Name: defining hsp33's redox-regulated chaperone activity and mapping conformational changes on hsp33 using hydrogen-deuterium exchange mass spectrometry
Uploaded: 2018-06-07T15:00:00
Description: Watch this Scientific Journal Video about Defining Hsp33's Redox-regulated Chaperone Activity and Mapping Conformational Changes on Hsp33 Using Hydrogen-deuterium Exchange Mass Spectrometry at JoVE.co

The authors are thankful to Meytal Radzinski for her helpful discussions and critical reading of the article, and to Patrick Griffin and his lab members for their unlimited assistance while establishing the HDX analysis platform. The authors are grateful to the German-Israel Foundation (I-2332-1149.9/2012), the Binational Science Foundation (2015056), the Marie-Curie integration grant (618806), the Israel Science Foundation (1765/13 and 2629/16), and the Human Frontier Science Program (CDA00064/2014) for their financial support.

1. Preparation of Fully Reduced and Fully Oxidized Proteins
<ol>
	<li>Preparation of a fully reduced protein 
	Note: Here, we describe the reduction of a zinc-containing protein, and use a ZnCl2 solution to restore the Zn-incorporated, reduced protein state. The ZnCl2 solution can be replaced or discarded. Note that the time and temperature of the reduction process depends on the protein stability and function, and is thus specific per protein.
	<ol>
		<li>Thaw the protein sa...

In this paper, we provided protocols for the analysis of redox-dependent chaperone activity and the characterization of structural changes upon the binding of a client protein. These are complementary methodologies to define potential chaperone-substrate complexes and analyze potential interaction sites.
Here, we applied these protocols for the characterization of a complex between the redox-regulated chaperone Hsp33 with a well-studied chaperone substrate CS. We presented two different types ...

