Peroxiredoxin is a Versatile Self-Assembling Tecton for Protein

Apr 21, 2014 - Callaghan Innovation Research Limited, Lower Hutt, New Zealand .... Biosystems 3130xl Genetic Analyzer at University of Canterbury...
0 downloads 0 Views 2MB Size
Download PDF
Article pubs.acs.org/Biomac

Peroxiredoxin is a Versatile Self-Assembling Tecton for Protein Nanotechnology Amy J. Phillips,†,‡,§ Jacob Littlejohn,† N. Amy Yewdall,† Tong Zhu,† Céline Valéry,† F. Grant Pearce,† Alok K. Mitra,§ Mazdak Radjainia,§ and Juliet A. Gerrard*,†,‡,§,∥ †

Biomolecular Interaction Centre and School of Biological Sciences, University of Canterbury, Christchurch, New Zealand MacDiarmid Institute for Advanced Materials and Nanotechnology, Victoria University, Wellington, New Zealand § School of Biological Sciences, University of Auckland, Auckland, New Zealand ∥ Callaghan Innovation Research Limited, Lower Hutt, New Zealand ‡

S Supporting Information *

ABSTRACT: The potential for protein tectons to be used in nanotechnology is increasingly recognized, but the repertoire of stable proteins that assemble into defined shapes in response to an environmental trigger is limited. Peroxiredoxins (Prxs) are a protein family that shows an amazing array of supramolecular assemblies, making them attractive tectons. Human Prx3 (hPrx3) forms toroidal oligomers characteristic of the Prx family, but no structure has been solved to date. Here we report the first 3-D structure of this protein, derived from single-particle analysis of TEM images, establishing a dodecameric structure. This result was supported by SAXS measurements. We also present the first detailed structure of a double toroidal Prx from a higher organism determined by SPA. Guided by these structures, variants of the protein were designed to facilitate controlled assembly of protein nanostructures through the association of the toroids. We observed an enhanced population of stacked toroids, as seen by TEM; nanocages and interlocked toroids were also visible. Low pH was successfully predicted to generate long ordered nanotubes. Control over the length of the tubes was gained by adding ammonium sulfate to the assembly buffer. These versatile assembly properties demonstrate the considerable potential of hPrx3 as a tecton for protein nanotechnology.

■

INTRODUCTION Proteins are finding utility in a host of nanotechnological applications. Their potential for creating designer materials and intricate nanoscale devices is unquestioned, since biology is filled with examples of proteins acting in these roles. This promise has been heralded for the past decade or more,1−3 but the challenge of designing and, particularly, controlling protein−protein interactions in vitro to engender useful function is considerable.4 In the past decade, our understanding of how to create artificial protein constructs with predictable assembly properties has increased exponentially5 and a handful of useful protein tectons, or building blocks, are emerging.6−12 However, a bottleneck in the development of the field is the paucity of stable protein tectons endowed with the ability to self-assemble into a variety of nanoarchitectures. The peroxiredoxin (Prx) protein family can adopt an intriguing repertoire of supramolecular assemblies.13 They typically exist as obligate homodimers, which associate to form toroidal shaped oligomers. While a variety of factors are known to promote oligomerization,14−16 many aspects of Prx selfassembly and its role in function remain unknown. While there is no crystal or solution structure for human Prx3 (hPrx3), the crystal structure of the bovine homologue, which shares a 93% sequence homology with hPrx3, has been solved.17 However, there has been some debate as to the precise nature of the © 2014 American Chemical Society

bovine toroidal oligomer structure, with some results suggesting that the protein exists as a decamer,18 while others suggest that it is dodecameric.17 Since the crystal structure of bovine Prx3 shows an unusual concatenated arrangement of dodecameric toroids, which may be only representative of a subpopulation of the species in solution,13,17 it was not clear if the oligomeric state of isolated hPrx3 toroids is a dodecamer or a decamer, analogous to all other human Prxs. Such a knowledge is mandatory if rational design of “on-demand” higher order assemblies of hPrx3 are to be pursued. An attractive feature of the Prxs is the propensity of the protein toroids to associate into high molecular weight (HMW) complexes. Although the structural characterization of these HMW forms is sparse, the most reported assembly is a stack of toroidal oligomers,14,19,20 but other more complex structures, including tubes, clusters, and cages, have been reported.19,21,22 While the role or even the existence of these higher order structures in vivo is still very much under question, understanding the conditions that promote them in vitro attracted our interest from a nanotechnological perspective. Protein nanostructures are attracting increasing attention for their Received: February 19, 2014 Revised: April 15, 2014 Published: April 21, 2014 1871

dx.doi.org/10.1021/bm500261u | Biomacromolecules 2014, 15, 1871−1881

Biomacromolecules

Article

potential applications,23 and those that can assemble in response to an environmental trigger are of particular interest. Bovine Prx3 has been seen to form HMW assemblies, with stacked toroids reported in some instances.18 In this study, we report the detailed biophysical characterization of hPrx3 species in solution and propose a solution structure for two oligomeric forms, the toroid and double toroid, which were previously undetermined. We also present promising results toward designing novel structures with altered self-assembly properties, including the formation of interlocked toroids, similar to those observed for bovine Prx3 by X-ray crystallography,17 nanocages, and long ordered nanotubes.

