Rotational Dynamics of Laterally Frozen Nanoparticles Specifically

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2009, 113, 11179–11183 Published on Web 06/04/2009

Rotational Dynamics of Laterally Frozen Nanoparticles Specifically Attached to Biomembranes Sebastien Pierrat, Eva Hartinger, Simon Faiss, Andreas Janshoff, and Carsten So¨nnichsen* Physical Chemistry, UniVersity of MainzJakob-Welder-Weg 11, 55128 Mainz, Germany ReceiVed: April 2, 2009; ReVised Manuscript ReceiVed: May 19, 2009

The strongly polarized light scattering of gold nanorods at their longitudinal plasmon frequency allows for the tracking of single gold nanorod lateral positions and orientations via optical dark-field microscopy. We monitor both lateral and rotational diffusion of polymer-coated gold nanorods attached to artificial biomembranes on solid supports. The attachment is mediated by the biotin-streptavidin receptor-ligand system, but weak interaction is also observed in the absence of streptavidin. In the latter case, we observe a twodimensional lateral diffusion of the nanorods with a diffusion coefficient of 0.5 µm2 s-1. This lateral motion is strongly reduced with the addition of streptavidin. However, the particles are still able to rotate, and we study their rotational motion using polarization contrast microscopy. The rotational diffusion time in the range of 100 ms depends on the biotin concentration in the membrane, hence the number of anchor points, and on the temperature. Cooling the membrane beyond its gel-fluid transition point leads to a reduction of the rotational motion. The experimental results can be understood in terms of dragging forces introduced by the surface viscosity of the membrane. A quantitative analysis shows that entire patches of the membrane move with the particles. Introduction Rotational and lateral Brownian motion of anisotropic (rodor needle-shaped) particles in contact with two-dimensional (2D) membranes plays a crucial role in many biological phenomena, for example the cellular uptake of nanoparticles or viruses.1 Moreover, two-dimensionally restricted Brownian motion of rodshaped colloidal particles is a complex phenomenon in itself since lateral and rotational diffusion are intrinsically coupled. Even though this fact was emphasized more than fifty years ago by Perrin,2,3 Tom Lubensky and his co-workers only recently published a high-profile experimental study of the twodimensional Brownian motion of a micrometer-sized ellipsoid.4 Coupling the Brownian motion of a particle with a membrane or lipid bilayer adds the complicating viscoelastic properties of membranes to the overall system. Recent experimental results suggest that the commonly used Saffman-Delbruck model5,6 describing diffusion in membranes might not always be valid.7 In particular, it has been suggested that local deformation of the membrane would introduce additional hydrodynamic stress leading to a strong reduction of mobility.8,9 Single particle tracking with optical microscopy has been used extensively to study Brownian motion and translational diffusion.10-12 Rotational diffusion, which is many orders of magnitude faster for nanoparticles, requires very fast detection systems and an anisotropic optical signal of the tracked particles. Rod-shaped gold nanoparticles (nanorods) show strong light scattering cross sections at the longitudinal plasmon resonance,13 which allows the high contrast observation of single nanorods with dark-field optical microscopy.14 * To whom correspondence should be addressed. E-mail: soennichsen@ uni-mainz.de.

10.1021/jp9030333 CCC: $40.75

Because the light scattered by gold nanorods is polarized, it can be used to monitor their orientation over time to probe rotational diffusion on very small length scales.15 Initial studies used gold nanorods to track material deformations16 and to characterize the rotational motion of F1-ATPase molecules with high temporal resolution.17 Here, we study the two-dimensional lateral and rotational diffusion of gold nanorods attached to supported membranes and observe remarkable differences between nonspecific and specific interactions. Loosely bound nanoparticles show noticeable translational diffusion, whereas the presence of specific linkers significantly reduces lateral diffusion and slows down rotational diffusion. The single particle orientation time traces give strong support for a microscopic model of the nanorodmembrane system, where entire patches of the membrane rotate together with the particle. Experimental Section Synthesis and Biotinylation of Gold Nanorods. Gold nanorods are produced according to the seeded growth method18,19 briefly explained in the Supporting Information. The gold nanorods used in this study have a size of 15.5 × 41.5 nm and an optical extinction maximum at 650 nm (Supporting Information, Figure S1). The gold nanorods are coated with heterobifunctional mercapto-PolyEthyleneGlycol (PEG, MW 5000, Iris Biotech GmbH), which stabilizes the particles and provides chemical anchor points.20,21 Here, we use a mixture of methoxyPEG (SH-PEG-OCH3) and amino-PEG (SH-PEG-NH2) in a ratio of 1:1. The amino groups are used to attach a biotin derivative (Sulfo-NHS-biotin, Pierce).  2009 American Chemical Society

