This work was funded by the National Institute of General Medical Sciences (R43GM125420 and R44GM125420).

1. Installing the capillary tube
<ol>
	<li>Using a silica cleaving stone, cleave 250 &#181;m inner diameter (ID) silica capillary to 27 inches. Check the capillary ends for a clean, straight cut.</li>
	<li>Create two windows roughly 15 mm in length by burning away the polyimide coating. Starting from the &#8220;lower end&#8221; make the first photolysis window 90 mm away from the &#8220;lower end&#8221; and the second dosimeter window 225 mm away from the &#8220;lower end&#8221;. 
	NOTE: Once the coating is burned away the capillary is very fragile.</li>
	<li>Unscrew the nut and ferrule at port 5 and insert the &#8220;lower end&#8221; of the capillary just beyond the conical end of the ferrule (Figure 1A).</li>
	<li>On the photolysis module, remove the photolysis cell cap by pulling it straight out and then remove the magnetically mounted metal mask which will hold the capillary in place. 
	CAUTION: Inside the photolysis cell cap is a curved mirror, do not allow anything to touch the mirror.</li>
	<li>On the dosimeter module, open the dosimeter cell by pushing on the tab on the left and swing the dosimeter cell open to the right. Remove the magnetically mounted clips which will hold the capillary in place.</li>
	<li>Place the capillary into the grooved channel in the base of the photolysis cell. Center the capillary window with the photolysis cell window. While holding the capillary in position with one hand, add the magnetic mask to hold the capillary in place. Place the photolysis cap back in position.</li>
	<li>Place the capillary into the grooved channel in the base of the dosimeter cell. Center the second capillary window on the small aperture at the center of the dosimeter cell base. While holding the capillary in position with one hand, place the two magnetic clips in position to hold the capillary in place. Close the dosimeter cell until it clicks closed.</li>
	<li>Insert the capillary through the knurled nut atop the capillary lift of the product collector (Figure 1B). Extend the capillary to just above the bottom of the vial. 
	NOTE: It is important for the capillary to reach the bottom of the vial to ensure that the capillary is submerged in the quench solution while running an experiment.</li>
</ol>
2. Installing an injection loop
<ol>
	<li>Use Teflon tubing with an outer diameter 1/16&#8221; and an inner diameter of 0.015&#8221; (381 &#181;m). Follow equation 1 to calculate the length of tubing needed for the desired volume using equation 1. 
	<img alt="Equation 1" src="/files/ftp_upload/61861/61861equ01.jpg" /> 
	Where L is the length of the tubing in millimeters, V is the desired volume in microliters and d is the tube inner diameter in millimeters. For an injection volume of 25 &#181;L and an inner diameter of 381 &#181;m, the tubing length needs to be roughly 219.3 mm. 
	NOTE: For volumes less than 20 &#181;L, use PTFE tubing with an inner diameter of 0.01&#8221;.</li>
	<li>Cut the Teflon tubing to the necessary length using a tube cutter. Check the ends of the tube for a clean, straight cut.</li>
	<li>Insert one end of the new injection loop through one of the nuts and place a new ferrule onto the end of the tube. Hold the ferrule and nut in place and insert the tube into port 3 of the injection valve until it bottoms out in the valve. Hold the tube firmly and tighten the nut finger tight. Using a wrench, tighten the nut &#188; turn further. Remove and inspect the assembly. 
	NOTE: If the ferrule is not fixed in position, reinstall and tighten 1/8 turn further. Repeat with the other end of the loop.</li>
	<li>Once both ends have a nut and fixed ferrule, loosely screw one end to port 3 and the other end to port 6 (Figure 1C). Once in position tighten both sides to finger tight then &#188; turn past finger tight with a wrench.</li>
</ol>
3. Initialize the photolysis system
<ol>
	<li>Turn on the photolysis modules in the following order: (1) Fluidics Module (2) Photolysis Module (3) Dosimetry Module (4) Product Collector, and lastly (5) the system computer and launch the control software 
	NOTE: Allow the Dosimeter Module at least a half-hour to warm up from a cold start.</li>
	<li>Fully submerge tubing from the &#8220;Valve&#8221; position on the syringe pump (Figure 1D) into 10 mL of buffer. Place the tubing from the &#8220;Waste&#8221; position (Figure 1D) and tubing from port 2 on the syringe port (Figure 1C) to an empty container to collect waste. 
