This work was supported by funds from the Roy J. Carver Charitable Trust to Julien Roche. We thank J. D. Levengood and B. S. Tolbert for kindly providing the RRM2 sample.

NOTE: The protocol described here requires (i) a high-pressure pump and cell with a 2.5 kbar rated aluminum-toughened zirconia tube18, (ii) the software SPARKY19 for analysis of the NMR spectra, and (iii) a curve fitting software.
1. Sample preparation, assembly of the high-pressure cell, and setting up the experiments.
<ol>
	<li>Choice of buffer: Use equal mixture of anionic and cationic buffers, such as phosphate and Tris20,21. 
	NOTE: The pKa of anionic buffers such as phosphate and MES is associated with a substantial reaction volume (i.e., the difference in the partial molal volumes of the acid and the ionized products). The pH of such buffers can therefore be significantly affected by a change of pressure (~0.25-0.5 pH unit/kbar).</li>
	<li>Ensure that the required sample volume is similar to that of a standard 3 mm diameter NMR tube (~300&#956;L).</li>
	<li>Introduce the 15N-labeled sample with a glass pipette into the zirconia tube. Make sure the sample seats at the bottom of the tube. Complete with 200 &#956;L of mineral oil to prevent the sample from mixing with the transmission liquid (e.g., water). Fill the rest of the tube with transmission liquid.</li>
	<li>Put a single-use O-ring on top of the zirconia tube and slide the tube into the base (Figure 1A,B). Then, connect the tube to the high-pressure tether line and tighten the base to the cell first by hand. Then, apply 14.7 Nm of torque to prevent leaks at lower pressure (Figure 1C,D).</li>
	<li>To check the integrity of the pressure cell assembly, pressurize the tube up to 300 bar outside the spectrometer using cell support and containment vessel. Wait for 15 min, reset the pressure to 1 bar and check for leaks with a clean lint-free wipe.</li>
	<li>Insert the unpressurized tube into the spectrometer by carefully guiding the tether line. Slide the tube in the spectrometer until reaching the sample sitting position (Figure 1E).</li>
	<li>Lock, shim, match, and tune the 1H and 15N channels as usual. 
	NOTE: Shims for high-pressure rated zirconia tubes are very different from standard NMR tubes. It is recommended to save the optimized shims for future use.</li>
	<li>Set up a 1H-15N-HSQC or TROSY-HSQC and record a reference experiment at atmospheric conditions (1bar).</li>
</ol>
2. Recording high-pressure NMR experiments
<ol>
	<li>Gradually increase the pressure from 1 bar to 2.5 kbar with 500 bar increments to test the overall stability of the protein. Set the speed of the pressure pump by default at ~18 bar/s. If the precise folding/unfolding rates are not known, let the sample equilibrate 15-20 min after each 500-bar increment. Record a spectrum at 2.5 kbar.</li>
	<li>Gradually decrease the pressure back to 1 bar with 500 bar steps to test the reversibility of the pressure perturbation. Record another spectrum at atmospheric conditions and compare the chemical shifts and peak intensities with that of the reference spectrum previously recorded in the same conditions. 
