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Atomic Force Microscopy of the Proteasome
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[34] Atomic Force Microscopy of the Proteasome By PAWEL A. OSMULSKI and MARIA GACZYNSKA Abstract
The proteasome should be an ideal molecule for studies on large enzymatic complexes, given its multisubunit and modular structure, compartmentalized design, numerous activities, and its own means of regulation. Considering the recent increased interest in the ubiquitin‐proteasome pathway, it is surprising that biophysical approaches to study this enzymatic assembly are applied with limited frequency. Methods including atomic force microscopy, fluorescence spectroscopy, surface plasmon resonance, and high‐pressure procedures all have gained popularity in characterization of the proteasome. These methods provide significant and often unexpected insight regarding the structure and function of the enzyme. This chapter describes the use of atomic force microscopy for dynamic structural studies of the proteasome. Introduction
The size of proteasomal assemblies and their degree of complication make them desirable but often challenging objects for biophysical studies. One such challenge stems from the fact that not only is the proteasome built from modules, but every module is also a multisubunit protein complex (Zwickl et al., 2001). The 19S cap or 11S activator attaches to the face of the eukaryotic catalytic 20S core. A gating mechanism guarding access to the catalytic chamber of the core, missing in the archaebacterial 20S proteasome, is also formed by the ring (Forster et al., 2003; Groll and Huber, 2003). The crystal structures of eukaryotic and archaebacterial core particles of the human activator complex and a structure of a hybrid core‐ activator complex have been solved; however, the most physiologically relevant 26S assembly continues to elude successful X‐ray analysis (Groll et al., 1997; Lowe et al., 1995; Whitby et al., 2000; Unno et al., 2002). It is reasonable to predict that giant protein assemblies such as the proteasomes exhibit a significant degree of structural dynamics, essential for their biological activity. The rotational symmetry of the proteasome resembles that of membrane channels or chaperonins. It has been established both experimentally with cation channels and theoretically using Monte Carlo simulations that multiprotomeric proteins exhibiting rotational symmetry demonstrate rapid, synchronized transitions between METHODS IN ENZYMOLOGY, VOL. 398 Copyright 2005, Elsevier Inc. All rights reserved.
0076-6879/05 $35.00 DOI: 10.1016/S0076-6879(05)98034-8
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conformations of all of their protomers (Duke et al., 2001). Apparently, the proteasome also exhibits these stability‐promoting transitions, although their amplitude is not as impressive as in the chaperonins (Horovitz et al., 2001). Analysis of crystal structures, despite all of the fundamental knowledge it offers, often falls short of providing a wealth of dynamic data. Although the giant proteasome remains beyond the reach of the nuclear magnetic resonance (NMR) technique, it is a convenient object for atomic force microscopy because its size is actually an advantage and its global conformation changes can be monitored relatively easily. Fluorescence spectroscopy and high‐pressure methods can also be used successfully with the proteasome. It is hoped that future research will see increased use of biophysical methods that are helpful to understanding the work of a biological nanodevice called the proteasome. Atomic Force Microscopy: An Overview of the Method
The atomic force microscope (AFM) was invented in 1986, the same year one of its inventors, Gerd Binnig, was awarded the Nobel Prize in Physics for his development of the scanning tunneling microscope (STM) in the early 1980s (Binnig, 1982, 1986). STM and AFM belong to a group of techniques currently growing in popularity, collectively known as scanning probe microscopy (SPM). Scanning probe microscopy is not strictly microscopy: it does not use lenses, and radiation is not transmitted through or reflected from the sample. Consequently, the wavelength of the radiation does not limit the resolution of the method. Instead, an SPM probe, which consists of a small and very sharp tip mounted on a cantilever, ‘‘feels’’ the surface of the sample. In the case of AFM, mechanical properties of the cantilever restrict the imaging accuracy. Deflection of a cantilever depends on the proximity of the sample and thus reflects the sample topography. Locking a sample/tip positioning system and a piezo element in a feedback loop facilitates constant corrections of the tip‐sample distance in a z (vertical) direction and allows for the creation of an image of the surface scanned in an x–y plane (Fotiadis et al., 2002; Yang et al., 2003). The changes in tip position are monitored in modern AFMs by a laser beam deflection method (Fig. 1). What makes AFM so attractive for biological applications is that the sample can be immersed in liquid during scanning. Atomic force microscopy has three major modes of operation: contact, oscillating, and force. In the contact, or DC mode (‘‘deflection of cantilever’’ or ‘‘deflection change’’), the tip and the atoms of the sample are in direct contact, causing the cantilever to deflect. The DC mode is rarely used with soft biological objects due to a potentially damaging shear force.
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FIG. 1. Schematics of AFM in the oscillating mode with a laser beam deflection detection method.
