Dynamic Regulation of Histone Chaperone Nucleoplasmin

Small-angle X-ray scattering (SAXS) analysis of nucleoplasmin (Npm) Core+A2 truncation (1-145) bound to five H2A/H2B dimers. (Top) SAXs envelope of the pentameric complex (pink) with the best nuclear magnetic resonance (NMR)–restrained SAXS hybrid model inside. (Bottom) SAXS curve of the complex (purple dots). Simulated SAXS curve (black line) from the best-scoring structural model. [From Warren, C., et al. “Dynamic intramolecular regulation of the histone chaperone nucleoplasmin controls histone binding and release.” Nat. Commun. 8, 2215 (2017). DOI:10.1038/s41467-017-02308-3. Reused under a Creative Commons license (CC BY 4.0, https://creativecommons.org/licenses/by/4.0/​).]
Histones are eukaryotic cell nuclei proteins that package and order DNA into structural units called nucleosomes. Chromatin is the complex of DNA and proteins comprising the genome’s physiological form. As chromatin’s chief protein components, histones act as spools around which DNA winds, playing a role in gene regulation. A chaperone protein assists in the folding and unfolding of macromolecules, such as in the assembly of nucleosomes from folded histones and DNA. Nucleoplasmin (Npm) is a highly conserved embryonic histone chaperone, responsible for the maternal storage and zygotic release of histones H2A and H2B. Npm contains a pentameric N-terminal Core domain and an intrinsically disordered C-terminal Tail domain. Although intrinsically disordered regions are common among histone chaperones, their roles in histone binding and chaperoning have remained unclear.

This study, using the Xenopus laevis Npm Tail domain, unveils the architecture of the Npm histone complex and a mechanism of histone chaperone regulation. It demonstrates that intramolecular regulation of the histone chaperone Npm controls histone binding and release—a key process in the earliest stages of embryonic development. Structural analyses enabled model constructions of both the Npm Tail domain and the pentameric complex, revealing that the Tail domain controls the binding of histones through specific, electrostatic interactions. Functional analyses demonstrated that these competitive interactions negatively regulate Npm histone chaperone activity in vitro. Data from these studies establish a potentially generalizable mechanism of histone chaperone regulation via dynamic and specific intramolecular shielding of histone interaction sites.

Warren, C., et al. “Dynamic intramolecular regulation of the histone chaperone nucleoplasmin controls histone binding and release.” Nat. Commun. 8, 2215 (2017). DOI:10.1038/s41467-017-02308-3.

Instruments and Facilities Used: Bruker 600 nuclear magnetic resonance (NMR) and Inova 600 NMR instruments in the Albert Einstein College of Medicine (AECM) Einstein Structural NMR Resource; and bio–small-angle X-ray scattering (bio-SAXS) beamline 4-2, SLAC National Accelerator Laboratory’s Stanford Synchrotron Radiation Lightsource (SSRL).

Funding Acknowledgements: Supported by The American Cancer Society (ACS)Robbie Sue Mudd Kidney Cancer Research Scholar Grant (124891-RSG-13-396-01-DMC) and National Institutes of Health (NIH) grant R01GM108646 (both to D.S.) and training grants T32GM007491 and F31GM116536 (to C.W). J.M.K. supported by Einstein Medical Scientist Training Program Grant (T32 GM007288). Bruker 600 nuclear magnetic resonance (NMR) instrument purchased using funds from NIH award 1S10OD016305 and supported by Albert Einstein College of Medicine (AECM). Inova 600 NMR instrument in the Einstein Structural NMR Resource purchased using funds from NIH award 1S10RR017998 and National Science Foundation (NSF) award DBI0331934 and supported by the AECM. Use of Stanford Synchrotron Radiation Lightsource (SSRL), SLAC National Accelerator Laboratory (SLAC), supported by the Office of Basic Energy Sciences (OBES), U.S. Department of Energy (DOE) Office of Science, under Contract No. DE-AC02-76SF00515. SSRL Structural Molecular Biology Program supported by the Office of Biological and Environmental Research (OBER), DOE Office of Science, and by National Institutes of Health’s (NIH) National Institute of General Medical Sciences (NIGMS; including P41GM103393).

