Bacterial Resistance and Prostate Cancer Susceptibility Toward Metal

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Cite This: ACS Appl. Mater. Interfaces XXXX, XXX, XXX−XXX

Bacterial Resistance and Prostate Cancer Susceptibility Toward Metal-Ion-doped DNA Complexes Srivithya Vellampatti,† Gopalakrishnan Chandrasekaran,‡ Sekhar Babu Mitta,† Sreekantha Reddy Dugasani,† Vinoth-Kumar Lakshmanan,*,‡,§ and Sung Ha Park*,† †

ACS Appl. Mater. Interfaces Downloaded from pubs.acs.org by UNIV OF SOUTH DAKOTA on 12/16/18. For personal use only.

Sungkyunkwan Advanced Institute of Nanotechnology (SAINT) and Department of Physics, Sungkyunkwan University, Suwon 16419, Korea ‡ Department of Biomedical Sciences, Chonnam National University Medical School, Gwangju 61469, Korea § Department of Biomedical Engineering, Sri Shakthi Institute of Engineering and Technology, Coimbatore 641062, India S Supporting Information *

ABSTRACT: DNA nanotechnology has laid a platform to construct a variety of customshaped nanoscale objects for functionalization of specific target materials to achieve programmability and molecular recognition. Herein, we prepared DNA nanostructures [namely, synthetic DNA rings (RDNA) and DNA duplexes extracted from salmon (SDNA)] containing metal ions (M2+) such as Cu2+, Ni2+, and Zn2+ as payloads for delivery to exterminate highly pathologic hospital bacterial strains (e.g., Escherichia coli and Bacillus subtilis) and prostate cancer cells (i.e., PC3, LNCaP, TRAMP-C1, 22Rv1, and DU145). Morphologies of these M2+-doped RDNA were visualized using atomic force microscopy. Interactions between M2+ and DNA were studied using UV−vis and Fourier transform infrared spectroscopy. Quantitative composition and chemical changes in DNA without or with M2+ were obtained using X-ray photoelectron spectroscopy. In addition, M2+-doped DNA complexes were subjected to antibacterial activity studies. They showed no bacteriostatic or bactericidal effects on bacterial strains used. Finally, in vitro cellular toxicity study was conducted to evaluate the effect of pristine DNA and M2+-doped DNA complexes on prostate cancer cells. Cytotoxicities conferred by M2+-doped DNA complexes for most cell lines were significantly higher than those of M2+ without DNA. Cellular uptake of these complexes was confirmed by fluorescence microscopy using PhenGreen FL indicator. On the basis of our observations, DNA nanostructures can be used as safe and efficient nanocarriers for delivery of therapeutics. They have enhanced therapeutic window than bare metals. KEYWORDS: DNA, metal-ion-doped DNA, spectroscopy, bacterial resistance, prostate cancer

1. INTRODUCTION Although metals are used extensively in physical applications, they are inextricably quintessential components of various chemical and biomolecular processes and therapy. Metals have exceptional properties such as charge, redox activity, and ability to interact with the ligands that impart utility to biology.1,2 Especially, transition-metal ions enact a major role in the development of new metal-based drugs due to their diverse coordination numbers, net charges, and redox potentials that qualify them as free radicals with immense contribution to medicinal therapeutics. Metal ions are very reactive under normal conditions. They might be essential or noxious. Therefore, appropriate intracellular metal-ion concentration must be maintained to retard any pathological disorders. Metal ions can bind some ligands to study their pragmatic knowledge of interactions. DNA is a self-assembling biopolymer built from phosphate backbone, deoxyribose sugar, and bases. It is a central molecule indulged in conservation, expression, and transmission of genetic information. Apart from its role as a cellular repository, DNA has attracted usages not only in chemistry and biology but © XXXX American Chemical Society

also in physics, material sciences, and nanotechnology due to its molecular recognition and spontaneous self-assembly driven by hybridization of complementary base pairs.3−5 Owing to drawbacks observed in sequestering metal ions to cancer cells, DNA nanotechnology has triggered an impetus for the advancement of metal-based drugs. This has grown into a main field of interest worldwide without freaking out the public. Although various drug carriers made of inorganic substances (e.g., mesoporous silica, quantum dots, and carbon nanotubes) have been reported, organic DNA molecules provide unique advantages as drug carriers due to their biocompatibility and sequence specificity.6−14 Positively charged metal ions can bind to electron donor atoms of hydrogen-bonded DNA bases through intercalation. They can also bind to negatively charged phosphate backbone through electrostatic interaction.15,16 Divalent metal ion (M2+)doped DNA duplexes and nanostructures are gradually being Received: October 14, 2018 Accepted: December 4, 2018