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Figure 1. (a) Schematic representation of the experimental setup. The sample is illuminated by a dark-field condenser (oil immersion) and the light scattered by the gold nanorods is collected by a water immersion objective. A two-channel polarization system (Dual-View represented in the gray-dashed box) is then mounted on the microscope, allowing the separation of two orthogonal polarization directions. The two signals are then recorded by a electron-multiplying CCD camera on a single frame, half of the picture corresponding to one polarization direction, the other half to the orthogonal one. (b) Time trace of the reduced intensity recorded by the first channel (blue dots), the second channel (black dots), and reduced linear dichroism (red dots). (c) Power spectral density (PSD) of the reduced linear dichroism fitted by a Lorentzian curve (red line) to determine the characteristic time.

Membrane Preparation. Supported lipid bilayers are prepared by spreading unilamellar vesicles22 in microfluidic channels23 (details are described in the Supporting Information). Single Particle Tracking. The gold rods are visualized using dark-field microscopy. Tracking of single particle motion is performed using a fast camera (PhotonMAX, Princeton Instrument) operating at 50 fps with a field of view of 70 µm × 20 µm. The time trace of the particle positions is determined by an image analysis script (Matlab). Single Particle Orientation Tracking. Taking advantage of the strongly polarized light scattered by gold nanorods, the nanorods’ orientation is determined using two-channel polarization contrast microscopy (DualView, Roper Bioscience) as sketched in Figure 1a. A beam splitter and two polarization filters allow the separation of two orthogonal polarization directions I⊥ and I| of the same area on the sample. The two signals are recorded by a fast camera operating at 200 fps for up to 30 s, each of them being projected on half of the EM-CCD chip. The field of view is reduced to 100 µm × 15 µm. A Matlab script has been developed to identify the same particle on both channels and to determine the time trace of the respective intensities I⊥(t) and I|(t). Analysis of Orientation Time Traces. The dynamics of the rotational diffusion of a single gold nanorod can be analyzed from the correlation of the fluctuating polarization signal. A typical signal of a rotating single particle shows anticorrelated intensities in the two channels (Figure 1b, blue and black dots).

We consider a particle to be rotating if the correlation coefficient of the time dependent intensities I⊥(t) and I|(t) is smaller than -0.25. The reduced linear dichroism A(t)24

A(t) )

I//(t) - I⊥(t) I//(t) + I⊥(t)

is computed (Figure 1b, red dots) in order to reduce the noise arising from experimental artifacts such as overall fluctuations of the light intensity or out-of-plane rotations. We fit the power spectral density PSD of the reduced linear dichroism with a Lorentzian function to obtain the rotational diffusion times τdif (Figure 1c, details in the Supporting Information). Results and Discussion As a first step, we study the specificity of the interactions between biotinylated gold nanorods and lipid membranes covered by streptavidin. We observe single particles attached to supported POPC lipid bilayers using dark-field optical microscopy. The lipid bilayers are created on the substrate in two separate microfluidic channels with one channel always containing 2% wt biotin-DOPE, whereas in the second channel, the biotin-DOPE concentration is varied from 0 to 1% wt (Supporting Information, Figure S2). The density of rods immobilized on the lipid bilayer increases with the concentration of biotin-DOPE as expected (Figure 2). Controls with no biotinylated lipids or no streptavidin show approximately the same low coverage of immobilized rods than the BSA passivated

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Figure 2. Characterization of the interactions between biotinylated gold nanorods and streptavidin-coated biomembranes. (a) Dark-field microscopy picture of two phosphocholine (POPC) membranes containing either 0% (left channel) or 2% biotinylated lipids (right channel). After passivation of the glass substrate with bovine serum albumin (BSA) and incubation of streptavidin, biotinylated gold nanorods are immobilized on the membrane. On the membrane containing 2% biotinylated lipids, about 100 rods per 100 µm2 are found whereas less than 30 rods per 100 µm2 are found on the membrane containing no biotinylated lipids. The scale bar is 40 µm. (b) Numbers of gold nanorods counted on membranes containing different biotinylated lipids concentrations relative to the number of rods on a 2% biotin membrane.

