This work was supported by the NSF CAREER Award (MCB1701692) for LMJ.

1. Set up IC-FPOP flow system
<ol>
	<li>To begin the assembly of the flow system, cut the fused silica using a cleavage stone to size. The IC-FPOP flow system requires four 12 cm and one 17 cm fused silica with an inner diameter (ID) of 450 &#181;m and outer diameter (OD) of 670 &#181;m, two 24 cm and one 40 cm with an ID of 75 &#181;m and an OD of 360 &#181;m, and finally two 57 cm with an ID of 150 &#181;m and an OD of 360 &#181;m. 
	NOTE: When cutting the silica tubing, gently scrape away the polyimide coating, and bend to get the cleanest cut. Check to make sure it is a straight cut (this is necessary to ensure no blockages or leaks form).</li>
	<li>Set up 15 connections using nano-tight sleeves (0.0155&#34; ID X 1/16&#34; OD for 360 &#181;m OD silica tubing and 0.027&#34; ID X 1/16&#34; OD for the 670 &#181;m OD silica tubing) with super flangeless nut PEEK 1/4-28 flat-bottom for 1/16&#34; OD and super flangeless ferrule w/SST ring, Tefzel (ETFE), 1/4-28 flat-bottom, for 1/16&#34; OD. Construct connections as per the manufacturer&#39;s protocol (Figure 1).</li>
	<li>Place 6 small cylindrical magnets in one 500 &#181;L syringe. Fill this syringe along with another 500 &#181;L syringe and two 5 mL syringes with buffer and remove air. Position on syringe pump as shown on Figure 2A. 
	NOTE: The 5 mL syringes are larger than the 500 &#181;L syringes, so a spacer is needed to tighten all the syringes simultaneously in place (Figure 2B).</li>
	<li>Tighten syringe pump stopper so that the cell syringe has roughly 50 &#181;L left when the motor stalls. This will leave room for the magnetic stirrers. (Figure 2C-D). 
	CAUTION: If the syringe pump puts pressure on the magnets, they will jam the syringe and can cause the syringe to break.</li>
	<li>Using a Luer adapter, connect a manual valve to each syringe. Assemble the silica tubing as shown in Figure 3. 
	NOTE: Thread the line with the cells + H2O2 all the way through the cross to the other side. Then insert it into the 450 &#181;m ID silica tubing. The sheath buffers in the 5 mL syringe will be flowing at a faster rate than the cells and H2O2. With the sheath buffers on both sides, the cells will be hydrodynamically focused into a single line for irradiation.</li>
	<li>Position flow system next to laser. Using a lighter, burn away the silica coating on the 450 &#181;m ID tubing to make a window for laser irradiation.</li>
	<li>Place a magnetic stirrer above the cell syringe containing the six magnets.</li>
	<li>Set syringe pump to 492.4 &#181;L/min for a final flow rate of 1,083.3 &#181;L/min. Flow buffer through the system three times to flush the system and test for any leaks.</li>
	<li>Focus the excimer laser onto the silica tubing using a convex lens. Once focused, test the irradiation window by placing a small piece of paper behind the silica tubing and turn the laser on. Measure the region burnt from the irradiation. Calculate the needed laser frequency by using the irradiation window and flow rate to obtain an exclusion fraction of zero. 
	NOTE: The recommended laser energy for an IC-FPOP experiment is &#8805;120 mJ. To get an exclusion fraction of 0 with an irradiation window of 2.58 mm and a flow rate of 1083.3 &#181;L/min, the frequency needs to be 44. Below are the equations that are needed to calculate the laser frequency if the irradiation window, flow rate, and exclusion fraction is known. 
	<img src="/files/ftp_upload/60911/60911eq1.jpg" /> 
	where w is the volume in nL, x is the laser spot width in mm, y is the silica tubing ID in &#181;m, v is the total volume in &#181;L, b is the exclusion fraction, a is the flow rate in &#181;L/min, and f is the frequency in Hz.</li>
</ol>
2. Make quench and H2O2
<ol>
	<li>Make quench containing 100 mM N-tert-Butyl-alpha-phenylnitrone (PBN) and 100 mM N,N&#39;-dimethylthiourea (DMTU). Aliquot 11 mL of quench for each sample into a 50 mL conical tube.</li>
	<li>Dilute H2O2 to 200 mM. Each sample requires 500 &#181;L of H2O2. 
