Highly Brominated Quinolines: Synthesis, Characterization, and Investigation of Anticancer Activities Supported by Molecular Dynamics

ABSTRACT

In this study, we synthesized and characterized novel brominated methoxyquinolines (7 and 11) and nitrated bromoquinoline (17) derivatives with potential antiproliferative activity against cancer cell lines. Starting from 1,2,3,4‐tetrahydroquinoline (THQ, 1), a series of brominated quinoline compounds was obtained via regioselective bromination and subsequent reactions. The structure of the key compound, 3,5,6,7‐tetrabromo‐8‐methoxyquinoline (7), was confirmed using 1D and 2D NMR techniques. Additionally, unexpected bromination of 3,6,8‐trimethoxyquinoline (5) yielded 5,7‐dibromo‐3,6‐dimethoxy‐8‐hydroxyquinoline (11), allowing functionalization of both rings in the quinoline. The direct nitration of 6,8‐dibromoquinoline (6) yielded the corresponding 5‐nitro derivative (17), a precursor to amino derivatives that activate the bromine group on the ring. Antiproliferative activities of these derivatives (7, 11, 17) were assessed against C6, HeLa, and HT29 cancer cell lines using the BCPE assay. Compounds 7, 11, and 17 exhibited significant inhibitory effects, with compound 11 showing the highest activity (IC₅₀ values of 5.45–9.6 μg/mL). Furthermore, the cytotoxicity of these compounds was evaluated using the LDH assay, indicating lower cytotoxic effects compared to the control drug 5‐FU. The ability of compounds 11 and 17 to induce apoptosis was confirmed through DNA laddering, while compound 7 showed no such effect. Compounds 7 and 11 inhibited human topoisomerase I, a critical enzyme for DNA replication and repair, with significant binding energies determined by MM‐PBSA studies. The wound healing assay demonstrated that compound 17 effectively inhibited the migration of HT29 cells. These findings highlight the potential of these novel quinoline derivatives as effective anticancer agents, warranting further investigation into their mechanisms of action and therapeutic applications.

Further bromination of methoxy quinoline furnished novel polyfunctionalized quinoline derivatives. Additionally, nitration of dibromoquinoline gave 5‐nitrated quinoline. Their significant anticancer activities were determined and also supported by in silico studies.

1. Introduction

Quinoline and its derivatives have gained significant attention as important heterocyclic pharmacophores with diverse biological effects (Köprülü et al. 2021; Özcan et al. 2020; Li and Zhu 2021; Mittal et al. 2023). On the other hand, 8‐hydroxyquinoline (8‐HQ) derivatives are known for their strong metal ion chelating properties and have a wide range of pharmacological applications (Navarro et al. 2006; Li and Xu 2008; Song et al. 2015; Ökten et al. 2017a; Zhou et al. 2023; Guan et al. 2023; Bissani and Pilger 2023). The quinoline and 8‐HQ scaffolds possess versatile binding properties, and through the modification of their functional groups, they can function as potent and specific ligands for a variety of biotargets (Saadeh et al. 2020). Consequently, these scaffolds are regarded as privileged structures in medicinal chemistry. They are extensively utilized as initial compounds for creating substances with a wide range of pharmacological activities (Ökten et al. 2017b).

Recent studies have extensively demonstrated using bromoquinolines as precursors for preparing multifunctional quinoline compounds (Ökten et al. 2015; Şahin et al. 2008). Additionally, brominated tetrahydroquinolines and quinolines have been shown to play an essential role in synthesizing corresponding derivatives with cyano, methoxy, phenyl, and amino groups (Ökten et al. 2013). The straightforward bromination procedure of 1,2,3,4‐tetrahydroquinoline (1) allows for the easy preparation of monobromo, dibromo, and tribromoquinoline derivatives, which can be subsequently transformed into their respective derivatives for antiproliferative activity against various cancer cell lines (Ökten et al. 2013; Çakmak and Ökten 2017; Ökten and Çakmak 2015; Zemtsova et al. 2015; Thomas et al. 1994). Quinolines derived from bromoquinolines with different functional groups have shown promising anticancer activities in several studies (Köprülü et al. 2019, 2021; Özcan et al. 2020; Ökten et al. 2017a). Recent research has highlighted the potent inhibitory effects of 8‐substituted quinolines containing hydroxyl and methoxy groups at the C‐8 position against HT29 (human adenocarcinoma), HeLa (human cervical cancer), and C6 (rat glioblastoma) cell lines (Ökten et al. 2017a). Additionally, derivatives of bromo‐ and cyano‐substituted 8‐hydroxyquinoline quinoline have demonstrated the ability to induce apoptosis in C6, HT29, and HeLa cell lines. Specifically, 5,7‐dicyano‐8‐hydroxyquinoline and 5,7‐dibromo‐8‐hydroxyquinoline have been found to inhibit the recombinant human DNA topoisomerase I enzyme (Ökten et al. 2017a).

We have developed new methods for modifying quinolines using nitrated bromoquinolines. The nitro group on the quinoline activates the bromo group for nucleophilic substitution, allowing us to efficiently produce 6‐piperazinyl and 6‐morpholinyl quinolines from 6‐bromo‐5‐nitroquinoline through S_NAr nucleophilic substitution (Scheme 1). Notably, 6‐bromo‐5‐nitroquinoline displayed significant antiproliferative and apoptotic effects with low cytotoxicity against various cancer cell lines (Ökten et al. 2020).

SCHEME 1.

Synthesis of starting compounds (3, 5, 6).

Our ongoing research is focused on the innovative synthesis of polyfunctional quinolines, using methoxy 1,2,3,4‐tetrahydroquinoline as our key starting material. We have meticulously designed and synthesized over 50 novel quinoline derivatives, followed by comprehensive in silico studies to evaluate their bioactivities, with particular emphasis on their potential to inhibit the Topo 1 enzyme and bind to cancer cell DNA. Among the compounds examined, 3,5,6,7‐tetrabromo‐8‐methoxyquinoline (7) and 5,7‐dibromo‐3,6‐dimethoxy‐8‐hydroxyquinoline (11) emerged as the most promising candidates. To achieve these target compounds, we developed and refined a new synthesis methodology, which enabled us to obtain them in a purified form with high efficiency. This advanced method allows for the streamlined synthesis of a library of brominated derivatives at positions C‐3, C‐5, and C‐7, which we intend to further investigate for their biological activities by experimental and structure‐based drug design methods. The initial studies of these synthesized quinoline derivatives have revealed significant anticancer potential. By in silico docking, molecular dynamics, and MM‐PBSA (Molecular Mechanics‐Poisson Boltzmann Surface Area) binding affinity studies, we aim to uncover critical insights into the mechanisms of action of potential brominated quinoline derivatives with anticancer activities against topoisomerase‐I, paving the way for the design of even more potent and selective anticancer agents as topoisomerase‐I inhibitors.