Structure of a Flavoenzyme Assembly Intermediate

(a) The structure of the FrdA-SdhE flavoenzyme assembly intermediate: flavoprotein subunit FrdA (cyan), assembly factor SdhE (green), flavin adenine dinucleotide FAD (orange sticks), and malonate (yellow sticks). The boxed region highlights the covalent interaction between the FAD and the enzyme. (b) Overlay of the flavin-binding domains of the FrdA subunit from the FrdA-SdhE intermediate (cyan) and the FrdA subunit from the mature assembled FrdABCD complex (gray). A rotation of 10.8° is observed in the capping domain of the assembly intermediate when compared to assembled FrdABCD. [From Sharma, P., et al. “Crystal structure of an assembly intermediate of respiratory Complex II,” Nat. Commun. 9, 274 (2018). DOI:10.1038/s41467-017-02713-8. Reused under a Creative Commons license (CC BY 4.0, https://creativecommons.org/licenses/by/4.0/​.)]
Enzymes frequently depend on an electron transport cofactor for executing catalytic functions such as reduction-oxidation (redox) reactions. For flavoenzymes, the cofactor is flavin adenine dinucleotide (FAD), whose binding type with the enzyme impacts the redox potential and thus reaction chemistry, such as for metabolism and detoxification. Researchers in this study discovered that the structure of an assembled flavoenzyme intermediate reveals the mechanism of covalent flavin binding in respiration. Assembly factors include SdhAF2 in humans, SdhE in Escherichia coli, and Sdh5 in yeast. Other revelations include that mitochondrial flavoenzymes drive both noncovalent and covalent redox reactions and that the assembly factor (SdhE, a small protein of ~90 to 140 amino acids, conserved in all kingdoms) in the structure of the SdhE:FrdA complex with covalent FAD stabilizes a conformation of the flavoprotein subunit FrdA that favors succinate oxidation.

Researchers fixed the E. coli FrdA-SdhE intermediate via site-specific crosslinking, resolving the structure to 2.6 angstroms (Å). This study identified that SdhE stabilizes an FrdA conformation that likely enables the mechanism of autocatalytic covalent flavinylation. FrdA’s FAD-binding domain and capping domain both interact with SdhE, but structural data revealed a 10.8° difference in their angles. The investigators believe that domain rotation affects flavinylation, showing that enzymes are tuned to catalyze reactions in different ways and that conformational diversity can directly relate to catalytic mechanism diversity.

Sharma, P., et al. “Crystal structure of an assembly intermediate of respiratory Complex II.” Nat. Commun. 9, 274 (2018). DOI:10.1038/s41467-017-02713-8.

Instruments and Facilities Used: Small angle X-ray scattering (SAXS) and diffraction and mass spectrometry analysis using beamline 9-2 at Stanford Synchrotron Radiation Lightsource (SSRL) at SLAC National Accelerator Laboratory (SLAC).

Funding Acknowledgements: Supported by Department of Veterans Affairs (DVA; BX001077 to G.C.) and National Institutes of Health (NIH; GM061606 to G.C. and T.M.I.). G.C. (recipient of a Senior Research Career Scientist award, #IK6B004215 from DVA). Vanderbilt University crystallization facility support: S10 RR026915. Use of Stanford Synchrotron Radiation Lightsource (SSRL), SLAC National Accelerator Laboratory (SLAC), supported by Office of Basic Energy Sciences (OBES), U.S. Department of Energy (DOE) Office of Science, under Contract No. DE-AC02-76SF00515. SSRL Structural Molecular Biology Program supported by Office of Biological and Environmental Research (OBER), DOE Office of Science, and by the NIH National Institute of General Medical Sciences (NIGMS; including P41GM103393).