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DOI: 10.1021/acsami.8b17013 ACS Appl. Mater. Interfaces XXXX, XXX, XXX−XXX

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Figure 1. Experimental procedures showing the synthesis of ring DNA (RDNA) and salmon DNA (SDNA) modified with divalent metal ions (M2+) such as Cu2+, Ni2+, and Zn2+. (a) Free solution annealing procedure for RDNA followed by modification with metal ions (M-RDNA). (b) M2+ modification on SDNA duplexes (M-SDNA). The zoomed-in DNA model shows plausible coordinations of M2+ to the DNA duplex (phosphate backbone by electrostatic interaction and the place between base pairs via intercalation).

Figure 2. AFM images of Cu2+-, Ni2+-, and Zn2+-doped Ring DNA (marked as Cu-RDNA, Ni-RDNA, and Zn-RDNA, respectively). (a−d) AFM images of RDNAs with fixed concentration of 500 nM without M2+ or with [Cu2+] of 4 mM, [Ni2+] of 2 mM, and [Zn2+] of 1 mM, respectively. The ring morphology remains unchanged even after the addition of M2+. Scan size of all images is 1 × 1 μm2.

available DNA duplexes extracted from salmon fish (SDNA) to deliver M2+ as an active drug. Owing to their significance in biological and medicinal applications such as altering the nature of cancer cells and producing reactive oxygen species,30−34 we used distinct M2+ (i.e., Cu2+, Ni2+, and Zn2+) with carefully chosen concentrations to enhance their function. Structural

used in electronics, optics, photonics, and magnetism.17−29 However, utilization of DNA as a carrier for functionalized materials (especially M2+ as a potent anticancer drug) has not been discussed intensively. To shed a light on this, we proposed vehicles made of sequence-designed synthetic ring DNA (RDNA) and naturally B


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Figure 3. Spectral measurements of pristine SDNA and M2+-doped SDNA (M-SDNA) at room temperature. (a) UV−vis absorption spectra of pristine SDNA (0.1 wt %) and M-SDNA at [Cu2+] of 4 mM, [Ni2+] of 2 mM, and [Zn2+] of 1 mM. (b) Attenuated total reflectance FT-IR spectra of pristine SDNA and M-SDNA in the range of 1800−800 cm−1. Hypochromism was observed for Ni- and Zn-SDNA in the whole range of wavenumbers. For clarity, spectrum of pristine SDNA was highlighted in gray.

stability of RDNA with the addition of M2+ was examined by atomic force microscopy (AFM). Binding of M2+ to DNA and element mapping of M2+-doped DNA was tested by ultraviolet− visible (UV−vis) and Fourier transform infrared (FT-IR) spectroscopies and X-ray photoelectron spectroscopy (XPS). To address the effect of M2+-doped DNA on prognosis, we tested its antibacterial activity against Escherichia coli (E. coli) and Bacillus subtilis (B. subtilis) species through live−dead assay and human prostate cancer cell lines (e.g., PC3, LNCaP, TRAMP-C1, 22Rv1, and DU145) through 3-(4,5-dimethylthiazol-2-yl)-5(3-carboxymethonyphenol)-2-(4-sulfophenyl)-2Htetrazolium (MTS) assay. Cellular uptake of M2+-doped DNA complexes by PC3 cells was studied using PhenGreen FL by fluorescence microscopy.