In the oscillating mode (AC, ‘‘amplitude of cantilever’’ or ‘‘amplitude change’’), the probe vibrates vertically while scanning. The proximity of the atoms of the sample causes changes in the amplitude of oscillations due to van der Waals and electrostatic forces. These vibrations are activated acoustically (‘‘tapping mode’’) or magnetically. The oscillating mode is well suited for imaging biological objects because the tip touches the sample only very briefly and with a low force (Hansma et al., 1994). Both contact and oscillating modes produce topography images with information about the height of the scanned objects, allowing the three‐dimensional map of the objects to be reconstructed and analyzed. The third AFM mode of operation, the force mode, also called force spectroscopy, does not provide an image of the sample. In this mode, the strength of the interaction between the tip and the sample is measured in the order of piconewtons (Bustamante et al., 1997; Santos and Castanho, 2004). For biological applications, the tip is usually modified with a receptor‐specific ligand. For successful imaging or force curve plotting, the sample must be immobilized on an atomically flat surface (‘‘AFM substrate’’). Muscovite
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mica is used as a substrate for most bioapplications, as a clean, flat, and negatively charged surface is obtained easily by peeling this layered mineral. Most proteins in physiological pH have enough surface positive charges to bind to mica by electrostatic forces, which are gentle enough to leave the protein in a native state and strong enough to keep it in place during oscillation mode scanning. Alternatively, the molecules can be attached to a modified surface of a substrate by cross‐linking, His‐tag binding, affinity binding, or embedding in a prepared lipid bilayer (see later). Atomic force microscopy samples can be imaged either in air (‘‘dry mode’’) or in liquid (‘‘liquid’’ or ‘‘wet’’ mode). For the dry mode, which is often used with nucleic acids, a small droplet is deposited on the substrate, washed with water after a few minutes, and dried in a stream of clean air or nitrogen. For ‘‘wet’’ imaging, both the probe and the sample are immersed in a liquid during scanning and permitting the addition or removal of ligands. Although sharpness of the images is somewhat compromised in liquid, it is still possible to achieve the practical lateral resolution of 1–2 nm and vertical resolution of 0.1–0.2 nm. With such a resolution, proteins up to 50 kDa will generally be imaged as rods or granules without additional structural features recognizable. Larger proteins may show some structural features, but it takes multisubunit protein complexes to enjoy the full power of AFM imaging. The unique advantage of the liquid mode is studying biologically active, native molecule. One limitation of AFM imaging is that a standard tip of 10 nm diameter at the very end will obviously not penetrate deep channels in protein assemblies, imaging holes as craters or shallow depressions. The temporal resolution of AFM reaches milliseconds for force measurements. When scanning, a tip needs milliseconds to run across a molecule (one‐dimensional imaging). A two‐dimensional image of a single molecule is created in a time scale of seconds to minutes. Examples of some studies on native proteins include GroEL/GroES chaperone, ion pumps and membrane channels, antibodies, ATP synthases, toxins, and photosystem proteins, in addition to others (Czajkowsky et al., 2000; Ehrenhofer et al., 1997; Fotiadis et al., 2002; Viani et al., 2000; Vie et al., 2001; Yang et al., 2003). Imaging of Proteasomes by Tapping Mode AFM in Liquid
Proteasomal assemblies, with their large sizes and easily recognizable shapes, are perfect subjects for AFM imaging. The major advantage of AFM is the possibility to observe biologically active molecules in real time with lateral resolution close to electron microscopy and with excellent vertical resolution. The limitations of AFM technique include (i) lateral resolution, poorer than in X‐ray crystallography or NMR spectroscopy;
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(ii) temporal resolution of standard topography imaging in the range of seconds and minutes; fortunately, the force mode does not suffer from this limitation; and (iii) inability to explore the molecule beyond its surface. Nevertheless, the relative ease of imaging and the availability of a dynamic aspect make the AFM a very valuable technique for studies on the proteasome. The method to image randomly dispersed, electrostatically attached to mica 20S particles is described later. Such particles are perfect subjects for studies of conformational dynamics. Equipment and Supplies We use a Nanoscope IIIa microscope (Digital Instruments) with an E scanner and a glass cantilever holder for scanning in fluids (‘‘liquid cell,’’ ‘‘wet chamber’’). The Nanoscope IIIa uses acoustically activated vibrations. Molecular Imaging Corp. offers similar microscopes with magnetically activated vibrations. Muscovite mica glued with a two‐component epoxy resin (available from hardware or craft stores) to a steel disk serves as the AFM substrate. The steel disks and the mica in sheets or ready‐to‐ use disks are available from electron microscopy suppliers. If the mica is purchased as inexpensive sheets, it can be cut with scissors into squares of roughly the size of steel disks, or mica disks can be punched out from the sheet with a large‐diameter hole puncher. The steel disks with mica should be prepared in advance. It is convenient to glue several dozens of mica pieces at a time, wait for at least 24 h for the resin to cure, and store the ready‐to‐use substrates in a dust‐free container. The fresh surface of a mica should be exposed just before the experiment, no more than about a minute before deposition of a protein solution. Removal of an external layer of mica can be achieved by peeling it out with a sticky tape (Scotch‐ type, pressure‐sensitive). If the exposed surface is cracked, the next layer should be peeled out. The substrate can be reused for as long as a fresh surface of mica can be exposed. When the mica is used up, the steel disk should be cleaned with acetone to remove remaining glue before attaching another piece of mica. For imaging proteasomes in the tapping mode in liquid, we use oxide‐ sharpened silicon probes (NP‐ST or NP‐STT) on cantilevers with a nominal spring constant of 0.32 N/m (Digital Instruments). Other probes of similar parameters include OTR8 and ORC8 (Olympus) and CSC (MicroMash). Not all tips on probes are of equal quality, which is apparent when imaging an AFM standard grid or any sample of known features. It is advisable to discard blunt tips and to use only the sharpest for the best results. Standard grids, or calibration gratings (several kinds are available from MicroMash), are usually made from silicon and represent arrays of steps, tips, or
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pillars of well‐defined, micrometer‐scale dimensions. They are used for tip evaluation and for calibration of piezoelectric scanners. Sample Preparation and Scanning For AFM imaging, purified proteasomes are diluted in 5 mM Tris‐HCl buffer, pH 7.0 (imaging buffer), to give a nanomolar concentration. A 50 mM buffer can be used with similar results. It is advisable to filter all buffers with a 0.2‐m filter and to perform all manipulations in a clean air hood (PCR hood) to avoid contamination with dust particles. The protein preparation has to be as pure as possible, as any contaminant protein will be imaged as well as the proteasome. A 2‐l droplet of the diluted proteasomes is deposited on a freshly cleaved mica surface. After a few minutes of binding, but before drying of the droplet, the sample should be overlaid with about 30 l of the imaging buffer and mounted in the head of the microscope. Additional buffer, enough to fill a cell, is needed when a liquid cell is used with O rings to seal off the liquid and to enable an easy exchange of buffers. Resonant frequency of the tip is manually tuned to 9–10 kHz, with the best results obtained for frequency slightly below the actual resonant peak. The amplitude of the tip ranges from 200 to 500 mV and set point ranges from 1.4 to 2 V. The higher the set point, the lower the force of tapping. Integral gain should be set at the highest value not causing distortion of the image (‘‘ringing’’), usually 0.2–0.4. Fields of 1 m2 or smaller are scanned at rates of 2–3 Hz with the highest available resolution of 512 512 pixels. It is advisable to collect both trace and retrace images (the probe moving left to right or right to left while scanning), as sometimes objects blurred in the image created in one direction will be acceptable in the other image. Ideally, the proteasome particles should be abundantly present in a scanned field, but should not touch each other. If desired, a monomolecular layer of proteasomes can be obtained by increasing the concentration of sample deposited on a mica. The proteasomes are stable during subsequent scans; they do not change orientation and location. The lateral drift reaches only about 10 nm per scan and it is possible to image the same particle for more than an hour. Ligands (substrates, inhibitors) or extra buffer can be added to the sample with a pipette through openings in the cantilever holder, even if O rings are not used. The ligands are not detected by the probe unless they are large enough (at least in the range of a few thousands daltons) and immobilized on the surface of a studied particle or, rather unlikely, on the surface of mica (Fig. 2B). After prolonged scanning, the quality of the image may decline. This will be most likely the effect of a tip losing its sharpness due to wearing down or contamination. A worn‐down tip should be discarded. A contaminated tip
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FIG. 2. Examples of AFM images of proteasomes. (A) Side plots (tilted topography images) of 20S proteasomes from fission yeast (Schizosaccharomyces pombe). (Left to right) Top‐view particle in the closed conformation, side‐view particle in the barrel conformation, open particle in the top view, and side‐view cylinder. (B) Images of the human 26S proteasomes. (Left) A typical particle with two caps and the 20S core recognizable. (Right) A particle touching, presumably bound to, the 19S cap is ubiquitinated lysozyme, which was added to the mica‐attached 26S proteasomes. The ubiquitinated protein appears unusually large, most likely because it was not attached stably to the mica and shifted during scanning. (C) A side plot of a fragment of a field of 20S proteasomes from Saccharomyces cerevisiae. The particles were distributed randomly on a mica substrate. The lightest shades of gray correspond to the highest‐positioned areas.
