Didymin Induces Apoptosis by Inhibiting N-Myc and Upregulating RKIP in Neuroblastoma

Neuroblastomas are the neural crest–derived malignant tumors which constitute most common solid tumors in infants outside the central nervous system (1). More than 90% of the neuroblastomas are diagnosed before 5 years of age with greater occurrence of refractory and relapsed neuroblastomas in pediatric population over 18 months. Neuroblastomas metastasize to various sites such as skull, bones, spine, and retro-orbital tissues, which lead to challenging clinical presentations such as proptosis and retro-orbital ecchymosis in children (2–4). The major genetic causes for the incidence and therapeutic refractoriness of neuroblastomas include amplification of the oncogene N-Myc and loss of tumor suppressor p53 (5, 6). Raf-kinase inhibitory protein (RKIP) is a novel protein that interacts with MAP/ERK kinases and acts as an inhibitor of mitogen-activated protein kinase (MAPK) pathway (7). RKIP is an established metastasis suppressor protein and, given the role of MAPK pathway in regulating the survival and metastatic potential, RKIP has been the focus of recent investigations in assessing the effects of novel anticancer agents (8, 9). Loss of p53 leads to increased activity of multifunctional proteins, such as RLIP76, which mediate enhanced proliferation, invasion, and drug/radiation resistance in neuroblastoma (10, 11). In this regard, effective strategy for targeting the incidence of neuroblastoma in genetically identified risk groups should decrease the incidence of neuroblastoma, as well as improve the survival following initial diagnosis.

Given the limited 3-year event-free survival of 15% in the incident population and the nature of symptoms in affected children, validation of novel compounds becomes a vital focus of translational research to effectively target neuroblastomas. Also, due to the developing nervous and organ systems in children, the interventional choices will naturally favor active anticancer ingredients of safe dietary components, such as fruits, vegetables, and the compounds derived from them. Didymin is a flavonoid that is richly expressed in citrus fruits such as oranges, lemons as well as other dietary compounds such as mandarin and bergamot. Recently, it was reported that didymin can cause cell death in non–small cell lung cancer in a p53-independent manner (12). The use of chemotherapeutic drugs such as cisplatin, doxorubicin, and vincristine is highly limited in refractory and relapsed neuroblastomas due to frequent loss-of-function mutations in the tumor suppressor p53 (13, 14). Hence, the elucidation of signaling pathways through which didymin acts in neuroblastomas would have potential implications toward the effective management of neuroblastomas. In this regard, we investigated the efficacy and mechanisms of action of didymin in vitro cultures and in vivo mice xenograft models of neuroblastomas.

Our studies first focused on testing the effect of didymin on the survival and clonogenic potential of both p53 wild type and p53 mutant neuroblastomas. Next, we investigated the induction of apoptosis followed by analyzing the impact on key nodes of proliferation in neuroblastomas. Cell-cycle analysis revealed a mechanism of action of didymin that includes its ability to regulate cyclin signaling downstream of p53. Importantly, we investigated the effect of didymin on N-Myc by Western blot, mRNA, and N-Myc luciferase promoter–reporter assays followed by in vivo mice xenograft studies. Our studies revealed the ability of didymin to regulate important nodes of signaling along with being effective, irrespective of p53 status in neuroblastomas.

Didymin, hesperidin, and 2′-hydroxyflavanone (2HF; purity >98%) were purchased from Indofine Chemical Company. PARP, cyclin B1, cyclin D1, CDK4, Akt, pAkt (S⁴⁷³), glyceraldehyde-3-phosphate dehydrogenase (GAPDH), N-Myc, Ki67, CD31, and RKIP antibodies were purchased from Santa Cruz Biotechnology, Upstate Cell Signaling, and Cell Signaling Technologies. The siGENOME smart pool siRNA for RKIP and control scrambled siRNA were purchased from Dharmacon. APO LOGIX carboxyfluorescein (FAM) Caspase Detection Kit was purchased from Cell Technology. Lipofectamine 2000 and fluorescein isothiocyanate (FITC)-labeled Annexin V Conjugate Detection Kit were purchased from Invitrogen. The thymidine kinase promoter–driven Renilla luciferase (pTK-RL) and terminal deoxynucleotidyl-transferase dUTP nick-end labeling (TUNEL) Fluorescence Kit were obtained from Promega. Avidin/biotin complex (ABC) Detection Kit was purchased from Vector.