	NOTE: Use the buffer your protein will be suspended in. A recommended buffer is 10 mM NaPO4.</li>
	<li>On the Product Collector carousel, place 1.5 mL microcentrifuge tubes at the position marked H and 6.</li>
	<li>If using 250 &#181;m ID capillary, set the load flow rate to 500 &#181;L/min, processing flow rate to 7.5 &#181;L/min, and waste flow rate to 500 &#181;L/min.</li>
	<li>Calculate the flash delay, product time, and waste time to use the Semi-Automatic Mode using equation 2. 
	<img alt="Equation 2" src="/files/ftp_upload/61861/61861equ02.jpg" /> 
	Where t is the time in minutes, s is the capillary distance in millimeters, d is the inner diameter of the capillary in millimeters, and f is the flow rate in &#181;L/min.
	<ol>
		<li>For the flash delay, the distance is from the &#8220;lower end&#8221; of the capillary to the first window, which should be roughly 90 mm. Using 250 &#181;m ID capillary and a processing flow rate of 7.5 &#181;L/min the sample will arrive to the photolysis window in about 35 seconds. Allow two flashes at 1 Hz to occur before the samples arrive, so set the flash delay to 33 seconds.</li>
		<li>For the product time, enter the amount of time injected solution will take to flow from the injection valve to the end of the capillary. For a 27&#8221; long 250 &#181;m ID capillary, set the product time to 4.5 minutes.</li>
		<li>For the waste time, enter the amount of time the total volume of injected solution will take to be collected. At this time, the product collector will move from the product position to the waste vial. For an injection volume of 25 &#181;L and a flow rate of 7.5 &#181;L/min, set the waste time to 7.8 minutes.</li>
	</ol>
	</li>
	<li>Rinse out the injection loop five times by injecting 25 &#181;L of HPLC-grade water into the injecting port with the injection valve set to the load position.</li>
	<li>Manually turn the injection valve up to &#8216;inject position&#8217; to flush the rest of the system. Select Process (Out) on the control software to begin flowing water. While flowing, raise the product collector capillary lift by selecting Up under the product collector so that you can see the end of the capillary. Flow water through the capillary until a droplet forms. 
	NOTE: If a droplet does not form, check the injector valve for leaks. If there is a leak, loosen the nut, reseat the capillary, and retighten.</li>
</ol>
4. Determine actual &#9642;OH yield to test radical scavenging effects from the buffer.
<ol>
	<li>In the Control Software, start the flow by selecting Process (Out). In the settings tab, set the flash voltage to 0 V. Under the Manual Control tab in the dosimeter data section, select Start Data + AutoZero.</li>
	<li>Select the position for the product vial (H) and waste vial (6).</li>
	<li>Select Ready, manually turn the injection valve down to the load position, and inject 25 &#181;L of 1 mM Adenine with 100 mM H2O2 into the injection port. Once injected, manually turn the injection valve up to the inject position. 
	NOTE: This automatically triggers the system to begin the flow, turn on the plasma source, and acquire the dosimetry data.</li>
	<li>Ramp the voltage up by navigating to the settings tab, and changing the flash voltage. Repeat steps 4.2 and 4.3 using 500 V, 750 V, 1000 V, and 1250 V. Perform the reading of adenine absorbance at each voltage in triplicates.</li>
	<li>Select calculations tab to determine the average absorbance of each sample. First, click Select, then manually select the beginning and end of the peak absorbance. In the available space, write in a description of the sample. Repeat for all acquired data. 
	NOTE: Bubbles may form causing a spike in dosimeter reading. When selecting data to determine average absorbance, omit areas with bubbles.</li>
	<li>Copy and paste data in Excel to calculate the average change in adenine absorbance for each voltage, thus determining the effective &#9642;OH yield.</li>
	<li>Repeat steps 4.1-4.6 if multiple sample conditions (different buffer/additives) are being used to normalize the effective &#9642;OH yield for each condition.</li>
</ol>
5. Modification of protein to detect changes in higher order structure.
<ol>
	<li>Mix 4 mM adenine with 20 &#181;M of protein in a one-to-one ratio to make a solution containing 2 mM adenine and 10 &#181;M protein.</li>
	<li>Make the quench solution using 0.3 mg/mL catalase to break down excess hydrogen peroxide and 35 mM methionine amide to quench any remaining radicals. Aliquot 25 &#181;L of quench solution into a 200 &#181;L microtube so that an equal volume of quench and the labeled product is mixed.</li>