	NOTE: If the native crosspeaks are more intense after the pressure run, it may indicate that small aggregates present in solution at atmospheric pressure may have dissociated and properly refolded. On the other hand, a loss in the intensity or significant chemical shift changes suggest that the protein may experience a non-reversible misfolding in high-pressure conditions.</li>
	<li>Record a series of 2D experiments from 1 bar to 2.5 kbar every 500 bar. It is recommended to record additional experiments near the inflection point of the folding/unfolding transition to improve the precision of the fit.</li>
</ol>
3. Analyzing peak intensity changes
<ol>
	<li>Process all the spectra and transfer the backbone assignment from the reference spectrum at 1 bar to the spectrum recorded at 500 bar, and then transfer the assignment from 500 bar to 1 kbar and so forth. 
	NOTE: Because pressure induces a non-uniform shift of 1H and 15N chemical shifts, do not simply copy the backbone assignment from one spectrum to the next. Adjust it manually.</li>
	<li>In Sparky menu, click on Peak &#62; Peak List (lt). In the Peak List window, click on Options and select the option to display both Frequencies (ppm) and Data Height. Save the list obtained for each spectrum.</li>
	<li>In a curve fitting software, copy the crosspeak identity and peak intensity values to have the pressure values (in bar) as the X-axis variable and the intensity as the Y-axis variable.</li>
	<li>If a complete or near complete (&#62;80%) unfolding is observed, fit the individual peak intensity profiles to extract the free energy and volume change upon unfolding, respectively, using a simple two-state model: 
	<img alt="Equation 1" src="/files/ftp_upload/62701/62701eq01.jpg" />&#160; &#160; &#160; &#160; &#160; &#160; &#160; &#160; &#160; &#160; &#160; &#160; &#160; &#160; &#160; &#160; &#160; &#160; &#160;Eq. 1 
	Where, &#34;I&#34; is the observed intensity of a crosspeak at a given pressure p and IF is the intensity of the same crosspeak in a fully folded state. R is the gas constant, T is the absolute temperature, &#916;GU0 the standard Gibbs free energy difference between the unfolded and folded states at atmospheric pressure p0 (1 bar), and &#916;Vu the volume change upon unfolding. With the pressure p in bar and temperature T in kelvin, R = 1.987 cal/K, &#916;GU0 is in cal/mol, and &#916;VU is in cal/mol/bar. Multiply the &#916;VU values obtained from the fit by 41.84 to convert into mL/mol. &#916;VU values are typically in the range of -50 to -150 mL/mol for globular proteins22. Here, all the parameters are expressed with regard to the unfolding reaction but can easily be converted into folding reaction parameters (&#916;GU0 = -&#916;GF0 and &#916;VU = -&#916;VF).</li>
</ol>
4. Analyzing chemical shift changes
<ol>
	<li>Arrange the columns in the software in order to have pressure points as variable and the 1H chemical shifts extracted from Sparky lists as the Y-axis.</li>
	<li>Fit the pressure dependence of the 1H chemical shifts to a simple quadratic equation: 
	&#948;(p) = &#948;0(p0) + B1(p-p0) + B2 (p-p0)2&#160; &#160; &#160; &#160; &#160; &#160; &#160; &#160; &#160; &#160;&#160;Eq. 2 
	Where, &#948;(p) is the measured 1H chemical shift of a crosspeak at a given pressure p and &#948;0(p0) the 1H chemical shifts of the same crosspeak in the reference spectrum recorded at 1 bar. B1 and B2 represent the first and second-order parameters expressed in ppm/bar and ppm/bar2, respectively.</li>
</ol>

