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Abstract
Biomaterials based on deoxyribonucleic acid (DNA) have shown notable potential in optoelectronic and photonic devices. In order to further investigate the optical properties of a DNA-based lipid complex such as DNA-cetyltrimethylammonium (CTMA), which is widely used in current DNA thin film research, a new refinement process was developed to minimize the relative bound water content and control binding of CTMA onto the DNA backbone. The water contents and CTMA binding in the DNA-CTMA precipitates were identified by spectrometric measurements to quantify effects of our refinement process. Dissolving these refined DNA-CTMAs in organic solvents, thin solid films were deposited on Si and quartz substrate using the spin coating process. Their refractive indices and absorbance were measured to quantitatively assess the impact of our refinement process on the optical properties of the DNA-CTMA films. In addition, thermo-optic coefficients, dn/dT, were also measured in a temperature range from 30 to 100°C to observe differences among refined DNA-CTMAs. Detailed quantitative spectroscopic analyses and optical measurements are reported.
© 2017 Optical Society of America
1. Introduction
Deoxyribonucleic acid (DNA) is a basic building-block of living organisms and has been the fundamental basis in life-science and bio-technology research [1], which has enabled customized DNA sequence control and adding functional material at the level of the constituent units [2]. In addition, a new type of 2, 3-dimensional nano-structures have been realized controlling the neighboring DNA nucleobases [3]. Recently DNA has been further employed in macroscopic thin solid film technology, which has opened a new possibility in bio-compatible electro-optics and photonics such as optical amplifier based on dye-doped DNA optical waveguides [4], mode-locking of fiber laser using high optical nonlinearities [5], and efficiency enhancement in organic light emitting diode (OLED) [6], to name a few. DNA has been also investigated for optical memories, optical switches, and optical sensors that could replace conventional optical polymer materials [7–10].
However, in order to further apply DNA to practical photonic devices, optical properties of DNA thin solid films should be flexibly controlled in terms of the refractive index, optical loss, and their dependence on temperature. Conventional optical polymers [11] such as Poly methyl methacrylate (PMMA), and poly carbonate (PC) are technologically matured so that optical property control is well established, yet methods to systematically control optical properties of DNA thin solid films have not been fully investigated despite a high potential in bio-compatible photonics applications.
In recent studies, salmon testis DNA (stDNA) [12] has been widely used in DNA thin film development and various cationic lipid, or surfactants, have been dissolved with stDNA in aqueous solution to obtain DNA-lipid precipitates [13,14]. These precipitates have been dissolved in organic solvents for conventional spin-coating process to make thin solid films. Cetyltrimethylammonium chloride (CTMA-Cl) has been one of most well-known cationic lipid for forming DNA-CTMA precipitates in aqueous solutions. DNA-CTMA thin films have been fabricated and their basic optical properties have been reported [15–17]. In an attempt to remove residual CTMA, which is not tightly bound to DNA, rinsing of DNA-CTMA precipitates by applying Soxhlet-dialysis has been reported [18]. However, this rinsing alone cannot alter the fundamental precipitation process itself, which would significantly affect material properties of DNA-CTMA thin solid films. Detailed control of precipitation conditions and post-processing for the precipitates have not been systematically investigated yet, which could explain why there exist wide ranges of variation among optical properties in DNA-CTMA thin films in prior reports [19,20].
In the process of DNA-CTMA precipitation in aqueous solution, the amount of cationic lipid binding can vary in a wide range, and insufficient lipid binding could result in an excessive content of water, as schematically depicted in Fig. 1(a). We paid keen attention to this water content, or equivalently the amount of cation binding, in DNA-CTMA precipitates, and we developed post-processes to systematically control their amount and subsequently optical properties of DNA-CTMA thin solid films, for the first time to the best knowledge of the authors.
