Method Article

Contrast-Matching Detergent in Small-Angle Neutron Scattering Experiments for Membrane Protein Structural Analysis and Ab Initio Modeling

DOI:

10.3791/57901

⸱

October 21st, 2018

 ,  ,  ,  ,  , 
1Neutron Scattering Division, Oak Ridge National Laboratory, 2School of Chemistry and Biochemistry, Georgia Institute of Technology

In This Article

Summary

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This protocol demonstrates how to obtain a low-resolution ab initio model and structural details of a detergent-solubilized membrane protein in solution using small-angle neutron scattering with contrast-matching of the detergent.

Abstract

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The biological small-angle neutron scattering instrument at the High-Flux Isotope Reactor of Oak Ridge National Laboratory is dedicated to the investigation of biological materials, biofuel processing, and bio-inspired materials covering nanometer to micrometer length scales. The methods presented here for investigating physical properties (i.e., size and shape) of membrane proteins (here, MmIAP, an intramembrane aspartyl protease from Methanoculleus marisnigri) in solutions of micelle-forming detergents are well-suited for this small-angle neutron scattering instrument, among others. Other biophysical characterization techniques are hindered by their inability to address the detergent contributions in a protein-detergent complex structure. Additionally, access to the Bio-Deuteration Lab provides unique capabilities for preparing large-scale cultivations and expressing deuterium-labeled proteins for enhanced scattering signal from the protein. While this technique does not provide structural details at high-resolution, the structural knowledge gap for membrane proteins contains many addressable areas of research without requiring near-atomic resolution. For example, these areas include determination of oligomeric states, complex formation, conformational changes during perturbation, and folding/unfolding events. These investigations can be readily accomplished through applications of this method.

Introduction

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Membrane proteins are encoded by an estimated 30% of all genes1 and represent a strong majority of targets for modern medicinal drugs.2 These proteins perform a wide array of vital cellular functions,3 but despite their abundance and importance — only represent about 1% of total structures deposited in the Research Collaboratory for Structural Bioinformatics (RCSB) Protein Data Bank.4 Due to their partially hydrophobic nature, structural determination of membrane-bound proteins has been exceedingly challenging.5,6,7

As many biophysical techniques require monodisperse particles in solution for measurement, isolating membrane proteins from native membranes and stabilizing these proteins in a soluble mimic of the native membranes has been an active area of research in recent decades.8,9,10 These investigations have led to the development of many novel amphiphilic assemblies to solubilize membrane proteins, such as nanodiscs,11,12,13 bicelles,14,15 and amphipols.16,17 However, the use of detergent micelles remains one of the most common and straightforward approaches for satisfying the solubility requirements of a given protein.18,19,20,21,22,23,24,25 Unfortunately, no single detergent or magic mixture of detergents currently exists that satisfies all membrane proteins; thus, these conditions must be empirically screened for the unique requirements of each protein.26,27

Detergents self-assemble in solution above their critical micelle concentration to form aggregate structures called micelles. Micelles are composed of many detergent monomers (typically ranging from 20-200) with hydrophobic alkyl chains forming a micelle core and hydrophilic head groups arranged in a micelle shell layer facing the aqueous solvent. The behavior of detergents and micelle formation has been classically described by Charles Tanford in The Hydrophobic Effect,28 and sizes and shapes of micelles from commonly used detergents in membrane protein studies have been characterized using small-angle scattering.29,30 Detergent organization about membrane proteins has also been studied, and the formation of protein-detergent complexes (PDCs) is expected with detergent molecules surrounding the protein in an arrangement that resembles the neat detergent micelles.31

One added advantage in using detergents is that the resulting micelle properties can be manipulated by incorporating other detergents. Many detergents exhibit ideal mixing, and select properties of mixed micelles may even be predicted from the components and ratio of mixing.22 However, the presence of detergent can still present challenges for biophysical characterizations by contributing to the overall signal. For example, with X-ray and light scattering techniques, signal from detergent in the PDC is practically indistinguishable from protein.32 Investigations with single-particle cryo-electron microscopy (cryo-EM) typically rely on trapped (frozen) particles; structural details of the protein are still obscured by certain detergents or a high concentration of detergent which adds to the background.33 Alternate approaches toward interpreting the full PDC structure (including the detergent) have been made through computational methods which seek to reconstruct the detergent around a given membrane protein.34

