We acknowledge research funding from the National Institute of General Medical Sciences grant R43GM125420-01to support commercial development of a benchtop FPOP device and R01GM127267 for the development of standardization and dosimetry protocols for high-energy FPOP.

Joshua S. Sharp discloses a significant financial interest in GenNext Technologies, Inc., a small company seeking to commercialize technologies for protein higher order structure analysis including hydroxyl radical protein footprinting.

Mass spectrometry-based structural techniques, including hydrogen-deuterium exchange, chemical cross-linking, covalent labeling, and native spray mass spectrometry and ion mobility have been rapidly growing in popularity due to their flexibility, sensitivity, and ability to handle complex mixtures. FPOP boasts several advantages that has boosted its popularity in the area of mass spectrometry-based structural techniques. Like most covalent labeling strategies, it provides a stable chemical snapshot of protein topography ...

Fast photochemical oxidation of proteins (FPOP) is an emerging technique for the determination of protein topographical changes by ultra-fast covalent modification of the solvent-exposed surface area of proteins followed by detection by LC-MS1. FPOP generates a high concentration of hydroxyl radicals in situ by UV laser flash photolysis of hydrogen peroxide. These hydroxyl radicals are very reactive and short lived, consumed on roughly a microsecond timescale under FPOP conditions2. These hydroxyl radicals diffuse through water and oxidize various organic components in solution at kinetic rates generally ranging from fast (~106 M-1 s-1) to diffusion-controlled3. When the hydroxyl radical encounters a protein surface, the radical will oxidize the amino acid side chains on the protein surface, resulting in a mass shift of that amino acid (most commonly the net addition of one oxygen atom)4. The rate of the oxidation reaction at any amino acid depends on two factors: the inherent reactivity of that amino acid (which depends on the side chain and the sequence context)4,5 and the accessibility of that side chain to the diffusing hydroxyl radical, which closely correlates to the average solvent accessible surface area6,7. All of the standard amino acids except glycine have been observed as labeled by these highly reactive hydroxyl radicals in FPOP experiments, albeit at widely differing yields; in practice, Ser, Thr, Asn, and Ala are rarely seen as oxidized in most samples except under high radical doses and identified by careful and sensitive targeted ETD fragmentation8,9. After oxidation, samples are quenched to remove hydrogen peroxide and secondary oxidants (superoxide, singlet oxygen, peptidyl hydroperoxides, etc.) The quenched samples are then proteolytically digested to generate mixtures of oxidized peptides, where the structural information is frozen as a chemical &#8220;snapshot&#8221; in the patterns of oxidation products of the various peptides (Figure 1). Liquid chromatography coupled to mass spectrometry (LC-MS) is used to measure the amount of oxidation of amino acids in a given proteolytic peptide based on the relative intensities of the oxidized and unoxidized versions of that peptide. By comparing this oxidative footprint of the same protein obtained under different conformational conditions (e.g., ligand-bound versus ligand-free), differences in the amount of oxidation of a given region of the protein can be directly correlated with differences in the solvent-accessible surface area of that region6,7. The ability to provide protein topographical information makes FPOP an attractive technology for the higher-order structure determination of proteins, including in protein therapeutic discovery and development10,11.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/61580/61580fig01.jpg" /> 
