Bromoxib is cytotoxic in leukemia and lymphoma cells and activates the mitochondrial apoptosis pathway
To evaluate the cytotoxic potential, we treated several hematological cancer cell lines (Fig. 2A) and solid tumor cell lines (Fig. 2B) with the polybrominated diphenyl ether bromoxib for 24 h. Subsequently, cell viability was assessed by AlamarBlue® assay. Thus, bromoxib displayed a high cytotoxicity in Ramos lymphoma cells (IC50 2.38 µM), and leukemia cells, such as HL60 (IC50 10.40 µM), SUPB15 (IC50 6.50 µM), Jurkat (IC50 9.54 µM), MOLT4 (IC50 18.72 µM) and also in the glioblastoma cell lines SJ-GBM2 (IC50 2.50 µM) and TP365MG (IC50 6.91 µM), whereas other cell lines were less sensitive (Fig. 2A, B).
Next, we investigated which apoptosis signaling pathways were activated by bromoxib. There exist two central apoptosis pathways: The extrinsic death receptor pathway and the intrinsic mitochondrial death pathway. Death receptor-mediated cell death is activated by death receptors (such as CD95/Apo-1/Fas, TRAIL-R1, or TRAIL-R2) upon binding to their respective ligands (e.g., CD95L/Apo-1L/FasL or TRAIL), leading to the activation of initiator caspase-8, which subsequently activates effector caspase-3. The mitochondrial death pathway is instigated by cellular stress, such as DNA damage induced during radio- and chemotherapy. It commences with the release of cytochrome c from the mitochondria, mediated by proapoptotic Bcl-2 proteins like Bax or Bak. Antiapoptotic Bcl-2 proteins (e.g. Bcl-2, Bcl-xL, Mcl-1) counteract proapoptotic Bcl-2 members by inhibiting the release of mitochondrial cytochrome c and thereby blocking the mitochondrial death pathway [10]. Within the cytosol, cytochrome c binds to the adapter protein Apaf-1, prompting the activation of initiator caspase-9 within a high molecular weight signaling complex known as apoptosome. Upon activation, caspase-9 catalyzes the activation of effector caspase-3 and 7 [11, 12].
In order to study whether bromoxib induces apoptosis via the external death receptor pathway, we used caspase-8 deficient and proficient Jurkat cells [13] and assessed the apoptosis-related DNA degradation by flow-cytometric measurement of propidium iodide stained apoptotic hypodiploid nuclei using the protocol of Nicoletti et al. [14]. As shown in Fig. 2C, bromoxib induced apoptosis in both, caspase-8 positive and caspas-8 negative Jurkat cells, similar to the broad kinase inhibitor and potent apoptotic stimulus staurosporine (STS) and the DNA-damaging anticancer drug etoposide which are known to induce apoptosis independent of death receptor signaling [15, 16]. As expected, apoptosis-induction via the death receptor ligand TRAIL (tumor necrosis factor-related apoptosis-inducing ligand) was completely blocked in the absence of caspase-8 (Fig. 2C). Hence, the activation of external death receptor signaling could be ruled out in bromoxib-induced apoptosis. However, when we used Jurkat cells deficient for caspase-9 (as the central initiator-caspase of the mitochondrial death pathway), bromoxib-induced apoptosis was completely abrogated (Fig. 2D).
Usually, overexpression of antiapoptotic Bcl-2 proteins (such as Bcl-2, Bcl-xL, or Mcl-1) inhibits the activation of the mitochondrial death pathway [10]. However, since some natural products like viriditoxin can directly activate the mitochondrial apoptosis pathway even in the presence of antiapoptotic Bcl-2 [17], we investigated to what extent bromoxib-induced apoptosis was affected in Bcl-2 overexpressing Jurkat cells. As shown in Fig. 2E, overexpression of Bcl-2 strongly attenuated apoptosis induced by bromoxib and etoposide. As staurosporine is proficient in inducing apoptosis in Bcl-2 or Bcl-xL overexpressing cells [15], apoptosis induction was not affected by Bcl-2.
Activation of the mitochondrial apoptosis pathway is characterized by the translocation of the proapoptotic Bax protein to the mitochondria, which mediates the subsequent mitochondrial release of proapoptotic factors such as cytochrome c and Smac (also known as Diablo) [10, 12, 18]. To further confirm the involvement of the mitochondrial apoptosis pathway, we monitored the translocation of GFP-Bax to mitochondria and the mitochondrial release of Smac-mCherry in HeLa cells. Thus, we observed that bromoxib induced the mitochondrial translocation of Bax and mitochondrial release of Smac within 1–2 h (Fig. 3). Taken together, these data indicate, that bromoxib induced apoptosis by activation of the intrinsic mitochondria-dependent signaling pathway.