The CRISPR Target-Recognition Mechanism

Cas1-Cas2 complex
Surface representation of the Cas1-Cas2 complex, consisting of four Cas1 proteins (light and dark green) and two Cas2 proteins (yellow). Donor DNA (brown) is being integrated into the target DNA (blue), at a precise location in the CRISPR array, following a short leader sequence (red). [From Wright, A. V., et al.  “Structures of the CRISPR Genome Integration Complex,” Science 357(6356), 1113–1118 (2017). [DOI:10.1126/science.aao0679. Reprinted with permission from AAAS*.]
* Readers may view, browse, and/or download material for temporary copying purposes only, provided these uses are for noncommercial personal purposes. Except as provided by law, this material may not be further reproduced, distributed, transmitted, modified, adapted, performed, displayed, published, or sold in whole or in part, without prior written permission from the publisher.
Bacterial DNA is characterized by regions of clustered regularly interspaced short palindromic repeats (CRISPRs) and associated Cas proteins (CRISPR-associated endonucleases). The CRISPR-Cas system has revolutionized gene editing by vastly simplifying the insertion of short snippets of new (“donor”) DNA into very specific locations of target DNA. Researchers in this study have discovered how Cas proteins recognize their target locations with such great specificity. They used x-ray crystallography to solve the structures of Cas1 and Cas2—responsible for DNA-snippet capture and integration—as the proteins were bound to synthesized DNA strands designed to mimic different stages of the process. The research also demonstrated how the system works in its native context as part of a bacterial immune system and how Cas proteins act as general-purpose molecular recording devices—tools for encoding information in genomes.

Cas1 appears to have evolved from a more “promiscuous” (less selective) type of enzyme that catalyzes the movement of DNA sequences from one position to another (a transposase). At some point, Cas1 acquired an unusual degree of specificity for a particular location in the bacterial genome, the CRISPR array. This specificity is critical to the bacteria, both for acquiring immunity and for avoiding genome damage caused by the insertion of viral fragments at the wrong location. The researchers wanted to learn how Cas1-Cas2 proteins recognize the target sequence to enable comparison with previously studied transposases and integrases (i.e., enzymes that catalyze the integration of donor DNA into target DNA) and to determine whether the proteins can be altered to recognize new sequences for custom applications.

The researchers crystallized Cas1-Cas2 in complex with preformed DNA strands that mimicked reaction intermediates and products. X-ray crystallography revealed that the structures showed substantial distortions in the target DNA, but there were surprisingly few sequence-specific contacts with the Cas1-Cas2 complex, and the DNA’s resulting flexibility produced disorder in the crystals. Attempts to model the DNA across the disordered sections showed that the DNA had to be even more distorted. Cryoelectron microscopy experiments, coupled with the crystallography data, confirmed that an accessor protein called the integration host factor (IHF) introduces an additional sharp bend in the DNA, bringing an upstream recognition sequence into contact with Cas1 to increase both the specificity and efficiency of integration. The architecture of the CRISPR integration complex suggests that subtle adjustment of the distance between Cas1 active sites could reprogram the system to recognize different target sites. Changes in its architecture could be exploited, thereby, for genome tagging applications and also may explain the natural divergence of CRISPR arrays in bacteria.

Wright, A. V., et al. “Structures of the CRISPR Genome Integration Complex,” Science 357(6356), 1113–1118 (2017). [DOI:10.1126/science.aao0679].

Instruments and Facilities Used: X-ray macromolecular crystallography; beamline 8.3.1; protein crystallography (PX); and scattering/diffraction at the Advanced Light Source at Lawrence Berkeley National Laboratory; Stanford Synchrotron Radiation Light Source 9-2 beamline.

