pubs.acs.org/Langmuir © 2010 American Chemical Society
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Aggregation of Fullerol C60(OH)24 Nanoparticles as Revealed Using Flow Field-Flow Fractionation and Atomic Force Microscopy Shoeleh Assemi,*,† Soheyl Tadjiki,‡ Bogdan C. Donose,§, Anh V. Nguyen,§ and Jan D. Miller† †
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Department of Metallurgical Engineering, University of Utah, 135 South 1460 East, Room 412, Salt Lake City, Utah 84112, ‡Postnova Analytics, 230 South 500 East, Suite 120, Salt Lake City, Utah 84102, and §School of Chemical Engineering, The University of Queensland, Brisbane, Queensland 4072, Australia. Present address. Advanced Water Management Center, The University of Queensland, Brisbane, Queensland 4072, Australia Received March 8, 2010. Revised Manuscript Received August 27, 2010 The effects of solution pH and 1:1 electrolyte concentration on the aggregation behavior of fullerol C60(OH)24 nanoparticles were investigated using flow field-flow fractionation (FlFFF). Particle separations were confirmed by examining FFF fractions using atomic force microscopy (AFM). Results showed that fullerol C60(OH)24 nanoparticles remain stable at low salt concentration (0.001 M NaCl) and basic pH (pH 10). Changing the pH did not affect the size significantly, but increasing the salt concentration promoted some aggregation. Fullerol C60(OH)24 nanoparticles did not form large clusters and reached a maximum size of at most several nanometers. Particle interaction analysis using the colloid interaction theory as described by the energetics of electrostatic repulsion and van der Waals attraction explained the differences in the colloidal stability of the fullerol C60(OH)24 nanoparticles under different solution conditions.
Introduction Size is the fundamental property that defines nanoparticles. It is speculated that in nanoparticles, surface properties can be more dominant than in the bulk material because of the relatively higher proportion of atoms at the particle surface.1 Also, because of the proportionally higher curvature of the surface, defects, edges, and catalytically active sites might be more exposed than in the bulk material.2 Physicochemical properties such as color, light-scattering properties, thermal behavior, solubility, and conductivity can be different from those in the bulk matter.3 These unique properties result in a wide range of size-specific applications that include drug delivery, energy storage, personal care materials, and environmental applications. The toxicity and transport of nanoparticles are also size- and shape-dependent.4-6 Understanding the changes in the size of nanoparticles from origin to destination is essential for any nanoparticle application or study on the toxicity, fate, and transport of nanomaterials.7-9 In this regard, size measurements and predictions of the aggregation state of nanoparticles have become of particular importance. The limitations of the employed technique, the effect of the preparation steps (e.g., drying), and assumptions made in the data interpretation should be considered carefully. Depending on the nature of the nanoparticle, several techniques can be used for size analysis, including optical spectroscopy, fluorescence polarization anisotropy, transmission electron microscopy (TEM), and light-scattering techniques.10 Other methods *Corresponding author. E-mail:
[email protected]. Phone: þ1 (801) 581 8604. Fax: þ1 (801) 581 4937.
(1) Waychunas, G. A. In Nanoparticles and the Environment, Banfield, J. F.; Navrotsky, A., Eds. Mineralogical Society of America: Washington DC, 2001, p105. (2) Madden, A. S.; Hochella, M. F. Geochim. Cosmochim. Acta 2005, 69, 389. (3) Hochella, M. F. Earth Planet. Sci. Lett. 2002, 203, 593. (4) Darlington, T. K.; et al. Environ. Toxicol. Chem. 2009, 28, 1191. (5) Handy, R. D.; et al. Ecotoxicology 2008, 17, 287. (6) Hauck, T. S.; et al. Small 2010, 6, 138. (7) Oberdorster, G.; et al. Part. Fibre Toxicol. 2005, 2, 8. (8) Donaldson, K.; et al. Occup. Environ. Med. 2001, 58, 211. (9) Borm, P.; et al. Toxicol. Sci. 2006, 90, 23. (10) Jones, C. F.; Grainger, D. W. Adv. Drug Delivery Rev. 2009, 61, 438.
