Cyanobacterial Studies Examine Cellular Structure During Nitrogen Starvation

Researchers from Washington University in St. Louis and Oak Ridge National Laboratory (ORNL) are using neutrons to study what happens when cyanobacteria cell samples (pictured) are starved for nitrogen. They are especially interested in how this process affects phycobilisomes, large antenna protein complexes in the cells that harvest light for photosynthesis. A better understanding of this natural phenomenon could lead to improvements in artificial resources like solar panels. [Courtesy ORNL]
Using nondestructive neutron scattering techniques, scientists are examining how single-celled organisms called cyanobacteria produce oxygen and obtain energy through photosynthesis. Collaborators are conducting a series of experiments to study the behavior of phycobilisomes—large antenna protein complexes in cyanobacteria cells. Phycobilisomes harvest light to initiate photosynthesis, and a better understanding of this process could help researchers design more efficient solar panels and other artificial structures that mimic natural systems. Neutrons can analyze these delicate structures without damaging or killing the cyanobacteria and with more spatial accuracy than other techniques like microscopy. Bio-SANS allows observing what’s happening at the nanoscale level in real time in a living cell.

Phycobilisomes attach to cellular membranes where the light-dependent reactions of photosynthesis take place. Changing the antenna complexes of the phycobilisomes can have dramatic and far-reaching consequences in cyanobacteria. Artificially modifying phycobilisomes by deleting certain genes in the cells caused structural defects in the cellular membranes and other cell physiology, allowing scientists to observe the resulting structural changes.

Starving the cyanobacteria for nitrogen naturally modifies the antenna complexes, causing the antenna to decrease in size and leading to significant cellular membrane modifications, because the cells break down the phycobilisomes and use them as an alternative nitrogen source to survive. By determining the extent of these changes, the team hopes to better understand the structure-function relationship between cellular organization and natural modification. These processes can be immediately reversed by restoring nitrogen to the cells. The researchers plan to compare these results to those recorded from their genetic studies to explore the differences between artificial and natural modifications and their effects on the intracellular makeup of cyanobacteria.

Instruments and Facilities Used: Photosynthetic Antenna Research Center (PARC), a DOE BES-funded Energy Frontier Research Center based at Washington University at St Louis. Small angle neutron scattering (SANS) was performed at the DOE-BER supported Bio-SANS instrument, beamline CG‑3, at Oak Ridge National Laboratory’s High Flux Isotope Reactor.

Precise Control of Neutron Contrast in Surfactant Micelles Provides Platform for Membrane Structure Studies

Detergent Micelles
The scattering collected for detergent at its solution match point, where contrast still persists between core and shell, produces a non-flat scattering profile (red). Incorporating the non-ionic detergent DDM with deuterium-labelled chains allows matching of the core and shell contrast, producing the flat scattering profile shown in blue. [Reprinted with permission from Oliver, R. C., S. V. Pingali, and V. S. Urban. “Designing Mixed Detergent Micelles for Uniform Neutron Contrast.” J. Phys. Chem. Lett. 8, 5041–5046 (2017). [DOI: 10.1021/acs.jpclett.7b02149]. Copyright 2017 American Chemical Society.]
Scientists in this study have successfully demonstrated the ability to manipulate the neutron contrast of detergent micelles by incorporating a similar detergent with deuterium-labelled alkyl chains. The presence of excess detergent micelle scattering often has a detrimental influence on scattering data obtained for membrane protein–detergent complexes. Isolation of the scattering signal from the protein of interest can be accomplished by eliminating all scattering from the detergent. Using this approach enabled determination of the overall structure and oligomeric state of a small membrane protein enzyme.

Oliver, R. C., et al. “Designing Mixed Detergent Micelles for Uniform Neutron Contrast.” The Journal of Physical Chemistry Letters 8(20), 5041–5046 (2017). [DOI:10.1021/acs.jpclett.7b02149].

Instruments and Facilities Used: Small angle neutron scattering (SANS): Bio-SANS beamline (CG3) of the High-Flux Isotope Reactor at Oak Ridge National Laboratory (ORNL). Recorded scattering data using MantidPlot software. Neutron contrast studies: ModULes for the Analysis of Contrast (MULCh) Variation Data at University of Sydney (smb-research.smb.usyd.edu.au/NCVWeb/).

