Dispersion and Exfoliation of Nanotubes with Synthetic

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J. Phys. Chem. C 2010, 114, 11741–11747

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Dispersion and Exfoliation of Nanotubes with Synthetic Oligonucleotides: Variation of Dispersion Efficiency and Oligo-Nanotube Interaction with Base Type J. Marguerite Hughes, Helen Cathcart, and Jonathan N. Coleman* School of Physics and Centre for Research on AdaptiVe Nanostructures and NanodeVices (CRANN), Trinity College Dublin, Dublin 2, Ireland ReceiVed: March 1, 2010; ReVised Manuscript ReceiVed: May 7, 2010

Single walled carbon nanotubes (SWNTs) were dispersed and exfoliated in four different homopolymer oligonucleotides (dA15, dG15, dC15, and dT15). The dispersed nanotube concentration and degree of exfoliation were measured for each nucleobase. The nanotubes were more highly exfoliated and more temporally stable in dC15 and dT15. While the degree of exfoliation was relatively time independent, absorption and photoluminescence spectra showed definite changes over time after the initial sample preparation. In particular, photoluminescence signals appeared at well-defined times, consistent with previous evidence of time-dependent DNA wrapping followed by oxide removal. Analysis of the nanotubes’ optical properties, including circular dichroism, suggests that all bases except adenine stack onto the nanotube surface. In contrast, dA15 is unstable on the nanotube surface and eventually returns to a self-stacked arrangement. The order of the dispersion efficiencies was found to be T > C > G . A, where thymine produced the most intense NT optical signals and cytosine was seen to wrap SWNTs the fastest. Introduction Single walled carbon nanotubes (SWNTs) have been proposed for use in applications as varied as electronics,1 body armor,2 and sensors for DNA hybridization.3 A prerequisite for the realization of many of these applications is the development of a reliable method for preparing high-quality SWNT dispersions with large populations of individual nanotubes and few bundles. Much progress has been made in this area, and a range of different techniques is now available.4 Such techniques include theuseofsurfactants,5-8 certainsolvents,9-13 polymer-wrapping,14,15 strong acids,16 surface functionalization,17,18 and synthetic peptides19-22 in the preparation of liquid-phase SWNT dispersions. DNA has been used successfully to disperse SWNTs in water.23-28 Such dispersions have a high population of individual nanotubes and low mean bundle diameters. In addition, they are thought to be biocompatible and extremely promising for medical applications.29-33 Furthermore, DNA-wrapped SWNTs have been separated according to type/chirality by ion-exchange chromatography, or density gradient ultracentrifugation.27,34,35 DNA-based SWNT dispersions have been prepared using both natural and synthetic DNA. The accepted interaction mechanism is helical wrapping, whereby the hydrophobic DNA bases preferentially π-stack (noncovalent bonding) on the nanotube surface, leaving the hydrophilic backbone free to interact with the water.24,36 Cathcart et al. have shown that the DNA can then rearrange itself into an ordered helical pattern about the nanotube.28 This study was performed with the aim of investigating more closely the effect of nucleotide base choice on the interaction. To date, several experimental and theoretical studies have indicated that pyrimidine (cytosine (C) or thymine (T))-based oligonucleotides are superior dispersants to those containing only adenine (A) or guanine (G) (purines).24,37,38 In particular, it has been seen that poly(T) has a higher dispersion efficiency than other oligonucleotides, that is, that a greater proportion of nanotubes remain dispersed in solution with * To whom correspondence should be addressed. E-mail: [email protected].

poly(T) following centrifugation than with the other oligos, particularly poly(A).24,39 This is in spite of the fact that the purines are larger molecules and thus have a greater nanotube surface area with which to interact, offering the possibility that poly(A) and poly(G) would bind better to the nanotube and provide better nanotube coverage.40 However, such predictions are based on simulations involving monomer units of DNA. Detailed theoretical considerations suggest that when a DNA strand containing only purines is deformed, there is a large base-base repulsion which increases the strain energy and thus increases the formation energy of the DNA/SWNT hybrid, particularly for large-diameter nanotubes.41 Therefore, strands with only pyrimidines can wrap and solvate nanotubes more efficiently. Many of the aforementioned existing experimental studies on the interactions of different bases with SWNTs have focused primarily on the concentration of SWNTs remaining dispersed after centrifugation as the main method of comparing dispersion efficiencies of the different homopolymer oligonucleotides. However, this provides no information about the wrapping between the DNA and nanotube. Furthermore, the postcentrifugation nanotube concentration (PCC) is strongly dependent on the sonication intensity received during sample preparation, which can vary between batches (even those ostensibly prepared under identical conditions). Second, measurement of the PCC does not reveal whether the SWNTs are dispersed in small bundles, or whether the dispersion contains a high population of individual nanotubes; that is, it is not a true measure of the dispersion quality. The wrapping of DNA around nanotubes is strongly time dependent,28 and although nanotubes may rapidly be dispersed and debundled by DNA, the presence of individuals of different species cannot be accurately ascertained by absorption or photoluminescence (PL) spectroscopy until wrapping is completed. The reason for this is that surface oxides, which produce holes in the π-electron valence band and lead to the quenching of the PL and absorption signals, may be present on the surface of a nanotube dispersed in water, and if so to be