Fig. 1 (a) Schematic illustration of DNA-CTMA complex structure. Double helix represents DNA and CTMAs are represented as line segments. Before our proposed re-precipitation process, shown in the left, CTMAs are not fully bound to phosphate backbone of DNA to result in defects as shown in orange dots allowing water binding. After re-precipitation, binding of CTMAs to DNA is significantly increased to drive out residual water. (b) Schematics of refractive index control by changing the level of water binding, which can be realized in refinement process. As we decrease the water contents in an order from sample A1 to A2, B1, and B2, the refractive index of DNA-CTMA thin solid film decreases monotonically. Detailed processes to distinguish samples will be described in the text.
In this study, we experimentally demonstrated that our purification process can flexibly reduce the water content in DNA-CTMA precipitates and more importantly it can effectively enhance the cationic lipid binding to DNA, as schematically shown in Fig. 1(b). Four types of DNA-CTMA precipitates with different levels of cationic lipid binding were prepared and thin solid films were fabricated using them. Optical properties and thermo-optical coefficient of these thin films were characterized in conjunction with the cationic lipid binding, which lead us to a consistent and repeatable method to control the refractive index of DNA-CTMA thin solid films, for the first time.
2. Experimental details
It has been reported that DNA-CTMA thin solid films can be obtained with stDNA with a molecular weight level of higher than 1000 kDa [21], which have been applied various optical applications. Commercially available stDNA sodium salt has a molecular weight with an average value of 1000-1300 kDa (1500-2000 base pairs, 96% purity) [22]. We will restrict our investigations to this specific stDNA in the following discussions.
2.1 DNA-CTMA refinement process
DNA in aqueous solution will significantly increase non-Newtonian elastic properties because of its long chain-like structure, which leads to a longer relaxation time in comparison to conventional synthetic polymers [23]. On the while, C-16 chain of CTMA cation is expected to have a higher mobility in aqueous solution than DNA. And interactions between the hydrocarbon chain of CTMA and the DNA phosphate backbone, especially the number of effective collisions between them would be one of keys to forming DNA-CTMA precipitates [24,25]. In this study, we tried to vary the number of effective collisions by establishing refinement processes, schematically shown in Fig. 2(a).
Fig. 2 (a) Schematic flows of the proposed refinement process for DNA-CTMA precipitates. Utilizing re-precipitation process, a systematic control of water content and CTMA binding to DNA was achieved. (b) FE-SEM images of precipitates and their surface morphology are presented. The scale bar is 1μm respectively.
Firstly, we divided precipitation processes into two categories A, and B. Sample A was prepared using the stDNA aqueous solution as the reactant basis, into which drops of the CTMA aqueous solution were added. In contrast, sample B was using the CTMA aqueous solution as the reactant basis, into which drops of the stDNA aqueous solution were added. Note that sample A would represent a situation where less mobile DNA is dominantly rich and the interaction between CTMA and phosphate backbones of DNA would be allowed only around localized CTMA droplets. The sample B represents a contrasting situation where more mobile CTMA is dominant and the interaction between CTMA and DNA would occur along the entire DNA strands. After these processes, we obtained DNA-CTMA precipitate samples A1, and B1.
We further dissolved these A1 and B1, completely in a minimal volume of ethanol to make a supersaturated solution at an elevated temperature 55~60°C. Then deionized water with ~1/4 volume of ethanol was added, which was followed by vortexing to make the solution well-dispersed. The solutions were slowly cooled down to 3~5°C to produce re-precipitated powders, which was followed by the usual filtering and vacuum drying processes. After these processes, we obtained DNA-CTMA samples A2, and B2.
