Studies on copper (II) interaction with the (CCG) 12 repeats sequence: An insight into genomic instability in neurodegeneration

Abstract

Background

Studies indicate that a comprehensive understanding of the mechanisms underlying neurodegeneration in Alzheimer's disease, Parkinson's disease, and related disorders is still not clearly understood. Fragile X-associated tremor/ataxia syndrome (FXTAS) is one such late-onset neurodegenerative disorder characterized by premutation alleles (55–200 CGG repeats) in the genome. In parallel, dysregulation of trace metal homeostasis, particularly copper (Cu) imbalance, has been documented in Alzheimer's disease, Parkinson's disease, and FXTAS. However, the precise role of trace metal dyshomeostasis as a disease modifier in neurodegeneration, and its contribution to disease onset and progression, remains poorly understood.

Objective

To understand the interaction of Copper (II) with the (CCG)₁₂ repeats sequence using spectroscopic and computational modeling studies.

Methods

(CCG)₁₂ repeats were done by the phosphonamidite technique with a 380B ABI automated synthesizer. The binding of Cu to (CCG)₁₂ was studied using Circular Dichroism, Fluorescence, UV spectroscopy, and a molecular docking model.

Results

The current study shows conformational changes to the (CCG)₁₂ repeat sequence by abnormal interactions of Cu using CD and other spectroscopic methods. Circular dichroism results confirm binding of Cu to the base pairs of DNAs and alter the structure from B-DNA to an altered B-DNA conformation. Further, molecular docking studies reveal the Cu binding to DNA directly through the hydrogen bond formation between G-C base pairs.

Conclusions

The altered form of B-DNA will affect the integrity of DNA, which in turn modulates the replication and transcription processes, leading to genomic instability and cell dysfunction in neurodegenerative disorders.

Keywords

Alzheimer's disease CCG sequence copper Fragile X-associated tremor/ataxia syndrome (FXTAS)genomic instability neurodegeneration Parkinson's disease

Introduction

Growing evidence implicates genomic instability in neurological disorders like Fragile X-associated tremor/ataxia syndrome (FXTAS) through the presence of CGG repeats in the genome, contributing the cellular dysfunction.¹ In parallel, the imbalance in metal homeostasis and mitochondrial impairment have been implicated in FXTAS and could enhance oxidative damage initiated by CCG-repeat-induced genome instability.² Copper (Cu), a key player in redox biology and neuronal cell function, is assumed to be a modifier of disease biology. Still, its specific role in FXTAS disease biology and in neuronal cell dysfunction in brain disorders remains incompletely characterized.^3–8 The role of CCG repeats in genomic instability in FXTAS still needs to be understood.⁹ The FMR1 premutation consists of 55–200 CCG repeats in the 5′-UTR.¹⁰ The expanded CCG may have a role in multiple instability phenotypes, like (a) somatic and intergenerational repeat length instability, (b) chromosomal fragility at the FRAXA site (c) formation of unusual nucleic acid structures like hairpins, G-quadruplexes, R-loops that modify replication and transcription, and (d) increased DNA damage through strand breaks.¹¹ These genomic changes produce a genomic environment prone to mutations and altered gene expression in affected cells, contributing to cell dysfunction.^12–21

In the present study, we focused on understanding the role of Cu in modulating the genomic instability through Cu and (CCG)₁₂ interactions.

Methods

Synthesis of (CCG)₁₂ repeats

Synthesis of CCG repeats was done by the phosphonamidite technique with a 380B ABI automated synthesizer. Unprotected oligonucleotide bases were obtained in the presence of ammonia, and the process was carried out at 55°C for 12 h. Ethanol was used to precipitate the sample, and it was purified using FPLC and freeze-dried.

The synthesis of (CCG)₁₂ oligonucleotide was carried out at 55°C in the presence of ammonia to remove the phosphate and base protecting functional groups. This has been a routine experimental procedure as indicated by Beaucage and Caruthers.²² This becomes particularly important for GC-rich sequences. These conditions are routinely used in phosphoramidite-based DNA synthesis. They are particularly important for GC-rich sequences, such as CCG repeats, which tend to form secondary structural elements such as looped structures. This may be more resistant to deprotection under milder conditions.

The high temperature does not cause perturbations to the integrity of the oligonucleotide. While it may transiently affect the hydrogen bonding, the diester backbone of the DNA is retained. Further, any loss of base pairing is reversible upon cooling.

Preparation of DNA stock solution

A stock solution of double-stranded (CCG)₁₂ repeat. The DNA sequence was prepared by dissolving 1.0 mg of DNA in 200 µl MilliQ water (5 µg/µl), and further dilutions of DNA were prepared as required using 5 mM Tris-HCl of pH 7.4 for UV and CD studies. For measurement of Tm, the DNA samples were buffered in 10 mM HEPES (pH.7).

