Novel meriolin derivatives activate the mitochondrial apoptosis pathway in the presence of antiapoptotic Bcl-2

doi:10.21203/rs.3.rs-3527444/v1

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Novel meriolin derivatives activate the mitochondrial apoptosis pathway in the presence of antiapoptotic Bcl-2

https://doi.org/10.21203/rs.3.rs-3527444/v1

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published 09 Mar, 2024

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Meriolin derivatives represent a new class of kinase inhibitors with a pronounced cytotoxic potential. Here, we investigated a newly synthesized meriolin derivative (termed meriolin 16) that displayed a strong apoptotic potential in Jurkat leukemia and Ramos lymphoma cells. Meriolin 16 induced apoptosis in rapid kinetics (within 2 - 3 h) and more potently (IC₅₀: 50 nM) than the previously described derivatives meriolin 31 and 36 [1]. Exposure of Ramos cells to meriolin 16, 31, or 36 for 5 min was sufficient to trigger severe and irreversible cytotoxicity. Apoptosis induction by all three meriolin derivatives was independent of death receptor signaling but required caspase-9 and Apaf-1 as central mediators of the mitochondrial death pathway. The mitochondrial toxicity of meriolins was further confirmed by the breakdown of the mitochondrial membrane potential (ΔΨm), mitochondrial release of proapoptotic Smac, processing of the dynamin-like GTPase OPA1 and subsequent fission of mitochondria. Remarkably, all meriolin derivatives could directly activate the mitochondrial apoptosis pathway in Jurkat cells even in the presence of antiapoptotic Bcl-2 protein. In addition, meriolins were capable to induce cell death in imatinib-resistant K562 and KCL22 chronic myeloid leukemia cells as well as in cisplatin-resistant J82 urothelial carcinoma and 2102EP germ cell tumor cells. Since tumor cells frequently inactivate the mitochondrial death pathway (e.g. by overexpression of antiapoptotic Bcl-2 proteins) in order to acquire therapy resistance, meriolin derivatives might represent a promising therapeutic option for overcoming treatment resistance.

Biological sciences/Cancer/Cancer therapy/Drug development

Biological sciences/Drug discovery/Pharmaceutics

Biological sciences/Cell biology/Cell death/Apoptosis

Multikinase inhibitors interfere with cell proliferation, apoptosis, inflammation and metabolism and are therefore successfully applied in cancer therapy [2–5]. So far, 66 protein kinase inhibitors have been approved by the FDA that are predominantly applied in cancer treatment [6].

Meriolins (3-(pyrimidin-4-yl)-7-azaindoles) are synthetic hybrids of the naturally occurring (aza)indole alkaloid family of variolins and meridianins and exhibit potent kinase inhibitory features [7, 8] (Fig. 1A). They were first synthesized in 2001 [9] and termed meriolins by Meijer and co-workers [7, 10]. Similar to their parental natural products, meriolins potently inhibit a broad range of cyclin-dependent kinases (CDKs) and appear to be even more active than variolins and meridianins in vitro and in vivo [1, 7, 10–14]. Previous studies could show that meriolins act as multikinase inhibitors by inhibiting a variety of kinases such as CDKs (CDK1, 2, 4, 5 and 9) [7, 10, 15, 16] and to a lesser extent other kinases such as GSK-3, CK1, DYRK1A [10], sphingosine kinase 1 and 2 [1, 17], and PDK1 [18]. It has been shown that meriolins exhibit a pronounced cytotoxicity against several cancer cell lines with IC₅₀ values in nanomolar range [1, 7, 10, 12, 13]. In addition, it was demonstrated that meriolins induce cell cycle arrest [11, 19] and apoptosis [1, 7, 10, 11, 20] and consequently display antitumor activity in different xenograft cancer models (i.e. Ewing’s sarcoma, LS174T colorectal carcinoma and U87 glioblastoma) [7, 11]. Since dysregulation of cell cycle and aberrant proliferation represent key hallmarks of cancer [21, 22], targeting of CDKs represents a promising approach for anticancer therapy [4, 5]. In this context meriolin 3 (that inhibits CDK1, CDK2, CDK5 and CDK9) has found its way into preclinical trials [16].

We have previously shown that out of 13 newly synthesized meriolin derivatives, 6 displayed a pronounced cytotoxicity in Jurkat T cell leukemia and Ramos Burkitt B cell lymphoma cells. Among these tested meriolin derivatives meriolin 31 and 36 (Fig. 1A) showed the highest cytotoxic potential [1].

Here, we characterize the apoptosis signaling of the novel derivative meriolin 16 in comparison to the potent derivatives meriolin 31 and 36. Meriolin 16 was developed and synthesized with an additional methoxy group in order to enhance its cytotoxicity and accordingly displayed a higher cytotoxicity (IC₅₀: 50 nM) than meriolin 31 and 36 (IC₅₀: 90 nM and 170 nM) and even as meriolin 3 (IC₅₀: 70 nM), which is the most potent meriolin derivative described so far. Interestingly, a 5 min exposure to meriolin 16, 31 or 36 was sufficient to activate the endogenous suicide program upon 24 h incubation. Meriolin-induced apoptosis was independent of external death receptor signaling but relied on Apaf-1 and caspase-9 as central mediators of the mitochondrial apoptosis pathway. Activation of the intrinsic mitochondrial death pathway was further confirmed by the breakdown of the mitochondrial membrane potential (ΔΨm), mitochondrial release of proapoptotic Smac, processing of the dynamin-like GTPase OPA1 and subsequent mitochondrial fission. Intriguingly, all three meriolin derivatives could directly activate the mitochondrial apoptosis pathway in Jurkat cells even in the presence of antiapoptotic Bcl-2 protein. Since tumor cells frequently inactivate the mitochondrial death pathway in order to acquire therapy resistance (e.g. by elevated levels of antiapoptotic Bcl-2 proteins) these meriolin derivatives display a favorable option for overcoming therapy resistance. In this context we could show that meriolins were able to induce cell death in imatinib-resistant K562 and KCL22 chronic myeloid leukemia cells and in cisplatin resistant J82 urothelial carcinoma and 2102EP germ cell tumor cells. Thus, the tested meriolin derivatives represent a promising therapeutic option for cancer treatment.

Synthesis of meriolin 16 by Masuda borylation-Suzuki coupling sequence

Starting from 4-methoxy 7-azaindole, meriolin 16 was prepared in concise two step synthesis Fig. 1B). First, 4-methoxy 7-azaindole was iodinated and N-tosylated in a one-pot fashion according to our literature protocol [23] to give 3-iodo-4-methoxy-1-tosyl-1H-pyrrolo[2,3-b]pyridine in 94% yield. Then, Masuda borylation-Suzuki coupling sequence [8] of 3-iodo-4-methoxy-1-tosyl-1H-pyrrolo[2,3-b]pyridine and 4-bromopyridine-2,6-diamine followed by subsequent detosylation furnished meriolin 16 in 98% after three steps in a one-pot fashion [24].

