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Guanidino Anthrathiophenediones as G‑Quadruplex Binders: Uptake, Intracellular Localization, and Anti-Harvey-ras Gene Activity in Bladder Cancer Cells Susanna Cogoi,† Andrey E. Shchekotikhin,‡ Alexandro Membrino,†,⊥ Yuri B. Sinkevich,§ and Luigi E. Xodo*,† †

Department of Medical and Biological Science, P.le Kolbe 4, School of Medicine, 33100 Udine, Italy Gause Institute of New Antibiotics, Russian Academy of Medical Sciences, B. Pirogovskaya, 11, Moscow 119021, Russia § Mendeleyev University of Chemical Technology, 9 Miusskaya Square, Moscow 125190, Russia ‡

S Supporting Information *

ABSTRACT: We prepared a series of anthrathiophenediones (ATPDs) with guanidino-alkyl side chains of different length (compounds 1, 10−13). The aim was to investigate their interaction with DNA and RNA G-quadruplexes, their uptake in malignant and nonmalignant cells, and their capacity to modulate gene expression and inhibit cell growth. Flow cytometry showed that the ATPDs enter more efficiently in malignant T24 bladder cells than in nonmalignant embryonic kidney 293 or fibroblast NIH 3T3 cells. In T24 malignant cells, compound 1, with two ethyl side chains, is taken up by endocytosis, while 12 and 13, with respectively propyl and butyl side chains, are transported by passive diffusion. The designed ATPDs localize in the cytoplasm and nucleus and tightly bind to DNA and RNA G-quadruplexes. They also decrease HRAS expression, increase the cell population in G0/G1, and strongly inhibit proliferation in malignant T24 bladder cells, but not in nonmalignant 293 or NIH 3T3 cells.

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INTRODUCTION Gene expression is normally mediated by transcription factors that bind to cis-elements located in the promoter of the genes. In eukaryotes, several oncogenes contain guanine-rich cis-elements, often located upstream of the transcription start site.1a−d,e These sequences are composed of blocks of guanines that can fold into a quadruplex structure. Quadruplex-forming motifs (QFMs) have been identified in the promoters of numerous oncogenes, including c-MYC,2a KRAS,2b HRAS,2c c-KIT,2d BCL-2,2e and VEGF.2f It has been hypothesized that quadruplex DNA behaves as a repressor element for transcription, but not all the data reported in the literature are in keeping with this notion. For instance, transcription of murine KRAS, Myo-D, and ILPR is favored by G4-DNA.3a,b In a previous study we have demonstrated that the expression of HRAS is regulated by a mechanism involving two quadruplex structures formed by Grich elements overlapping binding sites for the transcription factors MAZ and Sp1.2c We have provided evidence that these two folded structures behave as transcription repressors. About 60% of bladder tumors harbor HRAS mutations,4 and in about half of these tumors HRAS is overexpressed.5 These two genetic alterations play a role in the pathogenesis of bladder cancer.6a It is known that mutated HRAS protein stimulates constitutively the MAPK/ERK pathway leading to cell proliferation.6b,c Therefore, an attractive therapeutic strategy to inhibit HRAS transcription might be using small molecules that bind to G4-DNA and repress transcription. We have previously reported that guanidinium © XXXX American Chemical Society

phthalocyanines (GPcs) bind to the HRAS quadruplexes and down-regulate transcription.2c,7 This is an interesting finding as the down-regulation of HRAS is useful to limit the aggressiveness of bladder cancer cells or, at least, to sensitize them to chemotherapy. However, GPcs could be stronger G4-DNA ligands if they would access the nucleus in a more efficient way. Confocal microcopy studies actually show that these molecules accumulate more in the cytoplasm than in the nucleus.7 We therefore searched for new G4-DNA ligands with a high affinity for HRAS quadruplexes and a high capacity to reach the nucleus. But as cancer cells acquire multidrug-resistance that reduces the efficacy of anticancer drugs, these G4-DNA ligands will have a greater potential in therapy if they tightly bind to their DNA target and escape the drug transporters expressed by cancer cells, i.e., ATP-binding cassette transporters and Pgp.8a−c Among the heteroareneanthracenediones previously prepared by Shchekotikhin et al.,9a−c the pharmacophore groups of the side chains turned out to play a critical role in circumventing multidrug resistance. Studies performed with anthracenedione-based compounds have led to the identification of 4,11-bis[(2aminoethyl)amino]anthra[2,3-b]thiophene-5,10-diones as anticancer agents. The most active molecules in this series were able to attenuate the function of topoisomerase I.9a Moreover, the introduction of guanidinium groups in the side chains of Received: August 31, 2012

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heteroareneanthracenediones significantly enhanced the inhibition of telomerase.9a We subsequently hypothesized that this effect could be due to the binding of the heteroareneanthracenediones to telomeric G4-DNA.10a,b In the present work we have synthesized a series of anthrathiophenediones with two alkyl side chains of different length, each terminating with a guanidinium group. We found that they tightly bind to biologically relevant DNA- and RNAquadruplexes. All the designed molecules showed a good quadruplex to duplex specificity. Since quite little is known about the cellular uptake of anthraquinone derivatives, we tried to understand, by FACS and confocal microscopy, how the designed anthrathiophenediones are transported into cancer and nonmalignant cells. We discovered that the uptake of these molecules by T24 cancer bladder cells depends on the length of the alkyl side chains: compound 1, with two ethyl groups, is transported by endocytosis, while compounds 12 and 13, with two propyl or butyl groups, are transported by passive diffusion. Furthermore, we studied by dual luciferase and Western blots assays the capacity of the anthrathiophenediones to repress HRAS transcription and p21HRAS expression in bladder cancer cells. Together, the results suggest that the designed ATPDs are specific G-quadruplex ligands useful for down-regulating HRAS expression in bladder cancer cells Finally, as the designed ATPDs localize both in the nucleus and cytoplasm of the cells, we asked if quadruplex RNA can also be a target for these molecules. It is known that certain genes are characterized by G-rich 5′-untranslated region (5′-UTR) forming local G-quadruplex structures. The first 5′-UTR quadruplex has been reported by Balasubramanian and coworkers, who discovered that an 18-mer sequence of NRAS mRNA produces a very stable quadruplex that inhibits translation.11 Other studies have demonstrated the presence of quadruplex-forming sequences in several other genes: this gave rise to the hypothesis that such unusual structures might be translation regulating factors.12 Actually, small molecules stabilizing G4-RNA structures located in the 5′-end of the UTR region have been reported to repress translation.13 In the light of this finding the search for ligands with a high affinity for G4-RNA structures may have an important impact on therapy. We interestingly found that the designed ATPDs bind to the 5′UTR G-quadruplex of NRAS with an even higher affinity than that observed with quadruplex DNA. Due to their binding and uptake properties, further investigation and in vivo studies are planned with these very promising compounds.

Scheme 1. Structure of 4,11-Bis[(ωguanidinoethyl)amino]anthra[2,3-b]thiophene-5,10-dione (Compound 1)

The synthesis of the ATPDs was accomplished by a nucleophilic substitution of the alkoxy groups in the periposition of 4,11-dibutoxyanthra[2,3-b]thiophene-5,10-dione with diaminoalkanes (Scheme 2).9c As the alkoxy groups show a different reactivity, the treatment of compound 2 with 1,2ethylendiamine at room temperature for 3 h gave a mixture of products 3−5, where the monosubstitution of the alkoxy groups predominated. This mixture was separated chromatographically on silica gel, and its main components were characterized. NMR showed that 4-amino ATPD 3 was the main product, whereas the 11-amino derivative 4 formed in lesser amounts (Supporting Information, S1). The mono(aminoethyl)amino derivatives 3 and 4 were used to prepare ATPDs with asymmetric side chains: 4-C2/11-C3 and 4-C3/11-C2, where C2 and C3 indicate ethyl and propyl spaces. The reaction between 3 and 4 with 1,3diaminopropane gave anthra[2,3-b]thiophene-5,10-diones 6 and 7, respectively. Instead, the reaction of anthra[2,3b]thiophene-5,10-dione 2 with 1,3-dimainopropane or 1,4diaminobutane gave derivatives containing 1,3-propyl or 1,4butyl (C4) spacers (compounds 8 and 9). The distal amino groups in anthra[2,3-b]thiophene-5,10-diones 6−9 were transformed into guanidino groups by treatment with pyrazolocarboxamidine, yielding the corresponding bis-guanidino derivatives 10−13. All compounds were purified by reprecipitation and gave good analytical and spectroscopic data in full accordance with their assigned structures. The purified compounds were used for physical and cell-based studies. Designed ATPDs Stabilize DNA G-Quadruplexes. To test the capacity of the designed ATPDs to bind to G4-DNA, we performed fluorescence resonance energy transfer (FRET) titration experiments with several quadruplex-forming sequences (QFS) labeled at the 5′ and 3′ ends with carboxyfluorescein (F, donor) and tetramethylrhodamine (T, acceptor), as previously described.7 We used the human telomere sequence (TTAGGG)4 (htelo) and QFSs of the ras genes (HRAS and KRAS) located in the promoter at critical regions controlling transcription.2c,3a In 100 mM KCl, labeled F-htelo-T folds into a mixed parallel− antiparallel quadruplex,14a−c characterized by an energy transfer P of 0.5 [P = ID/(ID + IA), where ID and IA are the emission intensities of the donor and acceptor] (under crowded conditions htelo adopts a parallel G-quadruplex14d,e). Since the ATPDs absorb between 500 and 650 nm (Supporting Information, S2), when they bind to quadruplex F-htelo-T, they quench the fluorescence, in particular the acceptor emission at 580 nm, and this effect was used to determine the affinity of each ligand for the quadruplex.7 A typical titration of quadruplex htelo with compound 13 is shown in Figure 1A, where the addition of increasing amounts of 13 reduces the emission of Fhtelo-T in a dose−response fashion. When a 10-fold excess of ligand was added to the quadruplex solution, the fluorescence