All of the samples were prepared from 4g/L solution of stDNA and 4g/L solution of CTMA-Cl. The prepared CTMA-Cl solution was made by diluting the 25wt.% CTMA-Cl stock solution by adding water. We added 250mL CTMA-Cl solution into 250mL of stDNA solution for the samples in group A, and added 200mL of stDNA solution into 200mL of CTMA-Cl solution for the samples in group B. The mixing process was carried out using buret for about 1 hour (4~5mL/min), followed by overhead stirring at room temperature for about 12 hours. The precipitate was separated from the reactants in a total volume of about 500mL through a 0.65μm PTFE filter and rinsed three times with deionized water. Then, it was dried in a vacuum oven at 55 to 60°C for 24 hours to obtain a primary product. Next, 300mg of the primary product is taken in a Falcon tube (50mL), and then a minimum amount of ethanol (about 6mL) is gradually added in the bath at 55 to 60°C. When it is completely dissolved, deionized water was added in a small amount of 500μL per reaction to make a supersaturation condition at an elevated temperature of 60 to 65°C. About 2.0mL of deionized water was added to dissolve the primary product transparently, followed by vortexing to disperse evenly. The aqueous solution was slowly cooled to room temperature at about 1°C/min, and maintained at about 4°C to re-precipitate. The resulting second-round precipitate was filtered and rinsed again and dried in a vacuum oven for 24 hours to obtain a secondary product. The yield was about 74%, with the weight of about 220mg.
FE-SEM image of DNA-CTMA precipitate samples are presented in Fig. 2 (b). Comparison between A1 and A2 and comparison between B1 and B2 clearly shows that the surface of the powder gradually increases in wrinkles, and the interface between the layer and the layer becomes more apparent after our refinement process. This morphology resembles the stratified structure pattern of DNA-CTMA as predicted by previous studies [19,25,26].
We used thin-layer chromatography (TLC) analysis to observe the remaining DNA or CTMA molecules from the solution after precipitation. The base pairs of the DNA contains an aromatic ring and exhibits absorption near λ = 254nm in UV. In addition, the nitrogen contained in CTMA is easily oxidized by an oxidizing agent and can be visualized as a dyed form. Therefore, by measuring the UV absorption at λ = 254nm, and KMnO4 staining test, we can estimate the trace amount of DNA and CTMA in the filtered solutions, respectively. For the separated solution from A2, and B2 samples, neither DNA nor CTMA was observed. This result indicates significant improvement of collision binding between anionic sites of the phosphate backbone of DNA and the cationic sites of CTMA through our re-precipitation process.
2.2 Thin-film fabrication
Thin film fabrication experiments were carried out under a constant temperature 20°C and humidity of 0.5mmH2O in a Class1000 clean room. A TFT-Tech Manual spin-coater was used to make thin films on fused silica substrate and dry oxide treated Si-wafers. All substrates were cleaned in the following the conventional procedure, 1 minute in acetone, 30 seconds in isopropyl alcohol, and followed by deionized water rinsing. Using the 4 types of DNA-CTMA precipitates, solutions with concentration of 2.0wt.% were prepared using 99.5% 1-Butanol solvent. Approximately 30μl of the solutions was dropped on the substrates and spin-coated with a sequence of 2 seconds at 500rpm, 3 seconds at 2000rpm, and 10 seconds at 4000rpm. Uniform film with the thickness of 60 ± 5 nm were obtained and these samples were then vacuum dried at 60°C temperature for 24 hours and annealed at 120°C for 2~3 minutes. The samples were stored in a desiccator.
In order to have a reference sample, we also fabricated pure stDNA thin film without adding CTMA. Using deionized water 0.5wt.% stDNA solution was prepared. After the identical cleaning procedures as above, the substrate was heated at 120°C for 5 minutes to facilitate water evaporation in spin coating process. Approximately 50μl of the stDNA aqueous solution was dropped on the substrate and spin-coated with a sequence of 10 seconds at 500rpm, 5 seconds at 2000rpm, and 10 seconds at 3000rpm, which produced a uniform film with the thickness of 160 ± 5nm. After vacuum drying at 60°C temperature over 24 hours, the samples were stored in a desiccator.
2.3 Calculation bound water and CTMA saturation
Elemental Analysis [27] was performed with Thermo-Scientific FLASH 2000 series CHNS/O (Eager Xperience S/W), 1H-NMR spectrums were measured using a Bruker Avance-300 FT-NMR Spectrometer. A sample volume of about 50mg was used. The measurement sample had to be dissolved in methanol-d4. The calculation of the relative bound water content versus CTMA saturation obtained by NMR analyses [19,28] and elemental analysis is summarized as follows.