Funding Acknowledgements: Advanced Light Source (ALS) 8.3.1 beamline, Lawrence Berkeley National Laboratory (LBNL), and Stanford Synchrotron Radiation Lightsource (SSRL) 9-2 beamline, SLAC National Accelerator Laboratory (SLAC), for assistance with data collection. ALS Beamline 8.3.1, is operated by University of California Office of the President, Multicampus Research Programs and Initiatives (grant MR-15-328599), and Program for Breakthrough Biomedical Research, partially funded by the Sandler Foundation. Use of SSRL supported by the Office of Basic Energy Sciences (OBES), U.S. Department of Energy (DOE) Office of Science, under contract no. DE-AC02-76SF00515. Electron microscopy (EM) data collected in Howard Hughes Medical Institute (HHMI) EM facility located at University of California, Berkeley. SSRL Structural Molecular Biology Program supported by DOE Office of Biological and Environmental Research (OBER) and the National Institutes of Health’s (NIH) National Institute of General Medical Sciences (NIGMS; including grant no. P41GM103393). Project funded by U.S. National Science Foundation (NSF) grant no. 1244557 (to J.A.D.) and NIGMS grant no. 1P50GM102706-01 (to J. H. Cate). A.V.W. and K.W.D. support: NSF Graduate Research Fellowship; G.J.K. funding: HHMI. J.A.D. and E.N.: HHMI investigators and members of the Center for RNA Systems Biology. Atomic coordinates and structure factors for the reported crystal structures deposited in the Protein Data Bank under accession codes 5VVJ (half-site–bound), 5VVK (pseudo–full-site–bound), and 5VVL (pseudo–full-site–bound with Ni2+). Cryo-EM structure and map deposited in the Protein Data Bank under accession code 5WFE and the Electron Microscopy Data Bank under accession code EMD-8827.

Probing S-layer Protein Structural Dynamics by SAXS

Biophysical Journal Cover Image
Calcium mediates the structural state of the Caulobacter crescentus surface layer protein, RsaA. Image featured on the cover of Biophysical Journal.

All archaea, and many bacteria, possess a protein shell referred to as a surface layer (S-layer), which usually consist of a single protein that self-assembles into a two-dimensional (2D) crystal lattice.  Studies have revealed the structural dynamics of this S-layer protein from the model bacterium Caulobacter crescentus, called RsaA. Using small angle scattering and diffraction (SAXS/D) techniques, multiple structural states of RsaA were successfully characterized including monomeric, aggregated, and crystalline states (see figure), with only monomeric Rssa forming 2D crystals. Enabling differentiation of the discrete states, these results rationalize physiological data implicating RsaA as a player in environmental adaptation of C. crescentus. The findings also provide a biochemical and physiological basis for RsaA’s calcium (Ca)-binding behavior, which extends far beyond Ca’s usual role in S-layer biology of aiding biogenesis or oligomerization, and demonstrate a connection to cellular fitness. Further characterization using slow and fast time-resolved SAXS/D methods is ongoing.

SAXS/D data
Small angle X-ray scattering and diffraction (SAXS/D) data of five solutions with different concentrations of the Caulobacter crescentus S-layer protein, RsaA, in the presence of calcium (Ca). Scattering profiles indicate concentration-dependent crystallization. Automatic indexing of the numbered peaks yielded a hexagonal crystal lattice consistent with predictions and denoted by Miller indices. (Top) The diffraction pattern obtained for the highest concentration used (8 mg/ml) shows powder rings. (Bottom) Transmission electron microscopy of the 8 mg/ml RsaA in the presence of 10 millimole per Liter (mm/L) of calcium chloride (CaCl2) (scale bar 200 nm). [Reprinted from Herrmann, J., et al. “Environmental Calcium Controls Alternate Physical States of the Caulobacter Surface Layer.” Biophys. J. 112(9), 1841–1851 (2017). DOI:10.1016/j.bpj.2017.04.003. Copyright 2017, with permission from Elsevier.]
Herrmann, J., et al. “Environmental Calcium Controls Alternate Physical States of the Caulobacter Surface Layer,” Biophys. J. 112(9), 1841–1851 (2017). [DOI:10.1016/j.bpj.2017.04.003].

Instruments and Facilities Used: Stanford ChEM-H Macromolecular Structure Knowledge Center,  Stanford Department of Structural Biology Electron Microscopy Center, Stanford Synchrotron Radiation Lightsource (SSRL) at SLAC National Accelerator Laboratory (SLAC). Beamlines or instruments used: transmission electron microscopy (TEM) and small angle X-ray scattering and diffraction (SAXS/D) at SSRL beamline 4-2 at SLAC.