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RESULTS Synthesis of the Anthrathiophenedione Derivatives. We have previously reported that compound 1 with two (guanidinoethyl)amino side chains (Scheme 1) was able to inhibit telomerase.9a This suggests that the tetracyclic thiopheneanthraquinone scaffold of 1 could interact with the quadruplex structure formed by the human telomeric sequence. In the present study we have addressed this hypothesis by designing a series of ATPDs with different combination of (guanidinoalky) amino side chains, which we used for physical and cell-based assays. We reasoned that if one increases the size of the planar chromophore, through the fusion of the anthraquinone core to a thiophene, this should provide more π−π interactions with a Gquartet, while guanidino groups at the end of the side chains should promote electrostatic interactions with the G-quadruplex. B

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Scheme 2. Synthesis of 4,11-Bis(ω-guanidinoalkylamino)anthra[2,3-b]thiophene-5,10-diones 10−13a

Reagents and conditions: (a) ethylenediamine, THF, 30 °C, 2.5 h; (b) 1,3-diaminopropane, THF, 50 °C, 2 h; (c) 1,4-diaminobutane, THF, 50 °C, 2 h; (d) pyrazole-1-carboxamidine hydrochloride, EDIA, DMSO, 60 °C, 5 h.

a

best-fitted to the Hill equation ( f = [L]n/(KD + [L]n), where f is the fraction of filled quadruplex sites, L is the ligand, KD the apparent dissociation constant, and n is the Hill coefficient describing cooperativity. KD of 0.18 ± 0.013 μM and n = 1.5 were obtained. The saturation of quadruplex sites with 13 was obtained at 2 equiv of ligand per quadruplex, suggesting the binding of one ligand to the external G-quartets. The same analysis was performed with all designed ATPDs, and the binding data obtained are collected in Table 1. Note that the ligands show Table 1. Binding data of the interaction between quadruplexes htelo and KRAS and designed ATPDs, in 50 mM Tris-HCl, pH 7.4, 100 or 10 mM KCl ligand

htelo KD (μM)

hteloa ΔTM (K)b, r = 5

KRAS KD (μM)

KRASc ΔTM (K),d r = 5

1 10 11 12 13

0.15 ± 0.01 0.32 ± 0.03 0.33 ± 0.05 0.13 ± 0.01 0.18 ± 0.01

14 21 23 21 34

0.25 ± 0.02 0.21 ± 0.01 0.23 ± 0.02 0.18 ± 0.01 0.27 ± 0.01

15 27 28 33 35

TM of quadruplex htelo in 100 mM KCl is 50 °C. bΔTM ± 0.5 K is the difference in 100 mM KCl of the quadruplex in the absence and presence of ligand (r = 5). cTM of quadruplex KRAS in 10 mM KCl is 60 °C. dΔTM ± 0.5 K at r = 5. a

Figure 1. (A) FRET-titration of 200 nM quadruplex htelo with increasing amounts of compound 13. Inset shows the fractions of quadruplex htelo bound to 13 and best-fit of the experimental points to the Hill equation. (B) FRET-melting curves of quadruplex htelo in the presence of increasing amounts of compound 13, at various r values (r = [ligand]/[quadruplex]). The experiments have been performed in 50 mM Tris-HCl, pH 7.4, 100 mM KCl.

a rather high affinity for quadruplex htelo, the KD’s being between 0.15 and 0.33 μM. It has been recently reported that, for the interaction of quadruplex htelo with bis-indole carboxamides, KD values are between 2 and 27 μM; for the interaction of htelo with a trisubstituted isoalloxazines, this value is 8.6 μM.15a,b The stabilization of quadruplex htelo by the ATPDs was examined by FRET-melting experiments. In 100 mM KCl, htelo melts with a cooperative transition with a TM of 50 °C. The stabilization induced by the ATPDs depends on the r value, where r= [ligand]/[quadruplex]. For instance, at r = 1, 2, 3, 4 and 5, compound 13 increases the TM from 50 to 58.5, 76.3, 79.1, 81.1, and 83.7 °C, respectively (Figure 1B). The ligand-induced change in quadruplex TM, ΔTM, at r=5 (1 μM ligand) caused by

emission was quenched. As previously demonstrated, the ligand is able to quench the fluorescence only when it is bound to the quadruplex, i.e., at a distance of few angstroms from the quadruplex fluorophores.7 From the fluorescence spectra we calculated the fraction of quadruplex sites filled with 13 as a function of ligand concentration. The binding data obtained were C

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Next, we wanted to investigate the affinity of the ligands for duplex DNA. We therefore performed FRET-melting competition assays, in which a dual-labeled quadruplex is melted in the presence of an ATPD ligand and an excess of duplex DNA. We first determined the binding stoichiometry of the ATPDs for duplex DNA by means of UV−vis titrations. A UV−vis titration of compound 1 with a 32mer duplex is reported in Supporting Information, S4. We found an average stoichiometry of 1 ATPD per 7 base pairs of duplex. We then performed FRET-melting completion experiments where the number of duplex sites were 37- and 75-fold in excess over the quadruplex sites. The results showed that the TM did not practically change upon addition of 37- and 75-fold duplex competitor (Supporting Information, S5). This finding suggests that all ligands are characterized by a good quadruplex over duplex specificity and can, therefore, be used to target quadruplex DNA in cell-based experiments. The binding data obtained can be summarized as follows: (i) The designed ATPDs form a family of ligands that strongly stabilize quadruplex DNA, with KD’s between 0.13 ± 0.01 and 0.37 ± 0.11 μM, and ΔTM from 14 to 34 °C. (ii) ΔTM with all G4-DNA roughly increases with the length of the alkyl side chains, which probably interact with the loops and grooves of the quadruplex. (iii) Compound 13 is the most efficient stabilizer, causing a ΔTM between 26 and 35 K (r = 4 or 5) with all the four quadruplexes; compound 1 is instead the least efficient stabilizer with ΔTM between 14 and 22 K. (iv) ATPDs show a good quadruplex over duplex specificity. (v) The binding curves are nicely described by the Hill equation, suggesting that ATPDs

1, 10-13 are 14, 21, 23, 21, and 34 K, respectively (Table 1). It should be borne in mind that strong stabilizers of the human telomeric G-quadruplex are pyridostatin (ΔTM 35 K),16 triarylpyridines with C4-pyrrolidine/C4-piperazione side chains (ΔTM up to 30 K),17a macrocyclic quinacridine-based compounds (ΔTM of 28 K)17b and phthalocyanine derivatives (ΔTM up to 40 K).17c The stabilizing capacity of the ATPDs was also assessed with two QFSs, hras-1 and hras-2, located in the HRAS promoter and behaving as transcription repressors,2c and with a critical QFS of the murine KRAS promoter3a (Supporting Information, S3) (Tables 1 and 2). Table 2. Binding Data of the Interaction between Quadruplexes hras-1 and hras-2 and the Designed ATPDs in 50 mM Tris-HCl, pH 7.4, 100 (or 10) mM KCl ligand

hras-1 KD (μM)

hras-1a ΔTM (K),b r = 5

hras-2c KD (μM)

hras-2 ΔTM (K),d r = 4

1 10 11 12 13

0.34 ± 0.07 0.36 ± 0.02 0.28 ± 0.01 0.30 ± 0.03 0.21 ± 0.04

12 and 22 13 and 23 16 and 26 19 and 29 20 and 32

0.27 ± 0.02 0.37 ± 0.11 0.28 ± 0.02 0.23 ± 0.08 0.28 ± 0.02

19 17 12 17 26

TM of quadruplex hras-1 in 100 mM KCl is byphasic 52 and 62 °C. ΔTM ± 0.5 K is the difference in 100 mM KCl of the quadruplex in the absence and presence of ligand (r = 5). cTM of quadruplex hras-2 in 10 mM KCl is 70 °C. dΔTM ± 0.5 K at r = 4. a b

Figure 2. (A) FACS analysis of T24 bladder cancer cells incubated for 8 h with increasing amounts of ATPDs (0.25, 1, 2.5, 5, and 10 μM). (B) (Top) Uptake in T24 cancer cells of compounds 1 and 10−13 (0.25, 1, 2.5, 5, and 10 μM) after 8 h incubation; (bottom) uptake in T24 cancer cells of compounds 1 and 10−13 (10 μM) at 0.25, 1, 2, 6, and 8 h incubation times. D