Meriolin 16, 31 and 36 are highly cytotoxic and induce apoptosis in rapid kinetics in Ramos and Jurkat cells

To evaluate the cytotoxic potential of the novel derivative meriolin 16 in comparison to meriolin 31 and 36, we performed viability assays. All three meriolin derivatives were extremely cytotoxic in Ramos lymphoma cells with IC₅₀ values ranging from 50 nM (meriolin 16) and 90 nM (meriolin 31) to 170 nM (meriolin 36) upon 24 h treatment (Fig. 2A). Thus, the IC₅₀ value of meriolin 16 was even lower than for meriolin 3 (IC₅₀: 70 nM; Suppl. Figure 1A), which is the most potent meriolin derivative described so far. In addition, we treated Ramos cells for only 5 min with meriolin 16, 31 or 36. Subsequently, cells were washed and further incubated for 24 h. Thereby, we could show that the exposure to the respective meriolin derivative for 5 min was sufficient to cause severe and irreversible cytotoxicity (Fig. 2A). Similarly, a 5 min exposure to meriolins was sufficient for the cleavage of the caspase substrate poly(ADP-ribose) polymerase-1 (PARP) after 24 h. Interestingly, PARP-cleavage upon 5 min exposure to all three meriolin derivatives was more pronounced than 5 min treatment with the broad kinase inhibitor and potent apoptotic stimulus staurosporine (STS) that was used as positive control (Fig. 2B).

To further evaluate the apoptotic potential, we monitored the apoptosis-related DNA degradation by the flow-cytometric measurement of propidium iodide stained apoptotic hypodiploid nuclei [25]. Again, meriolin 16 turned out to be more potent and to induce apoptosis at lower concentrations compared to meriolin 31 and 36 and to a similar extent in both, Jurkat leukemia and Ramos lymphoma cells. Meriolin-induced apoptosis was prevented by co-treatment with the pan-caspase inhibitor Q-VD-OPh (QVD), thereby indicating that cell death was caspase dependent (Fig. 3A-C). To measure the catalytic caspase activity, we used the profluorescent caspase-3 substrate Ac-DEVD-AMC to monitor the activity of caspase-3 upon treatment with different meriolin derivatives. The measurement of an 8-hour kinetics revealed a rapid activation of caspase-3 as early as 3–4 h in Ramos and Jurkat cells. However, meriolin-induced caspase activation was not quite as rapid as upon treatment with staurosporine (Fig. 3D-F). In addition, we observed a significant cleavage of the caspase substrate PARP in both cell lines with similar kinetics as in the caspase activity assay, which was again prevented by co-treatment with the pan-caspase inhibitor QVD (Fig. 3G-I).

Meriolins induce DNA damage which is a downstream event mediated by caspases

It has been demonstrated that CDK1 phosphorylates and activates the dynamin-related GTPase DRP1 during mitosis [26]. Conversely, loss of the fission protein DRP1 causes mitochondrial hyperfusion and induces ATM-dependent G2/M arrest and aneuploidy through DNA replication stress [27]. Therefore, we investigated whether the meriolin-induced CDK1 inhibition might trigger DNA damage, and subsequently initiate the DNA damage response. For this purpose, apoptosis induction upon treatment with meriolin 16 and 36 as well as the CDK1, 2 and 4 inhibitor R547 [28] was compared with the topoisomerase inhibitors and DNA damaging agents camptothecin, daunorubicin and etoposide. As shown in Suppl. Figure 2A, all tested apoptosis stimuli induced a pronounced caspase activity in similar range and kinetics. Next, we investigated the effect of CDK-inhibition by meriolins and R547 on the phosphorylation of the two checkpoint kinases CHK1 and CHK2. Both are responsible for cell cycle arrest upon DNA damage or during incomplete cell cycle progression. Upon DNA damage, CHK1 and CHK2 become phosphorylated by ATM or ATR [29]. ATR is activated by DNA single-strand breaks and other damage during the replication process and subsequently phosphorylates CHK1 at Ser345. CHK2 is phosphorylated at Thr68 by ATM, which is activated by DNA double-strand breaks. As depicted in Suppl. Figure 2B,C, all three CDK-inhibitors (meriolin 16, 36 and R547) induced a pronounced phosphorylation of CHK2 indicating the induction of DNA double-strand breaks, whereas CHK1 remained merely unaffected. Another biomarker for DNA double-strand breaks is the phosphorylation of γH2AX at Ser139 by ATM as a first step in the recruitment of DNA repair proteins [30]. In addition to γH2AX, ATM also phosphorylates KAP1 at Ser Ser824 [31] which subsequently co-localizes with γH2AX, TP53BP1 and BRCA1 at DNA damage foci [32]. Treatment of Ramos cells with meriolin 16 induced a substantial phosphorylation of γH2AX (at Ser139) and KAP1 (at Ser Ser824). However, pretreatment with the pan-caspase inhibitor QVD completely abolished meriolin 16-induced phosphorylation of γH2AX and KAP1, whereas etoposide-mediated phosphorylation was not affected (Suppl. Figure 2D, E). Thus, these data indicate that in contrast to etoposide, which inflicts DNA damage in a direct way, meriolin 16 induced DNA damage is rather an apoptotic downstream event mediated by caspases.

Meriolin 16, 31 and 36 activate the mitochondrial apoptosis pathway even in the presence of antiapoptotic Bcl-2

Next, we investigated whether the meriolin derivatives affected the different apoptotic signaling pathways. There exist at least two major pathways leading to the apoptotic demise of the cell – the extrinsic death receptor pathway and the intrinsic mitochondrial apoptosis pathway. In death receptor-mediated apoptosis, the stimulation of receptors (such as CD95/Apo-1/Fas, TRAIL-R1 or TRAIL-R2) via their respective ligands (e.g. CD95L/Apo-1L/FasL or TRAIL) leads to the activation of initiator caspase-8 that proteolytically activates effector caspase-3. The intrinsic mitochondrial cytochrome c/Apaf-1 pathway is usually activated by cell stress such as DNA damage inflicted during radio- and chemotherapy and is initiated by the release of cytochrome c from the mitochondria. The mitochondrial cytochrome c release is mediated by proapoptotic Bcl-2 proteins, such as Bax or Bak. Antiapoptotic members of the Bcl-2 family (such as Bcl-2, Bcl-xL, Mcl-1) can in turn inhibit the release of cytochrome c, thereby blocking the mitochondrial apoptosis pathway [33]. In the cytosol, cytochrome c binds to the adapter protein Apaf-1 and subsequently activates the initiator caspase-9 in a high molecular weight signal complex, termed apoptosome. Activated caspase-9 then cleaves and activates effector caspase-3 and − 7 [34, 35].

Since caspase-8 represents the central initiator caspase of the death receptor pathway we used caspase-8 deficient Jurkat cells [36] to investigate whether meriolin-induced apoptosis is mediated via the extrinsic apoptosis pathway. As shown in Fig. 4A, B, all three meriolins induced apoptosis and cleavage of the caspase substrate PARP in both, parental caspase-8 proficient Jurkat cells as well as in caspase-8 deficient Jurkat cells. Similarly, staurosporine and the anticancer drug etoposide, which have been shown to induce apoptosis independent of external death receptor signaling [37, 38], were also able to induce apoptosis in caspase-8 deficient cells. As expected, death receptor stimulation via TRAIL was consequently blocked in the absence of caspase-8. Thus, an involvement of the external death receptor pathway in meriolin-induced apoptosis could be excluded.