Figure 1: Overview of FPOP.&#160;The surface of the protein is covalently modified by highly reactive hydroxyl radicals. The hydroxyl radicals will react with amino acid side chains of the protein at a rate that is strongly influenced by the solvent accessibility of the side chain. Topographical changes (for example, due to the binding of a ligand as shown above) will protect amino acids in the region of interaction from reacting with hydroxyl radicals, resulting in a decrease in the intensity of modified peptide in the LC-MS signal. <a href="https://www.jove.com/files/ftp_upload/61580/61580fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Different constituents present in the FPOP solution (e.g., ligands, excipients, buffers) have different scavenging activity towards the hydroxyl radicals generated upon the laser photolysis of hydrogen peroxide3. Similarly, a small change in peroxide concentration, laser fluence, and buffer composition may change the effective radical dose, making the reproduction of FPOP data challenging across the samples and between different labs. Therefore, it is important to be able to compare the hydroxyl radical dose available to react with protein in each sample using one of several available hydroxyl radical dosimeters12,13,14,15,16. Hydroxyl radical dosimeters act by competing with the analyte (and with all scavengers in solution) for the pool of hydroxyl radicals; the effective dose of hydroxyl radicals is measured by measuring the amount of oxidation of the dosimeter. Note that &#8220;effective hydroxyl radical dose&#8221; is a function of both the initial concentration of hydroxyl radical generated and the half-life of the radical. These two parameters are partially dependent on one another, making the theoretical kinetic modeling somewhat complex (Figure 2). Two samples could have wildly different initial radical half-lives while still maintaining the same effective radical dose by changing the initial concentration of hydroxyl radical formed; they will still generate identical footprints17. Adenine13 and Tris12 are convenient hydroxyl radical dosimeters because their level of oxidation can be measured by UV spectroscopy in real-time, allowing for researchers to quickly identify when there is a problem with effective hydroxyl radical dose and to troubleshoot their problem. To solve this issue, an inline dosimeter located in the flow system directly after the site of irradiation that can monitor the signal from adenine absorbance changes in real-time is important. This helps in carrying out FPOP experiments in buffers or any other excipient with widely differing levels of hydroxyl radical scavenging capacity17. This radical dosage compensation can be performed in real-time, yielding statistically indistinguishable results for the same conformer by adjusting the effective radical dose.
In this protocol, we have detailed procedures for performing a typical FPOP experiment with radical dosage compensation using adenine as an internal optical radical dosimeter. This method allows investigators to compare footprints across FPOP conditions that have different scavenging capacity by performing compensation in real-time.
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/61580/61580fig02.jpg" /> 
Figure 2: Kinetic simulation of dosimetry-based compensation.&#160;1 mM adenine dosimeter response is measured in 5 &#181;M lysozyme analyte with a 1 mM initial hydroxyl radical concentration (&#9642;OH t1/2=53 ns), and set as a target dosimeter response (black). Upon the addition of 1 mM of the scavenger excipient histidine, the dosimeter response (blue) decreases along with the amount of protein oxidation in a proportional manner (cyan). The half-life of the hydroxyl radical also decreases (&#9642;OH t1/2=39 ns). When the amount of hydroxyl radical generated is increased to give an equivalent yield of oxidized dosimeter in the sample with 1 mM histidine scavenger as achieved with 1 mM hydroxyl radical in the absence of scavenger (red), the amount of protein oxidation that occurs similarly becomes identical (magenta), while the hydroxyl radical half-life decreases even further (&#9642;OH t1/2=29 ns). Adapted with permission from Sharp J.S., Am Pharmaceut Rev 22, 50-55, 2019. <a href="https://www.jove.com/files/ftp_upload/61580/61580fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

