Probing the Structure of the Caulobacter crescentus Ribosome with

Probing the Structure of the Caulobacter crescentus Ribosome with Chemical Labeling and Mass Spectrometry Richard L. Beardsley, William E. Running, and James P. Reilly* Department of Chemistry, Indiana University, Bloomington, Indiana 47405 Received April 13, 2006

The ribosomal proteins of Caulobacter crescentus were amidinated before and after disassembly of the organelle and the results analyzed by mass spectrometry. Comparison with structural information from previous X-ray crystal studies of other bacterial ribosomes provides insight about the C. crescentus ribosome. In total, 47 of the 54 proteins present in the ribosome of C. crescentus were detected after labeling. The extent of derivatization for each protein is strongly dependent on the solvent accessibility of its target residues. Proteins of the ribosome stalk, which are known to be largely solvent-accessible, were labeled quite extensively. In striking contrast, other proteins that are known to be highly shielded in their subunits were labeled at very few of their potential sites. Furthermore, evidence that protein L12 binds to the ribosome via its N-terminal domain is consistent with previous findings.1 Keywords: chemical labeling • proteins • ribosome structure • protein mass spectrometry

Introduction Elucidating the structure of large proteins and complexes is a necessary step toward understanding their function. Traditionally, the methods of choice for solving high resolution structures have been either X-ray crystallography2,3 or nuclear magnetic resonance (NMR).4-6 Although advances are continuously being made, growing viable crystals for X-ray crystallography studies tends to be challenging and time-consuming, particularly for large noncovalent assemblies of biomolecules. Although solution phase NMR obviates this step, this technique is best-suited for smaller proteins.7 Additionally, both of these techniques require the isolation of relatively large quantities of sample. As a consequence of these limitations, some researchers have turned to labeling strategies in combination with mass spectrometry to probe protein structure.8-14 These approaches do not provide the complete picture or the high resolution of X-ray crystallography or NMR, but it is possible to map solvent-accessible sites in native structures, as well as identify residues involved in ligand binding and the interface regions of large noncovalent complexes. Hydrogen/deuterium exchange (HDX) is the most widely used form of labeling for probing protein conformations.8,15-20 HDX occurs when a donor molecule (e.g., D2O) forms a hydrogen bonded pair with an accepting heteroatom leading to the exchange of 2H with backbone amide and nonaliphatic side-chain hydrogens. The premise of this method is that the exchange rates of labile hydrogens correlate with their solvent accessibilities. Since HDX increases the mass of the acceptor molecule by 1 Da per exchange, mass spectrometry is wellsuited for monitoring these reactions. In a typical study, a protein is digested by an enzyme (often pepsin) following HDX, * To whom correspondence should be addressed. E-mail: reilly@ indiana.edu. 10.1021/pr060170w CCC: $33.50

 2006 American Chemical Society

and the resulting fragments are analyzed in an LC-MS/MS experiment.15 Attempts are often made to map the specific sites of exchange, although protein coverage is typically incomplete due to the site-selective nature of low energy peptide fragmentation.21 A wide array of structural studies have been performed by HDX, including the monitoring of ligand binding, protein folding dynamics, and mapping of protein-protein interfaces in complexes.15 A number of covalent labeling alternatives to HDX have been developed that target specific residues.11,14,22-31 An advantage of these approaches is that the scrambling of labels that readily occurs with HDX is eliminated.32 Although HDX can reveal more extensive coverage of protein topology, the mass spectrometric data provided by other derivatization tactics can be much simpler to interpret due to their reaction specificity and the lack of back exchange. The choice of proper derivatization reagents is critical. Reaction conditions must not be too harsh (e.g., extreme pH, high temperature, or long reaction times) since preserving the native structure of proteins and protein complexes during labeling is essential if the derivatizations are to reflect solvent accessibility. Also, the added moieties should not drastically perturb the physiochemical properties of the target residues as this could induce structural changes that affect native conformations. Additionally, the target residues for modification should be relatively abundant so that substantial regions of a protein surface are represented. Labeling of only methionine residues, for instance, would not be of general utility, since this amino acid is relatively rare. Covalent labeling strategies demonstrated as elucidative probes of site-specific solvent accessibility in proteins include aminoacylation of lysine residues,11,23,25,28 the derivatization of arginine by 1,2-cyclohexanedione,11,25 carbethoxylation of histidine residues,24 and iodination of tyrosine.25,27 In these studies, site reactivity was assessed by mass spectrometry through the Journal of Proteome Research 2006, 5, 2935-2946

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research articles observation of proteolytically generated peptides. Recently, a promising new method involving photochemical oxidation was introduced by Hettich and co-workers and further developed by Hambly and Gross.14,31 In this approach, hydroxyl radicals generated by UV photolysis of hydrogen peroxide react with multiple protein side chains (i.e., Cys, Trp, Tyr, Met, Phe, His, Ile, Leu, and Pro). The utility of this technique was initially demonstrated with model proteins lysozyme and β-lactoglobulin A. Labeling results and solution phase NMR structures were found to be consistent. More recently, Hambly and Gross also utilized photochemical oxidation to determine the solvent accessibilities of residues in the heme binding pocket of apomyoglobin.31 With a pulsed excimer laser, they induced photolysis on submicrosecond reaction times. Aminoacetylation has been the most widely employed covalent attachment probe of surface topology. Ligand binding, protein-protein complexes, and small model proteins such as ubiquitin have all been investigated.23,28 Przybylski and co-workers applied this form of labeling to study noncovalent complexes of elongation factor proteins EF-Ts and EF-Tu from Thermus thermophilus.23 Using mass spectrometry, they were able to locate several lysine residues that were shielded as a result of the tetrameric complex (2 EF-Ts and 2 EF-Tu) formed by these proteins. Because of its central role in protein synthesis, the ribosome has been one of the most intensely studied cellular components.33 Despite years of investigation, high resolution structures of the ribosomal subunits have been solved for only a few organisms.1,34-36 The ribosome is comprised of small (30S) and large (50S) nucleoprotein subunits, each possessing ribosomal RNA and several dozen surface-bound proteins. Growth of crystals suitable for X-ray diffraction studies is challenging due to the heterogeneity, large size (>2 MDa), and internal flexibility of the intact ribosome. The main purpose of this work was to investigate the extent of labeling of specific proteins in the intact ribosome of Caulobacter crescentus and determine if it correlated with what is known about ribosomal structures. We have recently demonstrated that the amidination of lysine residues can be employed to probe their solvent accessibilities in individual proteins.37 Since the ribosome is a nucleoprotein complex that is stabilized by numerous electrostatic interactions, including charge-charge interactions between phosphate groups of the rRNA nucleotide backbone and protonated basic residues, this system is well-suited for a labeling strategy that targets lysines. Amidination can be performed under mild conditions, and it adds a moiety that retains the basicity of the lysine side chain. Therefore, it is not expected to greatly perturb the tertiary or quaternary structure of the ribosome as we previously demonstrated for several small proteins.37 The intact ribosome of C. crescentus is a ∼2 MDa complex containing 2 subunits (large and small) with a total of 54 individual proteins and 3 rRNA molecules. Ribosomal proteins are amidinated before and after disassembling the intact organelle. The proteins are separated using a 2D liquid chromatography system constructed in-house and then electrosprayed and mass-analyzed using a quadrupole time-of-flight (QTOF) mass spectrometer. The extent of protein labeling before and after ribosome disassembly is compared. In addition, labeled proteins are digested by Glu-C and the fragments analyzed in a LC-MS/MS experiment to map the labeled sites. Although the C. crescentus ribosome has not been studied by X-ray crystallography, known structures of other bacterial ribosomes provide useful sources for comparison. Our 2936