Cells frequently encounter an accumulation of reactive oxygen species (ROS) produced as byproducts of respiration1,2, protein and lipid oxidation3,4, and additional processes5,6,7. Despite ROS&#39; beneficial role in diverse biological processes such as cellular signaling8,9 and immune response10, an imbalance between ROS production and its detoxification might occur, leading to oxidative stress7. The biological targets of ROS are proteins, lipids, and nucleic acids, the oxidation of which affect their structure and function. Therefore, the accumulation of cellular oxidants is strongly linked to a diverse range of pathologies including cancer9,11, inflammation12,13, and aging14,15, and have been found to be involved in the onset and progression of neurodegenerative disorders such as Alzheimer&#39;s, Parkinson&#39;s, and ALS disease16,17,18.
Both newly synthesized and mature proteins are highly sensitive to oxidation due to the potentially harmful modifications of their side chains, which shape protein structure and function19,20. Therefore, oxidative stress usually leads to a widespread protein inactivation, misfolding and aggregation, eventually leading to cell death. One of the elegant cellular strategies to cope with the potential damage of protein oxidation is to utilize redox-dependent chaperones, which inhibit the widespread protein aggregation, instead of forming stable complexes with misfolded client proteins21,22,23. These first-line defense chaperones are rapidly activated by a site-specific oxidation (usually on cysteine residues) that converts them into potent anti-aggregation molecules24. Since oxidative stress results in the inhibition of respiration and in decreases in the cellular ATP levels25, canonical ATP-dependent chaperones are less effective during oxidative stress conditions25,26,27. Therefore, redox-activated ATP-independent chaperones play a vital role in maintaining protein homeostasis upon the accumulation of oxidants in bacteria and eukaryotes (e.g., Hsp3328 and RidA29 in bacteria, Get330 in yeast, peroxiredoxins31 in eukaryotes). The activity of these chaperones strongly depends on reversible structural conformational changes induced by a site-specific oxidation that uncovers hydrophobic regions involved in the recognition of misfolded client proteins.
Research of the anti-aggregation mechanism and the principles governing the recognition of the client proteins by chaperones is not easy due to the dynamic and heterogenic nature of chaperone-substrate interactions32,33,34,35,36,37. However, stress-regulated chaperones have an opportunity to advance our understanding of the anti-aggregation function due to their ability to: 1) obtain two different forms of the chaperone, active (e.g., oxidized) and inactive (e.g., reduced), with the introduction or removal of a stress condition easily switching between them (e.g., oxidant and reducing agent), 2) have a broad range of substrates, 3) form highly stable complexes with the client proteins that may be evaluated by different structural methodologies, and 4) focus solely on the substrate recognition and release, mediated by redox-dependent conformational changes, as the majority of these chaperones lacks the folding capability.
Here, we analyze the bacterial redox-regulated chaperone Hsp33&#39;s anti-aggregation activity, a vital component of the bacterial defense system against oxidation-induced protein aggregation28. When reduced, Hsp33 is a tightly folded zinc-binding protein with no chaperone activity; however, when exposed to oxidative stress, Hsp33 undergoes extensive conformational changes which expose its substrate binding regions38,39. Upon oxidation, the zinc ion that is strongly bound to four highly conserved cysteine residues of the C-terminal domain is released40. This results in the formation of two disulfide bonds, an unfolding of the C-terminal domain, and a destabilization of the adjacent linker region41. The C-terminal and linker regions are highly flexible and are defined as intrinsically or partially disordered. Upon return to non-stress conditions, the cysteines become reduced and the chaperone returns to its native folded state with no anti-aggregation activity. The refolding of the chaperone leads to a further unfolding and destabilization of the bound client protein, which triggers its transfer to the canonical chaperone system, DnaK/J, for refolding38. Analysis of Hsp33&#39;s interaction sites suggests that Hsp33 uses both its charged disordered regions as well as the hydrophobic regions on the linker and N-terminal domain to capture misfolded client proteins and prevent their aggregation38,42. In the folded state, these regions are hidden by the folded linker and C-terminal domain. Interestingly, the linker region serves as a gatekeeper of Hsp33&#39;s folded and inactive state, &#34;sensing&#34; the folding status of its adjacent C-terminal domain34. Once destabilized by mutagenesis (either by a point mutation or a full sequence perturbation), Hsp33 is converted to a constitutively active chaperone regardless the redox state of its redox-sensitive cysteines43.
The protocols presented here allow monitoring of Hsp33&#39;s redox-dependent chaperone activity, as well as mapping conformational changes upon the activation and binding of client proteins. This methodology can be adapted to research other chaperone-client recognition models as well as non-chaperone protein-protein interactions. Moreover, we present protocols for the preparation of fully reduced and oxidized chaperones that can be used in studies of other redox-switch proteins, to reveal potential roles of protein oxidation on the protein activity.
Specifically, we describe a procedure to monitor chaperone activity in vitro and define its substrate specificity under different types of protein aggregation (chemically or thermally induced) using light scattering (LS) measured by a fluorospectrometer44. During aggregation, light scattering at 360 nm increases rapidly due to the increasing turbidity. Thus, aggregation can be monitored in a time-dependent manner at this wavelength. LS is a fast and sensitive method for testing protein aggregation and thus the anti-aggregation activity of a protein of interest using nanomolar concentrations, enabling the characterization of protein aggregation-related kinetic parameters under different conditions. Moreover, the LS protocol described here does not require expensive instrumentation, and can be easily established in any laboratory.
Nevertheless, it is quite challenging to obtain &#34;clean&#34; kinetic curves and to derive a protein&#39;s kinetic parameters from such light scattering experiments, due to noise and the large number of outliers generated by air bubbles and large aggregates. To overcome this obstacle, we present a novel graphical tool, Kfits45, used for reducing noise levels in different kinetic measurements, specifically fitted for protein aggregation kinetic data. This software provides preliminary kinetic parameters for an early assessment of the results and allows the user to &#34;clean&#34; large quantities of data quickly without affecting its kinetic properties. Kfits is implemented in Python and available in open source at 45.
One of the challenging questions in the field relates to mapping interaction sites between chaperones and their client proteins and understanding how chaperones recognize a wide range of misfolded substrates. This question is further complicated when studying highly dynamic protein complexes which involve intrinsically disordered chaperones and aggregation-prone substrates. Fortunately, structural mass spectrometry has dramatically advanced over the last decade and has successfully provided helpful approaches and tools to analyze the structural plasticity and map residues involved in protein recognition46,47,48,49. Here, we present one such technique-hydrogen-deuterium exchange mass spectrometry (HDX-MS)-which allows the mapping of residue-level changes in a structural conformation upon protein modification or protein/Ligand binding35,50,51,52,53,54,55. HDX-MS uses the continuous exchange of backbone hydrogens by deuterium, the rate of which is affected by the chemical environment, accessibility, and covalent and non-covalent bonds56. HDX-MS tracks these exchange processes using a deuterated solvent, commonly heavy water (D2O), and allows measurement based on the change in molecular weight following the hydrogen to deuterium exchange. Slower rates of hydrogen-deuterium exchange can result from hydrogens participating in hydrogen bonds or, simply, from steric hindrance, which indicates local changes in structure57. Changes upon a ligand binding or post-translational modifications can also lead to differences in the hydrogen environment, with a binding resulting in differences in the hydrogen-deuterium exchange (HDX) rates46,53.
We applied this technology to 1) map Hsp33 regions which rapidly unfold upon oxidation, leading to the activation of Hsp33, and 2) define the potential binding interface of Hsp33 with its full-length misfolded substrate, citrate synthase (CS)38.
The methods described in this manuscript can be applied to study redox-dependent functions of proteins in vitro, defining anti-aggregation activity and the role of structural changes (if any) in protein function. These methodologies can be easily adapted to diverse biological systems and applied in the laboratory.

