Single Molecule Imaging of Protein Molecules in Nanopores

Dec 15, 2009 - For a given protein, the number of molecules found at a given depth decreased as the pore size decreased. In addition, as the depth inc...
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Anal. Chem. 2010, 82, 478–482

Letters to Analytical Chemistry Single Molecule Imaging of Protein Molecules in Nanopores Changbei Ma and Edward S. Yeung* Ames Laboratory-U.S. DOE and Department of Chemistry, Iowa State University, Ames, Iowa 50011 The interactions between single protein molecules and nanoporous polycarbonate membranes were investigated at the single molecule level. Entrapment of proteins was shown to be size selective and was dependent on the membrane pore diameter. A pore size that is only slightly larger than the maximum dimension of the proteins was inadequate for intrusion into the pores. For a given protein, the number of molecules found at a given depth decreased as the pore size decreased. In addition, as the depth increased, for a given size pore, the number of molecules decreased rapidly. The depth-dependent histograms nicely fit a one-dimensional diffusion model. However, a highly restricted motion was observed even when the pore diameter was 10 times the size of the protein, resulting in anomalously small diffusion coefficients. We also demonstrated the subtle differences in depth distribution among BSA and hemoglobin that have nearly the same molecular weight but slightly different molecular shapes. These results give unique insights into the detailed mechanism of size-exclusion chromatography and membrane filtration. Efficient methods for separating and purifying proteins from a complex mixture are of the utmost importance in biology and biomedical engineering. There are a number of different liquid chromatography methods that are commonly used for purification. Currently, nucleic acids, proteins, and industrial polymers are routinely separated by size exclusion chromatography (SEC) on the basis of molecular size in solution.1 This is achieved with a porous packing material that is compatible with the mobile phase. The smallest components in the sample migrate into the smallest pores of the packing while the molecular dimensions of the highermolecular-weight components prevent them from penetrating into the pores. Therefore, the largest molecules will elute first and the smallest molecules will elute last. However, SEC pores in commercial chromatographic materials are highly heterogeneous in size and shape, making it difficult to elucidate the exact contributions of pore size to retention time. There is also increasing interest in using synthetic nanoporous membranes which are extremely homogeneous to achieve protein * To whom correspondence should be addressed. E-mail: yeung@ ameslab.gov. (1) Striegel, A. M. Anal. Chem. 2005, 77, 104A–113A.

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separations if the proteins have significantly different molecular sizes.2-4 Nanoporous membranes capable of selectively separating molecules are interesting targets both in fundamental research and for real applications.5 Potential advantages of membrane-based protein separations include low cost, high speed, and high throughput. In addition, membrane-based separations can be easily scaled up for use in large-scale commercial production.2 In particular, the availability of membranes containing nanoscale cylindrical pores with well-defined structures and monodisperse sizes have made it possible to correlate the pore size with the separation performance.6,7 Membranes that have been employed for separating proteins include nanoporous anodic alumina membranes,8-10 polycarbonate track-etched membranes,11-16 gold nanotubule membranes,17-19 carbon nanotube membranes,20,21 and ultrathin nanoporous silicon membranes.22 For example, Yu et al. demonstrated that by controlling the diameters of the pores, Au nanotubule membranes can show good selectivity for the (2) (3) (4) (5) (6) (7) (8) (9) (10) (11) (12) (13) (14) (15) (16) (17) (18) (19) (20) (21) (22)