Bromoxib causes changes in mitochondrial morphology, resulting in fission that is induced by OMA1-mediated processing of OPA1
Next, we focused on mitochondria as the executioners of the intrinsic apoptosis pathway. Therefore, we assessed the impact of bromoxib on the mitochondrial membrane potential (ΔΨm) in Ramos cells. As depicted in Fig. 4A, bromoxib induced a rapid breakdown of the mitochondrial membrane potential (ΔΨm) within 1–2 min, which was as fast as the protonophore carbonyl cyanide m-chlorophenyl hydrazone (CCCP), which served as a positive control.
In addition to the breakdown of the mitochondrial membrane potential (ΔΨm), the mitochondrial death pathway is characterized by mitochondrial fragmentation (fission). Mitochondrial fission is partly regulated by the proteolytic cleavage of the dynamin-like GTPase OPA1 by OMA1 or YME1L1, thereby maintaining the equilibrium between mitochondrial fusion and fission. Consequently, OPA1 serves a pivotal role in mitochondrial homeostasis by sensing mitochondrial stress, such as the breakdown of the mitochondrial membrane potential (ΔΨm), and coordinating mitochondrial quality control and intrinsic apoptosis. Stress-induced cleavage of long isoforms of OPA1 (L-OPA1) and the subsequent formation of short isoforms (S-OPA1) tilts the balance towards mitochondrial fragmentation and increased sensitivity to proapoptotic stimuli [19, 20].
Therefore, we studied the impact of bromoxib on OPA1 processing and observed that bromoxib induced the cleavage of L-OPA1 in Ramos cells within 5 min (upper Western blot of Fig. 4B). The recovery of long OPA1 forms upon processing after drug removal serves as an indicator of the recovery of the fusion machinery. To measure L-OPA1 recovery, Ramos cells were treated with bromoxib or CCCP for 30 min, followed by drug removal. Similar to CCCP, the continuous incubation with bromoxib led to L-OPA1 depletion. However, the removal of bromoxib and CCCP allowed the recovery of the long OPA1 forms (Fig. 4B; lower panel), indicating that the effect on mitochondrial fusion is reversible. Usually, S-OPA1 arises through proteolytic cleavage of L-OPA1 by proteases such as OMA1 or YME1L1 [21, 22]. To address which protease (OMA1 or YME1L1) catalyzes the cleavage of L-OPA1 into S-OPA1 forms, mouse embryonic fibroblasts (MEFs) lacking either OMA1, YME1L1, or both were treated with bromoxib or CCCP. The short and long OPA1 forms were then analyzed by immunoblotting (Fig. 4C). The findings indicated that OMA1 is necessary for cleaving L-OPA1 into S-OPA1, as cleavage was absent in OMA1 knockouts. Conversely, YME1L1 knockout did not affect OPA1 cleavage (Fig. 4C). Thus, these data indicate that OMA1 is the responsible protease for bromoxib (and CCCP) induced OPA1 cleavage. In addition, we observed that L-OPA1 depletion coincided with the rapid and pronounced fragmentation of the mitochondrial network within 30 min of treatment with bromoxib or CCCP (used as positive control for mitochondrial fragmentation; Fig. 4D).
In addition, mitochondrial depolarization associated with a sustained increase in cytosolic Ca2+ can also induce mitochondrial fission through activation of the Ca2+-dependent phosphatase calcineurin and subsequent dephosphorylation of DRP1 [23, 24]. Therefore, we investigated whether bromoxib was able to induce the mobilization of Ca2+ in Ramos cells. As a positive control for Ca2+ influx, we used the ionophore ionomycin, which also induces mitochondrial permeability transition opening [25, 26]. Intriguingly, bromoxib induced an increase in cytosolic Ca2+ even when extracellular Ca2+ was chelated by EGTA (Fig. 4E and Fig. 4F; left graph). In addition, thapsigargin was employed to fully deplete the Ca2+ stores of the endoplasmic reticulum (ER). To determine the origin of the released Ca2+, thapsigargin was initially applied to release all Ca2+ stored within the ER. However, subsequent application of bromoxib was still able to mediate a residual release of intracellular Ca2+ (Fig. 4F; right graph), indicating that the Ca2+ mobilized by bromoxib originates from the ER and most likely from mitochondria.