10.1021/jp101834t  2010 American Chemical Society Published on Web 06/22/2010

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removed before stable luminescence is observed.42-45 When DNA forms an ordered coating around a nanotube, it is thought that these surface oxides are transferred from the nanotube to the DNA. Thus, formation of a complete DNA monolayer around the nanotube coincides with the return of the absorption and PL signals.28 This investigation examines the impact of different nucleobases in the dispersion of SWNTs, and the quality of such dispersions, by using four short-stranded homopolymer oligonucleotides (dA15, dC15, dT15 and dG15) to disperse nanotubes and then by comparing them based on several different metrics (not solely the proportion of material remaining following centrifugation). This type of study is unique in comparing the base types. It not only utilizes absorption and photoluminescence spectroscopy, atomic force microscopy, and circular dichroism spectroscopy to characterize the samples (and the size of the nanotube bundles and individual nanotubes they exfoliate in the dispersion), but also investigates their behavior over time and correlates the results obtained using each technique to obtain a clear picture of how each nucleobase interacts with a nanotube. The samples’ performances are also assessed with respect to natural double stranded DNA. Experimental Procedure Four different oligonucleotides (dA15, dG15, dC15, and dT15) were purchased from Sigma Genosys (http://www.sigmaaldrich.com/life-science/custom-oligos.html). These oligos were short, single-stranded DNA, 15 nucleotides in length, which contained only one base type: adenine (A); guanine (G); cytosine (C); or thymine (T). The oligo stock solutions were prepared by dissolving each oligo in Millipore water at a concentration of 1 mg/mL, by adding the required quantity of water to the oligonucleotide and agitating it using a vortex. In the case of dG15, it was necessary to dissolve the oligo in 10 µL of 10 mM, pH 7 phosphate buffer before adding the Millipore water. The oligo:SWNT samples were prepared according to the procedure outlined by Cathcart et al.25 Briefly, the oligo stock solution was added to HiPco SWNTs (Carbon Nanotechnologies Inc., lot number PO342) in a round-bottomed flask at the mass ratio of 1:1 oligo:SWNT. This was stirred and sonicated for two minutes in ice-water using a Branson 1510 sonic bath (frequency 42 kHz, with a power output of 80 W). A total of 500 µL of water was then added every 2 min until the dispersion was at the desired nanotube concentration of 0.1 mg/mL. The flask was positioned in the bath such that the sonication intensity was maximized and the dispersion was sonicated for 2 h in total. Crushed ice was added to the bath every 20 min to prevent heating in the sample. Absorbance spectra were recorded for all dispersions, and the samples were then subjected to mild centrifugation for 60 min at ∼3300g in order to remove any large nanotube aggregates which were not dispersed during sonication. The postcentrifuge concentrations were calculated from the ratio of the absorbances before and after centrifugation. The nanotube concentrations were found to be 5.5 × 10-2 mg/mL for the dT15:SWNT sample; 5.5 × 10-2 mg/mL for the dC15:SWNT sample; 6.8 × 10-2 mg/ mL for the dG15:SWNT sample; and 6.2 × 10-2 mg/mL for the dA15:SWNT sample. [The PCC has often been used as a measure of the dispersion efficiency of the dispersant. It is important to note, though, that in nanotube dispersions the PCC is strongly dependent on the sonication intensity received during sample preparation, which can vary somewhat between preparations. So, given that relatively few samples have been prepared, we cannot say with certainty that the variation in PCC observed