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such as UV absorption11 or mass spectrometry12 can be used to monitor the aggregation or quantify the nanoparticles as well. Field-flow fractionation (FFF) is an elution technique that analyzes ensembles of the sample that have a similar property (e.g., diffusion coefficient or buoyant mass) and produces a size distribution rather than an average size. In FFF, the separation takes place in an open channel (versus a packed column in chromatography). More importantly, particle size fractions can be isolated from the distribution and examined for different purposes.13,14 FFF has been successfully used in the separation and characterization of natural organic matter,15,16 carbon nanotubes,17,18 nanoparticles isolated from cells and tissue,19 and the analysis of DNA-nanoparticle complexes.20 Fullerols (C60(OH)x), the hydroxylated forms of fullerenes, are water-soluble, spherical in shape, and about 1.2 nm in size. Fullerols are available commercially, and some data on their size and aggregation is readily available for reference. Fullerols have several clinical applications as drug carriers or tumor inhibitors,21-23 and their toxicity is still undecided.24,25 Information on the solubility and aggregation behavior of fullerols as important fullerene derivatives can help in establishing new applications and advanced studies on their toxicity and environmental impact. There are limited studies on the aggregation of fullerol nanoparticles. Lecoanet et al.26 found that fullerols did not aggregate (11) (12) (13) (14) (15) (16) (17) (18) (19) (20) 549. (21) (22) (23) (24) (25) (26)
Vileno, B.; et al. Adv. Funct. Mater 2006, 16, 120. Chithrani, B. D.; Ghazani, A. A.; Chan, W. C. W. Nano Lett. 2006, 6, 662. Beckett, R. At. Spect. 1991, 12, 228. Ranville, J. F.; et al. Anal. Chim. Acta 1999, 381, 315. Beckett, R.; Zhang, J; Giddings, C. Environ. Sci. Technol. 1987, 21, 289. Lead, J. R.; et al. Environ. Sci. Technol. 2000, 34, 3508. Liu, J.; et al. Science 1998, 280, 1253. Moon, M. H.; et al. J. Sep. Sci. Technol. 2004, 27, 710. Tadjiki, S.; et al. J. Nanopart. Res. 2008, 11, 981. Ma, P. M.; Buschmann, M. D.; Winnik, F. M. Biomacromolecules 2010, 11, Jiadan, Z.; et al. Small 2008, 4, 1168. Prato, M. J. Mater. Chem. 1997, 7, 1097. Sitharaman, B.; et al. Nano Lett. 2004, 4, 2373. Roberts, J. E.; et al. Toxicol. Appl. Pharmacol. 2008, 228, 49. Gelderman, M. P.; et al. Int. J. Nanomed. 2008, 3, 59. Lecoanet, H; Weisner, M. R. Environ. Sci. Technol. 2004, 38, 4377.
Published on Web 09/17/2010
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and were very mobile in a column of glass beads when compared to C60 fullerenes. No size or charge data was reported for the fullerols used. Brant et al.27 reported an average size of about 100 nm for the same fullerol sample using DLS and TEM. The size measured for water-soluble fullerols was even larger than for the hydrophobic fullerene clusters, indicating that fullerols might in fact be hydrophobic and might stay in a supramolecular state even at low concentrations. Similar controversial findings about the size of fullerene derivatives have been reported in the literature.23 We report the separation and characterization of fullerol C60(OH)24 nanoparticles in solutions of varying pH and salt concentration, using flow field-flow fractionation (FlFFF) along with a validation of the separation by imaging with atomic force microscopy. Interactions between nanoparticles will be discussed in terms of DLVO theory.