2. Materials and Method

2.1. Synthesis of 3,5,6,7‐Tetrabromo‐8‐methoxyquinoline (7)

A solution of 6‐bromo‐8‐methoxy‐1,2,3,4‐tetrahydroquinoline (3, 100 mg, 0.41 mmol, 1 eq) in CHCl₃ (15 mL) was combined with a solution of bromine (347 mg, 2.17 mmol, 5.3 eq) in CHCl₃ (5 mL) over 5 min in the absence of light at room temperature. The mixture was left to react until all the bromine was used up (5 days). Afterward, the resulting mixture was washed with a solution of 5% aqueous NaHCO₃ (3 × 20 mL) and dried over Na₂SO₄. The solvent was removed by evaporation, and the crude material (153 mg) was purified by passing it through a silica column and eluting it with AcOEt/hexane (1:5, 100 mL). The crude product was then recrystallized in AcOEt/hexane (1:5) to obtain 3,5,6,7‐tetrabromo‐8‐methoxyquinoline (7). White powder solid (143 mg, 74% yield), Mp 134°C–136°C. ¹H NMR (400 MHz, CDC1₃): δ 8.76 (d, J ₂₄ = 1.6 Hz, 1H, H₂), 8.68 (d, J ₄₂ = 1.6 Hz, 1H, H₄), 4.02 (s, OCH ₃ ). ¹³C NMR (100 MHz, CDC1₃): δ 154.5 (q), 151.1 (q), 149.8, 137.3, 134.1 (q), 127.9 (q), 120.2 (q), 103.2(q), 102.8 (q), 61.2 (OCH ₃ ). HETCOR: 8.76–149.8, 8.68–137.3, 4.02–61.2; FT‐IR (cm‐1): 3063, 2984, 2943, 2863, 1605, 1572, 1467, 1450, 1420, 1341, 1314, 1286, 1231, 1198, 1130, 1093, 1066, 976, 941, 889, 837, 781, 724, 678, 655, 631, 596, 583, 471. Anal. Calcd for C₁₀H₅Br₄NO (470.71 g/mol): C, 25.30%, H, 1.06%, N, 2.95%. Found: C, 25.34%; H, 1.07%; N, 3.00%.

2.2. Synthesis of 5,7‐Dibromo‐3,6‐dimethoxy‐8‐hydroxyquinoline (11)

The reaction was carried out by adding a solution of bromine (361 mg, 2.02 mmol, 2 equivalents) in CH₂Cl₂ (10 mL) to a solution of 3,6,8‐trimethoxyquinoline (5, 200 mg, 0.11 mmol, 1 equivalent) in CH₂Cl₂ (30 mL) over 30 min in the dark at room temperature. The reaction was allowed to proceed until all bromine was consumed (2 days). The resulting mixture was then washed with a 5% aqueous NaHCO₃ solution (3 × 25 mL) and dried over Na₂SO₄. After evaporating the solvent, the crude material (132 mg) was passed through a silica column, eluting with AcOEt/hexane (1:1, 100 mL). The crude product was then recrystallized in AcOEt/hexane (1:1) to yield 5,7‐dibromo‐3,6‐dimethoxy‐8‐hydroxyquinoline (4). White powder solid (243.0 mg, 76% yield), Mp 156°C–158°C. ¹H NMR (400 MHz, CDCl₃): δ 8.47 (d, J ₂₄  = 2.4 Hz, 1H, H2), 7.70 (d, J ₄₂  = 2.4 Hz, 1H, H4), 4.85 (bs, 1H, ‐OH), 4.03 (s, 3H, ‐OCH ₃ ), 4.01 (s, 3H, ‐OCH ₃ ); ¹³C NMR (100 MHz, CDC1₃): δ 155.7, 154.0, 150.8, 142.1, 130.6, 127.9, 112.6, 103.0, 99.7, 61.1, 55.9; FT‐IR (cm⁻¹): 3353, 2939, 1610, 1483, 1450, 1387, 1362, 1294, 1221, 1074, 1030, 964, 866, 845, 783, 721, 665, 650. Anal. Calcd for C₁₁H₉Br₂NO₃ (363.00 g/mol): C, 36.40%; H, 2.50%; N, 3.86%. Found: C, 36.47%; H, 2.48%; N, 3.89%.

2.3. Synthesis of 6,8‐Dibromo‐5‐nitroquinoline (17)

In this study, 1.00 g (3.484 mmol) of 6,8‐dibromoquinoline (6) was dissolved in 10 mL of sulfuric acid and chilled to −5°C in a salt‐ice bath. A solution of 50% nitric acid in 20 mL of sulfuric acid was then slowly added to the 6,8‐dibromoquinoline (6) solution while stirring at −5°C. After 1 h in an ice bath, the resulting yellow mixture was allowed to reach room temperature. The red solution was then poured onto 50 g of crushed ice in a beaker. The ice was left to melt, and the mixture was extracted three times with 40 mL of CH₂Cl₂. The organic phase was neutralized using a 10% aqueous NaHCO₃ solution and then dried over Na₂SO₄. Finally, the solvent was removed under vacuum. As a result, yellow needle‐shaped crystals were obtained as the sole product 17, with a yield of 96% (1.106 g). Mp 160°C–162°C. ¹H NMR (400 MHz, CDCl₃): δ 9.17 (dd, J ₂₃  = 4 Hz, J ₂₄  = 1.2 Hz, 1H, H₂), 8.33 (s, 1H, H₇), 8.08 (dd, J ₄₂  = 1.2 Hz, J ₄₃ = 4 Hz, 1H, H₄), 7.69 (dd, J ₃₂  = 4 Hz, J ₃₄ = 4 Hz, 1H, H₃). ¹³C NMR (100 MHz, CDCl₃): δ 152.6, 147.3 (q), 143.9 (q), 135.7, 130.6, 128.8 (q), 124.6 (q), 121.9 (q), 112 (q). FT‐IR (cm⁻¹): 3083, 1583, 1536, 1519, 1471, 1371, 1340, 1290, 1199, 927, 873, 856, 806, 779. Anal. Calcd for C₉H₄Br₂N₂O₂ (331.95 g/mol): C, 32.56%; H, 1.21%; N, 8.44%. Found C, 32.52%; H, 1.23%; N, 8.46%.

2.4. Cell Culture

The HeLa (ATCCCCL‐2) human cervical adenocarcinoma, HT29 (ATCCHTB‐38) human colon adenocarcinoma, and C6 (ATCCCCL‐107) rat glioblastoma adherent cells were grown in Dulbecco's Modified Eagle's Medium (DMEM, Sigma‐Aldrich) with 10% heat‐inactivated fetal bovine serum (FBS, Sigma‐Aldrich, Germany) and 1% Penicillin–Streptomycin (PenStrep) Solution (Sigma‐Aldrich, Germany) at 37°C in a humidified atmosphere with 5% CO₂. Upon reaching 80% confluence, the culture medium was changed, and the cells were subcultured using 4 mL of trypsin–EDTA (Sigma‐Aldrich). The tested compounds and positive control 5‐Fluorouracil (5‐FU) were dissolved in DMSO to prepare a stock solution and then diluted with DMEM. DMSO served as a negative control for all experiments, and its final concentration in the cell culture medium did not exceed 0.5%–1%. All tests were performed in triplicate.

2.5. BrdU Cell Proliferation ELISA (BCPE)

In order to evaluate the impact of quinoline compounds on cell proliferation, we followed Roche's protocol for the BrdU Cell Proliferation ELISA kit. We seeded HeLa, HT29, and C6 cancer cells onto a 96‐well culture plate from COSTAR (Corning, USA) at a density of 3 × 10³ cells in 100 μL. These cells were then exposed to increasing concentrations of quinoline compounds and 5‐FU (ranging from 5 to 75 μg/mL), which were diluted in supplemented DMEM to attain a final well volume of 200 μL. Following overnight incubation at 37°C with 5% CO₂, we added 20 μL of BrdU labeling reagent to each well and allowed the cells to incubate for 4 h. Subsequently, the culture medium was aspirated, and 200 μL of FixDenat solution was added to fix the cells for 30 min at room temperature. After that, we added a 1:100 diluted anti‐BrdU‐POD to each well and incubated the cells for 90 min at room temperature, followed by three washes with a washing buffer. Substrate solution was then added, and the cells were incubated for 30 min in the dark at room temperature. We utilized an ELISA plate reader from BioTek (Epoch) to measure the optical density at 450–650 nm. Finally, using XLfit5 software (IDBS), we calculated the IC₅₀ of both the test and control compounds, expressed in μg/mL at 95% confidence intervals.