	<li>Dilute H2O2 to 200 mM and keep on ice.</li>
	<li>On the Control Software in the settings tab, start the flash voltage at 0 V to determine any background oxidation.</li>
	<li>In the manual control tab, select Start Data + AutoZero, followed by Process (Out), then Ready, and finally turn the injection valve down to the load position.</li>
	<li>Place a quench solution in position 1 on the product collect carousel. On the System Control Software change the product vial to 1.</li>
	<li>Immediately before injection, mix 12.5 &#181;L of the adenine and protein mixture with 12.5 &#181;L of H2O2 to make a final concentration of 1 mM Adenine, 5 &#181;M protein, and 100 mM H2O2. Gently pipette up and down to mix, quickly spin down, and inject 25 &#181;L using the injection port within 10 seconds of mixing.</li>
	<li>Switch the injection valve to the inject position and wait while sample is being processed.</li>
	<li>Repeat acquisition with 500 V, 750 V, 1000 V, and 1250 V. Perform each voltage measurement in triplicate.</li>
	<li>Calculate the average absorbance as described in step 4.5. Copy and paste all data to Excel.</li>
</ol>
6. Shut the system down
<ol>
	<li>After all samples have been collected, flush out the syringe port and sample loop by setting the injection valve down to the load position and inject 25 &#181;L of HPLC water five times.</li>
	<li>Turn the injection valve up to the inject position to flush the rest of the system with HPLC water.</li>
	<li>Stop the flow, close the system control software, and turn off the modules starting with the product collector, dosimeter module, photolysis module, then finally the fluidics module.</li>
</ol>
7. Sample preparation and liquid chromatography-mass spectrometry
<ol>
	<li>Denature the protein by incubating samples at 80 &#176;C for 20 min in the presence of 50 mM Tris and 1 mM CaCl2. Cool samples to room temperature and add 1:20 trypsin to protein. Digest the protein overnight at 37 &#176;C with sample mixing. Next morning, terminate trypsin digestion by heating samples to 95 &#176;C for 10 minutes.</li>
	<li>Detect peptides using a high-resolution LC-MS/MS system connected with a UPLC system.</li>
	<li>Load the sample first onto trap column (300 &#181;m ID X 5 mm 100 &#197; pore size, 5 &#181;m particle size) and wash at 5.0 &#181;L/mL for 3 min with water containing 2% solvent B (acetonitrile and 0.1% formic acid).</li>
	<li>Separate the peptides on a C18 nanocolumn (0.75 mm x 150 mm, 2 &#181;m particle size, 100 &#197; pore size) at a flow rate of 300 nL/min with a gradient between solvent A (water containing 0.1% formic acid) and solvent B. The gradient for peptide elution consists of a linear increase from 2 to 35% B over 22 min, ramped to 95% B over 5 min and held for 3 min to wash the column, and then returned to 2% B over 3 min and held for 9 min to re-equilibrate the column.</li>
	<li>Acquire the data in positive ion mode. Set the spray voltage to 2400 V, and the temperature of the ion transfer tube to 300 &#176;C.</li>
	<li>Acquire the full MS scans from 250-2000 m/z followed by eight subsequent data-dependent MS/MS scans on the top eight most abundant peptide ions. Use collision-induced dissociation at 35% normalized energy to fragment the peptides.</li>
	<li>Identify all the unmodified peptides detected from MS/MS analysis using an available protein analysis software against the necessary FASTA file containing the protein sequence and the relevant proteolytic enzyme.</li>
	<li>Search and quantify modified peptides using a HRPF Data Processing Software. The extent of oxidation for each identified peptide is calculated by dividing the summed chromatographic peak area of a modified peptide by the total chromatographic peak area of that peptide modified and unmodified using equation 3. 
	P = [I(singly oxidized) X 1 + I (doubly oxidized) X 2 + I(triply oxidized) X 3 + .../[Iunoxidized + I(singly oxidized) + I(doubly oxidized) + I(triply oxidized) ...] 
	where P denotes the average number of oxidation events per peptide molecule, and I represents the peak area of the unoxidized peptide (Iunoxidized) and the peptide with n oxidation events.</li>
</ol>
8. For a differential study, repeat steps 5-7 on the second condition.
NOTE: To confirm reproducibility, two biological replicates in addition to technical triplicates for each condition are recommended.