Hughes et al. here is not within the statistical variation expected for samples prepared with only one type of oligonucleotide. To use this concentration for comparisons in the future, it would be advisable to prepare several samples using each oligo, which was not feasible here due to the small amounts of each oligionucleotide which were available.] The samples were then serially diluted, first to a common nanotube concentration of 4 × 10-2 mg/mL and then to a final concentration of 2.5 × 10-2 mg/mL using Millipore water. This was in order to properly compare the dispersion efficiency of the different nucleotide bases. Each sample received 30 min sonication in an ice-water sonic bath after each dilution step. They were subsequently analyzed using a range of techniques in order to compare nanotube bundle sizes and DNA coverage in each sample. The concentration of 2.5 × 10-2 mg/mL was chosen for a number of reasons. First, in similar natural DNAbased dispersions, the partial concentration of individual nanotubes (Mi/V) was found to be maximized at this concentration.25 Second, it was found to be a suitable concentration for atomic force microscope (AFM) characterization, as it produced samples with an appropriate number of SWNTs per unit area following deposition. Third, the concentration was low enough to prevent effects such as reabsorption and inner-filter effects from distorting the NIR-photoluminescence results.46 (It should be noted that throughout this paper the word “concentration” refers to the nanotube concentration unless stated otherwise.) Each sample was characterized regularly over ∼42 days to monitor any changes in the interaction between the oligos and the SWNTs. First, to indicate that the DNA was wrapping around the nanotubes and stabilizing the dispersions according to Derjaguin-Landau-Verwey-Overbeek (DLVO) theory,4,47 the zeta potential of the samples was regularly monitored. It was consistently found to be more negative than -20 mV for all samples concerned, indicating that they were indeed stabilized by electrostatic repulsion (although we cannot rule out a contribution from steric stabilization).48 [See also the Supporting Information.] Four other methods were employed to examine the oligo: SWNT hybrids. AFM analysis allowed examination of the hybrids and the calculation of statistical data pertaining to the nanotube/bundle diameters as a function of time. Samples were prepared by placing a drop of the sample on freshly cleaved mica and allowing it to dry under ambient conditions. The samples were then characterized using a Veeco Nanoscope IIIa AFM in tapping mode, with NCH-50 Pointprobe silicon cantilevers (quoted tip radius < 10 nm). Several images were taken of each sample at various points on the mica surface, and the diameter distribution of the sample was determined by measuring the heights of ∼150 objects (bundles and SWNTs/ hybrids) above the mica surface. The absorbance of the samples was measured regularly using a Cary 6000i UV-vis-NIR spectrophotometer. The samples were placed in a 2 mm path length quartz cuvette and analyzed over a wavelength range of 200-1300 nm. NIR-PL spectra were recorded using an Edinburgh Instruments FLS920 fluorescence spectrometer with a Hamamatsu R5509 near-IR photomultiplier tube. The samples were placed in a 2 mm × 2 mm quartz cuvette, thus minimizing reabsorption and inner-filter effects.46 The samples were excited at a wavelength of 650 nm and NIRPL emission was recorded over a 900-1300 nm wavelength range. Finally, circular dichroism (CD) spectra were recorded using a Jasco spectropolarimeter. Samples were placed in a 2 mm quartz cuvette and examined over a 200-350 nm wavelength range.

Dispersion and Exfoliation of Nanotubes

Figure 1. (A) AFM image of oligo dispersed SWNTs deposited on mica. (B,C) Statistical analysis of AFM data for all oligonucleotideSWNT hybrids. (B) Mean hybrid (bundle or individual) diameter as a function of time after sample preparation. (C) Population of individual nanotubes as a function of time.

Results and Discussion AFM images were recorded regularly, allowing monitoring of the distribution of diameters of the oligo:SWNT hybrids over time (Figure 1A). We expect these hybrids to consist of both nanotube bundles and individual nanotubes, stabilized by the oligos. From these distributions, one can determine the mean hybrid diameters and estimate the number of individuals per unit volume (among other things) as a function of time. The number of individuals per unit volume is expressed as

Ni 4CNT Ni Nt Ni ) ≈ V Nt V Nt F π〈D2〉L NT bun where Ni is the number of individual nanotubes measured (objects with diameter Di < 1.4 nm) and Nt is the total number of nanotubes/bundles measured. CNT and FNT are the concentration and mass density of nanotubes, respectively, while 〈D2〉 is the mean square diameter of nanotubes in the sample. Lbun is the average nanotube bundle length as calculated using the AFM, and it is assumed that SWNT bundles and individual nanotubes are roughly the same length.12 Both the mean diameter and the number of individuals per unit volume are used as metrics of the sample quality. Figure 1B shows the mean hybrid diameter as a function of time after sample preparation. This shows that all four oligonucleotides were initially very efficient at dispersing/exfoliating SWNTs, displaying mean hybrid diameters of between 1.2 and 2 nm on day 1; this is impressive, remembering that an uncoated individual HiPCO nanotube usually has a diameter < 1.4 nm.