Activation of the mitochondrial apoptosis pathway can be blocked by overexpression of anti-apoptotic Bcl-2 proteins (such as Bcl-2, Bcl-xL, or Mcl-1). Intriguingly, meriolin 16, 31 and 36 mediated apoptosis induction and PARP-cleavage was only attenuated but not blocked in Jurkat cells overexpressing Bcl-2 (Fig. 4C, D). As to be expected, the apoptosis induction by the DNA-damaging anticancer drug etoposide was prevented in the presence of Bcl-2. Since staurosporine is proficient to induce apoptosis in Bcl-2 or Bcl-xL overexpressing cells [37], apoptosis induction was not affected by Bcl-2 (Fig. 4C, D). Because the activation of the mitochondrial apoptosis pathway relies on the Apaf-1 mediated activation of caspase-9 within the apoptosome, we investigated whether meriolins were capable of inducing apoptosis in the melanoma cell line SK-Mel-94, which was previously shown to be resistant to anticancer drugs due to the loss of Apaf-1 expression [39]. As shown in Fig. 4E and F, treatment of SK-Mel-94 cells with meriolin 16, 31 and 36 could not induce caspase-activation or PARP cleavage. Since anticancer drugs induce apoptosis via the mitochondrial apoptosis pathway, etoposide was also unable to induce apoptosis in SK-Mel-94 cells. As staurosporine is capable to activate caspase-9 in the absence of Apaf-1 [37], caspase-activation and PARP-cleavage was not obstructed (Fig. 4E, F). The observation that meriolin 16, 31 and 36 obviously required apoptosome formation was further corroborated by the observation that meriolin-induced apoptosis and PARP-cleavage was completely blocked in caspase-9 deficient Jurkat cells (Fig. 4G, H).

Activation of the mitochondrial death pathway is initiated by the release of proapoptotic factors such as cytochrome c and Smac (also known as Diablo) from the mitochondria [35]. To further corroborate the involvement of the mitochondrial apoptosis pathway, we monitored the mitochondrial release of Smac-mCherry in HeLa cells. Thus, we could show that within 2–3 h all three meriolin derivatives initiated the release of Smac from the mitochondrion (Fig. 5). Taken together, these data indicate that all three meriolin derivatives can activate the mitochondrial apoptosis pathway in a direct way that could not be blocked by Bcl-2 overexpression.

Meriolin 16, 31 and 36 impair mitochondrial function and structure

In the following, we focused on mitochondria as the executioners of the intrinsic apoptosis pathway. First, we measured the effect of meriolin 16, 31 and 36 on the mitochondrial membrane potential (ΔΨm). As shown in Fig. 6A, all three meriolin derivatives induced a substantial breakdown of the ΔΨm after 6 h – however not as fast as the protonophore carbonyl cyanide m-chlorophenyl hydrazone (CCCP), which was used as positive control.

Another hallmark of the mitochondrial apoptosis pathway is the fragmentation (fission) of mitochondria. In this context, the dynamin-like GTPase OPA1 plays a major role. OPA1 is a central regulator in the maintenance of mitochondrial homeostasis, serves as a sensor of mitochondrial stress (such as impaired mitochondrial membrane potential) and interconnects mitochondrial quality control and intrinsic apoptosis. The activity of OPA1 is regulated via proteolytic processing through OMA1 and YME1L1 and balances the fusion and fission of the mitochondrial network. Stress-induced cleavage of OPA1 and the associated loss of its long isoforms (L-OPA1) shifts the balance towards increased fission, leading to mitochondrial fragmentation and increased sensitivity to proapoptotic stimuli [40, 41]. Consequently, we investigated the impact of meriolin 16 and 36 on OPA1 and observed that both meriolin derivatives caused the cleavage of the long OPA1 isoforms (L-OPA1) in Ramos and Jurkat cells within 6–8 h (Fig. 6B). In addition, we observed that the loss of long OPA1 isoforms was accompanied by substantial fragmentation (fission) of the mitochondrial network after 6–8 h (Fig. 6C).

Meriolin 16 and 36 induce cytotoxicity in imatinib and cisplatin resistant cancer cells

The major detrimental feature of tumors is their potential to acquire resistance to radio- and chemotherapy. Consequently, preexisting or acquired resistance to anticancer drugs like imatinib or cisplatin treatment represents a major drawback that consequently results in tumor progression. In order to evaluate the potential to overcome therapy resistance, we tested the meriolin derivatives in imatinib-resistant sublines of K562 and KCL22 chronic myeloid leukemia (CML) cells and cisplatin-resistant sublines of J82 urothelial cancer cells and 2102EP germ cell tumor cells in comparison to the respective cisplatin-sensitive parental cell line [42–45]. As shown in Fig. 7, meriolin 16 and meriolin 36 displayed a pronounced cytotoxicity after 72 h treatment in imatinib-resistant K562 and KCL22 cells as well as in cisplatin-resistant J82 and 2102EP cells and appeared to be even more cytotoxic in the respective resistant cell line compared to the respective sensitive parental line.

As 7-azaindoles, meriolins represent structural hybrids of meridianins and variolins that have been isolated from marine invertebrates (Fig. 1A) [8, 46]. Like their parental natural compounds, meriolins potently inhibit a broad range of CDKs and appear to be even more active than variolins and meridianins in vitro and in vivo [1, 8, 10–14]. Thus, it has been shown that meriolins display a strong cytotoxic potential in nanomolar range in various tumor cell lines of different origin, such as colon cancer (HCT116, LS-174T), hepatoma (Huh7, F1), cervix carcinoma (HeLa), breast carcinoma (MCF-7), glioma (GBM, SW1088, U87), neuroblastoma (SH-SY5Y), leukemia (Jurkat, Molt-4), lymphoma (Ramos), and myeloma (KMS-11) [1, 7, 11, 19, 20, 47]. However, nontransformed cells as human foreskin fibroblasts appear to be more resilient [7]. In recent years, this substance class underwent derivatization and optimization in order to enhance their efficacy in cancer therapy. In this context, we have previously shown that 6 out of 13 newly synthesized meriolin derivatives displayed a pronounced cytotoxicity with meriolin 31 and 36 comprising the highest cytotoxic potential [1].

Here, we investigated the bioactivity of the novel derivative meriolin 16 and compared its cytotoxic potential to meriolin 31 and 36. Thus, we could show that meriolin 16 was extremely cytotoxic (IC₅₀: 50 nM) in Ramos lymphoma cells as compared to meriolin 31 (IC₅₀: 90 nM) and meriolin 36 (IC₅₀: 170 nM) upon 24 h treatment. Meriolin 16 was even more potent than meriolin 3 (IC₅₀: 70 nM) that has already been tested in preclinical trials [16]. Since meriolin 3 exhibits the highest cytotoxic potential amongst all meriolines described so far, this renders meriolin 16 as the most potent meriolin derivative to date.