Preparation of copper chloride stock solution

Copper chloride (CuCl₂.2H₂0) dihydrate was purchased from Merck Schuchard, Germany. A 50 mM Stock solution was obtained by dissolving 8.5 mg of copper chloride in 990 ml of MilliQ water. Dilutions were prepared depending on the needs of the experiment. We used the Cu concentration equivalent to 11–22 µmol/L as observed in Parkinson's disease^3–8

Ethidium bromide stock solution

Ethidium bromide (C₂₁H₂₀BrN₃) (EB) was purchased from Amersham Life Sciences, UK. The 5 mg/ml stock solution of EtBr was obtained by dissolving EtBr in milliQ water and was stored at 4°C in the dark. Dilutions were prepared depending on the needs of the experiment.

Circular dichroism studies

A Jasco-J-715 Spectropolarimeter was used to measure the changes in (CCG)₁₂ sequence DNA conformation at 25°C using a 1 mm path length quartz cuvette between 200 and 320 nm wavelength. The spectra were recorded by taking an average of four scans at 20 nm /min. (CCG)₁₂ sequence samples (20 µg/ml) were prepared in Tris–HCl buffer of 7.4 pH and 5 mM with and without Cu. The buffer background was subtracted from the spectra using the built-in feature of Jasco software.

Thermal denaturation studies of (CCG)₁₂ repeat sequence with and without Cu were recorded using the Jasco J-715 Spectropolarimeter with the temperature control system (Peltier type Model PTC-348WI). Spectrum was recorded between 200 nm and 320 nm wavelength, between a temperature range of 20°C to 90°C. The CD spectral characterization of DNA conformations is analyzed.²³

UV/VIS absorption studies

A Jasco V-530 Spectrophotometer (Jasco, Japan) was used to study the absorption spectra of (CCG)₁₂ and its binding ability with Cu, (CCG)₁₂ sample (10 µg) were dissolved in 400 µl of Tris–HCl buffer in the absence and presence of increasing Cu concentrations (25 µM to 5 mM) (Total volume of sample used for UV/VIS absorbance studies was 400 µl). The gradient absorbance spectra were measured between 220 nm and 320 nm wavelength using a 1-cm pathlength quartz cuvette. The buffer baseline was subtracted using the built-in feature of Jasco software.

Fluorescence studies

To study the dynamics of DNA, fluorescence emission studies were carried out using equimolar concentrations of (CCG)₁₂ repeat sequence and EB (1:1). The effect of different concentrations of Cu on the EB-DNA complex was analyzed. Excitation of DNA/EtBr solutions was carried out at 530 nm, and emission spectra were recorded between 550 nm to 650 nm using a Jasco J-600 spectrofluorometer.

In silico molecular docking studies

Molecular docking, a valuable computational method for rational drug design, can be used to predict the binding orientation and conformations of drugs, small molecules, peptides, metal Ions, and proteins through binding affinity by scoring functions. The docking study defines and evaluates the key DNA- Cu (II) complexes with nucleotide bases and reveals detailed information on conformational change and binding patterns. Initial Molecular docking studies were carried out with Cu and the crystal structure of two models with (CCG) repeats in duplex DNA, PDB ID 5XEW (A chain -TTCCGCCGCCGAA), B-chain (TTCCGCCGCCGAA), a duplex model for (CCG)₁₂, and PDB ID 1NOQ [A chain: (CCGCCG) B chain (CCGCCG)] from the Protein Data Bank.²³ Cu ligands were retrieved from the Pubchem database.^24,25

Further, our study also involved inspecting how copper (Cu²⁺) binds differently to various DNA forms, hence B-, and Z-DNA conformations using crystallographic structures PDB IDs: 1BNA, (B-form) 4XSN, (Z-form) 8OE8, (Z-form), and molecular docking results. Minimization of the retrieved structures was done using the conjugate gradient protocol using the CHARMM force field and ligands minimized with the Max-Steps protocol incorporated in the Discovery Studio 3.5 software.²⁵ The binding sites were identified using the define and edit binding site tool for forms of DNA. The docking was performed with CDOCKER.(Discovery Studio 3.5 (BIOVIA, San Diego, CA, USA).²⁶

Molecular docking was performed using a two-stage strategy involving blind docking followed by focused docking. Initially, blind docking was carried out to identify potential Cu²⁺ binding regions across the entire DNA structure without any prior bias. The search space was defined to encompass the whole DNA molecule, allowing unrestricted sampling of both major and minor grooves as well as phosphate backbone regions. This approach ensured unbiased identification of preferred metal-binding hotspots, particularly within CCG repeat motifs. Subsequently, focused docking was performed by restricting the grid box to the regions identified from blind docking, specifically around guanine N7, cytosine O2, phosphate oxygen atoms, and groove-accessible sites. This refinement allowed accurate estimation of interaction energies and coordination geometries.

The Cu²⁺ ion has a + 2 formal charge, consistent with its physiologically relevant oxidation state. Partial charges and non-bonded parameters were assigned using the CHARMm force field, which has been widely employed for metal–biomolecule interactions. Before docking, all DNA structures and Cu²⁺ ions were subjected to energy minimization using the conjugate gradient method to remove steric clashes and optimize geometry. For each docking experiment, multiple initial ligand conformations were generated, and up to 1000 MD-based docking trials were performed per system to ensure adequate conformational sampling. The resulting poses were ranked using CDOCKER interaction energy, which accounts for van der Waals and electrostatic contributions. The lowest-energy and chemically plausible poses were selected for further analysis.