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separation of proteins on the basis of molecular size.17 More recently, Striemer et al. separated two differently sized proteins using ultrathin silicon membranes under physiological conditions.22 Understanding molecular motions inside the nanopores is essential for developing size-selective separations based on these membranes. So far, the three-dimensional dynamical behavior of proteins in these membranes at the single-molecule level has not been directly imaged in real time. Single-molecule spectroscopy has proven itself to be a powerful tool in chemistry and the biosciences.23 Observation of single molecules allows their dynamic behaviors to be studied and provides insight into molecular genetics,24-26 biochip assembly,27-29 biosensor design,30-32 DNA biophysics,33-42 enzyme dynamics,43-47 DNA/protein detection,48-50 and basic separation theory.51-63 In (23) Moerner, W. E. Proc. Natl. Acad. Sci. U.S.A. 2007, 104, 12596–12602. (24) Herrick, J.; Michalet, X.; Conti, C.; Schurra, C.; Bensimon, A. Proc. Natl. Acad. Sci. U.S.A. 2000, 97, 222–227. (25) Herrick, J.; Bensimon, A. Chromosome Res. 1999, 7, 409–423. (26) Lyubchenko, Y. L.; Shlyakhtenko, L. S. Proc. Natl. Acad. Sci. U.S.A. 1997, 94, 496–501. (27) Hill, E. K.; De Mello, A. J. Analyst 2000, 125, 1033–1036. (28) Turner, S. W. P.; Levene, M.; Korlach, J.; Webb, W. W.; Craighead, H. G. Proceedings of the Micro Total Analysis System, Monterey, CA, October 2125, 2001; pp 259-261. (29) Shivashankar, G. V.; Libchaber, A. Curr. Sci. 1999, 76, 813–818. (30) Chan, V.; Mckenzie, S. E.; Surrey, S.; Fortina, P.; Graves, D. J. J. Colloid Interface Sci. 1998, 203, 197–207. (31) Chan, V.; Graves, D. J.; Fortina, P.; McKenzie, S. E. Langmuir 1997, 13, 320–329. (32) Jordan, C. E.; Frutos, A. G.; Thiel, A. J.; Corn, R. M. Anal. Chem. 1997, 69, 4939–4947. (33) Bensimon, A.; Simon, A.; Chiffaudel, A.; Croquette, V.; Heslot, F.; Bensimon, D. Science 1994, 265, 2096–2098. (34) Xue, Q.; Yeung, E. S. Nature 1995, 373, 681–683. (35) Houseal, T. W.; Bustamante, C.; Stump, R. F.; Maestre, M. F. Biophys. J. 1989, 56, 507–516. (36) Fan, F.-R. F.; Bard, A. J. Science 1995, 267, 871–874. (37) Funatsu, T.; Harada, Y.; Tokunaga, M.; Saito, K.; Yanagida, T. Nature 1995, 374, 555–559. (38) Chiu, D. T.; Zare, R. N. J. Am. Chem. Soc. 1996, 118, 6512–6513. (39) Nie, S.; Chiu, D. T.; Zare, R. N. Science 1994, 266, 1018–1021. (40) Xu, X.; Yeung, E. S. Science 1997, 275, 1106–1109. (41) Dickson, R. M.; Norris, D. J.; Tzeng, Y.-L.; Moerner, W. E. Science 1996, 274, 966–969. (42) Ma, Y.; Shortreed, M. R.; Yeung, E. S. Anal. Chem. 2000, 72, 4640–4645. (43) Kettling, U.; Koltermann, A.; Schwille, P.; Eigen, M. Proc. Natl. Acad. Sci. U.S.A. 1998, 95, 1416–1420. (44) Li, H. W.; Yeung, E. S. Anal. Chem. 2005, 77, 4374–4377. (45) Lu, H. P.; Xun, L. Y.; Xie, X. S. Science 1998, 282, 1877–1882. (46) Gorris, H. H.; Rissin, D. M.; Walt, D. R. Proc. Natl. Acad. Sci. U.S.A. 2007, 104, 17680–17685. (47) Li, J. W.; Yeung, E. S. Anal. Chem. 2008, 80, 8509–8513. (48) Li, Z. H.; Hayman, R. B.; Walt, D. R. J. Am. Chem. Soc. 2008, 130, 12622– 12623. (49) Lee, J. Y.; Li, J. W.; Yeung, E. S. Anal. Chem. 2007, 79, 8083–8089. (50) Li, J. W.; Xie, W. J.; Fang, N.; Yeung, E. S. Anal. Bioanal. Chem. 2009, 394, 489–497. (51) Xu, X.-H.; Yeung, E. S. Science 1998, 281, 1650–1653. (52) Shortreed, M. R.; Li, H.; Huang, W.-H.; Yeung, E. S. Anal. Chem. 2000, 72, 2879–2885. (53) Smith, S. B.; Aldridge, P. K.; Callis, J. B. Science 1989, 243, 203–206. (54) Zullli, S. L.; Kovaleski, J. M.; Zhu, X. R.; Harris, J. M.; Wirth, M. J. Anal. Chem. 1994, 66, 1708–1712. (55) Swinton, D. J.; Wirth, M. J. Anal. Chem. 2000, 72, 3725–3730. (56) Wirth, M. J.; Swinton, D. J. Anal. Chem. 1998, 70, 5264–5271. (57) Honcius, A.; Schwartz, D. K. J. Am. Chem. Soc. 2009, 131, 5973–5979. (58) Kang, S. H.; Shortreed, M. R.; Yeung, E. S. Anal. Chem. 2001, 73, 1091– 1099. (59) Kang, S. H.; Yeung, E. S. Anal. Chem. 2002, 74, 6334–6339. (60) Isailovic, S.; Li, H. W.; Yeung, E. S. J. Chromatogr. 2007, 1150, 259–266. (61) Li, H. W.; Park, H. Y.; Porter, M. D.; Yeung, E. S. Anal. Chem. 2005, 77, 3256–3260.