can be cleaned by rinsing in ultrapure water or by irradiation with ultraviolet light for a few hours; however, the cleaning is not always successful. An obvious sign of tip contamination is a ‘‘double image’’: all topographical features are repeated in regular intervals. This is the result of a contaminating particle attached to the AFM tip and acting as a second tip. This ‘‘second tip’’ is often removable by ‘‘shaking off’’ the contaminant during several seconds of a fast scanning (at least 4 Hz) without engagement with the sample. Buffers with high ionic strength are sometimes used to improve imaging due to a reduction of electrostatic forces between a tip and a sample. Addition of NaCl up to 200 mM during scanning, however, does not influence the results for proteasomes (Osmulski and Gaczynska, 2000).
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Image Processing Each company producing STM microscopes also supplies specialized software capable of collecting and, to varying degrees, manipulating and analyzing data. There are also numerous stand‐alone software packages for reading data in several formats, processing the images, performing feature analysis (e.g., contour recognition, particle counting, profiling). and preparing images for publication. None of them is perfect in all of the aforementioned aspects, and the reader is strongly encouraged to test several sources to find the software that fits the particular needs. Although the existence of multiple data formats does not help this endeavor, most provide free trials and there are several good packages in the public domain. The most comprehensive collection of the imaging software with short descriptions of their capabilities is provided by the following Web pages: http://www.microscopy.info (MC Services) http://www.weizmann.ac.il/Chemical Research Support/surflab/peter/ software/index.html (Weizmann Institute of Science, Israel) http://web.mit.edu/cortiz/www/Software.html (Ortiz Laboratory at MIT, Cambridge, MA) http://www.uksaf.org/software.html (UK Surface Analysis Forum) These sites also list ample academic and commercial links to groups and companies working with STM images. From our perspective, two packages are the best for both the beginner and the advanced user: Image SXM by Steve Barrett (http://reg.ssci.liv.ac.uk) and ImageJ (http://rsb.info. nih.gov/ij/). They are available in most platforms, simple to use, comprehensive in their capabilities, provide an open source code, and are freely available. The topographical images are presented in false color scale, where a particular color or shade corresponds to a height of the sample. The amber and gray‐scale color palettes are the most popular. Tilted images (side plots; see Fig. 2) are often presented to better show particular features of objects. The collected images often require processing, as they show a certain level of tilting of a background surface and electronic and thermal noise. The most pronounced but easy to remove artifacts arise from peculiar properties of a piezo element and shape of the tip. For this purpose, a standard image processing consisting of flattening and plain fit provided by the Nanoscope software is applied. The lateral dimensions of particles are measured with the help of a cross‐section option (Nanoscope, others). Because the tip cannot reproduce steep banks of small particles well, it is necessary to consider that the dimensions will be enlarged due to a
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tip‐broadening effect. This effect can be removed conveniently and additional processing performed with software such as Image SXM. Section analysis in Nanoscope software, SXM Image, and others used for measuring particles. The Proteasome under AFM: Observations
Proteasome molecules imaged by the aforementioned method revealed a high level of dynamics when imaged with the practical resolution of 1–2 nm (Fig. 2). After analysis of AFM images performed under different conditions, it was possible to correlate the dynamic behavior of the proteasomes with their catalytic actions, to monitor a ligand binding, and to develop a model of allosteric signaling between the active centers and the gate (Gaczynska et al., 2003; Osmulski and Gaczynska, 2000, 2002). The rings of particles in the top‐view orientation were either cone shaped or had a crater‐like cavity in the middle, positioned where the entrance to the central channel was expected to be guarded by the gate. Therefore, we named the conformants ‘‘closed’’ and ‘‘open,’’ respectively, hypothesizing that they represent the closed and open gate to the proteasomal central channel (Fig. 2A; Osmulski and Gaczynska, 2000). This hypothesis was supported by the observation that images of top‐view proteasomes from Thermoplasma acidophilum, which do not have a gate in the ring, showed exclusively open particles (P. A. Osmulski et al., submitted; proteasomes kindly provided by A. L. Goldberg). In contrast, when eukaryotic proteasomes were analyzed, both the closed and the open conformants were present in all scans, and all of the particles were able to switch conformations between subsequent scans. Addition of peptide or protein substrates changed the partition between closed and open forms (Osmulski and Gaczynska, 2000). The conformational changes were observed in side‐ view proteasomes as well, with a partition between barrel‐shaped and cylinder‐shaped molecules closely following the partition between closed and open conformants (Osmulski and Gaczynska, 2002). These results prompted us to hypothesize that a ‘‘closed barrel’’ and an ‘‘open cylinder’’ represent allosteric forms analogous to R (relaxed) and T (tense) forms in a classical two‐state model of allosteric transitions. We have found that gate opening observed under AFM is coupled with a tetrahedral transition state in any of the active centers. Interestingly, kinetic parameters calculated on the basis of the open/closed partition as a function of substrate concentration were in perfect agreement with the parameters obtained in the traditional ‘‘macroscopic’’ way, with the increase in a product concentration measured instead of the abundance of conformers (P. A. Osmulski et al.,
