genetic mechanisms have been identified that result in biased PP2A heterotrimer formation, resulting in the inhibition of the formation of tumor-suppressive holoenzymes of PP2A (8). PP2A B56- and B55-containing PP2A complexes direct most of the tumor suppressive phosphatase activity against signaling pathways associated with cell growth and proliferation, such as MAPK and PI3K/AKT (9). The c-Myc oncoprotein is one of the best-characterized PP2A substrates. The PP2A-B56α complex directly dephosphorylates the S62 residue marking c-Myc for ubiquitin-mediated proteasomal degradation (10, 11, 12). Our group and others have previously demonstrated that utilizing small molecules targeting PP2A drives c-Myc degradation in MYC-driven cancers (13, 14, 15). The small molecule DT-061 biases PP2A heterotrimers towards those containing the B56α regulatory subunit by binding in a pocket formed by the A, C, and B56α subunits, prolonging the off-rate of B56α from the heterotrimeric complex (16). This biases the pool of PP2A complexes toward those containing B56α and its dephosphorylation activity toward B56α-dependent substrates. Treatment with DT-061 has been shown to directly dephosphorylate the S62 c-Myc residue, regulating its protein stability, and driving proteasome-mediated degradation (13, 14, 15, 16). Although there is high sequence homology between the phosphodegron of c-Myc and N-Myc, it has not been determined to date whether the PP2A-B56α heterotrimer is directly responsible for N-Myc dephosphorylation and degradation.

Given the level of conservation of the phosphodegron between the Myc family of proteins, we hypothesize that PP2A-B56α is capable of directly regulating N-Myc degradation in NB models. Here, we demonstrate a significant decrease in N-Myc phosphorylation and expression across multiple independent N-Myc-dependent NB cell lines treated with DT-061. To determine if the observed effects were a result of the PP2A specific effects of DT-061, we utilized the neutral competitive antagonist DT-766, a structurally similar, but inactive analog of DT-061 that has been previously demonstrated to occupy the same binding pocket of DT-061 without promoting PP2A-B56α heterotrimerization (15, 16, 17, 18). We also tested the PP2A selective, but not specific, inhibitor, okadaic acid (OA). Treatment with either DT-766 or OA rescued the loss of N-Myc expression induced by DT-061 (19). The DT-061-mediated loss of N-Myc expression was accompanied by a loss of cell viability and inhibition of colony formation, resulting from an increase in apoptotic cell death in the treated NB cells. These effects on N-Myc protein stability were abrogated when the N-Myc S62 and T58 phosphosites were mutated. In an NB xenograft mouse model, a significant difference in tumor volumes between DT-061 and vehicle treatment was observed, along with a reduction of N-Myc protein expression and its transcriptional activity in the DT-061 treated tumor lysates.

Our findings suggest that PP2A-B56α regulates N-Myc stability by direct dephosphorylation of its S62 residue. The data also indicate that N-Myc can be therapeutically targeted by PP2A modulating compounds, resulting in loss of NB cell growth in vitro and in vivo. Collectively, our work highlights the importance of the PP2A tumor suppressor in regulating N-Myc function and opens new potential treatment regimens for high-risk NB patients.

Results

DT-061 treatment of NB cells drives proteasome-mediated N-Myc degradation

To investigate the potential regulation of N-Myc phosphorylation and protein stability by the serine/threonine phosphatase PP2A, we screened a panel of NB cell lines for N-Myc and c-Myc expression and selected the cell lines with the highest N-Myc protein expression for our studies, specifically SKNBE2, Kelly, and CHP212 (Fig. 1A). We next correlated N-Myc protein expression with functional dependence on N-Myc using the Cancer Dependency Map (DepMap) database (20), which demonstrated that the cell lines with the highest N-Myc expression were highly dependent on the protein for their survival (Fig. S1) (21).Figure 1Small-molecule activation of PP2A drives N-Myc protein degradation in MYCN amplified NB cells.A, NB cell lines are selected by expression of N-Myc and not c-Myc. B, N-Myc overexpressing cell lines, Kelly, SKNBE2, and CHP212 were treated with DMSO, 20, 40, or 80 μM DT-061 for 1 to 6 h, and whole cell lysates were analyzed by immunoblotting for phosphorylated N-Myc (pS62 N-Myc), total N-Myc, and Vinculin. C, quantitative RT-PCR analysis of MYCN mRNA in NB cells following treatment with 20 μM DT-061 for 6 h. Data are normalized to Actin housekeeping and represented as mean fold change ± S.D. (n = 3). D, NB cells were treated with DMSO, 80 μM DT-766 alone, 20 μM DT-061 alone, or a combination of DT-766 and DT-061 for 6 h. Quantification of blots from *panelE*, data represented as mean ± S.D. (n = 3 or 4). p-values were calculated from three independent biological replicates using a two-tailed t-test for *panelC* and one-way ANOVA with Dunnett’s correction for multiple comparisons. ∗p < 0.05, ∗∗p < 0.01, ∗∗∗p < 0.001. E, representative images of N-Myc and Vinculin expression analyzed in panel**D by immunoblotting from whole cell lysate. F, representative images of N-Myc expression by immunoblotting from NB cells treated with DMSO, 20 μM DT-061, 10 μM MG-132, or combination of DT-061 and MG-132 for 6 h.