Funding Acknowledgements: Part of work performed at Stanford University’s ChEM-H Macromolecular Structure Knowledge Center and Department of Structural Biology Electron Microscopy Center. Support: U.S. Department of Energy (DOE), SLAC National Accelerator Laboratory (SLAC) Laboratory Directed Research and Development (co-PI: John Bargar), under contract No. DE-AC02-76SF00515. Material based on work supported by the Office of Biological and Environmental Research (0BER) Mesoscale to Molecules: Bioimaging Science Program, DOE Office of Science. J.H. support: National Science Foundation (NSF) Graduate Research Fellowship Program (NSF-GRFP) and DOE Office of Science Graduate Student Research Program (DOE-SCGSR). J.S. support: grant from Natural Sciences and Engineering Research Council of Canada. L.S. support: National Institutes of Health’s (NIH) National Institute of General Medical Sciences (NIGMS; R35118072A). Use of Stanford Synchrotron Radiation Lightsource (SSRL), SLAC, support: Office of Basic Energy Sciences (OBES), DOE Office of Science, under contract No. DE-AC02-76SF00515. SSRL Structural Molecular Biology Program support: DOE OBER and NIH NGMIS (including grant No. P41GM103393).

Direct Measurement of Protein Dynamics In Vivo

quasi-elastic neutron scattering (QENS)
Labeling strategy for probing live cells using quasi-elastic neutron scattering (QENS). Graph: QENS spectra of uninduced cells (blue) and cells with expressed GroEL protein (red) in the deuterium oxide (D2O) buffer. [Reprinted with permission from Anunciado, D. B., et al. “In Vivo Protein Dynamics on the Nanometer Length Scale and Nanosecond Time Scale.” J. Phys. Chem. Lett. 8(8), 1899–1904 (2017). DOI: 10.1021/acs.jpclett.7b00399. Copyright 2017 American Chemical Society.]
Quasi-elastic neutron scattering was used to study the protein GroEL in vivo. Protiated GroEL was over-expressed in deuterated Escherichia coli cells by addition of protiated amino acids during induction of GroEL. The data showed retardation factors of ∼2 and ∼4 for the internal dynamics and global diffusion of the protein, compared to those of the protein in solution at the same concentration. Comparison with literature values suggests that the effective diffusivity of proteins depends on the length or time scale probed. Selective isotope labeling of biomolecules in cells opens up new lines of biological research using “in-cell neutron scattering” to extract information on dynamics of biomolecular systems that is unobtainable by other analysis techniques.

Anunciado, D. B., et al. “In Vivo Protein Dynamics on the Nanometer Length Scale and Nanosecond Time Scale.” J. Phys. Chem. Lett. 8(8), 1899–1904 (2017). [DOI:10.1021/acs.jpclett.7b00399].

Instruments and Facilities Used: Quasi-elastic neutron scattering, BASIS neutron backscattering spectrometer at Spallation Neutron Source at Oak Ridge National Laboratory.

Funding Acknowledgements: Qiu Zhang: technical assistance. Manuscript authored by UT-Battelle, LLC, under Contract No. DE-AC05-00OR22725 with the U.S. Department of Energy (DOE Neutron scattering experiments at Oak Ridge National Laboratory’s (ORNL) Spallation Neutron Source (SNS) support: Office of Basic Energy Sciences (OBES) Scientific User Facilities Division, DOE Office of Science. Additional funding: Adaptive Biosystems Imaging project (ERKP851) and Center for Structural Molecular Biology (Project ERKP291) supported by DOE Office of Biological and Environmental Research (OBER).

