6628
Langmuir 1998, 14, 6628-6631
Scanning Force Microscopy Images of Cytochrome c Oxidase Immobilized in an Electrode-Supported Lipid Bilayer Membrane James D. Burgess,† Vivian W. Jones, and Marc D. Porter* Microanalytical Instrumentation Center, Ames LaboratorysUSDOE, and Department of Chemistry, Iowa State University, Ames, Iowa 50011
Melissa C. Rhoten and Fred M. Hawkridge* Department of Chemistry, Virginia Commonwealth University, Box 842006, Richmond, Virginia 23284 Received June 3, 1998. In Final Form: September 21, 1998 Scanning force microscopic images of cytochrome c oxidase immobilized within an electrode-supported lipid bilayer membrane are reported. These images represent the first direct evidence that the microscopic architecture of this important model system effectively mimics the microstructure proposed for the inner mitochondrial membrane. The images reveal (1) that the oxidase is present as monomers and small aggregates within the supported membrane and (2) that the oxidase constitutes ∼20% of the lipid bilayer membrane. The latter finding allows an estimation of the lower limit for the minimum turnover rate (∼0.5 electrons/s) required for the transition of the oxidase from its resting to pulsed kinetic state.
Introduction Mammalian cytochrome c oxidase is the terminal enzyme in oxidative phosphorylation.1 This vital molecular machine is embedded within the inner mitochondrial membrane and pumps protons against a concentration gradient to support adenosine triphosphate production.2 As such, this oxidase derives its energy through catalyzing the reduction of oxygen by cytochrome c, a small watersoluble protein that resides between the outer and inner mitochondrial membranes.3 Despite a number of elegant investigations using, for example, vesicles as biomimetic hosts,1 the development of critical insights into the factors that control the performance of the membrane-confined oxidase has proven elusive. This situation reflects, in large part, the lack of analytical methodologies to interrogate effectively the oxidase in a lipid bilayer membrane (LBM) environment. To address this limitation, Hawkridge and co-workers have devised an approach that couples the self-assembly of thiol-based adlayers with deoxycholate dialysis for the creation of oxidase-containing LBMs that are supported on metal electrodes.4-7 Figure 1 presents the conceptualization of this model system. The model has the enzyme oriented within the LBM in a manner similar to that envisioned at the inner mitochondrial membrane.1 As such, the larger hydrophilic end of the oxidase protrudes * To whom correspondence should be addressed: Marc D. Porter (e-mail:
[email protected]) or Fred M. Hawkridge (email:
[email protected]). † Present address: Longwood College, Department of Natural Sciences, 201 High St., Farmville, VA 23909. (1) Malatesta, F.; Antonini, G.; Sarti, P.; Brunori, M. Biophys. Chem. 1995, 54, 1. (2) Wikstro¨m, M. Nature 1977, 266, 271. (3) Babcock, G. T.; Wiksto¨m, M. Nature 1988, 356, 301. (4) Cullison, J. K.; Hawkridge, F. M.; Nakashima, N.; Yoshikawa, S. Langmuir 1994, 10, 877. (5) Burgess, J. D.; Hawkridge, F. M. Langmuir 1997, 13, 3781. (6) Burgess, J. D.; Rhoten, M. C.; Hawkridge, F. M. Langmuir 1998, 14, 2467. (7) Burgess, J. D.; Rhoten, M. C.; Hawkridge, F. M. J. Am. Chem. Soc. 1998, 120, 4488.