To assess the effects of PP2A modulation on N-Myc expression in these models, we treated these same cell lines with DT-061, a compound that we have previously demonstrated specifically stabilizes the PP2A-B56α heterocomplex (16). Western blot detection of N-Myc protein expression after 1, 3, and 6 hours of treatment demonstrated dose-dependent loss of N-Myc expression following DT-061 treatment (Figs. 1B and S2). To determine if the loss of N-Myc expression was a result of its transcriptional downregulation, we measured N-Myc mRNA expression by RT-PCR after 6 hours of DT-061 treatment. N-Myc mRNA levels remained unchanged upon DT-061 treatment in both Kelly and SKNBE2 and increased in the CHP212 cell line (Fig. 1C), suggesting a post-transcriptional mechanism for the loss of N-Myc expression seen after DT-061 treatment. Next, to determine the PP2A-B56α specificity of DT-061 modulation on loss of N-Myc expression, we co-treated these cells with the neutral competitive antagonist, DT-766 (also known as TRC-766), which binds in the same pocket as DT-061 but does not promote PP2A-B56α heterotrimerization (15, 17, 22, 23). This resulted in the complete abrogation of the total N-Myc protein expression previously observed (Fig. 1, D and E). Further, we used the pan-PP2A and -PP1 inhibitor OA in combination with DT-061 to determine if the loss of N-Myc expression was phosphatase specific (24, 25). Here, the inhibition of the catalytic activity of PP2A abrogated the loss of N-Myc expression upon DT-061 treatment, suggesting that the downregulation of N-Myc protein expression is dependent on PP2A phosphatase activity (Fig. S3A). Finally, to determine if the loss of N-Myc protein expression was mediated through proteasomal degradation, we co-treated cells with the proteasome inhibitor MG-132 and DT-061. These results showed that this combination treatment abrogated the effects of DT-061-driven N-Myc degradation, suggesting that the observed loss of expression is a result of PP2A-mediated proteasomal degradation (Figs. 1F and S4). Collectively, these results indicate that in NB cells, DT-061 treatment results in degradation of the N-Myc protein through a post-transcriptional mechanism, which is specific to the protein phosphatase activity of PP2A.

Modulation of PP2A inhibits NB clonogenicity and induces apoptosis

The DepMap CRISPRi data indicates that these MYCN-amplified lines are highly dependent on this gene for their survival. Therefore, we next explore if the DT-061-driven loss of N-Myc protein expression affected NB cell viability in vitro. The cytotoxic effect of DT-061 was determined using an 3-(4,5-dimethylthiazol-2-yl)-5-(3-carboxymethoxyphenyl)-2-(4-sulfophenyl)-2H-tetrazolium (MTS) cell viability assay, with a calculated growth inhibitory concentration of 50% (GI50) of around 15 μM after 48 hours of treatment (Fig. 2A). While treatment with the neutral competitive antagonist DT-766 had no effect on the cell viability of NB cells, co-treatment with DT-061 partially abrogated the cytotoxic effects, suggesting the observed effect of DT-061 treatment on cell viability is PP2A-dependent (Figs. 2A, and S5, A and B). In long-term cell growth assays, we utilized colony formation assays (CFAs), where NB cells were plated at low density and treated for 2 weeks with DT-061, resulting in a dose-dependent inhibition of colony formation with a calculated GI_50_ of ∼5 μM (Fig. 2, B and C).Figure 2**DT-061 inhibits clonogenicity and induces apoptosis.**A, cell viability of NB cells was measured by MTS post-treatment with increasing concentrations of 0 to 80 μM DT-061 for 48 h. GI_50_ values were determined with or without co-treatment with 80 μM DT-766. Data represented as mean GI_50_ ± S.D. Experiments were performed with technical triplicates within three independent biological replicates (n = 3) and normalized to DMSO control; statistical significance determined by two-way ANOVA with Bonferroni’s post hoc analysis; ∗∗p < 0.01. B, representative images of colony formation assays of NB cells plated at low density and treated with 0 to 10 μM DT-061 for 14 days. E, colonies were counted using ImageJ and normalized to DMSO control to determine GI_50_ (n = 3). D, N-Myc and apoptotic markers, cleaved PARP and cleaved caspase-3, were detected by immunoblotting after 24 h treatment with 20 μM DT-061 in CHP212 and Kelly (n = 3).