As severe mitochondrial damage and oxidative stress are often interdependent, we investigated the effect of bromoxib on cellular reactive oxygen species (ROS) generation. However, measurements with the fluorescent dye H2-DCF-DA revealed no increase in ROS production (Fig. 5A, B). Accordingly, treatment with the antioxidant N-acetylcysteine (NAC) did not affect bromoxib-induced toxicity (Fig. 5C).
Bromoxib inhibits glycolysis and mitochondrial metabolism
Given that one of the primary functions of mitochondria is the supply of cellular ATP, we examined the extent to which bromoxib affects cellular ATP levels. To distinguish the effects on glycolysis from oxidative phosphorylation (OXPHOS), cells were provided with either glucose or galactose as the sole sugar source within the medium. The glycolytic consumption of galactose instead of glucose does not result in a net ATP gain, thereby forcing the cell to rely exclusively on OXPHOS for ATP production. This renders the cell particularly susceptible to inhibitors of the electron transport chain (ETC) [27]. As shown in Fig. 5D, bromoxib induced a marked reduction in ATP levels under both glucose and galactose conditions, which was distinct from all other ETC inhibitors tested. To investigate whether bromoxib could directly inhibit one of the five electron transport chain (ETC) complexes, we analyzed the activity of each complex in purified mitochondria following bromoxib treatment. Intriguingly, bromoxib exhibited marked inhibitory activity and selectively targeted ETC complexes II, III, and the ATP-synthase (complex V) (Fig. 5E).
To confirm bromoxib's role as an inhibitor of mitochondrial respiration, a mitochondrial stress test was performed using the Agilent Seahorse system. Therefore, HeLa cells were treated with bromoxib and mitochondrial respiration was monitored by oxygen consumption rate (OCR) for 100 min. To induce inhibition of ATP-synthase (complex V) and restrict the electron flow along the ETC, oligomycin was applied, thereby leading to a decline in mitochondrial respiration, as indicated by the OCR. This reduction in OCR correlates with cellular ATP production. Subsequently, a decoupling agent, FCCP (carbonyl cyanide-4 (trifluoromethoxy) phenylhydrazone; a CCCP analog), was administered. This agent induces the collapse of the proton gradient and disrupts the mitochondrial membrane potential. As a result, electron flow through the ETC becomes unrestricted and oxygen consumption by complex IV reaches its maximum capacity, resulting in a plateau effect with DMSO between 60 and 80 min (Fig. 5F).
The FCCP-augmented OCR is then used to calculate the spare respiratory capacity (SRC), defined as the difference between maximal and basal respiration (Fig. 5G). The SRC provides a measure of the cell's ability to respond to escalated energy demands or stress conditions. The subsequent application of a combination of rotenone (inhibitor of complex I) and antimycin A (inhibitor of complex III) effectively stops mitochondrial respiration and facilitates the calculation of non-mitochondrial respiration driven by mechanisms outside the mitochondria. As shown in Fig. 5F, treatment with 10 µM bromoxib immediately caused a transient increase in OCR before mitochondrial respiration dropped to almost zero. Interestingly, subsequent addition of FCCP did not result in an increase in OCR, consistent with the observation that bromoxib impaired ETC complexes II and III. Subsequently, the non-mitochondrial respiration, basal respiration, acute response, post-treatment respiration, maximal respiration, spare respiratory capacity (SRC), ATP production and proton leak were calculated from the OCR measurements (Fig. 5G). Thus, bromoxib induced an acute response and triggered a pronounced reduction of respiration after treatment, maximal respiration, and spare respiratory capacity (Fig. 5G). In addition, bromoxib provoked a proton leak and completely abrogated the ATP production (Fig. 5D, G). These findings support the notion that bromoxib, like CCCP, might function as a protonophore and in addition blocks OXPHOS complexes, thereby impairing mitochondrial morphology and respiratory metabolism.
Bromoxib acts as a mitochondrial uncoupler
In general, a protonophore can be defined as a proton translocator or ionophore that facilitates proton transfer across membranes. Protonophores typically consist of an aromatic structure and a negative charge, which is distributed over the atoms via π-orbitals upon proton attachment, leading to charge delocalization. This chemical property justifies the term ‘protonophore’, as positively charged protons characterized by their hydrophilic nature, typically require channels, transporters, or passive diffusion facilitated by protonophores to traverse membranes [28, 29]. Prominent examples of protonophores are compounds like CCCP, 2,4-dinitrophenol, or triphenylphosphonium, which are known for their cytotoxic potential in various cancer cell types, including breast cancer, lung cancer, neuroblastoma, and glioblastoma cells [30–32]. These agents function by disrupting the normal proton gradient across the inner mitochondrial membrane, thereby uncoupling oxidative phosphorylation (OXPHOS) and diminishing ATP production.