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Beardsley et al. Scheme 1. Derivatization of Primary Amine Groups in Proteins with S-Methyl Thioacetimidate

results suggest that the ribosome of C. crescentus is very similar in structure to its counterparts in other bacteria.

Experimental Procedures Materials. Tris-(hydroxymethyl)aminomethane (Trizma base) and magnesium acetate were provided by Sigma (St. Louis, MO). Anhydrous diethyl ether, thioacetamide, and ammonium bicarbonate were purchased from Fisher (Fair Lawn, NJ). Iodomethane, iodoacetamide, and formic acid were obtained from Aldrich (Milwaukee, WI). Thiopropionamide was supplied by TCI America (Portland, OR). Acetonitrile, glacial acetic acid, urea, and trifluoroacetic acid (TFA) were purchased from EM Science (Gibbstown, NJ). Endoprotease Glu-C and Glu-C reaction buffer were purchased from New England BioLabs (Ipswich, MA). All water used in this research was purified by an E-pure water filtration system from Barnstead Thermolyne Co. (Dubuque, IA). Preparation of S-Methyl Thioacetimidate. S-Methyl thioacetimidate was prepared as described previously by Beardsley and Reilly.38 A total of 11 g of thioacetamide was stirred into 1 L of diethyl ether at ambient temperature. A volume of 8.8 mL of iodomethane was added to this solution, and the reaction mixture was allowed to stand at ambient temperature for 14 h. The precipitate that formed was isolated by suction filtration and stored over desiccant at room temperature. The dried powder was used in all experiments without further purification. Amidination of Ribosomal Proteins. Intact ribosomes, including all proteins and rRNA, were isolated from C. crescentus via the method of Spedding that was adapted by Arnold and Reilly.39,40 The concentration of proteins in the stock sample (i.e., before ribosome disassembly) was 8.5 mg/mL as determined by a Bradford assay. Ribosomes were stored at pH 8.0 in 10 mM Tris-HCl, 10 mM magnesium acetate, 60 mM NH4Cl, and 3 mM β-mercaptoethanol. A sample (100 µL) of this solution was mixed with an equal volume of 43.4 g/L S-methyl thioacetimidate buffered by 250 mM Trizma base to begin amidination reactions (Scheme 1). This was incubated for 1 h at ambient temperature before adding glacial acetic acid and 1 M MgCl2 such that the final solution contained 3:6:1 (v/v/v) reaction mixture/glacial acetic acid/1 M MgCl2. In addition to terminating the reaction, these reagents also precipitated rRNA, which was necessary to avoid fouling chromatography columns during subsequent 2D-LC separations. The rRNA precipitate was separated by centrifugation at 14 100 rpm for 5 min. The supernatant, containing ribosomal proteins, was used in 2DLC-MS experiments without further modification. Ribosomal proteins were also amidinated after precipitation of rRNA and complete disassembly of the ribosome structure. To the above-mentioned 100 µL aliquot of the ribosome stock solution, 100 µL of glacial acetic acid and 33 µL of 1 M MgCl2 were added. Precipitated rRNA was separated from proteins by centrifugation at 14 100 rpm for 5 min. The excess acetic acid in the supernatant was partially evaporated using a speedvac. It was found that complete removal of acetic acid led to the precipitation of ribosomal proteins. Therefore, the amidi-

Probing the Structure of the Caulobacter crescentus Ribosome

nation reaction conditions included 7.5% (v/v) acetic acid. Because of the presence of this acid, the reaction was buffered by 4 M Trizma base instead of 250 mM as described for amidination of the intact ribosome. A pH of 8 was measured for both sets of conditions. With the exception of the higher buffer concentration and presence of acetic acid, the reaction conditions (i.e., S-methyl thioacetimidate concentration and incubation time) were the same as described above. This reaction was terminated by adding an equal volume of 10% (v/v) TFA.

research articles Digestion and LC-MS/MS of Labeled Ribosomal Proteins. Prior to enzymatic digestion, it was necessary to remove excess reagents (i.e., Trizma base and S-methyl thioacetimidate) from the amidinated proteins. This was accomplished using 3 kDa molecular weight cutoff (MWCO) filters (Centricon, Millipore, Billerica, MA). Reaction mixtures were exchanged with water in three centrifugation cycles (13 000 rpm for 20 min each). Purified fractions were evaporated to dryness using a speedvac (Jouan, Winchester, VA) and reconstituted in digestion buffer (50 mM Tris-HCl and 0.5 mM diglutamate, pH 8). A volume of 100 µL of this solution was combined with 2 µg of lyophilized Glu-C (New England Biolabs, Beverly, MA) and incubated for 18 h at 37 °C. The digestion was terminated by adding TFA to a concentration of 1% (v/v). Glu-C was chosen for this work because the residues it targets for cleavage (i.e., Glu and Asp) were not affected by the labeling procedures. Since many lysine residues were amidinated, trypsin would have provided a limited number of peptides.

