Investigation of the activity of a novel tropolone in osteosarcoma

Abstract

Osteosarcoma is a primary malignant bone tumor characterized by frequent metastasis, rapid disease progression, and a high rate of mortality. Treatment options for OS have remained largely unchanged for decades, consisting primarily of cytotoxic chemotherapy and surgery, thus necessitating the urgent need for novel therapies. Tropolones are naturally occurring seven-membered non-benzenoid aromatic compounds that possess anti-proliferative effects in a wide-array of cancer cell types. MO-OH-Nap is an α-substituted tropolone that has activity as an iron chelator. Here we demonstrate that MO-OH-Nap activates all three arms of the unfolded protein response (UPR) pathway and induces apoptosis in a panel of human OS cell lines. Co-incubation with ferric chloride or ammonium ferrous sulfate completely prevents the induction of apoptotic and UPR markers in MO-OH-Nap-treated OS cells. MO-OH-Nap upregulates transferrin receptor 1 (TFR1) protein levels, as well as TFR1, divalent metal transporter 1 (DMT1), iron-regulatory proteins (IRP1, IRP2), ferroportin (FPN), and zinc transporter 14 (ZIP14) transcript levels, demonstrating the impact of MO-OH-Nap on iron-homeostasis pathways in OS cells. Furthermore, MO-OH-Nap treatment restricts the migration and invasion of OS cells in vitro. Lastly, metabolomic profiling of MO-OH-Nap-treated OS cells revealed distinct changes in purine and pyrimidine metabolism. Collectively, we demonstrate that MO-OH-Nap-induced cytotoxic effects in OS cells are dependent on the tropolone’s ability to alter cellular iron availability and that this agent exploits key metabolic pathways. These studies support further evaluation of MO-OH-Nap as a novel treatment for OS.

Introduction.

Osteosarcoma (OS) is the most common primary bone cancer in children, and it is the second leading cause of cancer-related deaths in adolescents (1). OS primarily occurs in the metaphysis of long bones and is most often found in the proximal tibia, proximal humerus, or proximal/distal femur. Initial treatment consists of multi-agent cytotoxic chemotherapy and surgery. Since the introduction of systemic chemotherapy, five-year survival rates for localized disease have largely plateaued at around 70% (2). OS cells have a high propensity to metastasize, and metastatic lesions can be found in almost any organ, with the most common being lungs, followed less frequently by the bone and lymph nodes. The five-year survival rate for patients with distant metastatic disease is a dismal 29% (3). Although research efforts have greatly expanded over the past decades, the development of targeted agents and immunotherapies for OS have not come to fruition. Unfortunately, patients who either do not reach remission or subsequently experience relapse have limited treatment options and poor prognosis. Thus, there is an urgent need for novel therapies for OS.

Tropolones are seven-membered non-benzenoid aromatic compounds derived from plant monoterpenes (4). Unlike many natural products, tropolones are small and possess a simple backbone that allows for extensive structural modification, making them ideal scaffolds for drug development. Several tropolones demonstrate anti-proliferative effects in cancer cell lines, leading many researchers to investigate the use of tropolones as anti-cancer agents (5–8). For instance, β-thujaplicin, a representative and well-studied tropolone, displays anti-cancer activity in several mouse tumor models, including breast cancer, melanoma, and lung cancer (9–11). Tropolones act as divalent cation chelators (8), and thus it has been suggested they may inhibit enzymes which utilize zinc as a co-factor, including the histone deacetylases (HDAC) (12).

MO-OH-Nap is an α-substituted tropolone (Fig. 1a) that potently induces apoptosis in multiple myeloma (MM) cells (13). The cytotoxic effects of MO-OH-Nap in MM cells are attributed to the drugs ability to activate the unfolded protein response pathway (UPR), with subsequent induction of caspase-mediated cell death (13). Our previous studies indicate that inhibition of HDACs is not responsible for the cytotoxic effects of MO-OH-Nap in MM cells. Subsequent work revealed that co-incubation with exogenous iron completely prevented MO-OH-Nap-mediated induction of the UPR and apoptosis (14). MO-OH-Nap’s ability to chelate ferrous iron was confirmed with a ferrozine assay. Together, these studies demonstrated that MO-OH-Nap-induced cytotoxic effects in MM cells are dependent on the tropolone’s ability to alter cellular iron availability.

Figure 1. MO-OH-Nap induces apoptosis in OS cells.