Materials and Methods Samples. Fullerol C60(OH)24 nanoparticles were obtained in powder form from the MER Corporation (Tucson, AZ) and were used as received. According to the manufacturer, the fullerols were synthesized from 99.99% fullerene C60 in dry form using bromination followed by addition of NaOH. The purity of the sample (as deposited on an alumina optical window) was verified using energydispersive X-ray spectroscopy (FEI Novanano, Hilsboro, OR). Only signals for carbon, oxygen, sodium, and aluminum were obtained, confirming the absence of impurities in the fullerols. Samples were prepared by dissolving the powder in the electrolyte of choice with a concentration of 1.3 mg mL-1 and were used without filtration. Millipore double-deionized water with a resistivity of >18 MΩ 3 cm was used in all experiments. All glassware was soaked in a solution of 14% KOH, 75% ethanol, and 11% deionized water overnight and washed with copious amounts of deionized water. The plasticware was soaked in 1% detergent overnight and rinsed with deionized water/ethanol/acetone/water. Electrophoretic Mobility Measurements. Electrophoretic mobilities of the fullerol nanoparticles were measured using phase analysis light scattering (Zeta PALS, Brookhaven Corporation, Holtsville, NY). Samples were prepared by dispersing the powdered particles in the electrolyte under the desired conditions and equilibrating overnight. Briefly, a solution of about 0.8 mg 3 mL-1 fullerol at the desired ionic strength was prepared. The solution had a pH of about 10, which was changed by the addition of 0.1 M HCl. The final concentration of HCl in the 5 mL solution was less than 1 10-3 M, which did not change the ionic strength of the solution significantly. Zeta potential values were calculated from the measured electrophoretic mobilities using H€ uckel’s equation for the case of nanoparticles (κa , 1).28 Flow Field-Flow Fractionation (FlFFF). A complete description of FFF techniques can be found in publications by Giddings29 and Giddings and Caldwell.29,30 In flow FFF (FlFFF), the applied field is a crossflow and the measured parameter is the diffusion coefficient. The lower frit (the accumulation wall) is covered with a membrane to prevent sample loss. The separation is thus the result of the interaction between a field (cross flow) that is applied perpendicular to the stream carrying the sample, and the sample species. In asymmetrical FlFFF, the channel is trapezoidal (Figure 1) and the upper frit is replaced by a poly(methylmethacrylate) (PMMA) plate. The inlet flow is divided into two parts; the field, which goes out through the lower frit, and the outlet flow stream. Similar to other FFF techniques, separation is achieved by the interaction of the sample particles with the applied field and their Brownian motion. As a result, smaller particles with a higher (27) Brant, J. A.; et al. J. Colloid Interface Sci. 2007, 314, 281. (28) Hunter, R. J. Foundations of Colloid Science; Oxford University Press: New York, 1987; Vol. 1, p 673. (29) Giddings, J. C. Chem. Eng. News 1988, 66, 34. (30) Giddings, J. C.; Caldwell, K. D. In Physical Methods of Chemistry; Rositer, B. W., Hamilton, J. F., Eds.; John Wiley & Sons: New York, 1989; p 867.
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Figure 1. Schematic representation of separation in an asymmetrical flow FFF channel. The field, applied perpendicular to the channel, pushes the sample species toward the lower wall (accumulation wall). The sample will equilibrate under the applied field for a certain time. The sample species form clouds with a center of mass depending on the diffusion coefficient. Smaller samples with larger diffusion coefficients form clouds with a center of mass closer to the center of the channel. The elution starts after sample equilibration. Smaller sample components will elute with the faster component of the parabolic flow, and the larger ones elute later. diffusion coefficient will form clouds with a center of mass closer to the middle of the channel and will elute with the faster component of the laminar flow (Figure 1). In asymmetrical FlFFF, when a constant field is applied, the hydrodynamic diameter of the particle (dh) can be calculated directly from the retention time (tr) without calibration: 2kTtr
dh ¼
πηw2 ln 1 þ a
Vc
!
ð1Þ
V3
where k is the Boltzmann constant, T is the absolute temperature, tr is the retention time, η is the viscosity of the carrier fluid at the given temperature, w is the channel thickness, V_ is the channel flow rate, Vc is the cross-flow rate of the carrier fluid, and a is the geometric factor of the channel.31 The diffusion coefficient (Dh) corresponding to each size (dh) can be calculated from the Stokes-Einstein equation:28 Dh ¼
kT 3πηdh
ð2Þ
The FlFFF experiments were performed using a Postnova Analytics AF2000 FOCUS asymmetrical flow field-flow fractionation instrument with UV detection (absorption) at 254 nm (Postnova Analytics, Salt Lake City, UT). A constant field of 3 mL 3 min-1 and an injection volume of 0.2 mL were used in all experiments. The channel void volume was measured as 1 mL using bovine serum albumin.32 Samples were introduced into the channel via a Rheodyne injection system. The channel was flushed with the carrier solution after each run, and the response of the detector was recorded. An observed peak would indicate physical adsorption of the sample to the membrane. Such a peak was not observed for fullerol samples. The fractograms (UV signal vs retention time) were converted to a size distribution using the Postnova FFF Analysis software. Atomic Force Microscopy. Fractions obtained from FFF size distributions were examined by AFM imaging performed in air and in situ. Samples were imaged in AC mode (tapping mode) with an Asylum Research MFP-3D AFM mounted on a Herzan (31) Schimpf, M.; Caldwell, K.; Giddings, J. C., Eds. Field-Flow Fractionation Handbook, John Wiley & Sons: New York, 2000. (32) Giddings, J. C.; Benincasa, A. M.; Williams, P. S. J. Chromatogr. 1992, 627, 23.