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submitted). Core particles isolated from human and mouse cultured cells, rabbit muscle, Saccharomyces cerevisiae, or Schizosaccharomyces pombe were undistinguishable in AFM images and revealed the same dynamic behavior (Gaczynska et al., 2003; P. A. Osmulski and M. Gaczynska, unpublished observations). Analysis of AFM images is very useful for the characterization of ligand binding to the proteasomes. This was shown on the example of proline‐ and arginine‐rich peptides (PR peptides; see Gaczynska and Osmulski, 2005). These peptides bind to the 20S core particles, presumably to the ring. AFM images revealed that the 20S proteasome treated with the peptides was unable to switch between open and closed conformers and remained in a quasi‐open form permanently (Gaczynska et al., 2003). The 20S core particles are not the only proteasomal assemblies imaged under AFM. Studies of the interactions of human 26S complexes with PR peptides showed dramatic conformational changes in the inhibitor‐treated proteasomes (Gaczynska et al., 2003). The yeast 26S assemblies (Fig. 2B) and human proteasomes complexed with the 11S activator or the PI31 inhibitor (kindly provided by G. DeMartino) were visualized as well (P. A. Osmulski and M. Gaczynska, unpublished observations). The power of AFM, however, extends well beyond simple topography imaging, and more applications are waiting to be implemented in proteasomal studies. The force measurements, for example, will be very useful to study binding of the ubiquitinated substrates to 19S cap, their defolding, and translocation to the core particle. In addition to the aforementioned method of imaging electrostatically attached particles, several studies have explored the immobilized and organized 20S complexes under AFM. For example, proteasomes from archaeon T. acidophilum with histidine tags introduced at the C terminus of the subunit were specifically attached to a mica‐supported, ultraflat chelator lipid membrane (Dorn et al., 1999). As a result, images were obtained of a dense layer of the 20S particles oriented in a side‐view position. Interactions of the core particles with different supported lipid bilayers were explored for bovine proteasomes (Furuike et al., 2003). In another study, contact mode AFM was used to successfully image the shear‐resistant two‐dimensional crystals of T. acidophilum proteasomes (Thess et al., 2002). The 20S proteasomes His6‐tagged at the N termini of subunits crystallized readily on a nickel‐chelating lipid bilayer prepared on mica and optimized their packing in a crystal by interlocking both laterally and vertically (Thess et al., 2002). The immobilized and organized proteasomes constitute very useful research objects: however, the dynamic information may be, in most parts, lost.
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Acknowledgments The work was funded by R01 GM069819 (M.G.) and San Antonio Cancer Institute grants (P. A. O., M. G.).
References Binnig, G., Quate, C. F., and Gerber, C. H. (1986). Atomic force microscope. Phys. Rev. Lett. 56, 930–933. Binnig, G., and Rohrer, H. (1982). Scanning tunneling microscopy. Helv. Phys. Acta 55, 726–735. Bustamante, C., Rivetti, C., and Keller, D. J. (1997). Scanning force microscopy under aqueous solutions. Curr. Opinion Struct. Biol. 7, 709–716. Czajkowsky, D. M., Iwamoto, H., and Shao, Z. (2000). Atomic force microscopy in structural biology: From the subcellular to the submolecular. J. Electr. Micr. 49, 395–406. Dorn, I. T., Eschrich, R., Seemuller, E., Guckenberger, R., and Tampe, R. (1999). High‐ resolution AFM‐imaging and mechanistic analysis of the 20 S proteasome. J. Mol. Biol. 288, 1027–1036. Duke, T. A., Novere, N. L., and Bray, D. (2001). Conformational spread in a ring of proteins: A stochastic approach to allostery. J. Mol. Biol. 308, 541–553. Ehrenhofer, U., Rakowska, A., Schneider, S. W., Schwab, A., and Oberleithner, H. (1997). The atomic force microscope detects ATP‐sensitive protein clusters in the plasma membrane of transformed MDCK cells. Cell Biol. Int. 21, 737–746. Forster, A., Whitby, F. G., and Hill, C. P. (2003). The pore of activated 20S proteasomes has an ordered 7‐fold symmetric conformation. EMBO J. 22, 4356–4364. Fotiadis, D., Scheuring, S., Muller, S. A., Engel, A., and Muller, D. J. (2002). Imaging and manipulation of biological structures with the AFM. Micron 33, 385–397. Furuike, S., Hirokawa, J., Yamada, S., and Yamazaki, M. (2003). Atomic force microscopy studies of interaction of the 20S proteasome with supported lipid bilayers. Biochim. Biophys. Acta 1615, 1–6. Gaczynska, M., and Osmulski, P. A. (2005). Characterization of noncompetitive regulators of proteasome activity. Methods Enzymol. 398[35] 2005 (this volume). Gaczynska, M., Osmulski, P. A., Gao, Y., Post, M. J., and Simons, M. (2003). Proline‐ and arginine‐rich peptides constitute a novel class of allosteric inhibitors of proteasome activity. Biochemistry 42, 8663–8670. Groll, M., and Huber, R. (2003). Substrate access and processing by the 20S proteasome core particle. Int. J. Bioch. Cell Biol. 35, 606–616. Groll, M., Ditzel, L., Lowe, J., Stock, D., Bochtler, M., Bartunik, H. D., and Huber, R. (1997). Structure of 20S proteasome from yeast at 2.4 A resolution. Nature 386, 463–471. Hansma, P. K., Cleveland, J. P., Radmacher, M., Walters, D. A., Hillner, P. E., Bezanilla, M., Fritz, M., Vie, D., Hansma, H. G., Prater, C. B., Massie, J., Fukunaga, L., Gurley, J., and Elings, V. (1994). Tapping mode atomic force microscopy in liquids. Appl. Phys. Lett. 64, 1738–1740. Horovitz, A., Fridmann, Y., Kafri, G., and Yifrach, O. (2001). Review: Allostery in chaperonins. J. Struct. Biol. 135, 104–114. Lowe, J., Stock, D., Jap, B., Zwickl, P., Baumeister, W., and Huber, R. (1995). Crystal structure of the 20S proteasome from the archaeon T. acidophilum at 3.4 A resolution. Science 268, 533–539.