For the case of neutron scattering, the core-shell arrangement of detergent in the micelle produces a form factor which contributes to the observed scattering. Fortunately, solution components can be altered such that they do not contribute to the net observed scattering. This "contrast matching" process is achieved by substituting deuterium for hydrogen to achieve a scattering length density that matches that of the background (buffer). A judicious choice of detergent (with available deuterated counterparts) and their ratio of mixing must be considered. For detergent micelles, this substitution can be performed using a detergent with the same head group but having a deuterated alkyl chain (d-tail instead of h-tail). Since the detergents are well-mixed,35 their aggregates will have a scattering length density that is the mole-fraction weighted average of the two components (h-tails and d-tails). When this average contrast is consistent with that of the head group, the uniform aggregate structures can be fully matched to remove all contributions to observed scattering.

We present here a protocol to manipulate the neutron contrast of detergent micelles by incorporating chemically identical detergent molecules with deuterium-labeled alkyl chains.19,36,37 This permits complete simultaneous contrast matching of micelle core and shell, which is a unique capability of neutron scattering.35,38 With this significantly refined level of detail, contrast matching can enable otherwise unfeasible studies of membrane protein structures. Additionally, this contrast-matching approach could be extended to other systems involving detergent, such as polymer exchange reactions39 and oil-water dispersants,40 or even other solubilizing agents, such as bicelles,41 nanodiscs,42 or block copolymers.43 A similar approach as outlined in this manuscript, but employing a single detergent species with partial deuterium substitutions on the alkyl chain and/or head group, was recently published.37 While this can be expected to improve the random distribution of hydrogen and deuterium throughout the detergent compared to the approach presented here, the limited number of available positions on the detergent for substitution and two-step detergent synthesis required poses additional challenges for consideration.

Steps 1 and 2 of the protocol detailed below often overlap since initial experiment planning must be done to submit a quality proposal. However, proposal submission is considered here as the first step to emphasize that this process should be started well in advance of a neutron experiment. It should also be noted that a prerequisite step, which should be demonstrated by the proposal, is to have biochemical and physical characterization (including purity and stability) of the sample supporting the need for neutron studies. A general discussion of small-angle neutron scattering (SANS) is beyond the scope of this article. A brief but thorough introduction is available in the reference work Characterization of Materials by Kaufmann,44 and a comprehensive textbook focused on biological small-angle solution scattering has recently been published.45 Further recommended reading is given in the Discussion section. Small angle scattering uses the so called scattering vector Q as the central quantity that describes the scattering process. This article uses the widely accepted definition Q = 4π sin(θ)/λ, where θ is half the angle between incoming and scattered beam and λ is the wavelength of the neutron radiation in Angstroms. Other definitions exist that use different symbols such as 's' for the scattering vector, and that may differ by a factor 2π or by using nanometers in place of Angstrom (see also discussion of Figure 10).

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Protocol

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1. Prepare and Submit a Neutron Facility Beam Time and Instrument Proposal

  1. Consult online resources for identifying neutron scattering facilities that provide general user neutron beam time access, such as Oak Ridge National Laboratory (ORNL). A map of neutron facilities and information about neutron research worldwide is available online.46 Be aware that these facilities typically have regular calls for proposals; this determines when the next beam time will be available. Neutrons are used for a variety of applications; search for small-angle neutron scattering (SANS) instruments, particularly those with capabilities for biological samples.
  2. Use resources provided by the neutron facilities to help navigate the neutron beam time proposal submission process. Consult with a Neutron Scattering Scientist (NSS) that is associated with the instrument for which you will apply. Review the neutron facility’s current information regarding proposal calls, deadlines, and advice for submission; for Oak Ridge this is available online.47 Submit the beam time proposal following all guidelines for a successful submission.
  3. If deuterated protein is required (see 2.2), neutron scattering facilities are often supported by laboratories and expertise dedicated to the production of deuterated materials. To request access to the Bio-Deuteration Lab (BDL) at ORNL, select the BDL as the second instrument in the proposal system. While the SANS proposal is reviewed for feasibility and by a scientific review committee, BDL requests are only screened for feasibility. To help with the BDL feasibility review, submit a completed information request form48 that describes the protein expression protocol and estimated yield (available online).
  4. After successful peer-review of the beam time proposal, confirm the awarded date of beam time in advance of the experiment. Ensure that all sample and buffer details and formulas are listed correctly in the proposal for proper review. Any changes to the proposed experimental plan, including changes in sample and buffer composition, require notification of the facility.
    NOTE: Changes should be discussed as soon as possible with the assigned NSS.