Funding Acknowledgements: Neutron scattering studies at CG-3 Bio-SANS instrument at the High-Flux Isotope Reactor (HFIR), Oak Ridge National Laboratory (ORNL), sponsored by the Office of Biological and Environmental Research (OBER) and Office of Basic Energy Sciences (OBES) Scientific User Facilities Division (SUF), U.S. Department of Energy (DOE) Office of Science. Work benefited from use of the SasView application, originally developed under National Science Foundation (NSF) Award DMR-0520547. SasView contains code developed with funding from the European Union’s Horizon 2020 Research and Innovation Programme under the SINE2020 project, Grant Agreement No 654000. Manuscript authored by UT-Battelle, LLC, under Contract No. DE-AC05-00OR22725 with DOE. DOE will provide public access to these results of federally sponsored research in accordance with the DOE Public Access Plan (http://energy.gov/downloads/doe-public-access-plan).

Dynamics on Cellulose Show Two Important Populations from Neutron Scattering and Simulations

Elastic Intensity Scans
Elastic intensity scans of dry and hydrated cellulose. Dashed lines denote inflection points in the curves at 220 and 260 K, the temperatures at which the surface water (nonfreezing) and interfibrillar water (freezing) become mobile in the hydrated cellulose sample, respectively. Inset illustration depicts water populations associated with cellulose. [Adapted from O’Neill, H., et al. “Dynamics of Water Bound to Crystalline Cellulose.” Sci. Rep. 7, 11840 (2017). [DOI:10.1038/s41598-017-12035-w]. Reused under a Creative Commons license (CC BY 4.0, https://creativecommons.org/licenses/by/4.0/). Curve colors were modified, and additional labels and inset added.]
Biomass pretreatment is necessary to make cellulose accessible to hydrolysis for conversion to biofuels. Understanding water’s role in cellulose reactivity will aid discovery of the underlying processes that change biomass morphology and reactivity during different pretreatment regimes for biofuels production. In this study, cellulose-water interactions were examined using neutron scattering supported by molecular dynamics simulation. The data show two distinct populations of water molecules—one tightly “bound” to the surface and the other interfibrillar and translationally mobile. Accurate models of hydration water in the cell wall can address fundamental questions about cellulose-water interactions. The mobility of the interfibrillar water is also important to enzyme and chemical attack and is distinct from the bound and bulk water.

O’Neill, H. M., et al. “Dynamics of Water Bound to Crystalline Cellulose.” Sci. Rep. 7, Article 11840 (2017). [DOI:10.1038/s41598-017-12035-w].

Instruments and Facilities Used: Deuterium labeling, neutron scattering, and molecular dynamics simulation; quasi-elastic neutron scattering (QENS) using BASIS, the Backscattering Spectrometer, at Oak Ridge National Laboratory (ORNL) Spallation Neutron Source (SNS). Structural characterization using small-angle neutron scattering (SANS) CG-3 Bio-SANS instrument at High Flux Isotope Reactor (HFIR) facility of ORNL and X-ray diffraction (XRD) analysis combined with SANS. Wide-Angle X-ray Diffraction (WAXD) using theta-theta goniometer Bruker D5005 instrument; quasi-elastic neutron scattering (QENS) using BASIS, the Backscattering Spectrometer at ORNL SNS; Molecular Dynamics (MD) Simulations (GROMACS software and the CHARMM C36 carbohydrate force field); and TIP4P water model.

Funding Acknowledgements: Deuterium labeling, neutron scattering, and molecular dynamics simulation; quasi-elastic neutron scattering (QENS) using BASIS, the Backscattering Spectrometer, at Oak Ridge National Laboratory (ORNL) Spallation Neutron Source (SNS). Structural characterization using small-angle neutron scattering (SANS) CG-3 Bio-SANS instrument at High Flux Isotope Reactor (HFIR) facility, ORNL, and x-ray diffraction (XRD) analysis combined with SANS. Wide-angle x-ray diffraction (WAXD) using theta-theta goniometer Bruker D5005 instrument; QENS; molecular dynamics (MD) simulations (GROMACS software and CHARMM C36 carbohydrate force field); and TIP4P water model. H.O’N., J.H., B.E., J.C.S., P.L. and B.H.D. support: U.S. Department of Energy (DOE) Genomic Science Program, Office of Biological and Environmental Research (OBER), DOE Office of Science, under Contract FWP ERKP752, for sample preparation and QENS studies. SANS studies on Bio-SANS by S.V.P. and V.U. support: OBER-funded Center for Structural Molecular Biology (CSMB) under Contract FWP ERKP291, using facilities supported by the Office of Basic Energy Sciences (OBES), DOE Office of Science. Molecular dynamics (MD) simulations performed by L.P. supported by the Center for Lignocellulose Structure and Formation, an Energy Frontier Research Center, funded by DOE OBES, under Award DE-SC0001090. Research used resources of the National Energy Research Scientific Computing Center (NERSC), a DOE Office of Science User Facility supported under Contract No. DE-AC02-05CH11231. E.M. support: Oak Ridge National Laboratory’s (ORNL) Spallation Neutron Source (SNS), funded by the DOE OBES Scientific User Facilities Division. Manuscript authored by UT-Battelle, LLC, under Contract No. DE-AC05-00OR22725 with DOE.