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			<td height="21" style="height:21px;width:792px;">Acetonitrile HPLC plus</td>
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			<td>solvent</td>
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			<td>solvent</td>
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			<td height="20" style="height:20px;width:792px;">Tris(hydroxymethyl)aminomethane</td>
			<td style="width:256px;">Sigma Aldrich</td>
			<td style="width:119px;">252859</td>
			<td>buffer</td>
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			<td height="20" style="height:20px;width:792px;">Trifluoroacetic acid</td>
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			<td style="width:119px;">76-05-1</td>
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			<td style="width:256px;">Sigma Aldrich</td>
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			<td>solvent</td>
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			<td height="20" style="height:20px;">ZnCl2, Zinc Chloride</td>
			<td style="width:256px;">Merck</td>
			<td style="width:119px;">B0755416 308</td>
			<td>reagent</td>
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			<td height="20" style="height:20px;">DTT</td>
			<td>goldbio</td>
			<td>27565-41-9</td>
			<td>reducing agent</td>
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			<td height="20" style="height:20px;">PD mini trap G-25 columns GE healthcare</td>
			<td>GE healthcare</td>
			<td>29-9180-07</td>
			<td>desalting column</td>
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			<td height="20" style="height:20px;">Potassium Phosphate</td>
			<td>United states Biochemical Corporation</td>
			<td>20274</td>
			<td>buffer</td>
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			<td height="40" style="height:40px;">Hydrogen peroxide 30%</td>
			<td style="width:256px;">Merck</td>
			<td style="width:119px;">K46809910526</td>
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			<td height="20" style="height:20px;">citrate synthase</td>
			<td style="width:256px;">sigma aldrich</td>
			<td style="width:119px;">C3260</td>
			<td>substrate</td>
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			<td height="20" style="height:20px;">HEPES acid free</td>
			<td style="width:256px;">sigma aldrich</td>
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			<td>denaturant</td>
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			<td height="20" style="height:20px;">Deuterium Chloride Solution</td>
			<td style="width:256px;">sigma aldrich</td>
			<td style="width:119px;">543047-10G</td>
			<td>buffer</td>
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			<td height="20" style="height:20px;">Deuterium Oxide 99%</td>
			<td style="width:256px;">sigma aldrich</td>
			<td style="width:119px;">151882-100G</td>
			<td>solvent</td>
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			<td height="21" style="height:21px;width:792px;">Jasco FP-8500 Fluorospectrometer</td>
			<td>Jasco</td>
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			<td style="width:256px;">Sartorius</td>
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			<td height="23" style="height:23px;">AffiPro Immobilized Pepsin column (20mm length, 2.0mm diameter).</td>
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			<td height="20" style="height:20px;">Waters Pre-column (ACQUITY UPLC BEH C18 VanGuard 130 &#197;, 1.7um, 2.1mmx5mm)</td>
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			<td style="width:119px;"></td>
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			<td height="20" style="height:20px;width:792px;">Vinyl Anaerobic chamber with Airlock door</td>
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			<td height="22" style="height:22px;width:792px;">PAL system LHX - robotic system for handling HDX samples</td>
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			<td colspan="2">https://www.palsystem.com/index.php?id=840</td>
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			<td height="22" style="height:22px;width:792px;">Dionex Ultimate 3000, XRS pump</td>
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			<td height="22" style="height:22px;width:792px;">Dionex AXP-MS auxiliary pump</td>
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			<td height="21" style="height:21px;width:792px;">Kfits: Fit aggregation Data</td>
			<td>http://kfits.reichmannlab.com/fitter/</td>
			<td style="width:119px;"></td>
			<td></td>
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		<tr height="40">
			<td height="40" style="height:40px;">Thermo Scientific Xcalibur software</td>
			<td style="width:256px;">https://www.thermofisher.com/order/catalog/product/OPTON-30487</td>
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			<td style="width:256px;">https://tools.thermofisher.com/content/sfs/brochures/WS-MS-Q-Exactive-Calibration-Maintenance-iQuan2016-EN.pdf</td>
			<td style="width:119px;"></td>
			<td></td>
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			<td height="60" style="height:60px;">Chronos software (Axel Semrau)</td>
			<td style="width:256px;">http://www.axel-semrau.de/en/Software/Software+Solutions/Chronos-p-966.html</td>
			<td style="width:119px;"></td>
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			<td height="40" style="height:40px;">Proteome Discoverer V1.4 software</td>
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			<td height="24" style="height:24px;">HDX workbench software</td>
			<td colspan="3">http://hdx.florida.scripps.edu/hdx_workbench/Home.html</td>
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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.

The two methods presented make it possible to follow the kinetic activity and dynamics of protein interactions between a chaperone and its substrate. Moreover, the reduction-oxidation protocol allows the preparation of a fully reduced and fully oxidized chaperone, giving a more in-depth understanding of the activation mechanism of redox-dependent disordered chaperones.
First, we used light scattering in order to examine the redo...