J. Phys. Chem. C, Vol. 114, No. 27, 2010 11743 Including the DNA coating, objects in this size range can only consist of individual nanotubes or very small bundles. However, as time went on, the pyrimidine (cytosine and thymine) stabilized nanotubes remained well exfoliated, while for the purine (adenine and guanine) stabilized nanotubes the mean hybrid diameter appeared to increase after ∼25 days. This suggests the occurrence of aggregation, indicating that the purine-dispersed nanotube dispersions are unstable. It should be noted that, by day 42, small nanotube aggregates were visible in the dA15: SWNT sample, suggesting that this sample was particularly unstable. These aggregates were too big to be imaged with AFM and were not picked up by the AFM results. Such aggregates were not seen in the other samples. In addition, the samples prepared with dC15 and dT15 displayed lower average hybrid diameters at all times than those prepared with dA15 and dG15. This confirms the predictions that pyrimidines disperse and exfoliate SWNTs more effectively than purines. It should be noted that hybrids with diameters between 0.4 and 0.7 nm were observed in all samples. HiPCO SWNTs, though, typically have diameters of between 0.7 and 1.4 nm. This shift can be attributed to a chemical contrast effect between the hybrid and the mica. Nonetheless, although contrast effects may cause a slight shift in the diameters (and populations of individuals), the samples can still be compared relative to each other and analyzed for relative changes in the mean diameters over time. The mean nanotube diameters did not display any clear progression that would indicate a transition from an uncoated to a coated state over time. This is unlike samples prepared with double-stranded DNA (ds-DNA), where the mean diameters were observed to increase by ∼1 nm over time, as the DNA first frayed and unzipped onto the nanotube, forming a disordered coating, and then rearranged itself into a complete, ordered coating over time.28 However, in these samples, the oligonucleotides were already in single-stranded form and did not need to unzip in order to coat the nanotube. In addition, the oligos are very short, at just 15 bases long, which would both facilitate the rapid formation of a disordered DNA coating on the nanotube and also allow the DNA to rearrange itself into an ordered coating on a much faster time scale. Therefore, the ss-DNA can interact rapidly with the nanotube, suggesting that they form a disordered DNA coating on the nanotube walls even before the first AFM measurements are taken, and as a consequence few changes in the overall diameters of the nanotubes were observed over time, but other methods of analysis are needed to provide further clarification on the interaction between the oligos and the nanotubes. Another way of presenting the hybrid diameter data is to calculate the number of individual nanotubes per unit volume of dispersion as described above. This data is presented in Figure 1C. All dispersions had roughly 10 individuals µm-3, a value typical for good quality biomolecule stabilized nanotube dispersions.4 Individual populations were slightly higher for dT15 and dC15 dispersions at all times as expected. Importantly, the dT15 and dC15 dispersions were extremely stable, while the dG15 and dA15 dispersions showed evidence of aggregation at longer times. Absorption spectra in the wavelength range 200-1200 nm (due to the strong absorption of water above 1350 nm) were taken regularly over the course of the study to evaluate changes in the resolution and intensity of the nanotubes’ characteristic van Hove peaks (particularly in the Es11 region) and to look for changes in the intensity of the characteristic oligonucleotide absorption peak (∼250-270 nm). The Es11 van Hove absorption

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Figure 2. (A) Evolution in background subtracted absorption spectra for dT15:SWNTs with time. (B) Growth of (7, 6) peak with time for all hybrids. (C) Background corrected absorption spectra for all oligo: SWNT solutions, day 42.

peaks were initially flat and broad, and poorly defined. However, all samples showed progressive growth and sharpening of the Es11 van Hove absorption peaks over time. As a representative example, Figure 2A shows the background corrected absorption spectra for the dT15:SWNT sample at different times. The correction was performed by subtracting a k/λb background (k and b were empirically calculated from the spectrum itself) from the original absorption spectrum. This subtraction was carried out to approximate the removal of absorption due to scattering and/or plasmon effects following the procedure of Nair et al.49 It is clear that the Es11 peaks grow and become more resolved over time. Sharpening of van Hove absorption peaks in SWNT dispersions usually reflects debundling of nanotubes in the sample. However, these samples already contain large populations of individual nanotubes, and AFM measurements indicate that nanotube diameters remain fairly constant over time. Thus, we suggest that changes in the van Hove peaks are consistent with the formation of an oligo-DNA coating on the nanotube surface, followed by the transfer of surface oxides (which produce holes in the π-electron valence band and thus quench both photoluminescence and absorption signals42-44) from the nanotube to the DNA, that is, optical bleaching followed by oxide removal. This phenomenon has previously been reported for ds-DNA:SWNT samples.28 Thus, the rate at which the van Hove peaks switch on can be used to infer the rate of wrapping and oxide transfer for the different oligos. In order to compare different oligonucleotides, the intensity of the main peak between 1133 and 1151 nm, attributed to the (7, 6) nanotube (where the height is measured relatiVe to the local minimum at 1090 nm in each graph, to compensate for any overall changes in the extinction coefficient), has been plotted in Figure 2B for each dispersion as a function of time.