Chromatography and Mass Spectrometry of Labeled Ribosomal Proteins. Protein separations were performed following the labeling experiments using an in-house-designed 2D-LC system.41 In brief, this system consists of a strong cation exchange (SCX) column (TosoHaas, SP NPR) followed by 20 reversed-phase columns (Javelin, Thermo, 20 × 1.0 mm, C4) arranged serially and controlled by a Labview interface. In all experiments, approximately 50 µg of protein, contained in the acidified reaction mixtures described above, was injected onto the SCX column. SCX mobile phase A was composed of 20 mM acetic acid, adjusted to pH 5.1 with 1% (v/v) methylamine in water, and 6 M urea. SCX mobile phase B was identical to A, but also included 500 mM NaCl. Prior to starting the gradient, protein solutions were loaded onto the SCX column under isocratic conditions for 20 min using 100% A at a flow rate of 150 µL/min. After sample loading, the flow rate was increased to 300 µL/min, and the concentration of SCX mobile phase B was increased linearly from 0 to 10% for 5 min. Next, B was linearly increased to 35% over a period of 21 min (i.e., 25-46 min). From 46 to 95 min, B was increased to 70% (i.e., 350 mM NaCl). Finally, the concentration of B was ramped to 100% and held constant for the final 15 min of the separation. During this gradient, the reversed-phase columns (i.e., columns 1-20) were “loaded” with the eluting proteins from the SCX column via a valve-switching apparatus (controlled by LabView) that positioned the columns in-line with the effluent at fixed intervals. Columns 1 and 2 were loaded for 10 min each during the first 20 min. Subsequently, traps 3-20 were loaded for 5 min apiece during the remainder of the gradient.

Glu-C digests were analyzed by LC-MS/MS using a C18 reversed-phase column (100 × 0.254 mm, Biobasic18, Thermo Hypersil-Keystone, Bellefonte, PA) and an ion trap mass spectrometer (LCQ-Deca XP Plus, Thermo, San Diego, CA). Approximately 20 µg of ribosomal protein digest were injected onto the column in each experiment. Mobile phase A contained 0.1% formic acid in water, while mobile phase B was composed of 0.1% formic acid in acetonitrile. A flow rate of 5 µL/min was established by a “T” splitter positioned before the injection loop. Samples were injected using an autosampler (Surveyor, Thermo, San Jose, CA) and loaded onto the column using 95% A and 5% B. The gradient was implemented by linearly increasing mobile phase B from 5% to 40% over 240 min. The effluent was directed to the ESI source of the mass spectrometer. CID MS/MS spectra were acquired from the three most intense precursor ions in a single MS scan. Following the acquisition of one MS scan and the three subsequent MS/MS spectra, this sequence was repeated. Dynamic exclusion limited the number of fragmentation spectra acquired from a single m/z precursor ion to two. Using helium as the CID target gas, all precursor ions were activated for 30 ms by resonance excitation employing a normalized collision energy of 35%.

Twenty LC-MS experiments were performed following the loading steps. The effluent from each trap was directed through a C4 column (Thermo Hypersil Keystone Pioneer, 1 × 100 mm) before being electrosprayed and mass-analyzed in a quadrupole time-of-flight mass spectrometer (QTOF, Waters, Manchester, U.K.). Mobile phase A was buffered by 0.1% (v/v) formic acid and contained 5% acetonitrile, while B contained the same buffer and 95% acetonitrile. In all steps described below, the flow rate was maintained at 50 µL/min. Prior to beginning the gradient, salts from the SCX step were purged with 100% mobile phase A for 7 min. Proteins were eluted over the next 30 min by linearly increasing mobile phase B from 20% to 70%. The effluent from each column was split prior to the mass spectrometer to establish a flow rate of 10 µL/min directed to the electrospray ionization (ESI) source of the QTOF. The remaining effluent was directed to waste. A voltage of +3.0 kV was applied to the ESI needle. Mass spectra were acquired over a range of 600-1800 m/z. The molecular weights of detected proteins were calculated from charge state distributions using a maximum entropy algorithm. The experimental deconvoluted protein mass errors were typically (2 Da. Mass spectra used for these calculations were manually selected by averaging the scans within a chromatographic peak.

Comparing Labeling to Crystal Structures of the Ribosome. The structures of 30S and 50S ribosomal subunits were obtained from the Protein Data Bank and analyzed in Protein Explorer.42 The 30S subunit was derived from Escherichia coli (Protein Data Bank, 1PNX),34 while the 50S counterpart was taken from both Deinococcus radiodurans (Protein Data Bank, 1NKW)36 and T. thermophilus (Protein Data Bank, 1GIY).1 The structure from T. thermophilus was used to consider ribosomal stalk proteins that were not included in the 50S D. radiodurans structure. The 50S subunit of bacterial ribosomes contains two stalk-like structures that extend away from the nucleoprotein core of the complex. The structure from T. thermophilus was not used for other comparisons, since the D. radiodurans structure included more of the ribosomal proteins. The structures of proteins in the 50S and 30S subunits are displayed as van der Waals radii. The coordinates of each amino acid residue in the 50S subunits were only available from the PDB as single points centered at the R-carbon. Therefore, the proteins of the 50S subunit are pictured as spheres representing this position for each residue. The images of the 30S subunit show van der Waals radii of all atoms except hydrogens, unless otherwise noted. The protein sequences of these ribosomes were compared to the predicted sequences of C. crescentus ribosomal Journal of Proteome Research • Vol. 5, No. 11, 2006 2937

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Figure 1. QTOF mass spectra of ribosomal protein L22 acquired during 2D-LC experiments. The spectra shown here were taken from unmodified samples (A), and those that were acetamidinated with the ribosome intact (B) or disassembled (C).

proteins43 using the Web-based LALIGN algorithm.44,45 Alignments were calculated using the default settings of this algorithm. Solvent-Accessible Surface Area Calculations. The experimentally determined extent to which each ribosomal protein was labeled was compared to its solvent-accessible surface area calculated from the crystal structures of the 30S and 50S subunits. The calculations were performed using a Web-based algorithm, parameter optimized surfaces for complexes (POPSCOMP).46,47 Solvent-accessible surface area is defined as the area that is accessible to a probe with the diameter of a water molecule rolling over the surface.48 All calculations done here were performed using the coarse-grained function of the 2938


software that treats each residue as a single sphere centered at the R-carbon.