2.6. Lactate Dehydrogenase Assay

We utilized the LDH Cytotoxicity Detection Kit from Roche (USA) to assess the cytotoxicity of quinoline compounds. LDH is an enzyme present in the cytoplasm that plays a crucial role in converting pyruvate to lactate. Its activity can be used to measure the release of cytoplasmic enzymes from damaged cells. Cells from HeLa, HT29, and C6 cell lines were seeded in triplicate in 96‐well culture plates and treated with quinoline compounds at concentrations equivalent to their IC₅₀. Following a 24‐h incubation, Triton X was added to the high‐control wells to degrade the cell membranes, and supernatant from all wells was transferred to the assay plate. A reaction mixture was then added to each well, and after a period of incubation, the absorbances at 492–630 nm were detected using a plate reader. To calculate the percentage difference in cytotoxicity between the treated and control wells, the following formula was used: Cytotoxicity % = (Experimental Value—Low Control) / (High Control‐Low Control) × 100.

2.7. DNA Laddering Assay

The modified DNA laddering assay, based on Gong's method (Gong et al. 1994), involved culturing HeLa, HT29, and C6 cells in 25 cm² culture flasks at a density of 7.5 × 10⁵ cells/well for 24 h. They were then treated with IC₅₀ concentrations of various quinoline compounds, with Camptothecin (CPT) from Sigma‐Aldrich, Germany, used as a positive control, known for inducing apoptosis. Subsequently, cells were detached by scraping, centrifuged at 6000 rpm for 5 min, and fixed with 70% ethanol at −20°C for 36 h. The DNA was then extracted using 0.2 M phosphate–citrate buffer at pH 7.8, followed by the addition of RNase A and proteinase K. The resulting DNA fragmentation was visualized by ethidium bromide staining after loading total DNA onto a 2% agarose gel.

2.8. Topoisomerase I Inhibition

The impact of quinoline compounds on the activity of DNA topoisomerase I (Topo I) enzyme was studied using a TopoGen Topo I assay kit (TG1015‐2; TopoGen, Buena Vista, CO). The assay is based on the relaxation of supercoiled DNA, which has different electrophoretic mobility compared to completely relaxed DNA. A reaction mixture (20 μL) containing plasmid DNA (supercoiled pHOT1 DNA), 10XTGS buffer, and Topo I was prepared and dispensed into 0.2 mL PCR tubes. Quinoline compounds and the positive control (CPT) were added to their respective tubes, and all samples were then incubated at 37°C for 30 min. The reaction was stopped by adding 4 μL of stop solution (5×), and the samples were loaded onto a 2% agarose gel and visualized using ultraviolet–visible transillumination.

2.9. Wound‐Healing Assay

For the wound‐healing assay, silicone culture inserts from ibidi GmbH, Germany, were used to evaluate cell migration in vitro. The inserts were placed in a 3 cm dish plate, and each compartment was filled with a cell suspension. After the cells covered the surface area, the inserts were removed, and the cells were treated with quinoline compounds at IC₅₀ concentrations. The gap between the two wells in the culture inserts was observed at various time points and captured using an inverted microscope (Leica DMIL, Germany) to evaluate the closure of the gap.

2.10. Computational Studies

2.10.1. Initial Structures and System Setup

X‐ray structure coordinates for human topoisomerase‐I in complex with a 22 bp DNA duplex, possessing a single‐strand nick between nucleotides T10 and C11, were obtained from Protein Data Bank (PDB ID: 1t8i) (Staker et al. 2005). In the X‐ray structure, DNA is in complex with camptothecin (an anticancer agent) to prevent resealing of the single‐strand nick. Amino acid sequence alignment of human topoisomerase‐I present in its X‐ray structure (PDB ID: 1t8i) and at UNIPROT database (Uniprot gene code: TOP1_HUMAN, Uniprot ID: P11387, 765 aa in length) presents 100% sequence identity.

Ligands were constructed, minimized, and saved in SYBYL mol2 format by Avogadro2 v0.8.0 (Hanwell et al. 2012). MGLTools v. 1.5.7 (Sanner 1999) handled the addition of missing protons, Gasteiger partial atomic charges, merging nonpolar C‐H protons onto the covalently attached C atoms, assignment of rotatable bonds in the ligand, and determination of the grid box sizes in the DNA binding site. Coordinates for the ligand and receptor species were then saved in PDBQT format.

Molecular dynamics (MD) computations involved the determination of AM1‐BCC partial atomic charges (at semiempirical quantum chemistry level) for the docked coordinates of the ligand (with zero net charge) by the antechamber module of AMBER v22 (Case et al. 2023). The LeaP module of AMBER v22 parameterized human topoisomerase 1, DNA, and ligands (Case et al. 2023) using the AMBER‐ff19SB force field (Tian et al. 2020), AMBER‐DNA.bsc1 force field (Ivani et al. 2016), and the general AMBER force field‐2 (GAFF2) (He et al. 2020), respectively. Receptor+ligand complexes were neutralized by the addition of 24 Na⁺ cations in LeaP, which were then submerged into octahedral OPC waterboxes (Izadi et al. 2014) with dimensions of 109.743 Å × 109.743 Å × 109.743 Å. Additionally, 16 Cl⁻ and 16 Na⁺ ions were added to the aqueous solutions of the complexes under physiological salt concentration (150 mM).

2.11. Docking

A 64‐bit Linux version of autodock_Vina v.1.1.2 was used to obtain docked coordinates of the ligands in the camptothecin binding site of DNA in complex with human topoisomerase 1. A configuration file (including the parameters: exhaustiveness = 8, num_modes = 20, center_x = 20.77, center_y = − 4.25, center_z = 28.01, size_x = 18.50, size_y = 24.71, size_z = 16.52, energy_range = 3) along with PDBQT files for the receptor and ligand species was used for docking computations in triplicate. The docking results were analyzed by MGLTools v1.5.7 (Salomon‐Ferrer et al. 2013) and saved in SYBYL mol2 format for the highest affinity docked coordinates of the ligand.

2.12. Molecular Dynamics (MD) Simulations

A restrained minimization procedure using the steepest descent method through 1 fsec time steps over 1 psec (1000 iterations) was applied to the complex system by the sander. MPI module of AMBER v22 (2022) suite of programs (Case et al. 2023), in which amino acid (aa) residues (aa 201–765 in PDB file 1t8i) and DNA nucleotides (nt 1–22 and nt 101–122 in PDB file 1t8i) of the receptor were restrained by a force constant of 100 kcal/mol.Ų, using four intel7 CPU cores running in parallel. Heating, pressure equilibration, and production molecular dynamics (MD) simulations were carried out by the pmemd.cuda module (Sindhikara et al. 2009) of AMBER v22 (2022) suite of programs (Case et al. 2023), using a Nvidia Tesla P100 Graphics Card running under 64‐bit Centos v7.3 OS at the Turkish Science e‐Infrastructure (TRUBA, Turkiye).

The temperature of the relaxed complex systems was gradually increased from 100 K and equilibrated at 300 K using a Langevin Dynamics thermostat (Åqvist et al. 2004) over 1 nsec of MD (with 1 fsec time steps over 1,000,000 iterations) and a collision frequency of 1/ps while applying a force constant of 100 kcal/mol.Ų to restrain aa residues of the protein and nt residues of the DNA. A Langevin Dynamics thermostat with a collision frequency of 1/ps was applied from here on in all steps of pressure equilibration and production MD computations to keep the temperature of the complex systems at 300 K. Initially, box densities of the complex systems were relaxed in two steps (each running over 1,000,000 steps at 1 fsec time intervals) using a Monte Carlo barostat fixed at 1 bar (Darden et al. 1993), in which aa residues in the protein and nt residues of the DNA were restrained by force constants of 100 kcal/mol.Ų in the first step and 10 kcal/mol.Ų in the second step of the first pressure equilibration phase.