a few nanometers from the bilayer with its cytochrome c binding site exposed at the outer LBM surface.1,8 We note that the model, which is supported by the findings of several electrochemically based investigations,4-7 depicts the architecture developed from investigations of the oxidase crystal structure8 and from characterizations of the oxidase reconstituted into vesicles.9-13 Microstructural characterizations of oxidase-containing LBMs that are supported at solid substrates have yet to be carried out. The electrode-supported LBMs have provided a much needed avenue for testing and extending fundamental elements of the functional characteristics of the oxidase that were heretofore largely conjecture.7 Indeed, recent work by Hawkridge and co-workers has established that (1) the embedded oxidase exhibits a transition from its resting kinetic state to its pulsed kinetic state and (2) a minimum turnover rate is required to trigger this transition.7 This dependence was first proposed in 1977 by Antonini and co-workers,14 testifying to the long standing need for a viable approach to probe the operational characteristics of this important enzymatic system. Interestingly, the oxidase-containing LBM also appears to function as a proton pump in the presence of oxygen7 and exhibits a catalytic selectivity for cytochrome c derived, for example, from horse as opposed to tuna.15 Furthermore, the immobilized oxidase exhibits a dependence of (8) Tsukihara, T.; Aoyama, H.; Yamashita, E.; Tomizaki, T.; Yamaguchi, H.; Shinzawa-Itoh, K.; Nakashima, R.; Yaono, R.; Yoshikawa, S. Science 1995, 269, 1069. (9) Henderson, R.; Capaldi, R. A.; Leigh, J. S. J. Mol. Biol. 1977, 112, 631. (10) Deatherage, J. F.; Hederson, R.; Capaldi, R. A. J. Mol. Biol. 1982, 158, 487. (11) Deatherage, J. F.; Hederson, R.; Capaldi, R. A. J. Mol. Biol. 1982, 158, 501. (12) Valpuesta, J. M.; Henderson, R.; Frey, T. G. J. Mol. Biol. 1990, 214, 237. (13) Tihova, M.; Tattrie, B.; Nicholls, P. Biochem. J. 1993, 292, 933. (14) Antonini, E.; Brunori, M.; Colosimo, A.; Greenwood, C.; Wilson, M. T. Proc. (15) Rhoten, M. C.; Burgess, J. D.; Hawkridge, F. M. Manuscript in preparation.
10.1021/la980648d CCC: $15.00 © 1998 American Chemical Society Published on Web 11/10/1998
Letters
Langmuir, Vol. 14, No. 23, 1998 6629
Figure 1. Model of the cytochrome c oxidase modified electrode (not to scale). Darker color of the oxidase indicates the small and large hydrophilic ends of the molecule.
electron-transfer rate on exposure to cyanide,6 a respiratory inhibitor that blocks the oxygen binding site of the oxidase.1 The oxidase therefore appears to be fully functional when embedded in the electrode-supported LBM. In view of the potential significance of these findings, it is essential that the oxidase-containing LBM be established as a viable biomimetic model16 at a microstructural level. This paper examines this issue by applying tapping mode scanning force microscopy (TMSFM) to image oxidase-containing and oxidase-devoid LBMs. The resulting images, which are presented herein, support the proposed architecture of the LBM by revealing the presence of monomers and small aggregates of the oxidase sequestered within the electrode-supported LBM. These results also provide a basis for an estimate of oxidase coverage, which is then applied to an evaluation of the earlier kinetic data7 related to the turnover rate that is required for the transition of the enzyme from its resting to pulsed kinetic state. Experimental Section Sample Preparation. Details of the sample preparation and oxidase purification17 have been reported earlier.4-6 Briefly, a gold-coated quartz crystal microbalance (QCM) electrode is covered with 1.6 monolayers of electrodeposited silver to create a surface at which an octadecanethiolate submonolayer can be formed. The response of the QCM is used to control the extent of the deposition of the atomically thick silver film and of the organic adlayer.5 The silver-sulfur bond accommodates an orientation of the hydrocarbon tail of the resulting alkanethiolate that is near normal with respect to the surface plane.18-21 (16) Sackman, E. Science 1996, 271, 43. (17) Soulimane, T.; Buse, G. Eur. J. Biochem. 1995, 227, 588. (18) Walczak, M. M.; Chung, C.; Stole, S. M.; Widrig, C. A.; Porter, M. D. J. Am. Chem. Soc. 1991, 113, 2370. (19) Bryant, M. A.; Pemberton, J. E. J. Am. Chem. Soc. 1991, 113, 8284. (20) Laibinis, P. E.; Whitesudes, G. M.; Allara, D. L.; Tao, Y.; Parikh, A. N.; Nuzzo, R. G. J. Am. Chem. Soc. 1991, 113, 7152.