Hollow Pyramid Unlocks Principles of Protein Architecture

symmetric crystal structure
Designed and observed cage symmetry. (A) Left: Schematic diagram of the symmetry principles used to design the 12-subunit tetrahedral cage by fusing two oligomeric domains (green and orange) by a semirigid linker (magenta). Right: Single point mutation distinguishing our PCtrip construct from PCquad replaces tyrosine (black sticks) with alanine in the trimeric domain that makes contact with the linker. (B) Side-by-side view of the theoretically designed (perfectly symmetric) model of the protein cage (left) and the most symmetric crystal structure obtained in this work for the PCquad variant (right). (C) Walleyed stereo view of three crystal structures of PCquad (yellow, magenta, and blue) overlaid on the ideal model (green ribbon), showing the agreement of the observed structures and the design. [[From Lai, Y.-T., et al., “Designing and Defining Dynamic Protein Cage Nanoassemblies in Solution.” Sci. Adv. 2(12), e1501855 (2016). DOI:10.1126/sciadv.1501855. Reused under a Creative Commons license (CC BY NC-4.0, https://creativecommons.org/licenses/by-nc/4.0/.]
Large biological macromolecules (up to 10,000 atoms) can catalyze chemical reactions in ways that are difficult to replicate inorganically. Their size allows for the complexity needed to perform such functions, but it also makes them more susceptible to misfolding and aggregating—thus, the importance of understanding the fundamental architectural principles that cause large proteins to favor specific conformations. At the nanoscale, where organic chemical groups interact with solvent water molecules, these principles are very different from the ones used to build houses and cars. To investigate these principles, a group of researchers designed a protein that would self-assemble into a hollow pyramid (or tetrahedron). Upon crystallizing the macromolecule, the group found that they had indeed been successful in creating the assembly, but it was unexpectedly warped and collapsed in an asymmetric manner, with some edges bent inward—an asymmetric tetrahedron. To ensure that this was not an artifact of crystallization, they investigated the protein’s behavior in solution using small-angle x-ray scattering (SAXS), which showed that the collapse could be controlled by adjusting the solution’s salt concentration; the structure was disassembled by varying the pH.

The flexibility of this macromolecule suggests that it could be useful for the controlled capture and release of smaller compounds. Overall, the researchers expect that, with the tools and techniques developed here, the combination of SAXS with crystallography or electron microscopy could be increasingly useful in analyzing and optimizing designed protein assemblies and understanding their behavior in solution.

Lai, Y.-T., et al. “Designing and Defining Dynamic Protein Cage Nanoassemblies in Solution.” Sci. Adv. 2(12), e1501855 (2016). [DOI:10.1126/sciadv.1501855].

Instruments and Facilities Used: Small-angle X-ray scattering (SAXS) at Advanced Light Source (ALS) with SIBYLS Beamline 12.3.1 (a joint crystallography and SAXS beamline) at Lawrence Berkeley National Laboratory.

Funding Acknowledgements: Berkeley Center for Structural Biology (BCSB), Advanced Light Source (ALS), Lawrence Berkeley National Laboratory (LBNL), work performed as collaboration between the Joint BioEnergy Institute (JBEI; https://www.jbei.org) and Great Lakes Bioenergy Research Center (GLBRC; https://www.glbrc.org). JBEI support: Office of Biological and Environmental Research (OBER), U.S. Department of Energy (DOE) Office of Science, through contract DE-AC02-05CH11231 between LBNL and DOE. GLBRC support: OBER, DOE Office of Science, through Grant DE-FG02-07ER64495. BCSB support in part: National Institutes of Health’s (NIH) National Institute of General Medical Sciences (NIGMS). ALS support: Director, Office of Basic Energy Sciences (OBES), DOE Office of Science, under Contract DE-AC02-05CH11231. Support for part of work: National Science Foundation (NSF) under Cooperative Agreement 1355438.

Designing Cyclic Oligomers: Greater than the Sum of Their Parts

Computational oligomer design.
Computational oligomer design. Scientists used small angle solution studies and X-ray scattering to prove that synthetic design matched the computational design. [Reprinted by permission from Springer Nature: Fallas, J. A., et al. “Computational Design of Self-Assembling Cyclic Protein Homo-Oligomers.” Nat. Chem. 9, 353–360 (2017). DOI:10.1038/nchem.2673. Copyright 2017.]
Cyclic proteins that assemble from multiple identical subunits (homo-oligomers) play key roles in many biological processes, including enzymatic catalysis and function and cell signaling. Researchers in the Molecular Biophysics and Integrated Bioimaging (MBIB) Division worked with University of Washington’s David Baker, who led a team to design in silico and crystallize self-assembling cyclic homo-oligomer proteins.

A strategy was developed that designs interfaces onto idealized proteins aimed to direct their assembly into multimeric complexes. Researchers used structural characterization, both X-ray crystallography and small angle X-ray scattering (SAXS), to show that many of the designs adopted the target oligomerization state and predicted structure. Not only does the work demonstrate that scientists have a basic understanding of what determines oligomerization, but that they could design proteins with tunable shape, size, and symmetry for a variety of biological applications.