Western blot analysis showed an induction of apoptotic protein markers, including cleaved PARP and cleaved caspase-3 in CHP212 and Kelly cell lines upon DT-061 treatment, which correlated with loss of N-Myc protein expression in these cells (Fig. 2D). Collectively, these findings support a model in which the DT-061-driven loss of N-Myc expression results in the inhibition of NB cell survival and the induction of apoptosis.

Regulation of N-Myc expression is dependent on the PP2A heterotrimeric complex containing B56α

We have previously published that DT-061 binds to a specific pocket in PP2A-B56α heterotrimer using a solved cryo-EM structure (16). Additionally, the PP2A-B56α subunit directly dephosphorylates c-Myc, resulting in its degradation. To determine if the DT-061-mediated effects on N-Myc were dependent on the PP2A-B56α holoenzyme, we generated cell knockouts of B56α, B55α (a structurally distinct PP2A B regulatory subunit family member), and an empty vector (EV) control in the SKNBE2 cell line using CRISPR/Cas9 (Fig. 3, A and B). In SKNBE2 cells lacking B56α, DT-061 had no effect on N-Myc expression, while in the SKNBE2 cells with either the empty vector control or cells lacking B55α, there is consistent loss of N-Myc expression to similar levels as that seen in the control parental cell line (Fig. 3, C and D). Collectively, these findings suggest that the N-Myc dephosphorylation and degradation are primarily mediated by PP2A-B56α reactivation.Figure 3**CRISPR knock-down of the B56α regulatory subunit abrogates the effect of DT-061.**A, immunoblotting validation of the protein expression loss of B56α by stable CRISPR-Cas9 expression along with an empty vector (no guide) and B55α as controls in SKNBE2. B, normalized protein expression of B56α and B55α in the knock-down SKNBE2 cells compared to EV (n = 4) represented as mean ± S.D. Statistical significance determined by one-way ANOVA with Dunnett’s post hoc multiple comparisons test ∗p < 0.05, ∗∗∗p < 0.001. C, representative image of N-Myc immunoblot from SKNBE2 EV, B56α, and B55α cells treated with DMSO or 20 μM DT-061 for 6 h. D, quantification of blots from panel B was normalized to Vinculin and then relative to the DMSO treatment of the corresponding knock-down cells (n = 4) and analyzed using two-way ANOVA with Tukey’s post hoc multiple comparisons test ∗p < 0.05, ∗∗p < 0.01, ∗∗∗p < 0.001; ns, not significant.