	NOTE: The quench can be made the day before and stored at 4 &#176;C overnight, protected from light. H2O2 should be made fresh the day of experimentation.</li>
</ol>
3. Collect cells
<ol>
	<li>Grow cells in a T175 flask to about 70-90% confluency.</li>
	<li>Remove media and rinse with buffer. 
	NOTE: Typical buffers to use are cell culture grade Dulbecco&#39;s phosphate-buffered saline (DPBS) or Hank&#39;s balanced salt solution (HBSS).</li>
	<li>Detach cells using either trypsin-EDTA or with a scraper.</li>
	<li>Once detached resuspend in 10 mL of buffer and count the cells.</li>
	<li>Spin down, remove the buffer and trypsin-EDTA, and resuspend to make 2 x 106 cells/mL.</li>
	<li>Aliquot 500 &#181;L of the cells per sample. 
	NOTE: For each condition, make a minimum of 3 laser samples, and 3 no laser controls.</li>
</ol>
4. Performing IC-FPOP
<ol>
	<li>Fill the two 5 mL syringes with buffer, the 500 &#181;L syringe containing the magnets with the cells, and the final 500 &#181;L syringe with H2O2. Turn the magnetic stirrer on.</li>
	<li>Spike in 220 &#181;L of dimethyl sulfoxide (DMSO) to one aliquot of quench, gently mix, and place behind flow system to collect irradiated samples. The addition of DMSO will inhibit endogenous methionine sulfoxide reductase.</li>
	<li>Turn on laser, wait 7 s, and then turn on flow system.</li>
	<li>Once the sample finishes flowing, turn the laser off, and gently mix the quench with the collected sample. Place this to the side during steps 4.5 and 4.6.</li>
	<li>Fill all four syringes with the buffer the cells are suspended in and flow it through the flow system.</li>
	<li>After the system finishes flushing, repeat steps 4.1 and 4.2. Start the flow without irradiation. This is the no laser control to account for background oxidation in the cells.</li>
	<li>While the next sample is running, spin down the previous sample at 450-800 x g for 5 min, remove the solvent, and resuspend in 100 &#181;L of a cell lysis buffer like radioimmunoprecipitation assay (RIPA) buffer.</li>
	<li>Transfer the sample to a microcentrifuge tube and flash freeze in liquid nitrogen.</li>
	<li>When all samples have finished running, disassemble the flow system for cleaning. Discard the used silica tubing and clean all the other connections by sonicating for 1 h in 50% water: 50% methanol. Clean the syringes as per manufacturer&#39;s instructions.</li>
</ol>
5. Digest
<ol>
	<li>Digest the whole cell lysate. Begin by thawing the samples and heat at 95 &#176;C for 10 min.</li>
	<li>After heating, cool the lysate on ice for 15 min.</li>
	<li>Add 25 units of nuclease to digest DNA and RNA and incubate at room temperate for 15 min.</li>
	<li>Spin samples using a table-top centrifuge at 16,000 x g for 10 min at 4 &#176;C.</li>
	<li>Collect the supernatant and transfer it to a clean microcentrifuge tube.</li>
	<li>Check the protein concentration using a protein assay kit.</li>
	<li>Transfer 20-100 &#181;g of sample to a clean microcentrifuge tube and bring to 100 &#181;L.</li>
	<li>Reduce samples with 20 mM dithiothreiotol (DTT) at 50 &#176;C for 45 min.</li>
	<li>Cool the samples at room temperature for 15 min.</li>
	<li>Alkylate with 20 mM iodoacetamide (IAA) at room temperature for 20 min protected from light.</li>
	<li>Add pre-chilled acetone at a 1:4 ratio protein: acetone. Mix samples and place in -20 &#176;C overnight.</li>
	<li>The next morning, spin samples at 16,000 x g for 10 min at 4 &#176;C.</li>