2. Determine Neutron Contrast Match Points and Necessary Contrast for Protein Measurement

  1. Gather information (from supplier’s product information, online databases, published values, etc.) relating to atomic compositions and volumes for the detergent system components to be contrast-matched. This information is used to determine neutron scattering length densities (SLDs) and thus contrast match points (CMPs) in solution, which is critical to obtaining quality SANS data and structural information for the membrane protein of interest in the context of a PDC. A table summarizing the physical properties and contrast match points for detergents commonly used in biophysical studies of membrane proteins is included (Table 1).
  2. Determine neutron SLDs using the web application MULCh: ModULes For The Analysis Of Contrast Variation Data49 (available online50 free-of-charge). A link to the user manual is located on the home page. Access the website and read the accompanying documentation. Figure 1 and Figure 2 provide an overview of Contrast module input and output for two related examples.
    NOTE: Other websites for calculating SLDs are available, such as the one maintained by the National Institute of Standards and Technology (NIST) Center for Neutron Research.51
    1. Open the Contrast module by clicking ‘Contrast’ located in the left navigation pane. Provide text for a project title and enter details below for two components (e.g., detergent head group and tail) as ‘subunit 1’ and ‘subunit 2’.
    2. Enter the molecular formula for each component in the ‘Formula’ text box.
      NOTE: The radio button ‘M’ must be selected under ‘Substance Type’ to enter a molecular formula. Additionally, use the letter ‘X’ in the formula to indicate readily exchangeable hydrogen atoms. Protein, RNA, or DNA sequences can be entered if the corresponding ‘P’, ‘R’, or ‘D’ radio buttons are selected. If multiple copies of a component exist within the subunit, the value for ‘Nmolecules’ can be changed accordingly. A practical example here relates to lipid-like detergents containing two identical tails, allowing the formula for one tail to be entered with Nmolecules equal to 2.
    3. Enter the volume in cubic Angstroms for each component in the box below ‘Volume (Å3)’. Use Tanford’s formula28 for alkyl chains of length n, Vtail = 27.4 + n * 26.9, to calculate approximate tail volume, or obtain molecular volumes from product information provided or published values in the literature.
    4. Enter details for buffer components. Change the ‘Number dissolved species in the solvent’ using the dropdown menu to add or remove rows for each buffer component. In the case of micelle-forming detergents, include the free detergent monomers as separate buffer components at the detergent’s critical micelle concentration (CMC).
    5. Click ‘Submit’ to perform the neutron contrast calculations and generate a results page that provides a table of scattering length density and contrast match points as well as formulae for determining these parameters at any given percentage of D2O in the buffer. Review the scattering parameters with particular attention to the component CMPs. If the match points are similar (within 10% D2O) and the free micelle concentration is low, then the average CMP can be used for contrast-matching the detergent contributions. In most cases, the CMP of the detergent head group and tail will differ by more than 10-15%, producing a core-shell form factor in the SANS data which obfuscates direct collection of the scattering signal from the protein alone.
  3. Assess the contrast between detergent head group and tail. When the detergent head group and alkyl chain tail CMPs are not well-matched, the availability and incorporation of a deuterium-substituted detergent can permit scattering length densities of select components to be manipulated. In most cases, this will require forming a mixed micelle through the incorporation of an identical detergent with deuterium-substituted alkyl chains. The addition of deuterium raises the scattering length density of the core formed by the alkyl chain tails. The desired endpoint, in this case, is such that the head group shell CMP and alkyl chain core CMP have approximately equal values.
    1. Raise the CMP of the detergent micelle core to be approximately equal to the CMP of the head group shell by determining the appropriate ratio of mixing between h-tails and d-tails that achieves complete contrast matching with the head groups. A prerequisite for this determination is knowledge of the CMP for a deuterium-substituted alkyl chain tail. A commercially-available counterpart to n-Dodecyl β-D-maltoside (DDM) is available with all alkyl chain hydrogen replaced by deuterium (d25-DDM). Repeat the contrast calculations outlined in step 2.1 for a ‘d-tail’ in place of the ‘h-tail’.
    2. Calculate the mole ratio of h-tail to d-tail required for contrast-matching with the head group CMP. The following formula can assist with this determination:
      CMPhead = CMPh-tail * χh-tail + CMPd-tail * χd-tail
      where CMP is the scattering length density and χ is the volume fraction for the detergent head, h-tail, or d-tail component. For buffered solutions of DDM, incorporation of 44% by weight (43% by mole) of the total detergent as d25-DDM with deuterium-substituted alkyl chains produces a single CMP in 48.5% D2O for both head and tail components.
  4. Consider the degree of deuterium substitution for the membrane protein. To measure sufficient scattering signal from the protein, the CMP for the protein should be at least 15% D2O in solution away from the detergent CMP and measurement conditions. The signal increases with the square of this% difference. Since the CMP of an unlabeled protein typically occurs with ~42% D2O in solution (a CMP similar to many detergent head groups), deuterium labeling of the protein is almost always a necessity.52