Watch this Scientific Journal Video about Defining Hsp33's Redox-regulated Chaperone Activity and Mapping Conformational Changes on Hsp33 Using Hydrogen-deuterium Exchange Mass Spectrometry at JoVE.co

Defining Hsp33&#039;s Redox-regulated Chaperone Activity and Mapping Conformational Changes on Hsp33 Using Hydrogen-deuterium Exchange Mass Spectrometry

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.

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 ...

The workflow presented here can help extend our understanding of the cellular role of chaperones during oxidative stress conditions. Specifically, it can help to understand how do chaperones prevent protein aggregation during unfolding conditions. For these two backs of different techniques provide insight into the activity of flag segregated chaperones.It can also be used to study other protein for interaction such as enzyme social interactions and protein dynamics. Visual demonstration of this method is critical as it involves the usage of different instruments which may be challenging to new users. First, spin down a previously thawed protein sample to remove aggregates.Then add five millimolar DTT and 20 micromolar zinc chloride and incubate the sample for at least 1.5 hours at 37 degrees Celsius. To remove the DTT equilibrate a desalting column with at 40 millimolar potassium phosphate buffer by filling the column completely with the buffer and letting it drip out. Using tweezers, gently push down on the white disc filter in the column and remove it.Refill the column with potassium phosphate buffer and centrifuge at 1000 times G for three minutes. Following centrifugation transfer the column into a clean tube and add the protein sample slowly to the middle of the column. Centrifuge the column at 1000 times G for two minutes so that the DTT-free protein is in the flow-through.Now check the protein concentration and measure its absorbance at 280 nanometers. Distribute half of the protein samples into aliquots. Incubate the aliquots under anaerobic conditions for 20 minutes for complete removal of oxygen.Then seal the tubes with plastic film and store the samples under cold conditions. Next, add freshly-prepared five millimolar hydrogen peroxide to the remaining protein sample and incubate for three hours at 40 degrees Celsius while shaking. Following centrifugation check the protein concentration then divide the oxidized proteins into aliquots and store them under cold conditions.Prepare the denatured substrate by incubating 12 micromolar citrate synthase overnight in 40 millimolar HEPES and 4.5 molar guanidinium chloride. On the following day, open the fluorospectrometer software and go to time course measurement. Set the excitation bandwidth to 2.5 nanometers.The excitation wavelength to 360 nanometers. The emission wavelength to 360 nanometers. The emission bandwidth to five nanometers.And the data interval to 0.5 seconds. Set the temperature to 25 degrees Celsius. Prepare the sample by adding 1600 microliters of 40 millimolar HEPES to a quartz cuvette.Insert the cuvette into the sample holder of a fluorospectrometer allowing the sample to reach the desired temperature. After setting the stirring to 600 rpm begin the measurement until the baseline is established. To measure the citrate synthase aggregation in the absence of a chaperone, add 10 microliters of denatured citrate synthetase, 120 seconds into the measurement.Then continue the measurement for 1200 seconds. To measure the citrate synthase aggregation in the presence of Hsp33 add Hsp33 60 seconds into the measurement. After an additional 60 seconds add 10 microliters of denatured citrate synthetase and continue the measurement for 1200 seconds.Next, upload the data file to perform data analysis and noise removal by Kfits. To remove any noise choose analysis parameters use automatic best model and select noise is always above signal. Manually remove obvious outliers using the green and red adjustable lines which will filter the noises above the green line and below the red line.Following this, set the baseline and the fit curve. After applying the noise threshold, download the processed data. Prepare protein substrate complex samples by adding citrate synthetase to Hsp33 at a ratio of 1 to 1.5 at 43 degrees Celsius.Use at least four steps and incubate the samples for 15 minutes at 43 degrees Celsius after every addition to allow complex formation. Centrifuge the samples at 16000 times G at four degrees Celsius for 30 minutes to remove any aggregates. Then place the samples in 150 microliter glass inserts and transfer them into vials.Then place them in the proper trays. Hold buffers H and D at 25 degrees Celsius and hold the samples and quenching buffer at zero to two degrees Celsius. Next, manually turn on all cooling units of the mass spectrometer.Once all cooling systems reach their target temperatures manually switch on both the HPLC and loading pumps. Open the software which controls both pumps as well as the software which controls the mass spectrometer and make sure that the MS is on standby. Following this, disconnect the HPLC outlet valve from the MS source.Wash the system first with a triple cleaner solution. Then with buffer B and finally with buffer A.Then insert the pepsin column into the system. After washing the system with a steady flow and steady pressure, insert the HPLC outlet valve into the MS source and turn the MS on.Now open the software. Build a running sequence and enter all time points, sample names and locations in the trays as well as the buffer names and locations. After the run is complete analyze the peptide coverage of the sample by running the non-deuterated control samples through the software.Transfer data into the HDX Workbench software and analyze the results using the software. Save the selected peptides and continue further for analysis of the deuterium uptake. Chemical denaturation leads to a rapid citrate synthetase aggregation induced by the re-folding of a fully denaturized protein in unfavorable buffer conditions.On the other hand, the thermal-induced aggregation results from a relatively slow unfolding of the natively folded substrate. In both forms of denaturation, the addition of oxidized Hsp33 completely abolished aggregation, lowering the 360 nanometer readings to negligible values. The presence of a reduced Hsp33 had no effect on the citrate synthetase stability and gave an aggregation curve similar to the one detected in the absence of a chaperone.The deuteration pattern between the bound and unbound chaperone Hsp33 to its substrate citrate synthetase reveals a client interaction site within the C-terminal redox switch domain of the chaperone. Reduced Hsp33 has a completely ordered structure but becomes functional when it senses oxidative stress. While the reduced bound and unbound Hsp33 produces similar peptide curves the oxidized Hsp33 shows a 30%deuteration difference between the bound and unbound chaperone with specific areas of the bound protein indicating an allosteric hindrance.Once established the scattering assay can be completed within a few hours while the DHEX methodology can be completed in two to four days if performed properly. While attempting this procedure it is important to plan ahead. Remember that the condition described such as temperature, incubation times and buffer solutions are protein-specific and must be optimized for each protein.Following this procedure other methodologies such as cross-linking, or precipitation can be performed in order to complement studies of chaperone substrates interactions. In conclusion, this technique has paved the way for researchers in the field of structural biology to explore protein plasticity in molecular chaperones.