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anti-vibration table, placed in a TMC soundproof enclosure. Gold-coated NT-MDT HA/NC (Etalon) cantilevers with a nominal spring constant of 3.4 N/m and a tip diameter of ∼8 nm (reported by the manufacturer) were used for imaging. The integrity of the tips was checked using Tipcheck (Budgetsensors, Bulgaria), which produces reverse images of the apex of the tip, and by obtaining force curves between the tip and the uncoated HOPG substrate before each experiment. Prior to imaging, 10 μL of the fullerol suspension was placed on freshly cleaved highly oriented pyrolytic graphite (SPI, grade I). We allowed 2 h for the adsorption of nanoparticles on the graphite substrate. For images obtained in air, the liquid was removed with a stream of highpurity nitrogen in a laminar flow cabinet and the images were obtained immediately in air (humidity maintained at 50%) under semi-moist conditions. Typically, images were acquired at an 800 mV amplitude set point with a scan rate between 0.3 and 0.5 Hz. The images were processed using Asylum Research MFP-3D software. The experiments were repeated on three different sets of samples. For images obtained in situ, the solution was replaced with a blank electrolyte (no fullerol) after 2 h of adsorption time. Interaction Energy Profiles. The stability of colloidal particles can usually be described as the sum of attractive van der Waals and repulsive electrical double-layer forces by the Derjaguin-Landau-Verwey-Overbeek (DLVO) theory. The interaction energy between two curved particles is commonly calculated from the corresponding interaction energy per unit area between parallel surfaces using the Derjaguin approximation,33 which relates the force between two spherical particles to the energy per unit area between two flat parallel plates E(D). The Derjaguin approximation is valid as long as the interaction range and the separation distance, D, between the parallel surfaces are much smaller than the particle radius (R). Therefore, for nanoparticles, this approach tends to overestimate the interaction energy.34,35 The most accurate approach to predicting the nanoparticle interactions would involve direct numerical solutions of the equations governing the van der Waals and electrical double-layer interactions between the nanoparticles. However, this numerical approach would be tedious for the current analysis. Therefore, an alternative SEI (surface element integration) method34 was employed. The method improves on the Derjaguin approximation by replacing infinity in the integration of the Derjaguin approximation by a finite upper limit, as confirmed by excellent agreement with the numerical solutions.34 Using the approach outlined by Bhattacharjee and Elimelech,34 we solved for the total interaction energy, ET, between two spheres with radius R as Z ET ðhÞ ¼ 2π
R
h
pffiffiffiffiffiffiffiffiffiffiffiffiffiffiffi pffiffiffiffiffiffiffiffiffiffiffiffiffiffiffi i Eðh þ 2R - 2 R2 - r2 Þ - Eðh þ 2R þ 2 R2 - r2 Þ r dr
0
ð3Þ EðDÞ ¼ -
A 2εε0 Kζ2 þ 2 12πD expðKDÞ - 1
ð4Þ
where ζ is the surface potential of single (noninteracting) nanoparticles, κ is the inverse Debye length, h is the separation distance, ε is the permittivity of the water, ε0 is the permittivity of a vacuum, and A is the Hamaker constant. D = h þ 2R - 2(R2 - r2)1/2 is the distance between the surface elements and r is the radial distance from the surface elements to the axis of symmetry (the center-to-center line). The surface potential was assumed to be equal to the zeta potential of fullerol nanoparticles. Equation 4 describes the (van der Waals and double-layer) interaction energy per unit area between two parallel surface elements. The second term on the right-hand side of eq 4 describes the double-layer interaction energy between parallel (33) Israelachvili, J. N. Intermolecular and Surface Forces; Academic Press: London, 1992. (34) Bhattacharjee, S.; Elimelech., M. J. Colloid Interface Sci. 1997, 193, 273. (35) Todd, B. A.; Eppel., S. J. Langmuir 2004, 20, 4892. (36) Parsegian, V. A.; Gingell, D. Biophys. J. 1972, 12, 1192.