CHLA-90 and SK-N-BE2 (p53 mutant) as well as SMS-KCNR and LAN-5 (p53 wild-type) human neuroblastoma cell lines were kindly authenticated and provided in May 2010 by Dr. Patrick Reynolds, Children's Neuroblastoma Cancer Foundation/Children Oncology group, Texas Tech University, School of Medicine, Lubbock, TX. The authentication of cell lines was done by analyzing 15 different human short-tandem repeat (STR; carried out by Center for Investigative Genetics core facility at University of North Texas Health Science Center, Fort Worth, TX) to test for interspecies contamination. The cell lines were last tested in April 2011. Cells were grown in complete medium consisting of Iscove's modified Dulbecco's medium supplemented with 3 mmol/L l-glutamine, 5 μg/mL each of insulin and transferrin, 5 ng/mL of selenous acid (ITS culture supplement), and 20% FBS at 37°C in a humidified 5% CO₂ atmosphere. Cell lines were subcultured by detaching without trypsin from culture plates by using a modified Puck's Solution A plus EDTA (Puck's EDTA), which contain 140 mmol/L NaCl, 5 mmol/L KCl, 5.5 mmol/L glucose, 4 mmol/L NaHCO3, 0.8 mmol/L EDTA, 13 mmol/L phenol red, and 9 mmol/L HEPES buffer (pH 7.3). All the cells were also tested for Mycoplasma once every 3 months.

Cell density measurements were done by a hemacytometer to count dye-excluding cells resistant to staining with trypan blue. Approximately 2 × 10⁴ cells were plated into each well of a 96-well flat-bottomed microtiter plate 24 hours prior to addition of medium containing varying concentrations of indicated flavonoids. After 12 hours of incubation, indicated flavonoids (didymin, hesperidin, 2HF; ranging from 0–200 μmol/L each) were added to 8 replicate wells. After 48 hours of incubation, 20 μL (5 mg/mL stock) MTT was added to each well and incubated for 2 hours at 37°C. The plates were centrifuged and medium was removed. Formazan dye trapped in cells was dissolved by addition of 100 μL dimethyl sulfoxide with gentle shaking for 2 hours at room temperature, followed by measurement of absorbance at 570 nm (15–17).

The desalted RKIP siRNA (Dharmacon) was resuspended in 1× universal buffer provided by Dharmacon Research laboratory. Cells were transfected with scrambled and RKIP siRNA at the concentration of 1 μg/mL in serum-free medium, using Lipofectamine 2000 (Invitrogen) for 3 hours, according to the manufacturer's instructions. Excess siRNA was washed off with PBS and complete medium (i.e., with FBS) was added. After 24-hour silencing, cells were exposed with 50 μmol/L didymin. After 48-hour incubation, MTT assays as well as Western blot analyses for RKIP and N-Myc expression were done as described previously (18–20).

Cell survival was evaluated using a standard colony-forming assay. A total of 1 ×10⁵ cells/mL were incubated with 1 of the 3 drugs (didymin, hesperidin, 2HF-50 μmol/L each) for 48 hours, and aliquots of 50 or 100 μL were added to 60-mm size Petri dishes containing 4 mL culture medium. After 10 days, adherent colonies were fixed, stained with 0.5% methylene blue for 30 minutes, and counted using the Innotech Alpha Imager HP (21).

Detection of active caspase-3 in live cell cultures was carried out using an APO LOGIX carboxyfluorescein (FAM) Caspase Detection Kit (Cell Technology). The kit detects active caspase-3 in living cells through FAM-labeled DEVD fluoromethyl ketone (FMK), which irreversibly binds to active caspase-3. The inhibitor is cell permeable and noncytotoxic. The assay was carried out according to manufacturer's instructions (22).

For measurement of apoptosis by TUNEL assay, 1 × 10⁵ cells were grown on the cover slips for approximately 12 hours followed by treatment with test compounds (didymin, hesperidin, and 2HF 50 μmol/L each) for 24 hours. Apoptosis was determined by the labeling of DNA fragments with TUNEL using Promega fluorescence apoptosis detection system. Slides were analyzed under a fluorescence microscope using a standard fluorescein filter set to view the green fluorescence at 520 nm and red fluorescence of propidium iodide at more than 620 nm (17).