single-molecule experiments with R-phycoerythrin, Kang et al. analyzed the motion and adsorption/desorption behaviors of these molecules at the fused-silica/water interface at various pH and ionic strengths within the evanescent-field layer.59 Li et al. used λ-DNA as a single-molecule probe to study chromatographic retention on various self-assembled monolayer surfaces based on hydrogen bonding ability and hydrophobicity of the unpaired purine and pyrimidine bases at the ends of the molecule.61 However, so far all of these studies have been limited to optically flat surfaces. Here we monitor the interactions between single protein molecules in aqueous solution and porous polycarbonate membranes. The porous surface more closely resembles real chromatographic packing materials and may provide additional insights beyond the optically flat self-assembled monolayers. Such information should prove valuable for understanding of SEC and membrane separations. EXPERIMENTAL SECTION Buffer Solutions. Na2CO3/NaHCO3 buffer solutions (pH 10.0) were prepared from 0.1 M solutions of Na2CO3 and NaHCO3 (ACS grade or higher, Fisher Scientific, Fair Lawn, NJ) dissolved in ultrapure (18 MΩ) water. The molarity and the nominal ionic strength of the buffer was 5.0 mM. Phosphatebuffered saline (PBS) buffer (10×, pH 7.2) was purchased from Invitrogen (Carlsbad, CA) and diluted to 1× without further adjustment. Ultrapure water from a Milli-Q system (Millipore, Billerica, MA) was used throughout the experiments. All solutions were photobleached overnight using a mercury UV lamp and filtered through a 0.2 µm filter prior to use. Protein Samples. Five proteins of different molecular weights (MW) and hydrodynamics radii (Rh) were used in these experiments: myoglobin (MW ) 16 900, Rh ) 2.05 nm (pH 7.4), pI ) 7.0), bovine hemoglobin (MW ) 65 000, Rh ) 3.20 nm (pH 7.1), pI ) 7.0), bovine serum albumin (BSA, MW ) 67 000, Rh ) 3.51 nm (pH 6.8), pI ) 4.9), immunoglobulin (IgG, MW ) 150 000, Rh ) 5.34 nm (pH 7.4), pI ) 7.0), and fibrinogen (MW ) 340 000, Rh ) 10.95 nm (pH 7.4), pI ) 5.5) were purchased from SigmaAldrich (St. Louis, MO). Protein stock solutions were dissolved in 1× PBS buffer. The proteins were labeled at 2 mg/mL with a microscale Alexa Fluor 532 protein labeling kit from Molecular Probes (Carlsbad, CA). The dyes react with primary amines, forming stable covalent bonds. Alexa Fluor 532 labeled goatantimouse IgG was purchased from Molecular Probes. For singlemolecule imaging, these samples were further diluted to 50 pM at low temperature with appropriate buffer solutions immediately prior to the start of the experiment. These proteins are multiply labeled (4-8 dyes/molecule) and do not photobleach in the 1 s period (100 × 10 ms) in our experiments. Polycarbonate Membranes. Polycarbonate track-etched nanoporous membranes (6 µm thick) with 10, 30, 50, and 80 nm diameter pores were obtained from Sterlitech Corporation (Kent, WA). The pore densities were 6 × 108 (10, 30, 50 nm) or 4 × 108 pores cm-2 (80 nm). The typical scanning electron microscope images of 10, 30, 50, and 80 nm pore-sized polycarbonate (62) Park, H. Y.; Li, H. W.; Yeung, E. S.; Porter, M. D. Langmuir 2006, 22, 4244–4249. (63) Fang, N.; Zhang, H.; Li, J. W.; Li, H. W.; Yeung, E. S. Anal. Chem. 2007, 79, 6047–6054.