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D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Characterization+of+enzyme+motions+by+solution+NMR+relaxation+dispersion.">Characterization of enzyme motions by solution NMR relaxation dispersion.</a> Accounts of Chemical Research. 41, 214-221 (2008).</li><li>Ishima, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=CPMG+relaxation+dispersion.">CPMG relaxation dispersion.</a> Methods in Molecular Biology. 1084, 29-49 (2014).</li><li>Longo, D. 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All the authors have read and approved the manuscript. They declare no conflicts of interest.

This&#160;study details a protocol implemented to probe protein structural and thermodynamics responses to pressure perturbation. The high-pressure experiments recorded here on RRM2 demonstrate that large variations in &#916;VU values, indicative of non-fully cooperative unfolding, can be found in a relatively small single domain protein. A similar picture emerges from the analysis of 1H chemical shift changes under pressure. It should be noted Kalbitzer and coworkers have demonstrated that a more i...

It has long been recognized that higher-energy, sparsely populated conformational states of proteins and protein complexes play a key role in many biological pathways1,2,3. Thanks to experiments based on Carr-Purcell-Meiboom-Gill (CPMG)4, Chemical Exchange Saturation Transfer (CEST)5, and dark-state exchange saturation transfer (DEST)6 pulse sequences (among others), solution NMR spectroscopy has emerged as a method of choice for characterizing transient conformational states7. Along with these experiments, perturbations such as temperature, pH, or chemical denaturants can be introduced to increase the relative population of higher energy conformational substates. Similarly, protein equilibria can also be perturbed by applying high hydrostatic pressure. Depending on the magnitude of the volume change associated with the corresponding conformational changes, an increase of pressure by a few hundred to a few thousand bars can significantly stabilize a higher energy state or cause a protein to completely unfold8,9,10. Protein NMR spectra typically display two types of changes with hydrostatic pressure: (i) chemical shift changes and (ii) peak intensity changes. Chemical shift changes reflect changes at the protein surface-water interface and/or local compression of the protein structure on a fast time scale (relative to NMR time scale)11. Crosspeaks exhibiting large non-linear chemical shifts pressure dependence can indicate the presence of higher energy conformational states12,13. On the other hand, peak intensity changes point to major conformational transitions on a slow time scale, such as changes in folded/unfolded state populations. The presence of folding intermediates or higher energy states can be detected from large variations in the magnitude of the volume change upon unfolding measured for different residues of a given protein14,15,16,17. Based on our experience, even small proteins that are typically classified as two-state folders exhibit non-uniform responses to pressure, which provides useful information about their local folding stability. Described here is a protocol for the acquisition and analysis of amide peak intensity and 1H chemical shifts pressure dependence, using as a model protein the isolated RNA recognition motif 2 (RRM2) of the heterogeneous nuclear ribonucleoprotein A1 (hnRNPA1).

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Bruker Nmr Cell 2.5 Kbar</td>
			<td>Daedalus Innovations LLC</td>
			<td>NMRCELL-B</td>
			<td></td>
		</tr>
		<tr>
			<td>Sparky3</td>
			<td>University of California San Francisco, CA</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>Xtreme-60 Syringe pump</td>
			<td>Daedalus Innovations LLC</td>
			<td>XTREME-60</td>
			<td></td>
		</tr>
	</tbody>
</table>

high-pressure nmr experiments for detecting protein low-lying conformational states

High-pressure is a well-known perturbation method that can be used to destabilize globular proteins and dissociate protein complexes in a reversible manner. Hydrostatic pressure drives thermodynamical equilibria toward the state(s) with the lower molar volume. Increasing pressure offers, therefore, the opportunities to finely tune the stability of globular proteins and the oligomerization equilibria of protein complexes. High-pressure NMR experiments allow a detailed characterization of the factors governing the stability of globular proteins, their folding mechanisms, and oligomerization mechanisms by combining the fine stability tuning ability of pressure perturbation and the site resolution offered by solution NMR spectroscopy. Here we present a protocol to probe the local folding stability of a protein via a set of 2D 1H-15N experiments recorded from 1 bar to 2.5 kbar. The steps required for the acquisition and analysis of such experiments are illustrated with data acquired on the RRM2 domain of hnRNPA1.

The protocol described here was used to probe the pressure dependence of RRM2, the second RNA recognition motif of hnRNPA1 (residues 95-106), which is almost completely unfolded within the 2.5 kbar range (&#62;90%). 1H-15N spectra were collected at 1 bar, 500 bar, 750 bar, 1 kbar, 1.5 kbar, 2 kbar, and 2.5 kbar&#160;(Figure 2). Since none of the native crosspeaks were visible above the noise level at 2.5 kbar, all corresponding residues were attributed an intensity valu...

Watch this Scientific Journal Video about High-Pressure NMR Experiments for Detecting Protein Low-Lying Conformational States at JoVE.com

High-Pressure NMR Experiments for Detecting Protein Low-Lying Conformational States

We provide a detailed description of the steps required to assemble a high-pressure cell, set up and record high-pressure NMR experiments, and finally analyze both peak intensity and chemical shift changes under pressure. These experiments can provide valuable insights into the folding pathways and structural stability of proteins.

High-pressure is a well-known perturbation method that can be used to destabilize globular proteins and dissociate protein complexes in a reversible ...