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Figure 3. (A) Uptake in malignant T24 bladder cells, nonmalignant murine NIH 3T3, and human 293 cells of ATPDs (0.25, 1, 2.5, 5, and 10 μM) after 8 h incubation. (B) Uptake in malignant T24 and nonmalignant NIH 3T3 and 293 cells of compound 1 (10 μM) at various incubation times, up to 8 h. (C) FACS analysis of T24 cancer cells treated with 80 μM dynasore and 5 μM compounds 1 and 10−13 for 4 h. Peaks from left to right: nontreated cells; cells treated with dynasore alone; cells treated with compound 1 and dynasore; cells treated with anthrathiophenedione. (D) Uptake in T24 cancer cells of 1 and 13 at 4 and 37 °C. Peaks from left to righ: nontreated cells at 37 °C, nontreated cells at 4 °C; cells treated with compound 1 or 13 at 4 °C; cells treated with compound 1 or 13 at 37 °C.

incubation of 15 min, 1, 2, 6, and 8 h. The result shows that the uptake reaches its peak in 6 h. Flow cytometry experiments were also performed with nonmalignant murine NIH 3T3 and human embryonic 293 cells. The uptake in these cells was significantly lower than in malignant T24 bladder cells, up to 10-fold with compound 1 (Figure 3A,B). The unexpected higher uptake of compound 1 compared to the more hydrophobic 12 and 13, particularly in T24 cancer cells, led us to hypothesize that the latter may aggregate in solution. But when we observed that 1 and 13 obey the Lambert−Beer law over a large concentration range (1−100 μM), we ruled out this possibility (Supporting Information, S7). As a second step we wanted to see if the ATPDs enter into the cells by passive diffusion or by active transport. To investigate whether the uptake occurs by endocytosis, T24 cancer cells were treated with dynasore, an inhibitor of the GTPase activity of dynamin which blocks the budding of the endocytotic vesicles from the membrane,20 before the uptake of the ATPDs was assessed by flow cytometry. As illustrated in Figure 3C, the uptake of compound 1 was clearly inhibited by dynasore (FL3-H was reduced from 60 to 20), which was not the case with the other compounds, except for 11. This suggests that the uptake of 1, the

bind to the quadruplexes in a cooperative manner, with a Hill coefficient varying from 1.5 to 3.2. ATPD Uptake Occurs by Endocytosis and/or Passive Diffusion. Recent data support the notion that G-quadruplex structures are involved in the regulation of transcription and translation.18a−c Ligands that specifically bind to G4-DNA can be used to modulate transcription,19 providing that they efficiently penetrate cell membranes and access the nucleus. To this aim we investigated the uptake of ATPDs in human T24 bladder cancer cells and nonmalignant murine NIH 3T3 as well as human 293 embryonic cells. As ATPDs emit red fluorescence when they are excited at 567 nm (Supporting Information, S2), their uptake was investigated by flow cytometry. Figure 2A,B shows the ATPD uptake in T24 cancer cells incubated for 8 h with increasing amounts of 1 and 10−13 (0.5, 1, 2.5, 5, 10 μM). The uptake increases with ligand concentration and, surprisingly, decreases with the length of the side chains (C2, C3, or C4) as follows: 2C2 < C3/C2 ∼ C2/C3 < 2C3 < 2C4. Compounds 1, 10, and 11, with C2 or C2/C3 side chains, are taken up to 10-fold more than 12 and 13, with C3 and C4 side chains, respectively. The uptake was investigated also as a function of time (Supporting Information, S6). Figure 2B shows the uptake of compound 1 (0.5, 1, 2.5, 5, 10 μM) in T24 cancer cells after an E

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the nucleus and cytoplasm of T24 cancer cells. As the ATPDs emit red fluorescence, their intracellular distribution was followed by irradiating the cells with a laser at 543 nm. Panels D−I show a typical distribution of compounds 1 and 11 (2.5 μM) in T24 cancer cells after an incubation of 2 h. Sectioning at 2 μm intervals, the Z-axis images of cells treated with each ATPD showed that the fluorescence was intracellular, localized in the cytoplasm and nucleus. No significant difference was observed between cancer T24 cells and nonmalignant NIH 3T3 cells (and 293 cells, not shown). Compared to propidium iodide which appears homogenously distributed in the nucleoplasm (panel B), compounds 1, 11 (panels E, H) and 10, 12 (Supporting Information, S8) show a granular nuclear distribution with accumulation of ATPD in the nucleoli. In contrast, compound 13 shows a more homogeneous distribution in the nucleoplasm. Confocal microscopy experiments have been performed also with nonmalignant 293 and NIH 3T3 cells. In the latter, the appearance of nucleolar bodies appears particularly evident (Figure 5) (Supporting Information, S9).

least hydrophobic molecule with C2 side chains, should occur mainly by clathrin- or caveola-mediated endocytosis, while the uptake of the other, more lipophilic compounds 10, 12, 13 should occur primarily by passive diffusion. The uptake of compound 11, which has 4-C3/11-C2 side chains and is slightly inhibited by dynasore, is likely to occur by both endocytosis and passive diffusion. The data show that endocytosis is a more efficient transport than passive diffusion, as 1 is taken up in T24 cancer cells, respectively, 3-, 5- and 10-fold more than 10, 12, and 13. Both types of passive and active transport should be temperature-dependent, because at temperatures under 37 °C both the membrane fluidity and the ATP level are reduced. We analyzed the uptake of the ligands at 4 and 37 °C and found indeed that it significantly decreased at a lower temperature. Figure 3D reports a typical FACS analysis at 4 and 37 °C of T24 cancer cells treated with 1 (with 2 C2) and 13 (with 2 C4), which enter into the cells by, respectively, endocytosis and diffusion. It can be seen that at 4 °C the uptake of both molecules decreases: endocytosis of 1 is more temperature-dependent than passive diffusion of 13. The uptake of the ATPDs by nonmalignant NIH 3T3 cells did not respond to dynasore or cytochalasin D, another inhibitor of endocytosis.21 Our conclusion was therefore that it occurs by passive diffusion. ATPD Intracellular Localization. To investigate the intracellular localization of the ligands, we used confocal microscopy. Figure 4 shows T24 cancer cells in which the cytoplasm was stained by immunofluorescence with a primary antibody specific for dynamin-related protein 1 (DRP1) (panel A), while the nucleus was stained with propidium iodide (PI) (panel B). Panel C shows the merge and gives a clear picture of

Figure 5. Confocal microscopy of nonmalignant NIH 3T3 cells treated for 4 h with 2.5 μM of ATPDs. Panels A−C show the cells with the cytoplasm stained with DRP1, the nucleus with propidium iodide. Panels D−I show NIH 3T3 cells treated for 2 h with 2.5 μM ATPDs 1 and 11 which emit red fluorescence upon excitation at 543 nm and with the cytoplasm stained in green with DRP1.

ATPDs Bind to RNA G-Quadruplex. Since the designed ATPDs accumulate in nucleous and cytoplasm, we asked if these molecules are also able to bind to RNA quadruplexes. Recent studies suggest that the identification of ligands that efficiently penetrate the cell membrane and tightly bind to G4-RNA may be therapeutically important. In fact, these molecules can be used to target the 5′ untranslated region (5′-UTR) of certain genes that contains quadruplex-forming sequences that are involved in the regulation of translation. NRAS11 and KRAS22 are protooncogenes whose mRNA contains UTRs with G-rich quadruplex forming sequences. To assess the binding to quadruplex RNA, we considered the quadruplex-forming sequence present in the 5′-UTR of NRAS. The probable involvement of this structure in translation regulation has been demonstrated.11 The NRAS

Figure 4. Confocal microscopy of T24 bladder cancer cells treated for 2 h with 2.5 μM of ATPDs. Panels A−C show the cells in which the cytoplasm was stained with a fluorescein-labeled secondary antibody recognizing a primary antibody specific for cytoplasmatic DRP1 protein (A), the nucleus was stained with propidium iodide (B). Panel C shows the merge of panels A and B. Panels D−I show T24 cells treated for 2 h with 2.5 μM ATPDs 1 and 11 which emit red fluorescence upon excitation at 543 nm; the cytoplasm was stained in green with DRP1. F

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Figure 6. FRET-titration in 50 mM Tris-HCl, pH 7.4, 10 mM KCl of 200 nM RNA G-quadruplex NRAS with increasing amounts of compound 1 (r = 1−5). Right panel shows the fractions of RNA G-quadruplex bound to 1 and the relative best-fit to the Hill equation. Bottom panel shows FRET-melting curves of the NRAS quadruplex in the presence of compound 1 at r values of 0, 1, 3, and 5.