	<li>Remove the supernatant and add 50 &#181;L of 90% pre-chilled acetone. Mix samples and spin down at 16,000 x g for 5 min at 4 &#176;C.</li>
	<li>Remove acetone and let samples dry by leaving the caps of the microcentrifuge open with a lint free wipe covering the top. After the samples have dried, resuspend protein pellet with 10 mM Tris buffer pH 8.</li>
	<li>Resuspend 20 &#181;g of trypsin in 40 &#181;L of 10 mM Tris buffer pH 8. Add 2 &#181;g of trypsin (mass: mass ratio of 1 trypsin: 50 sample). Incubate samples at 37 &#176;C overnight.</li>
	<li>The next morning check the peptide concentration using a peptide assay. After the sample has been removed for the peptide assay, quench the trypsin digestion by adding formic acid to the samples (final concentration is 5% formic acid).</li>
	<li>Once final peptide concentration is determined, transfer 10 &#181;g of each sample to a clean microcentrifuge tube. This ensures the same amount of each sample is analyzed. Dry the sample using a vacuum centrifuge. Once dried resuspend with 20 &#181;L of mass spectrometry grade 0.1% formic acid. Transfer samples to autosampler vials.</li>
</ol>
6. Liquid Chromatography-Tandem Mass Spectrometry
<ol>
	<li>To localize FPOP modifications analyze the digested cell lysate using LC-MS/MS analysis.</li>
	<li>Use mobile phases of 0.1% formic acid in water (A) and 0.1% formic acid in acetonitrile (B).</li>
	<li>Load 0.5 &#181;g of sample onto a 180 &#181;m x 20 mm C18 (5 &#181;m and 100 &#197;) trapping column and wash sample with 99% (A) and 1% (B) for 15 min.</li>
	<li>Using a 75 &#181;m x 30 cm C18 (5 &#181;m and 125 &#197;) analytical column, run the analytical separation method with a flow rate of 0.300 &#181;L/min starting at 3% (B) for one min then ramp to 10% (B) from 1-2 min. Next ramp to 45% (B) from 2-100 min then 100% (B) from 100-110 min. Clean the column by holding at 100% (B) from 110-115 min. Recondition the column by ramping down to 3% (B) from 115-116 min and hold at 3% (B) from 116-130 min.</li>
	<li>Set the MS acquisition method to have a resolution of 60,000 with an m/z scan range of 375-1500. Set the automatic gain control (AGC) target to 5.0 x 105 with a maximum injection time of 50 ms.</li>
	<li>During the MS acquisition, select the precursor ions with charge states 2-6 for isolation via data dependent acquisition (DDA) with an isolation window of 1.2 m/z and a cycle time of 4 s. Select the peptides with an intensity threshold of 5.0 x 104 for HCD activation with a normalized energy set to 32%. Exclude the peptides after 1 MS/MS acquisition for 60 s. Set the MS/MS resolution to 15,000 with an AGC target of 5.0 x 104 and a maximum injection time of 35 ms.</li>
</ol>
7. Data Processing
<ol>
	<li>Search the RAW files on an available protein analysis software against a relevant protein database and the relevant digest enzyme. Here, use the Swiss-Prot Homo Sapiens database and trypsin.</li>
	<li>Set the precursor mass to search between 350 to 5,000 Da and a mass tolerance of 10 ppm. There can be at most 1 missed cleavage site with a peptide length between 6 to 144 residues. These limitations facilitate database searching.</li>
	<li>Set the fragment ions maximum mass tolerance to 0.02 Da with the carbamidomethyl (+57.021) as a static modification and all FPOP modifications from 17 amino acids as a dynamic modification (Figure 4 is an example of the protein analysis workflow used to detect FPOP modifications). 