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2.6K Views.  Iowa State University. L'alta pressione è il metodo di scelta per modificare la popolazione relativa di un campione conformazionale proteico. È estremamente utile per isolare e caratterizzare stati conformazionali ad alta energia. Rispetto a tutti i metodi di perturbazione, come pH o temperatura, la perturbazione a pressione è facile da applicare e completamente reversibile.È anche una perturbazione locale, che colpisce principalmente le regioni con cavità di vetro o volumi vuoti. Per iniziare, introdur...

Video: High-Pressure NMR Experiments for Detecting Protein Low-Lying Conformational States

Using Scaffold Liposomes to Reconstitute Lipid-proximal Protein-protein Interactions In Vitro

Studies of integral membrane proteins in vitro are frequently complicated by the presence of a hydrophobic transmembrane domain. Further complicating these studies, reincorporation of detergent-solubilized membrane proteins into liposomes is a stochastic process where protein topology is impossible to enforce. This paper offers an alternative method to these challenging techniques that utilizes a liposome-based scaffold. Protein solubility is enhanced by deletion of the transmembrane domain, and these amino acids are replaced with a tethering moiety, such as a His-tag. This tether interacts with an anchoring group (Ni2+ coordinated by nitrilotriacetic acid (NTA(Ni2+)) for His-tagged proteins), which enforces a uniform protein topology at the surface of the liposome. An example is presented wherein the interaction between Dynamin-related protein 1 (Drp1) with an integral membrane protein, Mitochondrial Fission Factor (Mff), was investigated using this scaffold liposome method. In this work, we have demonstrated the ability of Mff to efficiently recruit soluble Drp1 to the surface of liposomes, which stimulated its GTPase activity. Moreover, Drp1 was able to tubulate the Mff-decorated lipid template in the presence of specific lipids. This example demonstrates the effectiveness of scaffold liposomes using structural and functional assays and highlights the role of Mff in regulating Drp1 activity.

Using Scaffold Liposomes to Reconstitute Lipid-proximal Protein-protein Interactions In Vitro

This paper describes a method for assessing the interactions and assemblies of integral membrane proteins in vitro with various partner factors in a lipid-proximal environment.

Studies of integral membrane proteins in vitro are frequently complicated by the presence of a hydrophobic transmembrane domain. Further complicating ...

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

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.

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

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.

The biological small-angle neutron scattering instrument at the High-Flux Isotope Reactor of Oak Ridge National Laboratory is dedicated to the ...

A Fluorescence Fluctuation Spectroscopy Assay of Protein-Protein Interactions at Cell-Cell Contacts

A variety of biological processes involves cell-cell interactions, typically mediated by proteins that interact at the interface between neighboring cells. Of interest, only few assays are capable of specifically probing such interactions directly in living cells. Here, we present an assay to measure the binding of proteins expressed at the surfaces of neighboring cells, at cell-cell contacts. This assay consists of two steps: mixing of cells expressing the proteins of interest fused to different fluorescent proteins, followed by fluorescence fluctuation spectroscopy measurements at cell-cell contacts using a confocal laser scanning microscope. We demonstrate the feasibility of this assay in a biologically relevant context by measuring the interactions of the amyloid precursor-like protein 1 (APLP1) across cell-cell junctions. We provide detailed protocols on the data acquisition using fluorescence-based techniques (scanning fluorescence cross-correlation spectroscopy, cross-correlation number and brightness analysis) and the required instrument calibrations. Further, we discuss critical steps in the data analysis and how to identify and correct external, spurious signal variations, such as those due to photobleaching or cell movement.
In general, the presented assay is applicable to any homo- or heterotypic protein-protein interaction at cell-cell contacts, between cells of the same or different types and can be implemented on a commercial confocal laser scanning microscope. An important requirement is the stability of the system, which needs to be sufficient to probe diffusive dynamics of the proteins of interest over several minutes.

This protocol describes a fluorescence fluctuation spectroscopy-based approach to investigate interactions among proteins mediating cell-cell interactions, i.e. proteins localized in cell junctions, directly in living cells. We provide detailed guidelines on instrument calibration, data acquisition and analysis, including corrections to possible artefact sources.

