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.

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]

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.

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.

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