(A) Chemical structure of MO-OH-Nap (B) MTT assays were performed following a 72-hour incubation with MO-OH-Nap in four OS cell lines (n = 4, data are shown as mean ± standard deviation). (C) Annexin V and propidium iodide (PI) flow cytometric studies were performed following a 72-hour incubation with MO-OH-Nap (MOOH). Data are expressed as the average percentage of early apoptotic (Annexin V+/PI-) and late apoptotic (Annexin V+/PI+) (n = 3 biological replicates, data are displayed as mean ± standard deviation, *denotes p < 0.05 **denotes p < 0.01 ***denotes p < 0.001, two tailed t-test). (D) Immunoblot analysis of cleaved PARP (denoted by arrow) and cleaved caspases 3, 8, 9 in OS cells incubated in the presence or absence of MO-OH-Nap for 72 hours. β-tubulin is shown as a loading control. Immunoblots are representative of three independent experiments.

OS cells exploits various mechanisms to increase intracellular iron pools to promote proliferation (15). Therapeutic strategies targeting iron metabolic pathways for the treatment of OS have not been well studied. In the present study, we hypothesized that MO-OH-Nap would induce cytotoxic effects in OS cells. Our studies show that MO-OH-Nap mediates activation of all three arms of the UPR, as well as induction of caspase-mediated apoptosis in a variety of human OS cell lines. As predicted, MO-OH-Nap-mediated cytotoxicity was fully abrogated by the addition of exogenous ferric or ferrous iron. In addition, MO-OH-Nap altered the expression of key regulators of iron homeostasis, including transferrin receptor 1 (TFR1), divalent metal transporter 1 (DMT1), iron-regulatory proteins (IRP1, IRP2), ferroportin (FPN), and zinc transporter 14 (ZIP14). MO-OH-Nap treatment also inhibited cellular migration and invasion in vitro. Furthermore, using liquid chromatography-tandem mass spectrometry (LC-MS/MS) we identified key metabolic changes that occur due to MO-OH-Nap treatment. Collectively, our findings illustrate that MO-OH-Nap may serve as a novel treatment option for OS.

Material and Methods.

Cell culture.

MG-63, HOS, 143B, and SaOS-2 cell lines were purchased from ATCC (Manassas, Virginia, USA), while CAL-72 cells were acquired from DSMZ (Braunschweig, Germany). Cells were grown in media (MEM for 143B, MG-63, HOS; McCoys 5A for SaOS-2; DMEM for CAL-72) supplemented with heat-inactivated fetal bovine serum (10% for MG-63, HOS, 143B, CAL-72; 15% for SaOS-2), BrDU (143B), insulin (CAL-72) and penicillin-streptomycin at 37°C and 5% CO2. MycoAlert mycoplasma detection kit (Lonza, Basel, Switzerland) was utilized to test for mycoplasma contamination.

Chemicals.

MO-OH-Nap (purity ≥ 95% by high-performance liquid chromatography and confirmed by nuclear magnetic resonance) was synthesized as previously described and kindly provided by Dr. Dennis Wright, University of Connecticut (6, 12). Stock solutions of MO-OH-Nap were prepared in DMSO (10 mM) and stored at −20°C. Ammonium ferrous sulfate (AFS) and ferric chloride (FC) were purchased from Sigma-Aldrich (St. Louis, MO, USA).

MTT.

OS cells were plated in 96-well plates and incubated in the presence or absence of MO-OH-Nap for 48 or 72 hours (5,000 cells/100 μL, not serum starved). In selected experiments, AFS was added concurrently with MO-OH-Nap. MTT (3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyl-2 assays were performed as previously described (16). Data were normalized to that of solvent treated cells, which were defined as having an MTT activity of 100%.

Immunoblot.

Cells were incubated in the presence or absence of MO-OH-Nap for 24, 48, or 72 hours. AFS or FC (50 μM) was added concurrently with MO-OH-Nap or after 24 hours (sequential treatment). Cells were washed with phosphate-buffered saline (PBS) and lysed in RIPA buffer (1% sodium deoxycholate, 0.15 M NaCl, 0.1% SDS, 1% (v/v) Triton X-100, 0.05 M Tris HCl, pH 7.4) supplemented with protease and phosphatase inhibitors. Protein concentration was quantified using the BCA method. Protein (15 μg) was run on an SDS-PAGE gel and transferred to a polyvinylidene difluoride membrane. Blots were incubated in primary antibody overnight at 4°C and in secondary antibody for 1 hour at room temperature. Blots were imaged using a chemiluminescence detection kit and a Bio-Rad Chemidoc MP imaging system. Antibodies used for these studies are listed in Supplemental Table 1.