The oxidase-containing LBM is formed at the thiolate-modified electrode via deoxycholate dialysis,4,6 a procedure used for reconstituting the oxidase into vesicles.22 The dialysis is accomplished by attaching the thiolate-modified substrate to a dual-chambered electrochemical cell.6 The cell contains a sample chamber and a flow chamber that are separated by a dialysis membrane. The substrate is attached to the sample chamber, which is filled with a solution containing deoxycholate, biological amphliphiles, phosphate buffer (pH 7.4), and the oxidase enzyme. Buffer is passed through the flow chamber of the cell to dialyze the enzymatic solution. The dialysis drives the formation of an LBM whereby the oxidase is sequestered within the chain structure of the alkanethiolate adlayer.6 We note that (1) the partial octadecanethiolate monolayer stabilizes the lipid bilayer through hydrophobic interactions that occur between the alkyl chains of the organic adlayer and the lipid hydrocarbon tails6 and (2) the unmodified regions of the silver surface serve as sites that accommodate the smaller hydrophilic end of the oxidase along with the polar lipid headgroups and trace levels of water.6,7 After dialysis, the sample chamber is extensively flushed with buffer. Instrumentation. All images were acquired in air using a Digital Instruments Nanoscope IIIa (Santa Barbara, CA), equipped with a phase extender module and a 15-µm scanner. The silicon-based SFM probes (Nanosensors) were 124 µm long, with resonance frequencies between 298 and 365 kHz. In the acquisition of these images, the set point amplitude of the cantilever was maximized relative to its free space amplitude. Samples were rinsed with water and dried using high-purity nitrogen prior to imaging.
Results and Discussion Electrochemical investigations4,6,7 have indicated that the oxidase is confined within the LBM in a manner that functionally mimics its performance when incorporated within the inner mitochondrial membrane. The TM-SFM images (500 × 500 nm) in Figures 2 and 3, which are (21) Sellers, H., Ulman, A.; Shnidman, Y.; Eilers, J. E. J. Am. Chem. Soc. 1993, 115, 9389. (22) Hinkle, P. C.; Kim, J. J.; Racker, E. J. Biol. Chem. 1972, 247, 1338.
6630 Langmuir, Vol. 14, No. 23, 1998
Letters
Figure 2. TM-SFM images of cytochrome c oxidase immobilized in a lipid bilayer membrane on an electrode: (A) topographical image, (B) phase contrast image.
Figure 3. TM-SFM images of a lipid bilayer membrane modified electrode devoid of cytochrome c oxidase: (A) topographical image, (B) phase contrast image.
representative of characterizing several locations on the four samples that were examined, provide structural insights that qualitatively support this assessment. Figure 2 presents respective topographic and phase contrast images collected simultaneously for an oxidasecontaining LBM, whereas Figure 3 contains the analogous images for an LBM without confined oxidase. We note that the subsequent analysis of these images reflects several attributes of TM-SFM as one of the many variants of SFM. In particular, the small contact time between the tip and sample while imaging with TM-SFM is a much less invasive means (e.g., reduced tip-sample interactions) to characterize compliant surfaces (e.g., protein surfaces) in comparison with that when using contact mode SFM.23-25 Indeed, attempts to image these surfaces by contact mode SFM were not successful because of tipinduced sample deformation. Moreover, the detected phase shifts for the tip response in TM-SFM are strongly sensitive to differences in the chemical composition and/ or mechanical properties (e.g., elasticity) of surfaces but much less so to differences in topology.26 Phase contrast (23) Fritz, M.; Radmacher, M.; Cleveland, J. P.; Allersma, M. W.; Stewart, R. J.; Gieselmann, R.; Janmey, P.; Schmidt, C. F.; Hansma, P. K. Langmuir 1995, 11, 3529. (24) Tamayo, J.; Garcı´a, R. Langmuir 1996, 12, 4430. (25) Ikai, A. Surf. Sci. Rep. 1996, 26, 261.
imaging therefore provides a means to probe for differences in the material heterogeneity of a surface at nanometer length scales without significant contributions from surface topology. The topographic images show that the surfaces of both samples are microscopically rough, reflecting in large part the topology of the underlying electrode surface.5 The topographical image of the oxidase-containing LBM (Figure 2A), however, has several cone-shaped features. These features, many of which protrude by as much as 2 nm from the surrounding architecture, are not evident in images of oxidase-devoid LBMs (e.g., Figure 3A). We note that the difference in the roughness of the two sets of different images is representative of the range of the topologies found with these samples. There is also a notable difference in the phase contrast images for the two types of LBMs. The phase contrast image for the oxidase-containing membrane (Figure 2B) reveals the presence of large, continuous regions of a lower phase shift that surround small domains of a significantly larger phase shift. These small domains are absent in the phase contrast image for the oxidase-devoid LBM (Figure 3B). Furthermore, the locations of the small (26) Finot, M. O.; McDermott, M. T. J. Am. Chem. Soc. 1997, 119, 8564.