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Time-resolved ElectroSpray Ionization Hydrogen-deuterium Exchange Mass Spectrometry for Studying Protein Structure and Dynamics

Intrinsically disordered proteins (IDPs) have long been a challenge to structural biologists due to their lack of stable secondary structure elements. Hydrogen-Deuterium Exchange (HDX) measured at rapid time scales is uniquely suited to detect structures and hydrogen bonding networks that are briefly populated, allowing for the characterization of transient conformers in native ensembles. Coupling of HDX to mass spectrometry offers several key advantages, including high sensitivity, low sample consumption and no restriction on protein size. This technique has advanced greatly in the last several decades, including the ability to monitor HDX labeling times on the millisecond time scale. In addition, by incorporating the HDX workflow onto a microfluidic platform housing an acidic protease microreactor, we are able to localize dynamic properties at the peptide level. In this study, Time-Resolved ElectroSpray Ionization Mass Spectrometry (TRESI-MS) coupled to HDX was used to provide a detailed picture of residual structure in the tau protein, as well as the conformational shifts induced upon hyperphosphorylation.

Conformational flexibility plays a critical role in protein function. Herein, we describe the use of time-resolved electrospray ionization mass spectrometry coupled to hydrogen-deuterium exchange for probing the rapid structural changes that drive function in ordered and disordered proteins.

Intrinsically disordered proteins (IDPs) have long been a challenge to structural biologists due to their lack of stable secondary structure elements. ...

Resolving Affinity Purified Protein Complexes by Blue Native PAGE and Protein Correlation Profiling

Most proteins act in association with others; hence, it is crucial to characterize these functional units in order to fully understand biological processes. Affinity purification coupled to mass spectrometry (AP-MS) has become the method of choice for identifying protein-protein interactions. However, conventional AP-MS studies provide information on protein interactions, but the organizational information is lost. To address this issue, we developed a strategy to unravel the distinct functional assemblies a protein might be involved in, by resolving affinity-purified protein complexes prior to their characterization by mass spectrometry. Protein complexes isolated through affinity purification of a bait protein using an epitope tag and competitive elution are separated through blue native electrophoresis. Comparison of protein migration profiles through correlation profiling using quantitative mass spectrometry allows assignment of interacting proteins to distinct molecular entities. This method is able to resolve protein complexes of close molecular weights that might not be resolved by traditional chromatographic techniques such as gel filtration. With little more work than conventional AP-geLC-MS/MS, we demonstrate this strategy may in many cases be adequate for obtaining protein complex topological information concomitantly to identifying protein interactions.

Here we present protocols for affinity purification of protein complexes and their separation by blue native PAGE, followed by protein correlation profiling using label free quantitative mass spectrometry. This method is useful to resolve interactomes into distinct protein complexes.

Most proteins act in association with others; hence, it is crucial to characterize these functional units in order to fully understand biological ...