	NOTE: FPOP modifications on serine and threonine are not included in the search due to their lower reactivity with hydroxyl radicals.</li>
	<li>Search with a decoy database with a false discovery rate of 1% and 5%.</li>
	<li>Once files are finished searching calculate the extent of FPOP modification. Open Proteome Discoverer 2.2, export sequence, modification locations, protein accession, spectrum file, precursor abundance, and retention time information. Calculate the extent of modification from equation 4: 
	<img src="/files/ftp_upload/60911/60911eq2.jpg" /> 
	The EIC area modified is the chromatographic area of a specific peptide with a hydroxyl radical modification. The EIC area is the total chromatographic area of both modified and unmodified areas of that specific peptide. The extent of oxidation is calculated in the samples with and without irradiation. The sample omitting irradiation (control) accounts for any background oxidation that could have been present in the cells before and after H2O2 exposure. The controls are subtracted from the irradiation samples to help isolate FPOP specific modifications.</li>
</ol>

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Lisa Jones is an inventor on the flow assembly for cells patent (publication number: 20180079998).

Several mass spectrometry-based techniques have been developed to study protein structure and protein-ligand complexes in a proteome-wide manner in whole cells or cell lysates. These techniques include but are not limited to stability of proteins from rate of oxidation (SPROX), thermal proteome profiling (TPP), chemical cross-linking (XL-MS), and hydroxyl radical protein footprinting (HRPF). Each technique has unique limitations and advantages compared to one another, which have been extensively reviewed12. Each of these methods have been used for proteome wide structural biology to elucidate protein structure and ultimately function within the complex cellular environment. IC-FPOP is a HRPF technique that utilizes hydroxyl radicals to oxidatively modify solvent exposed side chains of amino acids, probing protein structure and protein-ligand interactions within viable cells13. IC-FPOP is an improvement to initial HRPF in live cells that used Fenton chemistry to generate radicals on the minutes timescale14. In this study, structural changes in an integral membrane protein in response to lowering the pH or ionic strength of the buffer were successfully characterized with good oxidation coverage across the protein. Compared to Fenton chemistry, IC-FPOP is much faster, modifying proteins on the microsecond timescale, thus enabling the native protein conformation to be studied.
A key test for IC-FPOP is to confirm the viability of the cells following exposure to H2O2. Using a 40 cm mixing line, the cells are incubated in H2O2 for roughly 3 s before irradiation. This time can be adjusted by changing the length of this silica tubing. It is noteworthy that although the use of trypan blue to test cell viability shows the integrity of the cells are sustained following H2O2 incubation, the cells could potential be under stress effecting signaling pathways that interact with H2O2. Fortunately, the short incubation time is faster than protein synthesis providing confidence the proteins present are not induced by H2O2.
The next important step is to confirm proper assembly of the flow system. Once assembled, ensure there are no leaks present after flushing the system with the desired buffer. If leaks are present, make sure that the silica tubing was cut properly and is flush against the ferrule to make a proper seal once tightened down. All parts are hand-tightened, so no tools are necessary. During each IC-FPOP experiment, make sure that the magnetic stirrers in the cell syringe remain in motion. This small agitation limits the number of cells that settle at the bottom of the syringe but is not harsh enough to shear the cells. After one run, there is roughly 50 &#181;L of cells left in the syringe. Always make sure to dilute this out with a rinsing step to limit the number of cells that carry over to the next experiment. It is recommended to use a fresh cell syringe if multiple cell treatments are being compared. It is also important to select an appropriate buffer for the cells being tested. Some buffers quench the hydroxyl radical leading to fewer modifications on proteins. Xu et al. have shown that some commonly used buffers decrease the hydroxyl radical lifetime11. DPBS and HBSS are common buffers used for IC-FPOP experiments.
Following IC-FPOP, optimize the digestion protocol based on the parameters needed. Since FPOP produces irreversible covalent modifications, there is ample time available for a thorough digestion and clean-up without losing the labeling coverage. Always test and normalize the protein concentration so uniform peptide concentrations are loaded for tandem mass spectrometry. Finally, be mindful that an immunoprecipitation cannot be performed in conjunction with IC-FPOP. If an FPOP modification targets the region of interaction the affinity of the antibody will be lowered. To help increase the identification of FPOP modifications, 2D-chromatographic separation steps have shown to more than triple the number of oxidized peptides detected15.