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Figure 2. Pore size effect on the immobilization of various proteins across the membrane surface under capillary flow. The total number of stationary molecules in a series of images was counted (mean ( SD, n ) 3).

Figure 1. Scanning electron microscope image of the (a) 10, (b) 30, (c) 50, and (d) 80 nm pore-sized polycarbonate membrane used in our experiments.

membranes are shown in Figure 1. Prior to the measurements, polycarbonate nanoporous membranes were immersed in a buffer at room temperature overnight. Epifluorescence Microscopy and EMCCD Camera. An upright Nikon Eclipse 80i microscope was used to perform singlemolecule experiments. A 100× objective lens (Nikon Plan Apo/ 1.40 oil) was used. The fine focusing adjustment of the microscope is 0.1 mm per rotation. A motorized rotary stage from Sigma Koki (model no. SGSP-60YAM) was coupled to the fine-adjustment knob on the microscope. The motor is controlled by Intelligent Driver, CSG-602R (Sigma Koki). The travel for each pulse is 0.0025° (0.69 nm). An Andor Ixon DV 897 electron multiplying CCD (EMCCD) (cooled at -55 °C, Game 151, SNAP ES, Photometrics, Tucson, AZ) camera was mounted on the microscope. An X-Cite 120 PC (EXPO Photonic Solution) 120 W metal halide short arc lamp was used as a light source. The frame transfer EMCCD camera was operated in the external synchronization mode. Exposure timing for the EMCCD camera and the laser shutter was synchronized by a Uniblitz mechanical shutter (model LS2Z2, Vincent Associates, Rochester, NY) and a driver (model T132, Vincent Associates). The EMCCD acquisition frequency was set at 5 Hz, with the shutter driver set to 10 ms exposure and 190 ms delay for each frame. Winview 32 (Roper Scientific, Princeton, NJ) was used for image collection and data processing. RESULTS AND DISCUSSION Single Protein Molecule Dynamics on the Nanoporous Polycarbonate Membrane Surface. We first investigated the effect of pore size on trapping a single protein on the polycarbonate membrane surface. In order to demonstrate the effect of the pore size on the steric interactions of single protein molecules with the polycarbonate membranes, the surface must not adsorb protein. The sizes of the high-molecular weight proteins lgG and fibrinogen are large compared to 10 nm, so it should be difficult for these proteins to go into the pores. At the same time, at pH 10.0, the electrostatic repulsion between the negatively charged protein and the negatively charged surface is large, preventing adsorption. As can be seen from Figure 2, there were no IgG or fibrinogen molecules immobilized on the 10 nm membrane 480