in our laboratory, carrying f iref ly luciferase driven by the HRAS promoter (pHRAS-luc).2c As illustrated in Figure 7A, the HRAS promoter, upstream of the transcription start site (TSS), contains two quadruplex-forming G-elements, hras-1 and hras-2, which behave as transcription repressors.2c DMS-footprinting and CD showed that hras-1 folds into an antiparallel structure composed by three G-tetrads and 4−4−3 loops, while hras-2 folds into a parallel quadruplex with 1−4−1 loops (Supporting Information, S10).2c We also demonstrated by ChIP and EMSA that these two quadruplexes are bound by MAZ and Sp1, which synergistically activate transcription.2c These two proteins recognize two contiguous blocks of guanines separated by C/A (MAZ) and C (Sp1).23There are two perfect “GGGCGGG” sites within hras2 and one site in hras-1 where however one G is replaced by C, “GGGCGGC”. To account for transfection efficiency, the cells were cotransfected with plasmid pRL-CMV, in which Renilla luciferase was driven by the cytomegalovirus promoter. The optimal pHRAS-luc/pRL-CMV ratio used for the cotransfections was 25 parts of pHRAS-luc and 1 part of pRL-CMV. We used as a control a mutant plasmid (pHRAS-mut), generated by the wild-type pHRAS-luc, where we introduced 4 point mutations in hras-1/hras-2, in order to prevent their folding into quadruplex structures. In choosing the sites to introduce the point mutations, we left intact one MAZ/Sp1 binding site in each G-element, allowing us to assume that the difference in luciferase expressed by the two plasmids is a consequence of quadruplex formation. We have already reported that guanidino phthalocyanines bind to the HRAS quadruplexes and down-regulate HRAS promoter activity.2c Plasmid pHRAS-luc is therefore an appropriate system for evaluating the capacity of the designed ATPDs to inhibit HRAS transcription. To this end, we performed dual-luciferase assays following the scheme illustrated in Figure 7B. The cells were first treated for 24 h with increasing amounts of ATPDs (0.5, 2.5, and 5 μM), and then transfected with a 25:1 mixture of pHRAS-luc and pRL-CMV, in the presence of jet-PEI (Polyplus) as a transfectant agent. 48 h after transfection, f iref ly

sequence, 5′-GGGAGGGGCGGGUGGG, labeled with FAM and TAMRA, formed in 10 mM KCl a very stable quadruplex, characterized by a P-value of 0.67 and TM =77.2 °C. Figure 6 shows a representative FRET-titration of quadruplex NRAS with ATPD 1, the relative binding curve (giving a KD of 0.38 ± 0.07 μM) and FRET-melting at r = 1, 2, 3, and 5 giving TM’s of 81.9, 86.3, and 89.2 °C. The binding curves indicate a binding stoichiometry of 1 ligand per quadruplex (for G4-DNA we found two ligands/quadruplex). Due to the high TM of the NRAS quadruplex in 10 mM KCl (TM =77.2 °C), we measured ΔTM in the presence of an equimolar amount of each ligand (at r = 1) and found that ΔTM varied from 5 (compound 1) to >13 K (compound 13). These values are higher than those observed with hras-1 and hras-2 DNA quadruplexes (Table 3). ATPDs Efficiently Modulate Gene Expression. A first evaluation of the ATDPs’ capacity to modulate gene expression was obtained by dual-luciferase reporter assays. For these experiments we used a reporter plasmid, previously constructed Table 3. Binding Data of the Interaction between NRAS G4RNA and the Designed Anthrathiophenedione Derivatives, in 50 mM Tris-HCl, pH 7.4, 10 mM KCl

ligand

NRAS G4RNA KD (μM)

NRASa G4-RNA TM(°C), r =1

NRAS G4RNA ΔTM (K),b r = 1

hras-2 G4DNA ΔTM(K),b r=1

hras-1 G4DNA ΔTM (K),b,c r = 1

1 10 11 12 13

0.38 ± 0.07 0.12 ± 0.01 0.09 ± 0.01 0.11 ± 0.01 0.06 ± 0.01

82 86 88 88 >90

5 9 11 11 >13

3 4 3 3 7

5 5 8 5 7

TM of NRAS in 10 mM KCl is 77.2 °C. bΔTM ± 0.5 K is the difference of the quadruplex in the absence and presence of ligand (r = 1). cThe ΔTM is relative to the second transition of the quadruplex (TM = 62 °C). a

G

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Figure 7. (A) Sequence of HRAS promoter upstream of TSS. The QFSs hras-1 and hras-2 are indicated with brackets, while the MAZ binding sites are indicated with gray rectangles. The G > T point mutations in hras-1 and hras-2 that abrogate quadruplex formation at these G-elements are indicated. (B) Transfection scheme of the dual-luciferase assays. Dual luciferase assay performed at 72 h post-transfection with wild-type plasmid pHRAS-luc, in which f iref ly luciferase was driven by the human HRAS promoter, in the presence of the ATPDs 1 and 10−13 at 0.5, 2.5, and 5 μM. Ordinate reports the “Relative luciferase”, i.e., T/C × 100 where T is (f iref ly luciferase)/(Renilla luciferase) in the treated cells, while C is the same ratio in untreated cells. (C) (Top) Dual luciferase assay as in B but with the mutant vector pHRAS-mut; (bottom) dual luciferase assay as in B but with pIL-luc, where f iref ly luciferase is driven by the interleukin-8 promoter that does not contain QFSs, and ATPDs (5 μM). (D) Western blot showing the level of HRAS and βactin proteins in T24 bladder cancer cells treated with 5 μM ATPDs for 48 and 72 h. The HRAS protein level is given by T/C × 100, where T = p21HRAS/ β-actin in treated cells and C = p21HRAS/β-actin in nontreated cells.

and Renilla luciferases were measured with a luminescence plate reader. The results obtained are reported in a histogram. When T24 cells are pretreated with ATPD (0.5, 2.5, and 5 μM), the luciferase expression by wild type pHRAS-luc is down-regulated, roughly in a dose−response manner (Figure 7B). Under the

experimental conditions adopted (72 h post transfection), compounds 1, 10, and 13, which show the highest uptake, reduce luciferase to ∼25% of the control (untreated cells), while compounds 12 and 13, showing a lower uptake, reduce it to ∼50% of the control. As expected, the effect of the designed H

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ATPDs on luciferase expression by mutant pHRAS-mut was insignificant, as the G-elements could not fold into G4-DNA. As a second control, we used an expression vector, pIL-luc, carrying f irefly luciferase driven by the interleukin-8 promoter, whose sequence does not contain quadruplex-forming sequences. Again, the designed ATPDs showed no repression of luciferase (Figure 7C). Together, our data demonstrate that the designed ATPDs down-regulate the activity of the HRAS promoter by stabilizing the hras-1 and hras-2 quadruplexes. Next, we asked if the designed ATPDs are capable to reduce the level of protein HRAS (p21HRAS) in malignant T24 bladder cells. To address this question we measured p21HRAS by immunoblotting, using a monoclonal anti-HRAS antibody (HRAS-Ab). The cells were treated with ATPD (5 μM), and the extracted proteins were run in SDS-PAGE and analyzed by immunoblotting with HRAS-Ab. Since the half-life of human p21RAS varies from 20 to 36 h,24 the immunoblot was carried out at 24, 48, and 72 h. At 24 h the level of p21RAS was roughly the same in the treated and untreated T24 cells (not shown). But at 72 h, the protein was reduced to a level between 50% and 20% of the control (untreated cells) (Figure 7D). Instead, the immunoblot at 48 h after treatment showed that only compounds 1 and 13 reduced the protein. This correlates with the higher uptake of the former and the higher affinity for G4DNA of the latter. Considering that the designed ATPDs enter more efficiently into malignant T24 bladder cells than in nonmalignant 293 and NIH 3T3 cells, and that they efficiently down-regulate the expression of mutant HRAS (the genetic lesion responsible for the malignant phenotype of T24 bladder cells), we feel that they have potential as anticancer drugs for bladder tumors. We therefore investigated the ATPD capacity to impair proliferation in T24 cancer cells. ATPDs Inhibit the Growth of Bladder Cancer Cells. Mutant HRAS encodes for a hyperactivated protein, p21HRAS, that constitutively stimulates the MAPK/ERK pathway giving a selective growth advantage to T24 bladder cancer cells. A decrease in p21HRAS should therefore result in less proliferation. This has indeed been observed in ovarian cancer cells after silencing of HRAS gene expression by retrovirus mediated siRNA,25a and in pancreatic cancer cells (Panc-1) after KRAS suppression by G4-DNA decoy molecules.25b To test their effect on cell growth, the ATPDs were incubated, at the concentrations of 1, 2.5, 5, 10, 20, and 30 μM, with mutant HRAS T24 cancer cells as well as with wild-type HRAS 293 and NIH 3T3 nonmalignant cells for 72 h: at this time point p21HRAS is reduced to 20% of the control by the quadruplex-interactive drugs. The results of a resazurin assay are reported in Figure 8. The drug concentration that reduces proliferation by 50% compared to nontreated cells (IC50) is reported in Table 4. It can be seen that the designed ATPDs significantly reduce the proliferation of malignant T24 bladder cells but not of nonmalignant 293 and NIH 3T3 embryonic cells. The IC50 in T24 cancer cells of compounds 1, 10, and 11 varies from 4 to 7 μM, whereas in 293 and NIH 3T3 cells, IC50 is ∼30 μM or even higher. These results are in keeping with literature data25 and with the fact that ATPDs are taken up more by malignant than by nonmalignant cells. We next investigated if the inhibition of cell growth is associated with changes in the cell cycle. We carried out FACS experiments with mutant HRAS T24 and wild-type HRAS 293 cells, at various time points following ATPD treatment. At 24 h, the designed ATPDs did not affect the cell cycle of wild-type HRAS 293 cells, causing however a slight arrest in G2/M of

Figure 8. Percent viability of malignant T24 bladder cells and nonmalignant murine NIH 3T3 and human 293 cells treated with increasing amounts of ATPDs (0, 1, 2.5, 5, 10, 20, and 30 μM). Ordinate reports % viability of treated cells compared to nontreated cells. The data are the average of three independent measurements.