Cell migration was determined using a wound healing assay (23). A total of 2 × 10⁴ CHLA-90 and SMS-KCNR cells were seeded in 6-well plates to reach 100% confluence within 24 hours and then treated with 10 μmol/L mitomycin C for 2 hours. Subsequently, a similarly sized scratch was made with a 200-μL pipette tip across the center of each well and immediately imaged at baseline and then at 24 hours in control- and didymin-treated groups by using an Olympus Provis AX70 microscope. The rate of cell migration was determined by comparing the sizes of scratch area using ImageJ software.

The control and 50 μmol/L didymin-treated (at 24 hours) cells were lysed and analyzed by Western blot analyses for PARP cleavage, phosphoinositide 3-kinase (PI3K; Y^458/199), pAkt (S⁴⁷³), AKT, cyclin D1, cyclin B1, CDK4, N-Myc, RKIP, and vimentin by using specific antibodies. Briefly, lysates containing approximately 50 μg of proteins were subjected to SDS-PAGE and proteins were transferred onto nitrocellulose membrane. After blocking with 5% non-fat dry milk, the membrane was incubated overnight with the desired primary antibody (1:1,000 dilution). Subsequently, the membrane was incubated with appropriate secondary antibody for 2 hours, and the immune-reactive bands were visualized using the Enhanced Chemiluminescence Kit from Perkin-Elmer according to the manufacturer's instructions. The same membrane was reprobed with the antibody against GAPDH (1:5,000 dilution) as an internal control for equal protein loading. The intensity of immune-reactive bands on Western blots was determined using a densitometer (Molecular Dynamics) equipped with Image QuaNT software.

A total of 2 × 10⁵ cells were treated with didymin (50 μmol/L) for 18 hours at 37°C. After treatment, floating and adherent cells were collected, washed with PBS, and fixed with 70% ethanol. On the day of flow analysis, cell suspensions were centrifuged; counted, and same numbers of cells were resuspended in 500 μL PBS in flow cytometry tubes. Cells were then incubated with 2.5 μL of RNase (stock 20 mg/mL) at 37°C for 30 minutes after which they were treated with 10 μL of propidium iodide (stock 1 mg/mL) solution and then incubated at room temperature for 30 minutes in the dark. The stained cells were analyzed using the Beckman Coulter Cytomics FC500, Flow Cytometry Analyzer. Results were processed using CXP2.2 analysis software from Beckman Coulter (17).

Expression of N-Myc mRNA in neuroblastomas was evaluated by reverse transcriptase PCR (RT-PCR) analysis. The RNA from control and experimental groups was prepared 24 hours after didymin treatment using an Rneasy Kit (Qiagen) was quantified and 2 μg RNA was subjected to 1% agarose gel electrophoresis in MOPS-formaldehyde buffer. N-Myc gene-specific primers were used for RT-PCR (Biosynthesis Inc.; ref. 24). RT-PCR one step kit was used according to the manufacturer's instructions (Qiagen). Level of N-Myc protein in control- and didymin-treated cells were also measured by immunoassay using anti-NMyc IgG. Aliquots of crude cell extracts containing approximately 50 μg protein were applied to SDS-PAGE and Western blot analyses were done.

CHLA-90 and SMS-KCNR cells were cultured as adherent monolayers in a 96-well plate at a density of 2 × 10⁴ cells per well. The Firefly luciferase reporter construct (0.4 μg, pEB-Luc-N-Myc), kindly gifted by Prof. William L. Carroll, New York University Medical Center, NY, was cotransfected with 100 ng of Renilla luciferase construct (thymidine kinase promoter–driven Renilla luciferase; pTK-RL) by Lipofectamine Reagent according to the manufacturer's instructions (Invitrogen). After 24 hours of incubation, transfected cells were treated with 50 μmol/L of didymin. At 24-hour posttreatment, promoter activity was measured as luciferase activity by mixing cells with 50 μL of passive lysis buffer (Promega), followed by 15 minutes of shaking at room temperature. On a luminometer plate, 20 μL of the lysate was then mixed with 100 μL of luciferase assay reagent followed by 100 μL of Stop N' Glo Buffer (Promega). Luciferase activity was then determined as a ratio of Firefly to Renilla luciferase. Renilla luciferase activity was thus used as an internal control.