Results and Discussion Comparing the Labeling of Intact and Disassembled Ribosomes. With the methods discussed above, ribosomal proteins were amidinated before and after disassembling the C. crescentus ribosome. Following derivatizations, protein masses were measured by ESI-TOF mass spectrometry. Figure 1 displays very typical examples of these data. ESI mass spectra of ribosomal protein L22 appearing on the left were produced by summing the scans acquired, while this protein eluted from the reversed-phase column. The charge state distribution was


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Figure 2. Deconvoluted ESI QTOF mass spectra of acetamidinated ribosomal proteins. All units are in molecular weight. Spectra of proteins labeled as part of the intact ribosome are shown above their counterparts that were labeled following disassembly of the ribosome. Spectra of ribosomal proteins (A) S16, (B) S14, (C) S6, (D) S19, (E) S20, (F) L33, (G) L28, (H) L17, (I) L19, and (J) L27 are displayed here.

deconvoluted using the MaxEnt algorithm, and resulting spectra are displayed on the right.49 Underivatized samples were previously analyzed to identify these ribosomal proteins and their post-translational modifications.50 Many ribosomal proteins lose their N-terminal methionines, and some are methylated or acetylated in vivo. These latter modifications are of particular relevance to the present work, since they typically target potential amidination sites such as N-termini and lysine residues. The 14 110 Da molecular weight of L22 is consistent with the cleavage of methionine from this protein (calculated MW is 14 108). The spectrum of L22, labeled prior to disassembly of the ribosome (Figure 1B), demonstrates that this protein was partially derivatized. Although L22 contains 10 potential modification sites (i.e., nine lysines and one free N-terminus), the base peak is consistent with incorporation of only five acetamidine groups. There are also significant peaks in this spectrum that correspond to the addition of four (MW 14 272) and six (MW 14 355) acetamidine groups, as well as minor peaks that match three (MW 14 231) and seven (MW 14 396) added labels. This distribution of reaction products suggests that the reaction may be hindered but not completely blocked at some sites. The mass spectrum of L22, derivatized after disassociation of the ribosome, appears in Figure 1C. The deconvoluted spectrum contains a single dominant peak consistent with a completely labeled mass of 14 520 Da (expected MW is 14 519). This result indicates that some of the potential reaction sites of L22 are blocked by the structure of the intact ribosome complex.

Although we recently detected 53 of the 54 underivatized ribosomal proteins of C. crescentus, in the present work, 47 out of the 54 were observed in both labeled samples and their solvent accessibility was assessed. It is not entirely clear why seven of the ribosomal proteins were not observed in this 2DLC experiment. However, it is likely that these were missed due to matrix effects that are typical of analyses of complicated mixtures. We find that very high coverage of all components of a protein mixture often requires multiple experimental runs under different operating conditions. Representative deconvoluted ESI-QTOF mass spectra of 10 of these proteins are shown in Figure 2. Spectra of the remaining 37 observed proteins are provided in Supporting Information. For each part of Figure 2, the top spectrum was generated by labeling the intact ribosome, while the bottom counterpart was obtained by derivatizing the disassembled proteins. These spectra demonstrate the labeling characteristics observed in this study. A remarkable feature of these data is that spectra of ribosomal proteins that were labeled after removal of rRNA (i.e., after ribosome disassembly) exhibit only one dominant peak. This trend was observed for every protein that was detected in this study (see also spectra in Supporting Information). In all such cases, the measured protein mass was consistent with the labeling of all available sites (i.e., every free lysine and the N-terminus). In a few instances, a low-intensity peak corresponding to the labeling of all but one available site was also observed. One such case was L28 (Figure 2G), where the peak at molecular weight 10 912 corresponds to the addition of only Journal of Proteome Research • Vol. 5, No. 11, 2006 2939

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Figure 3. (A) Deconvoluted ESI-QTOF mass spectra of amidinated proteins from the L1 and L7/L12 stalks of the 50S subunit. Each spectrum was acquired after labeling the intact subunit. (B) The 50S subunit of T. thermophilus depicting the positions of stalk proteins. L1 is shown red, and a dimer of L7/L12 is depicted in white and orange. Since L10 was not included in this crystal structure, it is not displayed here. All other proteins are shown in blue with the 23 and 5S rRNA molecules in green and violet, respectively. (C) Ion trap tandem mass spectrum of the single amidinated N-terminal peptide (-SKLE) from L12.

10 out of 11 acetamidine groups. In other examples like this one, the n - 1 peak (n ) total available sites) was of similarly low intensity. Nevertheless, the labeling of proteins in the intact complex nearly always resulted in incomplete modification of the available sites. In many cases, a distribution of 3 or 4 peaks corresponding to varying degrees of partial labeling was observed. For example, the labeling of ribosomal protein S16 (Figure 2A) as part of the intact complex led to products with molecular weights of 18 056, 18 097, and 18 138 consistent with addition of 11-13 acetamidine groups. Interestingly, some proteins that were labeled when the ribosome was intact revealed a narrow distribution of reaction products. For example ribosomal proteins S6 (Figure 2C), S19 (Figure 2D), S20 (Figure 2E), L19 (Figure 2I), and L27 (Figure 2J) show only one prominent peak when labeled in the intact ribosome. The differences in labeling distributions in these cases should provide information about the environments of the available labeling sites. For example, the observation of a single dominant peak suggests that all of the unlabeled sites are directly involved in a binding interface, while those that are labeled are freely exposed and solvent-accessible. Alternatively, the observation of a broad distribution of reaction products suggests that some reaction sites are partially solvent-exposed or may be involved in more dynamic interactions that are in equilibrium between bound and unbound states. Therefore, the labeling reaction can occur at such sites, albeit more slowly. Future work involving time-dependent labeling experiments will be aimed at elucidating the relative reactivities of such sites. Stalk Proteins of the 50S Subunit. A few of the proteins detected in this study compose stalk-like structures that extend from the core of the 50S subunit.1,34,36,51 These proteins include L7/L12, L10, and L1. In general, the bacterial ribosomal stalk 2940


complex is known to be a pentamer containing two copies each of L7 and L12 (L7 is the N-acetylated form of L12) along with a single copy of L10 to which the L7/L12 proteins bind.52 However, Robinson and co-workers recently discovered that the stalk complex of T. thermophilus is a heptamer composed of one L10 molecule along with six L7/L12 proteins.53 This highly flexible structure responds to elongation factors during protein synthesis as it undergoes significant movements upon their binding.54,55 The L7/L12 stalk has been identified in lowresolution electron microscopy studies.51 Additional insights were also recently provided in the 70S crystal structure (i.e., the complex of the small and large subunit) of T. thermophilus as a compact dimer of L7/L12 was included in this map.1 The structures of the remaining L7/L12 proteins were likely not included due to their significant conformational flexibility. The other stalk of the 50S subunit is formed by L1, which is bound to an extended region of the 23S rRNA. Unlike many of the proteins found in the ribosome, which often bind within the folds of the 23S rRNA, stalk proteins share relatively few interactions with the rRNA and are more solvent-exposed. Their loosely bound nature was recently investigated by Robinson and co-workers.56 In this work, it was demonstrated that stalk proteins are more easily released than others from the 50S subunit of E. coli in response to changes in the pH. This behavior was attributed to their limited interaction with rRNA. The deconvoluted LC-ESI-QTOF mass spectra of amidinated L7/L12, L10, and L1 are displayed in Figure 3A. In each case, the spectrum was acquired from the sample that was labeled with the ribosome intact (i.e., amidination before rRNA precipitation). For each spectrum, the number of labels added is indicated. The extents of labeling for all of these proteins can also be found in Table 1. As shown here, the derivatization of