<table><tbody><tr><td>Adenine</td><td>Acros Organics</td><td>147440250</td><td>Soluble in water upto 3.5 mM</td></tr><tr><td>Aperture</td><td>Edmund Optics</td><td>39-905</td><td>1000 &amp;mu;m Aperture Diameter, Gold-Plated Copper Aperture</td></tr><tr><td>Aperture holder</td><td>Edmund Optics</td><td>53-287</td><td>25.8mm Outer Diameter, Precision Pinhole Mount</td></tr><tr><td>Catalse</td><td>Sigma Aldrich</td><td>C-40</td><td>Catalase from bovine liver, lyophilized powder, &amp;ge;10,000 units/mg protein</td></tr><tr><td>COMPex Pro laser</td><td>Coherent</td><td>1113836</td><td>COMPexPRO 102, F-Vversion, KrF laser, No XeCl</td></tr><tr><td>Dithiotheitol (DTT)</td><td>Promega</td><td>V3151</td><td>DTT, Molecular Grade (DL-Dithiothreitol)</td></tr><tr><td>Fraction collector</td><td>GenNext Technologies, Inc.</td><td>N/A</td><td>Automated fraction collector</td></tr><tr><td>Fused silica capillay</td><td>Molex</td><td>1068150023</td><td>Polymicro Flexible Fused Silica Capillary Tubing, Inner Diameter 100 &amp;micro;m, Outer Diameter 375 &amp;micro;m, TSP100375</td></tr><tr><td>Glutamine</td><td>Acros Organics</td><td>119951000</td><td>L(+)-Glutamine, 99%</td></tr><tr><td>Holder for lens</td><td>Edmund Optics</td><td>03-668</td><td>53 mm Outer Diameter, Three-Screw Adjustable Ring Mount</td></tr><tr><td>Hydrogen peroxide</td><td>Fisher Scientific</td><td>H325-100</td><td>Hydrogen Peroxide, 30% (Certified ACS), Fisher Chemical</td></tr><tr><td>LC-MS/MS system</td><td>Thermo Scientific</td><td>IQLAAEGAAPFADBMBCX</td><td>Dionex Ultimate 3000 coupled to Orbitap Fusion Tribrid mass spectrometer</td></tr><tr><td>Mas spec grade Acetonitrile</td><td>Fisher Scientific</td><td>A955-1</td><td>Acetonitrile, Optima LC/MS Grade, Fisher Chemical</td></tr><tr><td>Mass spec grade formic acid</td><td>Fisher Scientific</td><td>A117-50</td><td>Formic Acid, 99.0+%, Optima&amp;trade; LC/MS Grade, Fisher Chemical</td></tr><tr><td>Mass spec grade water</td><td>Fisher Scientific</td><td>W6-4</td><td>Water, Optima LC/MS Grade, Fisher Chemical</td></tr><tr><td>MES buffer</td><td>Sigma Aldrich</td><td>M0164</td><td>MES hemisodium salt</td></tr><tr><td>Methionine amide</td><td>Bachem</td><td>4000594.0005</td><td>H-met-NH&lt;sub&gt;2&lt;/sub&gt;.HCl</td></tr><tr><td>Micro V clamp</td><td>Thor Labs</td><td>VK250</td><td>Micro V-clamp with stainless steel blades</td></tr><tr><td>Motorized stage</td><td>Edmund Optics</td><td>68-638</td><td>50mm Travel Motorized Stage System with Manual Control</td></tr><tr><td>Nano C18 colum</td><td>Thermo Scientific</td><td>164534</td><td>Acclaim PepMap 100 C18 HPLC Columns</td></tr><tr><td>Optical bench</td><td>Edmund Optics</td><td>56-935</td><td>18&quot; x 18&quot; breadboard</td></tr><tr><td>Pioneer FPOP Module System</td><td>GenNext Technologies, Inc.</td><td>N/A</td><td>Inline FPOP Radical Dosimetry System</td></tr><tr><td>Post holder</td><td>Edmund Optics</td><td>58-979</td><td>3&quot; Length, &amp;frac14;-20 Thread, Post Holder</td></tr><tr><td>Sodium phosphate dibasic</td><td>Fisher Scientific</td><td>BP331-500</td><td>Sodium Phosphate Dibasic Heptahydrate (Colorless-to-White Crystals), Fisher BioReagents</td></tr><tr><td>Sodium phosphate monobasic</td><td>Fisher Scientific</td><td>BP330-500</td><td>Sodium Phosphate Monobasic Monohydrate (Colorless-to-white Crystals), Fisher BioReagents</td></tr><tr><td>Syringe</td><td>Hamilton</td><td>81065</td><td>100 &amp;micro;L, Model 1710 RN SYR, Small Removable NDL, 22s ga, 2 in, point style 3</td></tr><tr><td>Syringe pump</td><td>KD Scientific</td><td>788101</td><td>Legato&amp;nbsp;101 syringe pump</td></tr><tr><td>Trap C18 column</td><td>Thermo Scientific</td><td>160454</td><td>Thermo Scientific&amp;nbsp;Acclaim PepMap 100 C18 HPLC Columns</td></tr><tr><td>Tris</td><td>Sigma Aldrich</td><td>252859</td><td>Tris(hydroxymethyl)aminomethane</td></tr><tr><td>Trypsin</td><td>Promega</td><td>V5111</td><td>Sequencing Grade Modified Trypsin</td></tr><tr><td>UV plano convex lens</td><td>Edmund Optics</td><td>84-285</td><td>30 mm Dia. x 120 mm FL Uncoated, UV Plano-Convex Lens</td></tr></tbody></table>