Measuring and Modeling Poplar Root Water Extraction After Drought Using Neutron Imaging

 poplar seedling
Composite images of 16 radiographs of 11-week-old poplar seedling in sand. Intensity indicates water content. [Reprinted with permission of Springer from Dhiman, I., et al. “Quantifying Root Water Extraction After Drought Recovery Using Sub-mm In Situ Empirical Data.” Plant Soil 47, 1–17 (2017). [DOI:10.1007/s11104-017-3408-5]. © U.S. Government (outside the USA) 2017.]
Neutron radiography was used to measure soil water movement and water uptake by individual roots in situ. Root water uptake was linked to root traits; smaller-diameter roots had greater water uptake per unit surface area than larger-diameter roots. Model analysis based on root-free soil hydraulic properties indicated unreasonably large water fluxes among the vertical soil layers during the first 16 hours after wetting. These results suggest problems with common soil hydraulic or root surface area modeling approaches, indicating the need to further investigate and understand the impacts of roots on soil hydraulic properties. This work highlights the team’s ability to link root water uptake to characteristic root traits, thus enabling performance assessment of common water uptake models

Dhiman, I., et al. “Quantifying Root Water Extraction After Drought Recovery Using sub-mm In Situ Empirical Data.” Plant Soil 417, 1–17 (2017). [DOI:10.1007/s11104-017-3408-5].

Instruments and Facilities Used: Sequential neutron radiography using CG1-D beam line using cold neutrons, at High Flux Isotope Reactor (HFIR) at Oak Ridge National Laboratory. Neutron attenuation by plant samples was detected with a 25-μm lithium fluoride / zinc sulfide (LiF/ ZnS) scintillator linked to a charge coupled detector (CCD) camera system (iKon – L 936, Andor Technology plc., Belfast, U.K.). Roots scanned and dimensions measured using WinRhizo software (Regent Instruments Inc., Quebec, Canada.

Funding Acknowledgements: Research sponsored by Laboratory Directed Research and Development Program of Oak Ridge National Laboratory (ORNL), managed by UT-Battelle, LLC, for the Office of Biological and Environmental Research (OBER), U.S. Department of Energy (DOE) Office of Science , and by Office of Workforce Development for Teachers and Scientists, DOE Office of Science Graduate Student Research (SCGSR) program. SCGSR program administered by Oak Ridge Institute for Science and Education (ORISE) for DOE. ORISE is managed by Oak Ridge Associated Universities under contract number DE-AC05-06OR23100. ORNL is managed by UT-Battelle, LLC, for DOE under contract DE-AC05-1008 00OR22725. Research used resources at the High Flux Isotope Reactor (HFIR), a DOE Office of Science User Facility operated by ORNL. Manuscript authored by UT-Battelle, LLC, under Contract No. DE-AC05-00OR22725 with DOE.

Brown Rot Fungi Reveal a New Approach for Biomass Conversion to Fuels and Chemicals