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L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fast+photochemical+oxidation+of+proteins+for+epitope+mapping.">Fast photochemical oxidation of proteins for epitope mapping.</a> Analytical chemistry. 83 (20), 7657-7661 (2011).</li><li>Li, J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mapping+the+Energetic+Epitope+of+an+Antibody/Interleukin-23+Interaction+with+Hydrogen/Deuterium+Exchange,+Fast+Photochemical+Oxidation+of+Proteins+Mass+Spectrometry,+and+Alanine+Shave+Mutagenesis.">Mapping the Energetic Epitope of an Antibody/Interleukin-23 Interaction with Hydrogen/Deuterium Exchange, Fast Photochemical Oxidation of Proteins Mass Spectrometry, and Alanine Shave Mutagenesis.</a> Analytical chemistry. 89 (4), 2250-2258 (2017).</li><li>Liu, X. R., Zhang, M. M., Rempel, D. L., Gross, M. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+Single+Approach+Reveals+the+Composite+Conformational+Changes,+Order+of+Binding,+and+Affinities+for+Calcium+Binding+to+Calmodulin.">A Single Approach Reveals the Composite Conformational Changes, Order of Binding, and Affinities for Calcium Binding to Calmodulin.</a> Analytical Chemistry. 91 (9), 5508-5512 (2019).</li><li>Kiselar, J. G., Janmey, P. A., Almo, S. C., Chance, M. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Structural+analysis+of+gelsolin+using+synchrotron+protein+footprinting.">Structural analysis of gelsolin using synchrotron protein footprinting.</a> Molecular and Cellular Proteomics. 2 (10), 1120-1132 (2003).</li><li>Chea, E. E., Deredge, D. J., Jones, L. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Insights+on+the+Conformational+Ensemble+of+Cyt+C+Reveal+a+Compact+State+during+Peroxidase+Activity.">Insights on the Conformational Ensemble of Cyt C Reveal a Compact State during Peroxidase Activity.</a> Biophysical Journal. 118 (1), 128-137 (2020).</li><li>Poor, T. A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Probing+the+paramyxovirus+fusion+(F)+protein-refolding+event+from+pre-+to+postfusion+by+oxidative+footprinting.">Probing the paramyxovirus fusion (F) protein-refolding event from pre- to postfusion by oxidative footprinting.</a> Proceedings of the National Academy of Sciences U S A. 111 (25), 2596-2605 (2014).</li><li>Roush, A. E., Riaz, M., Misra, S. K., Weinberger, S. R., Sharp, J. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Intrinsic+Buffer+Hydroxyl+Radical+Dosimetry+Using+Tris(hydroxymethyl)aminomethane.">Intrinsic Buffer Hydroxyl Radical Dosimetry Using Tris(hydroxymethyl)aminomethane.</a> Journal of American Society of Mass Spectrometry. 31 (2), 169-172 (2020).</li><li>Everett, E. A., Falick, A. M., Reich, N. O. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Identification+of+a+critical+cysteine+in+EcoRI+DNA+methyltransferase+by+mass+spectrometry.">Identification of a critical cysteine in EcoRI DNA methyltransferase by mass spectrometry.</a> Journal of Biological Chemistry. 265 (29), 17713-17719 (1990).</li><li>Sanderson, R. J., Mosbaugh, D. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Identification+of+specific+carboxyl+groups+on+uracil-DNA+glycosylase+inhibitor+protein+that+are+required+for+activity.">Identification of specific carboxyl groups on uracil-DNA glycosylase inhibitor protein that are required for activity.</a> Journal of Biological Chemistry. 271 (46), 29170-29181 (1996).</li></ol>