In the NMR analysis, δ is a value corresponding to the chemical shift, and the scale of δ is expressed in ppm in units of Hz/MHz. In this regard, the relative distribution of specific functional nuclei in the material can be calculated from the area integral of the measured NMR spectrum intensities as follow;
Considering the condition of having four phosphates with two DNA base pairs (A-T, C-G), the occupation number of nitrogen (N) and carbon (C) in, NDNA = 15 and CDNA = 39 are obtained. In the four CTMA ligands, we have NCTMA = 4 and CCTMA = 76. Using these occupation numbers, the composition ratio of carbon to nitrogen was assumed to be S0 = CDNA/NDNA = 2.60, when there is no conjugated CTMA ligand. In contrast, when all of phosphate groups of DNA construct electrostatic interactions with CTMA ligands, the ratio becomes S = CDNA-CTMA/NDNA-CTMA = 6.05. Therefore, difference of the ideal composition ratio is ΔS = S - S0 = 3.45. The experimental mean occupation number, [N] and [C] were obtained by dividing the mass percentage from elemental analysis by the atomic weight of each element. Therefore, difference of the experimentally measured composition ratio is Δ[S] = [S] - [S0] and results from elemental analysis of sample A1, A2, B1, and B2 could interpreted as saturation degree as
The value [S] of each element was obtained as follows, sample A1: 4.5588/0.7141 = 6.384, A2: 4.4362/0.7616 = 5.825, B1: 4.4280/0.6998 = 6.327, and B2: 4.4511/0.7301 = 6.097, respectively.
3. Results and discussion
We investigated chemical properties of our powder samples using FT-IR and NMR to quantify the impacts of our re-precipitation process. 1H-NMR spectrums were measured using a Bruker Avance-300 FT-NMR Spectrometer.
Comparison of the FT-IR spectra of stDNA and CTMA-Cl is summarized in Fig. 3(a). Two types of stDNA with a molecular weight of ~9kDa (gray solid curve) and ~1,000kDa (red dotted curve) are shown in the upper graphs. Note that stDNA with ~1,000kDa was used in our study. Major peaks were observed near 1093 and 1238cm−1, which correspond to symmetric stretching and asymmetric stretching of the phosphate backbone (PO2)-. On the while, peaks related with nucleobases were observed in the spectral range 1500 to 1800cm−1. As the molecular weight increases, the DNA chain extends in a longer length and the water binding probability also increases [29,30], which is attributed to the spectral shifts in the nucleobase related peaks. In addition, the water remaining in the phosphate group may affect the binding pattern of the base to base, which may also alter the behavior of DNA nucleotides. We assume that the intensity changes in the peaks 1604, 1654 and 1695cm−1 could serve as an indicative of how the bound waters are affecting the vibration modes of nucleobase [31–33]. On the while, CTMA-Cl showed major peaks near 2850 and 2918cm−1, which correspond to vibration and stretching of C-H bonds.
Fig. 3 FTIR spectra of (a) stDNA and CTMA-Cl, stDNA with a molecular weight of ~9kDa (gray solid curve) and stDNA with ~1,000kDa (red dotted curve) are shown. Pb, Cy, Ad, Th and Gu represent Phosphate backbone, Cytosine, Adenine, Guanine and Thymine, respectively. (b) FTIR spectra of DNA-CTMA samples. The dotted lines in (a) and (b) indicate the spectral locations of phosphate backbone, nucleobases of DNA, and C-H bond of CTMA. (c) 300MHz FT-NMR spectra. Peaks correspond to the chemical shifts of H2O, –CH2 and -CH3 from CTMA ligand.
FT-IR spectra of DNA-CTMA samples prepared by our proposed re-precipitation method are shown in Fig. 3(b). During the refinement process, the relative peak intensity at 1093cm−1 decreased in comparison to the 1238cm−1 peak. This tendency is attributed to the well-bound cationic lipid in the phosphate group. The peak of symmetric stretching of (PO2)- at 1093cm−1 reduced by the long chain form of CTMA, while the asymmetric stretching at 1238cm−1 increased in the order from A1 to A2, and B1, B2. This is also attributed to the C-H elasticity of CTMA, which serves as a direct evidence that our proposed re-precipitation method did increase the CTMA binding to DNA [33,34].