enabling real-time compensation in fast photochemical oxidations of proteins for the determination of protein topography changes

Fast photochemical oxidation of proteins (FPOP) is a mass spectrometry-based structural biology technique that probes the solvent-accessible surface area of proteins. This technique relies on the reaction of amino acid side chains with hydroxyl radicals freely diffusing in solution. FPOP generates these radicals in situ by laser photolysis of hydrogen peroxide, creating a burst of hydroxyl radicals that is depleted on the order of a microsecond. When these hydroxyl radicals react with a solvent-accessible amino acid side chain, the reaction products exhibit a mass shift that can be measured and quantified by mass spectrometry. Since the rate of reaction of an amino acid depends in part on the average solvent accessible surface of that amino acid, measured changes in the amount of oxidation of a given region of a protein can be directly correlated to changes in the solvent accessibility of that region between different conformations (e.g., ligand-bound versus ligand-free, monomer vs. aggregate, etc.) FPOP has been applied in a number of problems in biology, including protein-protein interactions, protein conformational changes, and protein-ligand binding. As the available concentration of hydroxyl radicals varies based on many experimental conditions in the FPOP experiment, it is important to monitor the effective radical dose to which the protein analyte is exposed. This monitoring is efficiently achieved by incorporating an inline dosimeter to measure the signal from the FPOP reaction, with laser fluence adjusted in real-time to achieve the desired amount of oxidation. With this compensation, changes in protein topography reflecting conformational changes, ligand-binding surfaces, and/or protein-protein interaction interfaces can be determined in heterogeneous samples using relatively low sample amounts.

Comparison of the heavy chain peptide footprint of the adalimumab biosimilar in phosphate buffer and when heated at 55 &#176;C for 1 h show interesting results. Student&#8217;s t-test is used for the identification of peptides that are significantly changed in these two conditions (p &#8804; 0.05). The peptides 20-38, 99-125, 215-222, 223-252, 260-278, 376-413, and 414-420 show significant protection from solvent when the protein is heated to form aggregates (Figure 5)30</s...

Watch this Scientific Journal Video about Enabling Real-Time Compensation in Fast Photochemical Oxidations of Proteins for the Determination of Protein Topography Changes at JoVE.com

Enabling Real-Time Compensation in Fast Photochemical Oxidations of Proteins for the Determination of Protein Topography Changes

Fast photochemical oxidation of proteins is an emerging technique for the structural characterization of proteins. Different solvent additives and ligands have varied hydroxyl radical scavenging properties. To compare the protein structure in different conditions, real-time compensation of hydroxyl radicals generated in the reaction is required to normalize reaction conditions.

Fast photochemical oxidation of proteins (FPOP) is a mass spectrometry-based structural biology technique that probes the solvent-accessible surface area ...

enabling-real-time-compensation-fast-photochemical-oxidations

Research

JoVE Journal

Biochemistry

5.2K Views.  University of Mississippi. De methode vergemakkelijkt het meten van eiwitbevestiging en eiwitinteracties. De buffersamenstelling, monsterzuiverheid en eiwitconcentratievereisten zijn flexibel waardoor de methode uitdagende structurele problemen kan aanpakken. Deze techniek maakt een compressie van monsters in een breed scala van verschillende buffersystemen.Met behulp van onze dosimetriemethode kunnen onderzoekers in realtime controleren hoe goed hun etiketteringsreacties werken. Een toepassing van deze technolo...