PP2A-mediated dephosphorylation of S62 of N-Myc regulates its expression

Phosphorylation of S62 on the c-Myc protein is stabilizing, and subsequent phosphorylation at the T58 site is required for its degradation. However, the c-Myc protein is not ubiquitinated until PP2A dephosphorylates the S62 residue, leaving T58 solely phosphorylated (10). Mutations in these residues have been used as tools to study the mechanisms of phosphorylation-dependent signaling on the regulation of c-Myc protein stability. Based upon these previous observations, we generated phospho-mutants at these sites to determine the importance of these post-translational modifications on the observed PP2A-mediated effects on N-Myc protein stability (11, 12, 26). Specifically, the threonine 58 to alanine (T58A) mutant results in a non-phosphorylatable residue, thus blocking T58 phosphorylation by GSK3β, a step required for PP2A-dependent dephosphorylation of the S62 residue (27). The serine 62 to aspartic acid (S62D) mutation mimics a constitutively phosphorylated residue that cannot be dephosphorylated in a PP2A-dependent manner, thus preventing a critical ubiquitination step by the E3 ligase, Fbw7 (5, 10, 12). Therefore, we generated Kelly and SKNBE2 cell lines expressing the V5-tagged WT) S62D, or T58A mutant forms of N-Myc (Table S1). We performed cycloheximide chase experiments to determine the stability of the WT and mutant N-Myc isoforms at baseline (Fig. 4, A and B). Consistent with previous observations related to c-Myc, the N-Myc^S62D^ mutant and N-Myc^T58A^ mutant proteins were more stable compared to the N-Myc^WT^ protein (Fig. 4, A and B). The calculated half-life of the N-Myc^WT^ protein in this assay was approximately 13 min, whereas the half-lives of either single mutant were greater than 60 min. Similar results were observed in the SKNBE2 cell line (Fig. S6).Figure 4**PP2A activity is dependent on the N-Myc phosphorylation sites S62 and T58.**A, Kelly cells were transduced with pLenti6.3 expressing V5-tagged WT N-Myc and mutant S62D or T58A N-Myc. Cells were treated with 25 μg/ml cycloheximide in a chase experiment to assess protein stability over 60 min by immunoblot of whole cell lysate for V5-tagged N-Myc. B, quantification of V5-tagged N-Myc plotted after normalization to Vinculin and the untreated control. Data plotted as mean ± S.D. are from four independent experiments (n = 4), and half-lives were determined from exponential decay curve fitting. C, Kelly cells expressing the WT, S62D, and T58A N-Myc are treated with DMSO or 20 μM DT-061 for 3 h and immunoblotted for V5-tagged N-Myc, representative images are presented. D, quantification of V5-tagged N-Myc after normalization to Vinculin and DMSO controls from three independent experiments (n = 3). Statistical significance was determined by two-way ANOVA; ∗p < 0.05; ns, not significant. E, representative images of clonogenic assays of Kelly WT and S62D N-Myc plated at low density and treated with DT-061 for 14 days. F, colony formation assays from panel E were quantified by absorbance of dissolved crystal violet stained colonies, and plotted data represent the mean ± S.D. of three independent biological replicates (two technical replicates for each biological replicate).

We next selected a time point in which DT-061 treatment resulted in robust loss of N-Myc expression before the induction of cell death, to determine whether destabilization of N-Myc by DT-061 treatment was dependent on S62 dephosphorylation. In cells treated with DT-061 for 3 hours, we saw a marked reduction of N-Myc^WT^ protein expression, but not the N-Myc^S62D^ or N-Myc^T58A^ mutants (Fig. 4, C and D). This suggests that the DT-061 mediated degradation of N-Myc is dependent on the MB1 regulatory element and parallels the mechanism described for c-Myc, where the stability of N-Myc is regulated by the phosphorylation of T58 and subsequent PP2A-mediated dephosphorylation of S62. Consistent with this model, when we challenged Kelly N-Myc^WT^ and N-Myc^S62D^ with DT-061 in CFA, we found that N-Myc^S62D^ expressing cells were more resistant to drug treatment, implying that the growth inhibition we observed could be at least partially explained by the PP2A-driven N-Myc degradation. (Fig. 4, E and F). Interestingly, despite differences in N-Myc stability, there were no significant alterations in expression of N-Myc downstream targets AP2β, GATA3, and PHOX2B between N-Myc^WT^ and N-Myc^S62D^ expressing cells (Fig. S7A). Similarly, baseline clonogenic capacity did not differ between the two cell populations, where the mutant was not associated with an increased colony-forming ability despite further stability (Fig. S7B). Overall, these experiments indicate that PP2A is a regulator of N-Myc expression and function, which had not been previously shown in the literature.