E.E.C., J.S.S., and S.R.W. disclose a significant financial interest in GenNext Technologies, Inc., an early stage company seeking to commercialize technologies for higher-order protein structure analysis. The representative data provided has been reviewed by S.K.M., who has no financial conflict of interest.

There are several critical steps to ensure proper labeling of proteins during any HRPF experiment. First, an appropriate flow rate and source flash rate are selected to make certain each bolus of the sample is irradiated once. This ensures that the protein is exposed to a single bolus of newly formed &#9642;OH. Once a protein is oxidized, the higher order protein structure can be altered. To be confident the native protein structure is probed, each protein molecule must be modified in a single instant. Dosimet...

Protein conformation and associated higher order structure (HOS) are the principal determinants of proper biological function and aberrant behavior1. The same applies to biopharmaceuticals, whose structure and functional activity is dependent on various aspects of their production and environment. Biopharmaceutical change in HOS have been linked to adverse drug reactions (ADR) attributed to undesirable pharmacology and patient immunological response2,3. The appearance of ADRs has alerted the biopharmaceutical industry to the critical role that protein HOS plays in the safety and efficacy of biotherapeutics, and they have established the need for new and improved HOS analytics4.
Hydroxyl Radical Protein Footprinting (HRPF) is a promising technique to track the change in protein HOS. HRPF involves the irreversible labeling of a protein&#8217;s exterior with &#9642;OH followed with mass spectrometry (MS) analysis to identify the solvent accessible surface of the protein5,6,7. HRPF has successfully been used to detect defects in protein HOS and its function8,9, characterize the HOS of monoclonal antibodies (mAb)10,11,12,13, determine the binding Kd of a ligand14, and much more15,16,17,18,19. A common method to generate the &#9642;OH for HRPF is Fast Photochemical Oxidation of Proteins (FPOP), which employs high-energy, fast UV lasers to produce &#9642;OH from photolysis of H2O2. For the most part, FPOP uses expensive excimer lasers employing hazardous gas (KrF) that demands substantial safeguards to avoid respiratory and eye injury20. To avoid inhalation hazards, others have used frequency quadrupled neodymium yttrium aluminum garnet (Nd:YAG) lasers21, which eliminates the use of toxic gas but are still costly, require significant operational expertise, and demand extensive stray light controls to protect users from eye injury.
Although ample information can be obtained using HRPF, broad adoption in biopharma has not been met. Two barriers for the limited HRPF adoption include: 1) the use of dangerous and expensive lasers that demand substantial safety precautions20; and 2) the irreproducibility of HRPF caused by background scavenging of &#9642;OH that limit comparative studies22. To supplant laser use, a high-speed, high energy plasma flash photolysis unit was developed to safely perform FPOP in a facile manner. To improve on the irreproducibility of HRPF experiments, real-time radical dosimetry is implemented.
The practice of HRPF has been limited by irreproducibility attributed to background scavenging of &#9642;OH22. While &#9642;OH are excellent probes of protein topography, they also react with many constituents found in preparations, making it necessary to measure the effective concentration of radical available to oxidize a target protein. Variations in buffer preparation, hydrogen peroxide concentration, ligand properties, or photolysis can result in oxidation differences between control and experimental groups that create ambiguity in HOS differential studies. The addition of real-time radical dosimetry enables the adjustment of the effect &#9642;OH load and therefore increases the confidence and reproducibility during an HRPF experiment. The use of radical dosimetry in FPOP has been described elsewhere23,24,25, and is further discussed in detail in a recent publication26. Here, we describe the use of a novel flash photolysis system and real-time dosimetry to label equine apo-myoglobin (aMb), comparing levels of peptide oxidation in an FPOP experiment to that of obtained when using an excimer laser.

<table><tbody><tr><td>15 mL Conical Centrifuge Tubes</td><td>Fisher Scientific</td><td>14-959-53A</td><td>any brand is sufficient</td></tr><tr><td>50 &amp;micro;L SGE Gastight Syringes</td><td>Fisher Scientific</td><td>SG-00723</td><td></td></tr><tr><td>Acclaim PepMap 100 C18 nanocolumn (0.75 mm X 150 mm, 2 &amp;micro;m)</td><td>Thermo Scientific</td><td></td><td></td></tr><tr><td>Acetonitrile with 0.1% Formic Acid (v/v), LC/MS Grade</td><td>Fisher Scientific</td><td>LS120-500</td><td></td></tr><tr><td>Apomyoglobin</td><td>Sigma-Aldrich</td><td></td><td></td></tr><tr><td>Catalase</td><td>Sigma-Aldrich</td><td>C9322</td><td></td></tr><tr><td>Centrifuge</td><td>Eppendorf</td><td>022625501</td><td></td></tr><tr><td>Delicate Task Wipers</td><td>Fisher Scientific</td><td>06-666A</td><td></td></tr><tr><td>Hydrogen Peroxide</td><td>Fisher Scientific</td><td>H325-100</td><td>any 30% hydrogen peroxide is sufficient</td></tr><tr><td>Methionine amide</td><td>Chem-Impex</td><td>03109</td><td></td></tr><tr><td>Microcentrifuge</td><td>Thermo Scientific</td><td>75002436</td><td></td></tr><tr><td>Orbitrap Fusion Lumos Tribrid Mass Spectrometer</td><td>Thermo Scientific</td><td>Orbitrap Fusion Lumos Tribrid Mass Spectrometer</td><td>other high resolution instruments (e.g. Q exactive Orbitrap or Orbitrap Fusion) can be used</td></tr><tr><td>Pierce Trypsin Protease, MS Grade</td><td>Thermo Scientific</td><td>90058</td><td></td></tr><tr><td>Polymicro Cleaving Stone, 1&quot; x 1&quot; x 1/32&amp;rdquo;</td><td>Molex</td><td>1068680064</td><td>any capillary tubing cutter is sufficient</td></tr><tr><td>UPLC</td><td>Thermo Scientific</td><td></td><td></td></tr><tr><td>Water with 0.1% Formic Acid (v/v), LC/MS Grade</td><td>Fisher Scientific</td><td>LS118-500</td><td></td></tr><tr><td>Water, LC/MS Grade</td><td>Fisher Scientific</td><td>W6-4</td><td></td></tr></tbody></table>

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.