The increased cytotoxicity of meriolin 16 compared to meriolin 31 and 36 might be attributed to the additional methoxy group at the aromatic pyridine ring of meriolin 16 (see Fig. 1A). The methoxy group was intentionally added in order to modify its pharmacokinetic properties and to enhance its biological activity. Besides affecting the polarity and solubility of a molecule, an additional methoxy group can increase potential reactivity by enhancing the electron density of the aromatic ring, thus rendering it more nucleophilic [48]. Since meriolin 3 also contains a methoxy group at the same position as meriolin 16 (Suppl. Figure 1B), the methoxy group might be of importance for the structure-activity relationship of these derivatives.

Most interestingly, a 5 min exposure to meriolin 16, 31 or 36 was sufficient to activate the endogenous suicide program in Ramos lymphoma cells after 24 h (Fig. 2). We further investigated the apoptosis signaling pathways activated by meriolins. Using cell lines deficient for caspase-8, -9 or Apaf-1, we could show that meriolin-induced apoptosis was independent of external death receptor signaling but required Apaf-1 and caspase-9 as central mediators of the mitochondrial apoptosis pathway (Fig. 4). Activation of the mitochondrial apoptosis pathway was further confirmed by the mitochondrial release of proapoptotic Smac, breakdown of the mitochondrial membrane potential (ΔΨm), processing of the dynamin-like GTPase OPA1 and subsequent mitochondrial fission (Fig. 4,5).

The question however remains, how meriolins activate the mitochondrial apoptosis pathway. We initially observed that meriolin 16 and 36 and other CDK inhibitors such as R547 induce the ATM-mediated phosphorylation of CHK at Thr68. In addition, meriolin 16 induced the ATM-mediated phosphorylation of γH2AX (at Ser139) and KAP1 (at Ser824). However, in contrast to DNA damaging agents as the topoisomerase II inhibitor etoposide, inhibition of caspases completely blocked the meriolin 16 induced phosphorylation of γH2AX and KAP1. Thus, these data indicate that meriolin-inflicted DNA damage is rather an apoptotic downstream event mediated by caspases. This effect might be attributed to the caspase-mediated degradation of the inhibitor ICAD during apoptosis that subsequently releases and activates the caspase-activated DNase (CAD) which then produces the characteristic internucleosomal DNA cleavage [49, 50]. Apoptotic internucleosomal DNA cleavage might in turn activate ATM and induce the phosphorylation of γH2AX and KAP1, which in case of meriolin 16, could be inhibited upon caspase-inhibition but not in case of direct DNA damage inflicted by etoposide. Thus, meriolin-induced activation of the mitochondrial apoptosis pathway is obviously not initiated upon DNA damage – as in case of radio- and chemotherapy.

Bettayeb et al. could demonstrate that meriolin 3 inhibits the kinase activity of various cyclin dependent kinases (such as CDK1–7 and CDK9) and prevents the phosphorylation of CDK1, CDK4, CDK9, retinoblastoma protein and RNA polymerase II. In addition, they observed that meriolin 3 induced the downregulation the antiapoptotic Bcl-2 protein Mcl-1 [7]. Consequently, they proposed that meriolin 3-mediated inhibition of CDK9 reduces RNA polymerase II activity, which in turn downregulates short-lived Mcl-1 proteins. However, the authors did not include caspase-inhibitors in their study to exclude a potential caspase-mediated degradation of Mcl-1, as previously shown [51, 52]. Usually, antiapoptotic Bcl-2 members (such as Bcl-2, Bcl-xL or Mcl-1) inhibit the activation of the mitochondrial apoptosis pathway by counteracting proapoptotic Bcl-2 members (such as Bax, Bak or BH3-only proteins). However, since we observed that meriolin 16, 31 and 36 were able to induce apoptosis in the presence of antiapoptotic Bcl-2 (Fig. 4C, D), it is rather unlikely that meriolins induce apoptosis via downregulation of antiapoptotic Bcl-2 proteins. The observation that meriolin 16, 31 and 36 induce apoptosis even in the presence of Bcl-2 rather suggests that they activate the mitochondrial apoptosis pathway in a direct way downstream of pro- and antiapoptotic Bcl-2 members.

Antiapoptotic members of the Bcl-2 family, such as Bcl-2, Bcl-xL, or Mcl-1, were first identified in leukemia and lymphoma and are frequently overexpressed in other types of neoplasia to prevent apoptotic cell death during tumorigenesis. Moreover, since the major mechanism of genotoxic radio- and chemotherapy is the p53-mediated activation of the mitochondrial apoptosis pathway, tumor cells gain therapy resistance by inactivating this route e.g. via overexpression of anti-apoptotic Bcl-2 proteins [33, 53]. Thus, the observation that meriolins are capable of directly activating the mitochondrial apoptosis pathway even in the presence of Bcl-2 renders them as valuable agents for the treatment of therapy-resistant tumors. This feature is corroborated by our observation that the tested meriolin derivatives were capable of inducing cell death in imatinib-resistant K562 and KCL22 CML cells as well as in cisplatin-resistant J82 urothelial carcinoma and 2102EP germ cell tumor cells (Fig. 7).

Beside cell death induction, it has also been shown that meriolins as multikinase inhibitors target kinases such as CDKs (e.g. CDK1-7 and CDK9) [7] and induce cell cycle arrest in G2/M-phase [11]. Thus, meriolins act as a double-edged sword, since on the one hand they inhibit CDKs (i.e. proliferation) and on the other hand they can also induce cell death (i.e. apoptosis). This renders meriolins as valuable anticancer drugs since they simultaneously target two Achilles' heels of the tumor – i.e. unlimited proliferation and inhibition of cell death [21, 22].

Synthesis of meriolin 16, 31 and 36

The synthesis of meriolin 31 and meriolin 36 have been previously described [1]. Meriolin 16 has been synthesized in a similar fashion [8] and the experimental details and characterization are collated in the supplemental information.

Reagents

Meriolin 3 (#445821) and carbonyl cyanide m-chlorophenyl hydrazone (CCCP, #C2759) were obtained from Sigma (Munich, Germany). Etoposide (#1043) was purchased from Biovision (Waltham, MA, USA), staurosporine (STS, #9300) from LC Laboratories (Woburn, MA, USA) and N-(2-Quinolyl)valyl-aspartyl-(2,6-difluorophenoxy)methyl ketone (Q-VD-OPh, QVD, #S7311) from Selleckchem (Houston, TX, USA). TRAIL (#ALX-201-115-C010) was purchased from Enzo Life Sciences (Farmingdale, NY, USA). All other substances for which a manufacturer is not explicitly specified were obtained from Carl Roth.