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Figure 2. Zeta potential of fullerol nanoparticles at different concentrations of NaCl and 0.001 M NaNO3. The zeta potentials were calculated from the measured electrophoretic mobilities using H€ uckel’s equation. surface elements,36 and its disjoining pressure, (εε0 κ2/2)/sinh2(κD/2), which in the limit of small κD asymptotically approaches 1/D2 as in the case of the Langmuir equation.37 Equation 3, presenting an extension of the SEI for two spheres, was numerically integrated using a VBA (Visual Basic for Application) macro written in Microsoft Excel. Because there are no Hamaker constants available for fullerol nanoparticles, an approximate nonretarded Hamaker constant of 1.51 10-19 J was calculated using the Lifshitz-van der Waals constant for diamond-diamond in water (3.95 eV).38 This value is very close to the Hamaker constant calculated for carbon black dispersions in water (1.15 10-19 J),39 experimental values for carbon black ((0.5-1.0) 10-19 J),40 and theoretical calculations for diamond in water (1.3 19-19 J).41
Results and Discussion Electrophoretic Mobility Measurements. The zeta potentials of fullerol nanoparticles in solutions of different pH and salt concentrations were calculated from the measured electrophoretic mobilities, and the results are presented in Figure 2. The nanoparticles remained negatively charged even at very low pH, and the zeta potential did not change significantly via changes in solution pH. To investigate the effect of chloride ion adsorption on the surface charge of fullerols, we measured the electrophoretic mobility of the fullerols in 0.001 M NaNO3. The results (Figure 2) showed that the zeta potential of the fullerols was independent of the salt type and is perhaps the result of the fullerol structure. Potentiometric titration (Supporting Information) demonstrated that there were available sites for proton adsorption even up to pH 3, confirming the electrokinetic data showing that the fullerols are negatively charged regardless of the pH. Electrokinetic and titration data by other researchers11,27 have shown similar results (negative charge regardless of pH) and a multiple number of regions of high proton affinity ranging from pH∼9 to ∼3.6. These results suggest that several different sites for proton adsorption should be available on the fullerol surface. At least two different types of hydroxyl groups can exist because of the existence of pentagons and hexagons in the C60 structure.42 Different methods of synthesis can also break the fullerol cage and result in the production of hemiketals. According to the manufacturer, the dry synthesis from pure (99.99%) C60, through bromination, results in a final product of C60(OH)x(ONa)y, with x þ y = 24. Therefore, some oxide groups can contribute to the negative charge as well. (37) Langmuir, I. J. Chem. Phys. 1938, 6, 873. (38) Butt, H.-J., Graf., K., Kappl, M., Eds. Physics and Chemistry of Interfaces; Wiley-VCH: Weinheim, Germany, 2003. (39) Dagastine, R. R.; Prieve, D. D.; White, L. R. J. Colloid Interface Sci. 2002, 249, 78. (40) Hartley, P. A.; Parfitt, G. D. Langmuir 1985, 1, 651. (41) Fernandez-Varea, J. M.; Garcia-Molina, R. J. Colloid Interface Sci. 2000, 231, 394. (42) Taylor, R. The Chemistry of Fullerenes; World Scientific Publishing: Singapore, 1995.