To validate the docking protocol, Cu²⁺ docking was performed not only on the (CCG)₁₂ duplex model (PDB ID: 5XEW) but also on experimentally resolved Cu-bound DNA structures, including canonical B-DNA (1BNA) and Z-DNA structures (4XSN and 8OE8). The docking results reproduced known Cu²⁺ coordination patterns, including binding to guanine bases and phosphate oxygens, with coordination distances comparable to reported crystallographic data. This cross-validation confirms the reliability of the docking methodology and supports the biological relevance of the predicted Cu²⁺–(CCG)₁₂ interactions.

Results

Circular dichroism studies

Figure 1(a) shows the CD spectra of the (CCG) ₁₂ sequence, indicating a characteristic feature of the right-handed B-form helicity with a negative band at 254 nm and a positive band at 283 nm. These two bands are very sensitive to the interaction of small molecules. The effect of Cu (5–25 µM) (physiological concentration) on DNA shows a decrease in the intensity of the positive band with a blue shift to 279 nm (4 nm) and disappearance of the negative peak at 254 nm up to the concentration of 50 µM. With higher concentrations of Cu from 100 µM and 250 µM, it has further decreased in the positive band with a red shift to 288 nm (5 nm), with a complete disappearance of the peaks at 254 and 207 nm as shown in Figure 1(b)-(e). The observed change in the intensity and an evident blue and red shift is indicative of disruption of the stacking between the bases of the DNA sequence, due to the interaction of Cu with specific binding sites of DNA. Cu binding to DNA may push the strands of DNA apart to accommodate the incoming Cu ions. The insertion of Cu ions between G-C base pairs disrupts the H bonding and destabilizes the double helix.^27–29 A decrease in the intensity of the peaks could be related to an increase in the winding angle of the DNA helix due to the super-helical tension caused by Cu.²⁶ Loss of negative bands is evidence for the degradation of the right-handed form helicity of DNA.^26–29 The interaction of copper with DNA causes perturbations of the DNA structures of each strand and disturbs the secondary structure of DNA due to unwinding of the double helix, and further, the winding angle increases, the magnitude of the band decreases, and thus starts to remodel the secondary structure, which drives the conformational changes from B-DNA to altered B-DNA like B-C-A mixed conformation.^26–30

Figure 1.

(a) Circular dichroism studies showing the conformational changes of (CCG)₁₂ sequence in the presence of Cu. (a) (CCG)₁₂ alone, (b) 10 μM CuCl₂, (c) 20 μM CuCl₂, (d) 50 μM, and (e) 100 μM. (b) Circular dichroism studies showing the conformational changes of (CCG)₁₂ sequence in the presence of Cu. (a) (CCG)₁₂ alone, (b) 100 μM CuCl₂, (c) 250 μM CuCl₂.

UV absorption studies

UV-vis absorption is an effective method for detecting the interaction between Cu and the (CCG)₁₂ sequence. The absorption spectra show that the peak maximum at 259 nm is due to the strong absorption of purine and pyrimidine bases in DNA, as shown in Figure 2(a). With the gradual addition of Cu, the absorbance of the peak increases. The hyperchromic originates from DNA duplex breakage due to the interactions of Cu and disturbs the base pair of DNAs, causing significant changes in the electronic environment of the DNA-Cu complex.^29,30 Cu binding affects the H bonding in the guanine–cytosine pair and stacking pattern that results in damage to the DNA double helix, driving the conformational change in the DNA molecule. The binding of Cu is further supported by fluorescence studies.

Figure 2.

Effect of Cu on the Absorption spectra of (CCG)₁₂ in 5 mM Tris-HCl buffer (pH.7.4), (CCG)12 sequence, with 10,25,100 and 250 μM CuCl_2, showing an increase in order.

Fluorescence studies

To better understand the Cu binding to DNA, the competitive binding between fluorescent probes and Cu for DNA was done by using equimolar concentrations of (CCG)₁₂ and EB. Fluorescence spectra of (CCG)₁₂-EtBr complex show an emission maximum at 590 nm, as shown in Figure 3(a). The fluorescence intensity of the DNA-EB complex decreased gradually with the increasing concentrations of Cu. The percentage of decrease is shown in Figure 3(b). The decrease in the fluorescence intensity is due to Cu substituted for EB and dissociated EB molecules from (CCG) ₁₂ –EB complex into the solution. Cu binds with bases and forms a new non-fluorescent complex Cu-(CCG) ₁₂ –EB, which causes the fluorescence quenching of DNA–EB complex.^19,27–29^,31 The combined studies of fluorescence and CD studies show that the molecular conformation of DNA was affected, and the process of quenching will address the destabilization of DNA and transform the B-DNA to altered B-DNA.

Figure 3.