Hughes et al. It clearly shows that samples prepared with different oligos switch on at different times, suggesting that the rate of DNA wrapping (and subsequent oxide transfer) is base dependent. The sample prepared with dC15 switched on first, demonstrating a saturated absorption signal after just 5 days. This was followed by dG15 and dT15, which took 8 and 15 days respectively for the absorbance to appear. The dispersion prepared with dA15 was by far the slowest, taking a total of 26 days. This indicates that either ordered DNA wrapping was slowest for the dA15: SWNT hybrids or else oxide transfer was less favorable for this oligonucleotide. This said, it should be noted even the dA15: SWNT sample switched on faster than samples prepared with natural ds-DNA which took ∼35 days to switch on. This supports the hypothesis that short ss-DNA both covers the nanotubes and forms an ordered coating faster than the longer double stranded natural DNA. With regard to the absolute magnitude of the (7, 6) absorption peak, however, it would be misleading to suggest that a larger peak by itself indicates superior debundling, as the appearance over time of a shoulder peak at 1190 nm (due to the (8, 6) nanotube) corresponds to a relative decrease in the (7, 6) peak height. For example, this shoulder peak did not resolve at all for dA15:SWNT, and in spite of the (7, 6) peak intensity for dG15/SWNT on day 42, the (8, 6) peak was barely present therein either. In this respect, the dT15 performed best, followed by the dC15, then dG15, and, as mentioned, the dA15. Figure 2C displays background corrected absorption spectra for all hybrids on day 42. First, the shape of the dA15:SWNT spectrum between 400 and 900 nm following subtraction is noticeably different from that of the other complexes; this is primarily due to the spectral shape in the NIR region, which affects the empirical calculations. Nonetheless, this supports the evidence that the dA15 behaves differently from the other oligonucleotides and may aid the confirmation of AFM trends.50 Subtler shape differences between the different dispersions can be seen clearly in the Es11 region. Between 950 and 1200 nm, the dT15:SWNT and dG15:SWNT hybrids display several wellresolved peaks; in comparison, the dA15 produces flatter, broader peaks. The dC15:SWNT spectrum was broader generally than either the dG15 or dT15 spectra but yet displayed the resolved shoulder peak between 1120 and 1200 nm. Furthermore, the peak in the dA15:SWNT hybrid is red-shifted with respect to the same peak for the guanine and pyrimidine complexes (and also red shifts over time with respect to its own original position, unlike the other three). Shifts in the peak positions are consistent with changes in the nanotubes’ local dielectric constant due to different levels of shielding from the surrounding water by the DNA.51 Within the context of the overall results, this suggests that the dA15 produces a lower level of nanotube coverage than dT15, dC15, or dG15. Figure 3A shows the background corrected heights of the characteristic DNA absorption peak (around 260 nm) for each oligo as a function of time. In contrast to the van Hove peaks, the oligonucleotide peaks showed a steady decrease in intensity (hypochromicity) over the course of the study. Hughes et al. have shown that, compared to free DNA in solution, DNA bound to nanotubes exhibits hypochromicity as a result of stacking between the bases and the nanotube surface.39 Thus, the steady decrease in the peak intensity is consistent with increased stacking of the oligo bases on the nanotubes over time. On the other hand, the same result can occur if the intrabase stacking within the DNA molecules is increased; it is known that a single strand of DNA will absorb less than the sum of its nucleotides (and that double-stranded DNA in turn absorbs less

Dispersion and Exfoliation of Nanotubes

Figure 3. (A) Decrease in absorption intensity for characteristic oligonucleotide peak. (B) Circular dichroism (CD) spectrum of dC15: SWNT as a function of time. (C) Change in CD peak-peak heights over time, for all dispersions. (D) Oligonucleotide peak extinction coefficient versus CD peak-peak heights, each measured at various times.

than single-stranded DNA). In addition, sonication can disrupt the intrabase stacking in DNA. Thus, the observed hypochromicity in the π-π* transitions could be caused either by stacking between nanotube and nucleotide base or by recoVery of self-stacking between successive bases in the oligo after sonication and/or desorption.39,52-54 Thus, a measurement technique which is sensitive to stacking between nucleotide bases is needed to clarify these results. Such a technique is circular dichroism spectroscopy. Circular dichroism (CD) spectroscopy was used to monitor the proportion of free “self-stacked” DNA in solution. CD operates by measuring the differential absorption of left- and right-handed circularly polarized light, producing a characteristic spectrum for a given molecular structure and thus providing information on the secondary structure of a molecule. The intensity of the CD spectrum is proportional to the concentration of individual nucleotide bases in solution, but it is also amplified up to 10 times by the interactions between consecutive bases. Thus, if there is a high concentration of free self-stacked DNA in solution, a large CD peak-peak intensity will be observed. However, as the DNA wraps around the nanotube, it is favorable for the bases to rotate and π-stack onto the nanotube walls, for example, as described by Meng et al.55 This disrupts the base-base stacking and reduces the CD intensity. Thus, the time-dependent change in CD intensity allows one some insight into the processes occurring in the samples. If the CD intensity is found to decrease over time, it suggests that the base-base interactions are being disrupted in favor of base-nanotube