Lipid Droplet Isolation for Quantitative Mass Spectrometry Analysis

Lipid droplets are vital to the replication of a variety of different pathogens, most prominently the Hepatitis C Virus (HCV), as the putative site of virion morphogenesis. Quantitative lipid droplet proteome analysis can be used to identify proteins that localize to or are displaced from lipid droplets under conditions such as virus infections. Here, we describe a protocol that has been successfully used to characterize the changes in the lipid droplet proteome following infection with HCV. We use Stable Isotope Labeling with Amino Acids in Cell Culture (SILAC) and thus label the complete proteome of one population of cells with &#34;heavy&#34; amino acids to quantitate the proteins by mass spectrometry. For lipid droplet isolation, the two cell populations (i.e.&#160;HCV-infected/&#34;light&#34; amino acids and uninfected control/&#34;heavy&#34; amino acids) are mixed 1:1 and lysed mechanically in hypotonic buffer. After removing the nuclei and cell debris by low speed centrifugation, lipid droplet-associated proteins are enriched by two subsequent ultracentrifugation steps followed by three washing steps in isotonic buffer. The purity of the lipid droplet fractions is analyzed by western blotting with antibodies recognizing different subcellular compartments. Lipid droplet-associated proteins are then separated by SDS-polyacrylamide gel electrophoresis (SDS-PAGE) followed by Coomassie staining. After tryptic digest, the peptides are quantified by liquid chromatography-electrospray ionization-tandem mass spectrometry (LC-ESI-MS/MS). Using this method, we identified proteins recruited to lipid droplets upon HCV infection that might represent pro- or antiviral host factors. Our method can be applied to a variety of different cells and culture conditions, such as infection with pathogens, environmental stress, or drug treatment.

Lipid droplets are important organelles for the replication of several pathogens, including the Hepatitis C Virus (HCV). We describe a method to isolate lipid droplets for quantitative mass spectrometry of associated proteins; it can be used under a variety of conditions, such as virus infection, environmental stress, or drug treatment.

Lipid droplets are vital to the replication of a variety of different pathogens, most prominently the Hepatitis C Virus (HCV), as the putative site of ...

The Determination of Protease Specificity in Mouse Tissue Extracts by MALDI-TOF Mass Spectrometry: Manipulating PH to Cause Specificity Changes

Proteases have several biological functions, including protein activation/inactivation and food digestion. Identifying protease specificity is important for revealing protease function. The method proposed in this study determines protease specificity by measuring the molecular weight of unique substrates using Matrix-assisted Laser Desorption/Ionization Time-of-Flight (MALDI-TOF) mass spectrometry. The substrates contain iminobiotin, while the cleaved site consists of amino acids, and the spacer consists of polyethylene glycol. The cleaved substrate will generate a unique molecular weight using a cleaved amino acid. One of the merits of this method is that it may be carried out in one pot using crude samples, and it is also suitable for assessing multiple samples. In this article, we describe a simple experimental method optimized with samples extracted from mouse lung tissue, including tissue extraction, placement of digestive substrates into samples, purification of digestive substrates under different pH conditions, and measurement of the substrates&#39; molecular weight using MALDI-TOF mass spectrometry. In summary, this technique allows for the identification of protease specificity in crude samples derived from tissue extracts using MALDI-TOF mass spectrometry, which may easily be scaled up for multiple sample processing.

Here, we present a protocol to determine protease specificity in crude mouse tissue extracts using MALDI-TOF spectrometry.

Proteases have several biological functions, including protein activation/inactivation and food digestion. Identifying protease specificity is important ...

Proofreading and DNA Repair Assay Using Single Nucleotide Extension and MALDI-TOF Mass Spectrometry Analysis

The maintenance of the genome and its faithful replication is paramount for conserving genetic information. To assess high fidelity replication, we have developed a simple non-labeled and non-radio-isotopic method using a matrix-assisted laser desorption ionization with time-of-flight (MALDI-TOF) mass spectrometry (MS) analysis for a proofreading study. Here, a DNA polymerase [e.g., the Klenow fragment (KF) of Escherichia coli DNA polymerase I (pol I) in this study] in the presence of all four dideoxyribonucleotide triphosphates is used to process a mismatched primer-template duplex. The mismatched primer is then proofread/extended and subjected to MALDI-TOF MS. The products are distinguished by the mass change of the primer down to single nucleotide variations. Importantly, a proofreading can also be determined for internal single mismatches, albeit at different efficiencies. Mismatches located at 2-4-nucleotides (nt) from the 3' end were efficiently proofread by pol I, and a mismatch at 5 nt from the primer terminus showed only a partial correction. No proofreading occurred for internal mismatches located at 6 - 9 nt from the primer 3' end. This method can also be applied to DNA repair assays (e.g., assessing a base-lesion repair of substrates for the endo V repair pathway). Primers containing 3' penultimate deoxyinosine (dI) lesions could be corrected by pol I. Indeed, penultimate T-I, G-I, and A-I substrates had their last 2 dI-containing nucleotides excised by pol I before adding a correct ddN 5'-monophosphate (ddNMP) while penultimate C-I mismatches were tolerated by pol I, allowing the primer to be extended without repair, demonstrating the sensitivity and resolution of the MS assay to measure DNA repair.