remained unchanged. Inner and outer designed diameters of a RDNA were 13 and 29 nm, respectively. These values were in good agreement with experimental observation shown in Figure 2.42,43 To understand the binding of M2+ to DNA, UV−vis spectroscopy was performed for both pristine and M2+-doped DNA complexes. Results are shown in Figure 3a. UV−vis absorption spectra for RDNA and SDNA with various concentrations of Cu2+, Ni2+, and Zn2+ are shown in Figures S2 and S3 in Supporting Information (SI). Absorption peak at 260 nm in pristine DNA and M2+-doped DNA was clearly observable. Hypochromism (i.e., decrease in absorption peak intensity) was observed for bands at 260 nm, indicating intercalation of M2+ to nitrogen atoms of heterocyclic bases of DNA that exhibited a tolerance of DNA duplexes up to a critical [M2+]. Although Ni2+- and Zn2+-doped DNA complexes did not show any of their characteristic peaks or significant peak shifts in the measured wavelength range, characteristic peak position of Cu2+ was observed at ∼730 nm. Absorption peak position of the DNA also shifted from 260 to 275 nm for Cu2+-doped DNA complexes.44 Attenuated total reflectance FT-IR spectra of pristine SDNA and M-SDNA were obtained to probe the interaction and binding pattern of M2+ to SDNA (Figure 3b). When excited by IR radiation, specific wavelengths are absorbed, causing chemical bonds to undergo vibrations such as stretching and bending in the wavelength range of 800−4000 cm−1, giving rise to molecular fingerprint of the sample at molecular level. Noticeable changes of vibrational frequencies and peak intensities of M-SDNA compared to pristine SDNA were observed to realize the interaction of M2+ with SDNA. Peak assignments for pristine SDNA are summarized in Table S3 in the SI. Bands in 800−1000 and 1000−1300 cm−1 ranges were sensitive to sugar and phosphate backbone vibrations, respectively. These were reliable markers for various sugar puckering modes (N- and S-types). Bands in 1300−1800 cm−1 range were due to stretching vibrations of double bonds in DNA bases.45,46 In case of Cu2+, their binding was mainly with PO2− backbone (800−1300 cm−1), in which peaks were either decreased in intensity (858, 923 cm−1), shifted to lower wavelength (1050, 1222 cm−1), or disappeared (1083 cm−1). Such an observed increase in intensity was due to binding of M2+

2. RESULTS AND DISCUSSION Experimental procedures used to synthesize RDNA and SDNA doped with M2+ such as Cu2+, Ni2+, and Zn2+ are shown in Figure 1. Equimolar concentrations of RDNA strands and appropriate amount of SDNA was formed by annealing from 95 °C to room temperature and stirring of SDNA fibers overnight, respectively, followed by M2+ doping into DNA. Binding of M2+ to DNA can take place either at phosphate groups, sugar moieties, or various donor sites in nitrogenous bases. Two modes of interaction between M2+ and DNA (i.e., noncovalent and coordinate bonding) exist in M2+-doped DNA. Noncovalent bonding occurs through electrostatic attraction between M2+ and negatively charged phosphate backbone, whereas coordinate bonding occurs through intercalation between M2+ and DNA bases.14,15,35−39 Cu2+-, Ni2+-, and Zn2+-doped RDNA (labeled as Cu-RDNA, Ni-RDNA, and Zn-RDNA, respectively) were constructed and their morphologies were visualized by AFM. To obtain the maximum effect of M2+ without structure deformation of RDNA, concentrations of Cu2+ ([Cu2+]) at 4 mM, Ni2+ ([Ni2+]) at 2 mM, and Zn2+ ([Zn2+]) at 1 mM for a given RDNA concentration ([RDNA]) at 500 nM were carefully chosen. We named them critical concentration of a given M2+. Beyond the critical concentration, RDNA started deforming due to stress induced by excess M2+ in the DNA duplex as reported previously.40,41 RDNA with critical concentrations of Cu2+, Ni2+, and Zn2+ were well-formed and their morphologies C


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Figure 4. XPS analysis of M2+-doped SDNA (M-SDNA). (a) XPS survey spectra of pristine SDNA and M-SDNA samples labeled as SDNA, CuSDNA, Ni-SDNA, and Zn-SDNA. Core elements (i.e., Na, C, N, P, and O) present in the DNA and peaks produced by doped M2+ (i.e., Cu2+, Ni2+, and Zn2+) are indicated with dotted lines and dotted circles, respectively. (b−f) Deconvoluted spectra of Na 1s, C 1s, N 1s, P 2p, and O 1s core elements in pristine SDNA and M-SDNA samples designated with their corresponding binding energy peak positions. (g−i) Deconvoluted high-resolution core spectra and corresponding satellite (Sat) peaks of Cu 2p, Ni 2p, and Zn 2p in Cu-SDNA, Ni-SDNA, and Zn-SDNA, respectively, labeled with their peak positions. (j) Atomic concentration percentages of core elements and specific M2+ compositions for pristine SDNA and M-SDNA samples.

to DNA bases and PO2− sites.47−49 Ni2+- and Zn2+-doped DNA complexes exhibited minor changes in peak intensities due to strong base stacking and weak metal-base binding. The DNA marker band at 964 cm−1 corresponded to CC stretching of the DNA backbone. This was also present in M-SDNA.