A challenge of any FPOP experiment is the complicated level of data analysis due to the possible oxidation products that can arise. This is true for both in-cell or in vitro but is drastically increased with the added complexity of analyzing cell lysates. With further optimization of IC-FPOP more proteins with higher modification coverages are arising, thus expeditiously making analysis more arduous. The shear amount of data generated from a single IC-FPOP experiment limits the use of manual validation causing researchers to rely more heavily on software. Due to this, Rinas et al. developed a quantitation strategy for HRPF using Proteome Discoverer (PD)16. This method utilizes a multi-search node workflow on PD combined with a quantitative platform in a spreadsheet. Further improvements on the IC-FPOP platform are underway to increase the number of identified peptides with FPOP modifications along with increased reproducibility and quantitation accuracy.

Hydroxyl radical protein footprinting (HRPF) is a method that probes the solvent accessibility of a protein through covalent modifications produced from hydroxyl radicals. When protein structure or protein interactions change, it will alter the solvent exposure of amino acids, thus altering the extent of modification of residues. With HRPF, protein interactions1,2,3 and protein conformational changes4,5,6 have successfully been interrogated in vitro. There are several methods that generate hydroxyl radicals for HRPF experiments, one being fast photochemical oxidation of proteins (FPOP). FPOP was developed by Hambly and Gross in 2005 and utilizes a 248 nm excimer laser to produce hydroxyl radicals through the photolysis of hydrogen peroxide (H2O2)7.
Recently, Espino et al. extended the use of FPOP to probe protein structure in live cells, a method termed in-cell FPOP (IC-FPOP)8. In contrast to in vitro studies, studying proteins in cells accounts for molecular crowding along with various protein interactions that could potentially influence structure. Additionally, it presents the advantage of providing a snapshot of the full proteome potentially providing structural information of numerous systems at once to perform proteome wide structural biology. Furthermore, this technique is ideal for proteins that are difficult to study in vitro, like membrane proteins.
Initial studies of IC-FPOP successfully probed 105 proteins ranging in protein abundance and cellular localization. To improve the IC-FPOP method, Rinas et al. developed a microflow system for single cell flow9. The enhancement of the original flow system limits cell aggregation and increases the available H2O2 available for irradiation. In the initial flow system, cells clumping in the silica tubing resulted in clogs and uneven irradiation. The incorporation of two streams of a sheath buffer hydrodynamically focuses the cells, allowing them to flow individually past the laser. The incorporation of a separate syringe for the H2O2 enables more controlled and optimizable exposure time allowing higher H2O2 concentrations without adverse effects. Also, limiting the incubation time limits the breakdown of H2O2 by endogenous catalase. By incorporating this new flow system, the detected number of proteins with an FPOP modification increased 13-fold, thus expanding the capabilities of this method to probe a multitude of proteins in living cells. In this protocol a general IC-FPOP experiment is described focusing on the assembly of the IC-FPOP flow system.