3. Express and Purify the Membrane Protein of Interest

  1. Prepare LB (lysogeny broth) medium and grow Escherichia coli (E. coli) cells harboring an inducible expression vector for the target protein sequence.
    1. Prepare minimal media. In either H2O or D2O, dissolve 7.0 g/L (NH4)2SO4, 5.25 g/L Na2HPO4, 1.6 g/L KH2PO4, 0.50 g/L diammonium hydrogen citrate, 5.0 g/L glycerol, 1.0 mL/L of 20% w/v MgSO4·7H2O, and 1.0 mL/L of Holme trace metals (0.50 g/L CaCl2·2H2O, 0.098 g/L CoCl2, 0.102 g/L CuSO4, 16.7 g/L FeCl3·6H2O, 0.114 g/L MnSO4·H2O, 22.3 g/L Na2EDTA·2H2O, and 0.112 g/L ZnSO4·H2O).53,54
    2. Prepare appropriate antibiotic stock solutions using H2O or D2O.
    3. Prepare an isopropyl-β-D-1-thiogalactopyranoside stock solution using H2O or D2O.
    4. Sterile-filter all solutions into dry, sterile containers using bottle-top vacuum and syringe filters with a 0.22 micron pore size.
  2. Adapt E. coli cells to deuterium-labeled medium prior to scale-up for overexpression of a deuterated protein.
    1. Inoculate 3 mL of LB medium with an isolated colony from a Lysogeny Broth (LB) agar plate or cells from a frozen glycerol stock. Grow cells at 37 °C in a shaking incubator at 250 RPM or reduce the temperature to 30 °C or below to avoid overgrowth of the culture when incubating overnight.
      NOTE: The adaptation procedure can be varied.55,56,57
    2. Once the LB culture has grown to an optical density at 600 nm (OD600) of ~1, dilute it 1:20 in 3 mL of H2O minimal medium and grow to an OD600 of ~1. Repeat the 1:20 dilutions using minimal medium containing 50, 75 and 100% D2O (or up to the desired D2O percentage).
      NOTE: As the D2O content is increased, growth rates will decrease. The relationship between the D2O percentage in the growth media and deuterium substitution in the protein has been reported in the literature.58,59,60
    3. Continue growing the culture in a bioreactor to increase the yield of deuterated cell mass (Figure 3).
      NOTE: Step-by-step procedures for bioreactor operation have been reviewed elsewhere.61,62,63,64
  3. Harvest and lyse E. coli cells for membrane extraction and protein purification
    1. Pellet the cells by centrifugation at ~6,000 x g for ~30-45 min at 4 °C.
    2. Soften the cell pellet by soaking in buffer on ice for ~30 min. Then, gently resuspend the cell pellet in buffer by gently pipetting with a serological pipet, or gentle rocking on a 2D rocker, to a final concentration of ~1 g wet cell mass/10 mL buffer.
      NOTE: An example buffer is 50 mM HEPES (4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid), pH 7.5, 200 mM NaCl supplemented with a protease inhibitor tablet.
    3. Lyse resuspended cells with three passes through a high-pressure homogenizer at 10,000 psi while chilling the outflow.
      NOTE: Depending on the scale required, other lysis methods may be used (e.g., by French press or sonication).
    4. Pellet cell debris by centrifugation at ~4,000 - 5,000 x g for 15 min at 4 °C. Repeat this step until the supernatant is clarified.
    5. Ultracentrifuge (~150,000 x g for 30-45 min) the supernatant from the prior step, which contains the soluble and membrane fractions. The pellet after ultracentrifugation contains the membrane fraction.
    6. Resuspend the membrane pellet in a large Dounce homogenizer and isolate the membrane fraction again by ultracentrifugation (step 3.3.5). Repeat this step at least once to remove loosely bound proteins.
    7. Solubilize membranes by gentle rocking for ~30 min at 4 °C in a buffer containing detergent suitable for purification. Solubilization is apparent when the suspension turns from cloudy to transparent. Remove insoluble material by ultracentrifugation (~150,000 x g for 30-45 min).
      NOTE: An example buffer is 50 mM HEPES, pH 7.5, 500 mM NaCl, 20 mM imidazole and 4% (w/v) DDM for purification by Ni2+ affinity chromatography.
  4. Purify the protein following a protocol previously developed for protonated forms.
    NOTE: See Naing et al. for an example purification protocol for a hexahistidine tagged M. marisnigri JR1 intramembrane aspartyl protease (d-MmIAP).65 The final yield was ~ 1 mg of deuterated from 5 L deuterated cell culture growth, all of which was used in the SANS experiment (Figure 4).
  5. Exchange the purified protein into the “Final Exchange Buffer” for contrast matching.
    1. Prepare 200 mL of “Final Exchange Buffer” containing the correct mixture for contrast matching, e.g.,20 mM HEPES, pH 7.5, 250 mM NaCl, and 0.05% total DDM (0.028% DDM and 0.022% d25-DDM) in 49% D2O.
    2. Equilibrate a size exclusion column with >2 column volumes (CV) of “Final Exchange Buffer” in 3.5.1 using a Fast Protein Liquid Chromatography (FPLC) system.
    3. After injecting the sample, run the column and isolate fractions corresponding to the protein of interest.
    4. Concentrate the protein to the desired final concentration (2 - 5 mg/mL).
      NOTE: A volume of ~300 μL is required to fill a 1 mm cylindrical quartz cuvette used for SANS.
    5. Reserve an aliquot of matched buffer for SANS background subtraction.
      NOTE: The aliquot can be taken directly from the prepared buffer, or ideally from a clean elution fraction at the end of the FPLC run.