XPS measurement was conducted to analyze chemical compositions of M-SDNA complexes. Results are shown in Figure 4. Figure 4a shows the survey spectra of pristine SDNA and distinct M2+-doped SDNA complexes acquired in binding energy (BE) range of 0−1450 eV. The apparent binding energy D


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Figure 5. Bacterial growth curves of Escherichia coli (E. coli) and Bacillus subtilis (B. subtilis) after treatment with pristine R(S)DNA, M2+, and MR(S)DNA. (a, b) Growth pattern of E. coli treated with R(S)DNA, M2+, and M-R(S)DNA samples for 20 h. (c, d) Growth pattern of B. subtilis after treatment with R(S)DNA, M2+, and M-R(S)DNA for 20 h. The absorbance of optical density (OD) was measured at 600 nm. Each point in the curve represents the mean of three replicate measurements.

of oxygen Auger peak with an X-ray occurs at ∼980 eV, which can be seen in XPS survey spectra. Necessary elemental (Na, C, N, P, and O) peaks recorded were signified with dotted lines, whereas characteristic metal peaks of Cu2+, Ni2+, and Zn2+ were marked with dotted circles in the survey spectra of Cu-SDNA, Ni-SDNA, and Zn-SDNA. Deconvoluted high-resolution XPS spectra of Na 1s, C 1s, N 1s, P 2p, and O 1s in pristine SDNA and M-SDNA are illustrated in Figure 4b−f. Individual XPS spectra of Cu 2p, Ni 2p, and Zn 2p in Cu-SDNA, Ni-SDNA, and ZnSDNA are displayed (confirming the presence of M2+ in SDNA) in Figure 4g−i. Corresponding peak position BEs of each element were also presented to comprehend shifts in BE for MSDNA compared to SDNA. As seen from Figure 4b−f, peak position BEs (either upshifted or downshifted) and peak intensities (increased or decreased) of core elements in MSDNA respect to SDNA were clearly visible, providing clues for the formation of M-SDNA complexes. M2+ possessed high affinity of bindings at phosphate backbones and between bases through electrostatic binding and chemical intercalations, respectively.27,40,41,50,51 As shown in Figure 4d,e, SDNA had a doublet N 1s peak BEs at 398.78 and 397.63 eV. Upon addition of Cu2+, Ni2+, and Zn2+ into SDNA, Cu-SDNA BE peaks were upshifted to 406.53 and 399.48 eV, a broad singlet peak for Ni-SDNA was upshifted to 399.63 eV, whereas Zn-SDNA BE peaks were downshifted to 397.18, 394.13, and 392.93 eV. These results verified that M2+ could bind to nitrogenous bases via chemical intercalation. Besides, as

observed in deconvoluted spectra of P 2p shown in Figure 4e, SDNA had a phosphate peak BE at 131.73 eV. Similarly, central peaks for Cu-SDNA, Ni-SDNA, and Zn-SDNA complexes were detected at 133.18 (upshifted), 133.33 (upshifted), and 127.23 eV (downshifted), respectively, with decrement in peak intensities compared to SDNA. Therefore, it could be concluded that M2+ can bind to the phosphate backbone through electrostatic binding to SDNA. Finally, quantitative analysis for elemental composition of SDNA, and various M-SDNA in atomic concentrations shown in Figure 4j validated the existence of M2+ in SDNA (Figure 4g−i). M2+ in DNA could be used in important applications such as antibacterial and antiprostate cancer therapy. This will be discussed below. Figure 5 shows the bacterial growth curves of E. coli and B. subtilis upon treatment with pristine RDNA (and pristine SDNA), M2+, and M-RDNA (and M-SDNA). Antibacterial activities of M-DNA complexes against both Gram-positive (B. subtilis) and Gram-negative (E. coli) bacteria were studied. From these growth curves, it could be seen that neither the M-DNA complexes nor individual M2+ exhibited bacteriostatic or bactericidal effect on E. coli or B. subtilis strains. To further confirm that they showed no antibacterial activities, we performed live−dead cell viability assays for these bacterial cells treated with M-DNA. Live−dead bacterial viabilities of E. coli with M-RDNA and M-SDNA are shown in Figures 6 and 7, respectively. Cells with intact membranes will take up SYTO9 dye, which will stain E