■

column (GE Healthcare) previously equilibrated with running buffer (described above) and connected to a Viscotek 302−040 Triple Detector GPC/SEC system (ATA Scientific) operated at 28 °C. The 100 μL samples were injected onto the column and eluted at a flow rate of 0.5 mL/min. Absolute molecular weight, size, and size distributions were calculated using the refractive index, viscometry, and right- and low-angle light scattering measurements calibrated against a bovine serum albumin standard (66.5 kDa). All protein samples were run at 1 mg/mL for chromatographic analysis. Small Angle X-ray Scattering. Small angle X-ray scattering (SAXS) measurements were performed at the SAXS/WAXS beamline at the Australian Synchrotron. The radiation wavelength was set to 1.0332 Å and a Dectris-Pilatus detector used to record scattering patterns (1 M, 170 × 170 mm2, effective pixel size, 172 × 172 μm). The sample−detector distance was 1600 mm to provide a “q” (Fourier spacing) range of 0.0126−0.500 Å−1. Protein samples placed in a 1.5 mm glass capillary at 10 °C were previously centrifuged and filtered to remove any aggregates before data were collected. All samples were at concentrations of 0.5−2 mg/mL, and data were collected at 2 s intervals under continuous flow. 2D intensity plots were averaged using Scatterbrain software, and the background was subtracted. AUTOPOROD24 was used to calculate maximum particle size (Dmax) and Porod volume. Data sets were recorded in the form of 30 images, and scattering pattern analyses were carried out using PRIMUS software25 including Guinier fits. GNOM26 was used to perform indirect Fourier transforms and generate pair distribution function P(r) to indicate the relative probabilities of distances between scattering centers. Ab initio modeling was carried out using DAMMIN27 to generate ten bead models with imposed C6 symmetry, which were then averaged using DAMAVER and overlaid onto the crystal structure of the bovine homologue17 using SUPCOMB.28 An ab initio dummy-residue model was also generated using GASBOR29 with imposed C6 symmetry and overlaid onto the crystal structure of bovine Prx3. Transmission Electron Microscopy. hPrx3 preparations (0.05 mg/mL in reducing buffer: 20 mM HEPES, 150 mM NaCl, 2 mM TCEP, pH 8; or nonreducing buffer: 20 mM HEPES, 150 mM NaCl, pH 8; or ammonium sulfate buffer: 20 mM HEPES, 150 mM NaCl, 400 mM (NH4)2SO4) were applied onto copper mesh grids overlaid with continuous carbon. Briefly, 5 μL of the samples at 0.05 mg/mL were adsorbed onto glow-discharged grids for 1 min. Excess solution was removed with filter paper (Whatman #1), and the samples were negatively stained with NanoVan solution (Nanoprobes) for 1 min. Excess liquid was blotted with filter paper and the grid was allowed to air-dry. Images were acquired at a nominal magnification of 52000× using an FEI Tecnai 12 electron microscope operated at 120 kV. Electron micrographs recorded using Kodak ISO-163 film were developed for 10 min in D19 (Kodak) diluted 1:1 with deionized water. Single-Particle Image Processing. Electron micrographs were digitized using a Nikon Super Coolscan 9000 scanner at a raster step of 7.8 μm corresponding to a spacing of 1.5 Å on the specimen. A total of 6850 isolated tagged hPrx3 particles or 7130 cleaved hPrx3 particles prepared in reducing buffer were automatically selected using the boxer module of the EMAN v1.9 image processing suite30 and a window size of 200 × 200 pixels. To reduce high-and low-frequency noise components, a 20−200 Å band-pass filter was applied to the raw images, which were subsequently normalized. A total of 100 referencefree class averages were generated in nine rounds of iterative refinement by using the refine2d.py routine in EMAN v1.9. To initiate 3-D reconstructions of hPrx3 from conformationally homogeneous subsets of raw images, initial reference models were built using the startcsym routine of EMAN v1.9, which resulted in asymmetric toroidal-shape volumes. Five initial models were simultaneously refined by 12 iterative rounds of multimodel refinement of EMAN v1.9 with D6 symmetry imposed. The resulting models were separately refined with imposed D6 symmetry using only the selected set of raw images that the process of multimodel refinement had associated with a given reference model. Resolution of the reconstruction was determined by Fourier-shell correlation of two

MATERIALS AND METHODS

The hPrx3 cDNA (NCBI Accession No. NM_006793) was originally obtained from our collaborator Dr. Mark Hampton (University of Otago) and cloned into the pET151/D/LacZ vector (Invitrogen) without the mitochondrial leader sequence. The expression construct was transformed into E. coli DH5α cells for plasmid propagation and subsequently into Rosetta (DE3) cells (Novagen) for protein expression. A single colony was picked from the transformed cells and used to inoculate 10 mL of LB media supplemented with 100 μg/ mL ampicillin and 35 μg/mL chloramphenicol. This was grown overnight at 37 °C and then used to inoculate 1 L of LB medium plus antibiotic supplements and cultured at 37 °C to a cell density of OD600 = 0.6. The temperature was reduced to 26 °C, expression was induced with the addition of 0.6 mM isopropyl-β-D-thiogalactopyranoside (IPTG) and the culture was allowed to grow for a further 12 h at this temperature before harvesting. Cells were lysed in binding buffer (20 mM HEPES, 150 mM NaCl, 10 mM imidazole, pH 8.0) using a cell press (Microfluidics M110-P lab homogenizer) and the soluble fraction separated by centrifugation at 11000 rpm for 20 min. This was loaded onto a 5 mL HisTrap FF column (GE Healthcare) previously equilibrated with binding buffer (GE Healthcare). The column was washed with 20 mL binding buffer until the UV280 absorbance returned to baseline, and the protein was eluted using the gradient function to a final buffer composition of 20 mM HEPES, 150 mM NaCl, 500 mM imidazole, pH 8.0. Fractions containing the hPrx3 protein as identified by SDS-PAGE were pooled, further purified, and exchanged into running buffer (20 mM HEPES, 150 mM NaCl, pH 8.0) by gel filtration using a HiLoad 16/60 Superdex 200 column (GE healthcare). Unless otherwise stated, this buffer was used for all experiments; where reducing conditions were required, 2 mM TCEP was included in the running buffer. Where required, His-tag cleavage was achieved by incubation with 1 μg rTEV protease per mg hPrx3 protein at 4 °C for 24 h. rTEV protein, cleaved tags, and uncleaved protein were removed using a 1 mL HisTrap column (GE Healthcare) and fractions collected and checked with SDS-PAGE. For some TEM experiments, the His-tag was not cleaved, as images of the toroidal structure were more easily obtained in the presence of the tag. Only samples that were more than 95% pure were used for further experimental work. Mutagenesis. Cysteine-47 and serine-78 were exchanged respectively with serine and alanine using the Quik-Change sitedirected mutagenesis kit (Stratagene) to sequentially perform two single mutations. Plasmid pET151/D/LacZ (Invitrogen) containing the hPrx3 gene served as template. The following primer pairs (Invitrogen) were used: for the mutation of cys47 into serine, 5′-CCT TTG GAT TTC ACC TTT GTG AGT CCT ACA GAA ATT GTT GC-3′ (forward) and 5′-GCA ACA ATT TCT GTA GGA CTC ACA AAG GTG AAA TCC AAA GG-3′ (reverse); and for the mutation of ser78 into alanine, 5′-GTG GAT TCC CAC TTT GCC CAT CTT GCC TGG-3′ (forward) and 5′-CCA GGC AAG ATG GGC AAA GTG GGA ATC CAC-3′ (reverse). The fidelity of the amplification reactions was confirmed by DNA sequencing using an Applied Biosystems 3130xl Genetic Analyzer at University of Canterbury. Size Exclusion Chromatography (SEC)/Static Light Scattering. Purified hPrx3 samples were loaded onto a Superdex 200 10/300 1872

dx.doi.org/10.1021/bm500261u | Biomacromolecules 2014, 15, 1871−1881

Biomacromolecules

Article

reconstructions obtained from evenly split data sets using the FSC0.5 criterion.