Cell lines, primary cells and cell culture

Jurkat cells (human T cell leukemia; #ACC-282) were obtained from DSMZ. Ramos cells (human Burkitt B cell lymphoma) were kindly provided by Michael Engelke (Institute of Cellular and Molecular Immunology, University Hospital Göttingen, Göttingen, Germany). HeLa cells (human cervix carcinoma) stably expressing mito-DsRed were kindly provided by Aviva M. Tolkovsky (Department of Clinical Neurosciences, University of Cambridge, England, UK) and have been previously described [54]. Caspase-9-deficient Jurkat cells were kindly provided by Klaus Schulze-Osthoff (Interfaculty Institute for Biochemistry, University of Tübingen, Germany) [55] and retrovirally transduced with either empty pMSCVpuro (Clontech, Heidelberg, Germany) or pMSCVpuro containing cDNAs coding for untagged human wild-type caspase-9 as previously described [37]. Jurkat cells expressing wild-type Bcl-2 and respective vector control cells were kindly provided by Claus Belka (Ludwig-Maximilians University, Munich, Germany) and previously described [56]. Caspase-8 deficient Jurkat cells and the parental cell line A3 were kindly provided by John Blenis (Sandra and Edward Meyer Cancer Center, New York, NY, USA) [36]. The Apaf-1-deficient human melanoma cell line SK-Mel-94 was kindly provided by Maria S. Soengas (Molecular Oncology Program, Spanish National Cancer Research Centre (CNIO), Madrid, Spain) [39].

Transient expression of Smac-mCherry was achieved by lipofection at 70–80% confluence using Lipofectamine 2000 (Life Technologies, Darmstadt, Germany). Briefly, HeLa cells were incubated with 0.15 µl Lipofectamine 2000, 50 ng pcDNA3-Smac(1–60)mCherry and 25 µl optimem per well in glass bottom 8-well chambers (Ibidi, Planegg, Germany) for 16 h. The plasmid was kindly provided by Stephen Tait (Beatson Institute, University of Glasgow, Scotland, UK; plasmid Addgene ID 40880) and has been described previously [57]. The imatinib-sensitive and -resistant cell lines K562 (CML) and KCL22 (CML) were generated as previously described [42]. The cisplatin-sensitive and -resistant cell lines J82 (urothelial carcinoma) and 2102EP (germ cell tumor) were generated as previously described [44, 45]. HeLa, J82, Jurkat, K562, KCL22, Ramos, and SK-Mel-94 cells were cultivated in DMEM medium supplemented with 10% FCS, 100 U/ml penicillin, 100 µg/ml streptomycin, and 10mM HEPES at 37°C and 5% CO₂ in a humidity-saturated atmosphere. 2102EP germ cell tumor cells, cultivated in DMEM (1x) plus GlutaMAX-I medium supplemented with 10% FBS, 1% penicillin/streptomycin (10,000 U) and 1% L-Glutamin (200 mM) at 37°C and 7.5% CO₂, were kindly provided by Christoph Oing (University Medical Center Hamburg-Eppendorf, Germany) and Friedemann Honecker (Tumor and Breast Center ZeTuP, St. Gallen, Switzerland) [43].

Cytotoxicity measurements

For the measurement of cell viability in Ramos cells the resazurin reduction assay (also known as AlamarBlue® assay) was performed. In short, cells were seeded at a certain density depending on the intended incubation time (5 x 10⁵ cells/well for 8 or 24 h, 1 x 10⁵ cells/well for 72 h), treated with ascending substance concentrations and after the specified incubation time, resazurin (Sigma, #R7017) was added to a final concentration of 40 µM. After 120 min of incubation the fluorescence of resorufin (excitation: 560 nm, emission: 590 nm) was measured with a microplate spectrophotometer. Since the reduction of resazurin to resorufin is proportional to aerobic respiration it serves as a measure of cell viability. Alternatively, the triphenyl tetrazolium chloride (XTT) assay was used to evaluate cell viability of 2102EP germ cell tumor cells as described previously [58].

Fluorometric caspase-3 activity assay

Ramos or Jurkat cells were seeded at a density of 1 x 10⁶ cells/ml and treated with the respective agents for the indicated time. Briefly, cells were harvested by centrifugation at 600 g and lysed with 50 µl of ice-cold lysis buffer (20 mM HEPES, 84 mM KCl, 10 mM, MgCl₂, 200 µM EDTA, 200 µM EGTA, 0.5% NP40, 1 µg/ml leupeptin, 1 µg/ml pepstatin, 5 µg/ml aprotinin) on ice for 10 min. Cell lysates were transferred to a flat-bottom microplate and mixed with 150 µl of ice-cold reaction buffer (50 mM HEPES, 100 mM NaCl, 10% sucrose, 0.1% CHAPS, 2 mM CaCl₂, 13.35 mM DTT) containing 50 µM of the profluorescent caspase substrate Ac-DEVD-AMC (Biomol GmbH, Hamburg, Germany, #ABD-13402). The kinetics of AMC release were monitored by measuring AMC fluorescence intensity (excitation: 360 nm, emission: 450 nm) at 37°C in intervals of 2 min over a time course of 120 min, using a Synergy Mx microplate reader. The slope of the linear range of the fluorescence curves (Δrfu/min) was considered as corresponding to caspase-3 activity. Therefore, the mean slope of DMSO treated wells per time point and cell line was set to 100 and all other values were normalized to the corresponding DMSO mean.

FACS-based analysis of apoptotic cell death

The leakage of fragmented DNA from apoptotic nuclei was measured by the method of Nicoletti et al. [25]. Briefly, nuclei were prepared by lysing cells in a hypotonic lysis buffer (1% sodium citrate, 0.1% Triton X-100, 50 µg/ml propidium iodide) and subsequently analyzed by flow cytometry. Nuclei to the left of the 2N peak containing hypodiploid DNA were considered as apoptotic. All flow cytometric analyses were performed on an LSR-Fortessa™ (Becton Dickinson, Heidelberg, Germany).

Immunoblotting

Ramos or Jurkat cells were treated as specified and then harvested by centrifugation (900 g, 5 min) and quick-frozen in liquid nitrogen. Cell pellets were thawed on ice, incubated in lysis buffer [20 mM Tris-HCl, 150 mM NaCl, 1% v/v Triton X-100, 0.5 mM EDTA, 1 mM Na₃VO₄, 10 mM NaF, 2.5 mM Na₄P₂O₇, 0.5% sodium deoxycholate, protease inhibitors (Sigma, #P2714)] for 30 min. Cell lysates were purified from cell debris by centrifugation (17000 g, 15 min) and the protein concentration in the supernatant was determined by Bradford assay. The samples were diluted to a homogeneous protein concentration with sample buffer and both SDS-PAGE and Western blot analyses were performed. Finally, target protein specific primary antibodies [anti-OPA1 [59], anti-PARP1 (Enzo, #BML-SA250), anti-Caspase-8 (Cell Signaling Technology, #9746), anti-caspase-9 (Thermofisher, #PA5-16358), anti-GAPDH (glyceraldehyde 3-phosphate dehydrogenase; Abcam, #ab8245), or anti-tubulin (TUBA4A; Sigma, #T5168) were applied and fluorescence-coupled secondary antibodies (LI-COR Biosciences) were used for protein detection on a PVDF membrane using the LI-COR Odyssey® imaging system.