Since bromoxib displayed several features of a protonophore, we compared bromoxib with other protonophores concerning OCR, cytotoxicity, caspase-activation and apoptosis-induction. Using the Seahorse Mito Stress Test we analyzed the concentration-dependent mitochondrial uncoupling of bromoxib. For the protonophore FCCP it has been shown that it induces a concentration-dependent metabolic demand, leading to rapid oxidation of substrates such as sugars, lipids and amino acids [33]. In the case of bromoxib, we observed a striking similarity when comparing the concentration-dependent, plateau-like elevation of OCR in bromoxib-treated cells with the corresponding increase induced by FCCP-treatment, suggesting a similar cellular response. Thus, bromoxib appears to uncouple mitochondrial respiration in a similar way to FCCP by disrupting the proton gradient, leading to almost complete cessation of OXPHOS. (Fig. 6A). Likewise, bromoxib displayed a similar potential as the protonophore CCCP in terms of cytotoxicity, caspase-activation, and apoptosis rate (Fig. 6B, C and D). Remarkably, both protonophores displayed similar cytotoxic IC50 values in Ramos cells (bromoxib: 4.98 µM; CCCP: 4.96 µM; Fig. 6B). In addition, bromoxib and CCCP initiated the activation of caspases with similar kinetics (within 8 h) and to a similar extent in terms of caspase-activation and apoptosis-induction (Fig. 6C, D). A comparative analysis also included the hexokinase II-inhibitor 3-bromopyruvate (3-BP) to discern whether inhibition of glycolysis through suppression of glycolytic enzymes via 3-BP was sufficient to reduce cell viability and activate caspase-3 in Ramos cells. Although, 3-BP induced cytotoxicity (albeit with a higher IC50 value (11.02 µM) compared to bromoxib and CCCP) it did not provoke any caspase-activation (Fig. 6B, C). This proves that inhibition of glycolysis by 3-BP alone is not sufficient to trigger caspase-dependent apoptosis. In summary, we could show that bromoxib functions as a mitochondrial uncoupler and induces apoptosis similar to the protonophore CCCP in Ramos cells.
In thermal proteome profiling (TPP), bromoxib caused the stabilization of tubulin and proteins involved in fatty acid oxidation (FAO)
In order to identify proteins targeted by bromoxib, we conducted an unbiased proteomic analysis employing thermal proteome profiling (TPP). This method enables the detection of drug-protein interactions in living cells by identifying proteins whose thermal stabilities are altered, directly or indirectly, upon drug binding. Through mass spectrometry, the thermal profiles of these affected proteins can be determined, thereby enabling the successful identification of respective drug targets [34, 35]. Using the TPP-approach, 31 stabilized proteins could be identified in Ramos cells which were stabilized upon treatment with 40 µM bromoxib for 30 min (Fig. 7A and Suppl. Table 1). These proteins could be further subdivided into two main hit-clusters: (i) the COPI-mediated transport system, which is composed of tubulin isoforms and (ii) a protein-cluster involved in fatty acid metabolic processes (Fig. 7A, B and Suppl. Table 1).
Tubulin molecules are composed of heterodimers, consisting of α- and β-chains, which arrange themselves in a head-to-tail manner to form protofilaments running along the length of the microtubule structure. Following bromoxib treatment, two α-tubulins, namely TUBA1B and TUBA1C, in addition to the two β-tubulins TUBB and TUBB4B, exhibited enhanced stability in the TPP-approach. Therefore, we investigated whether the cytotoxic mechanism of bromoxib might be due to a disruption of tubulin polymerization. However, using a tubulin polymerization assay we detected no significant influence on the polymerization rate of tubulin and no disturbance of cell cycle progression as would be expected for an antimitotic toxin (Suppl. Figure 1). Consequently, we excluded a disturbed tubulin polymerization as the major mechanism for bromoxib-mediated cytotoxicity.
Next, we focused on the protein cluster associated with fatty acid and lipid metabolism, including the proteins ACSL4, HADHA, HADHB, ECH1, and ACADVL (Fig. 7B). Given that bromoxib predominantly targets mitochondria, it was expected that proteins localized within mitochondria would emerge as prominent candidates in the TPP results. As shown in Fig. 7C, these proteins were stabilized in TPP after only 30 min of exposure to bromoxib. To verify this finding, the bromoxib mediated stabilization of the candidate proteins (ACSL4, HADHB, ECH1, and ACADVL) was analyzed by cellular thermal shift assays (CETSA) [36] by subjecting the same temperature-treated samples analyzed by quantitative MS-based TPP to quantitative immunoblotting. This shows that treatment with bromoxib augmented the protein levels of these proteins upon increasing temperature treatment (Fig. 7D, E).