PP2A-B56α stabilization results in tumor growth inhibition in vivo

To investigate whether our observations translated to in vivo models of the disease, we injected the Kelly MYCN-amplified cell line subcutaneously into immunodeficient BALB/c nu/nu mice. Mice were randomized into Vehicle control or DT-061 (5 mg/kg BID) treatment groups. DT-061 treatment led to a significant decrease in the average tumor volume (Fig. 5A) with a terminal tumor growth inhibition of ∼39% (Fig. 5B). Western blot analysis of the tumor lysates revealed a significant reduction in N-Myc expression in the DT-061 treated group compared to the vehicle controls. (Fig. 5, C and D). The protein expression of the N-Myc transcriptional core regulatory circuit members, GATA3 and PHOX2B, was also significantly downregulated, further confirming the downstream effects of the PP2A-driven N-Myc loss seen in vivo (Fig. 5, C and D) (28). To further characterize whether these changes in N-Myc expression resulted in effects on downstream signaling, we processed the tumors for mRNA and measured the gene expression of transcriptional targets regulated by N-Myc. We found that in the DT-061-treated tumours, ASCL1, GATA3, HAND2, ISL1, PHOX2B, and TBX2 were all significantly downregulated compared to control, vehicle-treated mice, consistent with the loss of N-Myc expression seen upon PP2A modulation in vivo (Fig. 5E). These findings suggest that PP2A-driven N-Myc degradation inhibits tumor growth in vivo and establishes that single-agent DT-061 is tolerable and efficacious in N-Myc amplified NB models (Fig. 5F).Figure 5**PP2A activation results in tumor growth inhibition in a neuroblastoma xenograft mouse.**A, Kelly cells (5 × 10^6^) were subcutaneously injected into nude mice and randomized when tumors reached 100 mm^3^ into Control (n = 14) or 5 mg/kg DT-061 (n = 11) treated groups treated every other day. Tumors were measured by caliper every other day and are plotted as mean tumor volume ± S.E., two-way ANOVA with Dunnet’s multiple comparisons test; ∗p < 0.05, ∗∗p < 0.01, ∗∗∗p < 0.001. All mice were enrolled for 11 days and were sacrificed after reaching endpoint tumor size per the institutional guidelines. B, the terminal tumor volume by caliper was recorded and plotted for each individual mouse; one-way ANOVA with Dunnett’s multiple comparisons test ∗∗∗∗p < 0.0001. C and D, protein from Control (n = 14) or 5 mg/kg DT-061 (n = 10) were isolated from the tumor fragments and processed for immunoblotting for N-Myc and downstream effector proteins. Expression was first normalized to Vinculin and plotted for each individual mouse as relative expression to the average of the control group as fold change ± S.D; two-tailed t-test ∗∗p < 0.01, ∗∗∗p < 0.001. E, mRNA was purified from the isolated tumor fragments and converted to cDNA before quantitative PCR of downstream effector gene expression. Actin was used as a loading control. Data represent fold change ± S.D. relative to the control group; one-way ANOVA with Dunnett’s multiple comparisons test ∗∗∗p < 0.001, ∗∗∗∗p < 0.0001. F, body weight (grams) is plotted for the control (n = 14) and DT-061 treated (n = 11) mice as mean ± S.E.

Discussion

High-risk NB remains a disease with few targeted treatments and devastatingly poor outcomes. Despite the low-mutational burden of these cancers, the heterogeneity of NB, coupled with the pharmacological challenge of inhibiting Myc transcription factors, makes it a difficult disease to treat. Therefore, recent advances in targeting N-Myc have relied on downregulating MYCN gene expression or altering its protein stability (4, 29, 30, 31). Based upon the extensive work deciphering the relationship between PP2A and MYC-driven cancers done by our group and others, we hypothesized that N-Myc protein stability is regulated post-translationally by the PP2A serine/threonine phosphatase in a manner similar to c-Myc, due to their highly homologous MB1 element (13, 16, 17, 32, 33). Here, we report that the PP2A-B56α heterotrimer can negatively regulate N-Myc protein expression in NB and that PP2A-B56α selective reactivation can inhibit tumor growth both in vitro and in vivo.

Our findings suggest that small-molecule targeting of PP2A heterotrimerization is sufficient to promote N-Myc degradation and inhibit tumor growth. Indeed, these discoveries corroborate previous findings by Nazam et al. utilizing different PP2A activators (14, 34). For example, we observe similar decreases in the expression of both the total and S62 phosphorylated N-Myc upon treatment with DT-061, followed by an induction of cleaved PARP, along with tumor growth inhibition in our MYCN-amplified xenografts. Owing to the structural and regulatory complexity of PP2A, and in contrast to our study, Nazam et al. suggest that their PP2A-activating compounds decreased the activity of two endogenously expressed PP2A inhibitors, which have been characterized as tumor-promoting in NB (8, 35). To further expand this relationship between PP2A and NB, we investigated the importance of the tumor suppressive PP2A-B56α complex on regulating N-Myc stability by using DT-061 (16). The data we report suggest that PP2A-B56α is a major regulator of N-Myc stability and function; however, the effect is not completely abrogated when we genetically knocked down B56α expression by CRISPR. We speculate that this partial rescue of N-Myc expression upon B56α knockdown could be the result of incomplete polyclonal knockdown of this subunit by CRISPR or through compensatory upregulation of other B regulatory subunits in response to targeted reduction of B56α (16). Thus, while the role of the PP2A-B56α heterotrimer is clear, further investigation into other B regulatory members and the PP2A inhibitory families of proteins is necessary to fully understand the role of this family of serine/threonine phosphatases in regulating N-Myc expression and function.