■

RESULTS AND DISCUSSION hPrx3 Forms Dodecameric Toroids in Solution. In order to understand the assembly of hPrx3 toroids into higher order structures, it was first necessary to gain a detailed understanding of the single toroid structure and resolve whether it is a decamer or dodecamer, then to understand the nature of the interface between multiple toroids, known as the R interface.31 hPrx3 was expressed in E. coli and purified to homogeneity according to standard methods. Purified hPrx3 samples in reducing conditions were examined by transmission electron microscopy (TEM) after negative staining. (Details of biophysical characterization of each protein in dilute solutions, transmission electron microscopy, and single-particle analysis can be found in the Supporting Information.) hPrx3 particles were visible as ring-shaped toroidal assemblies (Figure 1a), which assumed a random orientation, and with clearly discernible hexagonal features. Projection class averages indicated rings with an external diameter of ∼16 nm and an internal diameter of ∼7 nm. The averaged images clearly showed the 6-fold symmetry of the oligomer (Figure 1b), with individual obligate dimers being easily distinguished. This suggests hPrx3 toroids are predominantly hexamers of the obligate protein dimers and therefore D6 symmetry was imposed for carrying out the 3-D reconstruction using the EMAN v1.9 image processing suite.30 The resulting reconstruction (Figure 1c) of the hPrx3 toroidal structure agrees well with X-ray crystal structure of bovine Prx317 (PDB ID: 1ZYE), which was docked into the 3-D reconstruction using UCSF Chimera.32 The resolution of the reconstruction was estimated to be 26 Å based on the FSC0.5 criterion, and shows density additional to the dodecameric Prx crystal structure. Raw images, projection averages, and the 3-D reconstruction of hPrx3 all clearly showed the hPrx3 oligomer to be a dodecamer. The 3-D reconstruction shows density additional to the dodecameric Prx crystal structure, which is well positioned to account for the 32 missing C-terminal residues from the crystal structure of the bovine analogue.17 The modeled C-terminal helices are angled toward the external face of the toroid, away from the dimer− dimer interface, suggesting that the missing residues may protrude from the toroid. These results resolve the uncertainty surrounding the toroidal form of hPrx3 and, in addition, were corroborated by solution studies by SAXS and SEC. Under reducing conditions, the SAXS intensity plot (Figure 2) showed multiple shoulders, indicating a multidomain particle with a number of intra- and interdomain distances. The volume of the particle calculated from the scattering curve was found to be consistent with the volume calculated for the bovine Prx3 dodecamer (Table S1). Data was collected for a range of concentrations, and the particle volume was not found to vary with protein concentration. This range is considered to be representative of physiological conditions, as the total concentration of hPrx3 within the cell is estimated at 2.5 mg/mL.38 This is also in good agreement with static light scattering results (Supporting Information, Figure S2), which gave a mass of 249 kDa. The simulated curve generated by the ab initio models showed excellent agreement to the scattering curve (Figure 2). The calculations and evaluation of particle size and volume (Supporting Information, Table S1) are consistent with the monodisperse dodecameric structure seen by TEM, which has a

Figure 1. TEM micrographs, projection averages, and 3-D reconstructions revealed a dodecameric toroid: (a) Electron micrograph showing a representative field of projected views of randomlyorientated hPrx3 particles, including top (black arrows) and side views (white arrows) of toroids. Samples were prepared in reducing conditions at 0.05 mg/mL. Scale bar is 150 nm. (b) A selection of enlarged projection raw images (first column) and class averages obtained from the classification of 7130 raw particles showing clearer en face profiles, as well as side views of rings (second and third columns). The fourth column shows class averages that were obtained from hPrx3 particles suspended in stain covering holes in a holey carbon film. These particles showed strong preferential orientation, and only top views were observed. Class averages showing top views of particles stained across holes (fourth column) show that the carbon film was not selected for dodecamers and biased the outcome. Box size corresponds to 30 nm; particle diameter is ∼16 nm. (c) A 26 Å 3-D reconstruction was generated by single-particle analysis with imposed D6 symmetry and contoured with a threshold of 5.5 (green) and 2 (transparent gray) sigma. The model is consistent with the crystal structure of a dodecameric Prx15 (PDB ID: 1ZYE), as evidenced by a good fit upon docking with the crystal structure (which gave a density cross-correlation coefficient of 0.94; right column). Scale bar is 5 nm.

theoretical mass of 257 kDa. Pair distribution function (P(r)) plots showed a nonhyperbolic curve and the ab initio models were consistent with a low electron density in the center of the oligomer, corroborating a toroidal structure (Figure 2b), which was supported by bead models generated from the scattering data (Figure 2c,d). Overlaying the model with the crystal structure of the bovine form in an analogous manner to that carried out for the TEM model again showed a good fit, with the additional C-terminal residues absent from the crystal structure present in a position consistent with the electron density in the TEM structure. 1873

dx.doi.org/10.1021/bm500261u | Biomacromolecules 2014, 15, 1871−1881

Biomacromolecules

Article

Figure 2. SAXS data of reduced hPrx3: Scattering data were used to generate intensity plots (a) and P(r) plots (b), which together supported a toroidal shaped species. Ab initio bead models generated by GASBOR (c) and DAMMIN (d) overlay well with the crystal structure and confirm the toroidal nature of the oligomer.

Under reducing conditions, the protein eluted as a single peak (249 kDa), consistent with the mass expected for the dodecamer. To determine the effect that the His-tag may be having on the assembly of hPrx3,39 wild-type His-tagged hPrx3 was also run on the size exclusion column under nonreducing conditions (Figure S3). While the tagged protein was predominantly dodecameric, at 1 mg/mL, the protein showed a small leading peak that had not been observed for cleaved hPrx3 (Figure S2). The molecular weight of this peak was determined using the inline scattering detectors, and it was found to have a mass of 470 kDa, consistent with the presence of stacked dodecamers in solution. At lower protein concentrations, smaller species consistent with dimers and tetramers were also observed. S78A-hPrx3 was found to be a dodecameric species under reducing conditions (Figure S4) and tended to form higher order species under nonreducing conditions. Both His-tagged and nontagged C47SS78A-hPrx3 formed a stable dodecamer under both reducing and nonreducing conditions, with a leading peak that is consistent with a small amount of stacked dodecamers in solution (Figure S5). Dodecameric Toroids Stack into Higher Order Assemblies. Using single particle analysis of images of 6850 particles, a 21 Å 3-D reconstruction was generated for the double toroid assembly (Figure 3). The 3-D model showed