The high-pressure plasma source coupled with real-time dosimetry allows better control of &#9642;OH yield to observe changes in higher-order protein structure more accurately. The addition of adenine allows for an effective real-time radical dosimeter. Upon oxidation, adenine loses UV absorbance at 265 nm (Figure 2A). The change in adenine absorbance is directly related to the concentration of radicals available for HRPF thus providing a means to effectively monitor changes in rad...

Watch this Scientific Journal Video about Laser-free Hydroxyl Radical Protein Footprinting to Perform Higher Order Structural Analysis of Proteins at JoVE.com

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

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 ...

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3.6K Views.  GenNext Technologies Inc.. Laservrije hydroxylradicale eiwitvoetafdruk maakt het identificeren van eiwitinteractieplaatsen en gebieden van bevestigingsverandering veel gemakkelijker, waardoor onderzoek in eiwitaggregatie-, structuur- en stabiliteitsstudies wordt versneld. Met inline radicale dosimetrie kunnen gebruikers de effectieve hydroxylradicale belasting in realtime aanpassen, en met realtime radicale dosimetrie kunt u experimentele tijd en kostbaar monster besparen terwijl de reproduceerbaarheid van etiketten wo...

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

Matrix-assisted Laser Desorption/Ionization Time of Flight (MALDI-TOF) Mass Spectrometric Analysis of Intact Proteins Larger than 100 kDa

Effectively determining masses of proteins is critical to many biological studies (e.g. for structural biology investigations). Accurate mass determination allows one to evaluate the correctness of protein primary sequences, the presence of mutations and/or post-translational modifications, the possible protein degradation, the sample homogeneity, and the degree of isotope incorporation in case of labelling (e.g. 13C labelling).
Electrospray ionization (ESI) mass spectrometry (MS) is widely used for mass determination of denatured proteins, but its efficiency is affected by the composition of the sample buffer. In particular, the presence of salts, detergents, and contaminants severely undermines the effectiveness of protein analysis by ESI-MS. Matrix-assisted laser desorption/ionization (MALDI) MS is an attractive alternative, due to its salt tolerance and the simplicity of data acquisition and interpretation. Moreover, the mass determination of large heterogeneous proteins (bigger than 100 kDa) is easier by MALDI-MS due to the absence of overlapping high charge state distributions which are present in ESI spectra.
Here we present an accessible approach for analyzing proteins larger than 100 kDa by MALDI-time of flight (TOF). We illustrate the advantages of using a mixture of two matrices (i.e. 2,5-dihydroxybenzoic acid and &#945;-cyano-4-hydroxycinnamic acid) and the utility of the thin layer method as approach for sample deposition. We also discuss the critical role of the matrix and solvent purity, of the standards used for calibration, of the laser energy, and of the acquisition time. Overall, we provide information necessary to a novice for analyzing intact proteins larger than 100 kDa by MALDI-MS.

Matrix-assisted Laser Desorption/Ionization Time of Flight MALDI-TOF Mass Spectrometric Analysis of Intact Proteins Larger than 100 kDa

Accurate mass measurement represents an important step during the investigation of proteins. Matrix-assisted laser desorption/ionization (MALDI) mass spectrometry (MS) can be used for such determination and its key advantage is the tolerance to salts, detergents and contaminants. Here we illustrate an accessible approach for the analysis of proteins larger than 100 kDa by MALDI-MS.

Effectively determining masses of proteins is critical to many biological studies (e.g. for structural biology investigations). Accurate mass ...

Chemistry

Analyzing Protein Dynamics Using Hydrogen Exchange Mass Spectrometry

All cellular processes depend on the functionality of proteins. Although the functionality of a given protein is the direct consequence of its unique amino acid sequence, it is only realized by the folding of the polypeptide chain into a single defined three-dimensional arrangement or more commonly into an ensemble of interconverting conformations. Investigating the connection between protein conformation and its function is therefore essential for a complete understanding of how proteins are able to fulfill their great variety of tasks. One possibility to study conformational changes a protein undergoes while progressing through its functional cycle is hydrogen-1H/2H-exchange in combination with high-resolution mass spectrometry (HX-MS). HX-MS is a versatile and robust method that adds a new dimension to structural information obtained by e.g. crystallography. It is used to study protein folding and unfolding, binding of small molecule ligands, protein-protein interactions, conformational changes linked to enzyme catalysis, and allostery. In addition, HX-MS is often used when the amount of protein is very limited or crystallization of the protein is not feasible. Here we provide a general protocol for studying protein dynamics with HX-MS and describe as an example how to reveal the interaction interface of two proteins in a complex. &#160;&#160;

Protein conformation and dynamics are key to understanding the relationship between protein structure and function. Hydrogen exchange coupled with high-resolution mass spectrometry is a versatile method to study the conformational dynamics of proteins as well as characterizing protein-ligand and protein-protein interactions, including contact interfaces and allosteric effects.