<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>5 mL Gas Tight Syringe, Removable Luer Lock</td><td>SGE Analytical Science</td><td>008760</td><td></td></tr><tr><td>50 mL Conical Centrifuge Tubes</td><td>Fisher Scientific</td><td>14-432-22</td><td>any brand is sufficient</td></tr><tr><td>500 &amp;micro;L SGE Gastight Syringes: Fixed Luer-Lok Models</td><td>Fisher Scientific</td><td>SG-00723</td><td></td></tr><tr><td>Acetone, HPLC Grade</td><td>Fisher Scientific</td><td>A929-4</td><td>4 L quantity is not necessary</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>ACQUITY UPLC M-Class Symmetry C18 Trap Column, 100 &amp;Aring;, 5 &amp;micro;m, 180 &amp;micro;m x 20 mm, 2G, V/M, 1/pkg</td><td>Waters</td><td>186007496</td><td></td></tr><tr><td>ACQUITY UPLC M-Class System</td><td>Waters</td><td>ACQUITY UPLC M-Class System</td><td></td></tr><tr><td>Aluminum Foil</td><td>Fisher Scientific</td><td>01-213-100</td><td>any brand is sufficient</td></tr><tr><td>Aqua 5 &amp;micro;m C18 125 &amp;Aring; packing material</td><td>Phenomenex</td><td>04A-4299</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>Dithiothreiotol (DTT)</td><td>AmericanBio</td><td>AB00490-00005</td><td></td></tr><tr><td>DMSO, Anhydrous</td><td>Invitrogen</td><td>D12345</td><td></td></tr><tr><td>EX350 excimer laser</td><td>GAM Laser</td><td>EX350 excimer laser</td><td></td></tr><tr><td>FEP Tubing 1/16&quot; OD x 0.020&quot; ID</td><td>IDEX Health &amp;amp; Science</td><td>1548L</td><td></td></tr><tr><td>Formic Acid, LC/MS Grade</td><td>Fisher Scientific</td><td>A117-50</td><td></td></tr><tr><td>HV3-2 VALVE</td><td>Hamilton</td><td>86728</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>Iodoacetamide (IAA)</td><td>ACROS Organics</td><td>122270050</td><td></td></tr><tr><td>Legato 210 syringe pump</td><td>KD Scientific</td><td>788212</td><td>Any syringe pump that can hold 4, 5 mL syringes, withdraw and expel liquid, and have a way to stale the motor should work</td></tr><tr><td>Luer Adapter Female Luer to 1/4-28 Male Polypropylene</td><td>IDEX Health &amp;amp; Science</td><td>P-618L</td><td></td></tr><tr><td>Methanol, LC/MS Grade</td><td>Fisher Scientific</td><td>A454SK-4</td><td>4 L quantity is not necessary</td></tr><tr><td>Microcentrifuge</td><td>Thermo Scientific</td><td>75002436</td><td></td></tr><tr><td>N,N&#039;-Dimethylthiourea (DMTU)</td><td>ACROS Organics</td><td>116891000</td><td></td></tr><tr><td>NanoTight Sleeve Green 1/16&quot; ID x .0155&quot; ID x 1.6&quot;</td><td>IDEX Health &amp;amp; Science</td><td>F-242X</td><td></td></tr><tr><td>NanoTight Sleeve Yellow 1/16&quot; OD x 0.027&quot; ID x 1.6&quot;</td><td>IDEX Health &amp;amp; Science</td><td>F-246</td><td></td></tr><tr><td>N-tert-Butyl-&amp;alpha;-phenylnitrone (PBN)</td><td>ACROS Organics</td><td>177350250</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>PE50-C pyroelectric energy meter</td><td>Ophir Optronics</td><td>7Z02936</td><td></td></tr><tr><td>Pierce Quantitative Colorimetric Peptide Assay</td><td>Thermo Scientific</td><td>23275</td><td></td></tr><tr><td>Pierce Rapid Gold BCA Protein Assay Kit</td><td>Thermo Scientific</td><td>A53225</td><td></td></tr><tr><td>Pierce Trypsin Protease, MS Grade</td><td>Thermo Scientific</td><td>90058</td><td></td></tr><tr><td>Pierce Universal Nuclease for Cell Lysis</td><td>Fisher Scientific</td><td>88702</td><td></td></tr><tr><td>Polymicro Cleaving Stone, 1&quot; x 1&quot; x 1/32&quot;</td><td>Molex</td><td>1068680064</td><td>any capillary tubing cutter is sufficient</td></tr><tr><td>Polymicro Flexible Fused Silica Capillary Tubing, Inner Diameter 150 &amp;micro;m, Outer Diameter 360 &amp;micro;m, TSP150350</td><td>Polymicro Technologies</td><td>1068150024</td><td></td></tr><tr><td>Polymicro Flexible Fused Silica Capillary Tubing, Inner Diameter 450 &amp;micro;m, Outer Diameter 670 &amp;micro;m, TSP450670</td><td>Polymicro Technologies</td><td>1068150025</td><td></td></tr><tr><td>Polymicro Flexible Fused Silica Capillary Tubing, Inner Diameter 75 &amp;micro;m, Outer Diameter 360 &amp;micro;m, TSP075375</td><td>Polymicro Technologies</td><td>1068150019</td><td></td></tr><tr><td>Potassium Phosphate Monobasic</td><td>Fisher Scientific</td><td>P382-500</td><td></td></tr><tr><td>Proteome Discover 2.2 (bottom-up proteomics software)</td><td>Thermo Scientific</td><td>OPTON-30799</td><td></td></tr><tr><td>Rotary Magnetic Tumble Stirrer</td><td>V&amp;amp;P Scientific, Inc.