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Figure 6. Live−dead bacterial assay of E. coli cells treated with M-RDNA complexes and visualized with fluorescent microscope. Nucleic acids of live bacterial cells with intact membranes were stained with SYTO9 (green, wavelength of 528 nm), whereas cells with damaged membranes (red, sometimes pale yellow during stain overlap) were stained with propidium iodide (red, 645 nm) only. Untreated and Triton X-100 treated cells were considered as positive and negative controls, respectively. Population of damaged cells (marked with the arrows) treated with M-RDNA was relatively less than that in the negative control sample.

nucleic acids to give bright green fluorescence. In contrast, propidium iodide (PI) stains nucleic acids of cells with damaged membranes. As a result, dead cells were stained bright red or pale yellow (in case of partially damaged membrane), whereas live intact cells were stained bright green. These cell-stained images suggested that the number of dead cells or cells with damaged membranes (red or pale yellow, spotted by arrow marks) was very low compared to that of live cells with intact membranes (green). These results of live−dead bacterial cell viabilities confirmed that M-DNA complexes (i.e., M-RDNA and MSDNA) had no antibacterial effect on Gram-positive or Gramnegative bacteria, in accordance with the results of bacterial growth curves shown in Figure 5. This can be attributed to the fact that bacteria possess adaptable mechanisms to efficiently metabolize M2+ from the surrounding environment, as reported elsewhere.52−57 To observe the effect of M-DNA complexes on viabilities of cancerous cells, five different prostate cancer cell lines (PC3, LNCaP, TRAMP-C1, 22Rv1, and DU145) of various etiologies

were treated with M-RDNA and M-SDNA. Results are shown in Figures 8 and 9. The overall toxicities conferred by M-DNA complexes for most cell lines (except LNCaP) were enhanced significantly compared to those of M2+ (i.e., Cu2+, Ni2+, and Zn2+) without DNA. Although cell viability data were collected twice (after 24 and 48 h), cell viabilities of M-DNA complexes were not significantly changed (i.e., differences of less than 6%) with respect to time. Interestingly, Cu− and Ni−DNA complexes showed much higher toxicities (>90%) to prostate cancer cells compared to controls (i.e., positive controls (PC) and negative controls (NC)), whereas Zn−DNA complexes exhibited 40−60% cytotoxicity. Due to delivery of M2+ by DNA (served as a biocompatible nanocarrier), cancerous cell might have malfunctioned by binding of M2+ to specific proteins. Reactive oxygen species (hydroxyl radical) produced by excess of M2+ might have been delivered into cells by DNA. This might be the main cause of DNA damage in cancer cells.33,34 Cellular uptake behavior of M-RDNA complexes in PC3 cells was studied. Figure 10 shows fluorescent images of cellular F


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Figure 7. Live−dead bacterial assay of E. coli cells treated with M-SDNA complexes and visualized with a fluorescent microscope. Nucleic acids of live bacterial cells with intact membranes were stained with SYTO9 (green), whereas cells with damaged membranes (red, sometimes pale yellow during stain overlap) were stained with propidium iodide (red) only. Untreated and Triton X-100 treated cells were considered as positive and negative controls, respectively. Population of damaged cells (marked with the arrows) treated with M-SDNA was much less than that in the negative control sample.

uptake of M-RDNA complexes using PhenGreen FL as an indicator to trace M2+. M-RDNA entered cells through endocytosis followed by the release of M2+ via pH control inside cells. M2+ was then transferred to secretory pathways by chaperone proteins present inside cells. M2+ was traced by PhenGreen FL dye, which gave a bright green color in fluorescence images (2nd column in Figure 10). Ni2+ in RDNA was found to be up-taken more effectively than Cu2+ or Zn2+ in RDNA by comparing the intensity of green fluorescence. Although degrees of cellular uptake between untreated cells and M-RDNA-treated cells were clearly distinguishable, mechanisms behind the cellular uptake of DNA with various M2+ (e.g., Cu2+, Ni2+, and Zn2+) in different cancer cells need to be studied systematically in the future.