Flow cytometry.

Cells were incubated in the presence or absence of MO-OH-Nap for 48 or 72 hours. Cells were processed for Annexin V/Propidium iodide staining using the eBioscience Annexin V staining kit per the manufacturer’s instruction. Early apoptotic cells are defined as Annexin V+/Propidium iodide- and late apoptotic as Annexin V+/Propidium iodide+. For TFR1 cell surface detection, cells were incubated in PBS supplemented with TFR-1 antibody at a concentration of 1 μg per 1 × 10⁶ cells for 15 minutes. Cells were washed twice with PBS before being incubated with secondary antibody for 15 minutes (Alexa fluor 488 donkey anti-mouse, Invitrogen, Waltham, MA, USA). For all flow cytometry experiments, ten-thousand cell events were recorded on a BD LSRII flow cytometer. Data analysis was performed using FlowJo software. All experiments were performed in triplicate.

qRT-PCR.

Cells were incubated for 24 or 48 hours in the presence or absence of MO-OH-Nap. In selected experiments, AFS or FC (50 μM) was added concurrently with MO-OH-Nap. RNA was isolated using the E.Z.N.A. HP total RNA kit from Omega (Norcross, GA, USA) and 1 μg of total RNA was reverse transcribed to cDNA using the i-Script cDNA synthesis kit (Bio-Rad, Hercules, CA, USA). cDNA, gene specific primers, and i-Taq Sybr green super mix (Bio-Rad) were mixed according to manufacturer’s instruction. qRT-PCR reactions were performed in triplicate in a CFX96 real time machine (Bio-Rad), and data was analyzed using the Bio-Rad CFX manager 3.1 software. Expression values were normalized to the house-keeping gene β-actin. Primer sequences used in these studies can be found in Supplemental Table 2.

Migration and Invasion assays.

All assays were performed as described previously (17). Cells were plated in serum-free media in the top compartment of 8.0-μm pore size trans-well chambers (Corning, Corning, NY, USA). For invasions assays, membranes were coated with 50 μL of matrigel (Corning; diluted 1:10 in serum free media) and allowed to solidify for 2 hours at 37°C. The bottom chambers were filled with complete media to serve as a chemoattractant. Cells were incubated in the presence or absence of MO-OH-Nap for 24 hours. At the end of the incubation period, cells on the top side of the membrane were removed with a cotton swap. Cells on the underside of the membrane were fixed and stained using the VWR hematology quick stain three step solution kit (Radnor, PA, USA). Cells were quantified using a light microscope (200X magnification) and by counting five random fields of view per membrane. Experiments were performed in triplicate.

Metabolomics Analysis.

Metabolic profiling of primary metabolites (123 metabolites) was performed using a LC-MS/MS 8060 system (Shimadzu Scientific, Columbia, Maryland, USA), equipped with an electrospray ionization source operated in both positive and negative mode. All LC-MS/MS experiments were performed using previously described methods (18, 19). The targeted metabolomics explored the changes of primary metabolites in OS cells incubated in the presence or absence of MO-OH-Nap (1 μM and 10 μM) for 48 hours in SaOS-2 and 143b cells. The metabolomics and pathway data analysis were performed by the software MetaboAnalyst 5.0 (https://www.metaboanalyst.ca/). Metabolomics data is available in supplemental file 2.

Statistical Analysis.

T-tests (two-tailed) were used to calculate statistical significance between control and treated groups for flow cytometry and qRT-PCR experiments. Metabolomics data was analyzed using the MetaboAnalyst 5.0 software. Univariate analysis method (t-test) were used for two-group datasets to calculate fold changes (FC) and p-value. For hierarchical clustering and heatmaps, distance was measured using Euclidean, and clustering algorithm using ward.D. CompuSyn software was used to analyze MTT data and calculate IC₅₀ values.

Results.

MO-OH-Nap induces apoptosis in OS cells.