Superposition of scaled scattering curves from a number of concentrations within the range tested (0.5−2 mg/mL; data not shown) revealed no interparticle interference, indicating that within this range the oligomeric state of reduced hPrx3 is not concentration dependent. Effect of Redox Conditions on hPrx3 Oligomeric State. The oligomeric state of Prx proteins shift upon oxidation. In the reduced state, the protein is generally toroidal, but in the absence of reducing agent, an equilibrium is established that favors a dimeric species.15,16,38 As well as being important in the biological function of the protein, this change in oligomeric state in response to an environmental trigger is an attractive feature of a protein used for nanostructure assembly in vitro.23 In nonreducing conditions, SAXS data showed a shift in particle size; however, the resulting mixture of species made definitive mass calculations problematic (data not shown). To obtain information about the size of hPrx3 assemblies under conditions where a monodisperse preparation could not be obtained, size exclusion chromatography with in-line light scattering was used to monitor the change in oligomeric state induced by reducing and nonreducing conditions (Figure S2). Under nonreducing conditions, the protein was predominantly dimeric (45 kDa) with a small peak observed eluting at 14.5 mL, which may be a tetrameric species that has previously been observed for a Prx derived from plant species,32 but not hPrx3. 1874

dx.doi.org/10.1021/bm500261u | Biomacromolecules 2014, 15, 1871−1881

Biomacromolecules

Article

Figure 3. TEM and docking of the crystal structure of the bovine homologue revealed the stacking interface of hPrx3: (a) Class averages from micrographs of the His-tagged hPrx3 sample showed the presence of stacks of toroids. (b) 3-D reconstruction of the two stacked toroids shows that contact between the dodecamers, rotation, and the gap seen in side-on views are likely caused by interactions being made only at the tips of dimers (left panel). The right panel shows our pseudoatomic interpretation of the hPrx3 stack reconstruction using atomic coordinates obtained for decameric SmPrx1stacks. Four adjacent dimers in the crystal structure of stacked decameric SmPrx131 (PDB ID: 3ZVJ, chains ABCDJKLM) were extracted and docked in three blocks [blue, red, and rainbow colors (rainbow colors are rendered from blue to red for residues from N-terminus to C-terminus; α6 helices are the C-most helices, colored red)]. The central panel shows the full crystal structure of the SmPrx131 (PDB ID: 3ZVJ) stacked decameric toroids that share the same height as the reconstructed EM map of stacked hPrx3 dodecamers.

Figure 4. TEM micrographs recorded from samples of (a) wild-type hPrx3 at 0.05 mg/mL in standard running buffer at pH 8.0, and (b) the same sample at 0.5 mg/mL dialyzed into an identical buffer, which had been adjusted to pH 4.0 with HCl. Higher order structures were not seen when the protein concentration was increased at pH 8.0; however, tubes formed most readily at a slightly increased concentration.

caused by conformational changes of secondary structure elements, which allow the necessary interactions for stacking to occur.31 A putative alternate model of stacking, featuring an orthorhomboid array of Prx2, was provided by Harris et al.20 Our 3-D reconstruction of stacked hPrx3, similar to the SmPrx1 model (Figure 3b), shows that toroids are rotated with respect to each other. The likely contacts at the α6 helices of alternate monomers are consistent with the large gap that can be seen

overlapping density envelopes indicating significant contact between the toroids, with interactions appearing to occur at the ends of the α6 helices of the protein.33 These helices project toward the top and bottom face of the toroid alternately, as the monomers are oriented as antiparallel dimers (Figure 3b). The two stacked decameric toroids that feature in the reported Xray crystal structure of the SmPrx1 “chaperone” species are rotated by 18° around the 5-fold symmetry axis, a rotation 1875

dx.doi.org/10.1021/bm500261u | Biomacromolecules 2014, 15, 1871−1881

Biomacromolecules

Article

Figure 5. Samples of wild-type hPrx3 dialyzed into a pH 4.0 buffer containing (a) 100 mM (NH4)2SO4; (b) 200 mM (NH4)2SO4; (c) 400 mM (NH4)2SO4. The size of the nanotubes formed decreased with increasing concentration of sulfate ions, resulting in fairly uniform stacks.

Figure 6. Positions of important residues and site-directed mutations. (a) Detail showing the A and B interfaces in context of the toroid. The α2 helix (light blue) and α6 helix (dark blue) positions are indicated for one 2-cys peroxiredoxin dimer (green). Within the active site are the spatially conserved P40, T44, and R123 residues (red) arranged around the CP (white spheres). The position of S78 (purple) at the A interface is also indicated. (b) Side view to further highlight the positioning of α2 and α6 helices that are involved in toroid stacking. The blue residues represent E20, K22, and A163 (as H164 cannot be seen in the crystal structure); these are hypothesized to mediate toroid stacking at the R interface. Figures were drawn using PyMOL Molecular Graphics System, Version 1.3 Schrödinger, LLC using the X-ray crystal structure of the bovine Prx3 protein PDB 1ZYE.

density map appears to be consistent with the stacking interface of the decameric crystal structure. Long Nanotubes, Cages, and Concatenated Structures Form in Response to Environmental Conditions. Our model indicates the R interface of hPrx3 to be characterized by salt bridges between key residues (Figure 6b): E20, K22, and H164, as judged by comparison to the crystal structure of Saccoccia et al.,31 which led us to hypothesize that changing the ionic strength and the pH of the surrounding buffer solution would alter the electrostatic interaction and, therefore, the propensity for stacking of

between toroids in class averages of side-on views of stacks. Our 3-D reconstruction of dodecameric hPrx3 and the decameric SmPrx1 are of similar height, around 8 nm at the farthest points. With the assumption that the mode of stacking that is seen in SmPrx1 is also present in our hPrx3 reconstruction, we docked three blocks of four adjacent dimers extracted from the SmPrx1 crystal structure31 (PDB ID: 3ZVJ) into the EM density map of stacked hPrx3 toroids. A perfect fit of these structures cannot be expected because of the decameric nature of SmPrx1; however, the stacking interface visualized in our EM 1876

dx.doi.org/10.1021/bm500261u | Biomacromolecules 2014, 15, 1871−1881

Biomacromolecules

Article

Figure 7. Samples of untagged S78A-hPrx3x in nonreducing conditions were seen to exist predominantly as toroidal oligomers, suggesting the dodecamer was stabilized by the mutation. The appearance of uniform, spherical structures (highlighted with boxes and shown in enlargement) suggested that the mutation encouraged the formation of nanocages, similar to that seen by other groups.22