</td><td>VP 710D3</td><td></td></tr><tr><td>Rotary Magnetic Tumble Stirrer, accessory kit for use with Syringe Pumps</td><td>V&amp;amp;P Scientific, Inc.</td><td>VP 710D3-4</td><td></td></tr><tr><td>Super Flangeless Ferrule w/SST Ring, Tefzel&amp;trade; (ETFE), 1/4-28 Flat-Bottom, for 1/16&quot; OD</td><td>IDEX Health &amp;amp; Science</td><td>P-259X</td><td></td></tr><tr><td>Super Flangeless Nut PEEK 1/4-28 Flat-Bottom, for 1/16&quot; &amp;amp; 1/32&quot; OD</td><td>IDEX Health &amp;amp; Science</td><td>P-255X</td><td></td></tr><tr><td>Super Tumble Stir Discs, 3.35 mm diameter, 0.61 mm thick</td><td>V&amp;amp;P Scientific, Inc.</td><td>VP 722F</td><td></td></tr><tr><td>Thermo Scientific Pierce RIPA Buffer</td><td>Fisher Scientific</td><td>PI89900</td><td></td></tr><tr><td>Tris Base</td><td>Fisher Scientific</td><td>BP152-500</td><td></td></tr><tr><td>Upchurch Scientific Low-Pressure Crosses: PEEK</td><td>Fisher Scientific</td><td>05-700-182</td><td></td></tr><tr><td>Upchurch Scientific Low-Pressure Tees: PEEK</td><td>Fisher Scientific</td><td>05-700-178</td><td></td></tr><tr><td>UV Fused Silica Plano-Convex Spherical Lenses</td><td>THORLABS</td><td>LA4052</td><td></td></tr><tr><td>V&amp;amp;P Scientific IncSupplier Diversity Partner TUMBL STIR DISC PARYLENE 1000</td><td>V&amp;amp;P Scientific, Inc.</td><td>VP724F</td><td></td></tr><tr><td>VHP MicroTight Union for 360&amp;micro;m OD</td><td>IDEX Health &amp;amp; Science</td><td>UH-436</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>

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.

IC-FPOP is a footprinting method to interrogate protein interactions in live cells. In the IC-FPOP flow system, the H2O2 exposure time is limited to roughly 3 s, warranting higher H2O2 concentrations without detrimental consequences for the cells. The flow system also incorporates two streams of sheath buffer, which hydrodynamically focuses the cells to the center of the tubing producing a single flow of cells to be uniformly irradiated (Figure 5)9. Fluorescence imaging of orthogonal YZ stacked images (Figure 5A) show a clear separation of the sheath buffer (containing a FITC fluorophore) from the cell solution (containing a TMRM fluorophore). To emphasize this separation, Figure 5B and Figure 5C show three-dimensional average heat maps of either the sheath buffer solution or cell solution, illustrating minimal mixing of the two solutions.
The use of the single cell flow system increases the number of oxidatively modified proteins by 13-fold (Figure 6A)9. In this method, proteins in many of the cellular compartments are labeled with membrane proteins, cytoplasmic proteins, and proteins within the nucleus being the most prevalent8,9. To ensure proteins were modified within intact cells, fluorescent images of CellROX treated cells were performed following H2O2 treatment and irradiation (Figure 6B)8. The stability of the cells throughout the labeling process further confirms the efficacy of IC-FPOP to probe proteins in their native cellular environment. By using tandem-mass spectrometry, these modifications can be localized to specific amino acids on a protein. Figure 7 represents a modification that takes place during IC-FPOP along with its extracted ion chromatogram. The shift observed in the extracted ion chromatogram translates to the change in hydrophobicity caused by the oxidized methionine in the modified peptide.