(a) Emission spectra of EtBr bound to (CCG)₁₂ in the absence and presence of Cu. (a) (CCG)₁₂-EtBr Complex, (b) 10 μM Cu, (c) 25 μM Cu, (d) 50 μM Cu, (e) 75 μM Cu, (f) 100 μM Cu, and (g) 250 μM Cu. (b) Percentage of decrease in the fluorescence intensity upon increasing the concentration of Cu.

Effect of temperature and pH on the (CCG) sequence conformation

To study the effect of temperature on DNA stability and conformation, we measured the CD spectra in the absence and presence of Cu at different temperatures, as shown in Figure 4(a) and 4(b). The CD spectrum of the (CCG)₁₂ sequence shows the characteristics of B-DNA. The decrease in the positive peak, accompanied by a blue shift to 279 nm (4 nm) and a complete loss of the negative peak at 245 nm, provides clues to the disruption of base pairs in the DNA structure as the temperature increases. It was saturated to 80°C. A similar pattern of decrease in the intensity of both the peaks with blue shift was seen and was saturated to at 70°C with 25 µM Cu as shown in Figure 4(b). The isodichroic point at 272 nm indicated that both bands are sensitive to Cu and temperature. The structural change is due to the distortion of the DNA double helix due to binding with Cu. Cu has a very high affinity towards G-C base pairs and tends to lower base pairing and stacking at higher temperature.^27–29 The hydrogen bonds between the complementary DNA strands are broken and disturb the secondary conformation that affects the thermal stability of the (CCG)₁₂ sequence.

Figure 4.

(a) Circular dichroism studies showing the conformational changes of the (CCG)₁₂ sequence in the presence and absence of Cu. (CCG)₁₂ alone with varying temperature: (A) 30, (B) 40, (C) 50, (D) 60, (E) 70, and (F) 80°. (b) (CCG)₁₂ with 25 μM CuCl₂. (CCG)₁₂ with (A) 30, (B) 40, (C) 50, (D) 60, and 70°. (c) CD spectra of (CCG)₁₂ sequence as a function of temperature and also with Cu. A.a.(CCG)₁₂ with 30°, A.b. (CCG)₁₂with 25 μM Cu at 30°. (d) B.a. (CCG)₁₂ alone at 70°, B.b. (CCG)₁₂ with 25 μM CuCl₂ at 70°. (e) CD spectra of (CCG)₁₂ sequence as a function of pH. (a) (CCG)₁₂ pH 7.4, (b) pH 6, (c) pH 5, (d) 6-h incubation, (e) 13-h incubation.

The UV-melting profiles have shown similarity with CD results. T_m of DNA depends on the percentage of GC content and hydrogen bonding between GC base pairs present; therefore, DNA with higher GC content will have a higher T_m.^31–33 (CCG) sequence contains higher GC content, hence, a higher T_m value of 80°C, and with 25 µM Cu, it is 70°C. The results show that the impact of incorporating copper into (CCG)₁₂ sequence causes structural perturbations, causing chemically modified structures to lower base pairing and stacking due to disturbances in DNA base stacking. The DNA-Cu complex is sensitive to higher temperatures and effectively disturbs the secondary structure of each strand due to unwinding of the helices. The Tm data show that Cu ions bind to phosphate and nucleotide bases of DNA, leading to unwinding of the two strands at higher temperatures, leading to its instability.^31,33

The conformational changes on varying the pH, ranging from pH 7 to pH5 is shown in Figure 4(c) CD spectra shows very little change at both positive and negative peaks on varying the pH. The changes have been observed at pH 5, and no changes at pH 7 and 6 after incubation at room temperature for 6 h. At pH 5, the spectra show an abrupt increase in intensity at the positive peak at 280 nm and a negligible change at the negative peak at 254 nm. The spectra show the stabilization of the DNA and an increase in the intensity without a change in the negative band is evidence for the change from B-DNA to A-DNA. The conformational changes and instability are due to the increased hydrogen ion concentration of a DNA solution results in proton binding to one of the cytosines in the C-C mismatched pair and stabilizes the double-handed DNA due to the winding of the DNA triggered due to the protonation of the cytosine.^33–36

Docking studies

Docking results reveal the coordination of Cu (II) complex on binding to DNA, indicating a covalent mode of DNA binding of the complex across Guanine of both A chain and B chain, with N7 nitrogen and 06 as the preferred coordination site of Cu. The docking results are shown in the Figure 5(a)-(d) and are tabulated in Tables 1 and 2. The results indicate that the Cu (II) ion is coordinated by one nitrogen atom of GUA 21 of B chain, then by a different nitrogen atom of GUA 8 from the opposite chain, as well as by the amide oxygen o6 atoms of GUA 21 B chain and GUA 8 of A chain. Revealing the significant prevalence of intrastrand GG cross-links. The N7 nitrogen of guanine is known to be an H-bond acceptor in the major groove, allowing the guanine-N7 to be the preferred coordination site of Cu2 + . From our studies, it is confirmed that Cu2 + binds via intrastrand bonds to two of the neighboring guanines Figure 5(a). Interaction of Cu causes perturbations in DNA structure by increasing the distance between the adjacent base pairs. This helix distortion causes one of the bases to rotate around its glycosidic bond. This results in a small tilt in the adjacent base pair, leading to DNA instability. This can further affect the H bonding between the guanine–cytosine pair, and with the greater degree of twisting of the chain, may affect sugar puckering and the stacking pattern. Through these events, there will be a distortion in G-C base pairs. Combined studies of CD spectroscopy and in silico Molecular docking results confirm the perturbations of the DNA helix due to cu binding. Our results revealed that metal binding to DNA plays an important role in determining its structure and chemical reactivity. Hence, further docking study explored the interesting behavior of copper (Cu²⁺) binding across different DNA forms, including B-, and Z-DNA, revealing how the nature of the helix influences its coordination pattern and binding strength.