J. Phys. Chem. C, Vol. 114, No. 27, 2010 11745 π-stacking, and the DNA is coating the nanotubes. However, if the CD intensity increases over time, it suggests that the bases in the DNA that were initially disrupted during sonication (and/ or stacking on the nanotube) are becoming detached from the surface and self-stacking in solution; thus, the DNA is not coating the nanotubes in a well-defined fashion. All of the oligonucleotides displayed strong CD signals consistent with a right-handed helical form, and their spectral shapes did not change appreciably over time. Figure 3B shows the CD spectra for the dC15:SWNT sample recorded throughout the experiment. It is clear that, in this sample, the CD intensity decreases over time. For easy comparison, the peak-peak intensity was measured for all samples and plotted as a function of time in Figure 3C. This clearly demonstrates a marked decrease in the CD intensity and, hence, a decrease in the proportion of free, “self-stacked”, DNA in solution for the pyrimidine-based dispersions and the dG15:SWNT dispersion. In all cases, it was found that the CD peak-peak intensity was less intense in the oligo:SWNT dispersions than for similar oligo-only samples of the same oligo concentration. This suggests that either sonication has disrupted the base-base stacking in a portion of the “free” DNA in solution or else some of the DNA has formed an irregular coating on the SWNT walls during the initial stages of the experiment, as suggested by the AFM results, since there is later a change in the DNA:SWNT optical properties that is not accompanied by a substantial change in diameter. Thus, it can be inferred that the DNA is continuously coating the nanotubes over time in these samples. However, the peak-peak intensity for dA15:SWNT increases over the course of the study, suggesting that the dA15 bases are self-stacking in solution rather than coating the nanotube. This suggests that while an irregular DNA coating forms on the nanotube during sample preparation allowing the nanotubes to be dispersed in water (indicated by AFM), a regular uniform dA15 coating does not readily form around the nanotube. This signifies that the slow “switch on” time for the van Hove absorbance peaks is due to the fact that the DNA does not readily form a uniform coating over the nanotube rather than because oxide transfer is unfavorable. It is also known that poly(A) has the greatest tendency to self-stack of poly(A), poly(T), poly(G), and poly(C). Its CD spectrum in aqueous solution is consistent with strong self-stacking, whereas poly(T), for example, is expected to be in a random coil form. Therefore, our results could directly represent competition between selfand cross-stacking in dA15.38,56 These findings support the previous results that demonstrate its lower capability to successfully disperse and debundle nanotubes in aqueous solution, and suggest a reason for the visible aggregation that was observed toward the end of the experiment. That the hypochromicity of the oligo absorption peak and the decrease in intensity of the CD peak for the poly(T), poly(G), and poly(C) samples are both controlled by adsorption of bases onto the nanotube surface is illustrated in Figure 3D. Here we show that the DNA peak intensity scales with the CD intensity, suggesting they share the same origin. Conversely, for the dA15 sample, these quantities are anticorrelated. This may be understood by attributing the origin of the oligo peak hypochromicity to self-stacking of a single strand of dA15 (instead of crossstacking onto SWNTs as in the other samples) and, as stated, the growth in the CD peak-peak heights also being attributable to self-stacking. Photoluminescence (PL) spectroscopy was the final tool employed to explore the wrapping of the oligonucleotides around the SWNTs. This is a useful tool for characterizing nanotube

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Hughes et al. different behavior was observed: for the dG15:SWNT samples, the PL turned on gradually, while for the dA15:SWNT samples a step increase was observed but at a much later time (∼25 days). This underlines the difference in dispersion performance between the respective oligonucleotides. These results strongly support the findings from AFM and spectroscopy that suggest the pyrimidines, and dT15 especially, are better dispersants of SWNTs than the purines. Conclusions

Figure 4. (A) PL spectra over time for dT15:SWNT. (B) Evolution in intensity for (7, 5) nanotube over time, for all samples.

dispersions, as primarily individual nanotubes are luminescent. However, as discussed previously, even individual nanotubes can have their PL quenched by surface oxides.42 Ordered wrapping by DNA is thought to remove these oxides, leading to the restoration of the nanotubes’ luminescence.28 Line scans were regularly performed on the samples at an excitation wavelength of 650 nm over a period of 42 days following sample preparation. Shown in Figure 4A are the PL spectra measured for the dT15:SWNT sample. These data clearly show the PL intensity increasing dramatically over the first 10 days before falling off slowly. Since predominantly individual semiconducting nanotubes fluoresce, the intensity of the photoluminescence signal is usually used to infer the level of debundling within a solution. The PL intensity, IPL, is directly proportional to the number of individual nanotubes: IPL ∝ Ni/V. However, the AFM results reported in Figure 1C show that Ni/V does not increase significantly in any sample over the course of the experiment. Thus, in this case, the change in PL intensity is not an indicator of debundling in the samples. It was previously shown that, for natural DNA:SWNT dispersions, individual semiconducting nanotubes only fluoresce when wrapped with an ordered DNA coating. Therefore, the rate at which the PL signal appears is related to the time taken for the DNA to form a complete, ordered coating around the nanotubes. In the case of ds-DNA and SWNTs, it was previously observed that the photoluminescence took ∼35 days to appear. Around this time, the PL intensity increased in a steplike manner to a much higher level. This signifies the completion of an ordered wrapping of at least a monolayer of DNA.28 It was expected that, for the oligonucleotides, this process would be substantially quicker due to the fact that the DNA was both single-stranded and much shorter, allowing it to quickly form a coating on the nanotube. Figure 4B shows the PL intensity as a function of time for all four samples. For both pyrimidines (dC and dT), we do indeed observe a steplike increase occurring during the first 10 days. However, for the purines, slightly