glass surface next to the lipid bilayer. The immobilization of the biotinylated gold nanorods on the membrane is therefore predominately caused by the specific streptavidin-biotin interaction. However, even in the absence of streptavidin, a few gold nanorods are attached to the membrane. On those nonspecifically attached particles, we observe noticeable 2D lateral diffusion on time scales of seconds (Figure 3a). By tracking single particle trajectories of 43 particles for 60 s and calculating the mean square displacements (MSD), we find a diffusive behavior following MSD(t) ) 4 Dtt with a lateral diffusion coefficient of Dt ) 0.52 ( 0.06 µm2 s-1 (Figure 3b). This value is much lower than the lateral diffusion coefficient calculated for the gold nanorods in pure water (Dt ) 8.08 µm2 s-1, see Supporting Information) and reflects the friction introduced by the membrane. However, considering the drag of PEG molecules penetrating the membrane (Figure 3c), using the SaffmanDelbruck model,5 leads to reasonable agreement with the experimental data (Supporting Information). The rotational diffusion of gold nanorods in the absence of streptavidin is too fast to be measured. The estimation of the rotational diffusion time leads to a value lower than 100 µs, well below the time resolution (5 ms) of our camera (Supporting Information). However, once streptavidin is added to the system, the interaction with the membrane becomes strong enough to freeze the lateral movement of the gold nanorods and to slow down the rotational motion. The density of PEG molecules on the gold

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Figure 3. Lateral diffusion of gold nanorods on a POPC membrane. (a) Trajectory of one single particle. (b) Average of the MSD calculated from the trajectory of 43 particles (gray dots) and linear fit (black line). The diffusion coefficient is estimated to be 0.52 ( 0.06 µm2 s-1. (c) Schematic representation of the interpenetration of the PEG molecules inside the membrane introducing a viscous drag that slows down the lateral diffusion of the gold nanorods.

nanorod surface is on the order of 5 PEG/100 nm2.20 Therefore, about 20 biotin molecules would be available to anchor the nanoparticles to the membrane, causing a local accumulation of streptavidin at the interface. We would therefore expect that the collective diffusion of the occluded area below the gold nanorods determines the mobility12 and results in the apparent trapping of the particles on any reasonable experimental time scale.25 Remarkably, we are still able to detect rotations of those laterally immobile particles. Depending on the concentration of biotinylated lipids, we find the rotational diffusion time varies from τdif ) 55 ( 4 ms to 217 ( 7 ms. The content of biotinylated lipids varies from 0.05 to 1% wt, and in each set of experiments about 100 particle traces are recorded (Figure 4a). The resulting rotational diffusion times show a linear dependency on the biotin concentration (Figure 4b), suggesting that the streptavidin molecules are acting as anchoring points for the gold nanorods on the membrane, which consequently reduces their mobility (Figure 5). Such an interpretation is confirmed by the reduction of the rotational diffusion time from 101 ( 5 to 42 ( 4 ms after exposing a membrane containing 0.2% wt biotinylated lipids for 45 min with an abundance of free biotin (Figure 4b and Supporting Information, Figure S3). The free biotin competes with the biotin moieties displayed by the gold nanorods and hence blocks potential anchor points on the membrane, leading to faster rotations of the particles on the bilayer.

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Figure 5. Schematic representation of the possible interactions between the gold nanorods and the lipid bilayer. The blue worms correspond to the biotinylated PEG molecules whereas the red balloons are the streptavidin attached to the membrane. (a) Low concentration of biotinylated lipids resulting in a few anchoring points of the gold nanorods on the membrane. The rotation of the rods is thought to be slowed by the drag of a small disk of lipids. (b) High concentration of biotinylated lipids resulting in a large number of anchoring points. Possible local deformation of the membrane might lead to a slower rotation of the gold nanorods.

Figure 4. Rotational diffusion of biotinylated gold nanorods on streptavidin coated biomembranes. (a) Histograms of the measured rotational diffusion time of individual gold nanorods on membranes containing different concentrations of biotinylated lipids. (b) Rotational diffusion time as a function of the biotinylated lipids concentration. The black points are the experimental data points and the dashed line is a guide for the eyes. The red point is the measured point after addition of an excess of free biotin. (c) Phase transition of a diC15-PC membrane containing 0.2% biotinylated lipids. Rotational diffusion time (black dots) and fraction of rotating rods (gray dots) as functions of the temperature. The transition temperature is found to be about 40 °C. The dashed lines are guides for the eyes.

A quantitative description of the surface shear viscosity experienced by spheres partially immersed in a membrane was recently reported by Fischer et al.26 They found the rotational drag fmemb of a sphere with radius r immersed in the membrane to a depth h to be

fmemb ) ηmr3(8π - 5 ln[tanh(3(2r - h)/2r)]) If f is the total drag on the particle defined as the sum of the water and the membrane contributions, the rotational diffusion coefficient is then given by

Dr ) kBT/f

with

f ) fmemb + fw

and the rotational diffusion times are obtained from τdif ) 1/2Dr.