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Fullerol C60(OH)24 Size Analysis. Figure 3a,b shows representative size distributions of fullerol nanoparticles for different NaCl concentrations obtained at pH ∼10 (Figure 3a) and pH ∼6.6 (Figure 3b). The diameter at the peak maximum and the corresponding diffusion coefficient for each run condition are presented in Table 1. The size of fullerol nanoparticles at the peak maximum varied from about 1.8 nm (0.001 M NaCl) to 6.7 nm (0.1 M NaCl). If the size of one fullerol nanoparticle is considered to be about 1.2 nm, then these results mean that the analyzed fullerol nanoparticles were perhaps in the form of doublets or larger. The increase in the salt concentration resulted in a shift (toward the larger size) in the peak maximum and a broadening of the size distribution. A 100-fold increase in the ionic strength resulted in about a 60% increase in the size of the nanoparticles at the peak maximum. The effect of the ionic strength seems to become more prominent at lower pH (Figure 3b,d), where more surface hydroxyl groups are neutralized. The representative size distributions of fullerol nanoparticles in acidic, basic, and neutral pH obtained in 0.001 M NaCl are presented in Figure 4. Changing the pH from 10 to 4 did not change the size distribution of fullerols significantly. These results are consistent with the electrokinetic measurements that showed a minimal change in the zeta potential by changing the pH, demonstrating that the fullerol nanoparticles were most stable at pH ∼10 and low salt concentration (0.001 M NaCl). However, even at acidic pH and high ionic strength, the agglomerates were not larger than several nanometers. Figure 3c,d shows the cumulative size distribution of the fullerol samples in different ionic strengths and solution pH values of 10 (Figure 3c) and 6.6 (Figure 3d). Increasing the ionic strength increased the median size (50% population) by 52-65%. Changing the pH changes the median size between 28 and 58% overall, depending on the ionic strength. We were not able to obtain a peak at pH 4 and ionic strengths higher than 0.001 M NaCl. Because pH 4 is very close to the point of zero charge (pzc) of the membrane covering the accumulation
Figure 3. Size distribution of fullerol C60(OH)24 nanoparticles obtained in different NaCl concentrations at pH 10 (a) and pH 6.6 (b) and cumulative size distributions at pH 10 (c) and pH 6.6 (d). The size distributions were obtained from the fractograms using eq 1.
Figure 4. Effect of pH on the size distribution of fullerol C60(OH)24 nanoparticles at an ionic strength of 0.001 M NaCl. The size distributions were obtained from the fractograms using eq 1.
Table 1. Comparison of the Diameter Obtained at the Peak Maxima and the Corresponding Diffusion Coefficients of Fullerol C60(OH)24 Nanoparticles Obtained by FlFFF at Different Solution pH and Ionic Strength (I)a pH ∼4 I (M)
dp (nm)
Dp (10-10 m2 3 s-1)
pH ∼7 dp (nm)
Dp (10-10 m2 3 s-1)
pH ∼10 dp (nm)
Dp (10-10 m2 3 s-1)
0.001 2.1 21 2.3 19 1.8 24 0.01 4.7 9.3 3.3 13 0.1 6.7 6.5 4.9 8.9 a dp is the diameter at the peak maximum (nm), and Dp is the diffusion coefficient at the peak maximum (10-10 m2 s-1) calculated from eq 2.
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wall, fullerol-membrane interactions are expected at higher ionic strengths when the thicknesses of the diffuse double layers are significantly reduced and other interparticle forces such as van der Waals or hydrophobic forces between the particles/aggregates and the membrane are able to dominate. Membrane-sample interaction at a solution pH close to the pzc of the membrane is a limitation of the FlFFF system and should be considered when designing experiments. To determine whether the obtained fractograms represented the entire injected sample, after each run the channel was flushed with the carrier and with the field off. No peak was observed, indicating that under most conditions the fullerol-membrane interaction was not significant. Furthermore, the area under the fractograms did not change from one condition to another, again indicating that the membrane-fullerol interaction was minimal under the employed experimental conditions. Excellent recovery was observed for all run conditions, except for the two conditions at pH 4 (0.01 and 0.1 M NaCl). Verification of Fullerol Size Separation. To verify the FlFFF results, fractions obtained from the peak maxima of the fractograms were examined by atomic force microscopy. Our attempts to obtain TEM electrographs of the fractions were unsuccessful, perhaps because of the destruction of the nanoparticles by the electron beam or the lack of contrast between the very small fullerol nanoparticles and the carbon grid. In AFM, the lateral resolution is limited by the number of pixels and the tip geometry but the height measurement can provide valuable information about the sample size. Figure 5 shows AC mode (tapping) images of the fractions obtained from the peak maxima of the fractograms at NaCl concentrations of 0.001 M (Figure 5a) and 0.01 M (Figure 5b) under moist conditions (where the electrolyte was blown with nitrogen and images were obtained immediately in air; see the Materials and Methods section). Images of the fractions could not be obtained at 0.1 M NaCl because of the presence of the salt crystals. Section analyses presented under each image in Figure 5 show the height of the adsorbed particles. The peak-to-valley