It seems likely that for hPrx3 also (NH4)2SO4 is limiting stack formation in a similar fashion, presumably binding in the active site and stabilizing the folded form of helix α2. The incomplete inhibition of nanotube formation would suggest sulfate ions bind somewhat more weakly to the native cysteine, than to the serine of C48S-SmPrxI, possibly due to the reduced electronegativity of sulfur compared to oxygen. We sought to probe the intertoroid interactions using sitedirected mutagenesis in order to stabilize the dodecamer without relying on the pH sensitive His-tag. The sites of mutation are illustrated in Figure 6. Previous studies had shown that single residue mutations can have a significant impact on Prx structure,14,34 and the mutation S78A-hPrx3 was therefore designed by analogy. Inspection of preparations of the untagged S78A-hPrx3 sample under nonreducing conditions by TEM revealed higher molecular weight species in the form of a nanocage, rather than stacked structures (Figure 7). Such species have previously been observed for human erythrocyte Prx220 and were seen to be present in dilute solution (see Supporting Information, Figure S4) in the case of S78A-hPrx3. Initial characterization by SEC/SLS and SAXS suggested that untagged C47SS78A-hPrx3 was primarily present as a single species with a volume of 569210 Å3 and a molecular mass of ∼300 kDa in nonreducing conditions, which would suggest a single toroidal ring (see Supporting Information, Table S1 and Figure S7). A shoulder peak is seen in the SEC/SLS elution profile, which is indicative of larger species. Molecular weight calculations suggest this shoulder to represent a ∼600 kDa species which would correspond to two stacked toroids (see Supporting Information, Figure S5). TEM analysis was carried out on His-tagged samples, which was seen to behave identically to the untagged construct in solution (Figure S5). Inspection of negatively stained specimens confirmed the

toroids. Such stacking has recently been observed at low pH for His-tagged SmPrx1.31 For the His-tagged hPrx3, lowering the pH did not lead to an increase in stacking of the toroids, which we propose is due to electrostatic repulsion between protonated histidine residues. With the His-tag removed, a range of pHs were examined and it was discovered that lowering the pH below 6 led to a dramatic increase in the propensity of the hPrx3 to form ordered nanotubes, with pH 4 producing especially long ordered structures (Figure 4). This is consistent with the notion that toroid stacking is mediated by contacts involving residues of the α6 helices, suggested by the 3-D structure of the double toroid described above. Lowered pH has been seen to encourage the movement of this helix to enable the R interface interactions to occur in the case of a decameric Prx,31 and these results indicate that the mechanism is conserved in the dodecameric hPrx3. Changing the ionic strength of the solution by adding ammonium sulfate (NH4)2SO4 was also investigated. While altering the ionic strength at standard pH (8.0) had no effect, the presence of (NH4)2SO4 in the low pH (4.0) buffer prior to exchanging the protein into the buffer appeared to impede nanotube formation (Figure 5). The length of the tubes was inversely correlated to the concentration of (NH4)2SO4. It is possible that this phenomenon is related to the (NH4)2SO4 altering starting the protein concentration in solution. However, this is similar to the result observed recently for C48S-SmPrxI, suggesting that it is a specific effect, although in the case of hPrx3, nanotube formation was not completely prevented. Whatever the cause, adjusting the (NH4)2SO4, provides a means to control the length of the tubes formed. For SmPrxI, Angelucci et al.14 hypothesized that sulfate ions stabilize the fully folded site of C48S by making polar contacts with the substituted serine and the conserved arginine (R124). 1877

dx.doi.org/10.1021/bm500261u | Biomacromolecules 2014, 15, 1871−1881

Biomacromolecules

Article

for the C-terminal residues that could not be modeled in the Xray crystal structure. Its location on the external face of the toroid is in line with the suggestion of Angelucci et al.14 that the unfolded C-terminal arm exposes a binding site, as it appears to move out of the way of the top face of the toroid. The reason as to why the formation of stacked toroids favors two in solution, as confirmed by examination of negatively stained specimens by TEM, requires further explanation, in view of the fact that assemblies of longer structures can be driven to form at low pH. When fibrillar structures assemble, competing events of nucleation, elongation, and termination occur, and the preponderance of any is critically dependent on the kinetics of each. In particular, the number of nucleation sites plays a large role in determining whether a few long structures, or many shorter structures occur. Our data suggest that lowering the pH reduces the number of nucleation sites, leading to longer structures. We have shown that, in the case of hPrx3, considerable versatility in quaternary structure is accessible by single mutations. S78A-hPrx3 forms nanocages, hinting at the presence of a small decameric subpopulation, whereas C47SS78A-hPrx3 forms concatenated structures and stacks. Introducing a mutation to disrupt the active site architecture drives the stacking and concatenation of toroids, in an analogous fashion to the SmPrx1 case. The formation of nanocages by S78A-hPrx3 may be due to unfolding of the active site and CP loop induced by this mutation. An analogous mutation in S. typhimurium AhpC34 stabilized the toroid, which was shown in later studies showed to coincide with increased movement around the CP loop.35 Thus, stabilizing the toroid and restricting the conformational changes that usually induce dissociation may force the CP loop to adopt a spatially different position, encouraging nanocage formation. Comparison to Previous Peroxiredoxins Studied in a Biomedical Context. Most examples of 2-cys Prxs in the literature have been shown to form decameric toroids,15,33,41 and in the case of other quaternary arrangements, the functional implications of the oligomeric structures are unclear. The tertiary structure of Prx monomers is highly conserved throughout the entire family, and within the 2-cys Prxs the differences seen between a decameric and dodecameric Prx are often surprisingly minor. This suggests that the size of the oligomer is finely tuned and governed by subtle sequence modifications. hPrx3 is shown here to be dodecameric. All other human 2-cys Prxs solved to date form decameric toroids, and interestingly, the only other eukaryotic Prx seen to be dodecameric is also mitochondrial.17 Overall, there is a strong correlation with the structure of Saccoccia et al.31 suggesting similar principles dictating interactions between the apposed toroids. Thus, in our deduced pseudoatomic model of hPrx3 double toroids, we find that three of the four residues proposed by Saccoccia et al.31 to be important in the interface are conserved in hPrx3 (E20, K22, and H164), which suggests that these interactions may be critical for optimal electrostatic stabilization of this intertoroid interface (Figure 6b). The fourth residue involved in the interface of SmPrx1 is H165, which is replaced by a threonine in hPrx3. In line with our finding, Saccoccia et al.31 observed that the R interface was stabilized at pH 4.2, at which the double decameric toroid structure was crystallized, in contrast to the single decameric toroid, crystallized at pH 7.6. They suggested