Table 4. IC50 Values of ATPDs in Malignant T24 and Nonmalignant 293 and NIH 3T3 Cell Lines

a

ligand

T24 IC50 (μM)a

293 IC50 (μM)a

NIH 3T3 IC50 (μM)a

1 10 11 12 13

4.4 7.9 6.5 19.5 19.1

>30 20−30 >30 >30 >30

20−30 >30 20−30 >30 >30

Values determined in cells incubated for 72 h with ATPD.

mutant HRAS T24 cancer cells (Supporting Information, S11). In contrast, at 72 h post-treatment (and also 48 h), the ATPDs induced a statistically significant increase of G0/G1 (from 50 ± 3% to 63 ± 3%) and concomitant decrease of G2/M (from 20 ± 3% to 12 ± 1%) in T24 cancer cells, but not in wild-type HRAS nonmalignant 293 cells (Figure 9A,B). These results are in agreement with the study by Liu and co-workers,25a who showed that the silencing of mutant HRAS in ovarian cancer cells increases the cell population in G0/G1. It is noteworthy that the changes in the cell cycle caused by the ATPDs at 24, 48, and 72 h correlate with the level of p21HRAS in the cells. The increase of cell population in G2/M occurs at 24 h, i.e., when the intracellular level of the p21HRAS is not yet reduced, as indicated by the immunoblots (the ATPDs may initially affect the cells by interfering with metabolizing enzymes such as topoisomerase 2, as etoposide does, Supporting Information, S11). Instead, at 48 and 72 h, i.e., when the ATPDs suppress the protein by blocking the HRAS promoter (see immunoblots), the cells appear arrested in G0/G1, as was expected. I

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Figure 9. Typical FACS analysis of malignant T24 bladder cancer cells (A) and nonmalignant 293 cells (B) untreated and treated for 72 h with 10 μM compounds 1−13. The histograms in the right show the percentage of cells in the G0/G1, S, and G2/M phases of the cell cycle. A standard t-test versus control was performed (*p < 0.05).

Finally, as the treated T24 cells remain substantially viable, as judged from the absence of a pre-G0/G1 peak (not shown), the ATPDs reduce cell proliferation by a cytostatic mechanism. To further support this conclusion, we measured by trypan blue the percentage of dead cells in ATPD-treated T24 cells and found that the drugs did not increase this percentage. We also found, on the basis of a caspase 3/7 assay, that the ADTPs did not induce apoptosis. These data together suggest that the designed ATPDs inhibit cell growth through a cytostatic mechanism mediated by a G0/G1 cell cycle arrest.

In this study, we designed ATPDs with alkyl side chains terminating with guanidino groups. Besides demonstrating that they bind with a high affinity to DNA- and RNA G-quadruplexes, we investigated their uptake, their intracellular localization, as well as their capacity to repress gene expression. To enhance the G-quadruplex affinity, we extended the aromatic surface of the chromophore by fusing a thiophene to the anthraquinione nucleus, and introduced alkyl side chains terminating with a guanidino group. We expected that the tetracyclic core should promote π−π stacking interactions with the external G-tetrads of the quadruplex, while the cationic side chains should bind to its loops or grooves. We actually found that the designed ATPDs enhanced strongly the affinity for G4-DNA, as we obtained ΔTM values for quadruplex htelo of 13 up to >30 K (ligand 1 μM), which is significantly higher than the values reported for aminoacyl anthraquinone conjugates (ΔT from 0 to 9.8 K, ligand 1 μM)32 and for disubstituted peptidyl anthraquinones with a lysyl terminal residue (ΔT up to 20 °C33). Wender et al. proposed that the guanidino group should play a role in cellular uptake, as they discovered that polyarginine is taken up more efficiently than polylysine.34 Against this background we investigated the uptake of a derivative of compound 1, where guanidine NH2 was replaced with a methyl chloride. Surprisingly, we found that this modification increased the uptake, leading us to conclude that the guanidino group cannot be a critical uptake determinant for ATPDs. Instead, a major determinant of the uptake is the hydrophobic character of the side chains. Indeed, most hydrophobic compounds with C3 or C4 side chains are transported into the cells by passive diffusion. Instead, compound 1 with C2 side chains and compound 11 with C2/ C3 side chains, being less hydrophobic, enter into malignant T24 cells by endocytosis: an active transport sensitive to dynasore and temperature. It is plausible that this behavior is common to other ligands bearing side chains with different hydrophobicity.

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DISCUSSION The basic anthraquinone core is present in some clinically used anticancer drugs including adryamicyn and mitoxantrone.26 Early studies have suggested that the anticancer activity of these molecules is associated to their capacity to bind to duplex DNA.27 The search for new anthraquinone derivatives with enhanced biological activity has continued over the past years, and recently Xie et al. reported that a marine anthraquinone (SZ685C) is a potent inducer of apoptosis with anticancer activity.28 In 1997 Sun et al. were the first to discover that human telomerase was inhibited by a diamidoanthraquinone.29 They provided evidence that the compound inhibited telomerase by a mechanism consistent with the interaction between the diamidoanthraquinone and the telomere G-quadruplex.30 Subsequent studies reinforced the G-quadruplex hypothesis showing that several other molecules, including acridines, fluorenones, phenanthrolines, and tetracyclic benzonaphthofurandiones, gave telomerase activity.31 Neidle and co-workers reported that, in addition to the anthraquionone core itself, important determinants for enhancing the binding to quadruplex DNA are side chains terminating with a cationic end group as well as the extension of the tricyclic chromophore.31 J

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Another interesting finding of our study is that malignant T24 bladder cells take up ATPDs up to 10-fold more efficiently than nonmalignant murine NIH 3T3 and human 293 embryonic cells. This suggests that the enhanced permeability and retention of tumors compared to normal tissues is due not only to the production of permeability factors and abnormalities in the tumor vasculature but also to different internalization mechanisms of malignant cells compared to nonmalignant cells.35 Considering that ATPDs show (i) a high affinity for DNA- and RNA-quadruplexes; (ii) a good quadruplex-to-duplex specificity; and (iii) a high uptake (in particular compound 1) and nuclear/ cytoplasm localization, they can be used to target DNA and RNA. We already had reported in a previous study that the transcription of HRAS is regulated by a mechanism involving two neighboring DNA quadruplexes formed by G-rich elements interacting with transcription factors MAZ and Sp1.2c We had also found, by a reporter assay, that guanidino phthalocyanines stabilize the quadruplexes and repress the activity of the HRAS promoter. In our present study, we describe that compounds 1, 10, and 11, which show a good uptake in T24 cancer cells, reduce the luciferase level in a dose−response manner, up to ∼20% of the control. Instead, compounds 12 and 13, which are taken up less efficiently, reduce luciferase to a lower extent (∼50% of the control). There is a clear correlation between ATPD uptake and luciferase expression. When the ATPDs were used to target HRAS at genomic level, they significantly repressed p21HRAS in malignant T24 bladder cells. Although compounds 12 and 13 showed a lower uptake in T24 cells than 1, 10, and 11, they also reduced p21HRAS. This might be because 12 and 13 stabilize quadruplexes hras-1 and hras-2 more than 1, 10, and 11. It clearly demonstrates that the activity of the ATPDs depends on several variables including binding affinity to G4-DNA, cellular uptake, and capacity to reach the target in the chromatin environment. The designed ATPDs were also found to stabilize a naturally occurring RNA G-quadruplex formed within the 5′ UTR of the human NRAS proto-oncogene with more efficiency than parental G4-DNA formed by a critical nuclease hypersensitive element of the murine KRAS promoter. Hence, thanks to its high uptake and cytoplasm accumulation, compound 1 can be used to repress the expression of the genes that are characterized by 5′UTR containing quadruplex forming G-rich elements as in NRAS11 and KRAS.22 Finally, ATPDs inhibited the proliferation of malignant T24 bladder cells more than nonmalignant murine NIH 3T3 and human 293 embryonic cells do. We found that the mechanism of action of these molecules is based on the arrest of the treated cells in the G0/G1 phase of the cell cycle. As ATDPs do not induce cell death and apoptosis, according to trypan blue and caspase 3/7 assays, they behave as cytostatic agents in T24 bladder cancer cells. The vast majority of anticancer drugs are cytotoxic agents that kill cancer cells and shrink the tumor.36 However, the substantial toxicity of cytotoxic agents and the repopulation of cancer cells escaping lethal chemotherapy are critical factors that often lower the success rate of treatment. Therefore, one strategy to limit tumor recurrence and increase survival is to associate chemotherapy with cytostatic drugs that suppress the growth of resistant cells.36 In this context our ATPD could be interesting cytostatic agents to treat bladder cancer.