As an initial assessment of the effects of MO-OH-Nap in OS cells, we performed MTT assays in five OS human cell lines (143B, CAL-72, HOS, MG-63 and SaOS-2). To varying degrees, 72-hour incubation with MO-OH-Nap induced concentration-dependent cytotoxicity in all five cell lines, with HOS being the most sensitive (IC₅₀ = 0.67 μM) and SaOS-2 being the least sensitive and (IC₅₀ = 5.93 μM, Fig. 1b, Table 1). Likewise, 48-hour incubation with MO-OH-Nap promoted cytotoxicity in OS cell lines in a concentration-dependent manner (Supplemental Fig. 1, Table 1). Annexin V and propidium iodide flow cytometric studies showed that MO-OH-Nap induces both early- and late-stage apoptosis at 48 and 72 hours in OS cells (Fig. 1c, Supplemental Fig. 2). To more directly assess the ability of MO-OH-Nap to induce apoptosis in OS cells, immunoblot analysis was performed to detect poly-ADP-ribose polymerase (PARP) cleavage and accumulation of cleaved caspases. All five cell lines show PARP cleavage and accumulation of at least two cleaved caspases at both 48- and 72-hour timepoints (Fig. 1d, Supplemental Fig. 3). Four out of five cell lines show accumulation of cleaved PARP and caspases 3 and 8 following a 24-hour incubation with 10 μM MO-OH-Nap (Supplemental Fig. 3). Collectively, these results demonstrate that MO-OH-Nap induces apoptotic cell death in a variety of human OS cell lines.

Table 1.

IC₅₀ values from the MTT cytotoxicity assays with MO-OH-Nap.

	IC50 (μM)
Cell line	48h	72h
143B	1.68	0.97
CAL-72	>10	3.38
HOS	0.81	0.67
MG-63	5.88	3.19
SaOS-2	>10	5.93

MO-OH-Nap activates the unfolded protein response pathway in OS cells.

To evaluate if MO-OH-Nap can induce ER stress in OS cells, we performed qRT-PCR analysis for UPR markers. Upregulation of ATF4, ATF6, CHOP, IRE1 and PERK transcript levels were observed in all tested cell lines following a 24-hour incubation with increasing concentrations of MO-OH-Nap (Fig. 2a). Likewise, we observed upregulation of ATF4, IRE1 and p-eIF2α protein levels in all five cell lines, albeit at varying timepoints amongst the cell lines (Fig. 2b, Supplemental Fig. 4). Together, these data illustrate that MO-OH-Nap induces activation of all three arms of the UPR pathway in OS cells.

Figure 2. MO-OH-Nap activates the unfolded protein response pathway in OS cells.

(A) qRT-PCR analysis of UPR markers in OS cells incubated in the presence or absence of MO-OH-Nap (MOOH) for 24 hours. Expression values were normalized to the house-keeping gene β-actin. Data are normalized so that control (DMSO treated) is equal to one (n = 3 biological replicates, data are displayed as mean ± standard deviation, *denotes p < 0.05 **denotes p < 0.01 ***denotes p < 0.001, two tailed t-test). (B) Immunoblot analysis of UPR markers in OS incubated in the presence or absence of MO-OH-Nap for 72 hours. β-tubulin is shown as a loading control. Immunoblots are representative of three independent experiments.

Exogenous iron abrogates the cytotoxic effects of MO-OH-Nap.

Previous work in MM cell lines demonstrated that MO-OH-Nap has activity as an iron chelator and that addition of exogenous iron reverses its cytotoxic effects (14). To determine whether effects on iron homeostasis underlie MO-OH-Nap-induced cytotoxicity in OS cells, we performed studies in which cells were incubated with MO-OH-Nap and iron. We found that the addition of ferrous iron prevents MO-OH-Nap-mediated cytotoxicity as demonstrated by MTT assays (Supplemental Figs. 5 and 6). Likewise, co-incubation with iron abrogates the induction of caspase and PARP cleavage in MO-OH-Nap-treated OS cells at 24, 48 and 72 hours (Fig. 3a and Supplemental Fig. 7). Furthermore, the addition of iron prevented MO-OH-Nap-mediated upregulation of UPR markers at both the level of protein (Supplemental Fig. 8) and gene expression (Fig. 3b, Supplemental Fig. 9). Conversely, the sequential addition of iron 24 hours after the start of MO-OH-Nap treatment only partially prevents the induction of apoptotic/UPR markers in SaOS-2 cells, but not MG-63 cells (Supplemental Fig. 10). Consistent with our previous studies, these results demonstrate that the ability of MO-OH-Nap to activate the UPR and induce apoptosis in OS cells is dependent on the ability of the drug to alter cellular iron availability.

Figure 3. Exogenous iron abrogates the cytotoxic effects of MO-OH-Nap.