A non-labeled, non-radio-isotopic method for DNA polymerase proofreading and a DNA repair assay was developed by using high-resolution MALDI-TOF mass spectrometry and a single nucleotide extension strategy. The assay proved to be very specific, simple, rapid, and easy to perform for proofreading and repair patches shorter than 9-nucleotides.

The maintenance of the genome and its faithful replication is paramount for conserving genetic information. To assess high fidelity replication, we have ...

Analyzing Dynamic Protein Complexes Assembled On and Released From Biolayer Interferometry Biosensor Using Mass Spectrometry and Electron Microscopy

In vivo, proteins are often part of large macromolecular complexes where binding specificity and dynamics ultimately dictate functional outputs. In this work, the pre-endosomal anthrax toxin is assembled and transitioned into the endosomal complex. First, the N-terminal domain of a cysteine mutant lethal factor (LFN) is attached to a biolayer interferometry (BLI) biosensor through disulfide coupling in an optimal orientation, allowing protective antigen (PA) prepore to bind (Kd 1 nM). The optimally oriented LFN-PAprepore complex then binds to soluble capillary morphogenic gene-2 (CMG2) cell surface receptor (Kd 170 pM), resulting in a representative anthrax pre-endosomal complex, stable at pH 7.5. This assembled complex is then subjected to acidification (pH 5.0) representative of the late endosome environment to transition the PAprepore into the membrane inserted pore state. This PApore state results in a weakened binding between the CMG2 receptor and the LFN-PApore and a substantial dissociation of CMG2 from the transition pore. The thio-attachment of LFN to the biosensor surface is easily reversed by dithiothreitol. Reduction on the BLI biosensor surface releases the LFN-PAprepore-CMG2 ternary complex or the acid transitioned LFN-PApore complexes into microliter volumes. Released complexes are then visualized and identified using electron microscopy and mass spectrometry. These experiments demonstrate how to monitor the kinetic assembly/disassembly of specific protein complexes using label-free BLI methodologies and evaluate the structure and identity of these BLI assembled complexes by electron microscopy and mass spectrometry, respectively, using easy-to-replicate sequential procedures.

Here we present a protocol to monitor the assembly and disassembly of the anthrax toxin using biolayer interferometry (BLI). Following assembly/disassembly on the biosensor surface, the large protein complexes are released from the surface for visualization and identification of components of the complexes using electron microscopy and mass spectrometry, respectively.

In vivo, proteins are often part of large macromolecular complexes where binding specificity and dynamics ultimately dictate functional outputs. In this ...

Nitropeptide Profiling and Identification Illustrated by Angiotensin II

Protein nitration is one of the most important post-translational modifications (PTM) on tyrosine residues and it can be induced by chemical actions of reactive oxygen species (ROS) and reactive nitrogen species (RNS) in eukaryotic cells. Precise identification of nitration sites on proteins is crucial for understanding the physiological and pathological processes related to protein nitration, such as inflammation, aging, and cancer. Since the nitrated proteins are of low abundance in cells even under induced conditions, no universal and efficient methods have been developed for the profiling and identification of protein nitration sites. Here we describe a protocol for nitropeptide enrichment by using a chemical reduction reaction and biotin labeling, followed by high resolution mass spectrometry. In our method, nitropeptide derivatives can be identified with high accuracy. Our method exhibits two advantages compared to the previously reported methods. First, dimethyl labeling is used to block the primary amine on nitropeptides, which can be used to generate quantitative results. Second, a disulfide bond containing NHS-biotin reagent is used for the enrichment, which can be further reduced and alkylated to enhance the detection signal on a mass spectrometer. This protocol has been successfully applied to the model peptide Angiotensin II in the current paper.

Proteomic profiling of tyrosine-nitrated proteins has been a challenging technique due to the low abundance of the 3-nitrotyrosine modification. Here we describe a novel approach for nitropeptide enrichment and profiling by using Angiotensin II as the model. This method can be extended for other in vitro or in vivo systems.

Protein nitration is one of the most important post-translational modifications (PTM) on tyrosine residues and it can be induced by chemical actions of ...

A Plasma Sample Preparation for Mass Spectrometry using an Automated Workstation

Sample preparation for mass spectrometry analysis in proteomics requires enzymatic cleavage of proteins into a peptide mixture. This process involves numerous incubation and liquid transfer steps in order to achieve denaturation, reduction, alkylation, and cleavage. Adapting this workflow onto an automated workstation can increase efficiency and reduce coefficients of variance, thereby providing more reliable data for statistical comparisons between sample types. We previously described an automated proteomic sample preparation workflow1. Here, we report the development of a more efficient and better controlled workflow with the following advantages: 1) The number of liquid transfer steps is reduced from nine to six by combining reagents; 2) Pipetting time is reduced by selective tip pipetting using a 96-position pipetting head with multiple channels; 3) Potential throughput is increased by the availability of up to 45 deck positions; 4) Complete enclosure of the system provides improved temperature and environmental control and reduces the potential for contamination of samples or reagents; and 5) The addition of stable isotope labeled peptides, as well as &#946;-galactosidase protein, to each sample makes monitoring and quality control possible throughout the entire process. These hardware and process improvements provide good reproducibility and improve intra-assay and inter-assay precision (CV of less than 20%) for LC-MS based protein and peptide quantification. The entire workflow for digesting 96 samples in a 96-well plate can be completed in approximately 5 hours.