Microscopy

Imaging of HeLa cells stably expressing mito-DsRed was performed using an Axio Observer 7 inverted microscope (Zeiss) equipped with a Plan-Apochromat 40x/1.4 Oil DIC (UV) VIS-IR M27 oil-immersion objective and a Colibri7 LED light source and using optical sectioning via Zeiss Apotome.2. The cells were seeded on glass bottom 8-well chamber slides (Ibidi, Planegg, Germany) and maintained in full growth medium supplemented with 10 mM HEPES during imaging. Images were taken once per hour and between measurements cells were incubated at 37°C.

Confocal microscopy live imaging of Hela wt cells transiently expressing SMAC-mCherry was performed using a SP8 Leica microscope (Inverse DMI 6000 AFC, Leica) with a C-Apochromat × 63 N.A. 1.2 water DIC objective. Cells were seeded onto µ-Slide 8 wells (IBIDI, 80826) and transfected with SMAC-mCherry as described before [60]. Cells were maintained in full growth medium at 37°C and 5% CO₂ during imaging and recorded every 30 min during 300 min with z-stack acquisition for correct signal detection. Images were analyzed and processed including smoothing and ROI detection at individual frames with Fiji [61]. For each individual time frame, the standard deviation (SD) of the fluorescence intensity of SMAC-mCherry was measured to monitor its subcellular distribution upon meriolin treatment (1 µM) supplemented with 10 µM QVD. A low SD is associated with a homogenous distribution (cytosolic), whereas a high SD is related to a confined accumulation in the mitochondria.

Measurement of mitochondrial membrane potential

To measure changes in mitochondrial membrane potential (ΔΨm), Ramos or Jurkat cells were loaded with the cell-permeable and positively charged dye tetramethylrhodamine ethyl ester (TMRE), which accumulates in active mitochondria characterized by a negative net charge, whereas depolarized mitochondria do not retain the dye. For this purpose, cells were resuspended in fresh medium containing 10 mM HEPES and supplemented with 100 nM TMRE (AAT Bioquest, Sunnyvale, CA, USA; #22220). After incubation for 15 min at 37°C, cells were washed twice with RPMI medium (plus HEPES) and incubated for another 15 min in order to allow the cells to recover and to ensure that the dye had accumulated in active mitochondria. Subsequently, the fluorescence of TMRE was measured by flow cytometry (excitation: 488 nm, emission: 575) with an LSR-Fortessa™ (Becton Dickinson, Heidelberg, Germany). Treatment with the protonophore CCCP served as positive control for complete mitochondrial depolarization.

Replicates and statistical analyses

Experiments were replicated at least three times, and representative data are shown, unless otherwise stated. Error bars indicate standard deviation. All statistical analyses were performed using Prism v7.01 (GraphPad Software, La Jolla, CA, USA).

Acknowledgments

The authors thank Claus Belka (Ludwig-Maximilians University, Munich, Germany), John Blenis (Sandra and Edward Meyer Cancer Center, New York, NY, USA), Michael Engelke (Institute of Cellular and Molecular Immunology, University Hospital Göttingen, Göttingen, Germany), Friedemann Honecker (Tumor and Breast Center ZeTuP, St. Gallen, Switzerland), Christoph Oing (University Medical Center Hamburg-Eppendorf, Germany), Klaus Schulze-Osthoff (Interfaculty Institute for Biochemistry, University of Tübingen, Germany), Maria S. Soengas (Molecular Oncology Program, Spanish National Cancer Research Centre (CNIO), Madrid, Spain), Stephen Tait (Beatson Institute, University of Glasgow, Scotland, UK) and Aviva M. Tolkovsky (Department of Clinical Neurosciences, University of Cambridge, England, UK) for providing valuable cell lines and reagents. This work was supported by grants from the Deutsche Forschungsgemeinschaft [DFG, German Research Foundation; GRK 2158 (to B.S., G.F., S.B., S.W. and T.J.J.M.), 417677437/GRK 2578 (to B.S., G.F. and S.W.), and STO 864/4-3, STO 864/5-1, STO 864/6-1 (to B.S.)] and the Düsseldorf School of Oncology (funded by the Comprehensive Cancer Center Düsseldorf/Deutsche Krebshilfe and the Medical Faculty of the Heinrich Heine University Düsseldorf; to B.S., G.F. and S.W.).

Conflict of interest statement

The authors declare no competing financial interests.

Authorship contributions

L.S. designed the experiments and together with I.H., J.H. and K.K. the cell viability assays, flow-cytometric analyses of apoptotic cell death and mitochondrial membrane potential, fluorometric caspase-3 activity assays, microscopy and immunoblot analyses were performed. D.D. synthesized meriolins 16, 31 and 36. H.F.-R. and A.J.G.-S. analyzed the mitochondrial release of Smac-mCherry in HeLa cells. M.S., G.F., M.A.S. and D.N. performed viability assays with cisplatin-sensitive and cisplatin-resistant J82 urothelial carcinoma and 2102EP germ cell tumor cells, respectively. S.B. provided imatinib-sensitive and -resistant cell lines K562 (CML) and KCI22 (CML). L.S., I.H., K.K., J.H., T.J.J.M., G.F., D.N., C.P., S.B., B.S. and S.W. analyzed and interpreted the data. S.W. wrote the manuscript. S.W. and B.S. supervised the project. All authors discussed the results and commented on the manuscript.

Ethics Statement

Does not apply.

Data availability statement

Data were generated by the authors and included in the article.