(A) Immunoblot analysis of apoptotic markers in OS cells incubated in the presence or absence of MO-OH-Nap (MOOH) for 72 hours. 50 μM ferric chloride (FC) or ammonium ferrous sulfate (AFS) was added concurrently with MO-OH-Nap. β-tubulin is shown as a loading control. Immunoblots are representative of three independent experiments. (B) qRT-PCR analysis of UPR markers in OS cells following a 48-hour incubation with MO-OH-Nap (10 μM). AFS or FC were added concurrently with MO-OH-Nap at a concentration of 50 μM. Expression values were normalized to the house-keeping gene β-actin. Data are normalized so that control (DMSO treated) is equal to one (n = 3 biological replicates, data are displayed as mean ± standard deviation, *denotes p < 0.05 **denotes p < 0.01 ***denotes p < 0.001, two tailed t-test).

MO-OH-Nap effects iron handling pathways.

To understand how MO-OH-Nap may alter intracellular iron homeostasis, we investigated the effects of MO-OH-Nap on the expression of TFR1, DMT1, IRP1, IRP2, FPN, and ZIP14. TFR1 enables the uptake of iron at the cell surface by internalizing iron-bound transferrin and is considered the primary regulator of iron uptake for the cell. Incubation with MO-OH-Nap induced upregulation in TFR1 protein levels in a concentration-dependent manner in all five OS cell lines as early as 24-hours (Fig. 4a). Likewise, MO-OH-Nap induced an increase in TFR1 surface protein as measured by flow cytometry (Fig. 4b). qRT-PCR analysis of key regulators of iron-homeostasis (TFR1, DMT1, IRP1, IRP2, FPN, and ZIP14) revealed upregulation in all six genes following a 24-hour incubation with MO-OH-Nap (Fig. 4c). Furthermore, 48-hour co-incubation with either ferric or ferrous iron prevents MO-OH-Nap-induced accumulation of TFR1 protein levels (Fig. 5a). Likewise, the addition of exogenous iron prevents MO-OH-Nap-mediated upregulation of TFR1, DMT1, IRP1, IRP2, FPN, and ZIP14 at the level of gene expression (Fig. 5b). The sequential addition of iron 24 hours after the start of MO-OH-Nap treatment only partially prevents the induction of TFR1 protein levels in SaOS-2 and MG-63 cells (Fig. 5c). Collectively, these results suggest that MO-OH-Nap alters cellular iron pools, leading to compensatory upregulation in genes that mediate iron homeostasis.

Figure 4. MO-OH-Nap affects iron handling pathways.

(A) Immunoblot analysis of transferrin receptor (TFR1) in OS cells following a 24-, 48-, and 72-hour incubation with MO-OH-Nap (MOOH). β-tubulin is shown as a loading control. Immunoblots are representative of three independent experiments. (B) Flow cytometric analysis of TFR1 cell surface levels in OS cells following a 24-, 48-, or 72-hour incubation with MO-OH-Nap (n = 3 biological replicates, data are displayed as normalized median fluorescent intensity ± standard deviation, *denotes p < 0.05. **denotes p < 0.01 ***denotes p < 0.001, two tailed t-test). (C) qRT-PCR analysis of TFR1, DMT1, FPN, IRP1, IRP2, and ZIP14 in OS cells following a 24-hour incubation with MO-OH-Nap. Expression values are normalized to the house-keeping gene β-actin. Data are normalized so that control (DMSO treated) is equal to one (n = 3 biological replicates, data are displayed as mean ± standard deviation, *denotes p < 0.05 **denotes p < 0.01 ***denotes p < 0.001, two tailed t-test).

Figure 5. MO-OH-Nap mediated activation of iron pathway genes is inhibited by the addition of exogenous iron.

(A) Immunoblot analysis of TFR1 in OS cells incubated in the presence or absence of 10 μM MO-OH-Nap (MOOH) with or without 50 μM ferric chloride (FC) or ammonium ferrous sulfate (AFS) for 48 hours. β-tubulin is shown as a loading control. Immunoblots are representative of three independent experiments. (B) RT-PCR analysis of TFR1, DMT1, FPN, IRP1, IRP2, and ZIP14 in OS cells in MG-63 and SaOS-2 cells following a 48-hour incubation with MO-OH-Nap (10 μM) with or without 50 μM FC or AFS. Expression values are normalized to the house-keeping gene β-actin. Data are normalized so that control (DMSO treated) is equal to one (n = 3 biological replicates, data are displayed as mean ± standard deviation, *denotes p < 0.05 **denotes p < 0.01 ***denotes p < 0.001, two tailed t-test). (C) Immunoblot analysis of TFR1 in OS cells incubated for 48 hours with MO-OH-Nap. Iron (50 μM) was added either concurrently or sequentially (24 hours after MO-OH-Nap addition, denoted with ‘*’).