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Osmulski, P. A., and Gaczynska, M. (2000). Atomic force microscopy reveals two conformations of the 20S proteasome from fission yeast. J. Biol. Chem. 275, 13171–13174. Osmulski, P. A., and Gaczynska, M. (2002). Nanoenzymology of the 20S proteasome: Proteasomal actions are controlled by the allosteric transition. Biochemistry 41, 7047–7053. Santos, N. C., and Castanho, M. A. (2004). An overview of the biophysical applications of atomic force microscopy. Biophys. Chem. 107, 133–149. Thess, A., Hutschenreiter, S., Hofmann, M., Tampe, R., Baumeister, W., and Guckenberger, R. (2002). Specific orientation and two‐dimensional crystallization of the proteasome at metal‐chelating lipid interfaces. J. Biol. Chem. 277, 36321–36328. Unno, M., Mizushima, T., Morimoto, Y., Tomisugi, Y., Tanaka, K., Yasuoka, N., and Tsukihara, T. (2002). Structure determination of the constitutive 20S proteasome from bovine liver at 2.75 A resolution. J. Biochem. 131, 171–173. Viani, M. B., Pietrasanta, L. I., Thompson, J. B., Chand, A., Gebeshuber, I. C., Kindt, J. H., Richter, M., Hansma, H. G., and Hansma, P. K. (2000). Probing protein‐protein interactions in real time. Nature Struct. Biol. 7, 644–647. Vie, V., Van Mau, N., Pomarede, P., Dance, C., Schwartz, J. L., Laprade, R., Frutos, R., Rang, C., Masson, L., Heitz, F., and Le Grimellec, C. (2001). Lipid‐induced pore formation of the Bacillus thuringiensis Cry1Aa insecticidal toxin. J. Membr. Biol. 180, 195–203. Whitby, F. G., Masters, E. I., Kramer, L., Knowlton, J. R., Yao, Y., Wang, C. C., and Hill, C. P. (2000). Structural basis for the activation of 20S proteasomes by 11S regulators. Nature 408, 115–120. Yang, Y., Wang, H., and Erie, D. A. (2003). Quantitative characterization of biomolecular assemblies and interactions using atomic force microscopy. Methods 29, 175–187. Zwickl, P., Seemuller, E., Kapelari, B., and Baumeister, W. (2001). The proteasome: A supramolecular assembly designed for controlled proteolysis. Adv. Prot. Chem. 59, 187–222.
[35] Characterization of Noncompetitive Regulators of Proteasome Activity By MARIA GACZYNSKA and PAWEL A. OSMULSKI Abstract
The success of bortezomib, a competitive proteasome inhibitor and a drug approved to treat multiple myeloma, spurred interest in compounds targeting catalytic sites of the enzyme. The aim of this chapter, however, is to focus attention on the small molecule, natural or synthetic compounds binding far away from the catalytic centers, yet modifying the performance of the proteasome. Defining allostery broadly as any kind of ligand‐ induced, long‐distance transfer of conformational signals within a molecule, most such compounds are allosteric effectors capable of regulating the proteasome in vitro and in vivo in a manner more diverse and precise than competitive inhibitors. Proline‐ and arginine‐rich peptides (PR peptides) are examples of such compounds and are currently being considered as METHODS IN ENZYMOLOGY, VOL. 398 Copyright 2005, Elsevier Inc. All rights reserved.
0076-6879/05 $35.00 DOI: 10.1016/S0076-6879(05)98035-X
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[34] Atomic Force Microscopy of the Proteasome By PAWEL A. OSMULSKI and MARIA GACZYNSKA Abstract
The proteasome should be an ideal molecule for studies on large enzymatic complexes, given its multisubunit and modular structure, compartmentalized design, numerous activities, and its own means of regulation. Considering the recent increased interest in the ubiquitin‐proteasome pathway, it is surprising that biophysical approaches to study this enzymatic assembly are applied with limited frequency. Methods including atomic force microscopy, fluorescence spectroscopy, surface plasmon resonance, and high‐pressure procedures all have gained popularity in characterization of the proteasome. These methods provide significant and often unexpected insight regarding the structure and function of the enzyme. This chapter describes the use of atomic force microscopy for dynamic structural studies of the proteasome. Introduction
The size of proteasomal assemblies and their degree of complication make them desirable but often challenging objects for biophysical studies. One such challenge stems from the fact that not only is the proteasome built from modules, but every module is also a multisubunit protein complex (Zwickl et al., 2001). The 19S cap or 11S activator attaches to the face of the eukaryotic catalytic 20S core. A gating mechanism guarding access to the catalytic chamber of the core, missing in the archaebacterial 20S proteasome, is also formed by the ring (Forster et al., 2003; Groll and Huber, 2003). The crystal structures of eukaryotic and archaebacterial core particles of the human activator complex and a structure of a hybrid core‐ activator complex have been solved; however, the most physiologically relevant 26S assembly continues to elude successful X‐ray analysis (Groll et al., 1997; Lowe et al., 1995; Whitby et al., 2000; Unno et al., 2002). It is reasonable to predict that giant protein assemblies such as the proteasomes exhibit a significant degree of structural dynamics, essential for their biological activity. The rotational symmetry of the proteasome resembles that of membrane channels or chaperonins. It has been established both experimentally with cation channels and theoretically using Monte Carlo simulations that multiprotomeric proteins exhibiting rotational symmetry demonstrate rapid, synchronized transitions between METHODS IN ENZYMOLOGY, VOL. 398 Copyright 2005, Elsevier Inc. All rights reserved.
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conformations of all of their protomers (Duke et al., 2001). Apparently, the proteasome also exhibits these stability‐promoting transitions, although their amplitude is not as impressive as in the chaperonins (Horovitz et al., 2001). Analysis of crystal structures, despite all of the fundamental knowledge it offers, often falls short of providing a wealth of dynamic data. Although the giant proteasome remains beyond the reach of the nuclear magnetic resonance (NMR) technique, it is a convenient object for atomic force microscopy because its size is actually an advantage and its global conformation changes can be monitored relatively easily. Fluorescence spectroscopy and high‐pressure methods can also be used successfully with the proteasome. It is hoped that future research will see increased use of biophysical methods that are helpful to understanding the work of a biological nanodevice called the proteasome. Atomic Force Microscopy: An Overview of the Method
The atomic force microscope (AFM) was invented in 1986, the same year one of its inventors, Gerd Binnig, was awarded the Nobel Prize in Physics for his development of the scanning tunneling microscope (STM) in the early 1980s (Binnig, 1982, 1986). STM and AFM belong to a group of techniques currently growing in popularity, collectively known as scanning probe microscopy (SPM). Scanning probe microscopy is not strictly microscopy: it does not use lenses, and radiation is not transmitted through or reflected from the sample. Consequently, the wavelength of the radiation does not limit the resolution of the method. Instead, an SPM probe, which consists of a small and very sharp tip mounted on a cantilever, ‘‘feels’’ the surface of the sample. In the case of AFM, mechanical properties of the cantilever restrict the imaging accuracy. Deflection of a cantilever depends on the proximity of the sample and thus reflects the sample topography. Locking a sample/tip positioning system and a piezo element in a feedback loop facilitates constant corrections of the tip‐sample distance in a z (vertical) direction and allows for the creation of an image of the surface scanned in an x–y plane (Fotiadis et al., 2002; Yang et al., 2003). The changes in tip position are monitored in modern AFMs by a laser beam deflection method (Fig. 1). What makes AFM so attractive for biological applications is that the sample can be immersed in liquid during scanning. Atomic force microscopy has three major modes of operation: contact, oscillating, and force. In the contact, or DC mode (‘‘deflection of cantilever’’ or ‘‘deflection change’’), the tip and the atoms of the sample are in direct contact, causing the cantilever to deflect. The DC mode is rarely used with soft biological objects due to a potentially damaging shear force.