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surface. However, for the low-molecular weight proteins myoglobin and BSA, we can see that those could be immobilized on the 10 nm membrane surface even though there adsorption can be ruled out based on observations for the larger proteins. Furthermore, there were more myoglobin than BSA molecules immobilized. This is because myoglobin has a smaller size than BSA, making it easier to enter the pores. As for the 30, 50, and 80 nm pore membranes, more and more molecules of each type were immobilized because of decreasing steric hindrance. This confirms that it was indeed due to the presence of the nanopores and not due to adsorption interactions of protein molecules with membranes. Figure 2 thus depicts the relative probabilities of protein molecules entering the nanopores as they move across the surface. Pore Size Effect on Protein Depth Distribution. The membrane pore size in relation to the size of the proteins in solution determines the extent of interaction between the solute and the membranes. Figure 3a illustrates the effect of membrane pore size on myoglobin depth distribution inside the pores. Such information was obtained by varying the focal plane of the microscope while counting the number of in-focus molecules at each distance starting from the membrane surface (see movie in the Supporting Information). The precision stage has a resolution of 0.69 nm. The rotation rate is set at 500 steps/s, and the acquisition frequency is set as 5 Hz. So, each frame travels 100 steps (500 × 0.2) or 69 nm (0.69 × 100). Each movie is 100 frames long, so it covers 6900 nm (69 × 100) for the full membrane thickness. We can determine the depth to the nearest frame, as can be seen from the movie in the Supporting Information. So, the horizontal scales in the histograms should be highly reliable. Since this is a dynamic process, all counts were obtained at the 8 min mark after the solution is introduced to the membrane surface by capillary flow. This choice was based on direct observation of particles as they accumulate on the membrane with the focal plane set at the surface. The protein could penetrate the 80, 50, 30, and 10 nm membrane pores to different depths of 5.5, 4.8, 2.8, and 1.4 µm, respectively. As the depth increased, for each size pore, the number of protein molecules inside the pores decreased rapidly. For example, for 50 nm pores, there were 78, 20, 13, 9, 5, 4, 1, and 0 molecules at a depth of 0, 1.4, 2.1, 2.8, 3.5, 4.1, 4.8, and 5.5 µm, respectively. At the same time, the number of protein molecules inside the pores at a given depth also decreased as the pore size decreased. For example, at a depth of 2.1 µm, there were 18, 13, 7, and 0 protein molecules in the 80, 50, 30, and 10 nm pores, respectively. No molecules were found inside the 10

Figure 3. Effect of membrane pore size on (a) myoglobin and (b) IgG proteins depth distribution inside the pores (mean ( SD, n ) 3).

nm membrane pores because the pore size was not large enough to accommodate the proteins. Ohya et al. found that no transfer of egg albumin occurred through track-etched nuclepore membranes of pore sizes less than 30 nm and concluded that the minimum pore size for any transfer of egg albumin to occur was 10 times its radius.64 However, Yu et al. argued that it did not mean that there was no protein transferred; it simply meant that the concentration was below the detection limit of their analytical method.17 Figure 3b illustrates the effect of pore size on IgG depth distribution inside the membrane pores. In general the results were similar to those for myoglobin. Molecular Size Effect on Protein Depth Distribution. Figure 4a illustrates the effect of protein molecular size on the depth distribution at pH 10.0 in the 80 nm pores. All of the proteins could enter the 80 nm pores, but at different depths the numbers of molecules were very different. The number of molecules inside the pores at the same depth also decreased as the protein molecular weight increased. At the depth of 1.4 µm, there were 29 myoglobin molecules, 21 BSA molecules, 8 IgG molecules, and 1 fibrinogen molecule. This indicates that high-molecular weight molecules moved more slowly than low-molecular weight molecules once they go into the pores. Figure 4b illustrates the effect of molecular size on depth distribution in 30 nm pores. In general the results were similar to those for 80 nm pores. Molecular Shape Effect on Protein Depth Distribution. Because hemoglobin and BSA have nearly the same molecular weight (65 000 vs 67 000), the free-solution diffusion coefficients for both molecules are nearly the same (BSA, 7.0 × 10-7 cm2 s-1, hemoglobin, 6.9 × 10-7 cm2 s-165). However, as can be seen (64) Ohya, H.; Kuromoto, M.; Watanabe, N.; Matsumoto, K.; Negishi, Y. Membrane 1989, 14, 329–336.

Figure 4. Effect of protein molecular size on depth distribution at pH 10.0 in (a) 80 nm pores and (b) 30 nm pores (mean ( SD, n ) 3).

Figure 5. Distribution of pore depth of individual protein molecules at pH 10.0 for BSA and hemoglobin (mean ( SD, n ) 3).

from Figure 5, the number of BSA in the membrane with 50 nm pores is more than that of hemoglobin at all depths of the pores. At the depths of 0 µm, 1.4 µm, 2.1 µm, 2.8 µm, 3.5 µm, 4.1 µm, and 4.8 µm, there were 56, 9, 4, 3, 2, 1, and 0 BSA molecules, respectively, while there were only 44, 6, 3, 1, 1, 1, and 0 hemoglobin molecules, respectively. Chun and Stroeve found that BSA diffuses faster than hemoglobin in gold nanotube membranes.18 They have discussed various explanations for this observation including the possibility that it is related to the difference in molecular shape. BSA is an elliptically shaped molecule, with approximate dimensions of 4 nm × 4 nm × 14 nm,66 whereas hemoglobin is more spherical, with approximate dimen(65) Mortimer, R. G. Physical Chemistry, 3rd ed.;Academic Press: Orlando, FL, 2008; p 474. (66) Musale, D. A.; Kulkarni, S. S. J. Membr. Sci. 1997, 136, 13–23.