Letters
domains in the phase contrast image coincide with those for the cone-shaped features in the topographic image. We therefore attribute the cone-shape features to monomers and small aggregates of embedded oxidase, findings similar to those observed in platinum/carbon replicas of freeze-fractured vesicles by electron microscopy.13 The proposed origin of the difference between the two samples is also supported by the size of the cone-shaped features. The size of the smallest feature (∼14 nm diameter) is comparable to the expected 7-8 nm diameter of an individual oxidase,8 after accounting for distortions (e.g., magnification) that occur because of the size and shape of the probe tip.27 Control experiments using the same tip show that 10-nm and 30-nm gold particles appear to have more than doubled their size to 20-25 nm and 60-70 nm, respectively, upon imaging. Last, estimates of the size and coverage of the features in images over much larger surface areas indicate that the oxidase occupies ∼20% of the LBM.28 A reevaluation, then, of earlier kinetic data7 provides the first assessment of a lower limit for the minimum turnover rate that is required for the transition of the oxidase from its resting kinetic state to its pulsed kinetic state. This transition involves an increase in the rate of intermolecular electron transfer through the enzyme, i.e., from the Cua site to the heme a3-Cub site.1,14,29 While Antonini and co-workers have demonstrated that the turnover rate must be kept low to convert gradually the enzyme from its resting to pulsed kinetic state,14,29 our earlier results,7 coupled with the coverage estimates, indicate that a turnover rate of only ∼0.5 electrons/s is the lower limit required for the transition.7 (27) Vesenka, J.; Miller, R.; Henderson, E. Rev. Sci. Instrum. 1994, 65, 2249. (28) Estimates of coverage are based on an analysis of the phase contrast images for several different samples and sample locations, using both the magnitude and contour of the topology as diagnostics for discriminating the signal from the background noise. The images used for this analysis were typically hundreds of micrometers in size. The limiting turnover rate for the oxidase was then computed using this approximate coverage and the rate data are shown and discussed in detail in Figure 2 of ref 7. (29) Brunori, M.; Colosimo, A.; Rainoni, G.; Wilson, M. T.; Antonini, E. J. Biol. Chem. 1979, 254, 10796.
Langmuir, Vol. 14, No. 23, 1998 6631
Conclusions Taken together, these images demonstrate that the architecture of the oxidase-containing electrode-supported LBM mimics that envisioned for the inner mitochondrial membrane of mammalian systems. This model system, which allows fixed concentrations of cytochrome c to be reacted with the immobilized oxidase, permits kinetic measurements of the reaction between cytochrome c and the oxidase under low turnover rate conditions. The images yield an estimation of the amount of oxidase immobilized within the electrode-supported LBM that, when coupled with earlier kinetic data,7 permits a lower limit estimation of the minimum turnover rate required for transition of the enzyme from its resting kinetic state to its pulsed kinetic state. We believe that the results reported herein will enhance the development of functional models30,31 for this complex but essential enzyme. This work will also stimulate explorations of this type of assembly as a model for other systems and for interrogations using other experimental probes.32 Acknowledgment. Dr. Bertha King, Dr. Zoia Nikolaeva, and Professor Mikhail Smirnov are gratefully acknowledged for providing isolated cytochrome c oxidase samples. We also acknowledge the National Science Foundation (Grant NSF CHE-9508640), the Microanalytical Instrumentation Center of Iowa State University, and the Office of Basic Energy Science, Chemical Sciences Division of the U.S. Department of Energy for support of portions of this research. J. D. Burgess and V. W. Jones acknowledge the support of Postdoctoral Fellowships through the Institute of Physical Research and Technology of Iowa State University. The Ames Laboratory is operated for the U.S. Department of Energy by Iowa State University under Contract No. W-7405-eng-82. LA980648D (30) Wilson, M. T.; Peterson, J.; Antonini, E.; Brunori, M.; Colosimo, A.; Wyman, J. Proc. Natl. Acad. Sci. U.S.A. 1981, 78, 7115. (31) Brunori, M.; Wilson, M. T. Trends Biochem. Sci. 1982, 7, 295. (32) Salamon, Z.; Tollin, G. Biophys. J. 1996, 71, 858.