MO-OH-Nap disrupts OS cell migration and invasion in vitro.

We performed migration and invasion assays in which OS cells were seeded on the top chamber of matrigel-coated (invasion) and non-coated (migration) trans-wells. Treatment with MO-OH-Nap for 24 hours significantly restricted the migration of 143B, MG-63 and SaOS-2 cells (Fig 6a). Likewise, MO-OH-Nap treatment disrupted invasion through matrigel in 143B and SaOS-2 cells (Fig 6b). In aggregate these data indicate that MO-OH-Nap treatment inhibits OS cell movement.

Figure 6. MO-OH-Nap disrupts OS cell migration and invasion in vitro.

Cells were seeded on (a) non-coated or (b) matrigel-coated 8.0-μM trans-wells and grown in the presence or absence of MO-OH-Nap (MOOH) for 24 hours. Cells on the bottom of the trans-wells were stained, imaged, and counted (n=5 fields per well). Data are normalized to control (n = 3 biological replicates, data are displayed as mean ± standard deviation, *denotes p < 0.05, **denotes p < 0.01, ***denotes p < 0.001, two tailed t-test).

MO-OH-Nap alters purine and pyrimidine metabolism.

To evaluate metabolic changes that occur as the result of MO-OH-Nap treatment, we simultaneously measured 123 metabolites using LC-MS/MS in SaOS-2 and 143B cell lines incubated in the presence or absence of MO-OH-Nap (1 μM and 10 μM) for 48 hours. Overall, 143B cells yielded the largest subset of differential metabolites. Relative to control cells, 143B cells treated with 1 μM and 10 μM MO-OH-Nap had 33 and 54 significantly altered metabolites, respectively (False discovery rate [FDR] < 0.05, Table 2, Supplemental Table 3). For SaOS-2 cells, treatment with 1 μM MO-OH-Nap yielded 2 significant changes, while 10 μM MO-OH-Nap producing 8 significant changes (Supplemental Table 4). Heatmaps with hierarchical clustering show perfect segregation of control and treatment samples for both 143B and SaOS-2 cells lines (Supplemental Figs. 11 and 12). Pathway analysis of the 54 metabolites deregulated in 10 μM MO-OH-Nap-treated 143B cells identified significant enrichment in purine metabolism (FDR<0.001, 15 metabolites, Table 2) and pyrimidine metabolism (FDR<0.001, 11 metabolites, Table 2). Purine metabolism was also one of the top deregulated pathways in 1 μM MO-OH-Nap-treated 143B cells (FDR<0.01, 7 metabolites, Supplemental Table 3). In addition, alanine, aspartate, and glutamate metabolism was shared among the top five enriched pathways in 1 μM and 10 μM MO-OH-Nap-treated 143B cells (Supplemental Tables 5 and 6). Taken together, these results demonstration disruption of several key metabolic pathways induced by MO-OH-Nap treatment in OS cell lines.

Table 2.

Metabolic changes in MO-OH-Nap treated 143B cells.