presence of double dodecamers and larger assemblies, revealing stacked toroids. In these specimens, the characteristic side views of a single toroid were almost completely absent, suggesting that specimen preparations for TEM promote the larger assemblies (see Supporting Information, Figure S6c). End-on views showed a different profile to those in the case of the native protein, seemingly thicker rings, suggesting the presence of two or more associated toroids. Projection averages revealed no side views of single toroids, suggesting that these comprise a relatively smaller population than in the wild-type samples, and stacks of up to six toroidal rings were seen in electron micrographs. Quantification of the number of stacked species observed by TEM could be complicated by the possibility that different species may vary in their propensity to attach to the carbon substrate. Mindful of this caveat, an approximation of the populations present for wild-type protein, with and without the tag, and for C47SS78A-hPrx3 was obtained by examining the number of particles in each class40 used to calculate the class averages. Figure S7 illustrates the difference in population classes. The His-tag clearly promotes a double toroidal structure of the wild-type protein, which is sparse in samples where the His-tag is removed, as evidenced by the absence of the side-views of stacks. Classification of the tagged specimen indicated projection averages that correspond to both sideviews and end-on views of stacks with side views of single toroids being absent. The C47SS78A mutation also changes the distribution of the toroid population strongly, favoring larger stacks. In contrast to the wild-type protein, presence of the His-tag appears to have no effect on the presence of high molecular weight assemblies of C47SS78A-hPrx3 (see Supporting Information, Figure S5). Furthermore, addition or removal of reducing agents and altering protein concentration did not alter the SEC/SLS elution profile or the SAXS results for this sample (Figure S5), with no dissociation to dimers being seen under any conditions. The second peak relating to a doubledodecameric species is seen in the SEC trace, which is not observed for untagged wild-type hPrx3. As with the His-tagged wild-type sample, it appears that specimen preparation for TEM is promoting the formation of larger assemblies, but the significant differences between images of C47SS78A-hPrx3 and His-tagged wild-type hPrx3 suggest that the CP of hPrx3 is essential for maintaining the active site structure. This is in line with the hypothesis suggested by Saccoccia et al. for SmPrx1.31 When mutated, the proposed stacking interactions are enabled, likely through the helix unwinding identified by Saccoccia et al.14,31 A number of conformations in addition to the toroid appear to be accessible to Prxs. In the C47SS78A-hPrx3 sample, a small percentage of the toroids were seen to interlock (Figure 7c). Concatenation of Prxs has been reported previously,17 and the agreement with this crystal structure leads us to believe that the apparent concatenated structures are promoted by the double mutation. Our interpretation of the assembly process is in line with the generally held view based on the atomic details of the Prx protein family, derived from X-ray crystallography (see Supporting Information). The role of C-terminal truncation inferred by Angeluci et al.14 supports our data, which suggest that a movement of the C-terminus occurs when the toroids stack. In both the TEM and the SAXS studies, a volume of density on the external face of the toroid is consistently seen. This is well placed to account 1878

dx.doi.org/10.1021/bm500261u | Biomacromolecules 2014, 15, 1871−1881

Biomacromolecules

Article

seen in other Prxs to have chaperone activity,31 a potential reason for the unusual dodecameric quaternary arrangement may be because the size of the toroid is tuned to provide an appropriately sized lumen for the substrate. The role of C-terminal truncation inferred by Angeluci et al.14 supports our data which suggest that a movement of the Cterminus occurs when the toroids stack. In both the TEM and SAXS studies, density on the external face of the toroid is consistently seen. This is well placed to account for the Cterminal residues that could not be modeled in the X-ray crystal structure. Its location on the external face of the toroid is in line with the suggestion of Angelucci et al.14 that the unfolded Cterminal arm exposes a binding site, as it appears to move out of the way of the top face of the toroid. The switching of Prxs between dimer and oligomer is known to be related to the oxidation state of CP, and changes in the redox state are proposed to induce helical unwinding around CP which in turn destabilizes the interface, causing the transition.15 Interestingly, in both cases where concatenation has been seen, the protein contains a mutation of a cysteine residue (the resolving cysteine, CR, in the work of Cao et al.15 and CP in this work). The native protein exists as a dimer in oxidizing conditions with a disulfide bond forming between the cysteine residues from adjacent monomers, requiring the Cterminal arm of one monomer to reach across the other, forming a domain swapped dimer. It has been suggested that dimers associate into toroids when the disulfide bond is broken in reducing conditions. However, an intermediate oxidized toroid has been seen, suggesting that the dimers may in fact associate while they are still disulfide-bonded.44,45 Earlier studies have reported that addition of ammonium sulfate decreases the propensity for toroids of Prx to stack when the CP was mutated to serine.18 This result is suggested to be due to the stabilization of the folded active site by the binding of the sulfate ion to the serine residue, and a nearby arginine. Increased ionic strength has been seen to encourage or prevent oligomerization of different Prxs, regardless of the nature of the ions.46−48 Angelucci et al.14 hypothesize that sulfate ions stabilize the fully folded site of C48S by making polar contacts with the serine and the conserved arginine (R124). In our case, it seems likely that (NH4)2SO4 is the limiting stack formation in a similar fashion, presumably binding in the active site and stabilizing the folded form of helix α2. The incomplete inhibition of nanotube formation would suggest sulfate ions bind more weakly to the native cysteine, possibly due to the reduced electronegativity of sulfur compared to oxygen. This result is also consistent with the suggestion14 that unfolding of the C-terminus alone is not sufficient for toroid stacking, and could provide an avenue to test Angelucci et al.’s hypothesis that the binding of substrates occur at the end of the tubes. This is noteworthy, as we present an approach to controlling the length of the tubes that does not involve mutation or other alterations to the primary structure. We speculate that in the case of C47SS78A-hPrx3, the absence of the conformational changes required for disulfide bond formation may allow the interactions that cause toroids to interlock to occur. It may also be that the mechanism for C47SS78A-hPrx3 subunit association differs from that of the native protein, as it does not rely on the reduction of CP. As the subunits, which are not in the disulfide-bonded, domain swapped state, as seen in native dimers, begin to associate, the newly forming toroid may have slightly different