Mice (n = 3 each for control and didymin treatment) were administered either 0.1 mL corn oil or 50 μg didymin/0.1 mL corn oil/mice (2 mg/kg b.w.) by oral gavage on alternate days for 8 weeks. On the last day, the blood was collected within 1 hour after final dosage. The 100 μL of serum was subjected to protein precipitation using 200 μL of methanol. The sample was centrifuged at 13,300 rpm for 15 minutes at 4°C and the clear supernatant was lyophilized. The resulting pellet was dissolved in 200 μL of methanol: 0.2% phosphoric acid buffer (58:42 v/v). The 25 μL of each sample was subjected to high-performance liquid chromatography (HPLC) analysis using C₁₈ reverse-phase column at wavelength of 295 nm on Agilent 1100 system with auto injector (Agilent Technologies). The mobile phase consisting of acetonitrile (ACN) and water at a flow rate of 1.0 mL/min with following gradient: 0–1 minute: ACN-10%, H₂O-90%; 1–2 minutes: gradually increasing to ACN-20%, H₂O-80%; 2–14 minutes: gradually changing to ACN-90%, H₂O-10%; 14–16 minutes: ACN-10%, H₂O-90%.

Hsd: Athymic nude nu/nu mice were obtained from Harlan. All animal experiments were carried out in accordance with a protocol approved by the Institutional Animal Care and Use Committee (IACUC). The 11-week-old mice (n = 10) were divided into 2 groups of 5 animals [(For treatment with (i) corn oil (vehicle), (ii) didymin (2 mg/kg b.w.)]. The animals were injected with 2 × 10⁶ SMS-KCNR neuroblastoma cell suspensions in 100 μL of PBS, subcutaneously into one flank of each nu/nu nude mouse. At the same time, animals were randomized into treatment groups as indicated in Figure 5. Treatment was started 10 days after the neuroblastoma cells implantation to see palpable tumor growth. Treatment consisted of didymin (2 mg/kg b.w.) in 100 μL corn oil by oral gavage alternate day. Control groups were treated with 100 μL corn oil by oral gavage alternate day. Animals were examined daily for signs of tumor growth. Tumors were measured in 2 dimensions using calipers. Photographs of animals were taken at days 1, 10, 20, 40, and 60 after subcutaneous injection, are shown for all groups. Photographs of tumors were also taken at day 60.

Tumors from SMS-KCNR neuroblastoma mice xenografts (control and didymin treated) were harvested on day 60. Tumor samples fixed in buffered formalin for 12 hours were processed conventionally for paraffin-embedded tumor sections (5-μm thick). Histopathologic analyses with ki67, anti-CD31, and N-Myc were carried out using Universal ABC Detection kit (Vector). The sections were examined under Olympus Provis AX70 microscope connected to a Nikon camera.

All data were evaluated with a 2-tailed unpaired Student t test and are expressed as mean ± SD. The statistical significance of differences between control and treatment groups was determined by ANOVA, followed by multiple comparison tests. Changes in tumor size and body weight during the course of the experiments were visualized by scatter plot. Differences were considered statistically significant when the P value was less than 0.05.

The cell survival study by MTT assay revealed that didymin is the most potent of the 3 flavonoids tested in 4 neuroblastomas cell lines [CHLA-90 and SK-N-BE2 (p53-mutant) as well as SMS-KCNR and LAN-5 (p53 wild-type)]. The other flavonoid hesperidin was less effective and 2HF was not much effective against the 4 neuroblastoma cell lines tested. The 50 μmol/L didymin caused approximately 75% cell death, whereas hesperidin induced only approximately 35% cell death and 2HF induced approximately 10% cell death in all tested neuroblastoma cell lines at similar concentrations. Didymin did not cause any cytotoxicity in normal human umbilical vein endothelial cells (HUVEC) in MTT assay (Fig. 1A). The ability of didymin to arrest the growth of neuroblastoma tumors was further evident in colony-forming assay in which 50 μmol/L didymin caused approximately 70% growth inhibition, as compared with hesperidin (approximately 30% growth inhibition) and 2HF (approximately 10% growth inhibition; Fig. 1B). Thus, didymin decreased both survival and clonogenic potential to a greater extent than hesperidin and 2HF in neuroblastomas, as revealed by MTT and colony-forming assays, respectively. The effect of didymin on apoptosis was further studied by TUNEL apoptosis assay. The didymin treatment induced enhanced cell death relative to hesperidin and 2HF in neuroblastoma, which was evident by increased green fluorescence, a measure of DNA strand breaks induced by apoptosis in TUNEL apoptotic assay (Fig. 2A; ref. 25). Didymin also effectively inhibited the migration of neuroblastoma cells in vitro wound healing assay relative to hesperidin and 2HF (Fig. 2B). These findings collectively confirmed the enhanced cytotoxic potential of didymin in targeting neuroblastomas relative to hesperidin and 2HF. Following initial assessment of the potent anticancer properties of didymin in neuroblastomas, we next carried out detailed mechanistic studies in CHLA-90 (p53 mutant) and SMS-KCNR (p53 wild type) neuroblastoma cells.