The 1H-NMR spectrum analyses of DNA-CTMA samples are summarized in Fig. 3(c), where the peaks corresponding to OH of water molecule and -CH2, -CH3 of CTMA are assigned according to prior reports [28]. The dipole of water molecule in the DNA-CTMA complex weakens the electron shielding to result in a chemical shift to the higher ppm. The water peak located at 4.88 ppm in sample A1 is higher than 4.83 ppm in sample B2, which indicates sample A1 has higher relative water content than B2. In addition to these shifts, the NMR spectrum integration values indicate that the water peak of B1,B2 decreased more than three times in comparison to sample A1. As expected from the schematic diagram in Fig. 1, we experimentally confirmed significant reduction of water content in DNA-CTMA can be achieved by using CTMA reactant basis with re-precipitation process.
We further analyzed impacts of our re-precipitation process in enthalpy and heat capacity of DNA-CTMA, using thermo-gravimetric analysis (TGA), and differential scanning calorimetry (DSC) [35]. The experimental results are summarized in Fig. 4.
Fig. 4 (a) TGA results for commercial stDNA, and DNA-CTMA precipitate samples prepared in this study. (b) DSC analyses of stDNA, DNA-CTMA samples.
As shown in Fig. 4(a), general trends of TGA curves of stDNA and DNA-CTMA A1 sample were similar to those in prior reports [14,36]. But it is noteworthy that in comparison to A2, B1 and B2 samples, stDNA and A1 sample showed significantly larger initial weight losses, which is attributed to relatively higher water contents. The initial weight loss due to water content in TGA is consistent to the NMR analyses such that we could confirm that water content in DNA-CTMA can be effectively reduced using CTMA reactant basis (B1) and re-precipitation process (A2, B2).
In DSC measurements shown in Fig. 4 b), the glass transition temperature, at which thermal molecular motion [37,38] begins to take place inside the complex, was observed around 148~155°C on prior reports [14,39]. Before reaching the temperature, endothermic behaviors are observed, which varied with the bound water content. In this endothermic reaction section, the volatile residue inside the material is heated and evaporated, so the weight of the material is also reduced. Interestingly, the samples A2, B1 and B2, which have less water contents than A1, showed significantly faster enthalpy relaxation than A1 and stDNA, as shown in Fig. 4(b). This is again consistent to other measurements to confirm that our re-precipitation process decreased the water contents in DNA-CTMA. However, the sample A2 showed the endothermic peak located in front of the other samples, because it has lowest saturation of CTMA than other samples (See the Table 1). Beyond the glass transition regions in DSC curves, we could observe the thermal decomposition regions which are conspicuously represented by exothermic reaction peaks. The decomposition temperatures of DNA-CTMA sample A1, A2 were about 230°C, which is about 5°C lower than those of sample B1, B2. When DNA-CTMA is stacked to each other, electrostatic interaction is formed by the CTMA chains. The more CTMAs bound to the DNA backbone in these materials, the higher temperature required for the thermal decomposition process. Therefore, even though the saturation of CTMA value of Sample A1 is relatively high, it can be predicted that there are more unbound cationic lipid remaining in the material from the DSC analysis result.
Table 1. Detailed comparison of refined DNA-CTMA samples
By the above material analyses, we could experimentally confirm that our proposed re-precipitation method could effectively control the binding of CTMA ligand, and water content in DNA-CTMA complexes, for the first time. It is noted that the method to make Group B sample made by using the CTMA aqueous solution as the reactant basis followed by the re-precipitation process is an optimal method to reduce the water content and increase CTMA binding in DNA-CTMA. As discussed above, CTMA saturation and relative bound water ratios in the prepared samples were estimated using elemental analysis [27] and NMR [19,28], respectively and the results are summarized in Table 1.