4. Make Final Preparations for Beam Time and Collect SANS Data

  1. After the proposal has been accepted and site access has been approved, complete all required training in advance of assigned beam time. This includes prearrival, web-based training as well as on-site radiological, laboratory, and instrument-specific training.
    NOTE: Training requirements may dictate advance arrival for first time users. More information is available online.66
  2. Load samples and buffers for data collection.
    NOTE: An overview of the biological small-angle neutron scattering (Bio-SANS) instrument sample environment area is shown in Figure 5.
    1. Load samples and buffers into quartz cells. Quartz cells are available from the instrument scientist or facility. Use gel-loading tips to access the narrow opening of most sample cells.
    2. Ensure that the beam shutter is closed, then approach the sample environment area and place the quartz cells in the sample changer. Record the sample changer position for each sample cell.
    3. Check the area and ensure the beam path is free from obstruction, then leave the sample environment area and open the beam shutter.
      NOTE: Carefully follow all facility rules and regulations, obey all postings, and heed advice of the instrument scientist. Samples may not be removed from the beam and manipulated after exposure to the beam without following special precautions. Consult with a Radiological Control Technician (RCT) prior to these procedures.
  3. Execute table scan for automated data collection
    1. Operate the instrument through custom control LabVIEW-based software, Spectrometer Instrument Control Environment (SpICE). Follow instructions for instrument operation provided by the Neutron Scattering Scientist. An overview of the table scan operation windows is provided in Figure 6.
    2. After entering the information required in the appropriate areas, execute the table scan and an automated data collection process will begin. Monitor progress via the ‘Table Scan Status’ tab.