measurements showed an average height of about 2.5 nm for the fraction obtained at 0.001 M NaCl and about 3.5 nm for the fraction obtained at 0.01 M NaCl. Imaging in air can induce aggregation due to partial drying. Besides, as mentioned before, obtaining images in high salt concentrations was almost impossible because of the crystallization of the electrolyte. Therefore, we obtained in situ AFM images from the fractions isolated from the peak maxima of fullerol size distributions at different NaCl concentrations (Figure 6). The height data corresponded very well to the peak diameter obtained from FlFFF and increased with an increase in ionic strength. Compared to the number of particles obtained in the air, the number of fullerol nanoparticles captured on the graphite surface was much less. Several larger-scale scans were needed to locate the fullerol nanoparticles, which were usually found at the atomic steps of the graphite surface. The lateral adsorption of fullerol aggregates on the HOPG surface is a feature induced by the interactions between the HOPG surface and fullerol aggregates and should be considered separately because it does not reflect the fullerol aggregation state in solution. Therefore, only the vertical size can give an indication of the real diameter of the aggregates as they are deposited on graphite from the solution. However, the deposition of fullerol aggregates on HOPG will be influenced by the solution conditions. As the ionic strength increases, the decrease in the thickness of the diffuse double layer should result in an increase in the interaction between the hydrophilic fullerol nanoparticles and the hydrophobic HOPG surface, which can be observed as an increase in the base size of the aggregates (Figure 6). Langmuir 2010, 26(20), 16063–16070
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Figure 5. AFM images of the fraction taken from the peak maximum of the fullerol size distribution in 0.001 N (a) and 0.01 M NaCl (b) under semi-moist conditions. Imaging at 0.1 M NaCl was hindered by the presence of salt crystals.
Our AFM results confirm the FlFFF data that fullerol C60(OH)24 nanoparticles can exist as very small clusters (doublets or triplets) at basic pH and low salt concentration. This is in contrast to some lightscattering data that reports sizes on the order of 100 nanometers for fullerol nanoparticles.27,43 In light scattering, the intensity of the scattered light varies with the sixth power of the diameter. Therefore, (43) Chen, K. L.; Elimelech, M. Langmuir 2006, 22, 10994.
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Figure 6. AFM images, of the fractions taken from the peak maximum of the C60(OH)24 fullerol size distribution in the electrolyte. 0.001 M NaCl (a), 0.01 M NaCl (b), and 0.1 M NaCl (c). In parts a-c, we show a large-scale height image (i), a smaller-scale height image that is an enlarged image of the particles (ii), a 3D image showing the C60(OH)24 particles on graphite (iii), the height of the particles from the section analysis of the image (iv).
the presence of aggregates, dust, or other particles in the solution can produce a significant variation in the results. Because FFF is a separation technique, such problems are eliminated because of the fact that 16068 DOI: 10.1021/la102942b
the impurities will not be retained and will be eluted at the beginning of the separating (void peak) and the detector always measures the signal of a monodispersed fraction eluting from the channel. Langmuir 2010, 26(20), 16063–16070
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Figure 7. Energy profiles for the interaction of two fullerol nanoparticles at different pH values (4-10) and salt concentration (0.001-0.1 M), as calculated from eq 3. R(particle radius) = 1 nm and A = 1.59-19 J. The zeta potentials (Figure 2) were assumed to be equal to the surface potentials.
Interaction Energy Profile for Fullerol Nanoparticles. Colloidal stability in aqueous media can be described by the sum of repulsive electrical double layer and attractive van der Waals forces as exemplified by the DLVO theory. Typically, aggregation should occur if the particles collide with sufficient kinetic energy to overcome the energy barrier and reach the primary minimum. The energy profiles were constructed as described in the Materials and Methods section. Our FlFFF results suggested that the fullerol C60(OH)24 nanoparticles existed in their smallest size as primary particles at basic pH and low ionic strength. Therefore, the size measured by FFF at 0.001 M NaCl and pH 10 was used for the calculations. Figure 7 shows interaction energy profiles between two fullerol nanoparticles (R = 1 nm) for dispersions at different pH values and ionic strengths. The height of the energy barriers decreased upon increasing the ionic strength and reached to below zero at 0.1 M NaCl. The height of the barriers is smaller than those reported for larger nanoparticles using classical DLVO,44 which could be due to several factors, including the difference in the model used. However, the trends do agree with those obtained from FlFFF experiments. For a given salt concentration, the interaction seemed not to be affected by a change in pH. The net interaction was repulsive for 0.001 M NaCl and 0.01 M NaCl at all pH values, suggesting that under these conditions the fullerols should remain stable. In fact, (44) Behrens, S. H.; Borkoves, M.; Schurtenberger, P. Langmuir 1998, 14, 1951.