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internal standard. Analytical TLC was performed on silica gel F254 plates (Merck) and column chromatography on Silica Gel Merck 60. Melting points were determined on a Buchi SMP-20 apparatus and are uncorrected. High-resolution mass spectra were recorded with electron-spray ionization on a Bruker Daltonics microOTOF-QII instrument. UV spectra were recorded on Hitachi-U2000 spectrophotometer. HPLC was performed using Shimadzu Class-VP V6.12SP1 system (Kromasil-100-5-mkm C-18 column, 4.6 mm × 250 mm), eluent A, H3PO4 (0.01M); B, MeCN. All solutions were dried over Na2SO4 and evaporated at reduced pressure on a Buchi-R200 rotary evaporator at the temperature below 45 °C. All products were vacuum-dried at room temperature. All solvents, chemicals, and reagents were obtained commercially and used without purifications. Anthra[2,3-b]thiophen5,10-dione 2 was prepared as described earlier.9c The purity of all anthra[2,3-b]thiophen-5,10-diones 1−13 was >95% as determined by HPLC analysis. The purity of compounds 11−13 was additionally confirmed by elemental analysis. Synthesis of Anthra[2,3-b]thiophene-5,10-diones 3-13. 4-[(2Aminoethyl)amino]-11-bytoxyanthra[2,3-b]thiophene-5,10-dione (3), 11-[(2-Aminoethyl)amino]-4-butoxyanthra[2,3-b]thiophene5,10-dione (4), and 4,11-Bis[(2-aminoethyl)amino]anthra[2,3-b]thiophene-5,10-dione (5). A mixture of anthrathiophenedione 29c (500 mg, 0.3 mmol), ethylenediamine (1.0 mL), and tetrahydrofuran (1.0 mL) was heated at 30 °C for 2.5 h. During this time the color of the reaction mixture changed from yellow to red-violet. The solution was quenched with water, and aqueous solution of HCl (1.0%) was added to reach pH 8.0. The product was extracted with n-butanol (3 × 30 mL), and the extract was washed twice with brine, dried, and evaporated. The products were separated by chromatographic fractionation with chloroform−methanol−NH4OH (10:1:0 → 10:4:1.5) as eluting solvent and after crystallization from corresponding solvents were isolated starting compound 2 (55 mg, 11%), products of monosubstitutions 3 (210 mg, 43%) and 4 (58 mg, 12%), and bis(ethylendiamine)-derivative 5 (120 mg, 26%). Data for 3 follow: Rf = 0.42 (chloroform−methanol, 10:3); mp 107− 108 °C (cyclohexane−benzene, 3:1). HPLC (LW = 261 nm, gradient B 20 → 70% (30 min)) tR = 20.40 min, purity 97.6%. 1H NMR (400 MHz, DMSO-d6) δ 11.67 (t, 1H, J = 5.0 Hz, NH), 8.05 (m, 2H, 6-H, 9-H), 7.92 (d, 1H, J = 5.6 Hz, 2-H), 7.89 (d, 1H, J = 5.6 Hz, 3-H), 7.71 (m, 2H, 7-H, 8-H), 3.97 (t, 2H, J = 6.4 Hz, OCH2), 3.73 (dd, 2H, 1J = 5.0 Hz, 2J = 5.8 Hz, HNCH2), 2.93 (t, 2H, J = 5.8 Hz, CH2NH2), 1.75 (m, 2H, OCH2CH2CH2), 1.50 (m, 2H, J = 6.0 Hz, CH2CH3), 0.97 (t, 3H, J = 7.3 Hz, CH3). 13C NMR (100 MHz, DMSO-d6) δ 181.59 (2CO), 148.19 (C), 146.26 (C), 145.00 (C), 134.54 (C), 133.36 (C), 131.78 (C), 118.50, (C), 107.64 (C), 133.44 (CH), 132.58 (CH), 129.62 (CH), 127.05 (CH), 125.94 (CH), 125.71 (CH), 72.96 (CH2), 49.07 (CH2), 41.51 (CH2), 32.08 (CH2), 18.81 (CH2), 13.87 (CH3). UV (ethanol) λmax (log ε) 260 (4.5), 310 (3.6), 480 sh (3.5), 521 (3.8), 560 sh (3.7) nm. HRMS (ESI) calculated for C22H23N2O3S [M + H]+ 395.1424, found 395.1417. Data for 4 follow: Rf = 0.51 (chloroform−methanol, 10:3); mp 126− 128 °C (cyclohexane−benzene, 3:1). HPLC (LW = 263 nm, gradient B 20 → 70% (30 min)) tR = 19.87 min, purity 99.4%. 1H NMR (400 MHz, DMSO-d6) δ 11.63 (t, 1H, J = 5.2 Hz, NH), 8.14 (d, 1H, J = 5.5 Hz, 2H), 8.08 (m, 2H, 6-H, 9-H), 7.77 (m, 2H, 7-H, 8-H), 7.95 (d, 1H, J = 5.5 Hz, 3-H), 3.95 (t, 2H, J = 6.5 Hz, OCH2), 3.89 (dd, 2H, 1J = 5.2 Hz, 2J = 6.0 Hz, HNCH2), 2.95 (t, 2H, J = 6.0 Hz, CH2NH2), 1.80 (m, 2H, OCH2CH2CH2), 1.51 (m, 2H, J = 6.0 Hz, CH2CH3), 0.96 (t, 3H, J = 7.3 Hz, CH3). 13C NMR (100 MHz, DMSO-d6) δ 182.25 (2CO), 147.82 (C), 147.14 (C), 142.57 (C), 134.50 (C), 133.64 (C), 130.75 (C), 120.12, (C), 106.26 (C), 133.58 (CH), 133.43 (CH), 132.75 (CH), 125.96 (CH), 125.86 (CH), 122.77 (CH), 73.75 (CH2), 47.75 (CH2), 41.77 (CH2), 31.89 (CH2), 18.82 (CH2), 13.94 (CH3). UV (ethanol) λmax (log ε) 260 (4.5), 320 (3.5), 480 sh (3.7), 511 (3.9), 545 sh (3.7) nm; HRMS (ESI) calculated for C22H23N2O3S [M + H]+ 395.1424, found 395.1413. Data for 5 follow: Rf = 0.01 (chloroform−methanol, 10:3); mp 143− 144 °C (ethanol−1,4-dioxane, 1:1) (mp 143−144 °C9a). 4-[(2-Aminoethyl)amino]-11-[(3-aminopropyl)amino]anthra[2,3b]thiophene-5,10-dione (6). A mixture of anthrathiophenedione 3 (70

EXPERIMENTAL SECTION

General Methods. NMR spectra were obtained with a 400 MHz (1H NMR) and 100 MHz (13C NMR) Varian VXR-400 instrument. Chemical shifts were measured in DMSO-d6 using tetramethylsilane as K