Metabolite	Fold Change (log2)	FDR
6-Phosphogluconic acid	2.93	0.009
Adenine †	−2.66	0.014
Adenosine †	−3.04	0.028
Adenosine 3’,5’-cyclic monophosphate †	−1.73	0.014
Adenosine diphosphate †	−1.72	0.015
Adenosine monophosphate †	−1.20	0.043
Adenosine triphosphate †	−1.81	0.042
Adenylsuccinic acid †	−5.65	0.009
Argininosuccinic acid	−2.91	0.015
Aspartic acid	−2.26	0.009
Citicoline	−4.12	0.009
Creatine	−1.49	0.015
Cystathionine	−2.91	0.009
Cytidine diphosphate ‡	0.59	0.022
Cytidine monophosphate ‡	−1.01	0.047
Cytidine triphosphate ‡	−1.82	0.027
Fructose 1,6-bisphosphate	−1.27	0.044
Fructose 6-phosphate	−1.37	0.011
Fumaric acid	−4.06	0.009
Glucose 1-phosphate	−1.28	0.016
Glutamic acid	−1.68	0.009
Glutamine † ‡	−1.47	0.029
Glutathione	−3.14	0.012
Glycerol 3-phosphate	2.28	0.006
Guanine †	−2.50	0.009
Guanosine †	−3.96	0.009
Guanosine monophosphate †	−2.75	0.016
Hypoxanthine †	1.71	0.030
Inosine †	−1.91	0.028
Lactic acid	−3.02	0.011
Lysine	−1.35	0.043
Malic acid	−3.50	0.009
methylsuccinic acid	−1.42	0.016
NAD	−2.67	0.009
NADP	−2.57	0.009
Nicotinamide ribotide	−2.56	0.009
Ornitine	−1.90	0.016
Oxidized glutathione	−3.26	0.011
Pantothenic acid	−1.90	0.021
Proline	−2.33	0.009
Pyruvic acid	−4.12	0.005
Ribose 5-phosphate †	1.86	0.009
Ribulose 5-phosphate	2.10	0.009
S-Adenosylhomocysteine	−2.96	0.009
Sedoheptulose 7-phosphate	3.62	0.014
Succinic acid	−2.12	0.010
Thymidine ‡	−1.89	0.016
Thymidine monophosphate ‡	2.40	0.009
Thymidine triphosphate ‡	−3.33	0.005
Uracil ‡	−3.51	0.009
Uridine ‡	−3.92	0.009
Uridine diphosphate ‡	−1.39	0.009
Uridine triphosphate ‡	−4.58	0.016
Xanthosine †	−3.62	0.009

Cells were incubated in the presence or absence of 10 μM MO-OH-Nap for 48 hours. Fold change (log 2) and FDR are shown (two tailed t-test). Metabolites denoted with a

^†represent those belonging to the purine metabolism pathway, while those denoted with a

^‡represent metabolites belonging to the pyrimidine pathway.

Discussion.

This work is the first to evaluate the anti-cancer activity of the novel tropolone MO-OH-Nap in OS cells. We show that MO-OH-Nap induces the UPR, activates apoptosis, deregulates iron homeostasis pathways, and alters the metabolic profiles of OS cells. Furthermore, we demonstrate that the anti-OS effects of MO-OH-Nap are abrogated by the addition of exogenous iron.

To the best of our knowledge only one other tropolone has been evaluated in OS cells. Hinokitiol, also known as β-Thujaplicin, induced cytotoxic effects in U-2 OS and MG-63 cells in MTT assays (20). Hinokitiol promoted cell cycle arrest at the S phase and a DNA damage response in both OS cell lines. In addition, they found that the cellular response to hinokitol varied based on p53 mutation status. In cells with wild-type p53, hinokitiol exposure resulted in senescence, whereas cells with mutated p53 primarily underwent apoptosis with cleaved-PARP expression. Our studies utilized cell lines with a range of p53 mutation status, including wild-type (CAL-72), mutated (143B, HOS, MG-63), and null (SaOS-2) (21). We did not observe a correlation between p53 mutation status and response to MO-OH-Nap. Treatment with MO-OH-Nap induced activation of all three arms of the UPR (PERK, IRE1 and ATF6) and initiated apoptotic cell death in a concentration-dependent manner in a variety of OS cell lines. Elevation of UPR-associated proteins serves as a protective mechanism in times of ER stress, but it also makes cells more susceptible to induction of apoptosis. Several studies have identified upregulation of the UPR as a common molecular feature of OS, suggesting activation of the pathway may serve as a novel therapeutic approach to induce OS cell death (22–24). Further investigation into the mechanism by which MO-OH-Nap mediates UPR activation will be the focus of future studies.

In addition to their anti-cancer effects, tropolones have been reported to have putative bone protective properties. In one study, hinokitiol was found to inhibit RANKL-induced osteoclast formation and bone resorption in vitro and protects against bone loss in vivo (25). Hinokitiol blocked activation of the ERK, p38, and JNK–MAPK pathways, which suppressed the activity and expression of downstream factors, such as c‐Jun, c‐Fos, and NFATC1. These results are consistent with those of a previous study in which hinokitiol inhibited periodontal bone loss caused by tooth ligation in mice (26). These findings may be relevant to OS, as stage one of the metastatic cascade of OS cells to the lungs involves the induction of osteoblast-secreted RANKL and activation of osteoclast-mediated bone resorption (27). Inhibition of this early stage of metastasis could have profound therapeutic implications for the treatment of OS. In fact, several studies have successfully used RANK-Fc to inhibit the RANKL-induced osteoclast formation and target metastasis in OS cells (28–30). To date, no one has evaluated the effects of tropolones on OS cell metastasis. However, we found that MO-OH-Nap restricted the migration and invasion of OS cells in vitro, and that this effect was apparent after only 24-hours with sub-cytotoxic concentrations. Future studies will assess MO-OH-Naps ability to inhibit osteoclast function in vitro and prevent lung metastasis in vivo.