Langmuir 2010, 26(20), 16063–16070
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minimal aggregation could be observed under these conditions (Figure 3). However, increasing the salt concentration reduced the decay length, and at 0.1 M NaCl, the net interaction became attractive, suggesting aggregation. FlFFF experiments showed a slight increase in the size of fullerol nanoparticles at pH 4 and in 0.001 M NaCl. No peaks could be observed at higher ionic strengths. The analysis suggests that the interactions between fullerol nanoparticles are governed by electrical double-layer forces, and the size of the fullerol aggregates might be tuned by changing the solution pH and ionic strength. Although the colloid interaction model presented here seems to agree with the FlFFF results based on the variables, future research should make a better estimate of the Hamaker constant for fullerol nanoparticles better. Our results demonstrate that (a) aggregation of the fullerol nanoparticles is suppressed by low ionic strength and basic pH because of electrostatic repulsion; (b) changing pH does not have a significant effect on the aggregation state of fullerol nanoparticles; and (c) fullerol nanoparticles do not form large clusters in solution at natural and basic pH values and with moderate or low salt concentration. At low and moderate ionic strengths and basic pH, aggregation is prevented by electrostatic repulsion. As discussed with regard to the EPM measurements, fullerol nanoparticles are in fact stable even at slightly acidic pH. At higher ionic strength, aggregation is promoted when the electrical double layer is significantly compressed. The hydroxyl groups on fullerol C60(OH)24 nanoparticles help disperse the fullerols perhaps by hydrogen bonding with the water molecules and possible shielding of the short-range hydrophobic forces. If the hydroxyl group is attached to a six-membered ring on the fullerol surface, then it can act as a phenol group and become negatively charged by losing the proton. The generated phenoxide ion can be stabilized as a result of charge delocalization, induced by the π bonds in the six-membered rings of the fullerol nanoparticle. This can explain the minimal change in the aggregation state of fullerol nanoparticles in response to changes in solution pH.
Conclusions Size distributions of fullerol nanoparticles are reported in aqueous solutions with varying pH and ionic strength using FlFFF. The particle size distribution and state of aggregation were verified by AFM. Our results show that the fullerols are in fact very small and exist in clusters of only a few nanometers in size. Fullerols can remain stable in aqueous solutions at basic pH and an ionic strength of 0.001 M NaCl or lower. Interactionenergy profiles for the fullerol nanoparticles, using surface element integration and an estimate of the Hamaker constant, matched fairly well with the experimental results obtained from FlFFF and atomic force microscopy. We demonstrated the utility of the FFF separation technique for the study of the aggregation of hydrophilic nanoparticles using fullerols as a model. This method eliminates measurement errors due to the interference between particles of different sizes. Our results show that fullerols are not large clusters with sizes over hundreds of nanometers but rather exist in the nanometer size range even at low pH . Our results also illustrate that by using basic parameters such as solution conditions and the zeta potential it was possible to predict the aggregation behavior and tune the size of fullerol nanoparticles. A subsequent study using this methodology will consider the stability of fullerene nanoparticles. The findings of this research may be useful in the nanoparticle DOI: 10.1021/la102942b
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industry, especially in biomedical and drug delivery research and in studies on toxicology, including the fate and transport of hydrophilic nanoparticles. Acknowledgment. This research was partially funded by grant no. 0227583 from the National Science Foundation and grant DEFG-03-93-ER14315 from DOE Basic Sciences, and by grant no.
16070 DOI: 10.1021/la102942b
Assemi et al.
DP0663688 from the Australian Research Council. We are grateful to four anonymous reviewers for their very helpful comments. Supporting Information Available: Potentiometric titration of fullerols as a function of acid concentration in 0.001 M NaCl. This material is available free of charge via the Internet at http://pubs.acs.org.
Langmuir 2010, 26(20), 16063–16070