Fallas, J. A., et al.Computational Design of Self-Assembling Cyclic Protein Homo-Oligomers.” Nat. Chem. 9, 353–360 (2017). [DOI:10.1038/nchem.2673].

Instruments and Facilities Used: Crystallography Collective program; beamline 5.0.2; small angle X-ray scattering (SAXS) on the SIBYLS BL12.3.1 beamline. Advanced Photon Source synchrotron beamline 24-ID-C at Berkeley Center for Structural Biology and Advanced Light Source at Lawrence Berkeley National Laboratory (LBNL).

Funding Acknowledgements: Support: Howard Hughes Medical Institute (HHMI), Air Force Office for Scientific Research (AFOSR FA950-12-10112), National Science Foundation (NSF MCB-1445201 and CHE-1332907), Bill and Melinda Gates Foundation (OPP1120319), and U.S. Department of Defense (DoD) Defense Threat Reduction Agency (HDTRA1-11-C-0026 AM06). R. Koga and L. Carter: assistance with size exclusion chromatography with multi-angle light scattering SEC-MALS. M. Collazo and M. Sawaya support: U.S. Department of Energy (DOE; Grant DE-FC02-02ER63421). M. Capel, K. Rajashankar, N. Sukumar, J. Schuermann, I. Kourinov, and F. Murphy at Northeastern Collaborative Access Team support: grants from National Center for Research Resources (5P41RR015301-10) and National Institutes of Health’s (NIH) National Institute of General Medical Sciences (NIGMS; P41GM103403-10). Use of Advanced Photon Source (APS), Argonne National Laboratory (ANL), support: Office of Biological and Environmental Research (OBER), DOE Office of Science, under Contract DE-AC02-06CH11357. X-ray crystallography and SAXS data collected at the Advanced Light Source (ALS, LBNL, Berkeley, CA, DOE Office of Science contract no. DE-AC02-05CH11231); SAXS data collected through the SIBYLS mail-in SAXS program under aforementioned contract no. and funded by DOE OBER Integrated Diffraction Analysis Technologies (IDAT), NIH Minocycline to Improve Neurologic Outcome in Stroke (MINOS; RO1GM105404), and ALS (K. Burnett and G. Hura). Berkeley Center for Structural Biology (BCSB) support in part: NIH NIGMS and HHMI. ALS support: Office of Basic Energy Sciences (OBES), Director, DOE Office of Science, under contract no. DE-AC02-05CH11231.

Cellulose Synthesis Complex

The plant cellulose synthesis complex is a large multi-subunit transmembrane protein complex responsible for synthesis of cellulose chains and their assembly into microfibrils. The image shows ab initio structures of CESA trimers calculated from small-angle scattering data represented by semi-transparent grey surface envelopes, superposed with the computational atomic models in orange. Image credits: Thomas Splettstoesser, scistyle.com, Berlin Germany

Vandavasi, V.G., D.K. Putnam, Q. Zhang, L. Petridis, W.T. Heller, B.T. Nixon, C.H. Haigler, U. Kalluri, L. Coates, and P. Langan. 2016. “A structural study of CESA1 catalytic domain of Arabidopsis cellulose synthesis complex: evidence for CESA trimers.” Plant physiology  170(1):123-35.  [DOI: 10.1104/pp.15.01356]

Funding Acknowledgements: Molecular biology and structural characterization: H.O., V.G.V., Q.Z., W.T.H., L.P., U.K., J.C.S., P.L., and L.C., supported by the Laboratory Directed Research and Development Program of Oak Ridge National Laboratory (ORNL), managed by UT-Battelle, LLC, for the U.S. Department of Energy (DOE) under Contract Number DE-AC05-00OR22725. Small-angle scattering (SAS) and computational analysis, performed by H.O., V.G.V., L.P., B.T.N., and C.H.H., supported by Center for Lignocellulose Structure and Formation, an Energy Frontier Research Center (EFRC) funded by the Office of Basic Energy Sciences (OBES). DOE Office of Science. J.M. and D.K.P. support: High Performance Computing Grant from Oak Ridge Associated Universities (ORAU). Bio-SANS is operated by the Center for Structural Molecular Biology at ORNL, supported by Office of Biological and Environmental Research (OBER), DOE Office of Science, Project ERKP291. EQ-SANS at Spallation Neutron Source (SNS) and High Flux Isotope Reactor (HFIR) sponsored by OBES Scientific User Facilities Division, DOE Office of Science, at ORNL. Sai Venkatesh Pingali: assistance with the operation of Bio-SANS and data reduction. Paul Abraham and the BioEnergy Science Center (BESC) proteomics facilities: validation of purified proteins. BESC supported by DOE OBER. Mass spectrometry (MS) analysis carried out by DOE OBERsupported Bioenergy Research Center proteomics pipeline. Use of the National Synchrotron Light Source (NSLS), Brookhaven National Laboratory (BNL), supported by OBES, DOE Office of Science, under Contract Number DE-AC02-98CH10886.