Figure 5.

(a) Docking interactions of Cu (II), with CCG DNA repeat 5XEW (A chain -TTCCGCCGCCGAA), B-chain (TTCCGCCGCCGAA), a duplex model for (CCG) showing octahedral coordination of Cu across N7, 06 of Guanine 8 A chain and Guanine 21, of B chain across 5XEW. Showing the overlay between Native DNA and Cu bound form. The figures highlight the differences in bonding distances between bound and unbound forms. (b) Docking interactions of Cu (II), with—B-DNA (Canonical) d(CGCGAATTCGCG)₂ (Dickerson dodecamer). (c) Docking interactions of Cu (II), with 4XSN Z-DNA d(CGCGCG)₂. (d) Docking interactions of Cu (II), with 8OE8, Z-DNA, d(CGCGCG)₂.

Table 1.

Showing distinct Cu²⁺–DNA interactions that depend strongly on the helical geometry and groove accessibility.

L. NO	PDB ID DNA form/Sequence / motif	Key structural insights	DNA CHAIN native with copper	Nucleotide atoms	Hydrogen bond distance in Å	Highest -CDOCKER energy(kcal·mol−1)	Highest –CDOCKER interaction energy (kcal·mol−1)	Relevance to cu²⁺ binding
1	1BNA-B-DNA (Canonical) d(CGCGAATTCGCG)₂ (Dickerson dodecamer)	Benchmark B-form double helix — wide major groove, narrow minor groove. Often used as a model in Cu²⁺ molecular dynamics (MD) studies.	A: Adenine 6	A:DA6:N173 - COPPER_(II)_ION:N1	2.32 Å	16.7786	12.7863	Analysis show Cu²⁺ bind near phosphate oxygens(O2) & T7, T19, Adenine N1 and Thymine N1 sites. Demonstrates inner-sphere coordination — key for understanding metal-driven B→Z transitions.
			A: Thymine7	COPPER_(II)_ION:N1 - A:DT7:N205	2.14 Å
			B: Thymine19	COPPER_(II)_ION:N1 - B:DT19:N584	2.13 Å
			B: Thymine 19	COPPER_(II)_ION:N1 - B:DT19:N586	2.1 Å
			B: Adenine 18	COPPER_(II)_ION:N1 - B:DA18:N1	2.30 Å
			B: Thymine 20	COPPER_(II)_ION:N1 - B:DT20	4.80 Å
2	4XSN Z-DNA d(CGCGCG)₂	Crystal structure showing Cu²⁺ binding to Z-DNA. Cu²⁺ directly coordinates N7 of guanine and phosphate oxygen atoms, forming square-planar geometry.	B: Guanine 4	COPPER_(II)_ION:Cu1 - B:DG4:OP2	3.84 Å	21.3621	9.43326	Analysis show Cu²⁺ bind G4OP2,N3,C5OP2,G4OP2
			B: Cytosine 5	COPPER_(II)_ION:Cu1 - B:DC5:OP2	3.58 Å
			B: Guanine 4	COPPER_(II)_ION:Cu1 - B:DG4:N3	2.86 Å
			B: Cytosine 5	COPPER_(II)_ION:Cu1 - B:DC5:O5'	2.77 Å
			B: Guanine 4	B:DG4:H21 - COPPER_(II)_ION:Cu1	2.37 Å
3	8OE8 Z-DNA d(CGCGCG)₂	Recent (2024) high-resolution structure of Cu-bound Z-DNA, confirming earlier 4XSN findings. Shows multiple Cu²⁺ coordination modes and increased Z-helical stability.	A: Cytosine 3	COPPER_(II)_ION:Cu1 - A:DC3:O2	2.30 Å	29.2873	28.8833	Provides details of Cu²⁺ coordination with C3-O2.
			B: Cytosine 3	COPPER_(II)_ION:Cu1 - B:DC3:O2	2.46 Å
			A: Guanine 4	A:DG4:H22 - COPPER_(II)_ION:Cu1	2.59 Å

Table 2.

Docking interaction details of copper with DNA(5XEW) with and without copper.