Drießen D, Stuhldreier F, Frank A, Stark H, Wesselborg S, Stork B, et al. Novel meriolin derivatives as rapid apoptosis inducers. Bioorg Med Chem 2019;27:3463-8.
Zhang J, Yang PL, Gray NS. Targeting cancer with small molecule kinase inhibitors. Nat Rev Cancer 2009;9:28-39.
Gross S, Rahal R, Stransky N, Lengauer C, Hoeflich KP. Targeting cancer with kinase inhibitors. J Clin Invest 2015;125:1780-9.
Zhang M, Zhang L, Hei R, Li X, Cai H, Wu X, et al. CDK inhibitors in cancer therapy, an overview of recent development. Am J Cancer Res 2021;11:1913-35.
Panagiotou E, Gomatou G, Trontzas IP, Syrigos N, Kotteas E. Cyclin-dependent kinase (CDK) inhibitors in solid tumors: a review of clinical trials. Clin Transl Oncol 2022;24:161-92.
Attwood MM, Fabbro D, Sokolov AV, Knapp S, Schioth HB. Trends in kinase drug discovery: targets, indications and inhibitor design. Nat Rev Drug Discov 2021;20:839-61.
Bettayeb K, Tirado OM, Marionneau-Lambot S, Ferandin Y, Lozach O, Morris JC, et al. Meriolins, a new class of cell death inducing kinase inhibitors with enhanced selectivity for cyclin-dependent kinases. Cancer Res 2007;67:8325-34.
Kruppa M, Müller TJJ. A Survey on the Synthesis of Variolins, Meridianins, and Meriolins-Naturally Occurring Marine (aza)Indole Alkaloids and Their Semisynthetic Derivatives. Molecules 2023;28:
Fresneda PM, Molina P, Bleda JA. Synthesis of the indole alkaloids meridianins from the tunicate Aplidium meridianum. Tetrahedron 2001;57:2355-63.
Echalier A, Bettayeb K, Ferandin Y, Lozach O, Clement M, Valette A, et al. Meriolins (3-(pyrimidin-4-yl)-7-azaindoles): synthesis, kinase inhibitory activity, cellular effects, and structure of a CDK2/cyclin A/meriolin complex. J Med Chem 2008;51:737-51.
Jarry M, Lecointre C, Malleval C, Desrues L, Schouft M-T, Lejoncour V, et al. Impact of meriolins, a new class of cyclin-dependent kinase inhibitors, on malignant glioma proliferation and neo-angiogenesis. Neuro Oncol 2014;16:1484-98.
Alexander A, Karakas C, Chen X, Carey JP, Yi M, Bondy M, et al. Cyclin E overexpression as a biomarker for combination treatment strategies in inflammatory breast cancer. Oncotarget 2017;8:14897-911.
Akli S, Van Pelt CS, Bui T, Meijer L, Keyomarsi K. Cdk2 is required for breast cancer mediated by the low-molecular-weight isoform of cyclin E. Cancer Res 2011;71:3377-86.
Mou J, Chen D, Deng Y. Inhibitors of cyclin-dependent kinase 1/2 for anticancer treatment. Med Chem 2020;16:307-25.
Bharate SB, Sawant SD, Singh PP, Vishwakarma RA. Kinase inhibitors of marine origin. Chem Rev 2013;113:6761-815.
Lukasik P, Baranowska-Bosiacka I, Kulczycka K, Gutowska I. Inhibitors of cyclin-dependent kinases: types and their mechanism of action. Int J Mol Sci 2021;22:2806.
Vogt D, Weber J, Ihlefeld K, Brüggerhoff A, Proschak E, Stark H. Design, synthesis and evaluation of 2-aminothiazole derivatives as sphingosine kinase inhibitors. Bioorg Med Chem 2014;22:5354-67.
Wucherer-Plietker M, Merkul E, Müller TJJ, Esdar C, Knöchel T, Heinrich T, et al. Discovery of novel 7-azaindoles as PDK1 inhibitors. Bioorg Med Chem Lett 2016;26:3073-80.
Chashoo G, Singh U, Singh PP, Mondhe DM, Vishwakarma RA. A marine-based meriolin (3-pyrimidinylazaindole) derivative (4ab) targets PI3K/AKT/mTOR pathway inducing cell cycle arrest and apoptosis in Molt-4 cells. Clin Cancer Drugs 2019;6:33-40.
Lukasik PM, Elabar S, Lam F, Shao H, Liu X, Abbas AY, et al. Synthesis and biological evaluation of imidazo[4,5-b]pyridine and 4-heteroaryl-pyrimidine derivatives as anti-cancer agents. Eur J Med Chem 2012;57:311-22.
Hanahan D. Hallmarks of Cancer: New Dimensions. Cancer Discov 2022;12:31-46.
Hanahan D, Weinberg RA. Hallmarks of Cancer : The Next Generation. Cell 2011;144:646-74.
Drießen D, Biesen L, Müller TJJ. Sequentially catalyzed three-component Masuda-Suzuki-Sonogashira synthesis of fluorescent 2-Alkynyl-4-(7-azaindol-3-yl) pyrimidines: Three palladium-catalyzed processes in a one-pot fashion. Synlett 2021;32:491-6.
Tasch BOA, Antovic D, Merkul E, Müller TJJ. One‐Pot Synthesis of Camalexins and 3, 3′‐Biindoles by the Masuda Borylation–Suzuki Arylation (MBSA) Sequence. European J Org Chem 2013;2013:4564-9.
Nicoletti I, Migliorati G, Pagliacci MC, Grignani F, Riccardi C. A rapid and simple method for measuring thymocyte apotosis by propidium iodide staining and flow cytometry. J Immunol Methods 1991;139:271-9.
Taguchi N, Ishihara N, Jofuku A, Oka T, Mihara K. Mitotic phosphorylation of dynamin-related GTPase Drp1 participates in mitochondrial fission. J Biol Chem 2007;282:11521-9.
Qian W, Choi S, Gibson GA, Watkins SC, Bakkenist CJ, Van Houten B. Mitochondrial hyperfusion induced by loss of the fission protein Drp1 causes ATM-dependent G2/M arrest and aneuploidy through DNA replication stress. J Cell Sci 2012;125:5745-57.
DePinto W, Chu X-J, Yin X, Smith M, Packman K, Goelzer P, et al. In vitro and in vivo activity of R547: a potent and selective cyclin-dependent kinase inhibitor currently in phase I clinical trials. Mol Cancer Ther 2006;5:2644-58.
Ciccia A, Elledge SJ. The DNA damage response: making it safe to play with knives. Mol Cell 2010;40:179-204.
Kuo LJ, Yang LX. Gamma-H2AX - a novel biomarker for DNA double-strand breaks. In Vivo 2008;22:305-9.
White D, Rafalska-Metcalf IU, Ivanov AV, Corsinotti A, Peng H, Lee S-C, et al. The ATM substrate KAP1 controls DNA repair in heterochromatin: regulation by HP1 proteins and serine 473/824 phosphorylation. Mol Cancer Res 2012;10:401-14.
White DE, Negorev D, Peng H, Ivanov AV, Maul GG, Rauscher III FJ. KAP1, a novel substrate for PIKK family members, colocalizes with numerous damage response factors at DNA lesions. Cancer Res 2006;66:11594-9.
Vogler M, Walter HS, Dyer MJS. Targeting anti-apoptotic BCL2 family proteins in haematological malignancies - from pathogenesis to treatment. Br J Haematol 2017;178:364-79.
Van Opdenbosch N, Lamkanfi M. Caspases in Cell Death, Inflammation, and Disease. Immunity 2019;50:1352-64.
Green DR, Llambi F. Cell death signaling. Cold Spring Harb Perspect Biol 2015;7:1-24.
Juo P, Kuo CJ, Yuan J, Blenis J. Essential requirement for caspase-8/FLICE in the initiation of the Fas-induced apoptotic cascade. Curr Biol 1998;8:1001-8.
Manns J, Daubrawa M, Driessen S, Paasch F, Hoffmann N, Löffler A, et al. Triggering of a novel intrinsic apoptosis pathway by the kinase inhibitor staurosporine: activation of caspase‐9 in the absence of Apaf‐1. FASEB J 2011;25:3250-61.