Video: Enabling Real-Time Compensation in Fast Photochemical Oxidations of Proteins for the Determination of Protein Topography Changes

Proton Transfer and Protein Conformation Dynamics in Photosensitive Proteins by Time-resolved Step-scan Fourier-transform Infrared Spectroscopy

Monitoring the dynamics of protonation and protein backbone conformation changes during the function of a protein is an essential step towards understanding its mechanism. Protonation and conformational changes affect the vibration pattern of amino acid side chains and of the peptide bond, respectively, both of which can be probed by infrared (IR) difference spectroscopy. For proteins whose function can be repetitively and reproducibly triggered by light, it is possible to obtain infrared difference spectra with (sub)microsecond resolution over a broad spectral range using the step-scan Fourier transform infrared technique. With ~102-103 repetitions of the photoreaction, the minimum number to complete a scan at reasonable spectral resolution and bandwidth, the noise level in the absorption difference spectra can be as low as ~10-4, sufficient to follow the kinetics of protonation changes from a single amino acid. Lower noise levels can be accomplished by more data averaging and/or mathematical processing. The amount of protein required for optimal results is between 5-100 &#181;g, depending on the sampling technique used. Regarding additional requirements, the protein needs to be first concentrated in a low ionic strength buffer and then dried to form a film. The protein film is hydrated prior to the experiment, either with little droplets of water or under controlled atmospheric humidity. The attained hydration level (g of water / g of protein) is gauged from an IR absorption spectrum. To showcase the technique, we studied the photocycle of the light-driven proton-pump bacteriorhodopsin in its native purple membrane environment, and of the light-gated ion channel channelrhodopsin-2 solubilized in detergent.

Key steps of protein function, in particular backbone conformational changes and proton transfer reactions, often take place in the microsecond to millisecond time scale. These dynamical processes can be studied by time-resolved step-scan Fourier-transform infrared spectroscopy, in particular for proteins whose function is triggered by light.

Monitoring the dynamics of protonation and protein backbone conformation changes during the function of a protein is an essential step towards ...

Engineering

Defining Hsp33's Redox-regulated Chaperone Activity and Mapping Conformational Changes on Hsp33 Using Hydrogen-deuterium Exchange Mass Spectrometry

Living organisms regularly need to cope with fluctuating environments during their life cycle, including changes in temperature, pH, the accumulation of reactive oxygen species, and more. These fluctuations can lead to a widespread protein unfolding, aggregation, and cell death. Therefore, cells have evolved a dynamic and stress-specific network of molecular chaperones, which maintain a "healthy" proteome during stress conditions. ATP-independent chaperones constitute one major class of molecular chaperones, which serve as first-line defense molecules, protecting against protein aggregation in a stress-dependent manner. One feature these chaperones have in common is their ability to utilize structural plasticity for their stress-specific activation, recognition, and release of the misfolded client. 
In this paper, we focus on the functional and structural analysis of one such intrinsically disordered chaperone, the bacterial redox-regulated Hsp33, which protects proteins against aggregation during oxidative stress. Here, we present a toolbox of diverse techniques for studying redox-regulated chaperone activity, as well as for mapping conformational changes of the chaperone, underlying its activity. Specifically, we describe a workflow which includes the preparation of fully reduced and fully oxidized proteins, followed by an analysis of the chaperone anti-aggregation activity in vitro using light-scattering, focusing on the degree of the anti-aggregation activity and its kinetics. To overcome frequent outliers accumulated during aggregation assays, we describe the usage of Kfits, a novel graphical tool which allows easy processing of kinetic measurements. This tool can be easily applied to other types of kinetic measurements for removing outliers and fitting kinetic parameters. To correlate the function with the protein structure, we describe the setup and workflow of a structural mass spectrometry technique, hydrogen-deuterium exchange mass spectrometry, that allows the mapping of conformational changes on the chaperone and substrate during different stages of Hsp33 activity. The same methodology can be applied to other protein-protein and protein-ligand interactions.

One of the most challenging stress conditions that organisms encounter during their lifetime involves the accumulation of oxidants. During oxidative stress, cells heavily rely on molecular chaperones. Here, we present methods used to investigate the redox-regulated anti-aggregation activity, as well as to monitor structural changes governing the chaperone function using HDX-MS.

Living organisms regularly need to cope with fluctuating environments during their life cycle, including changes in temperature, pH, the accumulation of ...

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 ...