PDB ID DNA form/Sequence / motif	DNA chain native with out copper	Nucleotide atoms	Hydrogen bond distance in Å	DNA chain native with copper	Nucleotide atoms	Hydrogen bond distance in Å	Estimated inhibition constant (Ki) for cu²⁺ binding to (CCG)\₁₂ repeats	Highest -CDOCKER energy(kcal·mol−1)	Highest –CDOCKER interaction energy (kcal·mol−1)	Structural implication
5XEW (CCG)\₁₂ duplex	GUANINE 8	N(7)-Cu (II)	5.279 Å	GUANINE 8	N(7)-Cu (II)	5.153 Å	1.6 × 10⁻¹⁹	22.6447	16.4999	Major groove (CCG tract)-Local helix distortion (Electrostatic interaction with GC-rich region)
	A chain	O(6)-Cu (II)	3.283 Å	A chain	O(6)-Cu (II)	3.152 Å		22.6447	16.4999
	GUANINE21,	N(7)-Cu (II)	4.379 Å	GUANINE21	N(7)-Cu (II)	4.550 Å		22.6447	16.4999	Base unstacking
	GUANINE21,	N(7)-Cu (II)	4.379 Å	GUANINE21	N(7)-Cu (II)	4.550 Å				Guanine-rich site
	B chain	O(6)-Cu (II)	2.287 Å	B chain	O(6)-Cu (II)	2.384 Å				Groove-based stabilization

In this study, Cu (II) coordination was examined across the B-, and Z-DNA conformations using crystallographic structures (PDB IDs: 1BNA, 4XSN, 8OE8,) and molecular docking results are summarized in Tables 1 and 2 and Figure 5(b)-(d). The analysis helps to understand how Cu (II) binding influences coordination geometry, binding affinity, and structural stabilization in different DNA forms. The results show that Cu (II) interacts differently with DNA depending on its helical structure and groove accessibility. The B-form DNA (PDB ID: 1BNA) represents the normal right-handed double helix found under physiological conditions. Docking results show Cu²⁺–ligand distances between 2.13 and 2.34 Å, involving phosphate oxygens (O2) and base atoms of T7, T19, and A1. These short distances indicate inner-sphere coordination, where Cu²⁺ forms direct electrostatic bonds with oxygen donors from the bases or backbone.

The CDOCKER interaction energy (12.78 kcal·mol⁻¹) suggests a moderate but specific binding affinity. Overall, the data indicates that in B-DNA, Cu (II) interacts mainly with phosphate oxygen and polar base edges, and its flexible, hydrated environment allows both direct and water-mediated binding. Such binding may play a role in metal-induced B→Z transitions. Similarly, docking interactions of Cu (II) Binding in Z-DNA (4XSN) were performed, and the results reveal that, in Z-DNA (PDB ID: 4XSN), Cu²⁺ binds directly to G4(OP2), N3, and C5(OP2) atoms with distances between 2.37 and 2.86 Å, indicating inner-sphere coordination. Longer distances (3.58–3.84 Å) suggest outer-sphere interactions or partial occupancy of Cu (II) in the lattice.

The negative docking energy (–21.36 kcal·mol⁻¹) points to stronger electrostatic stabilization and tighter metal association compared to B-DNA.

Likewise, Cu (II) Binding in Z-DNA (8OE8), Z-DNA structure (8OE8), shorter Cu–O distances (2.30–2.46 Å) confirm direct coordination, particularly with C3–O2 atoms. Compared to 4XSN, this structure shows more localized and uniform Cu²⁺ binding, indicating stronger site-specific interactions. The higher docking interaction energy (∼28.88 kcal·mol⁻¹) supports enhanced binding affinity, consistent with experimental observations. The zigzag phosphate backbone and alternating guanine conformations in Z-DNA provide a favorable electrostatic environment for Cu (II), allowing it to bridge nearby phosphate and base atoms and further stabilize the structure. Thus, the overall docking study shows that Cu (II) binds strongly and specifically to B-DNA, followed by Z-DNA forms. The differences arise from variations in groove geometry, hydration, and phosphate accessibility among these conformations.

Our docking results tabulated in Table 2 reveal the preferential binding of Cu²⁺ to Guanine 8 (A chain) and Guanine 21 (B chain) via N7 and O6 atoms of the major groove. This stable Guanine centric coordination and inner sphere interactions are indicated by the reduction of the Cu–DNA coordination distances. Further, the low Ki value (1.6 × 10⁻¹⁹ M) and favorable –CDOCKER energies indicate a strong affinity of Cu²⁺ for GC-rich CCG repeats. Table 2 indicates that the Ki values are computational estimates derived from CDOCKER interaction energies and are intended for relative comparison of Cu²⁺ affinity toward CCG repeat DNA. Cu–DNA interactions are predominantly electrostatic rather than classical hydrophobic interactions. Further, the –CDOCKER energy and –CDOCKER interaction energy were used as key metrics to evaluate molecular docking results. The –CDOCKER energy reflects the overall quality of the docking pose by accounting for both ligand–protein interactions and the internal strain of the ligand, ensuring energetically stable and structurally feasible conformations within the binding site. In contrast, the –CDOCKER interaction energy specifically represents the strength of protein–ligand binding by quantifying non-bonded interactions such as van der Waals and electrostatic forces. Ligands were primarily ranked based on –CDOCKER interaction energy to identify compounds with the most favorable binding affinities.