The didymin treatment increased caspase-3 activation in in situ caspase-3 cleavage assay as revealed by enhanced green fluorescence in the treated group compared with controls (Fig. 2C). Western blot revealed the enhanced PARP cleavage consequent to didymin treatment in neuroblastomas. Didymin treatment also decreased the levels of PI3K, phosphorylation, and protein levels of Akt. Didymin also effectively reduced the levels of vimentin, a major intermediate filament protein which regulates epithelial–mesenchymal transition, an essential process during oncogenic transformation as well as metastasis (Fig. 2D). These results provided the mechanistic rationale for the decreased survival and clonogenic potential induced by didymin, as revealed by our studies by revealing the inhibition of PI3K and pAkt by didymin. The activation of caspase-3 and enhanced PARP cleavage further validated the apoptosis observed by TUNEL apoptosis assay following didymin treatment. The expression of vimentin is associated with enhanced motility of tumor cells and hence inhibition of vimentin by didymin was significant in the context of antimigratory effects of didymin in neuroblastomas as revealed by in vitro migration assay (26, 27). Thus, these results provided further corroborative evidence for the observed antiproliferative and proapoptotic effects of didymin in initial cell survival, clonogenic, and apoptosis studies.

We next analyzed the effect of didymin on cell-cycle progression by fluorescence-activated cell sorting (FACS) analysis. Loss of p53 leads to decreased cell-cycle checkpoint regulation which favors enhanced proliferation of cells (28–30). Didymin caused significant G₂/M phase arrest in CHLA-90 (p53 mutant) and SMS-KCNR (p53 wild type) neuroblastoma cells, suggesting the inhibition of cell cycle as an additional mechanism for the antiproliferative effects of didymin (Fig. 3A). Western blot of didymin-treated neuroblastoma cells revealed decrease in cyclin B1, cyclin D1, and CDK4 (Fig. 3B). Cyclin D1 is a key G₂/M phase regulator whose increase during G₂ phase leads to progression to mitosis and hence the decrease of cyclin D1 by didymin impacts G₂/M phase progression (31–33). Another interesting finding was that we observed a decrease in the levels of CDK4, the kinase which is commonly associated with G₁ transition along with G₂/M phase arrest. CDK4 has multiple functions and has been also investigated for its role in G₂/M transition. It has been shown that overexpression of dominant-negative CDK4 leads to arrest of G₂ phase progression (33). Some of the anticancer compounds such as apigenin and thimerosal also cause inhibition of CDK4 along with cyclin B1, while causing G₂/M phase arrest (33–35). Cyclin B1, the regulatory subunit of cdc2 kinase, is required for mitotic phase initiation. Cyclin B1 is localized to kinetochores during cell division and regulates the alignment of chromosomes during mitosis (36). The p53 activation leads to G₂/M phase arrest by decreasing the levels of cyclin B1 levels as well as inhibiting the activity of cyclin B1 promoter, whereas the activation of cyclin B1 leads to cell-cycle progression even in the presence of p53 activation (37, 38). Thus, cyclin B1 is a critical requirement and mediator of G₂/M phase progression downstream of p53. In this context, the ability of didymin to downregulate cyclin B1 in both p53 mutant and p53 wild type neuroblastomas represents a potentially significant mechanism of action for the observed antiproliferative effects of didymin in neuroblastomas, irrespective of p53 status.