such as UV−visible, infrared, and X-ray photoelectron spectroscopies of M2+-doped DNA (M-DNA) complexes revealed binding modes and interactions between M2+ (i.e., Cu2+, Ni2+, and Zn2+) and DNA (i.e., RDNA and SDNA). These M-DNA complexes showed no antibacterial activities against Grampositive or Gram-negative bacterial strains. Cell viability and uptake analyses revealed that M-DNA complexes conferred cytotoxicity to prostate cancer cells through effective internalization. Further studies on these complexes are needed to understand cellular trafficking with in vivo studies. Our results suggest that DNA structures can function as therapeutic nanocarriers for efficient drug delivery applications that are readily feasible and economical.

3. CONCLUSIONS We developed a novel nanocarrier system by using natural and synthetic DNA with low immunogenicity, which actively delivered payload to the site of action. Spectral measurements

High-performance liquid chromatography purified sequence-designed DNA strands (i.e., ring 1-1 and ring 1-2) at 500 nM for RDNA purchased from IDT Technologies (CA) were added to a test tube containing 1× TAE/Mg2+ buffer [40 mM Tris, 20 mM acetic acid, 1 mM ethylenediaminetetraacetic acid (EDTA) (pH 8.0), and 12.5 mM

4. EXPERIMENTAL SECTION

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Figure 8. Cell viability assessments of (a) PC3, (b) LNCaP, (c) TRAMP-C1, (d) 22Rv1, and (e) DU145 prostate cancer cells by MTS assay (CellTiter96 AQueous One Solution Cell Proliferation Assay) after treatment with pristine RDNA, M2+, and M-RDNA samples for 24 and 48 h, respectively. Vertical axis indicates cell viabilities of testing samples (i.e., prostate cancer cells treated with pristine RDNA, M2+, and M-RDNA). Positive control (untreated, 100% viable) and negative control (treated with Triton X-100, 7−15% viable) samples are labeled as PC and NC, respectively. Although M2+ without RDNA revealed reduction, M-RDNA showed prominent effect in reducing prostate cancer cell viability. Samples were tested in triplicates.

Figure 9. Cell viability assessments of (a) PC3, (b) LNCaP, (c) TRAMP-C1, (d) 22Rv1, and (e) DU145 prostate cancer cells treated with pristine SDNA, M2+, and M-SDNA samples for 24 and 48 h, respectively. Vertical axis indicates cell viabilities of testing samples (prostate cancer cells treated with pristine SDNA, M2+, and M-SDNA). Positive (untreated, 100% viable) and negative control (treated with Triton X-100, 7−15% viable) samples are labeled as PC and NC, respectively. Samples were tested in triplicates. magnesium acetate]. Hybridization of RDNA was performed by placing the test tube in a Styrofoam box containing 2 L of boiling water. It was allowed to turn cold from 95 °C to room temperature for 24 h. Annealed RDNA structures were incubated at 4 °C for structure

stabilization (Figures 1a and S1, Tables S1 and S2 in the Supporting Information (SI)). SDNA solution was prepared by dissolving 0.05 g of commercially available DNA fibers extracted from salmon fish (GEM Corporation, Shiga, Japan) in 10 mL of deionized (DI) water and stirred overnight at H