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Table 1. Diffusion Coefficient of Proteins Inside Nanopores Determined from Gaussian Fitting of the Depth Histograms for 8 Minute Excursionsa D value (10-3 µm2/s) pore myoglobin (nm) 10 30 50 80

0.65 ± 0.06 1.48 ± 0.10 1.53 ± 0.09 1.88 ± 0.07

BSA

IgG

0 1.06 ± 0.08 1.19 ± 0.10 1.68 ± 0.13

0 0.66 ± 0.03 0.88 ± 0.06 1.28 ± 0.09

fibrinogen hemoglobin 0 0 0.67 ± 0.05 1.02 ± 0.06 0.71 ± 0.06

a Mean ± SD, n ) 3. The free solution D values65 are myoglobin, 110 µm2/s; BSA, 70 µm2/s; IgG, 37 µm2/s; fibrinogen, 22 µm2/s; hemoglobin, 69 µm2/s.

sions of 6.4 nm × 5.5 nm × 5 nm.67 The shapes of the protein molecules influence diffusion for BSA and hemoglobin in the nanopores. When there is electrostatic repulsion between the pore wall and the BSA, the prolate ellipsoid should be more aligned with the axis of the pore. This alignment of BSA within the pore should cause a reduction in hindered diffusion and cause BSA to diffuse faster compared to hemoglobin. One-Dimensional Diffusion Coefficients in Nanopores. The depth histograms above allow us to invoke a one-dimensional model for the diffusion of the proteins once they enter the nanopores. Since each pore is independent, the histograms represent separate measurements of the distance traveled by a molecule starting from the surface after a fixed time of 8 min. The fit is to a Gaussian curve with a half width s such that s2 ) 2Dt. This is different from fitting concentration curves in bulk measurements with the error function. The calculated diffusion coefficients are given in Table 1. The general trends of the D values are as expected, with the smaller pores giving smaller D values and the smaller proteins giving larger D values. However, the absolute values are orders of magnitude smaller than those for solution (bulk) measurements in the literature, even when the latter were converted to one(67) Hinz, H. J. Thermodynamic Data For Biochemistry and Biotechnology; Springer: Berlin, Germany, 1986; p 237.

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dimensional D values and even when accounting for uncertainties in selecting a diffusion time of 8 min. Since these proteins have been shown not to adsorb onto polycarbonate surfaces (from experiments performed with 10 nm pores), adsorption on the inner walls is not expected unless track etching modified the inner surface significantly. Steric effects are unlikely to be important, as the smooth-looking tracts in Figure 1 indicate that there are no features (roughness) extending into the pores from the walls that would significantly modify the stated diameters of the pores. Furthermore, the pore-size dependence (each column in Table 1) did not follow a diameter-squared relationship that would apply to adsorption mediated by the probability of molecules in solution encountering the walls. Also, the variation among proteins (each row in Table 1) was strictly size dependent rather than related to their pI values. It has been reported before40 that the viscosity of water is anomalously large near an interface even at the micrometer scale. This could explain the extremely small D values at the nannometer scale. Such D values seem to preclude the ability of any protein from passing through these membranes. However, in membrane separations, pressure is applied on one side to facilitate bulk flow rather than simply relying on diffusion. Future experiments with modified pore surfaces may be able to clarify the importance of each of these parameters. ACKNOWLEDGMENT The Ames Laboratory is operated for the U.S. Department of Energy by Iowa State University under Contract No. DE-AC0207CH11358. This work was supported by the Director of Science, Office of Basic Energy Science, Division of Chemical Sciences. SUPPORTING INFORMATION AVAILABLE Additional information as noted in text. This material is available free of charge via the Internet at http://pubs.acs.org.

Received for review November 30, 2009. AC902487C

November

1,

2009.

Accepted