The amplification of N-Myc contributes both to pathogenesis and aggressive progression of neuroblastomas, giving them a highly angiogenic phenotype (5, 39). Hence, we analyzed the impact of didymin on N-Myc expression. The didymin treatment decreased the levels of N-Myc protein in a time- and concentration-dependent manner as shown by Western blot analyses (Fig. 3C). The 50 μmol/L didymin treatment induced approximately 5-fold reduction in N-Myc mRNA levels as revealed by RT-PCR using N-Myc gene-specific primers (Fig. 3D). We further explored whether didymin regulates N-Myc promoter activity by luciferase promoter–reporter assays. The Firefly luciferase reporter construct (0.4 μg pEB-Luc-N-Myc) was cotransfected with 100 ng of Renilla luciferase plasmid construct (thymidine kinase promoter–driven Renilla luciferase; pTK-RL) by Lipofectamine reagent. After 24 hours of incubation, the transfected cells were treated with 50 μmol/L of didymin, further incubated for 24 hours, and the luciferase assay was done. Didymin treatment induced significant decrease in the activity of N-Myc promoter compared with controls (Fig. 4A).

PI3K inhibition mediated downregulation of N-Myc protein levels has been reported (40). Didymin caused significant inhibition of PI3K in neuroblastomas (Fig. 2D). RKIP can inhibit MAPK and Myc signaling (41–43). Interestingly, RKIP overexpression is also known to positively induce normal neuronal differentiation in neuroblastoma (44). Hence, we examined the effect of didymin on RKIP expression. Didymin significantly enhanced the levels of RKIP in both p53 mutant (CHLA-90) and p53 wild type (SMS-KCNR) neuroblastomas (Fig. 4B). Given the significant role of RKIP in tumor signaling, we further studied the impact of knockdown of RKIP in SMS-KCNR and CHLA-90 neuroblastoma cells. The MTT assay following knockdown of RKIP by siRNA treatment revealed partial reversal of the didymin-induced inhibition of neuroblastoma cell survival (data not shown). The knockdown of RKIP was also associated with an increase in the expression of N-Myc in neuroblastoma cells. The knockdown of RKIP also caused a partial reversal in the didymin-induced decrease of N-Myc expression as detected by Western blot analyses (Fig. 4C). This observed inhibitory effect of RKIP on N-Myc expression is in accordance with previous studies implicating the role of RKIP in regulation of Myc expression (42). Hence, RKIP upregulation, along with N-Myc repression, represents a potent mechanism of action of didymin of specific relevance in inducing its anticancer effects in neuroblastomas. The collective antiproliferative, antimigratory, and proapoptotic effects of didymin in vitro, along with its ability to inhibit N-Myc, PI3K/Akt, and MAPK signaling pathways which play a significant mechanistic role in the incidence, progression, and clinical refractoriness of neuroblastomas, provided strong rationale for the further investigation of didymin in in vivo mice xenograft studies.

We first assessed the absorption of orally administered didymin in mice. HPLC analysis of didymin-treated mice serum revealed that didymin is effectively absorbed after oral dosage and it reaches a serum concentration of 2.1 μmol/L (Fig. 4D). It is important to note that we quantified only didymin, but not its metabolites, to address whether didymin, a relatively new flavonoid in cancer research, is absorbed following oral administration.

We next assessed the regression of neuroblastoma mice xenografts following oral administration of didymin. The mice harboring SMS-KCNR neuroblastoma xenografts were treated with 50 μg/mice (equivalent to 2 mg/kg b.w. or 0.0002% w/w) of didymin in 100 μL corn oil alternate day for 60 days by oral gavage. Control group was treated with 100 μL corn oil. Treatment was started 10 days post neuroblastoma cells implantation after a palpable tumor growth was observed. Photographs of animals were taken at days 1, 10, 20, and 40 after subcutaneous injection are shown for both groups. Tumors were dissected at day 60, photographed and weighed. Didymin-treated animals had significant reductions in tumors as compared with control, whereas uncontrolled growth was observed in the control groups. The didymin-treated mice were active and weight gain was comparable with nontumor–bearing controls, and no overt toxicity was evident (Fig. 5A). The final tumor weight on day 60 was 0.91 g in didymin-treated group compared with 1.97 g in controls. Present studies, for the first time, showed sustained regression of xenografts of neuroblastomas by didymin (Fig. 5B and C).