After confirming impacts of reactant base and re-precipitation process over DNA-CTMA powders in the viewpoint of chemical analyses, we further investigated impacts on DNA-CTMA thin films for optical applications. Using these DNA-CTMA powders, 2.0wt.% solution was prepared using 1-Butanol solvent. Approximately 30μl solution was dropped on a cleaned dry oxide treated Si-wafer substrate and spin-coated to produce a uniform thin solid film with the thickness of 60 ± 5nm. The surface states of thin films were compared using atomic force microscopy (AFM), and the results are summarized in Fig. 5.
Fig. 5 AFM images of DNA-CTMA thin-films prepared by identical spin coating process, the scale bar is 1μm respectively.
In previous studies structures of the DNA-lipid complex has been graphically presented [40,41], yet this is the first systematic comparison of DNA-CTMA films with different water contents or equivalently different levels of CTMA lipid binding. It is noted that the DNA shrinking [42], shown as white dots in Fig. 5, was significantly reduced in thin films made from our re-precipitation process (B1, B2), and these films were found to be networked in a longer ranges than A1 and A2. These observation would be related with the water content and CTMA binding and further investigations are being pursued by the authors.
For the prepared DNA-CTMA thin films, optical characteristics were analyzed in terms of refractive index and the absorption coefficient and results are summarized in Fig. 6.
Fig. 6 (a) Refractive indices of thin solid films deposited on Si wafers for stDNA, DNA-CTMA samples prepared in this study. (b) UV absorption spectra of thin films deposited on silica substrates. Dot-dashed, dashed lines represent Tauc-Lorentz oscillators for electron transitions n- π* (4.47eV) and π- π* (4.74eV), respectively.
Refractive indices were estimated using an Alpha Spectroscopic Ellipsometer (Woollam Inc.) and it was based on isotropic Cauchy model similar to prior report [5,43,44]. As shown in Fig. 6(a), we included data for stDNA thin film with the thickness about 160nm which was fabricated by spin coating of stDNA aqueous solution on Si wafer substrate, and we found the refractive index of our DNA-CTMA thin films were lower than that of stDNA thin film, consistent with prior reports [45–47]. It was noted that among our DNA-CTMA thin films, the refractive index, n, was decreasing in the order of nA1 > nA2 > nB1 > nB2 in the spectral range from 400 to 900nm, and its values are nA1 = 1.527, nA2 = 1.521, nB1 = 1.519 and nB2 = 1.517 at λ = 589nm, respectively. The corresponding refractive index of stDNA film was n = 1.558. This order in the refractive index magnitude is consistent to above material analyses showing the similar order with which the water content is decreasing as summarized in Table 1. We experimentally found that both the water content and CTMA binding influenced the refractive index of DNA-CTMA films. In Table 1, the relative bound H2O contents were 1.4 in Sample B1, which is lower than 1.67 in Sample B2. In contrast, the CTMA saturation value was 1.080 in B1, which is about 0.07 higher than B2. The refractive indices of films made of these materials were n = 1.519 and 1.517 for B1 and B2. We found a trend experimentally that the lower water contents and the higher CTMA binding would result in a lower refractive index in DNA-CTMA films. But we could not determine which of the two would be a dominant factor and this issue is being investigated by the authors. Therefore, we confirmed that the water content and CTMA binding in DNA-CTMAs also played a critical role to vary their thin films’ refractive indices. It is noted that the index difference between A1 and B2 was Δn = 0.01, which is high enough to make an optical waveguide whose core and cladding are all DNA-CTMA. We experimentally observed that our proposed re-precipitation process serves as a unique method to systematically to control the refractive index of DNA-CTMA thin solid films, which could open a new potential toward all-DNA photonic device applications. As shown in Fig. 6(b), UV absorption spectra also showed consistent changes such that the absorption coefficient, α, corresponding to the π- π* electron transition decreased in the order of αA1 > αA2 > αB1 > αB2. Nucleobases of DNA are rich in π-electrons, and electron transitions can occur vertically or horizontally in nucleic acid plane structures. However, as CTMA binding increases, in-plane transitions (C = C) are known to be inhibited because the number of exposed nucleobases are reduced [48,49].