The order of these oligonucleotides’ dispersion efficiencies (from greatest to least) is found to be dT15 > dC15 > dG15 > dA15. Thymine-based oligo:SWNT dispersions produce the most intense photoluminescence and absorption signals, indicating that they may be the most efficient at removing surface oxides from nanotubes, although cytosine-based oligos produce the fastest change in optical signals, indicating quicker rates of DNA wrapping around nanotubes. In contrast, it appears that adenine does not readily stack on the nanotube surface, and adeninebased nucleotides instead tend to self-stack in water instead of forming an ordered coating around SWNTs, leaving the latter vulnerable to surface oxides and hampering spectroscopic analysis; that is, competition between self- and cross-stacking of DNA is a significant factor when considering the capacity of these ss-DNAs to wrap nanotubes. This study supports existing experimental and theoretical data indicating that dispersions of pyrimidine-stabilized nanotubes are superior to those stabilized with purines, and provides further evidence that although DNA molecules may disperse SWNTs effectively in water, allowing the DNA further time to wrap around the nanotube effectively is required for adequate characterization of the samples. This provides a solid foundation upon which to select specific types of DNA for SWNT dispersion and, in particular, sample characterization. In particular, this suggests that, when using DNA sequences for nanotube sorting,35 enough time should be given to allow the chosen sequence to wrap the selected nanotubes before separation. Acknowledgment. J.N.C. acknowledgments SFI funding under the PI award scheme, Contract Number 07/IN.1/I1772. Supporting Information Available: Plot showing the zeta potential over time for each oligo-SWNT hybrid. This material is available free of charge via the Internet at http://pubs.acs.org. References and Notes (1) Hu, L.; Hecht, D. S.; Gruner, G. Appl. Phys. Lett. 2009, 94, 081103. (2) Koziol, K.; Vilatela, J.; Moisala, A.; Motta, M.; Cunniff, P.; Sennett, M.; Windle, A. Science 2007, 318, 1892. (3) Onoa, B.; Zheng, M.; Dresselhaus, M. S.; Diner, B. A. Phys. Status Solidi A 2006, 203, 1124. (4) Coleman, J. N. AdV. Funct. Mater. 2009, 19, 3680. (5) Bergin, S. D.; Nicolosi, V.; Cathcart, H.; Lotya, M.; Rickard, D.; Sun, Z.; Blau, W. J.; Coleman, J. N. J. Phys. Chem. C 2008, 112, 972. (6) Bachilo, S. M.; Strano, M. S.; Kittrell, C.; Hauge, R. H.; Smalley, R. E.; Weisman, R. B. Science 2002, 298, 2361. (7) O’Connell, M. J.; et al. Science 2002, 297, 593. (8) Moore, V. C.; Strano, M. S.; Haroz, E. H.; Hauge, R. H.; Smalley, R. E.; Schmidt, J.; Talmon, Y. Nano Lett. 2003, 3, 1379. (9) Bergin, S. D.; Nicolosi, V.; Giordani, S.; de Gromard, A.; Carpenter, L.; Blau, W. J.; Coleman, J. N. Nanotechnology 2007, 18, 455705. (10) Bergin, S. D.; et al. AdV. Mater. 2008, 20, 1876. (11) Furtado, C. A.; Kim, U. J.; Gutierrez, H. R.; Pan, L.; Dickey, E. C.; Eklund, P. C. J. Am. Chem. Soc. 2004, 126, 6095. (12) Giordani, S.; Bergin, S. D.; Nicolosi, V.; Lebedkin, S.; Kappes, M. M.; Blau, W. J.; Coleman, J. N. J. Phys. Chem. B 2006, 110, 15708. (13) Landi, B. J.; Ruf, H. J.; Worman, J. J.; Raffaelle, R. P. J. Phys. Chem. B 2004, 108, 17089.