Here, we present an automated plasma protein digestion method for mass spectrometry-based quantitative proteomic analysis. In this protocol, the liquid transferring and incubation steps for protein denaturation, reduction, alkylation, and trypsin digestion reactions are streamlined and automated. It takes approximately five hours to prepare a 96-well plate with desired precision.

Sample preparation for mass spectrometry analysis in proteomics requires enzymatic cleavage of proteins into a peptide mixture. This process involves ...

Sample Preparation for Probe Electrospray Ionization Mass Spectrometry

Mass spectrometry (MS) is a powerful tool in analytical chemistry because it provides very accurate information about molecules, such as mass-to-charge ratios (m/z), which are useful to deduce molecular weights and structures. While it is essentially a destructive analytical method, recent advancements in the ambient ionization technique have enabled us to acquire data while leaving tissue in a relatively intact state in terms of integrity. Probe electrospray ionization (PESI) is a so-called direct method because it does not require complex and time-consuming pretreatment of samples. A fine needle serves as a sample picker, as well as an ionization emitter. Based on the very sharp and fine property of the probe tip, destruction of the samples is minimal, allowing us to acquire the real-time molecular information from living things in situ. Herein, we introduce three applications of PESI-MS technique that will be useful for biomedical research and development. One involves the application to solid tissue, which is the basic application of this technique for the medical diagnosis. As this technique requires only 10 mg of the sample, it may be very useful in the routine clinical settings. The second application is for in vitro medical diagnostics where human blood serum is measured. The ability to measure fluid samples is also valuable in various biological experiments where a sufficient volume of sample for conventional analytical techniques cannot be provided. The third application leans toward the direct application of probe needles in living animals, where we can obtain real-time dynamics of metabolites or drugs in specific organs. In each application, we can infer the molecules that have been detected by MS or use artificial intelligence to obtain a medical diagnosis.

This article introduces sample preparation methods for a unique real-time analytical method based on the ambient mass spectrometry. This method lets us perform real-time analysis of the biological molecules in vivo without any special pretreatments.

Mass spectrometry (MS) is a powerful tool in analytical chemistry because it provides very accurate information about molecules, such as mass-to-charge ...

Quantitative Analysis of the Cellular Lipidome of Saccharomyces Cerevisiae Using Liquid Chromatography Coupled with Tandem Mass Spectrometry

Lipids are structurally diverse amphipathic molecules that are insoluble in water. Lipids are essential contributors to the organization and function of biological membranes, energy storage and production, cellular signaling, vesicular transport of proteins, organelle biogenesis, and regulated cell death. Because the budding yeast Saccharomyces cerevisiae is a unicellular eukaryote amenable to thorough molecular analyses, its use as a model organism helped uncover mechanisms linking lipid metabolism and intracellular transport to complex biological processes within eukaryotic cells. The availability of a versatile analytical method for the robust, sensitive, and accurate quantitative assessment of major classes of lipids within a yeast cell is crucial for getting deep insights into these mechanisms. Here we present a protocol to use liquid chromatography coupled with tandem mass spectrometry (LC-MS/MS) for the quantitative analysis of major cellular lipids of S. cerevisiae. The LC-MS/MS method described is versatile and robust. It enables the identification and quantification of numerous species (including different isobaric or isomeric forms) within each of the 10 lipid classes. This method is sensitive and allows identification and quantitation of some lipid species at concentrations as low as 0.2 pmol/&#181;L. The method has been successfully applied to assessing lipidomes of whole yeast cells and their purified organelles. The use of alternative mobile phase additives for electrospray ionization mass spectrometry in this method can increase the efficiency of ionization for some lipid species and can be therefore used to improve their identification and quantitation.

Quantitative Analysis of the Cellular Lipidome of Saccharomyces Cerevisiae Using Liquid Chromatography Coupled with Tandem Mass Spectrometry

We present a protocol using liquid chromatography coupled with tandem mass spectrometry to identify and quantify major cellular lipids in Saccharomyces cerevisiae. The described method for a quantitative assessment of major lipid classes within a yeast cell is versatile, robust, and sensitive.

Lipids are structurally diverse amphipathic molecules that are insoluble in water. Lipids are essential contributors to the organization and function of ...