Thermally induced changes in DNA-CTMA thin films were measured using a temperature controlled ellipsometer, and the results are summarized in Fig. 7. Firstly we measured the thickness change under a temperature cycle from 30 to 100°C. The heating rate was about + 10°C/minute and cooling rate was −5°C/minute, which was controlled by a Peltier thermoelectric device. By the temperature cycles, DNA-CTMA thin films showed variations in the thickness as shown in Fig. 7(a)-7(d), which was similar to previous reports such that thickness decreased as the temperature increased in the first heating cycle, and the thickness showed a slightly thinner value than the initial thickness after the first cooling cycle [50]. However, meaningful differences were observed after the first heating/cooling cycle, which are shown in the dashed curves in Fig. 7(a)-7(d). Difference in the thickness before and after thermal cycles was significantly reduced in A2, B1, and B2 in comparison to A1. After the second cycle, the thickness variations of the samples were ~0.35nm in A1, ~0.10nm in A2, ~0.11nm in B1, ~0.07nm in B2. Especially sample with a less content of water and more CTMA binding than A1 did show less than 1/3 changes in the thickness after the second thermal cycle, which strongly indicates the water contents and CTMA binding did play an important role in thin film structuring.
Fig. 7 Thermally induced changes in DNA-CTMA thin-films. (a-d) Thickness changes as a function of temperature. (e) Refractive index changes as a function of temperature for vacuum dried films and (f) refractive index changes for pre-heated samples after the second thermal cycle.
In order to further investigate the impacts of water contents and CTMA binding to thermo-optic properties, we measured the refractive index changes as a function of temperature. The DNA-CTMA thin films were vacuum dried at 60°C temperature similar to [17,48] and then the pre-heated at 120°C [12] after the second thermal cycle. Note that the thickness of the film were fitted to the average of the values in the second thermal cycle shown in Fig. 7(a)-7(d), which was used to calculate the thermo-optic coefficient, dn/dT, in Fig. 7(e) and 7(f). The refractive index changes (Δn) were referenced to the refractive indices of the samples at 20°C, at λ = 633nm. The sample B1 and B2 showed a significantly less error in the linear fitting to obtain thermo-optic coefficients, dn/dT, than the samples A1 and A2. It is noted that the magnitude of dn/dT of B1, B2 significantly reduced when the films were preheated, which is attributed to removal of residual butanol solvent in the thin films. The refractive indices and the linear thermos-optic coefficient values for all the samples are summarized in Table 2, and dn/dT of these DNA-CTMA samples were comparable to conventional optical polymers [51].
Table 2. Refractive indices (n) and thermo-optical coefficient of DNA-CTMA (dn/dT × 104 [°C−1])
4. Conclusions
We proposed a new chemical refinement method for DNA-CTMA complex based on re-precipitation process that could systematically control both the water content and CTMA. The method subsequently allowed stable control of refractive index and thermos-optic coefficients of thin solid DNA-CTMA films. Materials analyses such as FT-IR, NMR, and FE-SEM, experimentally confirmed that our proposed method did provide a well-defined practical path to reduce the water contents by a factor of 3 in DNA-CTMA precipitate powders. Thin solid films were fabricated by spin coating of the prepared samples and the proposed method resulted in a refractive index Δn = 0.01 at λ = 633nm among DNA-CTMA films, which is high enough to make an optical waveguide whose core and cladding are all DNA-CTMA. Thermo-optic coefficient dn/dT was found to vary in the range from −1.383 to −2.592 × 10−4 °C−1. Further investigation on surface wetting properties would provide a new potential for all DNA based opto-electronic devices in bio-compatible photonic applications.
Funding
Institute of Physics and Applied Physics, Yonsei University, in part by Nano Material Technology Development Program through NRF funded by the MSIP (NRF-2012M3A7B4049800, NRF-2012M3A7B4049802).
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