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2H, CH2NH), 3.33 (m, 2H, CH2NH), 1.98 (m, 2H, CH2). HRMS (ESI) calculated for C23H27N8O2S [M + H]+ 479.1972, found 479.1969. 11-[(2-Guanidinoethyl)amino]-4-[(3-guanidinopropyl)amino]anthra[2,3-b]thiophene-5,10-dione Dihydrochloride (11). This compound was prepared from anthrathiophenedione 7 as described for 10. Dark blue solid, yield 69%, mp 208−209 °C (dec). HPLC (LW = 265 nm, gradient B 10 → 70% (30 min)) tR = 11.90 min, purity 98.1%. 1H NMR (400 MHz, DMSO-d6) δ 12.31 (br s, 2H, 2NH), 8.25 (m, 2H, 6H, 9-H), 8.23 (d, 1H, J = 5.6 Hz, 2-H), 8.07 (d, 1H, J = 5.6 Hz, 3-H), 8.04 (t, 1H, J = 5.3 Hz, NH), 8.00 (t, 1H, J = 5.4 Hz, NH), 7.76 (m, 2H, 7-H, 8-H), 7.47 (br s, 4H, 2NH2), 7.20 (br s, 4H, 2NH2), 4.03 (m, 2H, HNCH2), 3.85 (m, 2H, HNCH2), 3.55 (m, 2H, CH2NH), 3.3 (m, 2H, CH 2 NH), 1.95 (m, 2H, CH 2 ). HRMS (ESI) calculated for C23H27N8O2S [M + H]+ 479.1972, found 479.1966. Anal. Calcd for C23H26N8O2S·2HCl·2H2O: C, 47.02; H, 5.49; N, 19.07; S, 5.46. Found %: C, 47.18; H, 5.45; N, 18.92; S, 5.34. 4,11-Bis[(3-guanidinopropyl)amino]anthra[2,3-b]thiophene5,10-dione Dihydrochloride (12). This compound was prepared from anthrathiophenedione 8 as described for 10. Dark blue solid, yield 74%, mp 215−218 °C (dec). HPLC (LW = 265 nm, gradient B 10 → 90% (30 min)) tR = 11.64 min, purity 96.7%. 1H NMR (400 MHz, DMSO-d6) δ 12.48 (br s, 2H, 2NH), 8.22 (m, 3H, 2-H, 6-H, 9-H), 8.04 (m, 3H, 3-H, 2NH), 7.74 (m, 2H, 7-H, 8-H), 7.51 (br s, 4H, 2NH2), 7.18 (br s, 4H, 2NH2), 3.92 (m, 2H, HNCH2), 3.82 (m, 2H, HNCH2), 3.33 (m, 4H, 2CH2NH), 1.95 (m, 4H, 2CH2). UV (ethanol) λmax (log ε) 230 sh (4.3), 265 (4.6), 292 (4.0), 330 sh (3.6), 530 sh (3.8), 566 (4.1), 610 (4.2) nm. HRMS (ESI) calculated for C24H29N8O2S [M + H]+ 493.2129, found 493.2115. Anal. Calcd for C24H28N8O2S·2HCl·2H2O: C, 47.92; H, 5.70; N, 18.63; S, 5.33. Found %: C, 47.59; H, 5.84; N, 18.44; S, 5.24. 4,11-Bis[(4-guanidinobutyl)amino]anthra[2,3-b]thiophene-5,10dione Dihydrochloride (13). This compound was prepared from anthrathiophenedione 9 as described for 10. Dark blue solid, yield 71%, mp 201−203 °C (dec). HPLC (LW = 265 nm, gradient B 20 → 70% (30 min)) tR = 9.73 min, purity 97.3%. UV (ethanol) λmax (log ε) 230 sh (4.3), 265 (4.6), 292 (4.0), 336 sh (3.6), 528 sh (3.8). 1H NMR (400 MHz, DMSO-d6) δ 12.58 (br s, 2H, 2NH), 8.23 (m, 2H, 6-H, 9-H), 8.15 (d, 1H, J = 5.6 Hz, 3-H), 8.02 (d, 1H, J = 5.6 Hz, 3-H), 7.95 (br s, 2H, 2NH), 7.74 (m, 2H, 7-H, 8-H), 7.50 (br s, 4H, 2NH2), 7.15 (br s, 4H, 2NH2), 3.84 (m, 4H, 2HNCH2), 3.20 (m, 4H, 2CH2NH), 1.76 (m, 4H, 2CH2), 1.67 (m, 4H, 2CH2). 13C NMR (100 MHz, DMSO-d6) δ 178.59 (2CO), 169.31 (2C), 146.60 (C), 145.74 (C), 135.12 (2C), 134.16 (C), 133.97 (C), 106.04, (C), 104.88 (C), 131.56 (2CH), 131.03 (CH), 126.32 (CH), 125.58 (CH), 125.56 (CH), 47.25 (CH2), 45.55 (CH2), 41.29 (2CH2), 30.56 (2CH2), 27.56 (CH2), 27.54 (CH2). UV (ethanol) λmax (log ε) 230 sh (4.3), 264 (4.6), 293 (4.0), 340 sh (3.6), 533 sh (3.8), 567 (4.1), 611 (4.2) nm. HRMS (ESI) calculated for C26H33N8O2S [M + H]+ 521.2442, found 521.2432. Anal. Calcd for C26H32N8O2S·2HCl·2H2O: C, 49.60; H, 6.08; N, 17.80; S, 5.09. Found %: C, 49.68; H, 6.14; N, 18.14; S, 5.01. Oligonucleotides, Fluorophore-Labeled Oligonucleotides, and Plasmids. Oligonucleotides and oligonucleotides labeled at the 5′ and 3′ ends with FAM and TAMRA have been purchased from Microsynth (CH), as HPLC purified compounds. The fluorophorelabeled oligonucleotides used are the following: FAMTTAGGGTTAGGGTTAGGGTTAGGG-TAMRA (htelo); FAMTCGGGTTGCGGGCGCAGGGCACGGGCG-TAMRA (hras-1); FAM-CGGGGCGGGGCGGGGGCGGGGGCG-TAMRA (hras-2); FAM-GCGGGAGGGAGGGAAGGAGGGAGGGAGGGAGTAMRA (KRAS), FAM-GGGAGGGGCGGGUCUGGG-TAMRA (NRAS), where TAMRA = tetramethyl rhodamine; FAM = 6carboxyfluorescein. Oligonucleotide aliquots in milli-Q water were kept at −80 °C. Plasmid pHRAS-luc was previously constructed in our laboratory.2c Plasmid pHRAS-mut was obtained from pHRAS-luc by site-directed mutagenesis (GenScript, Piscataway, NJ). FRET Experiments. Fluorescence measurements were carried out with a microplate spectrofluorometer system (Perkin-Elmer 2300 Enspire) using a 96-well black plate, in which each well contained 50 μL of 200 nM dual-labeled oligonucleotide in 50 mM Tris HCl buffer, pH 8, and potassium chloride as specified in the figure captions. The emission

mg, 0.3 mmol), 1,3-diaminopropane (2.0 mL), and 1,4-dioxane (2.0 mL) was heated at 50 °C for 1.5−2 h. During this time the color of the reaction mixture changed from violet to dark blue, and after complete conversion of 3 (as determined by TLC) the solution was cooled and quenched with water. Aqueous solution of HCl (1.0%) was added to reach pH 8.0, the solution was saturated with NaCl, and the product was extracted with warm n-butanol (3 × 30 mL). The extract was washed twice with brine, dried, and evaporated. The residue was purified by column chromatography with chloroform−methanol−NH4OH (10:2:0 → 10:6:2) as eluting solvent. The solid residue obtained after evaporation was crystallized from ethanol−1,4-dioxane mixture (1:1) to afford 6 (60 mg, 85%) as dark blue crystals, mp 151−152 °C; dihydrochloride mp 242−243 °C (dec). HPLC (LW = 266 nm, gradient B 10 → 70% (30 min)) tR = 10.61 min, purity 98.3%. 1H NMR (400 MHz, DMSO-d6) δ 12.57 (t, 1H, J = 4.5 Hz, NH), 12.49 (t, 1H, J = 5.3 Hz, NH), 8.24 (m, 2H, 6-H, 9-H), 8.12 (d, 1H, J = 5.6 Hz, 2-H), 8.00 (d, 1H, J = 5.6 Hz, 3-H), 7.71 (m, 2H, 7-H, 8-H), 3.91 (dd, 2H, 1J = 4.5 Hz, 2 J = 6.0 Hz, HNCH2), 3.89 (dd, 2H, 1J = 5.3 Hz, 2J = 6.2 Hz, HNCH2), 2.88 (t, 2H, J = 6.0 Hz, CH2NH2), 2.76 (t, 2H, J = 6.2 Hz, CH2NH2), 1.82 (m, 2H, CH2). HRMS (ESI) calculated for C21H23N4O2S [M + H]+ 395.1536, found 395.1529. 11-[(2-Aminoethyl)amino]-4-[(3-aminopropyl)amino]anthra[2,3b]thiophene-5,10-dione (7). This compound was prepared from anthrathiophenedione 4 and 1,3-diaminopropane as described for 6 (50 °C, 1.5−2 h). Dark blue powder, yield 82%, mp 152−153 °C; dihydrochloride mp 231−234 °C (dec). HPLC (LW = 266 nm, gradient B 10 → 70% (30 min)) tR = 10.61 min, purity 96.8%. 1H NMR (400 MHz, DMSO-d6) δ 12.57 (m, 2H, 2NH), 8.26 (m, 2H, 6-H, 9-H), 8.14 (d, 1H, J = 5.6 Hz, 2-H), 8.09 (d, 1H, J = 5.6 Hz, 3-H), 7.74 (m, 2H, 7-H, 8-H), 3.86 (m, 4H, 2HNCH2), 2.93 (t, 2H, J = 6.0 Hz, CH2NH2), 2.75 (t, 2H, J = 6.5 Hz, CH2NH2), 1.81 (m, 2H, CH2). HRMS (ESI) calculated for C21H23N4O2S [M + H]+ 395.1536, found 395.1527. 4,11-Bis[(3-aminopropyl)amino]anthra[2,3-b]thiophene-5,10dione (8). This compound was prepared from anthrathiophenedione 29c and 1,3-diaminopropane as described for 6 (50 °C, 3−4 h). Dark blue powder, yield 77%, mp 155−156 °C; dihydrochloride mp 248−249 °C (dec). HPLC (LW = 266 nm, gradient B 10 → 70% (30 min)) tR = 10.96 min, purity 97.3%. 1H NMR (400 MHz, DMSO-d6) δ 12.58 (m, 2H, 2NH), 8.21 (m, 2H, 6-H, 9-H), 8.05 (d, 1H, J = 5.6 Hz, 2-H), 7.98 (d, 1H, J = 5.6 Hz, 3-H), 7.69 (m, 2H, 7-H, 8-H), 3.85 (dd, 2H, 1J = 5.0 Hz, 2 J = 6.4 Hz, HNCH2), 3.75 (dd, 2H, 1J = 4.7 Hz, 2J = 6.6 Hz, HNCH2), 2.72 (m, 4H, 2CH2NH2), 1.79 (m, 4H, 2CH2). HRMS (ESI) calculated for C22H25N4O2S [M + H]+ 409.1693, found 409.1690. 4,11-Bis[(4-aminobutyl)amino]anthra[2,3-b]thiophene-5,10dione (9). This compound was prepared from anthrathiophenedione 29c and 1,4-diaminobutane as described for 6 (60 °C, 4−5 h). Dark blue powder, yield 73%, mp 114−116 °C; dihydrochloride mp 245−248 °C (dec). HPLC (LW = 265 nm, gradient B 10 → 70% (30 min)) tR = 12.19 min, purity 98.2%. 1H NMR (400 MHz, DMSO-d6) δ 12.55 (m, 2H, 2NH), 8.19 (m, 2H, 6-H, 9-H), 8.01 (d, 1H, J = 5.6 Hz, 2-H), 7.91 (d, 1H, J = 5.6 Hz, 3-H), 7.67 (m, 2H, 7-H, 8-H), 3.76 (dd, 2H, 1J = 5.5 Hz, 2 J = 6.5 Hz, HNCH2), 3.66 (dd, 2H, 1J = 5.1 Hz, 2J = 6.2 Hz, HNCH2), 2.59 (m, 4H, 2CH2NH2), 2.59 (m, 4H, 2CH2NH2), 1.70 (m, 4H, 2CH2), 1.51 (m, 4H, 2CH2). HRMS (ESI) calculated for C24H29N4O2S [M + H]+ 437.2006, found 437.1992. 4-[(2-Guanidinoethyl)amino]-11-[(3-guanidinopropyl)amino]anthra[2,3-b]thiophene-5,10-dione Dihydrochloride (10). To a stirring solution of free base of anthrathiophenedione 6 (60 mg, 0.15 mmol) in DMSO (10.0 mL) were added ethyldiisopropylamine (1.0 mL, 6.0 mmol) and 0.5 g (3.4 mmol) hydrochloride of pyrazole-1carboxamidine. The mixture was stirred for 5 h at 60 °C and then cooled. The product was precipitated by treatment with acetone and collected by filtration. The blue solid was reprecipitated twice from hot water with acetone, washed with acetone, and, after drying, yielded dihydrochloride 10 (61 mg, 72%); mp 213-214 °C (dec). HPLC (LW = 265 nm, gradient B 10 → 70% (30 min)) tR = 11.94 min, purity 97.8%. 1H NMR (400 MHz, DMSO-d6) δ 12.41 (br s, 1H, NH), 12.14 (br s, 1H, NH), 8.25 (m, 3H, 2-H, 6-H, 9-H), 8.06 (t, 1H, J = 5.4 Hz, NH), 7.99 (d, 1H, J = 5.6 Hz, 3-H), 7.93 (t, 1H, J = 5.4 Hz, NH), 7.75 (m, 2H, 7-H, 8-H), 7.49 (br s, 4H, 2NH2), 7.17 (br s, 4H, 2NH2), 3.94 (m, 4H, 2HNCH2), 3.53 (m, L