The effect of didymin treatment on the neuroblastoma progression markers in resected mice xenograft sections was further examined. The histopathologic examination of paraffin-embedded tumor xenograft sections revealed the decreased levels of proliferation marker, Ki67, angiogenesis marker, CD31, and N-Myc expression in SMS-KCNR neuroblastoma after didymin treatment, which further supported our in vitro results (Fig. 5D).

Until today, there are no safe dietary choices to prevent the incidence and progression of neuroblastomas in children. The use of chemotherapeutic drugs, radiation, and surgical approaches are often coupled with recurrent and diverse clinical challenges such as limitations of young age, as well as amplification of off-target cytotoxicity on developing nervous and skeletal systems, over the course of time following initial therapy in children. In this regard, our current studies, for the first time, have revealed that the novel flavonoid didymin is effective against neuroblastomas both in vitro cell cultures and in vivo mice xenograft studies. Didymin effectively inhibited the proliferation of neuroblastomas and induced apoptosis by inhibiting PI3K, Akt, and increasing RKIP levels. Didymin also decreased the levels of vimentin, a marker of epithelial–mesenchymal transition, and decreased the migration of neuroblastoma cells in vitro. Importantly, didymin induced G₂/M cell-cycle arrest which was associated with decrease in the levels of cyclin B1, G₂/M phase regulator downstream of p53, and whose overexpression can override p53-induced G₂/M phase arrest (35). The downregulation of cyclin B1 explains one of the mechanisms responsible for the antiproliferative effect of didymin in both clinically sensitive p53-wild type and refractory p53 mutant neuroblastomas. Didymin did not cause any cytotoxicity in normal HUVEC cells as well as any overt toxicity in mice. Didymin strongly inhibited the proliferation (Ki67) and angiogenesis (CD31) markers as well as N-Myc expression in vivo mice xenograft studies.

Mechanistically, the inhibition of N-Myc by didymin, as elaborately confirmed at protein, mRNA, and promoter–reporter assays, was a hallmark of our studies. Amplification and overexpression of N-Myc is associated with both the incidence and refractoriness of neuroblastomas (5, 36). Hence, the ability of didymin to inhibit N-Myc at both transcriptional and translational levels reflects the significance of didymin toward developing novel interventional strategies for neuroblastomas. The nonbitter nature of didymin saves the time and effort spent on the investigations of taste-modifying additives for pediatric diets. As didymin is a highly palatable flavonoid, it can be possibly administered in milk or other liquid pediatric diets in the infants and adolescent children, who are usually not compatible with derivatized preparations such as pills and creams. Thus, didymin has both fundamental mechanistic and clinical as well as translational significance due to its potential to regulate critical nodes of neuroblastoma signaling which reflects its relevance to effectively prevent the incidence and progression of neuroblastomas (Fig. 6).

In summary, this study provides strong evidence for the “N-Myc inhibitory,” “antiproliferative,” and “antiangiogenic” properties of didymin with “no overt toxicity to normal cells and tissues” while characterizing potent anticancer effects in vitro cell cultures and in vivo mice xenograft models of neuroblastomas. This multispecific anticancer potential of didymin, which is of specific relevance to both the pathogenesis and progression of neuroblastomas, makes didymin a novel and highly relevant dietary agent for the treatment of neuroblastomas. Our studies also indicate that didymin is absorbed effectively following oral administration. In this context, the further studies focused on detailed absorption, distribution, metabolism, and excretion (ADME) kinetics of didymin should help to develop and test appropriate doses and formulations of didymin in milk and pediatric diets for use in children till 5 years, the high-risk window for the incidence of neuroblastomas. Collectively, didymin represents a novel and highly promising flavonoid with potential clinical significance to effectively prevent the incidence of neuroblastomas and also as an innovative approach for strategies proposed by new approaches for neuroblastoma therapy (NANT) consortium.

No potential conflicts of interest were disclosed.

The authors thank Dr. Sumihiro Suzuki, Department of Biostatistics, School of Public Health, UNTHSC, Fort Worth, TX, for his assistance in the statistical analyses of the data.

This work was supported in part by USPHS grant CA 77495, the Cancer Research Foundation of North Texas, and Institute for Cancer Research & the Joe & Jessie Crump Fund for Medical Education.

The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.