In the second phase of the pressure equilibration procedure, restraints on the receptor as well as on the DNA were lowered and applied only on the protein (CA, N, and C) and DNA (P, O5′, C5′, C4′, C3′, and O3′) backbone atoms. Firstly, a short minimization procedure with a mild restraint (10 kcal/mol.Ų) on the backbone atoms of the receptor of the complex systems was applied over 2000 steps. It was followed by another relaxation process at 300 K over 1 ns of simulation time (at 1 fs time intervals over 1,000,000 iterations) at a constant pressure (1 bar) maintained by Monte Carlo barostat (Darden et al. 1993) with backbone restraints set to 10 kcal/mol.Ų. The relaxation procedure was extended for two successive 2 × 1 nsec of MD simulations (each with 1 fs time intervals over 1,000,000 iterations) using the aforementioned thermostat (at 300 K) and barostat (at 1 bar) parameters, but this time, lowering the restraint force constants down to 1 kcal/mol.Ų and 0.1 kcal/mol.Ų, respectively, on the aforementioned protein and DNA backbone atoms. Finally, the restraints on the receptor backbone atoms were removed during ~1600 nsec of MD equilibration simulation (at 4 fsec time intervals over ~400 million iterations for the receptor+compound‐7 and receptor+compound‐11 MD systems and at 2 fsec time intervals over ~800 million iterations for the receptor+camptothecin MD system), using the previously mentioned thermostat (at 300 K) and barostat (at 1 bar) parameters. Electrostatic interactions up to a non‐bonded cutoff distance of 9 Å were handled by the Particle Mesh Ewald method during all MD simulations (Ryckaert et al. 1977). Heteronucleus‐H‐bond lengths were stabilized by the SHAKE algorithm (Gohlke and Case 2004) during heating and production MD simulations.

Three independent 3 × 250 nsec MD production simulations (at 4 fs time intervals over 62.5 million iterations for the receptor+compound‐7 and receptor+compound‐11 MD systems and at 2 fsec time intervals over ~125 million iterations for the receptor+camptothecin MD system) were carried out to sample more variable coordinates in the cellular environment, starting from the last snapshot coordinates from the previous 1600 nsec MD equilibration simulation. Frame (snapshot) coordinates were collected and appended into binary trajectory files every 50,000 steps (200 psec) for the receptor+compound‐7 and receptor+compound‐11 MD systems and 100,000 steps (200 psec) for the receptor+camptothecin MD system throughout the MD simulations.

2.13. Trajectory Analysis

The cpptraj module of AMBER v22 carried out RMSD, inter‐residual H‐bond analysis, imaging, and clustering throughout trajectories (Case et al. 2023).

2.14. Thermodynamic Computations

Molecular Mechanics Poisson Boltzmann Surface Area (MM‐PBSA) binding energies were previously determined (Andac et al. 2021) by the mmpbsa module of AMBER v22 (Duan et al. 2003). The parameterization of the complex, receptor, and ligand species was performed in LeaP using the AMBER‐FF03 force field and ‘pbradii mbondi2’ atomic parameters. Snapshots were extracted at 7.5 nsec intervals from three independent 3 × 250 nsec (750 nsec) MD production trajectories. All thermodynamic binding energy terms were computed in kcal/mol and averaged out of 100 snapshots. MM‐PBSA computes the energy of binding (ΔG ^o _tot) as the difference of enthalpy (ΔH ^o _tot) and entropy (T.ΔS^o _tot) energy terms (ΔG ^o _tot = ΔH ^o _tot—T.ΔS^o _tot), where T denotes temperature (300 Kelvin) and Δ represents the difference in energies between the complex and the reacting species (Δ = complex – {receptor + ligand}). ΔH ^o _tot is the sum of the gas phase (ΔH ^o _gas) and aqueous phase (ΔG ^o _solv) energies. ΔH ^o _gas is a sum of van der Waals (ΔE ^o _vdw), electrostatic (ΔE ^o _el), and internal energies (ΔE ^o _int) of binding in the gas phase, which is determined as in the Cornell et al. force field (Cornell et al. 1995). ΔG ^o _el and ΔG ^o _nonel are electrostatic and non‐electrostatic contributions to the solvation‐free energy of binding (ΔG ^o _solv), respectively. Normal mode entropy energies (ΔS^o _tot) were determined by the nabnmode module of AMBER v22 (Case et al. 2023) as the sum of rotational, vibrational, and translational entropy energies.

2.15. Statistical Analysis

The results were presented as the mean ± standard deviations (SD), and each experiment was performed at least three times. All statistical analyses were performed with SPSS and analyzed by one‐way analysis of variance. p‐value < 0.05 was considered statistically significant.

3. Results and Discussion

3.1. Synthesis and Structural Characterization

The starting compounds (3, 5, 6) utilized in this study were synthesized using procedures previously reported by our group, starting from 1,2,3,4‐tetrahydroquinoline (THQ, 1) (Scheme 1) (Ökten et al. 2013). The bromination reaction of substituted THQ was an effective method for functionalizing both rings. Our method's simple bromination reaction formed 6,8‐dibromTHQ 2 in a nearly quantitative yield. Aromatization with DDQ gave dibromoquinoline, while copper‐induced substitution of methoxide afforded mono methoxide. Additionally, a single‐step reaction of THQ 1 with molecular bromine resulted in the formation of tribromoquinoline, easily converted to trimethoxide using a single‐step reaction procedure.

In our latest research, we have meticulously built upon our prior work on the regioselective bromination of methoxyquinoline derivatives (Çakmak and Ökten 2017). Initially, we successfully synthesized 3,6‐dibromo‐8‐methoxide and 3,5,8‐tribromo‐8‐methoxyquinoline through the bromination of 6‐bromo‐8‐methoxyTHQ, employing different equivalents of molecular bromine. In our current study, we treated 6‐bromo‐8‐methoxyTHQ 1 with an excess amount of molecular bromine (5 eq) and obtained a novel product, 3,5,6,7‐tetrabromo‐8‐methoxyquinoline (7), as the sole product in high yield (78%) (Scheme 2).

SCHEME 2.

Synthesis of 3,5,6,7‐tetrabromo‐8‐methoxyquinoline (7).

The structure of compound 7 was elucidated using 1D and 2D NMR techniques, including ¹H NMR, ¹³C NMR, HETCOR, and APT. Detailed NMR spectra are provided in the Figures S1–S4. As anticipated, the ¹H NMR spectrum of 7 displayed two doublet signals with meta coupling (⁴ J = 1.6 Hz) at δ_H 8.76 and 8.68 ppm, consistent with the expected structure when compared to the starting material. The APT spectra of 7 revealed quaternary carbon signals at δ_C 154.5, 151.1, 134.1, 127.9, 120.2, 103.2, 102.8 ppm, along with signals at δ_C 149.8 (CH), 137.3 (CH) and 61.2 ppm, the latter assigned to the OCH ₃ group. These carbon signals suggest the presence of bromine substitutions at the C‐3, C‐5, and C‐7 positions of the quinoline ring. The HETCOR experiment corroborated these findings, showing correlations between the carbon signals at δ_C 149.8 and 137.3 ppm and the two doublets at δ_H 8.76 and 8.68 ppm, respectively. Additionally, the aliphatic carbon signal at δ_C 61.2 ppm correlated with the characteristic methoxide singlet at δ_H 4.02 ppm, further supporting the structural assignment.