Probing the Structure of the Caulobacter crescentus Ribosome Table 1. Comparison of Ribosomal Protein Labeling Efficiency: Intact versus Disassembled Ribosome observed proteins proteins

L1b

potential sitesa

number of labels added intact

dissassembled

L2 L4 L5 L6 L10 L12b,d L13 L14 L15 L16b L17 L18b L19 L20 L21 L22 L23 L24 L25 L27 L28 L29 L30 L31 L32b L33b L34 L35 L36

19 24 21 20 20 17 15 17 11 19 12 13 9 8 14 19 10 13 16 16 11 11 8 4 7 6 11 7 13 9

16-19 15-18 13-16 16-18 16-19 15 14, 15 9-11 7-9 10-13 3-5 2-5 7-9 7 3,4 13-15 3-7 4-6 11-13 16 8 6-9 6-7 3 6-7 4 7-9 0, 2-4 1-4 3-6

19 23, 24 21 20 19, 20 17 14, 15 16, 17 11 19 12 13 9 8 14 19 10 13 16 16 11 10, 11 8 4 7 6 11 7 13 9

S3 S4 S5b S6 S7 S8 S9c S10 S12 S13 S14 S15 S16 S17 S18c S19 S20

18 18 13 7 13 12 12 9 14 11 13 9 17 14 10 12 12

14 8-11 9,10 6,7 9-11 9-11 4-6 5-7 10-12 9-11 7-10 5-9 11-13 8-10 7-9 8,9 7-9

18 18 13 7 13 12 12 9 14 11 13 9 17 13, 14 10 12 12

a Potential reaction sites included all unmodified lysine residues and N-termini. b Indicates an observed methylation (i.e., +14 Da) in the spectrum of an unmodified protein. c Denotes acetylation of the N-terminus (i.e., + 42 Da). d Methylation observed but apparently does not modify a lysine or the N-terminus.

each of the stalk proteins in the intact ribosome was nearly complete. For example, L7/L12 contains 15 potential labeling sites, and the dominant peak in this spectrum matches the addition of 14 acetamidine groups. Furthermore, a peak at about 20% relative intensity corresponding to complete labeling was also observed. The derivatization of this protein following disassembly of the ribosome resulted in the dominant reaction product being completely modified (15 labels) with a less intense product corresponding to the incorporation of 14 tags also present. The measured mass of L12 was consistent with a single methylation, but this protein was labeled at all of its lysines and its N-terminus (15 total). Therefore, it appears that the protein must be methylated at a different residue. L7 could not be distinguished from L12 in these experiments, since an

research articles acetyl group is only 1 Da heavier than acetamidine. However, it is believed that only the unacetylated L12 is detected here, since L7 was not observed in analyses of unamidinated ribosomal proteins.50 Similarly, L10 and L1 were also derivatized quite extensively while still bound in the ribosome. The spectrum of L10 contains one peak that matches derivatization at all but 2 of its 17 available sites. The spectrum of L1 contains a distribution of four peaks; those corresponding to the addition of 17 and 18 acetamidine groups (out of 19 potential sites) are the most dominant. As with all proteins, L10 and L1 were completely labeled when the reaction was performed after rRNA precipitation. The extensive labeling of these ribosomal stalk proteins correlates very well with previous studies that suggested low interaction surface areas for these molecules.57,58 Furthermore, the structure of the 50S subunit of T. thermophilus, shown in Figure 3B, reveals that L1 and the L7/L12 complex are extended from the core of the ribosome and are therefore highly solvent-accessible. Since L10 was not included in this structure, it is not shown in this figure. Following amidination of the intact ribosome, the proteins were digested using Glu-C to provide peptides that could be fragmented to locate modified (or unmodified) sites. As shown in Figure 3A, the predominant labeling product of L12 matched derivatization at all but one of its 15 available sites. All lysinecontaining peptides that were observed from this protein exhibited mass and fragmentation characteristics consistent with the Lys being amidinated except for -SKLE whose ion trap MS/MS spectrum is displayed in Figure 3C. Neutral loss products contribute significantly to this spectrum. We have previously demonstrated that neutral loss of NH3 from singly protonated amidinated peptides is a favored process.38 This peptide contains two potential labeling sites (the N-terminus and Lys), and is labeled only once. The b3, a3, and b2 fragment ions that are in this spectrum are all consistent with the addition of only one amidine group. However, without the presence of a b1, y3, or an internal fragment ion, it is not possible to establish whether the lysine or the N-terminal amine is labeled. Nevertheless, these data do provide strong evidence that the N-terminal region of L12 is not very solvent-accessible. This interpretation is consistent with the known structures of L7/L12, as it is the N-terminal domain that anchors it to the ribosome and is involved in dimerization.52 In contrast, the C-terminal domain and a “hinge” region are relatively exposed.52 Comparing Limited Labeling to Crystal Structures. Ribosomal proteins L35 and L20 are examined more closely in Figure 4. The deconvoluted mass spectrum of L35 (Figure 4A), labeled with the ribosome intact, features a base peak of MW 7304 that corresponds to 2 of 13 potential sites being modified. In striking contrast to the stalk proteins just discussed, the limited derivatization of L35 indicates that many of its available sites are not solvent-accessible. The images displayed in Figure 4 were acquired from the Protein Data Bank and represent the crystal structure of the 50S ribosomal subunit of D. radiodurans.36 On the basis of the sequence homology shared by eubacterial ribosomal proteins, the structure of the ribosome of C. crescentus is expected to be similar. Figure 4A shows the position of L35 in the 50S complex. L35 is displayed both with and without the other molecules of the ribosome to demonstrate the extent of its burial in the complex. With the exception of L35 (colored white), all proteins are shown in blue, while 23S and 5S ribosomal RNA are shown in green and violet, respectively. The red spheres of L35 represent potential labeling Journal of Proteome Research • Vol. 5, No. 11, 2006 2941