A variety of biological processes involves cell-cell interactions, typically mediated by proteins that interact at the interface between neighboring ...

High Sensitivity Measurement of Transcription Factor-DNA Binding Affinities by Competitive Titration Using Fluorescence Microscopy

Accurate quantification of transcription factor (TF)-DNA interactions is essential for understanding the regulation of gene expression. Since existing approaches suffer from significant limitations, we have developed a new method for determining TF-DNA binding affinities with high sensitivity on a large scale. The assay relies on the established fluorescence anisotropy (FA) principle but introduces important technical improvements. First, we measure a full FA competitive titration curve in a single well by incorporating TF and a fluorescently labeled reference DNA in a porous agarose gel matrix. Unlabeled DNA oligomer is loaded on the top as a competitor and, through diffusion, forms a spatio-temporal gradient. The resulting FA gradient is then read out using a customized epifluorescence microscope setup. This improved setup greatly increases the sensitivity of FA signal detection, allowing both weak and strong binding to be reliably quantified, even for molecules of similar molecular weights. In this fashion, we can measure one titration curve per well of a multi-well plate, and through a fitting procedure, we can extract both the absolute dissociation constant (KD) and active protein concentration. By testing all single-point mutation variants of a given consensus binding sequence, we can survey the entire binding specificity landscape of a TF, typically on a single plate. The resulting position weight matrices (PWMs) outperform those derived from other methods in predicting in vivo TF occupancy. Here, we present a detailed guide for implementing HiP-FA on a conventional automated fluorescent microscope and the data analysis pipeline.

Here we present a novel method for determining binding affinities at equilibrium and in solution with high sensitivity on a large scale. This improves the quantitative analysis of transcription factor-DNA binding. The method is based on automated fluorescence anisotropy measurements in a controlled delivery system.

Accurate quantification of transcription factor (TF)-DNA interactions is essential for understanding the regulation of gene expression. Since existing ...

Identification of Functional Protein Regions Through Chimeric Protein Construction

The goal of this protocol encompasses the design of chimeric proteins in which distinct regions of a protein are replaced by their corresponding sequences in a structurally similar protein, in order to determine the functional importance of these regions. Such chimeras are generated by means of a nested PCR protocol using overlapping DNA fragments and adequately designed primers, followed by their expression within a mammalian system to ensure native secondary structure and post-translational modifications. 
The functional role of a distinct region is then indicated by a loss of activity of the chimera in an appropriate readout assay. In consequence, regions harboring a set of critical amino acids are identified, which can be further screened by complementary techniques (e.g. site-directed mutagenesis) to increase molecular resolution. Although limited to cases in which a structurally related protein with differing functions can be found, chimeric proteins have been successfully employed to identify critical binding regions in proteins such as cytokines and cytokine receptors. This method is particularly suitable in cases in which the protein's functional regions are not well defined, and constitutes a valuable first step in directed evolution approaches to narrow down the regions of interest and reduce the screening effort involved.

Structurally related proteins frequently exert distinct biological functions. The exchange of equivalent regions of these proteins in order to create chimeric proteins constitutes an innovative approach to identify critical protein regions that are responsible for their functional divergence.

The goal of this protocol encompasses the design of chimeric proteins in which distinct regions of a protein are replaced by their corresponding sequences ...

Single-Molecule Tracking Microscopy - A Tool for Determining the Diffusive States of Cytosolic Molecules