Further, our in-silico results corroborate with the CD and UV–vis studies, showing perturbations in base stacking and helix organization upon Cu²⁺ addition. The hyperchromic effect observed in UV absorption and the decrease in CD band intensities are consistent with the predicted base unstacking and helix distortion induced by Cu²⁺ binding.

Similarly, the docking-based prediction that Cu²⁺ competes for base-binding sites within the DNA duplex supports the fluorescence quenching of the DNA–EB complex. Structurally, Cu²⁺ binding is predicted to cause local helix distortion in the major groove of the CCG tract, leading to destabilization of G–C base pairing. Together, the docking and spectroscopic data provide a coherent mechanistic model in which Cu²⁺ binding to guanine-rich CCG repeats disrupts DNA secondary structure, potentially contributing to genomic instability in CCG-expanded regions.

The docking-derived interaction energies and binding modes of Cu²⁺ with CCG repeat DNA were further validated by comparison with experimentally resolved Cu²⁺–Z-DNA structures, reported computational studies³⁶ (Table 3). The agreement in coordination sites and relative binding strengths supports the reliability of the present docking protocol. The predicted Cu²⁺ coordination sites and interaction energies are closely interrelated with previously reported computational studies on metal DNA interactions, which consistently identify guanine N7, cytosine O2, and phosphate oxygen atoms as primary metal-binding sites. The similarity in binding modes and energy trends further supports the robustness of the present docking results. These findings highlight the functional importance of Cu as a structural modulator in DNA, particularly in CCG-rich sequences, where metal interactions can influence DNA stability and conformation.

Table 3.

Comparison of Cu²⁺–DNA docking results with reported experimental/computational affinities.

Ligand / metal	DNA with PDB id	Method (Literature)	Reported affinity / energy	This study	Key experimental / computational findings	Comparison with present study
Cu²⁺	Z-DNA d(CGCGCG)₂ (CCG-rich) (4XSN)	X-ray crystallography	−20 to −30 kcal·mol⁻¹	−21 to −29 kcal·mol⁻¹	Cu²⁺ directly coordinates guanine N7 and phosphate oxygen atoms, stabilizing Z-DNA conformation (PMID: 1939078)	Docking predicts identical coordination sites (G-N7, OP2) and similar binding geometry
Cu²⁺	GC- and CCG-rich DNA	Spectroscopic & biochemical studies			Cu²⁺ shows higher affinity for guanine and cytosine donor atoms (N7, O6, N3, O2) compared to Mg²⁺ (PMID: 39488948)	Strong interaction energies and repeated involvement of G and C atoms observed in docking

Discussion

The double-stranded Watson-Crick B-DNA is the principal genetic molecule and a gene-regulating element in biological systems. Any changes in B-DNA conformation will alter the properties and function of DNA that may have applications in the control of gene expression. Our results confirmed that Cu interacts with CCG sequences and induces changes in the conformation and stability of DNA. The altered conformation in the CCG repeat sequence will influence the transcription, leading to abnormalities in the gene expression that may have a role in genome biology/genetic instability in neurodegeneration.^37–43,44 Further investigations are needed to understand the genomic changes in neurodegenerative diseases and possibly to develop them as future drug targets.

Figure 6 illustrates a mechanistic framework describing how copper ions (Cu²⁺) interact with trinucleotide (CCG)₁₂ DNA repeats to drive genomic instability and contribute to neuronal dysfunction associated with FXTAS and the associated Parkinsonism disorders. Trinucleotide repeat–rich regions are intrinsically prone to forming non-canonical DNA structures, and their vulnerability is further exacerbated by metal ion dysregulation, a hallmark observed in several neurodegenerative disorders, including Parkinson's disease. This model integrates metal–DNA chemistry with replication stress, transcriptional interference, and neurodegenerative pathology.

Figure 6.

Proposed mechanistic model illustrating the interaction of Cu²⁺ with (CCG)₁₂ DNA repeats. Copper ions coordinate with guanine residues, inducing a transition from canonical B-DNA to distorted DNA conformations. These structural alterations stall replication and transcription machinery, promote R-loop formation, and cause DNA strand breaks and repeat instability. The resulting genomic instability contributes to neuronal dysfunction and Parkinsonism-associated neuropathology.

In the proposed mechanism, Cu²⁺ ions preferentially coordinate with guanine bases within (CCG)₁₂ repeats. Guanine-rich sequences provide high-affinity binding sites for divalent metal ions due to the presence of N7 and O6 atoms, facilitating stable Cu²⁺–DNA complexes. This coordination perturbs normal hydrogen bonding and base stacking interactions, driving a conformational transition from canonical B-DNA to distorted or alternative DNA structures. Such structural deviations include localized DNA bending, unwinding, or the formation of secondary structures that are incompatible with the smooth progression of DNA metabolic processes. These Cu²⁺-induced DNA conformational changes impose a significant barrier to replication and transcription machineries. During DNA replication, and polymerase stalling occurs at distorted repeat regions, leading to the generation of single- and double-strand DNA breaks. Similarly, during transcription, RNA polymerase encounters impediments at these altered DNA templates, leading to alterations in transcription. The coexistence of modulated replication and transcription complexes within the same genomic loci increases the probability of transcription–replication conflicts, leading to genomic instability.