Characterizing Cellular Proteins with In-cell Fast Photochemical Oxidation of Proteins

Fast photochemical oxidation of proteins (FPOP) is a hydroxyl radical protein footprinting method used to characterize protein structure and interactions. FPOP uses a 248 nm excimer laser to photolyze hydrogen peroxide producing hydroxyl radicals. These radicals oxidatively modify solvent exposed side chains of 19 of the 20 amino acids. Recently, this method has been used in live cells (IC-FPOP) to study protein interactions in their native environment. The study of proteins in cells accounts for intermolecular crowding and various protein interactions that are disrupted for in vitro studies. A custom single cell flow system was designed to reduce cell aggregation and clogging during IC-FPOP. This flow system focuses the cells past the excimer laser individually, thus ensuring consistent irradiation. By comparing the extent of oxidation produced from FPOP to the protein&#39;s solvent accessibility calculated from a crystal structure, IC-FPOP can accurately probe the solvent accessible side chains of proteins.

Here, we characterize protein structure and interaction sites in living cells using a protein footprinting technique termed in-cell fast photochemical oxidation of proteins (IC-FPOP).

Fast photochemical oxidation of proteins (FPOP) is a hydroxyl radical protein footprinting method used to characterize protein structure and interactions. ...

Laser-free Hydroxyl Radical Protein Footprinting to Perform Higher Order Structural Analysis of Proteins

Hydroxyl Radical Protein Footprinting (HRPF) is an emerging and promising higher order structural analysis technique that provides information on changes in protein structure, protein-protein interactions, or protein-ligand interactions. HRPF utilizes hydroxyl radicals (&#9642;OH) to irreversibly label a protein&#8217;s solvent accessible surface. The inherent complexity, cost, and hazardous nature of performing HRPF have substantially limited broad-based adoption in biopharma. These factors include: 1) the use of complicated, dangerous, and expensive lasers that demand substantial safety precautions; and 2) the irreproducibility of HRPF caused by background scavenging of &#9642;OH that limit comparative studies. This publication provides a protocol for operation of a laser-free HRPF system. This laser-free HRPF system utilizes a high energy, high-pressure plasma light source flash oxidation technology with in-line radical dosimetry. The plasma light source is safer, easier to use, and more efficient in generating hydroxyl radicals than laser-based HRPF systems, and the in-line radical dosimeter increases the reproducibility of studies. Combined, the laser-free HRPF system addresses and surmounts the mentioned shortcomings and limitations of laser-based techniques.

This protocol presents a method to use inline radical dosimetry and a plasma light source to perform flash oxidation protein footprinting. This method replaces the hazardous UV laser to simplify and improve the reproducibility of fast photochemical oxidation of protein studies.

Hydroxyl Radical Protein Footprinting (HRPF) is an emerging and promising higher order structural analysis technique that provides information on changes ...

Platform Incubator with Movable XY Stage: A New Platform for Implementing In-Cell Fast Photochemical Oxidation of Proteins

Fast Photochemical Oxidation of proteins (FPOP) coupled with mass spectrometry (MS) has become an invaluable tool in structural proteomics to interrogate protein interactions, structure, and protein conformational dynamics as a function of solvent accessibility. In recent years, the scope of FPOP, a hydroxyl radical protein foot printing (HRPF) technique, has been expanded to protein labeling in live cell cultures, providing the means to study protein interactions in the convoluted cellular environment. In-cell protein modifications can provide insight into ligand induced structural changes or conformational changes accompanying protein complex formation, all within the cellular context. Protein footprinting has been accomplished employing a customary flow-based system and a 248 nm KrF excimer laser to yield hydroxyl radicals via photolysis of hydrogen peroxide, requiring 20 minutes of analysis for one cell sample.To facilitate time-resolved FPOP experiments, the use of a new 6-well plate-based IC-FPOP platform was pioneered. In the current system, a single laser pulse irradiates one entire well, which truncates the FPOP experimental time frame resulting in 20 seconds of analysis time, a 60-fold decrease. This greatly reduced analysis time makes it possible to research cellular mechanisms such as biochemical signaling cascades, protein folding, and differential experiments (i.e., drug-free vs. drug bound) in a time-dependent manner. This new instrumentation, entitled Platform Incubator with Movable XY Stage (PIXY), allows the user to perform cell culture and IC-FPOP directly on the optical bench using a platform incubator with temperature, CO2 and humidity control. The platform also includes a positioning stage, peristaltic pumps, and mirror optics for laser beam guidance. IC-FPOP conditions such as optics configuration, flow rates, transient transfections, and H2O2 concentration in PIXY have been optimized and peer-reviewed. Automation of all components of the system will reduce human manipulation and increase throughput.