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Figure 4. Deconvoluted mass spectra of ribosomal proteins (A) L35 and (B) L20 acquired from the sample that was labeled with the ribosome intact. The structures to the right are of the 50S ribosomal subunit of Deinococcus radiodurans (PDB, 1NKW). The 23S rRNA is displayed in green, while the ribosomal proteins are in blue, and the 5S rRNA is shown in violet. L35 and L20 are highlighted in white. Residues that align with the lysines of L35 and L20 of C. crescentus are displayed in red.

sites. It is clear from these two images that the 23S rRNA almost completely buries L35. It is unlike most ribosomal proteins that have significant surface exposure (see proteins displayed in blue) but contain “fingerlike” structures that extend into the folds of the rRNA. The deconvoluted mass spectrum of L20 (labeled in the intact subunit) and images of the crystal structure of the 50S subunit of D. radiodurans are shown in Figure 4B. The base peak in this spectrum at MW 13 144 matches the molecular weight of L20 plus only 3 out of 14 acetamidine moieties. The red spheres are residues of L20 from D. radiodurans that align with the lysines and N-terminus of L20 from C. crescentus. Although a significant portion of this sequence appears as a globular structure that is exposed on the surface of the ribosome, a long extension is shielded by the 23S rRNA and protein L21. All but three of the aligned residues are in this shielded tail. The labeling results suggest that L20 of C. crescentus likely also contains a lysine-rich “tail” that permeates the 50S subunit. In cases where the extent of labeling is limited, a protein may either be largely buried in a complex or many of its available reaction sites may be positioned in a small area that is not solvent-accessible. Unfortunately, it is difficult to distinguish between these possibilities without the aid of a crystal structure. Summary of Labeling Results. Table 1 summarizes the extent of labeling for all ribosomal proteins. The potential reaction sites include all unmodified lysine residues and free N-termini, reduced by the number of methylations or acetylations that were identified in the unmodified sample.50 The numbers of derivatizations observed for the intact and disassembled samples are listed on the right. As described above, the extent of labeling was consistently lower for the intact ribosome, and results vary significantly from one protein to 2942


another. While every observed protein from the disassembled ribosome was completely modified, only a few of those from the intact ribosome (i.e., S6, L1, L10, L7/L12, L18, L19, and L25) were labeled extensively. It was common for a protein to be missing four or five labels, and in some cases (e.g., L35 and L20), nearly all potential sites were unlabeled. Extent of Labeling versus Calculated Solvent Accessibilities. As noted above, a crystal structure for the C. crescentus ribosome has not yet been determined. Therefore, our attempts to compare the extent of ribosomal protein labeling with calculated solvent accessibility predictions necessarily involved structural data from D. radiodurans and E. coli. The structures of these ribosomes are known to be similar based on X-ray diffraction experiments and high sequence similarities. The solvent shielding of corresponding ribosomal proteins of these organisms was calculated and compared to the labeling results for C. crescentus. The total buried surface area of an individual molecule in the complex was determined in the following manner. First, the solvent accessible surface area of each molecule in the complex was calculated without consideration of its binding partners. Next, the solvent-accessible surface areas of all possible pairwise combinations in the complex were calculated. Therefore, if two molecules share a binding interface, the solvent-accessible surface area of that pairwise complex will be less than the sum of their individual surface areas. This difference equals twice the surface area of the binding interface between two molecules in the complex. Since the total buried surface area of a pairwise complex is equal to the sum of the binding interfaces of each component, half of this total was taken to equal the interfacial (buried) surface area of a single protein. Last, the interfacial surface areas determined from each pairwise complex were summed to account for all binding interfaces, thus, providing the total buried surface area


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Figure 5. Comparison of ribosomal protein labeling efficiency in the intact ribosome to calculated solvent accessibility in the crystal structures of (A) the 50S subunit of D. radiodurans, and (B) the 30S subunit of E. coli. The gray bars show the calculated percentage of a proteins surface that is solvent-inaccessible, while the black bars indicate the percentage of the available reaction sites of a protein that were not labeled (weighted average). Only those proteins that were both included in the crystal structures and observed mass spectrometrically are represented here.

of each protein. Figure 5 compares the calculated percentage of each protein’s surface area that is buried from solvent in the complex (gray bars) with the percentage of available sites that were experimentally unamidinated (black bars). Figure 5A displays results for the large subunit proteins, while Figure 5B focuses on the small subunit. The calculations were performed on the large subunit of D. radiodurans and the small subunit of E. coli (since the crystal structure of the 30S subunit of D. radiodurans is not available). Because ribosomal proteins L1, L10, L7/L12, L25, and L28 were not included in the crystal structure of the large subunit, these proteins are not represented in Figure 5. The percentages of unlabeled sites are derived from the weighted average of the distribution of products observed in the deconvoluted mass spectrum of each protein. The calculated solvent accessibilities are in qualitative agreement with the labeling results. On the basis of the

calculations, the least accessible proteins in the 50S subunit are L35, L34, and L20. Although the percentages of unlabeled sites for these proteins do not exactly match the calculated burials, they are consistent in that these proteins were among the least labeled of any in this plot. Like these proteins, L16, L17, and L23 were also labeled at relatively few of their sites. However, their calculated solvent accessibilities suggest that they are not as solvent-shielded as the limited labeling indicates. This outcome suggests that the solvent-exposed domains of these proteins are not lysine-rich. Additionally, proteins that were predicted to be largely solvent-accessible (e.g., L5, L6, L29, S6, and S7) were more completely labeled. However, for a few proteins, the correlation between labeling and burial is not very good. Since the available labeling sites consist of lysine residues and the N-terminus, it is not surprising that there are some cases where the labeling does not reflect the overall burial of a Journal of Proteome Research • Vol. 5, No. 11, 2006 2943

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Beardsley et al.