Brown rot fungi

(A) Brown rot fungi mushrooms, (B) SANS profiles, (C) SFG spectra of brown rot fungi–mediated cellulose deconstruction, and (D) AFM images of repolymerized lignin in brown rot cell walls. [(A) Wikimedia Commons, Zinnmann; (B) Authors; (C) and (D) From Goodell, B., et al. “Modification of the Nanostructure of Lignocellulose Cell walls via a Non-Enzymatic Lignocellulose Deconstruction System in Brown Rot Wood-Decay Fungi.” Biotechnol. Biofuels 10, 179 (2017). [DOI 10.1186/s13068-017-0865-2]. Reused under a Creative Commons license (CC BY 4.0, https://creativecommons.org/licenses/by/4.0/).]
A multimodal approach used in this study examined wood decay by the brown rot fungi, Gloeophyllum trabeum or Rhodonia placenta, that degrade wood using a chelator-mediated Fenton (CMF) reaction. Small-angle neutron scattering (SANS) showed changes in microfibril bundling and lignin structure during biomass breakdown. Complementary approaches, sum frequency generation (SFG) spectroscopy, X-ray diffraction, atomic force microscopy (AFM), and transmission electron microscopy (TEM) also contributed information on nanoscale structural changes in wood over time. Woods studied were southern yellow pine (Pinus spp.) and birch (Betula verrucosa Ehr.). The data support a degradation mechanism in which sugars released by non-enzymatic action diffuse from the cell wall,  facilitated by increasing the porosity of the cell walls. This is a paradigm shift in understanding the mechanism of brown rot fungal degradation.

Goodell, B., et al. “Modification of the Nanostructure of Lignocellulose Cell Walls via a Non-Enzymatic Lignocellulose Deconstruction System in Brown Rot Wood-Decay Fungi.” Biotechnol. Biofuels 10(1), 179 (2017). [DOI:10.1186/s13068-017-0865-2].

Instruments and Facilities Used: Small angle neutron scattering (SANS) – Bio-SANS at High Flux Isotope Reactor (HFIR) at Oak Ridge National Laboratory (ORNL). Sum frequency generation (SFG) spectroscopy, broadband SFG system. Chelator-mediated Fenton treatments (CMF) and cellulase treatment. X-ray diffraction analysis (XRD) – PANalytical Empyrean diffractometer, The Netherlands, equipped with a Cu X-ray source. Attenuated total reflectance Fourier transform infrared analysis (ATR-FTIR): Nicolet 8700 FTIR Spectrometer (Thermo Scientific) equipped with a smart iTR diamond ATR unit, a KBr beam splitter, and a deuterated triglycine sulfate (DTGS) detector. Atomic force microscopy (AFM) of brown-rotted wood surfaces. Nanoscope IIIa AFM-Digital Instruments, Santa Barbara, California with three 5 µm scans. Transmission electron microscopy (TEM) – Philips CM12 TEM instrument (Philips, Eindhoven, The Netherlands); images recorded on Kodak 4489 negative film and the films subsequently scanned using an Epson Perfection Pro 750 film scanner.

Funding Acknowledgements: Dr. Zhu support: Chinese Forestry Industry Research Special Funds for Public Welfare Projects (#201204702-B2). Eastwood support: UK Natural Environment Research Council, award #NE/K011588/1. Daniel support: Formas Grant 2015-469. Support from ORNL-Proposal IPTS-12345/CG-3. Sum frequency generation (SFG) spectroscopy, x-ray diffraction (XRD), and infrared (IR) studies support: Center for Lignocellulose Structure and Formation, an Energy Frontier Research Center, funded by the Office of Basic Energy Sciences (OBES), U.S. Department of Energy (DOE) Office of Science, under Award Number DE-SC0001090. Pingali and O’Neill support: Biofuels SFA funded by the DOE Genomic Science Program, Office of Biological and Environmental Research (OBER), under Contract FWP ERKP752. Bio-SANS support: Center for Structural Molecular Biology, supported by DOE OBER under Contract FWP ERKP291. Neutron scattering facilities at Oak Ridge National Laboratory (ORNL) support: Scientific User Facilities Division, DOE OBES. Research also supported by U.S. Department of Agriculture’s (USDA) National Institute of Food and Agriculture (NIFA) and University of Massachusetts Amherst’s Center for Agriculture, Food and the Environment and the Microbiology Department: project # MAS00511.

Using Neutrons to Resolve Plasma Membrane Organization in Living Bacterial Cells

Bacillus subtilis cell wall

(Left) The cell wall (greenish brown, at top) of Bacillus subtilis is shown, along with the bacterium’s plasma membrane (blue and red) and a portion of cytoplasm (rust, at bottom). (Right) SANS used in conjunction with selective hydrogen/deuterium labeling techniques revealed the structure of the plasma membrane, including nanoscale lipid domains, while blocking interfering signals from other cellular features. [From Nickels, J. D., et al. “The In Vivo Structure of Biological Membranes and Evidence for Lipid Domains.” PLOS Biology 15(5), e2002214 (2017). [DOI:10.1371/journal.pbio.2002214]. Reused under a Creative Commons license (CC BY 4.0, https://creativecommons.org/licenses/by/4.0/).]
A new strategy devised by scientists at Oak Ridge National Laboratory used nondestructive small-angle neutron scattering (SANS) to reveal, for the first time, nanoscale membrane structures in living cells.