In our previous studies, we developed a selective synthetic method for 3,6‐dibromoquinoline (8) starting from 6‐bromoquinoline, which was subsequently converted into dimethoxide 9. Utilizing this approach, direct bromination of dimethoxide 9 with excess bromine resulted in the formation of 2,5‐dibromo‐3,6‐dimethoxyquinoline (10) with a high yield of 78% (Scheme 3) (Çakmak and Ökten 2017).

SCHEME 3.

Synthesis of 2,6‐dibromo quinoline and its bromination.

We have expanded our investigation of quinoline functionalization beyond the C‐5 position, leveraging the reactivity of the C2‐Br bromine group (Çakmak and Ökten 2017). Moreover, our successful synthesis of dimethoxide 9 from compound 8 inspired us to explore the bromination of trimethoxyquinoline 5. To achieve this, we prepared 3,6,8‐trimethoxyquinoline 5 from the corresponding tribromoquinoline 4 using previously reported methods (Scheme 1).

Unexpectedly, our research yielded surprising results. The bromination of trimethoxide 5 produced dibromide 11 instead of the anticipated tribromide 14. Moreover, we observed hydrolysis of the methoxide group at the C‐8 position, resulting in its conversion to a hydroxide group. This unexpected outcome suggests that the HBr generated during the electrophilic bromination reaction may have facilitated the degradation of the ether moiety (Lindley 1984; Doxsee et al. 1987) (Scheme 4).

SCHEME 4.

Synthesis of 5,7‐dibromo‐3,6‐dimethoxy‐8‐hydroxyquinoline (11).

The structure of compound 11 was characterized using various spectroscopic techniques, including ¹H NMR, ¹³C NMR, and IR spectroscopy. In the ¹H NMR, two aromatic doublet signals with meta coupling (⁴ J = 2.4 Hz) were observed at δ_H 8.47 and 7.70 ppm, which appeared further downfield than starting material 5 (Ökten et al. 2021). The disappearance of two doublets (Ökten et al. 2021) at δ_H 6.60 and 6.58 (⁴ J = 2.4 Hz) in ¹H NMR spectrum of compound 11 confirmed the substitution of bromine atoms at the C‐5 and C‐7 positions of the quinoline ring. Furthermore, the absence of a methoxide signal around δ_H 4.0 ppm, previously observed in compound 5, and the appearance of a new broad singlet at δ_H 4.85 ppm in the ¹H NMR spectrum of compound 11 indicated the conversion of the methoxide group into a hydroxide group. A broad absorption band further supported this transformation at 3353 cm⁻¹ in the IR spectrum of compound 11, characteristic of an O‐H stretching vibration. Additionally, the ¹³C NMR spectrum of 11 displayed methoxy and aryl carbon signals consistent with the proposed structure, providing further evidence for the successful transformation and bromination of compound 5.

We speculated that dibrominated trimethoxide (11) underwent ether cleavage in the presence of HBr, likely driven by its high strain energy. Computational calculations revealed that the total energy of compound 14 is significantly higher (45.069 kcal/mol) compared to quinolinol 11 (31.058 kcal/mol). This difference suggests that the C8‐OCH₃ group in compound 14 experiences substantial steric compression, primarily due to the spatial interference from the adjacent C7‐Br bromine atom and the lone electron pair of the nitrogen atom. This steric hindrance likely destabilizes the methoxy group, facilitating its cleavage and leading to the formation of compound 11.

We observed that the presence of nitro groups in the quinoline scaffold significantly reduces electron density through both inductive and resonance electron‐withdrawing effects. This reduction facilitates reactions with nucleophiles and enhances the ease of single‐electron transfer processes. For example, we successfully synthesized 6‐piperazinyl and 6‐morpholinyl quinolines from 6‐bromo‐5‐nitroquinoline via S_NAr nucleophilic substitution. These quinoline derivatives exhibited strong antiproliferative and apoptotic effects with low cytotoxicity across various cancer cell lines (Çakmak et al. 2020). Furthermore, we noted the significant inhibitory activity of these new piperazine‐ and morpholine‐substituted quinolines on human carbonic anhydrase isoenzymes (hCA I and II), cholinergic enzymes (AChE and BChE), and α‐glycosidase (Çakmak et al. 2020). These promising findings motivated us to extend our investigations to the nitration of 6,8‐dibromide 6, aiming to produce nitrated dibromoquinoline 17. These nitrated compounds hold great potential for the development of novel therapeutic agents targeting these crucial enzymes.

After the chemical reaction, 6,8‐dibromoquinoline (6) underwent selective nitration at the C‐5 position, yielding 6,8‐dibromo‐5‐nitroquinoline (17) as the sole product with an impressive yield of 96% (Scheme 5). In the ¹H NMR spectra of compound 17, the disappearance of two doublets with meta coupling, previously observed in the starting material 6, was noted. Instead, a new singlet appeared at δ_H 8.33 ppm in the aromatic region, confirming the attachment of the nitro group at the C‐5 position of the quinoline ring. Additionally, the chemical shift values of the remaining aromatic protons exhibited a downfield shift, consistent with the electron‐withdrawing effect of the NO₂ group. Notably, the characteristic H‐2 signal, which appeared at δ_H 9.04 (dd, ³ J = 4.2 Hz, ⁴ J = 1.6 Hz) in compound 6, shifted to δ_H 9.17 (dd, ³ J = 4 Hz, ⁴ J = 1.2 Hz) in compound 17, further supporting the successful nitration at the C‐5 position.

SCHEME 5.

Synthesis of 6,8‐dibromo‐5‐nitroquinoline (17) from starting 1,2,3,4‐tetrahydroquinoline and possible products.

It is hypothesized that the formation of dibromo nitroquinoline 17 during nitration is driven by the high strain energy of its precursor. Computational calculations revealed that the total energy of compound 18 is significantly higher (26.7086 kcal/mol) compared to the strain energy of nitroquinoline 17 (23.9001 kcal/mol) (Scheme 5). This difference suggests that the C‐7 position of the precursor experiences substantial steric compression due to the spatial proximity of the C6‐Br and C8‐Br bromine atoms, along with the lone electron pairs of the nitro group. Such steric hindrance likely makes the C‐5 position more accessible for nitration, leading to the selective formation of compound 17, which exhibits a more favorable and lower strain energy configuration.

3.2. Antiproliferative Activities of Compounds

In recent studies, it has been established that quinolines substituted with various groups demonstrate potent antiproliferative activity against specific cancer cell lines, as confirmed by sulphorodamine‐B dye (SRB), BCPE, and MTT assays (Ökten et al. 2013, 2020; Köprülü et al. 2021). In this investigation, we evaluated the antiproliferative potential of novel quinolines (7, 11, and 17) and their precursor compounds (3 and 5) against HeLa, C6, and HT29 cell lines using the BCPE assay at concentrations ranging from 5 to 75 μg/mL (Figure 1).

FIGURE 1.

The BCPE assay was utilized to examine the in vitro antiproliferative effects of quinoline derivatives (3, 5, 7, 11, 17) on C6, HeLa, and HT29 cancer cell lines. The cells were exposed to eight varying concentrations of quinoline compounds (p < 0.05). The provided data represent the mean results of three separate experiments conducted in triplicates.

Our findings revealed that all the novel quinoline derivatives (7, 11, and 17) exhibited significant inhibitory activity against all tested cancer cell lines when compared to the reference compound 5‐FU. In contrast, their precursor compounds (3, 5, and 6) (Ökten et al. 2013) showed no measurable antiproliferative effects against HT29, HeLa, and C6 cancer cell lines. Furthermore, neither 3,6,8‐trimethoxyquinoline (5) nor its precursor, 3,6,8‐tribromoquinoline (4) (Ökten et al. 2020), displayed any inhibitory activity against the C6, HeLa, and HT29 cell lines (Figure 1). These results highlight the importance of specific functionalization patterns on the quinoline scaffold in enhancing their biological activity.