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FIG. 1. Schematics of AFM in the oscillating mode with a laser beam deflection detection method.
In the oscillating mode (AC, ‘‘amplitude of cantilever’’ or ‘‘amplitude change’’), the probe vibrates vertically while scanning. The proximity of the atoms of the sample causes changes in the amplitude of oscillations due to van der Waals and electrostatic forces. These vibrations are activated acoustically (‘‘tapping mode’’) or magnetically. The oscillating mode is well suited for imaging biological objects because the tip touches the sample only very briefly and with a low force (Hansma et al., 1994). Both contact and oscillating modes produce topography images with information about the height of the scanned objects, allowing the three‐dimensional map of the objects to be reconstructed and analyzed. The third AFM mode of operation, the force mode, also called force spectroscopy, does not provide an image of the sample. In this mode, the strength of the interaction between the tip and the sample is measured in the order of piconewtons (Bustamante et al., 1997; Santos and Castanho, 2004). For biological applications, the tip is usually modified with a receptor‐specific ligand. For successful imaging or force curve plotting, the sample must be immobilized on an atomically flat surface (‘‘AFM substrate’’). Muscovite
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mica is used as a substrate for most bioapplications, as a clean, flat, and negatively charged surface is obtained easily by peeling this layered mineral. Most proteins in physiological pH have enough surface positive charges to bind to mica by electrostatic forces, which are gentle enough to leave the protein in a native state and strong enough to keep it in place during oscillation mode scanning. Alternatively, the molecules can be attached to a modified surface of a substrate by cross‐linking, His‐tag binding, affinity binding, or embedding in a prepared lipid bilayer (see later). Atomic force microscopy samples can be imaged either in air (‘‘dry mode’’) or in liquid (‘‘liquid’’ or ‘‘wet’’ mode). For the dry mode, which is often used with nucleic acids, a small droplet is deposited on the substrate, washed with water after a few minutes, and dried in a stream of clean air or nitrogen. For ‘‘wet’’ imaging, both the probe and the sample are immersed in a liquid during scanning and permitting the addition or removal of ligands. Although sharpness of the images is somewhat compromised in liquid, it is still possible to achieve the practical lateral resolution of 1–2 nm and vertical resolution of 0.1–0.2 nm. With such a resolution, proteins up to 50 kDa will generally be imaged as rods or granules without additional structural features recognizable. Larger proteins may show some structural features, but it takes multisubunit protein complexes to enjoy the full power of AFM imaging. The unique advantage of the liquid mode is studying biologically active, native molecule. One limitation of AFM imaging is that a standard tip of 10 nm diameter at the very end will obviously not penetrate deep channels in protein assemblies, imaging holes as craters or shallow depressions. The temporal resolution of AFM reaches milliseconds for force measurements. When scanning, a tip needs milliseconds to run across a molecule (one‐dimensional imaging). A two‐dimensional image of a single molecule is created in a time scale of seconds to minutes. Examples of some studies on native proteins include GroEL/GroES chaperone, ion pumps and membrane channels, antibodies, ATP synthases, toxins, and photosystem proteins, in addition to others (Czajkowsky et al., 2000; Ehrenhofer et al., 1997; Fotiadis et al., 2002; Viani et al., 2000; Vie et al., 2001; Yang et al., 2003). Imaging of Proteasomes by Tapping Mode AFM in Liquid
Proteasomal assemblies, with their large sizes and easily recognizable shapes, are perfect subjects for AFM imaging. The major advantage of AFM is the possibility to observe biologically active molecules in real time with lateral resolution close to electron microscopy and with excellent vertical resolution. The limitations of AFM technique include (i) lateral resolution, poorer than in X‐ray crystallography or NMR spectroscopy;
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(ii) temporal resolution of standard topography imaging in the range of seconds and minutes; fortunately, the force mode does not suffer from this limitation; and (iii) inability to explore the molecule beyond its surface. Nevertheless, the relative ease of imaging and the availability of a dynamic aspect make the AFM a very valuable technique for studies on the proteasome. The method to image randomly dispersed, electrostatically attached to mica 20S particles is described later. Such particles are perfect subjects for studies of conformational dynamics. Equipment and Supplies We use a Nanoscope IIIa microscope (Digital Instruments) with an E scanner and a glass cantilever holder for scanning in fluids (‘‘liquid cell,’’ ‘‘wet chamber’’). The Nanoscope IIIa uses acoustically activated vibrations. Molecular Imaging Corp. offers similar microscopes with magnetically activated vibrations. Muscovite mica glued with a two‐component epoxy resin (available from hardware or craft stores) to a steel disk serves as the AFM substrate. The steel disks and the mica in sheets or ready‐to‐ use disks are available from electron microscopy suppliers. If the mica is purchased as inexpensive sheets, it can be cut with scissors into squares of roughly the size of steel disks, or mica disks can be punched out from the sheet with a large‐diameter hole puncher. The steel disks with mica should be prepared in advance. It is convenient to glue several dozens of mica pieces at a time, wait for at least 24 h for the resin to cure, and store the ready‐to‐use substrates in a dust‐free container. The fresh