All cellular processes depend on the functionality of proteins. Although the functionality of a given protein is the direct consequence of its unique ...

Proteome-wide Quantification of Labeling Homogeneity at the Single Molecule Level

Cell proteomes are often characterized using electrophoresis assays, where all species of proteins in the cells are non-specifically labeled with a fluorescent dye and are spotted by a photodetector following their separation. Single molecule fluorescence imaging can provide ultrasensitive protein detection with its ability for visualizing individual fluorescent molecules. However, the application of this powerful imaging method to electrophoresis assays is hampered by the lack of ways to characterize the homogeneity of fluorescent labeling of each protein species across the proteome. Here, we developed a method to evaluate the labeling homogeneity across the proteome based on a single molecule fluorescence imaging assay. In our measurement using a HeLa cell sample, the proportion of proteins labeled with at least one dye, which we termed &#8216;labeling occupancy&#8217; (LO), was determined to range from 50% to 90%, supporting the high potential of the application of single molecule imaging to sensitive and precise proteome analysis.

Here, we present a protocol to assess the labeling homogeneity for each protein species in a complex protein sample at the single molecule level.

Cell proteomes are often characterized using electrophoresis assays, where all species of proteins in the cells are non-specifically labeled with a ...

In Vivo Hydroxyl Radical Protein Footprinting for the Study of Protein Interactions in Caenorhabditis elegans

Fast oxidation of proteins (FPOP) is a hydroxyl radical protein footprinting (HRPF) method used to study protein structure, protein-ligand interactions, and protein-protein interactions. FPOP utilizes a KrF excimer laser at 248 nm for photolysis of hydrogen peroxide to generate hydroxyl radicals which in turn oxidatively modify solvent-accessible amino acid side chains. Recently, we expanded the use of FPOP of in vivo oxidative labeling in Caenorhabditis elegans (C. elegans), entitled IV-FPOP. The transparent nematodes have been used as model systems for many human diseases. Structural studies in C. elegans by IV-FPOP is feasible because of the animal&#8217;s ability to uptake hydrogen peroxide, their transparency to laser irradiation at 248 nm, and the irreversible nature of the modification. The assembly of a microfluidic flow system for IV-FPOP labeling, IV-FPOP parameters, protein extraction, and LC-MS/MS optimized parameters are described herein.

In Vivo Hydroxyl Radical Protein Footprinting for the Study of Protein Interactions in Caenorhabditis elegans

In vivo fast photochemical oxidation of proteins (IV-FPOP) is a hydroxyl radical protein footprinting technique that allows for mapping of protein structure in their native environment. This protocol describes the assembly and set-up of the IV-FPOP microfluidic flow system.

Fast oxidation of proteins (FPOP) is a hydroxyl radical protein footprinting (HRPF) method used to study protein structure, protein-ligand interactions, ...

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. ...

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.

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 ...

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 ...

Covalent Labeling with Diethylpyrocarbonate for Studying Protein Higher-Order Structure by Mass Spectrometry

Characterizing a protein&#39;s higher-order structure is essential for understanding its function. Mass spectrometry (MS) has emerged as a powerful tool for this purpose, especially for protein systems that are difficult to study by traditional methods. To study a protein&#39;s structure by MS, specific chemical reactions are performed in solution that encode a protein&#39;s structural information into its mass. One particularly effective approach is to use reagents that covalently modify solvent accessible amino acid side chains. These reactions lead to mass increases that can be localized with residue-level resolution when combined with proteolytic digestion and tandem mass spectrometry. Here, we describe the protocols associated with use of diethylpyrocarbonate (DEPC) as a covalent labeling reagent together with MS detection. DEPC is a highly electrophilic molecule capable of labeling up to 30% of the residues in the average protein, thereby providing excellent structural resolution. DEPC has been successfully used together with MS to obtain structural information for small single-domain proteins, such as &#946;2-microglobulin, to large multi-domain proteins, such as monoclonal antibodies.