Our analysis of metabolomics data identified purine and pyrimidine metabolism as the top enriched pathways in MO-OH-Nap treated 143B cells. Purines and pyrimidines are critical for DNA replication, RNA synthesis, enzyme regulation, cell signaling, and metabolism (31). Across a wide range of cancer types, nucleotide metabolism is upregulated to allow for rapid cell proliferation, making it an attractive therapeutic target (32). Analysis of serum samples from 65 OS patient identified purine metabolism as one of the top deregulated metabolic pathways in OS (33). Profiling of IMPDH2 and HPRT in 127 patient samples revealed prognostic value in the expression patterns of purine-metabolizing enzymes for the pre- and post-chemotherapy period of OS treatment (34). Current treatment for OS includes methotrexate, which is a folic acid analog that inhibits purine and pyrimidine synthesis, consequently reducing nucleotide synthesis (35). We observed primarily downregulation of purine and pyrimidine pathway metabolites with MO-OH-Nap treatment, suggesting suppression of these pathways. Future studies will investigate the mechanism by which MO-OH-Nap alters purine and pyrimidine metabolism in OS cells, as well as explore the therapeutic potential of combining the dihydrofolate reductase inhibitor methotrexate with MO-OH-Nap.

Iron is required for ATP generation, DNA synthesis, antioxidant protein function, and DNA-damage repair. Owning to their higher rates of DNA synthesis and proliferation, cancer cells have an elevated requirement for iron relative to normal cells (36). Several studies have concluded that OS is characterized by abnormal iron metabolism. For instance, TFR1 protein levels are often upregulated in OS patient samples and the degree of TFR1 elevation serves as a prognostic indicator for survival (37). Upregulation of TFR1 increases the rate of iron uptake by OS cells, thereby promoting cell proliferation. In addition, TFR1 interaction with IKK leads to the activation the NF-Kappa-B signaling pathway, which in turn inhibits apoptosis and promotes OS cell survival (38). In addition, OS is characterized by low levels of ferritin light chain (FTL), and lower FTL expression is associated with shorter time to metastasis and decreased survival among patients (39). Lastly, TP53 is the most frequently mutated gene in OS, and recent research suggests mutated TP53 may act as a gain-of-function mutation capable of increasing intracellular ferrous iron pools, which in turn promotes OS cell growth and proliferation (40, 41). We previously demonstrated that MO-OH-Nap acts as a ferrous iron chelator and its ability to induce apoptosis in both MM and OS cells is abrogated by the addition of exogenous ferric or ferrous iron (14). Two iron chelators, deferoxamine and deferasirox, were shown to induce apoptosis in three OS cell lines (MG-63, MNNG/HOS and K7M2) via activation of MAPK signaling pathways and promotion of reactive oxygen species (ROS) (42). It is worth noting that MOH-OH-Nap did not induce ROS production in previous experiments (14). In another study, deferoxamine treatment induced growth inhibition and caspase-mediated apoptosis in SaOS-2 cells (43). Likewise, both deferoxamine and deferiprone were found to inhibit proliferation in MG-63 cells and this effect was reversed by co-incubation with iron citrate (44). To the best of our knowledge, iron chelators have not been demonstrated to induce the UPR in OS cells. However, deferoxamine does induce upregulation of UPR markers in MM cell lines (14). Additional research is needed to better understand the mechanism by which the ability of MO-OH-Nap to chelate iron translates to the induction of the UPR and apoptosis in OS cells.

Conclusions.

We demonstrate that MO-OH-Nap induces UPR-mediated apoptotic cell death in a variety of OS cell lines. Importantly, the cytotoxic effects of MO-OH-Nap are in part dependent on the ability of the drug to chelate cellular iron. Future studies will evaluate if MO-OH-Nap is efficacious in slowing tumor growth in mouse models of OS. Moreover, it will be important to evaluate the effects of combining MO-OH-Nap with clinically utilized chemotherapeutic drugs relevant to the treatment of OS.

Data availability statement.

The data that supports the findings of this study are available in the Supporting Information Material of this article.