A new static platform is used to characterize protein structure and interaction sites in the native cell environment utilizing a protein footprinting technique called in-cell fast photochemical oxidation of proteins (IC-FPOP).

Fast Photochemical Oxidation of proteins (FPOP) coupled with mass spectrometry (MS) has become an invaluable tool in structural proteomics to interrogate ...

Resin-Assisted Capture Coupled with Isobaric Tandem Mass Tag Labeling for Multiplexed Quantification of Protein Thiol Oxidation

Reversible oxidative modifications on protein thiols have recently emerged as important mediators of cellular function. Herein we describe the detailed procedure of a quantitative redox proteomics method that utilizes resin-assisted capture (RAC) in combination with tandem mass tag (TMT) isobaric labeling and liquid chromatography-tandem mass spectrometry (LC-MS/MS) to allow multiplexed stochiometric quantification of oxidized protein thiols at the proteome level. The site-specific quantitative information on oxidized cysteine residues provides additional insight into the functional impacts of such modifications. 
The workflow is adaptable across many sample types, including cultured cells (e.g., mammalian, prokaryotic) and whole tissues (e.g., heart, lung, muscle), which are initially lysed/homogenized and with free thiols being alkylated to prevent artificial oxidation. The oxidized protein thiols are then reduced and captured by a thiol-affinity resin, which streamlines and simplifies the workflow steps by allowing the proceeding digestion, labeling, and washing procedures to be performed without additional transfer of proteins/peptides. Finally, the labeled peptides are eluted and analyzed by LC-MS/MS to reveal comprehensive stoichiometric changes related to thiol oxidation across the entire proteome. This method greatly improves the understanding of the role of redox-dependent regulation under physiological and pathophysiological states related to protein thiol oxidation.

Protein thiol oxidation has significant implications under normal physiological and pathophysiological conditions. We describe the details of a quantitative redox proteomics method, which utilizes resin-assisted capture, isobaric labeling, and mass spectrometry, enabling site-specific identification and quantification of reversibly oxidized cysteine residues of proteins.

Reversible oxidative modifications on protein thiols have recently emerged as important mediators of cellular function. Herein we describe the detailed ...

F&#246;rster Resonance Energy Transfer Mapping: A New Methodology to Elucidate Global Structural Features

F&#246;rster resonance energy transfer (FRET) is an established fluorescence-based method used to successfully measure distances in and between biomolecules in vitro as well as within cells. In FRET, the efficiency of energy transfer, measured by changes in fluorescence intensity or lifetime, relates to the distance between two fluorescent molecules or labels. Determination of dynamics and conformational changes from the distances are just some examples of applications of this method to biological systems. Under certain conditions, this methodology can add to and enhance existing X-ray crystal structures by providing information regarding dynamics, flexibility, and adaptation to binding surfaces. We describe the use of FRET and associated distance determinations to elucidate structural properties, through the identification of a binding site or the orientations of dimer subunits. Through judicious choice of labeling sites, and often employment of multiple labeling strategies, we have successfully applied these mapping methods to determine global structural properties in a protein-DNA complex and the SecA-SecYEG protein translocation system. In the SecA-SecYEG system, we have used FRET mapping methods to identify the preprotein-binding site and determine the local conformation of the bound signal sequence region. This study outlines the steps for performing FRET mapping studies, including identification of appropriate labeling sites, discussion of possible labels including non-native amino acid residues, labeling procedures, how to perform measurements, and interpreting the data.

The study details the methodology of FRET mapping including the selection of labeling sites, choice of dyes, acquisition, and data analysis. This methodology is effective at determining binding sites, conformational changes, and dynamic motions in protein systems and is most useful if performed in conjunction with existing 3-D structural information.

F&#246;rster resonance energy transfer (FRET) is an established fluorescence-based method used to successfully measure distances in and between ...