Figure 6. Comparison of extent of derivatization with numbers of exposed potential labeling sites in the crystal structures of (A) the 50S subunit of D. radiodurans and (B) the 30S subunit of E. coli. Gray bars represent the experimentally determined weighted averages of acetamidine groups added to each protein. Black bars depict the number of solvent-accessible potential labeling sites.

protein. Since it is not likely for these sites to be evenly distributed on the protein surface, one cannot expect the extents of labeling to precisely correlate with the extent of solvent shielding. Furthermore, the distribution of labeling sites is skewed by the need for structural domains (e.g., fingerlike extensions) that must be rich in basic residues to stabilize binding to RNA, such as the case of L20 that was described above in Figure 4B. Another potential source of error is that the sequences on which the calculations were performed (i.e., ribosomes of E. coli and D. radiodurans) are not identical to the proteins in the ribosome of C. crescentus. In future work, the ribosomes of these organisms will be studied so that more definitive comparisons to their structures can be made. Comparison of Exposed Potential Labeling Sites with Extent of Labeling. As just described, relating the extent of labeling to calculated solvent accessibility may be too simplistic. Figure 6 shows an alternative comparison of the experimentally determined labeling efficiencies with the number of exposed potential labeling sites visible from the crystal structures of D. 2944


radiodurans and E. coli. Since the sequences of ribosomal proteins of these organisms are not identical to those of C. crescentus (between 40% and 60% sequence identity is typical), it was necessary to align the respective sequences to predict structurally homologous regions and to estimate the likely positions of potential labeling sites in the ribosome of C. crescentus. Comparisons for both the large and small subunit are displayed in Figure 6, panels A and B, respectively. Unfortunately, some of the proteins that were included in the solvent accessibility calculations of Figure 5 are excluded from this comparison because only a small number of their potential labeling sites could be aligned to the respective sequence in the crystal structure. These proteins include L15, L21, L30, L33, S14, and S16. It is evident in plots of Figure 6 that the number of exposed residues correlates very well with the extent of labeling nearly all proteins. These results reflect the similarity in the organization of the ribosomes of C. crescentus, E. coli, and D. radiodurans, and they also indicate that the overall


structure of the ribosomal subunits remains intact during the derivatization.

Conclusion The ribosome of C. crescentus was studied using a strategy based on amidination to determine the solvent accessibility of lysine residues and protein N-termini. The ribosomal proteins of C. crescentus were acetamidinated both as individual molecules and as part of the intact subunits. QTOF mass spectra of the labeled proteins demonstrate that the extent of labeling is highly dependent on whether the subunits remain intact or are disassembled. Furthermore, labeling results can be interpreted in light of what is known about the structures of other bacterial ribosomes. Ribosome stalk proteins, which are known to extend from the core of the organelle and be relatively solvent-accessible, were labeled quite extensively. Furthermore, fragmentation of the N-terminal peptide of L12 demonstrated that this region contained the lone unlabeled site of this protein. This result is consistent with previous reports indicating that this protein is anchored to the large subunit by its N-terminal domain. By comparison to the calculated solvent accessibilities of other bacterial ribosomes, it was found that the experimentally measured extent of labeling generally reflected the positions of proteins in the complex. However, by taking the precise positions of potential labeling sites into consideration, this comparison resulted in a better correlation. On the basis of these interpretations, it is concluded that the selectivity of amidination is remarkably consistent with the crystal structures. Thus, the overall conformation of the ribosomes of C. crescentus, D. radiodurans, and E. coli must be very similar.

Acknowledgment. This work has been supported by the National Science Foundation grant CHE0518234, the Indiana 21st Century Fund, and the Indiana Metacyte Initiative. This work was also supported in part by a fellowship from Merck Research Laboratories. Supporting Information Available: Spectra of the other 37 observed proteins. This material is available free of charge via the Internet at http://pubs.acs.org. References (1) Yusupov, M.; Yusupova, G.; Baucom, A.; Lieberman, K.; Earnest, T.; Cate, J.; Noller, H. Science 2001, 292, 883-896. (2) Blake, C.; Koenig, D.; Mair, G.; North, A.; Phillips, D.; Sarma, V. Nature 1965, 206, 757-761. (3) Schmidt, A.; Lamzin, V. Curr. Opin. Struct. Biol. 2002, 12, 698703. (4) Wuthrich, K. Acta Crystallogr., D 1995, 51, 249-270. (5) Garrett, D.; Seok, Y.; Liao, D.; Peterkofsky, A.; Gronenborn, A.; Clore, G. Biochemistry 1997, 36, 2517-2530. (6) Clore, G.; Gronenborn, A. Nat. Struct. Biol. 1997, 4, 849-853. (7) Christendat, D.; Yee, A.; Dharamsi, A.; Kluger, Y.; Savchenko, A.; Cort, J.; Booth, V.; Mackereth, C.; Saridakis, V.; Ekiel, I.; Kozlov, G.; Maxwell, K.; Wu, N.; Mcintosh, L.; Gehring, K.; Kennedy, M.; Davidson, A.; Pai, E.; Gerstein, M.; Edwards, A.; Arrowsmith, C. Nat. Struct. Biol. 2000, 7, 903-909. (8) Englander, S.; Mayne, L.; Bai, Y.; Sosnick, T. Protein Sci. 1997, 6, 1101-1109. (9) Englander, J.; Del Mar, C.; Li, W.; Englander, S.; Kim, J.; Stranz, D.; Hamuro, Y.; Woods, V. Proc. Natl. Acad. Sci. U.S.A. 2003, 100, 7057-7062. (10) Figueroa, I.; Russell, D. J. Am. Soc. Mass Spectrom. 1999, 10, 719731.