The experimental procedures for performing diethylpyrocarbonate-based covalent labeling with mass spectrometric detection are described. Diethylpyrocarbonate is simply mixed with the protein or protein complex of interest, leading to the modification of solvent accessible amino acid residues. The modified residues can be identified after proteolytic digestion and liquid chromatography/mass spectrometry analysis.

Characterizing a protein&#39;s higher-order structure is essential for understanding its function. Mass spectrometry (MS) has emerged as a powerful tool ...

Enrichment of Bacterial Lipoproteins and Preparation of N-terminal Lipopeptides for Structural Determination by Mass Spectrometry

Lipoproteins are important constituents of the bacterial cell envelope and potent activators of the mammalian innate immune response. Despite their significance to both cell physiology and immunology, much remains to be discovered about novel lipoprotein forms, how they are synthesized, and the effect of the various forms on host immunity. To enable thorough studies on lipoproteins, this protocol describes a method for bacterial lipoprotein enrichment and preparation of N-terminal tryptic lipopeptides for structural determination by matrix-assisted laser desorption ionization-time of flight mass spectrometry (MALDI-TOF MS). Expanding on an established Triton X-114 phase partitioning method for lipoprotein extraction and enrichment from the bacterial cell membrane, the protocol includes additional steps to remove non-lipoprotein contaminants, increasing lipoprotein yield and purity. Since lipoproteins are commonly used in Toll-like receptor (TLR) assays, it is critical to first characterize the N-terminal structure by MALDI-TOF MS. Herein, a method is presented to isolate concentrated hydrophobic peptides enriched in N-terminal lipopeptides suitable for direct analysis by MALDI-TOF MS/MS. Lipoproteins that have been separated by Sodium Dodecyl Sulfate Poly-Acrylamide Gel Electrophoresis (SDS-PAGE) are transferred to a nitrocellulose membrane, digested in situ with trypsin, sequentially washed to remove polar tryptic peptides, and finally eluted with chloroform-methanol. When coupled with MS of the more polar trypsinized peptides from wash solutions, this method provides the ability to both identify the lipoprotein and characterize its N-terminus in a single experiment. Intentional sodium adduct formation can also be employed as a tool to promote more structurally informative fragmentation spectra. Ultimately, enrichment of lipoproteins and determination of their N-terminal structures will permit more extensive studies on this ubiquitous class of bacterial proteins.

The enrichment of bacterial lipoproteins using a non-ionic surfactant phase partitioning method is described for direct use in TLR assays or other applications. Further steps are detailed to prepare N-terminal tryptic lipopeptides for structural characterization by mass spectrometry.

Lipoproteins are important constituents of the bacterial cell envelope and potent activators of the mammalian innate immune response. Despite their ...

Immunology and Infection

Monitoring Equilibrium Changes in RNA Structure by 'Peroxidative' and 'Oxidative' Hydroxyl Radical Footprinting

RNA molecules play an essential role in biology. In addition to transmitting genetic information, RNA can fold into unique tertiary structures fulfilling a specific biologic role as regulator, binder or catalyst. Information about tertiary contact formation is essential to understand the function of RNA molecules. Hydroxyl radicals (&bull;OH) are unique probes of the structure of nucleic acids due to their high reactivity and small size.1 When used as a footprinting probe, hydroxyl radicals map the solvent accessible surface of the phosphodiester backbone of DNA1 and RNA2 with as fine as single nucleotide resolution. Hydroxyl radical footprinting can be used to identify the nucleotides within an intermolecular contact surface, e.g. in DNA-protein1 and RNA-protein complexes. Equilibrium3 and kinetic4 transitions can be determined by conducting hydroxyl radical footprinting as a function of a solution variable or time, respectively. A key feature of footprinting is that limited exposure to the probe (e.g., 'single-hit kinetics') results in the uniform sampling of each nucleotide of the polymer.5
In this video article, we use the P4-P6 domain of the Tetrahymena ribozyme to illustrate RNA sample preparation and the determination of a Mg(II)-mediated folding isotherms. We describe the use of the well known hydroxyl radical footprinting protocol that requires H2O2 (we call this the 'peroxidative' protocol) and a valuable, but not widely known, alternative that uses naturally dissolved O2 (we call this the 'oxidative' protocol). An overview of the data reduction, transformation and analysis procedures is presented.

This protocol describes how to quantify the Mg(II)-dependent formation of RNA tertiary structure by two methods of hydroxyl radical footprinting.

RNA molecules play an essential role in biology. In addition to transmitting genetic information, RNA can fold into unique tertiary structures fulfilling ...

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

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