Dispersion and Exfoliation of Nanotubes (14) Manchado, M. A. L.; Valentini, L.; Biagiotti, J.; Kenny, J. M. Carbon 2005, 43, 1499. (15) Steuerman, D. W. J. Phys. Chem. B 2002, 106, 3124. (16) Penicaud, A.; Poulin, P.; Derre, A. J. Am. Chem. Soc. 2005, 127, 8. (17) Amiran, J.; Nicolosi, V.; Bergin, S. D.; Khan, U.; Lyons, P. E.; Coleman, J. N. J. Phys. Chem. C 2008, 112, 3519. (18) Koshio, A.; Yudasaka, M. Nano Lett. 2001, 1, 361. (19) Dalton, A. B.; et al. AdV. Funct. Mater. 2004, 14, 1147. (20) Dieckmann, G. R.; et al. J. Am. Chem. Soc. 2003, 125, 1770. (21) Nicolosi, V.; Cathcart, H.; Dalton, A. R.; Aherne, D.; Dieckmann, G. R.; Coleman, J. N. Biomacromolecules 2008, 9, 598. (22) Ortiz-Acevedo, A.; Xie, H.; Zorbas, V.; Sampson, W. M.; Dalton, A. B.; Baughman, R. H.; Draper, R. K.; Musselman, I. H.; Dieckmann, G. R. J. Am. Chem. Soc. 2005, 127, 9512. (23) Nakashima, N.; Okuzono, S.; Murakami, H.; Nakai, T.; Yoshikawa, K. Chem. Lett. 2003, 32, 456. (24) Zheng, M.; Jagota, A.; Semke, E. D.; Diner, B. A.; McLean, R. S.; Lustig, S. R.; Richardson, R. E.; Tassi, N. G. Nat. Mater. 2003, 2, 338. (25) Cathcart, H.; Quinn, S.; Nicolosi, V.; Kelly, J. M.; Blau, W. J.; Coleman, J. N. J. Phys. Chem. C 2007, 111, 66. (26) Gigliotti, B.; Sakizzie, B.; Bethune, D. S.; Shelby, R. M.; Cha, J. N. Nano Lett. 2006, 6, 159. (27) Zheng, M.; et al. Science 2003, 302, 1545. (28) Cathcart, H.; Nicolosi, V.; Hughes, J. M.; Blau, W. J.; Kelly, J. M.; Quinn, S. J.; Coleman, J. N. J. Am. Chem. Soc. 2008, 130, 12734. (29) Daniel, S.; Rao, T. P.; Rao, K. S.; Rani, S. U.; Naidu, G. R. K.; Lee, H. Y.; Kawai, T. Sens. Actuators, B 2007, 122, 672. (30) Heller, D. A.; Baik, S.; Eurell, T. E.; Strano, M. S. AdV. Mater. 2005, 17, 2793. (31) Heller, D. A.; Jeng, E. S.; Yeung, T. K.; Martinez, B. M.; Moll, A. E.; Gastala, J. B.; Strano, M. S. Science 2006, 311, 508. (32) Hu, C.; Zhang, Y.; Bao, G.; Zhang, Y.; Liu, M.; Wang, Z. L. J. Phys. Chem. B 2005, 109, 20072. (33) Karachevtsev, V. A.; Glamazda, A. Y.; Leontiev, V. S.; Lytvyn, O. S.; Dettlaff-Weglikowska, U. Chem. Phys. Lett. 2007, 435, 104. (34) Arnold, M. S.; Stupp, S. I.; Hersam, M. C. Nano Lett. 2005, 5, 713.

J. Phys. Chem. C, Vol. 114, No. 27, 2010 11747 (35) Tu, X. M.; Manohar, S.; Jagota, A.; Zheng, M. Nature 2009, 460, 250. (36) Johnson, R. R.; Johnson, A. T. C.; Klein, M. L. Nano Lett. 2008, 8, 69. (37) Haggenmueller, R.; et al. Langmuir 2008, 24, 5070. (38) Wang, Y. X. J. Phys. Chem. C 2008, 112, 14297. (39) Hughes, M. E.; Brandin, E.; Golovchenko, J. A. Nano Lett. 2007, 7, 1191. (40) Gowtham, S.; Scheicher, R. H.; Pandey, R.; Karna, S. P.; Ahuja, R. Nanotechnology 2008, 19, 125701. (41) Enyashin, A. N.; Gemming, S.; Seifert, G. Nanotechnology 2007, 18, 245702. (42) Dukovic, G.; et al. J. Am. Chem. Soc. 2004, 126, 15269. (43) Zheng, M.; Diner, B. A. J. Am. Chem. Soc. 2004, 126, 15490. (44) O’Connell, M. J.; Eibergen, E. E.; Doorn, S. K. Nat. Mater. 2005, 4, 412. (45) Strano, M. S.; et al. J. Phys. Chem. B 2003, 107, 6979. (46) Rickard, D.; Giordani, S.; Blau, W.; Coleman, J. N. J. Lumin. 2008, 128, 31. (47) Israelachvili, J. Intermolecular and Surface Forces; Academic Press: London, 1991. (48) White, B.; Banerjee, S.; O’Brien, S.; Turro, N. J.; Herman, I. P. J. Phys. Chem. C 2007, 111, 13684. (49) Nair, N.; Usrey, M. L.; Kim, W. J.; Braatz, R. D.; Strano, M. S. Anal. Chem. 2006, 78, 7689. (50) Ryabenko, A. G.; Dorofeeva, T. V.; Zvereva, G. I. Carbon 2004, 42, 1523. (51) Choi, J. H.; Strano, M. S. Appl. Phys. Lett. 2007, 90, 3. (52) Overberger, C. G.; Peng, L. J. Polym. Sci., Part A: Polym. Chem. 1986, 24, 3219. (53) Tinoco, I. J. Am. Chem. Soc. 1960, 82, 4785. (54) Bloomfield, V. A.; Crothers, D. M.; Tinoco, I. Nucleic Acids: Structures, Properties and Functions; University Science Books: CA, 2000. (55) Meng, S.; Wang, W. L.; Maragakis, P.; Kaxiras, E. Nano Lett. 2007, 7, 2312. (56) Ke, C.; Humeniuk, M.; S-Gracz, H.; Marszalek, P. E. Phys. ReV. Lett. 2007, 99, 018302.

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