Neutron scattering spectra confirmed that the plasma membrane of the bacterium Bacillus subtilis is lamellar with an average hydrophobic thickness of 24 Ångstroms. The data also revealed that the membrane contains lipid features of approximately 40 nm or less in size, consistent with hypothesized “lipid rafts” in biological systems. The observation of lipid segregation in the plasma membrane of a bacterium is consistent with the notion of nanoscopic lipid assemblies, often described as lipid rafts in mammalian systems, implying that lipid domains are integral features of all biological membranes. Among their functions, lipid rafts are thought to play a vital role in cell signaling and facilitate movement of essential biomolecules in and out of the cell.

In addition to these scientific findings, the methods developed provide a new experimental platform for pursuing additional areas of inquiry (e.g., systematic in vivo investigations of cell membrane structure and response to diverse environmental stimuli). This new approach also may prove valuable, for example, in biomass feedstock and biofuel production, where bacterial cell membranes play important roles, and in biomedicine, where bacterial membrane domains affect antibiotic resistance. Furthermore, the strategy for “visualizing” the membrane can be used with other physical characterization techniques to examine additional cell structures such as the cell wall.

Nickels, J. D., et al. “The In Vivo Structure of Biological Membranes and Evidence for Lipid Domains.” PLOS Biology 15(5): e2002214 (2017). [DOI:10.1371/journal.pbio.2002214]

Instruments and Facilities Used: Small-angle neutron scattering (SANS) at the High Flux Isotope Reactor (HFIR) at Oak Ridge National Laboratory (ORNL). EQ-SANS at Spallation Neutron Source at ORNL. Oak Ridge Leadership Computing Facility at ORNL.

Related ORNL News Feature: Neutrons provide the first nanoscale look at a living cell membrane

DOE Highlight: First Look at a Living Cell Membrane

The work described in this highlight builds on previous work including:

  • Heberle, F.A., et al. “Bilayer Thickness Mismatch Controls Domain Size in Model Membranes.” Journal of the American Chemical Society 135, 6853-6859 (2013). [DOI: 10.1021/ja3113615]
  • Nickels, J.D. et al. “Mechanical Properties of Nanoscopic Lipid Domains.” Journal of the American Chemical Society 137, 15772-15780 (2015). [DOI: 10.1021/jacs.5b08894]
  • Nickels, J.D. et al. “Lateral organization, bilayer asymmetry, and inter-leaflet coupling of biological membranes.” 192, 87-99 (2015). [DOI:10.1371/journal.pbio.2002214]

Funding Acknowledgements: Work sponsored by Laboratory Directed Research and Development Program (grant number 6988) of Oak Ridge National Laboratory (ORNL), managed by UT-Battelle, LLC, for the U.S. Department of Energy (DOE) under Contract No. DE-AC05-00OR22725. Support for J.K.: Office of Basic Energy Sciences (OBES) Scientific User Facilities Division, DOE Office of Science. R.F.S.: Office of Biological and Environmental Research (OBER), DOE Office of Science (grant number ERKP-851). Resources of Oak Ridge Leadership Computing Facility at ORNL, supported by Facilities Division of DOE Office of Advanced Scientific Computing Research (OASCR). Small-angle neutron scattering (SANS) performed at ORNL using Bio-SANS instrument at the High Flux Isotope Reactor (HFIR), supported by DOE OBER’s Biological Systems Science Division, through ORNL Center for Structural Molecular Biology, and EQ-SANS instrument at Spallation Neutron Source (SNS) at ORNL, supported by the DOE OBES Scientific User Facilities Division (grant number ERKP-SNX).

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).