The extent to which PP2A plays a tumor suppressive role in NB outside of its regulation of N-Myc remains an open and important question. While the silencing of N-Myc can induce NB cell death and senescence, PP2A is able to affect multiple pathways independent of N-Myc to induce apoptosis (17, 36, 37, 38). In fact, when we measured the colony-forming potential in cells expressing the S62D mutant N-Myc, these cells were not completely resistant to DT-061 treatment, implying N-Myc suppression alone is not responsible for all DT-061-mediated effects on cell viability and survival, which aligns with the knowledge of B56α′s broad substrate profile. On the contrary, we discovered an absence of enhanced clonogenic activity and altered differentiation marker expression in cells with stabilizing S62D N-Myc mutant compared to wild-type. These findings were surprising but may reflect the context in which these experiments were conducted. Because the Kelly cell line is a MYCN-amplified cell line, N-Myc expression is already pathologically high and is operating near its maximal threshold. Therefore, further stabilization of N-Myc through the S62D mutant may be insufficient to drive additional oncogenic output. Additionally, we demonstrated that the T58A mutant N-Myc affected half-life modestly, while this same mutation in the c-Myc phosphodegron has been previously shown to markedly improve stability as compared to the S62 mutant (6, 39). These differences between the regulation of N-Myc and c-Myc stability could be attributed to the existence of additional post-translational modifications on N-Myc that may affect its stability (40). To attribute the loss of N-Myc to its dephosphorylation by PP2A-B56α, we also attempted to measure the loss of phosphorylated S62 N-Myc upon DT-061 treatment but were unable to decouple it from the loss of total N-Myc expression. Further molecular analysis of this mechanism, ideally in a model that expresses no endogenous N-Myc, is necessary to determine the specificity of B56α to these post-translational modifications.

Downregulation of N-Myc expression, signaling, and function is a therapeutically relevant strategy, and approaches targeting this mechanism are now in clinical trials (41). Importantly, we are able to see single-agent efficacy in our xenograft model comparable to that of other therapies indirectly targeting N-Myc downregulation, such as AURKA, BET, and mTOR inhibition (42, 43, 44). However, these therapies are being explored clinically in combination with inhibitors of pathways like RAS/MAPK, ALK, and targeted immunotherapies as combination-based therapies to improve efficacy. For example, a phase II clinical trial investigating AURKA inhibition found that patients without MYCN amplification had better outcomes than those who did, suggesting that this inhibitor may be unable to overcome the increased N-Myc expression seen in those patients (45). Additional studies with PP2A modulators that could enhance the loss of N-Myc expression through a distinct mechanism would be of interest to the field. Collectively, these studies provide a rationale for investigating PP2A reactivation strategies in combination with other therapies for NB treatment.

While MYCN amplification is most prevalent in NB, and is commonly found in tumors of the developing nervous system, including medulloblastomas, retinoblastomas, and glioblastomas (2), it is also expressed in some subsets of prostate, small-cell lung, and hematologic cancers (46). Therefore, understanding how PP2A can modulate N-Myc stability and function could be beneficial for the treatment of other tumor types. Combined with the broad substrate repertoire and tumor suppressive activity of PP2A-B56α, small molecules targeting this mechanism can be advantageous for diseases such as NB, where cellular plasticity has been implicated in resistance, and in non-neuronal MYCN-driven cancers (46, 47). Additionally, NB subsets containing MYC overexpression and non-MYC-driven subtypes (1p/11q deletion, 17q gain) could benefit from this approach and are worth investigating (1). In conclusion, we demonstrate that PP2A is an important negative regulator of N-Myc expression and function and that B56α plays a major role in this process, proposing a compelling rationale for targeting this complex in *MYCN-*driven malignancies.

Experimental procedures

Cell lines and compounds

Human NB cell lines Kelly, SKNBE2, and CHP212 were obtained from the American Type Culture Collection. All cell lines were maintained in RPMI 1640 (Gibco, 11875119) supplemented with 10% fetal bovine serum, 100 units/ml penicillin and 100 μg/ml streptomycin (Gibco, 15140122) at 37 °C and 5% CO2. Cell cultures were mycoplasma tested at least every other month. DT-061 (HY-112929) and DT-766/TRC-766 (HY-131443) were purchased from MedChem Express, OA was purchased from P212121, and cycloheximide solution was purchased from Sigma-Aldrich (C4859). DT-061, DT-766, and OA were reconstituted in dimethyl sulfoxide (DMSO) and stored at −20 °C.