that the toroidal stacks formed due to the disruption of an ionic bond between the active site cysteine (CP) and a conserved arginine at low pH or upon oxidation of CP to the sulfinic or sulfonic acid form. These changes allow the unwinding of the helix containing CP and movement of the unfolded loop to form the appropriate intertoroid interactions. This hypothesis is supported by experimental evidence that mutation of CP promotes formation of HMW structures.14,18 However, in other Prxs, mutation of this residue has been reported to prevent HMW complex formation,42,43 suggesting that results from Prx from one species cannot necessarily be translated to another. The results presented here are consistent with the mechanism of R interface formation put forward by Saccoccia et al.31 Recently, Angelucci et al. published an elegant study in which various mutations were introduced at the active site cysteine of SmPrx1,14 to mimic different oxidation states of the wild-type structure. Of these, introduction of a serine in place of a cysteine residue, designed to mimic the protonated active site cysteine, was shown to induce the stacked structure.14 The active site architecture of Prxs is such that the pKa of CP in the wild-type protein is lowered to around 6,16,42 meaning that it is deprotonated at physiological pH and can form an ionic bond with the nearby arginine, R124 in the case of SmPrx1 (Figure 7a). The higher pKa of serine in this mutant does not allow deprotonation in our experimental conditions (pH 8.0), and therefore the ionic bond to R124 is predicted to be disrupted, facilitating stabilization of the R interface. Until very recently,31 there has been a paucity of structural information relating to the HMW “chaperone species” of Prx. Some groups have suggested the structures formed are clusters of particles,48 while others suggest stacks.18 Angelucci et al.14 have recently suggested that the high molecular weight species formed by different Prxs (nanocages or stacks) are specific to individual species. In contrast, we have shown that in the case of hPrx3, considerable versatility in quaternary structure is accessible by single mutations. S78A-hPrx3 forms nanocages, hinting at the presence of a small decameric subpopulation, whereas C47SS78A-hPrx3 forms concatenated structures and stacks. Introducing a mutation to disrupt the active site architecture drives the stacking and concatenation of toroids, in an analogous fashion to the SmPrx1 case. The formation of nanocages by S78A-hPrx3 may be due to unfolding of the active site and CP loop induced by this mutation. An analogous mutation in AhpC,34 which stabilized the toroid, was later shown to cause increased movement around the CP loop.35 Thus, stabilizing the toroid and restricting the conformational changes that usually induce dissociation may force the CP loop to adopt a spatially different position, encouraging nanocage formation. Angelucci et al.14 postulate that formation of stacks at low pH is relevant to the pathology of schistosomiasis, where low pH is encountered in the life cycle. Other Prxs shown to do this include a Prx2 formerly known as calpromotin, that has been seen to shift to a higher molecular weight species at pH 5,44 as well as the hPrx3 results that we present here. A systematic survey of Prxs from different organisms would be interesting to unravel the extent to which the pHs encountered in the natural biological environment influence the tendency of these proteins to assemble into higher order forms. Complementing previous work,31,44 we suggest that these structures are not an artifact of the crystal or TEM specimen preparation. Given that hPrx3 appears to be able to form HMW assemblies analogous to those 1879

dx.doi.org/10.1021/bm500261u | Biomacromolecules 2014, 15, 1871−1881

Biomacromolecules

Article

We also thank the staff at the Australian synchrotron for their assistance during data collection at the SAXS/WAXS beamLine, and the New Zealand Synchrotron Group for travel funding

dimensions that are more conducive to interlocking. This could explain why it has only been seen in Prxs with cysteine mutations and is supported by the absence of such structures in both tagged and untagged wild-type samples. Stacking has been seen to occur in a number of Prxs, influenced by the addition of metal ions, the presence of affinity tags, and the disruption of the active site architecture.13,31,37 Although the observation of HMW species in samples extracted from native cells31 suggests the stacks may be physiologically relevant, the absence of such structures in solutions of pure, native protein in vitro indicates that their formation must be controlled by other factors. Angelucci et al.14 suggest that unfolding of the α2 helix and the C-terminus are “strongly coupled and cannot occur independently” as the unfolded helix is predicted to have a steric clash with the C-terminus. Our TEM and SAXS data, which reveal the location of the Cterminal residues missing from the X-ray crystal structure, lend experimental credence to this hypothesis.

■

(1) Zhang, S. G. Biotechnol. Adv. 2002, 20, 321. (2) Waterhouse, S. H.; Gerrard, J. A. Aust. J. Chem. 2004, 57, 161. (3) Ellis-Behnke, R. G.; Liang, Y. X.; You, S. W.; Tay, D. K.; Zhang, S. G.; So, K. F.; Schneider, G. E. Proc. Natl. Acad. Sci. U.S.A. 2006, 103, 5054. (4) Clarke, J.; Regan, L. Curr. Opin. Struct. Biol. 2010, 20, 480. (5) Heddle, J. G.; Fujiwara, I.; Yamadaki, H.; Yoshii, S.; Nishio, K.; Addy, C.; Yamashita, I.; Tame, J. R. H. Small 2007, 3, 1950. (6) Miranda, F. F.; Iwasaki, K.; Akashi, S.; Sumitomo, K.; Kobayashi, M.; Yamashita, I.; Tame, J. R. H.; Heddle, J. G. Small 2009, 5, 2077. (7) Padilla, J. E.; Colovos, C.; Yeates, T. O. Proc. Natl. Acad., Sci. U.S.A. 98, 2217. (8) Lai, Y.-T.; Cascio, D.; Yeates, T. O. Science 2012, 336, 1129. (9) Medalsy, I.; Dgany, O.; Sowwan, M.; Cohen, H.; Yukashevska, A.; Wolf, S. G.; Koster, A.; Almog, O.; Marton, I. Nano Lett. 2008, 8, 473. (10) Ballister, E. R.; Lai, A. H.; Zuckermann, R. N. Proc. Natl. Acad. Sci. U.S.A. 2008, 105, 3733. (11) Brodin, J. D.; Carr, J. R.; Sontz, P. A.; Tezcan, F. A. Proc. Natl. Acad. Sci. U.S.A. 2014, 111, 2897−2902. (12) Buer, B.C.; Marsh, N. Protein Sci. 2012, 21, 453−462. (13) P. A., Karplus, Hall, A. Structural survey of the peroxiredoxins. In Peroxiredoxin Systems; Flohé, L., Harris, J. R., Eds.; Springer Science and Business Media Inc.: New York, NY, 2007; pp 41−60. (14) Angelucci, F.; Saccoccia, F.; Ardini, M.; Boumis, G.; Brunori, L.; Di Leandro, R.; Ippoliti, R.; Miele, A. E.; Natoli, G.; Scotti, S.; Bellelli, A. J. Mol. Biol. 2013, 425, 4556. (15) Barranco-Medina, S.; Lázaro, J.-J.; Dietz, K.-J. FEBS Lett. 2009, 583, 1809. (16) Wood, Z. A.; Schröder, E.; Harris, J. R.; Poole, L. B. Trends Biochem. Sci. 2003, 28, 32. (17) Cao, Z.; Roszak, A. W.; Gourlay, L. J.; Lindsay, J. G.; Isaacs, N. W. Structure 2005, 13, 1661. (18) Gourlay, L.; Bhella, D.; Kelly, S. M.; Price, N. C.; Lindsay, J. G. J. Biol. Chem. 2003, 278, 32631. (19) Kato, H.; Asanoi, M.; Nakazawa, T.; Maruyama, K. Zool. Sci. 1985, 2, 485. (20) Harris, J. R.; Schröder, E.; Isupov, M. N.; Scheffler, D.; Kristensen, P.; Littlechild, J. A.; Vagin, A. A.; Meissner, U. Biochim. Biophys. Acta 2001, 1547, 221. (21) Jang, H. H.; Lee, K. O.; Chi, Y. H.; Lee, J. R.; Lee, S. S.; Moon, J. C.; Yun, J. W.; Choi, Y. O.; Kim, W. Y.; Kang, J. S.; Cheong, G.-W.; Yun, D.-J.; Rhee, S. G.; Cho, M. J.; Lee, S. Y. Cell 2004, 117, 625. (22) Meissner, U.; Schröder, E.; Scheffler, D.; Martin, A. G.; Harris, J. R. Micron 2007, 38, 29. (23) Heddle, J. G. Nanotechnol. Sci. Appl. 2008, 1, 68. (24) Petoukhov, M. V.; Konarev, P. V.; Kikhney, A. G.; Svergun, D. I. J. Appl. Crystallogr. 2007, 40, s223. (25) Konarev, P. V.; Volkov, V. V.; Sokolova, A. V.; Koch, M. H. J.; Svergun, D. I. J. Appl. Crystallogr. 2003, 36, 1277. (26) Svergun, D. I. J. Appl. Crystallogr. 1992, 25, 495. (27) Svergun, D. I. Biophys. J. 1999, 76, 2879. (28) Kozin, M. B.; Svergun, D. I. J. Appl. Crystallogr. 2001, 34, 33− 41. (29) Svergun, D. I.; Petoukhov, M. V.; Koch, M. H. J. Biophys. J. 2001, 80, 2946. (30) Ludtke, S. J.; Baldwin, P. R.; Chiu, W. J. Struct. Biol. 1999, 128, 82. (31) Saccoccia, F.; Micco, P. D.; Bournis, G.; Brunori, M.; Koutris, I.; Miele, A. E.; Morea, V.; Sriratana, P.; Williams, D. L.; Bellelli, A.; Angelucci, F. Structure 2012, 20, 429. (32) Pettersen, E. F.; Goddard, T. D.; Huang, C. C.; Couch, G. S.; Greenblatt, D. M.; Meng, E. C.; Ferrin, T. E. J. Comput. Chem. 2004, 25, 1605.