Protein Cage

Using Small-angle X-ray scattering (SAXS) to efficiently and quickly image large protein molecular assemblies, scientists have designed a hollow, cube-shaped protein cage that has the potential for delivering proteins or other chemicals to specific locations for medical, energy, and other applications. The nanocage crystal structure was optimized at the Advanced Light Source (Image courtesy of Greg Hura, LBNL).

Lai, Y.T., E. Reading, G.L. Hura, K.L.Tsai, A. Laganowsky, F.J. Asturias,  J.A. Tainer, C.V. Robinson, and T.O. Yeates. 2014.“Structure of a Designed Protein Cage that Self-Assembles into a Highly Porous Cube,” Nature Chemistry 6, 1065–1071. doi:10.1038/nchem.2107

Feature Article at LBNL

Funding Acknowledgements: Work supported by National Science Foundation (NSF; grant CHE-1332907, T.O.Y.), Office of Biological and Environmental Research (OBER), U.S. Department of Energy (DOE) Office of Science, and the National Institutes of Health (NIH; grant R01GM067167, F.J.A.). SAXS data collection and analysis at BL12.3.1 at the Advanced Light Source (ALS), Lawrence Berkeley National Laboratory (LBNL) supported by the Integrated Diffraction Analysis Technologies (IDAT) program (DOE OBER), by DOE (contract DE-AC02-05CH11231) and by NIH Minocycline to Improve Neurologic Outcome in Stroke (MINOS; R01GM105404).

Spatial Organization of Lipid Domains in a Biomimetic Four-Lipid System

Floating freely in cell membranes, lipid rafts are organizing centers for membrane-mediated processes. Neutron scattering techniques were used to characterize lipid domain size transitions from nanometers to micrometers in a four-component biomimetic lipid mixture.  Results suggest that reversible changes in lipid composition may regulate the size of functional domains in lipid rafts. The BioSANS instrument at the ORNL Center for Structural Molecular Biology and the EQ-SANS instrument (BER) at the ORNL Spallation Neutron Source (BES) were used in this study.

Heberle F.A., Petruzielo R.S., Pan J., Drazba P., Kucerka N., Standaert R.F., Feigenson G.W., Katsaras J., “Bilayer thickness mismatch controls domain size in model membranes”, Journal of the American Chemical Society, 135, 18, 6853-6859 (2013). DOI: 10.1021/ja3113615

Funding Acknowledgements: Support: Laboratory Directed Research and Development Program of Oak Ridge National Laboratory (ORNL; to J.K. and R.F.S.), managed by UT-Battelle, LLC, for the U.S. Department of Energy (DOE), and from National Science Foundation (NSF) research award MCB 0842839 (to G.W.F.). Additional support: Office of Biological and Environmental Research (OBER), DOE Office of Science, for BioSANS instrument at ORNL Center for Structural Molecular Biology (CSMB), and from the DOE Office of Basic Energy Sciences (OBES) Scientific User Facilities (SUF) Division, for the EQ-SANS instrument at the ORNL Spallation Neutron Source (SNS). These facilities are managed for DOE by UT-Battelle, LLC, under Contract No. DE-AC05-00OR2275. A portion of this research conducted using resources of Cornell Center for Advanced Computing, which receives funding from Cornell University; NSF; and other leading public agencies, foundations, and corporations.