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Figure 10. Cellular uptake of M-RDNA complexes in PC3 prostate cancer cells using PhenGreen FL as an indicator (tracer dye) for bare M2+ (i.e., Cu2+, Ni2+, and Zn2+) detection visualized with differential interference contrast and fluorescent microscope. Cells untreated and treated with RDNAs (having a fixed concentration of 500 nM) with [Cu2+] of 4 mM, [Ni2+] of 2 mM, and [Zn2+] of 1 mM were examined. Cu-RDNA, Ni-RDNA, and ZnRDNA showed appreciable uptakes into cells compared to controls. 1000 rpm. Concentration of SDNA was found to be 1.0 wt %. For characterization, the SDNA solution was diluted to 0.1 wt % (Figure 1b). Cu2+, Ni2+, and Zn2+ solutions obtained from Cu(NO3)2, NiCl2, and ZnCl2, respectively, were added to test tubes containing the annealed RDNA in 1× TAE/Mg2+ buffer and dispersed SDNA in DI water followed by 24 h of incubation at room temperature. Final concentrations of Cu2+, Ni2+, and Zn2+ in both RDNA and SDNA were 4, 2, and 1 mM, respectively. Concentrations of RDNA and SDNA were fixed at 500 nM and 0.1 wt %, respectively (Figure 1). AFM imaging was executed on a Nanoscope III using NP-S oxidesharpened silicon nitride tips (Vecco, CA) in the fluid tapping mode. M2+-doped RDNA (M-RDNA) solution at 5 μL was then dropped onto mica substrate and left at room temperature for 1 min. Then, 20 μL and 5 μL of 1× TAE/Mg2+ buffer were dispensed on the mica substrate and AFM tip, respectively, before imaging (Figure 2). Ultraviolet−visible (UV−vis) spectroscopy was performed for SDNA samples with various concentrations of M2+ using Nano Drop 2000c (Thermo-Scientific, DE). Each sample at 2 μL was dispensed on the pedestal. Their absorption spectra were obtained in the range of 200−800 nm. FT-IR spectra of M2+-doped SDNA (M-SDNA) complexes placed on silicon wafer were recorded in the range of 4000−700 cm−1 using a spectrometer (TENSOR 27, Detector: MIR_ATR (ZnSe), Bruker, MA) with a resolution of 4 cm−1 and 32 scans (Figures 3, S2, and S3, and Table S3 in the SI). X-ray photoelectron spectroscopy (XPS) was performed using a spectrometer (ESCALAB 250Xi, Thermo Fisher Scientific, U.K.) equipped with a monochromatic Al Kα X-ray source at an energy of 1450 eV. X-ray spot size used here was 650 μm with a hemispherical analyzer. Surface composition and atomic-resolution structural maps of pristine SDNA and M-SDNA samples were obtained (Figure 4). Antibacterial activities of these prepared M-RDNA and M-SDNA complexes were determined using bacterial growth curve for pathological hospital strains (i.e., E. coli and B. subtilis). These bacterial strains from glycerol stocks were inoculated into Luria Broth and grown at 37 °C overnight in an orbital incubator shaker. The optical density at a wavelength of 600 nm (OD600) of the culture was measured using an

Epoch 2 Microplate Spectrophotometer (BioTek Instruments, VT). Culture with OD600 = 0.1 was used as the starter culture for bacterial experiment. For bacterial growth curve analysis, 100 μL of the culture was inoculated into a 96-well plate. The OD600 absorbance of the culture medium containing either M-RDNA or M-SDNA complexes was recorded over 20 h. Bacterial growth curve was plotted for OD600 against time. Measurement was carried out in triplicates. Error bars were added into the curve (Figure 5). For live−dead bacterial viability assay (LIVE/DEAD Bac Light Bacterial Viability Kits, Invitrogen, CA), equal volumes of SYTO9 and propidium iodide (PI) dye mixture were prepared. The dye mixture (3 μL) was then added to 1 mL of E. coli culture medium containing either M-RDNA or M-SDNA complexes followed by room-temperature incubation in the dark for 45 min. After the incubation, 2.5 μL of the bacterial suspension stained with the dye mixture was placed on a microscopy slide and covered with a cover slip. These cells were then visualized and imaged under a fluorescent microscope (EVOS FLoid Cell Imaging Station, Thermo Fisher Scientific, Korea) (Figures 6 and 7). Cellular studies were carried out using five different prostate cancer cells: PC3, LNCaP, TRAMP-C1, 22Rv1, and DU145. These cells were maintained in Roswell Park Memorial Institute (RPMI) 1640 medium supplemented with 10% fetal bovine serum and 1% antibiotic− antimycotic solution. Cells were cultured in 5% CO2 environment at 37 °C. Once these cells attained 80% confluency, they were treated with 0.25% trypsin-EDTA and centrifuged at 1200 rpm for 5 min. The cell pellet was then resuspended in a complete growth medium, counted using a hemocytometer, and used for further cell experiments. Prostate cancer cell viability was determined by 3-(4,5-dimethylthiazol-2-yl)-5(3-carboxymethonyphenol)-2-(4-sulfophenyl)-2H-tetrazolium (MTS) assay using CellTiter96 AQueous One Solution Cell Proliferation Assay Kit (Promega, WI). Five different prostate cancer cells were seeded into 96-well plates at cell density of 5000 cells/well. Final concentrations of Cu2+, Ni2+, and Zn2+ were 4, 2, and 1 mM, respectively. Concentrations of RDNA and SDNA were fixed at 500 nM and 0.1 wt %, respectively. They were prepared and stored at 4 °C. Appropriate controls such as TAE buffer, and DI water were also I