■

CONCLUSION The insights into the supramolecular assembly provided by this work provide a route to control the structures formed from selfassembling protein subunits and drive formation of novel structures. The observation that pH can trigger the assembly of long nanotubes and ammonium sulfate can facilitate the assembly of tubes of relatively homogeneous sizes augurs well for the use of Prxs as tectons for protein nanotechnology.36 Building directly on our detailed understanding of this system from the biomedical literature, work is underway to optimize this process since homogeneous preparations of ordered structures are attractive components for bionanotechnology. Examples of future applications include bionanocomposites, optical devices, biosensors, and drug delivery vehicles.31

■

ASSOCIATED CONTENT

S Supporting Information *

Additional material, including biochemical and biophysical characterization of hPrx3, is available. This material is available free of charge via the Internet at http://pubs.acs.org.

■

REFERENCES

AUTHOR INFORMATION

Corresponding Author

*E-mail: [email protected]. Funding

Funding for this work was received in part from the MacDiarmid Institute for Advanced Materials and Nanotechnology, the Biomolecular Interaction Centre, the New Zealand Synchrotron group, and the U.S. Army Research Office. Notes

The authors declare no competing financial interest.

■

ACKNOWLEDGMENTS The authors would like to thank collaborators at the University of Otago, in particular, Associate Professor Mark Hampton, for his kind gift of the hPrx3 cDNA, and for continuing valuable discussion. In addition, collaborators at the University of Auckland, particularly Dr. Shaun Lott and his research group, are thanked for the use of laboratory facilities and ongoing discussion. We thank Dr. Adrian Turner for the maintenance of the TEM facility at the School of Biological Sciences and Jackie Healy, University of Canterbury, for steadfast technical support. 1880

dx.doi.org/10.1021/bm500261u | Biomacromolecules 2014, 15, 1871−1881

Biomacromolecules

Article

(33) Wood, Z. A.; Poole, L. B.; Hantgan, R. R.; Karplus, P. A. Biochemistry 2002, 41, 5493. (34) Parsonage, D.; Youngblood, D. S.; Sarma, G. N.; Wood, Z. A.; Karplus, P. A.; Poole, L. B. Biochemistry 2005, 44, 10583. (35) Nirudodhi, S.; Parsonage, D.; Karplus, P. A.; Poole, L. B.; Maier, C. S. Int. J. Mass. Spectrom. 2011, 302, 93. (36) Gerrard, J. A. Protein Nanotechnology: What Is It? In Protein Nanotechnology: Protocols, Instrumentation and Applications, 2nd ed.; Gerrard, J. A., Ed.; Methods in Molecular Biology Series 996, Humana Press: New York, 2013; Chapter 1, pp 1−18. (37) Konig, J.; Galliardt, H.; Jutte, P.; Schaper, S.; Dittmann, L.; Dietz, K.-J. J. Exp. Bot. 2013, 64, 3483. (38) Cox, A. G.; Winterbourn, C. C.; Hampton, M. B. Biochem. J. 2010, 425, 313. (39) Cao, Z.; Bhella, D.; Lindsay, J. G. J. Mol. Biol. 2007, 372, 1022. (40) Radjainia, M.; Huang, B.; Bai, B.; Schmitz, M.; Yang, S. H.; Griffin, M. D. W.; Brimble, M. A.; Wang, Y.; Mitra, A. K. FEBS J. 2012, 279, 2495. (41) Schröder, E.; Littlechild, J. A.; Lebedev, A. A.; Errington, N.; Vagin, A. A.; Isupov, M. N. Structure 2000, 8, 605. (42) Nelson, K. J.; Parsonage, D.; Hall, A.; Karplus, A.; Poole, L. B. Biochemistry 2008, 47, 12860. (43) Moon, J. C.; Hah, Y.-S.; Kim, W. Y.; Jung, B. G.; Jang, H. H.; Lee, J. R.; Kim, S. Y.; Lee, Y. M.; Jeong, M. G.; Kim, C. W.; Cho, M. J.; Lee, S. Y. J. Biol. Chem. 2005, 280, 28775. (44) Kristensen, P.; Rasmussen, D. E.; Kristensen, B. I. Biochem. Biophys. Res. Commun. 1999, 262, 127. (45) Cao, Z.; Tavender, T. J.; Roszak, A. W.; Cogdell, R. J.; Bulleid, N. J. J. Biol. Chem. 2011, 286, 42257. (46) Kitano, K.; Kita, A.; Hakoshima, T.; Niimura, Y.; Miki, K. Proteins 2005, 59, 644. (47) Papinutto, E.; Windle, H. J.; Cendron, L.; Battistutta, R.; Kelleher, D.; Zanotti, G. Biochim. Biophys. Acta 2005, 1753, 240. (48) Chauhan, R.; Mande, S. C. Biochem. J. 2001, 354, 209.

1881

dx.doi.org/10.1021/bm500261u | Biomacromolecules 2014, 15, 1871−1881