Bromoxib increases fatty acid oxidation
Fatty acid β-oxidation (FAO) represents a central metabolic pathway responsible for the mitochondrial breakdown of long-chain acyl-CoA to acetyl-CoA. The process begins with the activation of long-chain fatty acids via thioesterification (CoA binding), facilitated by acyl-CoA synthetase (ACSL4). Subsequently, these long-chain fatty acids are transported from the cytosol to the inner mitochondrial area through carnitine-acylcarnitine translocase (CACT) and released as acyl-CoA [37]. Importantly, all five proteins that were stabilized by bromoxib treatment in the TPP-approach play a central role in FAO. Their distinct functions within the FAO pathway can be summarized as follows: acyl-CoA dehydrogenase (ACADVL) mediates the oxidation of the acyl while FAD is reduced to FADH2, enoyl-CoA hydratase (ECH1) facilitates hydration, 3-hydroxy acyl CoA dehydrogenase (HADHA) mediates an additional redox step where NADH will be released, and 3-ketoacyl CoA thiolase (HADHB) mediates thiolysis, consequently releasing acetyl-CoA into the TCA cycle and FADH2 and NADH into the ETC [37].
Therefore, we investigated the effect of bromoxib on fatty acid β-oxidation. First, we analyzed to what extent the protein levels of ACSL4, HADHA, HADHB, ACADVL, and ECH1 were affected upon treatment with bromoxib for 30 min or 24 h. Quantification of immunoblots revealed no significant change in protein levels of ACSL4, HADHB, ACADVL ECH1 after 30 min treatment with 40 µM bromoxib (Fig. 8A). However, protein level analysis upon 24 h treatment with bromoxib showed a noticeable reduction of ACSL4 and HADHB, an insignificant reduction of ACADVL levels, and a significant increase in ECH1 levels (Fig. 8B). This difference between short-term and long-term treatment might be attributed to distinct rapid and delayed response mechanisms to bromoxib.
Next, we investigated the effect of bromoxib on fatty acid metabolism by analyzing the total lipid content (Fig. 8C and D) and of the total and partial palmitate oxidation in Ramos cells via the detection of [14C]-CO2 (total palmitate oxidation) and [14C]-acid-soluble metabolites (partial palmitate oxidation) (Fig. 8E). Fatty acids (FAs) serve as building components of various types of lipids. They contain a carboxylic acid moiety coupled with a hydrocarbon chain characterized by varying carbon lengths and degrees of saturation [38]. By channeling FAs into different metabolic pathways, it is possible to produce more complex lipid species such as diacylglycerols (DGs) and triacylglycerols (TGs), and glycerophospholipids phosphatidylethanolamine (PE), phosphatidylinositol (PI), phosphatidylcholine (PC), lysophosphatidylcholine (LPC), and phosphatidylserine (PS), among others [39]. Additionally, FAs contribute to the production of triacylglycerols (TGs) and cholesteryl esters (CEs), which serve as reservoirs of energy in the form of lipid droplets. When needed, these lipid droplets are used as a fuel source for cellular bioenergetics via fatty acid oxidation (FAO) [40, 41]. Free FAs undergo esterification in the glycerol phosphate pathway, giving them amphiphilic properties similar to phospholipids and sphingolipids. In particular, cholesterol, phospholipids, and sphingolipids constitute the basic building blocks of biological membranes [40, 42].
Following treatment with 10 µM bromoxib for 30 min, a significant elevation was observed concerning the levels of lysophosphatidylcholine, phosphatidylcholine, phosphatidylethanolamine, and free cholesterol. Conversely, cardiolipin levels decreased significantly (Fig. 8C). Diacylglycerols, cholesteryl esters and triacylglycerols were unaffected, whereas free fatty acids were increased upon bromoxib treatment. (Fig. 8D). In terms of total and partial palmitate oxidation (as surrogate markers for FAO activity), a significant increase in [14C]-acid-soluble metabolites was observed following treatment with bromoxib, indicating an increase in partial palmitate oxidation (Fig. 8E). Thus, it appears that the cell attempts to compensate for the loss of ATP production (due to the breakdown of OXPOS) by rapidly upregulating FAO upon bromoxib-treatment.