Compound 11, derived from compound 5 through bromination, exhibited significantly higher antiproliferative activity on C6, HeLa, and HT29 cell lines, with IC₅₀ values ranging from 15.4 to 26.4 μM. In comparison, the reference drug 5‐FU displayed notably higher IC₅₀ values, ranging from 240.8 to 258.3 μM (Table 1). Among the newly synthesized compounds, compound 11 demonstrated the most potent inhibitory activity, with low IC₅₀ values of 15.4 μM (C6), 26.4 μM (HeLa), and 15.0 μM (HT29), highlighting its strong potential as an anticancer agent.

TABLE 1.

IC₅₀ values of compounds (μM).

Compounds	C6	HeLa	HT29
7	32.3	12.8	28.1
11	15.4	26.4	15.0
17	50.0	26.2	24.1
5‐FU	240.4	258.3	252.9

The substituents on the quinoline ring play a crucial role in determining their antiproliferative effects against cancer cell lines. Compounds 7 and 11, featuring bromine atoms at the C‐5 and C‐7 positions, demonstrated significant inhibition of C6, HeLa, and HT29 cell proliferation. In contrast, 3,6,8‐tribromoquinoline (4) and 3,6,8‐trimethoxyquinoline (5), with substitutions at the C‐3, C‐6, and C‐8 positions, exhibited no inhibitory activity.

Notably, the additional bromination of 3,6,8‐trimethoxyquinoline (5) at the C‐5 and C‐7 positions to yield compound 11 resulted in a substantial increase in antiproliferative activity, reflected by low IC₅₀ values against the tested cell lines. Furthermore, the conversion of the methoxy group at C‐8 into a hydroxyl group may have further enhanced the inhibitory potential of compound 11. This observation aligns with our previous findings, where brominated 8‐hydroxyquinolines demonstrated potent anticancer activity (Özcan et al. 2020; Ökten et al. 2017a, 2020).

Furthermore, the absence of antiproliferative activity in 6‐bromo‐8‐methoxy‐1,2,3,4‐tetrahydroquinoline (3), compared to the significant inhibitory effects observed in tetrabromoquinoline 7, which was produced through bromination of compound 3 at the C‐5 and C‐7 positions, strongly supports the critical role of bromine substitutions at these positions in enhancing antiproliferative activity.

Additionally, the introduction of a nitro group at the C‐5 position of the quinoline ring further amplified antiproliferative effects. Compound 17 (6,8‐dibromo‐5‐nitroquinoline) displayed remarkable inhibitory activity against C6, HT29, and HeLa cancer cell lines, with low IC₅₀ values of 50.0, 26.2, and 24.1 μM, respectively. In contrast, its precursor, 6,8‐dibromoquinoline (6), exhibited no inhibitory activity (Ökten et al. 2013).

These findings emphasize the synergistic impact of bromine and nitro substitutions on the quinoline scaffold in enhancing anticancer potency.

3.3. Cytotoxicity by LDH Assay

It is essential for compounds exhibiting high antiproliferative effects to also demonstrate low toxicity toward normal cells. To evaluate the cytotoxic potential of the compounds and assess indirect membrane damage, a lactate dehydrogenase (LDH) assay was performed (Ökten et al. 2017a).

The results revealed that compound 11 displayed a lower cytotoxic effect (15%–28%) across all tested cancer cell lines compared to the control drug 5‐FU (25%–42%) at its respective IC₅₀ concentrations (Figure 2).

FIGURE 2.

The cytotoxic activity of compounds 7, 11, and 17 was tested at their IC₅₀ concentrations on C6, HeLa, and HT29 cells. The LDH cytotoxicity test indicated a significant increase (p < 0.05) in the cytotoxicity of the cells. The percent cytotoxicity was reported as mean values of three independent assays with standard deviation (SD) included.

In contrast, compound 17 exhibited lower LDH release percentages (22% for HT29 and 20% for HeLa) compared to 5‐FU (24%–42%). However, both compound 17 (for HeLa) and compound 7 (for HT29) demonstrated higher cytotoxicities (~80%) relative to 5‐FU (Figure 2). Furthermore, for the C6 cell line, compounds 17 and 7 caused approximately 39%–43% membrane damage at their IC₅₀ concentrations.

When evaluating the balance between potency and cytotoxicity, the cytotoxic effects of the compounds at their IC₅₀ concentrations were largely comparable to 5‐FU, with the exception of compound 17 (for HeLa) and compound 7 (for HT29), which exhibited notably higher cytotoxicity.

3.4. The Evaluation of Apoptotic Potential With DNA Laddering

The induction of apoptosis is a crucial mechanism for eliminating diseased or abnormal cells. This process involves the activation of the enzyme caspase DNase, which causes specific cleavage of DNA at internucleosomal binding sites, resulting in fragments of approximately 200 base pairs, known as DNA ladders. The results of the DNA laddering test indicated that treatment with compounds 11 and 17 led to the formation of DNA ladders in HT29, HeLa, and C6 cell lines, while untreated cells did not show any DNA fragmentation (Figure 3). However, compound 7 did not induce apoptosis in any of the tested cell lines.

FIGURE 3.

(A) Agarose gel electrophoresis was utilized to detect DNA fragmentation in the HeLa cell line (left). The setup included: Lane 1 M: 1 kb DNA marker, Lane 2 C (+) (positive control, HeLa + CPT), Lane 3 (HeLa + compound 11), Lane 4 (HeLa + compound 7), and Lane 5 (HeLa + compound 17). (B) Agarose gel electrophoresis was also used to detect DNA fragmentation, this time in the HT29 cell line (middle). The setup consisted of Lane 1 M: 1 kb DNA marker, Lane 2 C (+) (positive control, HT29 + CPT), Lane 3 (HT29 + compound SO‐18), Lane 4 (HT29 + compound 7), Lane 5 (HT29 + compound 17). (C) Lastly, agarose gel electrophoresis was performed to detect DNA fragmentation in the C6 cell line (middle). The setup for this included: Lane 1 M: 1 kb DNA marker, Lane 2 C(+) (positive control, C6 + CPT), Lane 3 (C6 + compound 11), Lane 4 (C6 + compound 7), Lane 5 (C6 + compound 17).

3.5. Topoisomerase I Inhibition and Cell Migration

DNA topoisomerase I (Topo I) is a nuclear enzyme crucial for DNA replication, repair, and transcription, making it a key target for anticancer agents. The inhibitory activity of compounds 7, 11, and 17 on the Topo I enzyme was evaluated using a relaxation assay on supercoiled DNA via agarose gel electrophoresis. The supercoiled pHOT1 DNA incubated with Topo I led to relaxation (Lane 1), while Camptothecin (CPT), the control drug, and a quinoline derivative, displayed inhibition of the relaxation of supercoiled DNA by inhibiting the Topo I enzyme (Lane 2) (Figure 4). In a like manner, compounds 7 and 11 significantly inhibited the Topo I enzyme (Lane 4 and 5). However, compound 17 did not inhibit the Topo I enzyme (Lane 6) (Figure 4).

FIGURE 4.

The inhibition of human topoisomerase I (topo I) by 7, 11, and 17. SC DNA, supercoiled DNA; C(−), supercoiled DNA + Topo I; C(+), supercoiled DNA + Topo I + CPT; CPT: Camptothecin; Lane 4, DNA + Topo I + compound 11; Lane 5, DNA + Topo I + compound 7; Lane 6, DNA + Topo I + compound 17.