Immunoblotting

Whole-cell lysates were extracted by RIPA (Thermo Fisher Scientific, PI89901) supplemented with protease (Roche, 05892791001) and phosphatase (Roche, 4906837001) inhibitors. Total protein concentration was determined using Pierce BCA Protein Assay (Thermo Scientific, PI23225) and 10 to 20 μg of protein were separated by polyacrylamide gel electrophoresis (Bio-Rad) and transferred to nitrocellulose membranes using the Bio-Rad Trans-Blot Turbo system. PageRuler Prestained Protein Ladder (Fisher Scientific, PI26617) was used to confirm expected kDa of targets (Fisher Scientific, PI26617). Membranes were blocked in 5% non-fat milk/TBST for 1 hour and incubated overnight at 4 °C with primary antibodies: c-Myc (ABclonal, A19032, 1:1000), phospho S62 c-Myc (Abcam, ab185656, 1:1000), N-Myc (Abcam, ab16898, 1:1000), Vinculin (Santa Cruz Biotech, sc-73614, 1:5,000), GAPDH (Santa Cruz Biotech, sc-32233, 1:2000), V5-tag (Cell Signaling Technology, 13,202, 1:1000), AP2β (Cell Signaling Technology, 2509, 1:1000), GATA3 (Cell Signaling Technology, 5852, 1:1,000), PHOX2B (Santa Cruz Biotech, sc-376997, 1:1,000), PARP (Cell Signaling Technology, 9542L, 1:1000), Cleaved caspase-3 (Cell Signaling Technology, 9664L, 1:1,000), and PPP2R2A (Santa Cruz Biotech, sc-81606, 1:1000). PPP2R5A (3A6-D3, 1:500) antibody was provided by Dr Egon Ogris. After washing, membranes were incubated with HRP-conjugated secondary goat anti-rabbit (Cytiva, NA9341Ml) or goat anti-mouse IgG (Jackson ImmunoResearch, 115–035–003) at 1:10,000 for 1 hour at room temperature. Blots were developed using ECL (National Diagnostics, CL300200Ml) on a Bio-Rad ChemiDoc MP Imaging System.

Cell proliferation and viability assays

Cell viability was assessed using the CellTiter 96 AQueous One Solution Cell Proliferation MTS Assay (Promega, G1111) according to the manufacturer’s protocol. Briefly, cells were seeded into 96-well plates at a density of 5000 to 10,000 cells per well in 100 μl of culture medium. After the indicated treatments or time points, 20 μl of the MTS reagent was added directly to each well and incubated at 37 °C for 2 to 4 h. Absorbance was measured at 490 nm using a microplate reader (BioTek Synergy HTX), with background subtraction from blank wells. Cell viability was expressed as a percentage relative to DMSO-treated control wells.

Colony formation assays

CFAs were performed to assess the clonogenicity of NB cells following treatments. Kelly, CHP212, and SKNBE2 cells were seeded in six- or 12-well plates at a density of 500–1000 cells per well in their respective medium and allowed to adhere for 2 days. Cells were then treated with DMSO, DT-061, DT-766, or a combination. Fresh medium containing the corresponding treatments was replaced every 5 to 7 days. After ∼14 days, colonies were fixed with 100% methanol (Fisher Scientific, A452SK-4) for 15 min at room temperature, washed with DPBS (Gibco, 14190250), and stained with 0.5% crystal violet (Fisher Scientific, C581) for 30 min. Excess stain was removed by rinsing with water and air-dried.

For quantification, colony images were photographed with the Bio-Rad MP imaging system and colonies were counted using ImageJ. Alternatively, colony formation was quantified by solubilization of crystal violet in 10% acetic acid, and absorbance was measured at 590 nm using a BioTek Synergy HTX microplate reader. Colony formation efficiency was expressed as the percentage of colonies or absorbance relative to control conditions. All experiments were repeated in at least three independent biological replicates.

Site-directed mutagenesis

Site-directed mutagenesis of the human MYCN gene was performed using the QuikChange Lightning Site-Directed Mutagenesis Kit (Agilent Technologies, 210519) following the manufacturer’s protocol. The pLenti6.3/V5-DEST plasmid with V5-tagged MYCN (HsCD00860757, DNASU) or pLenti6.3/EV (Addgene, 120848) negative control was used. Two point mutations—T58A and S62D—were introduced individually using mutagenic primer pairs designed to substitute the corresponding codons (Table S1). PCR amplification was performed using PfuUltra High-Fidelity DNA polymerase included in the kit, followed by DpnI digestion to remove methylated parental plasmid DNA. The resulting plasmids were transformed into XL10-Gold ultracompetent cells, and colonies were screened by ampicillin resistance. Plasmid DNA was recovered and purified from multiple independent colonies by Mini- (Qiagen, 27106) and Maxiprep (Qiagen, 12663). All mutations were confirmed by whole-plasmid sequencing (Plasmidsaurus). The lentiviral packaging of the plasmid was performed by the University of Michigan Vector Core. Verified mutant MYCN constructs were lentivirally transduced into Kelly and SKNBE2 cells with 4 μg/ml polybrene (Santa Cruz Biotech, sc-134220) at 100,000 cells/well in 6-well plates. After 48 hours post-transduction, the medium was replaced with fresh growth medium containing 10 μg/ml blasticidin (Invivogen, ant-bl-05) for selection and maintained for at least 7 days to establish stable knockout populations. Successful transduction was measured by the detection of V5-tagged protein by immunoblotting.