Engels IH, Stepczynska A, Stroh C, Lauber K, Berg C, Schwenzer R, et al. Caspase-8/FLICE functions as an executioner caspase in anticancer drug-induced apoptosis. Oncogene 2000;19:4563-73.
Soengas MS, Capodieci P, Polsky D, Mora J, Esteller M, Opitz-Araya X, et al. Inactivation of the apoptosis effector Apaf-1 in malignant melanoma. Nature 2001;409:207-11.
MacVicar T, Langer T. OPA1 processing in cell death and disease – the long and short of it. J Cell Sci 2016;129:2297-306.
Xiao X, Hu Y, Quirós PM, Wei Q, López-Otín C, Dong Z. OMA1 mediates OPA1 proteolysis and mitochondrial fragmentation in experimental models of ischemic kidney injury. Am J Physiol Renal Physiol 2014;306:1318-26.
Bhatia S, Diedrich D, Frieg B, Ahlert H, Stein S, Bopp B, et al. Targeting HSP90 dimerization via the C terminus is effective in imatinib-resistant CML and lacks the heat shock response. Blood 2018;132:307-20.
Oing C, Verem I, Mansour WY, Bokemeyer C, Dyshlovoy S, Honecker F. 5-Azacitidine Exerts Prolonged Pro-Apoptotic Effects and Overcomes Cisplatin-Resistance in Non-Seminomatous Germ Cell Tumor Cells. Int J Mol Sci 2018;20:
Oechsle K, Honecker F, Cheng T, Mayer F, Czaykowski P, Winquist E, et al. Preclinical and clinical activity of sunitinib in patients with cisplatin-refractory or multiply relapsed germ cell tumors: a Canadian Urologic Oncology Group/German Testicular Cancer Study Group cooperative study. Ann Oncol 2011;22:2654-60.
Höhn A, Krüger K, Skowron MA, Bormann S, Schumacher L, Schulz WA, et al. Distinct mechanisms contribute to acquired cisplatin resistance of urothelial carcinoma cells. Oncotarget 2016;7:41320-35.
Bernardi P, Di F. The mitochondrial permeability transition pore: Molecular nature and role as a target in cardioprotection. J Mol Cell Cardiol 20141-7.
Huang S, Li R, Connolly PJ, Emanuel S, Middleton SA. Synthesis of 2-amino-4-(7-azaindol-3-yl) pyrimidines as cyclin dependent kinase 1 (CDK1) inhibitors. Bioorg Med Chem Lett 2006;16:4818-21.
Murrell JN. The Electronic Spectrum of Aromatic Molecules VI: The Mesomeric Effect. Proc Phys Soc 1955;68:969.
Enari M, Sakahira H, Yokoyama H, Okawa K, Iwamatsu A, Nagata S. A caspase-activated DNase that degrades DNA during apoptosis, and its inhibitor ICAD. Nature 1998;391:43-50.
Sakahira H, Enari M, Nagata S. Cleavage of CAD inhibitor in CAD activation and DNA degradation during apoptosis. Nature 1998;391:96-9.
Weng C, Li Y, Xu D, Shi Y, Tang H. Specific cleavage of Mcl-1 by caspase-3 in tumor necrosis factor-related apoptosis-inducing ligand (TRAIL)-induced apoptosis in Jurkat leukemia T cells. J Biol Chem 2005;280:10491-500.
Herrant M, Jacquel A, Marchetti S, Belhacene N, Colosetti P, Luciano F, et al. Cleavage of Mcl-1 by caspases impaired its ability to counteract Bim-induced apoptosis. Oncogene 2004;23:7863-73.
Roberts AW, Wei AH, Huang DCS. BCL2 and MCL1 inhibitors for hematologic malignancies. Blood 2021;138:1120-36.
Bampton ETW, Goemans CG, Niranjan D, Mizushima N, Tolkovsky AM. The dynamics of autophagy visualized in live cells: from autophagosome formation to fusion with endo/lysosomes. Autophagy 2005;1:23-36.
Samraj AK, Sohn D, Schulze-Osthoff K, Schmitz I. Loss of caspase-9 reveals its essential role for caspase-2 activation and mitochondrial membrane depolarization. Mol Biol Cell 2007;18:84-93.
Rudner J, Lepple-Wienhues A, Budach W, Berschauer J, Friedrich B, Wesselborg S, et al. Wild-type, mitochondrial and ER-restricted Bcl-2 inhibit DNA damage-induced apoptosis but do not affect death receptor-induced apoptosis. J Cell Sci 2001;114:4161-72.
Tait SWG, Parsons MJ, Llambi F, Bouchier-Hayes L, Connell S, Muñoz-Pinedo C, et al. Resistance to caspase-independent cell death requires persistence of intact mitochondria. Dev Cell 2010;18:802-13.
Müller MR, Burmeister A, Skowron MA, Stephan A, Bremmer F, Wakileh GA, et al. Therapeutical interference with the epigenetic landscape of germ cell tumors: a comparative drug study and new mechanistical insights. Clin Epigenetics 2022;14:1-22.
Barrera M, Koob S, Dikov D, Vogel F, Reichert AS. OPA1 functionally interacts with MIC60 but is dispensable for crista junction formation. FEBS Lett 2016;590:3309-22.
Böhler P, Stuhldreier F, Anand R, Kondadi AK, Schlütermann D, Berleth N, et al. The mycotoxin phomoxanthone A disturbs the form and function of the inner mitochondrial membrane. Cell Death Dis 2018;9:286-302.
Schindelin J, Arganda-Carreras I, Frise E, Kaynig V, Longair M, Pietzsch T, et al. Fiji: an open-source platform for biological-image analysis. Nat Methods 2012;9:676-82.
Franco LH, Joffé EBdK, Puricelli L, Tatian M, Seldes AM, Palermo JA. Indole alkaloids from the tunicate aplidium m eridianum. J Nat Prod 1998;61:1130-2.
Trimurtulu G, Faulkner DJ, Perry NB, Ettouati L, Litaudon M, Blunt JW, et al. Alkaloids from the antarctic sponge Kirkpatrickia varialosa. Part 2: Variolin A and N(3′)-methyl tetrahydrovariolin B. Tetrahedron 1994;50:3993-4000.
Perry NB, Ettouati L, Litaudon M, Blunt JW, Munro MHG, Parkin S, et al. Alkaloids from the antarctic sponge Kirkpatrickia varialosa.: Part 1: Variolin b, a new antitumour and antiviral compound. Tetrahedron 1994;50:3987-92.

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Novel meriolin derivatives activate the mitochondrial apoptosis pathway in the presence of antiapoptotic Bcl-2

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Journal Publication

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Abstract

Figures

Introduction

Results

Synthesis of meriolin 16 by Masuda borylation-Suzuki coupling sequence

Meriolins induce DNA damage which is a downstream event mediated by caspases

Meriolin 16, 31 and 36 impair mitochondrial function and structure

Meriolin 16 and 36 induce cytotoxicity in imatinib and cisplatin resistant cancer cells

Discussion

Materials and Methods

Synthesis of meriolin 16, 31 and 36

Reagents

Cell lines, primary cells and cell culture

Cytotoxicity measurements

Fluorometric caspase-3 activity assay

FACS-based analysis of apoptotic cell death

Immunoblotting

Microscopy

Measurement of mitochondrial membrane potential

Replicates and statistical analyses

Declarations

References

Additional Declarations

Supplementary Files

Status:

Journal Publication

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