Automated Two-dimensional Spatiotemporal Analysis of Mobile Single-molecule FRET Probes

Single-molecule F&#246;rster resonance energy transfer (smFRET) is a versatile technique reporting on distances in the sub-nanometer to nanometer range. It has been used in a wide range of biophysical and molecular biological experiments, including the measurement of molecular forces, characterization of conformational dynamics of biomolecules, observation of intracellular colocalization of proteins, and determination of receptor-ligand interaction times. In a widefield microscopy configuration, experiments are typically performed using surface-immobilized probes. Here, a method combining single-molecule tracking with alternating excitation (ALEX) smFRET experiments is presented, permitting the acquisition of smFRET time traces of surface-bound, yet mobile probes in plasma membranes or glass-supported lipid bilayers. For the analysis of recorded data, an automated, open-source software collection was developed supporting (i) the localization of fluorescent signals, (ii) single-particle tracking, (iii) determination of FRET-related quantities including correction factors, (iv) stringent verification of smFRET traces, and (v) intuitive presentation of the results. The generated data can conveniently be used as input for further exploration via specialized software, e.g., for the assessment of the diffusional behavior of probes or the investigation of FRET transitions.

This article presents a method for spatiotemporal analysis of mobile, single-molecule F&#246;rster resonance energy transfer (smFRET)-based probes using widefield fluorescence microscopy. The newly developed software toolkit allows the determination of smFRET time traces of moving probes, including the correct FRET efficiency and the molecular positions, as functions of time.

Single-molecule F&#246;rster resonance energy transfer (smFRET) is a versatile technique reporting on distances in the sub-nanometer to nanometer range. ...

Bioengineering

Time-Resolved Forster Resonance Energy Transfer for Monitoring Protein Phosphorylation in Cells

To perform the assay, dispense 50 microliters of cells at the pre-optimized density, into a 96-well tissue culture-treated plate in the appropriate culture medium. Then, incubate them overnight in a 37 degrees Celsius and 5% carbon dioxide incubator.
The following day, prepare intermediate two-fold and four-fold dilution series of the test compounds, by serially diluting the compound in a serum-free medium, across 12 wells of a polypropylene 96-well plate. For cell stimulation, add 50 microliters of serum-free medium, containing the stimulator at a 2X concentration, and incubate the cells for the pre-optimized time at either room temperature or 37 degrees.
For cell inhibition, add 25 microliters of serum-free medium containing the inhibitor at a 4X concentration, and incubate the cells for the pre-optimized time at either room temperature or 37 degrees. Then, add 25 microliters of serum-free medium containing the stimulator at a 4X concentration, and incubate for the pre-optimized time. Next, to lyse the cells, repair the 1X supplemented lysis buffer, and add phosphatase inhibitor cocktail to it.
After carefully removing and discarding the cell culture medium, immediately add 50 microliters of the prepared 1X supplemented lysis buffer, and incubate for 30 minutes at room temperature, under shaking with moderate agitation.
For TR-FRET detection, prepare the 4X antibody detection mix in 1X detection buffer. Then, prepare the Eu-Ab1 and FR-Ab2 antibody solutions, as well as the Eu-Ab3 and FR-Ab4 antibody solutions in 1X detection buffer. Finally, mix pre-diluted Eu-Ab1 with pre-diluted FR-Ab2 for detection of phosphoprotein, and pre-diluted Eu-Ab3 and pre-diluted FR-Ab4 for detection of total protein.
Then, carefully pipette 15 microliters of the cell lysate from the 96-well culture plate to a well of a white low-volume 384-well microplate. Next, add 15 microliters of the positive control lysate and 15 microliters of 1X lysis buffer as a negative control, to separate assay wells.
To wells containing 15 microliters of the lysate, add 5 microliters of the corresponding 4X antibody detection mix to detect the phosphoprotein or total protein. After covering the plate with a plate sealer, incubate it at room temperature for 1 hour up to overnight depending on the assay kit.
After the incubation is complete, remove the adhesive plate sealer, and read the plate on a TR-FRET-compatible microplate reader.

Enabling Real-Time Compensation in Fast Photochemical Oxidations of Proteins for the Determination of Protein Topography Changes

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