surface of a mica should be exposed just before the experiment, no more than about a minute before deposition of a protein solution. Removal of an external layer of mica can be achieved by peeling it out with a sticky tape (Scotch‐ type, pressure‐sensitive). If the exposed surface is cracked, the next layer should be peeled out. The substrate can be reused for as long as a fresh surface of mica can be exposed. When the mica is used up, the steel disk should be cleaned with acetone to remove remaining glue before attaching another piece of mica. For imaging proteasomes in the tapping mode in liquid, we use oxide‐ sharpened silicon probes (NP‐ST or NP‐STT) on cantilevers with a nominal spring constant of 0.32 N/m (Digital Instruments). Other probes of similar parameters include OTR8 and ORC8 (Olympus) and CSC (MicroMash). Not all tips on probes are of equal quality, which is apparent when imaging an AFM standard grid or any sample of known features. It is advisable to discard blunt tips and to use only the sharpest for the best results. Standard grids, or calibration gratings (several kinds are available from MicroMash), are usually made from silicon and represent arrays of steps, tips, or
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pillars of well‐defined, micrometer‐scale dimensions. They are used for tip evaluation and for calibration of piezoelectric scanners. Sample Preparation and Scanning For AFM imaging, purified proteasomes are diluted in 5 mM Tris‐HCl buffer, pH 7.0 (imaging buffer), to give a nanomolar concentration. A 50 mM buffer can be used with similar results. It is advisable to filter all buffers with a 0.2‐m filter and to perform all manipulations in a clean air hood (PCR hood) to avoid contamination with dust particles. The protein preparation has to be as pure as possible, as any contaminant protein will be imaged as well as the proteasome. A 2‐l droplet of the diluted proteasomes is deposited on a freshly cleaved mica surface. After a few minutes of binding, but before drying of the droplet, the sample should be overlaid with about 30 l of the imaging buffer and mounted in the head of the microscope. Additional buffer, enough to fill a cell, is needed when a liquid cell is used with O rings to seal off the liquid and to enable an easy exchange of buffers. Resonant frequency of the tip is manually tuned to 9–10 kHz, with the best results obtained for frequency slightly below the actual resonant peak. The amplitude of the tip ranges from 200 to 500 mV and set point ranges from 1.4 to 2 V. The higher the set point, the lower the force of tapping. Integral gain should be set at the highest value not causing distortion of the image (‘‘ringing’’), usually 0.2–0.4. Fields of 1 m2 or smaller are scanned at rates of 2–3 Hz with the highest available resolution of 512 512 pixels. It is advisable to collect both trace and retrace images (the probe moving left to right or right to left while scanning), as sometimes objects blurred in the image created in one direction will be acceptable in the other image. Ideally, the proteasome particles should be abundantly present in a scanned field, but should not touch each other. If desired, a monomolecular layer of proteasomes can be obtained by increasing the concentration of sample deposited on a mica. The proteasomes are stable during subsequent scans; they do not change orientation and location. The lateral drift reaches only about 10 nm per scan and it is possible to image the same particle for more than an hour. Ligands (substrates, inhibitors) or extra buffer can be added to the sample with a pipette through openings in the cantilever holder, even if O rings are not used. The ligands are not detected by the probe unless they are large enough (at least in the range of a few thousands daltons) and immobilized on the surface of a studied particle or, rather unlikely, on the surface of mica (Fig. 2B). After prolonged scanning, the quality of the image may decline. This will be most likely the effect of a tip losing its sharpness due to wearing down or contamination. A worn‐down tip should be discarded. A contaminated tip
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FIG. 2. Examples of AFM images of proteasomes. (A) Side plots (tilted topography images) of 20S proteasomes from fission yeast (Schizosaccharomyces pombe). (Left to right) Top‐view particle in the closed conformation, side‐view particle in the barrel conformation, open particle in the top view, and side‐view cylinder. (B) Images of the human 26S proteasomes. (Left) A typical particle with two caps and the 20S core recognizable. (Right) A particle touching, presumably bound to, the 19S cap is ubiquitinated lysozyme, which was added to the mica‐attached 26S proteasomes. The ubiquitinated protein appears unusually large, most likely because it was not attached stably to the mica and shifted during scanning. (C) A side plot of a fragment of a field of 20S proteasomes from Saccharomyces cerevisiae. The particles were distributed randomly on a mica substrate. The lightest shades of gray correspond to the highest‐positioned areas.
can be cleaned by rinsing in ultrapure water or by irradiation with ultraviolet light for a few hours; however, the cleaning is not always successful. An obvious sign of tip contamination is a ‘‘double image’’: all topographical features are repeated in regular intervals. This is the result of a contaminating particle attached to the AFM tip and acting as a second tip. This ‘‘second tip’’ is often removable by ‘‘shaking off’’ the contaminant during several seconds of a fast scanning (at least 4 Hz) without engagement with the sample. Buffers with high ionic strength are sometimes used to improve imaging due to a reduction of electrostatic forces between a tip and a sample. Addition of NaCl up to 200 mM during scanning, however, does not influence the results for proteasomes (Osmulski and Gaczynska, 2000).
