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-funded Energy Frontier Research Center based at Washington University. Small angle neutron scattering (SANS) 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 (

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, 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.

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.

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,]
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.

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,]
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]

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.