Cell migration, a vital process in various biological activities, was the focus of our study. We utilized the wound‐healing assay, a widely used technique, to investigate the impact of compounds 7 and 17 on the migration and proliferation of HT29 cancer cells. The cells were imaged at the beginning of the experiment (0 h) at 20, 46, and 62 h. After 62 h, it was found that compound 17 significantly inhibited the migration of HT29 cancer cells, while compound 7 showed no impact on the proliferation of HT29 cancer cell lines compared to untreated cells (Figure 5).

FIGURE 5.

Representation of the migration effect of compounds 7 and 17. C: Control, untreated HT‐29 cells; 7 and 17, HT‐29 cells + compounds 7 and 17 after 20 h, 46 h and 62 h.

These findings suggest that while compounds 7 and 11 effectively inhibit Topo I, only compound 17 exhibits anti‐migratory activity, highlighting their differential mechanisms of action in combating cancer cell proliferation and migration.

3.6. Dynamic Structures of Human Topoisomerase I‐DNA‐Ligand Complexes and MM‐PBSA Binding Studies

The average binding energies calculated by AutoDock Vina for camptothecin, compound 7, and compound 11 in complex with topoisomerase I (Topo I)‐DNA adduct were − 10.936 ± 0.023 kcal/mol, −7.715 ± 0.023 kcal/mol, and − 10.936 ± 0.023 kcal/mol, respectively. However, MM‐PBSA is considered a more reliable method for predicting binding energies consistent with experimental data (Andac et al. 2021). For MM‐PBSA analysis, molecular dynamics (MD) simulations were performed in three stages: relaxation, equilibration, and production phases. Following 1600 nsec of equilibration, three independent 250 nsec production runs (750 nsec total) were carried out. R.m.s.d curves revealed stable Topo I‐DNA complexes after 1000–1200 nsec, with ligands maintaining their positions in the DNA binding site throughout the simulations (Figure 6).

FIGURE 6.

R.m.s.d. curves determined through MD equilibrium (1‐ ~ 1600 nsec; black lines) and production (3 × 250 nsec between ~1600–1850 nsec; black, red, and green lines) trajectories of (A) compound‐7, (B) compound‐11, and (C) camtothecin in complex with Topo I.DNA adduct. R.m.s.d. curves on top, in the middle, and at the bottom represent Topo I, DNA, and the ligands, respectively.

Table 2 summarizes MM‐PBSA binding energies for Top1‐DNA‐ligand complexes, derived from 100 snapshots across three 250 nsec MD production runs. In the gas phase, compounds 7, 11, and camptothecin exhibited favorable electrostatic interaction energies (ΔE ^o _el = −5.58 ± 2.90 kcal/mol, −7.27 ± 3.02 kcal/mol, and − 30.31 ± 6.69 kcal/mol, respectively). In contrast, the unfavorable electrostatic energy contributions to the solvation of binding energies by compounds 7 and 11 (ΔG ^o _el = 19.98 ± 3.34 kcal/mol and 25.19 ± 4.08, respectively) outweigh the favorable ΔE ^o _el, resulting in net unfavorable electrostatic contributions (ΔE ^o _el + ΔG ^o _el) to the enthalpy of binding, calculated as 14.40 ± 3.12 kcal/mol and 17.92 ± 3.55 kcal/mol, respectively. This indicates that the binding of compounds 7 and 11 likely displaces all water molecules from the cavity of the nicked DNA binding site. The energy expended during binding is primarily used to align the ligands with the Topo I.DNA complex in a dual interaction process. Conversely, camptothecin exhibits a favorable electrostatic contribution to the enthalpy of binding (−6.78 ± 5.16 kcal/mol), as its favorable ΔE ^o _el predominates the unfavorable ΔG ^o _el (23.53 ± 3.62 kcal/mol). This suggests that not all water molecules are displaced from the binding site during camptothecin binding.

TABLE 2.

MM‐PBSA binding energies for camptothecin (CPT) and compounds 7 and 11 in complex with human Topo I.DNA dual. All energy values are given in kcal/mol. STD = standard deviation, n = 10.

Comp.	ΔH o gas			ΔG o solv		ΔH o tot
	ΔE o el	ΔE o vdw	ΔE o int	ΔG o el	ΔG o nonel	(ΔH o gas + ΔG o solv)	T.ΔS o tot	ΔG o bind	Kd (M)
7	−5.58 ± 2.90	−43.29 ± 2.02	0.00 ± 0.01	19.98 ± 3.34	−2.82 ± 0.09	−31.71 ± 2.65	−12.60 ± 14.82	−24.11	2.66 × 10−18
11	−7.27 ± 3.02	−41.78 ± 1.65	0.00 ± 0.01	25.19 ± 4.08	−3.00 ± 0.11	−26.86 ± 6.12	−9.89 ± 19.29	−16.97	4.27 × 10−13
CPT a	.‐30.31 ± 6.69	−56.14 ± 2.65	0.00 ± 0.00	23.53 ± 3.62	−3.99 ± 0.08	−36.60 ± 3.49	−12.98 ± 12.31	−23.62	6.06 × 10−18

Note: Mean enthalpy and entropy energy values were averaged out of 100 snapshots (n = 100).

ΔE ^o _el, ΔE ^o _vdw, and ΔE ^o _int are the electrostatic, Van der Waals, and internal energies, respectively, of binding in the gas phase, and the sum of them yields the energy of binding in the gas phase (ΔH ^o _gas).

ΔG ^o _el and ΔG ^o _nonel are electrostatic and non‐electrostatic contributions to the solvation‐free energy of binding (ΔG ^o _solv), respectively.

ΔH ^o _tot is the enthalpy energy of binding and is equal to [ΔH ^o _gas + ΔG ^o _solv].

ΔS ^o _tot is the entropy energy of binding and is a sum of translational (ΔS ^o _trans), rotational (ΔS ^o _rot), and vibrational (ΔS ^o _vib) entropies of binding.

ΔG ^o _bind is the binding free energy, which equals ΔH ^o _tot—T.ΔS ^o _tot, where T = 300 K.

K _d is the dissociation constant, which is determined by ΔG ^o _bind = R. T. ln K_d, where R = 1.987 cal/mol.K and T = 300 K.

^a CPT is Camptothecin, the control compound.

The binding of compounds 7 and 11, as well as camptothecin, results in favorable van der Waals interaction energies in the gas phase (ΔE ^o _vdw = −43.29 ± 2.02 kcal/mol, −41.78 ± 1.65 kcal/mol, −56.14 ± 2.65 kcal/mol, respectively). These are accompanied by favorable nonpolar contributions to the solvation energy (ΔG ^o _nonel = −2.82 ± 0.09 kcal/mol, −3.00 ± 0.11 kcal/mol, −3.99 ± 0.08 kcal/mol, respectively), leading to overall favorable hydrophobic/vdW energy contributions (ΔE ^o _vdw + ΔG ^o _nonel = −46.11 ± 1.06 kcal/mol, −44.78 ± 0.88 kcal/mol, −60.13 ± 1.37 kcal/mol, respectively) to the enthalpy of binding. The advantageous hydrophobic/vdW binding energy offsets the unfavorable effects of electrostatic interactions at the binding interface, resulting in favorable total enthalpy energies of binding (ΔH ^o _tot = −31.71 ± 2.65 kcal/mol, −26.86 ± 6.12 kcal/mol, −36.60 ± 3.49 kcal/mol, respectively) primarily driven by hydrophobic/vdW interaction forces in solution.