5. Reduce SANS Data from 2D Image to 1D Plot

  1. Identify each sample and buffer data file from the recorded scan numbers. Each measurement will be assigned a unique scan number. This information is helpful during data reduction to identify each corresponding sample and buffer pair.
  2. Use MantidPlot software and Python script for reduction
    1. Neutron Scattering Users are provided with account details to access their data on the Remote Analysis Cluster.67 Log in to the Remote Analysis Cluster and execute “MantidPlot” from the command line. Figure 7 and Figure 8 are provided to assist these steps.
    2. Obtain a User Reduction Script from the Neutron Scattering Scientist (this will usually happen during the experiment); this script is Python-based and will contain all the necessary calibration and scaling adjustments to reduce the 2D scattering image into a 1D plot of scattered intensities as a function of scattering angle, I(Q).
    3. Open the provided User Reduction Script in MantidPlot and place the corresponding scan numbers or identification for the sample and buffer pairs in the appropriate list.
    4. Execute the script to generate reduced data as a four-column text file (column order is Q, I(Q), I(Q) error, Q error) in the specified location. Right click the appropriate workspace in MantidPlot and select “Plot with errors…” for an initial examination of the scattering profiles.

6. Analyze Data for Structural Parameters of the Scattering Particle

  1. Transfer the reduced data file from the analysis server to a local computer using secure FTP file transfer. This can be performed with the use of software such as FileZilla or CyberDuck. Additional instructions and details for connecting to the analysis server file system are provided on the analysis.sns.gov login page.
  2. Download the ATSAS software suite65,68 (whole ATSAS suite).69 Individual programs70 are also available online for analyzing the SANS data, namely determining structural parameters and ab initio models.
    NOTE: Many options are available for SANS data analysis of biomacromolecules.45
  3. Use PRIMUS71 for data plotting, buffer scaling and subtraction, and Guinier analysis.
    1. Launch the ATSAS ‘SAS Data Analysis’ application and load the reduced data files corresponding to the sample and buffer pair.
      1. To scale the buffer properly, select a data range at high Q (Q > 0.5 Å-1) where both profiles are similar and flat, and click the ‘Scale’ button located under the ‘Operations’ tab. A scale factor will be applied to the buffer such that these two flat regions should overlap. Use caution: there should be a plausible physical reason that justifies scaling the buffer intensity to match the sample. If the discrepancy is large, consult an instrument scientist for valid approaches to background subtraction.44,45,72
      2. Increase the data range to view all points. Click ‘Subtract’ to perform this operation. Right click the buffer-subtracted datafile in the legend and save this file for subsequent analysis. Figure 9 outlines the use of the ‘Operations’ tab.
    2. Perform a Guinier analysis of the buffer-subtracted sample data using the ‘Analysis’ tab of the ‘SAS Data Analysis’ application.
      1. Be sure the proper file is selected in the list and click ‘Radius of Gyration’. An automated attempt to perform a Guinier fit will be provided by clicking on the ‘Autorg’ button.
      2. Expand the range of data used to include all of the low-Q data and begin narrowing the data range by taking away high-Q points until the Qmax*Rg limit is below 1.3. Use the plot of residuals to verify that data are linear in the fit range. The Guinier fit yields an approximate Rg and I0 for the scattering particles.
      3. Make small adjustments to the fit region and monitor the sensitivity of these values to the range of data used in the fit. Figure 10A demonstrates the use of the ‘Radius of Gyration’ tool.
  4. Obtain the probability distribution function (P(r)) in GNOM.73 The output file generated here will be used as an input for the ab initio modeling process.
    1. Start the Distance Distribution Wizard within the ‘Analysis’ tab. Obtaining a good fit to the data is essential to obtaining a quality model. For more information on obtaining accurate fits, please see the review74 by Putnam, et al.
      NOTE: The GNOM program provides additional information about the fit beyond the Distance Distribution Wizard. Figure 10B demonstrates proper use of the ‘Distance Distribution’ tool, and Figure 11 illustrates some of the common errors encountered.
    2. Determine Dmax, the maximum interatomic distance within the molecule.
      1. Estimate a value for Dmax by unchecking the box to force Rmax=0 and entering a large value for Dmax (150 Å, for example). The first x-intercept in the plot of P(r) yields this estimate.
      2. Make incremental changes to this Dmax value and the range of data used or number of points used in the fit to optimize the GNOM fit to the data and the resulting P(r) curve.
    3. Continue to refine the GNOM fit and save the GNOM output files with good fit parameters for subsequent ab initio modeling steps.
  5. Simulate SANS profiles from high resolution PDB models using the program CRYSON.75
    1. Open the program and select option ‘0’. Select the PDB file name in the popup file browser. Press enter to accept the default settings, except for specifying chain deuteration fractions and the fraction of D2O in the solvent to achieve the proper contrast parameters.
    2. Fit the model to an experimental data set by entering ‘Y’ and selecting the data file in the popup file browser. Accept the remaining default values, and output files will be written in the PDB file location. The ‘.fit’ file contains information for the predicted scattering profile from the high-resolution 3D model.