Neutrons Identify Oxygen Activation in LPMOs

Fungal lytic polysaccharide monooxygenases (LPMOs)
Scientists used neutrons to probe the structural details of the specialized LPMO fungal enzyme that relies on oxidation to digest molecules, aiming to improve the efficiency of enzymatic cellulose breakdown. [From O’Dell, W. B., P. K. Agarwal, and F. Meilleur. “Oxygen Activation at the Active Site of a Fungal Lytic Polysaccharide Monooxygenase.” Angew. Chem. Int. Ed. 56, 767–770 (2017). [DOI:10.1002/anie.201610502]. ©2017 The Authors. Published by Wiley-VCH Verla GmbH & Co. KGaA, Weinheim.]
Fungal lytic polysaccharide monooxygenases (LPMOs) are known to enhance the efficiency of cellulose-hydrolyzing enzymes through oxidative cleavage of the glycosidic bonds. For this study, PMO-2 from Neurospora crassa was heterologously expressed from Pichia pastoris, purified, and crystallized for high-resolution X-ray crystal structures that revealed “prebound” molecular oxygen in the resting state and a dioxo species in complex with the catalytic copper (Cu2+) ion, which is the first structural description of molecular oxygen (O2) activation by a LPMO. In addition, neutron diffraction studies and density functional theory calculations have identified a role for a conserved histidine in promoting oxygen activation. Extension of these studies to the enzyme-substrate complex could provide a complete picture of the enzymatic mechanism for the potential benefit of applications such as bioethanol production.

O’Dell, W. B., et al. “Oxygen Activation at the Active Site of a Fungal Lytic Polysaccharide Monooxygenase.” Angew. Chem. Int. Ed. 129(3), 785–788 (2017). [DOI:10.1002/anie.201610502].

Instruments and Facilities Used: Neutron crystallography. Joint X-ray/neutron refinement at Center for Structural Molecular Biology at Oak Ridge National Laboratory (ORNL). Diffraction data were collected at SER-CAT 22-ID at the Advanced Photon Source at Argonne National Laboratory and at CG-4D IMAGINE (NSF MRI 09229719) at the High Flux Isotope Reactor at ORNL.

Funding Acknowledgements: Protein expression and purification experiments at Center for Structural Molecular Biology (CSMB), a User Facility of the Office of Biological and Environmental Research (OBER), U.S. Department of Energy (DOE) Office of Science. Diffraction data collected at SER‐CAT 22‐ID at Argonne National Laboratory’s (ANL) Advanced Photon Source (APS) and at CG‐4D IMAGINE (National Science Foundation [NSF] magnetic resonance imaging [MRI] 09229719) at Oak Ridge National Laboratory’s (ORNL) High Flux Isotope Reactor (HFIR), both DOE Office of Biological Energy Sciences (OBES) User Facilities. W.B.O. support: NSF IGERT 1069091. F.M. support: U.S. Department of Agriculture (USDA National Institutes of Food and Agriculture (NIFA) Hatch 211001. P.K.A. support: NIH GM105978.

Description of Hydration Water in Green Fluorescent Protein Solution

green fluorescent protein
Graphic representation of hydration water surrounding green fluorescent protein. [Reprinted with permission from Perticaroli, S., et al. “Description of Hydration Water in Protein (Green Fluorescent Protein) Solution.” J. Am. Chem. Soc. 139(3), 1098–1105 (2017). [DOI:10.1021/jacs.6b08845]. Copyright 2016 American Chemical Society.]
Many unanswered questions remain about how macromolecules in solution perturb the water molecules in which they are immersed. Scientists used neutron scattering to provide an experimental description of the dynamical perturbation of water molecules surrounding proteins in solution, which play an important role in protein function and stability. Quantifying the magnitude of the perturbation of water around the green fluorescent protein (GFP), they found a systematic length-scale dependence of the dynamical retardation factor compared to the perturbation of bulk water. The findings deepen the understanding of water perturbation, which is of practical interest to researchers in food science, personal care, pharmaceutics, and protein dynamics.

Perticaroli, S., et al. “Description of Hydration Water in Protein (Green Fluorescent Protein) Solution.” J. Am. Chem. Soc. 139(3), 1098–1105 (2017). [DOI:10.1021/jacs.6b08845].

Instruments and Facilities Used: Neutron scattering at Center for Structural Molecular Biology and Spallation Neutron Source at Oak Ridge National Laboratory.

Funding Acknowledgements: H.O’N. and Q.Z. support: Center for Structural Molecular Biology, funded by the Office of Biological and Environmental Research (OBER), U.S. Department of Energy (DOE) Office of Science, under Contract FWP ERKP291. Research at Oak Ridge National Laboratory’s (ORNL) Spallation Neutron Source (SNS) sponsored by the Office of Basic Energy Sciences (OBES) Scientific User Facilities Division, DOE Office of Science. ORNL facilities sponsored by UT-Battelle, LLC, for the DOE under Contract No. DEAC0500OR22725.

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.