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spectra were obtained setting the excitation wavelength at 475 nm and recording the emission from 500 to 650 nm. Fluorescence melting experiments were performed on a real-time PCR apparatus (CFX96, BioRad, Hercules, CA). FRET-melting experiments were obtained by the following: equilibration step of 5 min at low temperature (20 °C); stepwise increase of the temperature of 1 °C/min for 76 cycles to reach 95 °C. All the samples in the wells were melted in 76 min. Confocal Microscopy. T24 bladder cancer and nonmalignant embryonic NIH 3T3 cells were plated (3 × 105) on coverslips (diameter 24 mm) and after 24 h treated with 2.5 μM anthrathiophenedione for 2 or 4 h. The cells were washed twice with PBS, fixed with 3% paraformaldehyde (PFA) in PBS for 20 min, after washing with 0.1 M glycine, containing 0.02% sodium azide in PBS to remove PFA and Triton X-100 (0.1% in PBS). The cells were incubated for 1 h with DRP1 antibody (Transduction Laboratories) diluted 1:75 in PBS and then 30 min with secondary mouse IgG FITC (Santa Cruz) diluted 1:250 in PBS. For some experiments the nuclei were stained for 30 min with propidium iodide (3 ng/μL, 0.4 μg/mL RNase A in PBS). The cells were analyzed using a Leica TCS SP1 confocal imaging system. FACS Analysis. T24, 293, and NIH 3T3 cells were plated in a 24well plate at density of 5 × 104 cells/well. After one day, the cells were treated with anthrathiophenedione: time and concentration as indicated in figure captions. For uptake studies the cells were treated for 30 min with 5 μM Cytochalasin D or 80 μM Dynasore before the addition of anthrathiophenedione. The cells were trypsinized and pelleted. The pellets were resuspended in 200 μL PBS and immediately analyzed by FACScan flow cytometer (Becton-Dickinson, San Jose) equipped with a single 488 nm argon laser. A minimum of 10 000 cells for each sample were acquired in list mode and analyzed using Cell Quest software. The cell population was analyzed by FSC light and SSC light. The signal was detected by FL3 (680 nm) channel in log scale. For the cell cycle analyses the cells have been harvested by trypsinization, fixed for 1 h at 4 °C in 70% ethanol, 30% PBS. After PBS washing, the cells were stained with 0.05 mg/mL propidium iodide in the presence of 0.1 mg/mL RNase A in PBS (30 min at room temperature). The samples were analyzed by FACS (FL 2 channel). The cell cycle data have been obtained with FLUOJO software (Tri Star, Inc., Ashland, OR). Cell Culture and Proliferation Assay. T24 urinary bladder cancer, 293 human kidney embryonic, and murine embryonic NIH 3T3 cells were maintained in exponential growth in Dulbecco’s Modified Eagle’s Medium (DMEM) containing 100 U/mL penicillin, 100 mg/mL streptomycin, 20 mM L-glutamine, and 10% fetal bovine serum (Euroclone, Milan, Italy). For proliferative assays, the cells were seeded (3000 cells/well) the day before anthrathiophenedione treatment in a 96-well plate. Cell viability was measured by resazurin assays following standard procedures at the time indicated. Dual Luciferase Assays. Transfection was performed by mixing vector (250 ng/well) pHRAS-luc or pHRAS-mut or pIL-luc with control plasmid pRL-CMV in which Renilla luciferase is driven by the CMV promoter (10 ng/well) using jet-PEI (Polyplus) as a transfecting reagent. Luciferase assays were performed 48 and 72 h after transfection with Dual-Glo Luciferase Assay System (Promega) following the supplier instructions. Firefly luciferase activity in cell lysates was measured and normalized for Renilla luciferase activity. Samples were read on a Turner Luminometer, and the relative luminescence was expressed as (T/C × 100) where T = firef ly luciferase/renilla luciferase in treated cells and C = firef ly luciferase/renilla luciferase in untreated cells. Western Blot. Total protein lysates (30 μg) were electrophoresed on 12% SDS-PAGE and transferred to a nitrocellulose membrane at 70 V for 2 h. The filter was blocked for 1 h with 5% BSA solution in TBST (10 mM Tris pH 7.9, 150 mM NaCl and 0.05% Tween) (Sigma-Aldrich, Milan, Italy) at room temperature. The primary antibodies (mouse monoclonal antiactin, Oncogene, diluted 1:10 000; mouse monoclonal c-HRAS Abcam, diluted 1:500) were overnight incubated with the samples at 4 °C. The expressions of β-actin was used as an internal control. The filters were washed with a 0.05% Tween in PBS and subsequently incubated for 1 h with the secondary antibody and antirabbit IgG for HRAS, diluited 1:10 000 (Calbiochem). For β-actin we used an antimouse IgM, diluted 1:10 000 (Calbiochem). Each

secondary antibody was coupled to horseradish peroxidase. For the detection of the proteins we used Super SignalWest PICO, and Super SignalWest FEMTO (Thermo Fisher Scientific Pierce). Exposure length depends on the antibodies used and was usually between 30 s to 5 min. The protein levels were quantified by Quantity ONE 4.6.5 software with Chemidoc (Bio Rad).

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ASSOCIATED CONTENT

S Supporting Information *

ATPD UV−vis and fluorescence spectra, UV−vis titrations of ATPDs with duplex DNA, confocal microscopy of ATPDs in T24 and NIH 3T3 cells, and putative structures of the HRAS promoter quadruplexes. This material is available free of charge via the Internet at http://pubs.acs.org.

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AUTHOR INFORMATION

Corresponding Author

*Phone: +39.0432.494395. E-mail. [email protected]. Present Address ⊥

System Biology Ireland-Conway Institute, University College Dublin, Belfast, Dublin 4, Ireland. Author Contributions

S.C. and A.S. contributed equally. Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS The authors are grateful to Yu. Luzikov, A. Korolev, M. Resnikova, N. Malutina for NMR, HRMS, and HPLS analyses and T. Pussini for FRET and FACS experiments. This work has been carried out with the financial support of AIRC-2010 (ref 10546, The Italian Association for Cancer Research) and PRIN 2009

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ABBREVIATIONS USED ATPDs, anthrathiophenediones; ChIP, chromatin immuno precipitation; TSS, transcription start site; EMSA, electrophoresis mobility shift assay; DMS, dimethyl sulfate; HRAS, Harvey ras; NRAS, neuroblastoma ras; FACS, fluorescenceactivated cell sorting; FRET, fluorescence-resonance energy transfer; PCR, polymerase chain reaction; DMEM, Dulbecco’s modified eagle medium; UTR, untranslated region; GPc, guanidinium phthalocyanines; QFS, quadruplex-forming sequences; DRP1, dynamin-related protein; FAM, 6-carboxyfluorescein; TAMRA, tetramethyl rhodamine

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REFERENCES

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