The experimentally found rotational diffusion times τdif ) 55-217 ms would correspond, according to this model, to spheres of radii r ) 45 - 70 nm immersed in the membrane to a depth of h ) 2.5 nm. A radius of 45 nm corresponds to the radius of gyration of the gold nanorods calculated from the size of the particles. The gold nanorods would drag a disk of lipids that form around the gold particles during their rotations.27 Increasing the biotin concentration in the membrane allows the formation of a larger cluster around the particle, that is, this dragging disk enlarges, which inherently increases the rotational diffusion time (Figure 5). A larger number of anchoring points may also result in the deformation of the membrane,11 perturbing the mobility of the lipids inside the bilayer as well as the motion of the nanoparticles on the membrane.9 The estimation of the lateral diffusion of the particles in similar conditions leads to a value about 50 times smaller than the value measured in the absence of streptavidin. This explains the absence of any lateral diffusion in the time scale of our experiments. In order to confirm that the dynamics of nanoparticles’ rotations are governed by the membrane’s physical properties, we measure the rotational diffusion on a membrane in the gel phase at room temperature. For this temperature-controlled experiment, we use diC15-PC lipids to prepare the membrane, which undergoes a gel-to-fluid phase transition around 35 °C.28 Varying the temperature from 30 to 50 °C, we extract the fraction of rotating rods, and for rotating particles, the rotational diffusion time (Figure 4c and Supporting Information, Figure S4). At high temperatures, the fraction of rotating rods is relatively high (60% at T ) 50 °C) and the rotational dynamics are fast (τdif ) 60 ( 4 ms). At low temperatures, the fraction of rotating rods (gray dots, Figure 4c) is significantly smaller (14% at T ) 30 °C) and the rotation (black dots, Figure 4c) is much slower (τdif ) 140 ( 8 ms). It should be emphasized that the rotational diffusion times are calculated only for the rotating gold nanorods and hence do not reflect the dynamics of the whole population. A transition in between those two regimes occurs at 40 °C, whic is in good agreement with the main phase transition temperature of diC15-PC on solid supports.28 Again, the measured diffusion times are in good quantitative agreement

Letters with the estimations using the model proposed by Fischer et al.26 A sphere with a radius of r ) 37 nm and viscosities of ηm ) 400 cP for the fluid29 and ηm ) 1000 cP for the gel phase30 produces rotational diffusion times similar to our measurements. Conclusions Using a single particle lateral and rotational tracking technique based on plasmonic light scattering, we have observed that the density of nanoparticles bound to the membrane increases with the density of biotinylated lipids. Particles interacting nonspecifically with the membrane are laterally mobile, whereas particles attached to it via streptavidin-biotin complexes are laterally fixed but able to rotate. The rotational diffusion time depends both on the density of streptavidin-biotin complexes and on the viscosity of the membrane. A gel-fluid phase transition is clearly observable in the fraction of rotating rods and in the rotational diffusion time. Taken together, these observations conclusively support the picture sketched in Figure 5. The mobility of rod-shaped gold nanoparticles on artificial supported biomembranes is governed by both the viscous properties of the membrane and the strength of the membranenanoparticle interaction. The dynamics of rotational motion are sensitive to the number of streptavidin molecules acting as anchoring points that determine the area of the membrane discs involved in the rotation. The measured values are in reasonable agreement with theoretical predictions based on models developed for spherical particle diffusion on biomembranes. The ability to observe separately lateral and rotational motion that occur on different time scales, as reported in this article, will give new insights on the diffusion of anisotropic molecules bound to membranes. In particular, appropriate functionalization of the gold nanorods and further characterization of the system would potentially provide information on the effects of clustering and adhesion on the mobility of anisotropic objects inside a membrane. We hope the experimental results presented in this report will trigger the development of a more detailed theoretical framework for the two-dimensional diffusion of anisotropic (rodlike) particles on membranes under the influence of surface shear viscosity. Acknowledgment. We thank Holger Adam for help with data analysis, Peter Boertz for help with the 3D graphic tools and Kurt Binder and Tanja Schilling for fruitful discussion. We acknowledge financial support by the DFG under the Emmy Noether program (SO 712/1-3). Supporting Information Available: Detailed descriptions of the experimental methods and the diffusion coefficients