Single-molecule localization microscopy probes the position and motions of individual molecules in living cells with tens of nanometer spatial and millisecond temporal resolution. These capabilities make single-molecule localization microscopy ideally suited to study molecular level biological functions in physiologically relevant environments. Here, we demonstrate an integrated protocol for both acquisition and processing/analysis of single-molecule tracking data to extract the different diffusive states a protein of interest may exhibit. This information can be used to quantify molecular complex formation in living cells. We provide a detailed description of a camera-based 3D single-molecule localization experiment, as well as the subsequent data processing steps that yield the trajectories of individual molecules. These trajectories are then analyzed using a numerical analysis framework to extract the prevalent diffusive states of the fluorescently labeled molecules and the relative abundance of these states. The analysis framework is based on stochastic simulations of intracellular Brownian diffusion trajectories that are spatially confined by an arbitrary cell geometry. Based on the simulated trajectories, raw single-molecule images are generated and analyzed in the same way as experimental images. In this way, experimental precision and accuracy limitations, which are difficult to calibrate experimentally, are explicitly incorporated into the analysis workflow. The diffusion coefficient and relative population fractions of the prevalent diffusive states are determined by fitting the distributions of experimental values using linear combinations of simulated distributions. We demonstrate the utility of our protocol by resolving the diffusive states of a protein that exhibits different diffusive states upon forming homo- and hetero-oligomeric complexes in the cytosol of a bacterial pathogen.

3D single-molecule localization microscopy is utilized to probe the spatial positions and motion trajectories of fluorescently labeled proteins in living bacterial cells. The experimental and data analysis protocol described herein determines the prevalent diffusive behaviors of cytosolic proteins based on pooled single-molecule trajectories.

Single-molecule localization microscopy probes the position and motions of individual molecules in living cells with tens of nanometer spatial and ...

Creating Highly Specific Chemically Induced Protein Dimerization Systems by Stepwise Phage Selection of a Combinatorial Single-Domain Antibody Library

Protein dimerization events that occur only in the presence of a small-molecule ligand enable the development of small-molecule biosensors for the dissection and manipulation of biological pathways. Currently, only a limited number of chemically induced dimerization (CID) systems exist and engineering new ones with desired sensitivity and selectivity for specific small-molecule ligands remains a challenge in the field of protein engineering. We here describe a high throughput screening method, combinatorial binders-enabled selection of CID (COMBINES-CID), for the de novo engineering of CID systems applicable to a large variety of ligands. This method uses the two-step selection of a phage-displayed combinatorial nanobody library to obtain 1) &#34;anchor binders&#34; that first bind to a ligand of interest and then 2) &#34;dimerization binders&#34; that only bind to anchor binder-ligand complexes. To select anchor binders, a combinatorial library of over 109 complementarity-determining region (CDR)-randomized nanobodies is screened with a biotinylated ligand and hits are validated with the unlabeled ligand by bio-layer interferometry (BLI). To obtain dimerization binders, the nanobody library is screened with anchor binder-ligand complexes as targets for positive screening and the unbound anchor binders for negative screening. COMBINES-CID is broadly applicable to select CID binders with other immunoglobulin, non-immunoglobulin, or computationally designed scaffolds to create biosensors for in vitro and in vivo detection of drugs, metabolites, signaling molecules, etc.

Creating chemically induced protein dimerization systems with desired affinity and specificity for any given small molecule ligand would have many biological sensing and actuation applications. Here, we describe an efficient, generalizable method for de novo engineering of chemically induced dimerization systems via the stepwise selection of a phage-displayed combinatorial single-domain antibody library.

Protein dimerization events that occur only in the presence of a small-molecule ligand enable the development of small-molecule biosensors for the ...

Native Cell Membrane Nanoparticles System for Membrane Protein-Protein Interaction Analysis

Protein-protein interactions in cell membrane systems play crucial roles in a wide range of biological processes- from cell-to-cell interactions to signal transduction; from sensing environmental signals to biological response; from metabolic regulation to developmental control. Accurate structural information of protein-protein interactions is crucial for understanding the molecular mechanisms of membrane protein complexes and for the design of highly specific molecules to modulate these proteins. Many in vivo and in vitro approaches have been developed for the detection and analysis of protein-protein interactions. Among them the structural biology approach is unique in that it can provide direct structural information of protein-protein interactions at the atomic level. However, current membrane protein structural biology is still largely limited to detergent-based methods. The major drawback of detergent-based methods is that they often dissociate or denature membrane protein complexes once their native lipid bilayer environment is removed by detergent molecules. We have been developing a&#160;native cell membrane nanoparticle system for membrane protein structural biology. Here, we demonstrate the use of this system in the analysis of protein-protein interactions on the cell membrane with a case study of the oligomeric state of AcrB.