Quantitative RT-PCR

Total RNA was extracted using the High Pure RNA Isolation Kit (Roche, 50–997–731) and quantified using the Take3 Microplate with the BioTek Synergy HTX. 1 μg of total RNA was used as a template for cDNA synthesis with the High-Capacity cDNA Reverse Transcription Kit (Applied Biosystems, 4374966) and diluted 1:20 prior to RT-PCR. RT-PCR was performed using a QuantStudio 5 Real-Time PCR System with Power SYBR Green PCR Master Mix (Applied Biosystems, 4367659). Primer sequences used are in Table S2.

CRISPR knock-out

CRISPR/Cas9-mediated gene knockout was performed using the pLentiCRISPRv2 all-in-one vector (GenScript), which encodes both Cas9 and the gene-specific single guide RNA. The sgRNA sequences targeting the gene of interest were designed using GenScript’s online CRISPR guide RNA design tool to minimize potential off-target effects and are available in Table S3. Oligonucleotides containing the sgRNA sequences were synthesized, annealed, and cloned into the pLentiCRISPRv2 plasmid according to the manufacturer’s protocol. An empty vector with no sgRNA sequence was used as a negative control. The lentiviral packaging of the plasmid was performed by the University of Michigan Vector Core. Kelly and SKNBE2 cells were transduced with the viral supernatant in the presence of 4 μg/ml polybrene (Santa Cruz Biotech, sc-134220). After 48 hours post-transduction, the medium was replaced with fresh growth medium containing 2 μg/ml puromycin (Invivogen, ant-pr-1) for selection and maintained for at least 7 days to establish stable knockout populations. Knockout efficiency was verified by immunoblotting for target protein.

In vivo xenograft model

Experiments on mice were performed with protocol approval by the Case Western University Institutional Animal Care and Use Committee (IACUC). BALB/c nu/nu mice were purchased from Charles River Laboratories. Kelly (5 × 10^6^) cells were mixed 1:1 with Matrigel (Corning 356230) and were injected subcutaneously into the right flanks of the mice. Mouse tumor weights were assessed by caliper measurement, and when the average tumor volumes reached 100 mm^3^, mice were randomized into twice-daily, oral gavage treatment of vehicle or 5 mg/kg DT-061, resuspended in a 10% N,N-Dimethylacetamide (Sigma-Aldrich, 271012), 10% solutol (Sigma-Aldrich, 42966), and 80% water. Tumor measurements and body weights were recorded every other day throughout the study. Tumor measurements were calculated by the following formula:

$$[eqn]$$

Treatment was administered for 11 days, and euthanasia was performed when the mice reached IACUC-approved terminal tumor volumes (∼2000 mm^3^). Mice in both treatment and control groups were taken down at the same treatment time point after recording final tumor and body weight measurements. Tumor fragments from each mice were collected for downstream analysis. Tumors dissociated by T-PER tissue lysis (Thermo Fisher Scientific, 78510) were processed for Western blot analysis according to above immunoblotting methods. For mRNA analysis, tumor fragments were homogenized with Trizol (Fisher Scientific, 15–596–026) and mRNA purified by phase separation with chloroform (Sigma-Aldrich, 288306) and isopropanol (Sigma-Aldrich, I9516). RT-PCR analysis was conducted following the protocol above.

Data availability

The dataset used for Figure S1 are sourced from The Cancer Dependency Map, (DepMap Public 24Q2) (48). All other data supporting the findings of this study are available in the main figures or supporting information.

Supporting information

This article contains supporting information.

Conflict of interest

The authors declare the following financial interests/personal relationships which may be considered as potential competing interests: C. M. O. reports receiving consulting fees from RAPPTA Therapeutics outside of the scope of the presented work. A. D. reports other support from RAPPTA Therapeutics during the conduct of the study. G. N. had an equity stake in RAPPTA Therapeutics and serves as a consultant for RAPPTA. No disclosures were reported by other authors.