At 300 K, the total entropy terms (−T.ΔS^o _tot) for the binding of compounds 7, 11, and camptothecin were determined to be unfavorable, with values of 12.60 ± 14.82 kcal/mol, 9.89 ± 19.29 kcal/mol, and 12.98 ± 12.31 kcal/mol, respectively. However, these unfavorable entropy contributions are outweighed by the favorable ΔH ^o _tot energies, resulting in overall favorable binding free energies (ΔG ^o _bind) of −24.11 kcal/mol, −16.97 kcal/mol, and − 23.62 kcal/mol, respectively. This indicates that the binding of compounds 7, 11, and CPT to the nicked DNA binding site of Top1.DNA is primarily enthalpy‐driven. According to Table PBSA, the dissociation constants (K_d) for compounds 7, 11, and CPT are 2.66 × 10⁻¹⁸ M, 4.27 × 10⁻¹³ M, and 6.06 × 10⁻¹⁸ M, respectively. As illustrated in Figure 4, agarose gel electrophoresis relaxation assays on supercoiled DNA confirmed that compounds 7, 11, and camptothecin effectively inhibit Topo I activity. The MM‐PBSA results presented here demonstrate a reliable ranking for Topo I inhibition. Compound 7 appears to be a slightly more effective Top1 inhibitor compared to camptothecin, while compound 11 remains a promising inhibitor with great ΔG ^o _bind and K_d values.

Hydrogen‐bonding (H‐bonding) analysis from three 250‐ns MD production trajectories revealed specific interactions involving the guanidino N‐H protons of ARG364 of Top1. These protons form H‐bonds with N1 of compound 7 (observed in 10.4% of the production trajectories), O3 of compound 11 (4.2%), and N1 and O20 of camptothecin (9.4%), as illustrated in Figure 7A–C, respectively. Additionally, O8 of compound 11 interacts through H‐bonding with the 5'‐OH group of G11 in the nicked DNA, as shown in Figure 7B. Further H‐bonding interactions were identified for camptothecin, specifically between its C20‐OH group and the γ‐COO⁻ group of ASP533 of Top1 (86.3% of the production trajectories). Another interaction was observed between the ω‐NH₃ ⁺ group of LYS532 of Top1 and the C20‐O and C21 = O groups of camptothecin (7.7%), as depicted in Figure 7C.

FIGURE 7.

Representative structures of (A) compound 7, (B) compound 11, and (C) camptothecin in complex with Top1.DNA dual (surface, stick, and slab views on the left), determined out of 3 × 250 nsec MD production simulations. VdW/Hydrophobic (light green/pink dashed lines) and H‐bonding (dark green dashed lines) interactions are shown on the right. Nucleotides and amino acids were numbered based on the X‐ray structure of the Topo I.DNA dual (PDB ID:1t8i).

In terms of hydrophobic interactions, base pairs DC112:::DG11 and DA113::DT10 participate in π‐π stacking and van der Waals interactions with the quinoline rings of compounds 7 and 11 (Figure 7A,B, respectively), as well as with rings A–D of camptothecin (Figure 7C).

3.7. Bromination Strategy and Structure–Activity Relationship of Quinoline

The bromination of quinoline at different positions is crucial for synthesizing new derivatives. Quinoline bromides, in particular, serve as key intermediates for generating bioactive derivatives (Ökten et al. 2015). Despite extensive research over the years, the desired success in brominating quinoline molecules has often been hindered by technical challenges. A review of the literature reveals that brominated derivatives are typically synthesized under drastic conditions, such as high temperatures and concentrated acid environments, yielding predominantly mono‐brominated products (Figure 8). Under milder conditions, the reactions often result in the formation of complex structures (Eisch 1962).

FIGURE 8.

Bromination strategy of quinoline derivatives and their relative anticancer activities.

To address these challenges, our research group successfully brominated the quinoline ring by targeting the bromination of 1,2,3,4‐tetrahydroquinoline (1), a compound more active in the bromination process (Figure 8). The initial compound synthesized, 6,8‐dibromotetrahydroquinoline (2), exhibited significant inhibitory activity against three distinct cancer cell lines (Ökten et al. 2013). This observation highlighted the critical importance of functionalizing the C‐6 and C‐8 positions in tetrahydroquinolines for biological activity. However, the bioactivity diminished when this molecule was converted into its aromatic counterparts, 6,8‐dibromoquinoline (6) and 3,6,8‐tribromoquinoline (4), both of which showed no inhibition against any cancer cells (Ökten et al. 2013, 2020). These findings suggest that the C‐6 and C‐8 positions in the aromatic quinoline ring have a limited contribution to bioactivity.

To further explore this observation, methoxy, and nitro derivatives were synthesized, leveraging the substitutive flexibility of the bromine atom (Figure 8). Analysis revealed that di‐ and tri‐methoxy quinoline derivatives, such as compounds 19 and 5, exhibited either selective inhibition or no activity against cancer cells (Ökten et al. 2020, Köprülü et al., 2018). This suggests that the loss of bioactivity in the aromatic ring is influenced more by the roles of the C‐6 and C‐8 positions than by the bromine atom itself.

Additionally, modifying the 6,8‐dibromotetrahydroquinoline (2) to a mono‐methoxy derivative (3) (Ökten et al. 2013) or converting 6,8‐dibromoquinoline (6) to a nitro derivative at the C‐5 position (Ökten et al. 2020) (Figure 8) produced varying effects on anticancer activity. For example, methoxylation of dibromotetrahydroquinoline (2) at the C‐6 position reduced its inhibitory activity. In contrast, nitration of dibromoquinoline (17) at the C‐5 position regained its bioactive properties. These findings indicate the critical roles of the bromine atom at the C‐6 position in tetrahydroquinoline derivatives and the nitro group at the C‐5 position in aromatic quinoline derivatives for bioactivity.

To evaluate the effects of the C‐5 and C‐7 positions on bioactivity, further bromination experiments were conducted on compounds 3 and 5. The results revealed that bromination at these positions significantly enhanced inhibitory activity (compounds 7 and 11) compared to the starting materials (3 and 5, respectively). These findings were confirmed by our earlier work, where bromination of 6,8‐dimethoxyquinoline (19) at the C‐5 position yielded compound 20, which exhibited similar bioactive properties (Ökten et al. 2020).

4. Conclusion

Our study presents efficient and selective synthetic methods for producing three polyfunctional quinoline compounds. We successfully developed two regioselective routes to prepare di‐ and tetrabrominated methoxyquinolines at specific positions under mild reaction conditions. These methods offer high yields, are cost‐effective, and enable large‐scale synthesis from commercially available tetrahydroquinoline (Scheme 1). The incorporation of bromine atoms into these quinoline derivatives provides versatile opportunities for further functionalization and structural diversification.

The brominated methoxy derivatives of 1,2,3,4‐tetrahydroquinoline (3) and 3,6,8‐trimethoxyquinoline (5) demonstrated high reactivity toward bromination, enabling the synthesis of a series of polysubstituted analogs .

The experimental biological evaluation and molecular dynamic studies revealed that 5,7‐dibromo‐3,6‐dimethoxy‐8‐hydroxyquinoline (11) and 3,5,6,7‐tetrabromo‐8‐methoxyquinoline (7) exhibited potent antiproliferative effects against various cancer cell lines and human topoisomerase I (Topo I) inhibition activity, with compound 11 displaying the highest efficacy. Moreover, these compounds showed lower cytotoxicity compared to conventional chemotherapy drugs, suggesting their potential as safer therapeutic alternatives.

The results from apoptosis induction and wound‐healing assays further emphasize the therapeutic promise of these quinoline derivatives. This study highlights the importance of regioselective synthesis strategies in developing structurally diverse compounds with significant anticancer potential.

Future research should focus on detailed mechanistic studies and in vivo evaluations to fully harness the potential of these compounds in cancer therapy.

Conflicts of Interest

The authors declare no conflicts of interest.

Supporting information

Data S1.

Data Availability Statement

Data sharing is not applicable to this article as no new data were created or analyzed in this study.