7. Create Ab Initio Models from the SANS Data.

NOTE: DAMMIF76 and DAMMIN77 within the ATSAS software suite is used to reconstruct dummy atom models (DAMs) using a simulated annealing process from the GNOM output, which contains P(r) data, or information about the probability or frequency of interatomic distances within the scattering particle. These programs may be run in batch mode or on the ATSAS-Online web server.

  1. Start the PRIMUS Shape Wizard from the ‘Dammif’ button on the ‘Analysis’ tab of ‘SAS Data Analysis’, and use a manual selection of parameters. The input for the wizard is illustrated in Figure 12.
    1. Define the Guinier range from the fit (step 6.3.2) and proceed to the next step using the navigation buttons. Define fit values from the P(r) plot (step 6.4) and proceed to the next step.
    2. Provide parameters for the ab initio modeling process. Enter a prefix for the model output filenames (e.g., SANSEnvelope), choose either ‘fast’ or ‘slow’ mode modeling, enter a value for the number of models (17 recommended for statistical significance), select from available options for particle symmetry, anisometry, and angular scale (if known). Ensure that the boxes are checked to average models with DAMAVER and refine the final model with DAMMIN.
    3. Initiate the process using the ‘commit’ button. Once the process is complete, view and save the working directory.
      NOTE: The typical modeling process for a single, small protein can take up to a few hours with a typical personal computer. Files are stored in temporary directories until saved.
  2. Overlay and superimpose the final SANS envelope with a related high-resolution model using SUPCOMB within ATSAS. Place a copy of the high-resolution PDB model to be fit to the SANS envelope in the working directory. Execute SUPCOMB from the command line using the PDB filenames as two arguments with the template structure listed first: $ supcomb HiRes.pdb SANSEnvelope.pdb
    NOTE: The second filename listed will be rewritten as a new file with an “r” appended to the end and having new coordinates for superimposition on the template structure.
  3. Visualize the ab initio model results using PyMOL (pymol.org), a 3D molecular graphics viewing program. Figure 13 provides an overview of the PyMOL operations.
    NOTE: There are publication guidelines for structural modeling of small-angle scattering data from biomolecules in solution.78
    1. Launch the PyMOL application and open the .pdb files corresponding to the 3D SANS envelope and the SUPCOMB-aligned high-resolution structure to be viewed.
    2. Visualize the SANS envelope by representing this model as a surface. Click the ‘S’ button next to the model and select ‘Show: As: Surface’.
    3. Visualize the high-resolution protein backbone structure by representing this model as a cartoon. Select the ‘S’ button next to this model and select ‘Show: As: Cartoon’. Display the chain as a rainbow from N- to C- terminus by selecting the ‘C’ button and ‘By Chain: Chainbows’.
    4. Modify the transparency of the surface representation for clarity. From the ‘Setting’ menu bar, select ‘Transparency » Surface » 40%’.
    5. Make the background white and non-opaque. From the ‘Display’ menu bar, select ‘Background »’ and select ‘Opaque’ to uncheck this option and ‘White’ to mark this color for background.
      NOTE: Additional operations and uses for PyMOL can be found at PyMolWiki.org.

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Results

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A beam time and instrument proposal should clearly convey all information needed to the review committee so that a valid assessment of the proposed experiment can be made. Communication with an NSS is highly suggested for inexperienced users. The NSS can assess initial feasibility and guide proposal submission to emphasize feasibility, safety, and the potential for high-impact science. The information provided in the proposal should include background information and context for the signi...

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Discussion

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Structural biology researchers take advantage of complementary structural techniques like solution scattering to obtain biochemical and structural details (such as overall size and shape) from biomolecules in solution. SANS is a particularly attractive technique for determining low resolution structures of membrane proteins, a core focus of modern structural biology and biochemistry. SANS requires quantities of purified proteins comparable to those of crystallographic trials (1 mg/sample). The recently expanding commerci...