A key downstream consequence highlighted in this model is the promotion of R-loop formation. Stalled transcription favors the re-annealing of nascent RNA transcripts to the complementary DNA strand, resulting in stable RNA:DNA hybrids and displacement of the non-template DNA strand. (CCG)₁₂ repeats are particularly susceptible to R-loop accumulation due to their GC-rich nature. Persistent R-loops act as physical obstacles to replication fork progression and serve as substrates for nucleases, thereby amplifying DNA damage and repeat instability.

The accumulation of DNA strand breaks and repeat length variability ultimately activates DNA damage response pathways. In neurons, which possess limited regenerative capacity and rely heavily on genomic integrity for long-term survival, chronic activation of these pathways can be especially detrimental. Unresolved DNA damage and repeat instability disrupt gene expression programs critical for neuronal maintenance, synaptic function, and mitochondrial homeostasis. Over time, this genomic stress contributes to neuronal dysfunction, the selective vulnerability of dopaminergic neurons, and the development of Parkinson's disease-associated neuropathology.

Collectively, Figure 6 highlights a unifying mechanism that links copper dyshomeostasis, trinucleotide repeat instability, and neurodegeneration. By demonstrating how Cu²⁺–(CCG)₁₂ DNA interactions can initiate a cascade of structural DNA alterations, replication and transcription stress, R-loop accumulation, and DNA damage, this model provides mechanistic insight into how environmental and metabolic factors converge on genomic instability pathways in neurodegeneration.

Footnotes

Acknowledgements

The authors thank the heads of IISC-Bangalore and KLEF-Vijayawada for facilities and support.

ORCID iDs

Lakshmi Sowmya Emani

Jayanth K Rao

Jagadeesha Kumar Dasappa

Priya Narayan

Govindaraju Mullur

Pavani Avula

Jagannatha Rao Kosagisharaf

Ethical considerations

Not applicable

Consent to participate

Not applicable

Consent for publication

Not applicable

Author contribution(s)

Lakshmi Sowmya Emani: Conceptualization; Data curation; Formal analysis; Formal analysis; Investigation; Methodology; Project administration; Supervision; Validation; Visualization; Writing – original draft; Writing – review & editing.

Jayanth K. Rao: Conceptualization; Data curation; Formal analysis; Formal analysis; Supervision; Validation; Visualization; Writing – original draft; Writing – review & editing.

Jagadeesha Kumar Dasappa: Data curation; Formal analysis; Investigation; Methodology; Software; Supervision; Validation; Visualization; Writing – original draft; Writing – review & editing.

Priya Narayan: Data curation; Formal analysis; Investigation; Methodology; Software; Supervision; Validation; Visualization; Writing – original draft; Writing – review & editing.

Govindaraju Mullur: Investigation; Methodology; Supervision; Validation; Visualization; Writing – original draft; Writing – review & editing.

Pavani Avula: Data curation; Software; Validation; Visualization; Writing – original draft; Writing – review & editing.

Jagannatha Rao Kosagisharaf: Conceptualization; Data curation; Formal analysis; Formal analysis; Investigation; Methodology; Project administration; Resources; Software; Supervision; Validation; Visualization; Writing – original draft; Writing – review & editing.

Funding

The authors would like to thank the UGC, DST, and DBT, Government of India, for their financial support to the Molecular Biophysics Unit, IISc, Bangalore. LSE is thankful to KLEF for financial support through a doctoral fellowship (KLEF-02).

DST, india, KLEF, (grant number Instrument Grant, KLEF-02).

Declaration of conflicting interests

The authors declared no potential conflicts of interest with respect to the research, authorship, and/or publication of this article.

Data availability statement

All data generated or analyzed during this study are included in this published article.

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Studies on copper (II) interaction with the (CCG) 12 repeats sequence: An insight into genomic instability in neurodegeneration

Abstract

Background

Objective

Methods

Results

Conclusions

Keywords

Introduction

Methods

Synthesis of (CCG)12 repeats

Preparation of DNA stock solution

Preparation of copper chloride stock solution

Ethidium bromide stock solution

Circular dichroism studies

UV/VIS absorption studies

Fluorescence studies

In silico molecular docking studies

Results

Circular dichroism studies

UV absorption studies

Fluorescence studies

Effect of temperature and pH on the (CCG) sequence conformation

Docking studies

Discussion

Footnotes

Acknowledgements

ORCID iDs

Ethical considerations

Consent to participate

Consent for publication

Author contribution(s)

Funding

Declaration of conflicting interests

Data availability statement

References

Synthesis of (CCG)₁₂ repeats