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Image Processing Each company producing STM microscopes also supplies specialized software capable of collecting and, to varying degrees, manipulating and analyzing data. There are also numerous stand‐alone software packages for reading data in several formats, processing the images, performing feature analysis (e.g., contour recognition, particle counting, profiling). and preparing images for publication. None of them is perfect in all of the aforementioned aspects, and the reader is strongly encouraged to test several sources to find the software that fits the particular needs. Although the existence of multiple data formats does not help this endeavor, most provide free trials and there are several good packages in the public domain. The most comprehensive collection of the imaging software with short descriptions of their capabilities is provided by the following Web pages: http://www.microscopy.info (MC Services) http://www.weizmann.ac.il/Chemical Research Support/surflab/peter/ software/index.html (Weizmann Institute of Science, Israel) http://web.mit.edu/cortiz/www/Software.html (Ortiz Laboratory at MIT, Cambridge, MA) http://www.uksaf.org/software.html (UK Surface Analysis Forum) These sites also list ample academic and commercial links to groups and companies working with STM images. From our perspective, two packages are the best for both the beginner and the advanced user: Image SXM by Steve Barrett (http://reg.ssci.liv.ac.uk) and ImageJ (http://rsb.info. nih.gov/ij/). They are available in most platforms, simple to use, comprehensive in their capabilities, provide an open source code, and are freely available. The topographical images are presented in false color scale, where a particular color or shade corresponds to a height of the sample. The amber and gray‐scale color palettes are the most popular. Tilted images (side plots; see Fig. 2) are often presented to better show particular features of objects. The collected images often require processing, as they show a certain level of tilting of a background surface and electronic and thermal noise. The most pronounced but easy to remove artifacts arise from peculiar properties of a piezo element and shape of the tip. For this purpose, a standard image processing consisting of flattening and plain fit provided by the Nanoscope software is applied. The lateral dimensions of particles are measured with the help of a cross‐section option (Nanoscope, others). Because the tip cannot reproduce steep banks of small particles well, it is necessary to consider that the dimensions will be enlarged due to a
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tip‐broadening effect. This effect can be removed conveniently and additional processing performed with software such as Image SXM. Section analysis in Nanoscope software, SXM Image, and others used for measuring particles. The Proteasome under AFM: Observations
Proteasome molecules imaged by the aforementioned method revealed a high level of dynamics when imaged with the practical resolution of 1–2 nm (Fig. 2). After analysis of AFM images performed under different conditions, it was possible to correlate the dynamic behavior of the proteasomes with their catalytic actions, to monitor a ligand binding, and to develop a model of allosteric signaling between the active centers and the gate (Gaczynska et al., 2003; Osmulski and Gaczynska, 2000, 2002). The rings of particles in the top‐view orientation were either cone shaped or had a crater‐like cavity in the middle, positioned where the entrance to the central channel was expected to be guarded by the gate. Therefore, we named the conformants ‘‘closed’’ and ‘‘open,’’ respectively, hypothesizing that they represent the closed and open gate to the proteasomal central channel (Fig. 2A; Osmulski and Gaczynska, 2000). This hypothesis was supported by the observation that images of top‐view proteasomes from Thermoplasma acidophilum, which do not have a gate in the ring, showed exclusively open particles (P. A. Osmulski et al., submitted; proteasomes kindly provided by A. L. Goldberg). In contrast, when eukaryotic proteasomes were analyzed, both the closed and the open conformants were present in all scans, and all of the particles were able to switch conformations between subsequent scans. Addition of peptide or protein substrates changed the partition between closed and open forms (Osmulski and Gaczynska, 2000). The conformational changes were observed in side‐ view proteasomes as well, with a partition between barrel‐shaped and cylinder‐shaped molecules closely following the partition between closed and open conformants (Osmulski and Gaczynska, 2002). These results prompted us to hypothesize that a ‘‘closed barrel’’ and an ‘‘open cylinder’’ represent allosteric forms analogous to R (relaxed) and T (tense) forms in a classical two‐state model of allosteric transitions. We have found that gate opening observed under AFM is coupled with a tetrahedral transition state in any of the active centers. Interestingly, kinetic parameters calculated on the basis of the open/closed partition as a function of substrate concentration were in perfect agreement with the parameters obtained in the traditional ‘‘macroscopic’’ way, with the increase in a product concentration measured instead of the abundance of conformers (P. A. Osmulski et al.,
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submitted). Core particles isolated from human and mouse cultured cells, rabbit muscle, Saccharomyces cerevisiae, or Schizosaccharomyces pombe were undistinguishable in AFM images and revealed the same dynamic behavior (Gaczynska et al., 2003; P. A. Osmulski and M. Gaczynska, unpublished observations). Analysis of AFM images is very useful for the characterization of ligand binding to the proteasomes. This was shown on the example of proline‐ and arginine‐rich peptides (PR peptides; see Gaczynska and Osmulski, 2005). These peptides bind to the 20S core particles, presumably to the ring. AFM images revealed that the 20S proteasome treated with the peptides was unable to switch between open and closed conformers and remained in a quasi‐open form permanently (Gaczynska et al., 2003). The 20S core particles are not the only proteasomal assemblies imaged under AFM. Studies of the interactions of human 26S complexes with PR peptides showed dramatic conformational changes in the inhibitor‐treated proteasomes (Gaczynska et al., 2003). The yeast 26S assemblies (Fig. 2B) and human proteasomes complexed with the 11S activator or the PI31 inhibitor (kindly provided by G. DeMartino) were visualized as well (P. A. Osmulski and M. Gaczynska, unpublished observations). The power of AFM, however, extends well beyond simple topography imaging, and more applications are waiting to be implemented in proteasomal studies. The force measurements, for example, will be very useful to study binding of the ubiquitinated substrates to 19S cap, their defolding, and translocation to the core particle. In addition to the aforementioned method of imaging electrostatically attached particles, several studies have explored the immobilized and organized 20S complexes under AFM. For example, proteasomes from archaeon T. acidophilum with histidine tags introduced at the C terminus of the subunit were specifically attached to a mica‐supported, ultraflat chelator lipid membrane (Dorn et al., 1999). As a result, images were obtained of a dense layer of the 20S particles oriented in a side‐view position. Interactions of the core particles with different supported lipid bilayers were explored for bovine proteasomes (Furuike et al., 2003). In another study, contact mode AFM was used to successfully image the shear‐resistant two‐dimensional crystals of T. acidophilum proteasomes (Thess et al., 2002). The 20S proteasomes His6‐tagged at the N termini of subunits crystallized readily on a nickel‐chelating lipid bilayer prepared on mica and optimized their packing in a crystal by interlocking both laterally and vertically (Thess et al., 2002). The immobilized and organized proteasomes constitute very useful research objects: however, the dynamic information may be, in most parts, lost.
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Acknowledgments The work was funded by R01 GM069819 (M.G.) and San Antonio Cancer Institute grants (P. A. O., M. G.).
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[35] Characterization of Noncompetitive Regulators of Proteasome Activity By MARIA GACZYNSKA and PAWEL A. OSMULSKI Abstract
The success of bortezomib, a competitive proteasome inhibitor and a drug approved to treat multiple myeloma, spurred interest in compounds targeting catalytic sites of the enzyme. The aim of this chapter, however, is to focus attention on the small molecule, natural or synthetic compounds binding far away from the catalytic centers, yet modifying the performance of the proteasome. Defining allostery broadly as any kind of ligand‐ induced, long‐distance transfer of conformational signals within a molecule, most such compounds are allosteric effectors capable of regulating the proteasome in vitro and in vivo in a manner more diverse and precise than competitive inhibitors. Proline‐ and arginine‐rich peptides (PR peptides) are examples of such compounds and are currently being considered as METHODS IN ENZYMOLOGY, VOL. 398 Copyright 2005, Elsevier Inc. All rights reserved.
0076-6879/05 $35.00 DOI: 10.1016/S0076-6879(05)98035-X