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Disclosures

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The authors Volker S. Urban, Sai Venkatesh Pingali, Kevin L. Weiss, and Ryan C. Oliver are contracted through UT-Battelle with US DOE to support the Neutron Science User Program and Bio-SANS instrument at Oak Ridge National Laboratory used in this Article.

This manuscript has been co-authored by UT-Battelle, LLC, under contract DE-AC05-00OR22725 with the US Department of Energy (DOE). The US government retains and the publisher, by accepting the article for publication, acknowledges that the US government retains a nonexclusive, paid-up, irrevocable, worldwide license to publish or reproduce the published form of this manuscript, or allow others to do so, for US government purposes. DOE will provide public access to these results of federally sponsored research in accordance with the DOE Public Access Plan.82

Acknowledgements

$$\rightleftharpoonup{xx}$$ $$\longleftharp{xx}$$, $$\longrightharp{xx}$$,

The Office of Biological and Environmental Research supported research at ORNL's Center for Structural Molecular Biology (CSMB) and Bio-SANS using facilities supported by the Scientific User Facilities Division, Office of Basic Energy Sciences, US Department of Energy. Structural work on membrane proteins in the Lieberman lab has been supported by NIH (DK091357, GM095638) and NSF (0845445).

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Materials

List of materials used in this article
NameCompanyCatalog NumberComments
Amicon Ultra MWCO 50KDa concentrator EMD MilliporeUFC905096labware
Ammonium citrate dibasicFisher ScientificA663medium component
Ammonium sulfateEMD Millipore2150medium component
Bioflo 310 Bioreactor SystemEppendorfM1287-2110equipment
Calcium chloride dihydrateAcros423525000medium component
CarbenicillinIBI ScientificIB02025antibiotic
ChloramphenicolEMD Millipore3130antibiotic
Cobalt (II) chlorideAcrosAC21413-0050medium component
Copper (II) sulfateAcrosAC19771-1000 medium component
Deuterium oxideSigma-Aldrich756822medium component
Drierite Gas PurifierW.A. Hammond Drierite Co. Ltd.27068
EDTA, disodium, dihydrateEMD Millipore4010medium component
Emulsiflex-C3AvestinEF-C3equipment
Äkta Purifier UPC100GE Healthcare equipment
GlycerolSigma-AldrichG5516medium component
HEPESSigma-AldrichH4034
HiPrep 16/60 Sephacryl S-300 HR columnGE Healthcare 17116701
ImidazoleVWR97064-622
IPTGTeknovaI3325
Iron(III) chloride hexahydrateMP BiochemicalsICN19404590medium component
LB Agar MillerFisher ScientificBP1425-2
Magnesium sulfate heptahydrateVWR97062-134medium component
Manganese(II) sulfate monohydrateAcrosAC20590-5000medium component
MaxQ 6000 Incubated/Refrigerated ShakerThermo ScientificSHKE6000-7 equipment
n-Dodecyl-d25-β-D-maltopyranosideAnatraceD310T
n-Dodecyl-β-D-maltopyranosideAnatraceD310A
Potassium phosphate monobasicVWR97062-346medium component
RC 6 Plus CentrifugeThermo Scientific Sorvall46910equipment
SIGMAFAST protease inhibitor cocktail tablets, EDTA-freeSigma-AldrichS8830
Sodium chlorideSigma-AldrichS3014
Sodium hydroxideSigma-Aldrich795429
Sodium phosphate dibasicSigma-AldrichS7907medium component
Sterile 25mm syringe filter with 0.2µm PES membraneVWR28145-501labware
Sterile disposable bottle top filter with 0.2µm PES membraneThermo Scientific596-4520 labware
Superdex 200 10/300 GL GE Healthcare 17517501
Superose-12 10/300 GL column GE Healthcare 17517301
Ultrospec 10 Cell Density MeterGE Healthcare 80211630equipment
Zinc sulfate monohydrateAcrosAC38980-2500 medium component

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Membrane Protein AnalysisDeuterium Labeled ProteinGuinier AnalysisSize Exclusion ChromatographyNeutron Contrast MatchingSANS Data ReductionPyMOL Visualization

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