J. Phys. Chem. C, Vol. 113, No. 26, 2009 11183 estimation, figures showing the size and optical spectra of the gold nanorods (Figure S1), the preparation method for the supported membrane (Figure S2), the rotational diffusion after incubation with free biotin (Figure S3), and the rotational diffusion on a diC15-PC membrane (Figure S4). This material is available free of charge via the Internet at http://pubs.acs.org. References and Notes (1) Seisenberger, G.; Ried, M. U.; Endress, T.; Buning, H.; Hallek, M.; Brauchle, C. Science. 2001, 294, 1929. (2) Perrin, F. J. Phys. Rad. 1934, 5, 497. (3) Perrin, F. J. Phys. Rad. 1936, 7, 1. (4) Han, Y.; Alsayed, A. M.; Nobili, M.; Zhang, J.; Lubensky, T. C.; Yodh, A. G. Science. 2006, 314, 626. (5) Saffman, P. G.; Delbruck, M. Proc. Nat. Acad. Sci. U.S.A. 1975, 72, 3111. (6) Peters, R.; Cherry, R. J. Proc. Nat. Acad. Sci. U.S.A. 1982, 79, 4317. (7) Gambin, Y.; Lopez-Esparza, R.; Reffay, M.; Sierecki, E.; Gov, N. S.; Genest, M.; Hodges, R. S.; Urbach, W. Proc. Natl. Acad. Sci. U.S.A. 2006, 103, 2098. (8) Guigas, G.; Weiss, M. Biophys. J. 2006, 91, 2393. (9) Naji, A.; Levine, A. J.; Pincus, P. A. Biophys. J. 2007, 93, L49. (10) Saxton, M. J.; Jacobson, K. Annu. ReV. Biophys. Biomol. Struct. 1997, 26, 373. (11) Lee, G.; Ishihara, A.; Jacobson, K. Proc. Nat. Acad. Sci. U.S.A. 1991, 88, 6274. (12) Zhang, L. F.; Granick, S. Proc. Natl. Acad. Sci. U.S.A. 2005, 102, 9118. (13) Perez-Juste, J.; Pastoriza-Santos, I.; Liz-Marzan, L. M.; Mulvaney, P. Coord. Chem. ReV. 2005, 249, 1870. (14) So¨nnichsen, C.; Franzl, T.; Wilk, T.; von Plessen, G.; Feldmann, J.; Wilson, O.; Mulvaney, P. Phys. ReV. Let. 2002, 88, 077402. (15) So¨nnichsen, C.; Alivisatos, A. P. Nano Lett. 2005, 5, 301. (16) Orendorff, C. J.; Baxter, S. C.; Goldsmith, E. C.; Murphy, C. J. Nanotechnology 2005, 16, 2601. (17) Spetzler, D.; York, J.; Daniel, D.; Fromme, R.; Lowry, D.; Frasch, W. Biochemistry 2006, 45, 3117. (18) Nikoobakht, B.; El-Sayed, M. A. Chem. Mater. 2003, 15, 1957. (19) Jana, N. R.; Gearheart, L.; Murphy, C. J. AdV. Mater. 2001, 13, 1389. (20) Pierrat, S.; Zins, I.; Breivogel, A.; So¨nnichsen, C. Nano Lett. 2007, 7, 259. (21) Hanauer, M.; Pierrat, S.; Zins, I.; Lotz, A.; So¨nnichsen, C. Nano Lett. 2007, 7, 2881. (22) MacDonald, R. C.; MacDonald, R. I.; Menco, B. P. M.; Takeshita, K.; Subbarao, N. K.; Hu, L. R. Biochim. Biophys. Acta 1991, 1061, 297. (23) Janshoff, A.; Kunneke, S. Eur. Biophys. J. 2000, 29, 549. (24) Wei, C. Y.; Lu, C. Y.; Kim, Y.; Vanden Bout, D. J. Fluoresc. 2007, 17, 797. (25) Saxton, M. J. Biophys. J. 1993, 64, 1766. (26) Fischer, T. M.; Dhar, P.; Heinig, P. J. Fluid. Mech. 2006, 558, 451. (27) Prasad, A.; Kondev, J.; Stone, H. A. Phys. Fluids 2007, 19. (28) Faiss, S.; Schuy, S.; Weiskopf, D.; Steinem, C.; Janshoff, A. J. Phys. Chem. B. 2007, 111, 13979. (29) Cherry, R. J.; Godfrey, R. E. Biophys. J. 1981, 36, 257. (30) Koan, M. M.; Blanchard, G. J. J. Phys. Chem. B. 2006, 110, 16584.

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