Presented here is a protocol for the determination of oligomeric state of membrane proteins that utilizes a native cell membrane nanoparticle system in conjunction with electron microscopy.

Protein-protein interactions in cell membrane systems play crucial roles in a wide range of biological processes- from cell-to-cell interactions to signal ...

Optimizing the Growth of Endothiapepsin Crystals for Serial Crystallography Experiments

Here, a protocol is presented to facilitate the creation of large volumes (&#62; 100 &#181;L) of micro-crystalline slurries suitable for serial crystallography experiments at both synchrotrons and XFELs. The method is based upon an understanding of the protein crystal phase diagram, and how that knowledge can be utilized. The method is divided into three stages: (1) optimizing crystal morphology, (2) transitioning to batch, and (3) scaling. Stage 1 involves finding well diffracting, single crystals, hopefully but not necessarily, presenting in a cube-like morphology. In Stage 2, the Stage 1 condition is optimized by crystal growth time. This strategy can transform crystals grown by vapor diffusion to batch. Once crystal growth can occur within approximately 24 h, a morphogram of the protein and precipitant mixture can be plotted and used as the basis for a scaling strategy (Stage 3). When crystals can be grown in batch, scaling can be attempted, and the crystal size and concentration optimized as the volume is increased. Endothiapepsin has been used as a demonstration protein for this protocol. Some of the decisions presented are specific to endothiapepsin. However, it is hoped that the way they have been applied will inspire a way of thinking about this procedure that others can adapt to their own projects.

The aim of this article is to give the viewer a solid understanding of how to transform their small-volume, vapor-diffusion protocol, for growing large, single protein crystals, into a large-volume batch micro-crystallization method for serial crystallography.

Here, a protocol is presented to facilitate the creation of large volumes (&#62; 100 &#181;L) of micro-crystalline slurries suitable for serial ...

NMR-Based Fragment Screening in a Minimum Sample but Maximum Automation Mode

Fragment-based screening (FBS) is a well-validated and accepted concept within the drug discovery process both in academia and industry. The greatest advantage of NMR-based fragment screening is its ability not only to detect binders over 7-8 orders of magnitude of affinity but also to monitor purity and chemical quality of the fragments and thus to produce high quality hits and minimal false positives or false negatives. A prerequisite within the FBS is to perform initial and periodic quality control of the fragment library, determining solubility and chemical integrity of the fragments in relevant buffers, and establishing multiple libraries to cover diverse scaffolds to accommodate various macromolecule target classes (proteins/RNA/DNA). Further, an extensive NMR-based screening protocol optimization with respect to sample quantities, speed of acquisition and analysis at the level of biological construct/fragment-space, in condition-space (buffer, additives, ions, pH, and temperature) and in ligand-space (ligand analogues, ligand concentration) is required. At least in academia, these screening efforts have so far been undertaken manually in a very limited fashion, leading to limited availability of screening infrastructure not only in the drug development process but also in the context of chemical probe development. In order to meet the requirements economically, advanced workflows are presented. They take advantage of the latest state-of-the-art advanced hardware, with which the liquid sample collection can be filled in a temperature-controlled fashion into the NMR-tubes in an automated manner. 1H/19F NMR ligand-based spectra are then collected at a given temperature. High-throughput sample changer (HT sample changer) can handle more than 500 samples in temperature-controlled blocks. This together with advanced software tools speeds up data acquisition and analysis. Further, application of screening routines on protein and RNA samples are described to make aware of the established protocols for a broad user base in biomacromolecular research.

Fragment-based screening by NMR is a robust method to rapidly identify small molecule binders to biomacromolecules (DNA, RNA, or proteins). Protocols describing automation-based sample preparation, NMR experiments &#38; acquisition conditions, and analysis workflows are presented. The technique allows for optimal exploitation of both 1H and 19F NMR-active nuclei for detection.

Fragment-based screening (FBS) is a well-validated and accepted concept within the drug discovery process both in academia and industry. The greatest ...