research articles (11) Suckau, D.; Mak, M.; Przybylski, M. Proc. Natl. Acad. Sci. U.S.A. 1992, 89, 5630-5634. (12) Calvete, J.; Campanero-Rhodes, M.; Raida, M.; Sanz, L. FEBS Lett. 1999, 444, 260-264. (13) Hager-Braun, C.; Tomer, K. Biochemistry 2002, 41, 1759-1766. (14) Sharp, J.; Becker, J.; Hettich, R. Anal. Chem. 2004, 76, 672-683. (15) Ehring, H. Anal. Biochem. 1999, 267, 252-259. (16) Zhu, M.; Rempel, D.; Du, Z.; Gross, M. J. Am. Chem. Soc. 2003, 125, 5252-5253. (17) Zhu, M.; Rempel, D.; Gross, M. J. Am. Soc. Mass Spectrom. 2004, 15, 388-397. (18) Zhang, Z.; Smith, D. Protein Sci. 1993, 2, 522-531. (19) Katta, V.; Chait, B. Rapid Commun. Mass Spectrom. 1991, 5, 214217. (20) Mandell, J.; Falick, A.; Komives, E. Proc. Natl. Acad. Sci. U.S.A. 1998, 95, 14705-14710. (21) Wysocki, V. H.; Tsaprailis, G.; Smith, L. L.; Breci, L. A. J. Mass Spectrom. 2000, 35, 1399-1406. (22) Schnaible, V.; Przybylski, M. Bioconjugate Chem. 1999, 10, 861866. (23) Glocker, M.; Nock, S.; Sprinzl, M.; Przybylski, M. Chem.sEur. J. 1998, 4, 707-715. (24) Kalkum, M.; Przybylski, M.; Glocker, M. Bioconjugate Chem. 1998, 9, 226-235. (25) Fiedler, W.; Borchers, C.; Macht, M.; Deininger, S.; Przybylski, M. Bioconjugate Chem. 1998, 9, 236-241. (26) Glocker, M.; Borchers, C.; Fiedler, W.; Suckau, D.; Przybylski, M. Bioconjugate Chem. 1994, 5, 583-590. (27) Santrucek, J.; Strohalm, M.; Kadlcik, V.; Hynek, R.; Kodicek, M. Biochem. Biophys. Res. Commun. 2004, 323, 1151-1156. (28) Novak, P.; Kruppa, G.; Young, M.; Schoeniger, J. J. Mass Spectrom. 2004, 39, 322-328. (29) Sharp, J.; Becker, J.; Hettich, R. Anal. Biochem. 2003, 313, 216225. (30) Zappacosta, F.; Ingallinella, P.; Scaloni, A.; Pessi, A.; Bianchi, E.; Sollazzo, M.; Tramontano, A.; Marino, G.; Pucci, P. Protein Sci. 1997, 6, 1901-1909. (31) Hambly, D.; Gross, M. J. Am. Soc. Mass Spectrom. 2005, 16, 20572063. (32) Smith, D.; Deng, Y.; Zhang, Z. J. Mass Spectrom. 1997, 32, 135146. (33) Garret, R. A., Douthwate, S. R., Matheson, A. T., Moore, P. B., Noller, H. F. The Ribosome: Structure, Function, Antiobiotics, and Cellular Interactions; ASM Press: Washington, DC, 2000. (34) Vila-Sanjurjo, A.; Ridgeway, W.; Seymaner, V.; Zhang, W.; Santoso, S.; Yu, K.; Cate, J. Proc. Natl. Acad. Sci. U.S.A. 2003, 100, 86828687. (35) Ban, N.; Nissen, P.; Hansen, J.; Moore, P.; Steitz, T. Science 2000, 289, 905-920. (36) Harms, J.; Schluenzen, F.; Zarivach, R.; Bashan, A.; Gat, S.; Agmon, I.; Bartels, H.; Franceschi, F.; Yonath, A. Cell 2001, 107, 679-688. (37) Janecki, D.; Beardsley, R.; Reilly, J. Anal. Chem. 2005, 77, 72747281. (38) Beardsley, R. L.; Reilly, J. P. J. Am. Soc. Mass Spectrom. 2004, 15, 158-167. (39) Spedding, G. In Ribosomes and Protein Synthesis, A Practical Approach; Oxford Univiversity Press: New York, 1990. (40) Arnold, R.; Reilly, J. Anal. Biochem. 1999, 269, 105-112. (41) Karty, J. A.; Reilly, J. P. manuscript in preparation, 2005. (42) Martz, E. Trends Biochem. Sci. 2002, 27, 107-109. (43) Nierman, W. C.; Feldblyum, T. V.; Laub, M. T.; Paulsen, I. T.; Nelson, K. E.; Eisen, J.; Heidelberg, J. F.; Alley, M. R. K.; Ohta, N.; Maddock, J. R.; Potocka, I.; Nelson, W. C.; Newton, A.; Stephens, C.; Phadke, N. D.; Ely, B.; Deboy, R. T.; Dodson, R. J.; Durkin, A. S.; Gwinn, M. L.; Haft, D. H.; Kolonay, J. F.; Smit, J.; Craven, M. B.; Khouri, H.; Shetty, J.; Berry, K.; Utterback, T.; Tran, K.; Wolf, A.; Vamathevan, J.; Ermolaeva, M.; White, O.; Salzberg, S. L.; Venter, J. C.; Shapiro, L.; Fraser, C. M. Proc. Natl. Acad. Sci. U.S.A. 2001, 98, 4136-4141. (44) Huang, X.; Miller, W. Adv. Appl. Math 1991, 12, 337-357. (45) Lalign is found at http://www.Ch.Embnet.Org/software/ lalign_form.Html, Accessed October, 2005 (46) Fraternali, F.; Cavallo, L. Nucleic Acids Res. 2002, 30, 2950-2960. (47) Kleinjung, J.; Fraternali, F. Nucleic Acids Res. 2005, 33, W342W346. (48) Lee, B.; Richards, F. M. J. Mol. Biol. 1971, 55, 379-400. (49) Evershed, R.; Robertson, D.; Beynon, R.; Green, B. Rapid Commun. Mass Spectrom. 1993, 7, 882-886. (50) Running, W. E.; Ravipaty, S.; Reilly, J. P. manuscript in preparation, 2006.

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research articles (51) Mueller, F.; Sommer, I.; Baranov, P.; Matadeen, R.; Stoldt, M.; Wohnert, J.; Gorlach, M.; Van Heel, M.; Brimacombe, R. J. Mol. Biol. 2000, 298, 35-59. (52) Wilson, D. N.; Nierhaus, K. H. Crit. Rev. Biochem. Mol. Biol. 2005, 40, 243-267. (53) Ilag, L.; Videler, H.; Mckay, A.; Sobott, F.; Fucini, P.; Nierhaus, K.; Robinson, C. Proc. Natl. Acad. Sci. U.S.A. 2005, 102, 81928197. (54) Frank, J.; Agrawal, R. K. Nature 2000, 406, 318-322. (55) Agrawal, R. K.; Penczek, P.; Grassucci, R. A.; Joachim, F. Proc. Natl. Acad. Sci. U.S.A. 1998, 95, 6134-6138.

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Beardsley et al. (56) Hanson, C.; Fucini, P.; Ilag, L.; Nierhaus, K.; Robinson, C. J. Biol. Chem. 2003, 278, 1259-1267. (57) Mueller, F.; Sommer, I.; Baranov, P.; Matadeen, R.; Stoldt, M.; Wohnert, J.; Gorlach, M.; Van Heel, M.; Brimacombe, R. J. Mol. Biol. 2000, 298, 35-59. (58) Hanson, C. L.; Fucini, P.; Ilag, L. L.; Nierhaus, K. H.; Robinson, C. V. J. Biol. Chem. 2003, 278, 1259-1267.

PR060170W

Probing the Structure of the Caulobacter crescentus Ribosome with

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