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ACS Applied Materials & Interfaces prepared for comparative analysis. After 24 h, the medium was removed and cells were washed with Dulbecco’s phosphate-buffered saline. MDNA (i.e., M-RDNA or M-SDNA) complexes (10 μL) and fresh RPMI media (90 μL) were added to each well and incubated for indicated time. Untreated cells (incubated with media alone) were considered as positive controls (PC), whereas cells treated with Triton X-100 (1%) were used as negative controls (NC). After 24 and 48 h of incubation, 20 μL of MTS solution was added into each well and incubated for 4 h to form formazan product. Cell viabilities were then obtained by measuring the OD of the culture at wavelength of 490 nm (OD490) using an iMark Microplate Reader (BioRad, CA). Cell viabilities were measured in triplicates. Average percentages with error bars were indicated (Figures 8 and 9). Cellular uptake studies of M-RDNA were carried out using PC3 cells. Briefly, PC3 cells were seeded into a confocal dish containing a cover slip (cell density: 5000 cells/cm2). For proper attachment, cells were incubated for 24 h. RPMI medium was then removed and the cells were washed twice with PBS. Fresh RPMI media along with M-RDNA complexes were added to the cells. PhenGreen FL (tracer dye for metal ions) was then added to the medium. Final concentration of PhenGreen FL was 5 μM. These cells were then incubated at 37 °C in a 5% CO2 incubator for 4 h. After the incubation, the cells were washed with PBS and fixed with 3.7% paraformaldehyde for 10 min. After washing thrice with PBS, the cells were permeabilized with 0.2% Triton X-100 for 5 min. These cells were again washed with PBS followed by addition of 4′,6-diamidino-2-phenylindole (300 nM) solution with 5 min of incubation. Finally, these cells were washed with PBS and imaged under a fluorescence microscope (EVOS FLoid Cell Imaging Station, Thermo Fisher Scientific, Korea) to study the uptake of M-RDNA complexes (Figure 10).

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necessary facilities. This work was supported by a grant from Chonnam National University Hospital (CRI17032-1 to V.K.L.).

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ABBREVIATIONS RDNA, DNA rings SDNA, salmon DNA AFM, atomic force microscope UV−vis, ultraviolet−visible FT-IR, Fourier transform infrared XPS, X-ray photoelectron spectroscopy BE, binding energies E. coli, Escherichia coli B. subtilis, Bacillus subtilis PI, propidium iodide MTS, 3-(4,5-dimethylthiazol-2-yl)-5(3-carboxymethonyphenol)-2-(4-sulfophenyl)-2H-tetrazolium DI, deionized TAE, Tris base, acetic acid and EDTA PC, positive controls NC, negative controls OD, optical density

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ASSOCIATED CONTENT

S Supporting Information *

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acsami.8b17013. The sequence and a schematic of ring DNA, UV absorption spectra of ring DNA and salmon DNA, and FT-IR absorption bands obtained from salmon DNA (PDF)

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REFERENCES

AUTHOR INFORMATION

Corresponding Authors

*E-mail: [email protected] (V.-K.L.). *E-mail: [email protected] (S.H.P.). ORCID

Sreekantha Reddy Dugasani: 0000-0001-5805-8713 Sung Ha Park: 0000-0002-0256-3363 Author Contributions

S.V. initiated the project, designed, and performed experiments, and analyzed data and wrote the paper. G.C., S.B.M., and S.R.D. analyzed data and wrote the paper. V.-K.L. and S.H.P. initiated and directed the project, designed experiments, analyzed data and wrote the paper. The manuscript was written through contributions of all the authors. All the authors have given approval to the final version of the manuscript. Funding

This work was supported by the National Research Foundation (NRF) of Korea (2018R1A2B6008094). Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS G.C. and V.-K.L. thank Chonnam National University and Chonnam National University Medical School for providing J


Research Article

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