To test if the FPOP modifications probe solvent accessibility inside the cells, in-cell labeled actin was compared to both an in vitro footprinting study and various crystal structures of actin (Figure 8)8. The in-cell labeled actin is represented in Figure 8A shows comparable extents of oxidation from the in vitro study by Guan et al.10 (Figure 8B), concluding actin has similar solvent accessibility for both in-cell and in vitro studies. To further confirm IC-FPOP was probing the solvent accessibility of actin, the extent of FPOP modifications were compared to the solvent accessibility of the labeled residues calculated from two actin crystal structures (Figure 8C). This correlation demonstrates that IC-FPOP probes the solvent accessibility of the monomeric protein well.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/60911/60911fig1.jpg" /> 
Figure 1: How to properly construct ferules. (A) Place ferrules, silica tubing, and sleeve together before tightening. (B) Tighten all components together. (C) Final product will produce a ferule that has been tightened down on the sleeve. <a href="https://www.jove.com/files/ftp_upload/60911/60911fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/60911/60911fig2.jpg" /> 
Figure 2: Setting up IC-FPOP flow system. (A) Image of a fully assembled IC-FPOP flow system positioned next to the laser. (B) Example of a spacer needed to increase the outer diameter of the 500 &#181;L syringes to successfully tighten all syringes down together. (C) Representative pictures showing the space required for the magnetic stirrers. (D) Stoppers are necessary to stall the syringe pump without breaking the syringes. <a href="https://www.jove.com/files/ftp_upload/60911/60911fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/60911/60911fig3.jpg" /> 
Figure 3: Schematic of the flow system developed for IC-FPOP. Blue lines represent silica tubing with a 450 &#181;m ID and 670 &#181;m OD, red lines have a 150 &#181;m ID and 360 &#181;m OD, and orange lines have a 75 &#181;m ID and 360 &#181;m OD. <a href="https://www.jove.com/files/ftp_upload/60911/60911fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/60911/60911fig4.jpg" /> 
Figure 4: Protein analysis software used to detect FPOP modifications. A typical workflow with the corresponding modifications searched in each node. <a href="https://www.jove.com/files/ftp_upload/60911/60911fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/60911/60911fig5.jpg" /> 
Figure 5: The single cell flow system hydrodynamically focuses the cells into a single stream. (A) Orthogonal YZ stack illustrating 3D focusing of the cellular analyte (red, TMRM fluorophore) surrounded by the sheath buffer (green, FITC fluorophore). Three-dimensional average intensity heat map of the sheath buffer (B) and cellular analyte (C). Lower intensities are blue and highest are red (B-C). This figure has been modified from Rinas et al.9 <a href="https://www.jove.com/files/ftp_upload/60911/60911fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/60911/60911fig6.jpg" /> 
Figure 6: Utilization of the IC-FPOP Flow System drastically increases FPOP modified proteins in intake cells. (A) Comparison of oxidized proteins identified with and without the flow system. The flow system identified 1391 FPOP modified proteins while only 105 proteins were identified with no flow system with an overlap of 58 modified proteins. This figure has been modified from Rinas et al.9 (B) Fluorescence imaging of CellROX treated cells after IC-FPOP show the cells are still intact after oxidative labeling. Cells were imaged using an Olympus Fluoview FV1000 MPE multiphoton microscope at 665 nm. Image shown is a single slice. This figure has been modified from Espino et al.8 <a href="https://www.jove.com/files/ftp_upload/60911/60911fig6large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 7" class="xfigimg" src="/files/ftp_upload/60911/60911fig7.jpg" /> 
Figure 7: Examples of tandem MS/MS spectra that take place during IC-FPOP. Product-ion (MS/MS) spectra of an (A) unmodified peptide, and an (B) oxidation detected on residue M8 found on that peptide. A representative EIC of the (C) unmodified peptide and (D) modified peptide. <a href="https://www.jove.com/files/ftp_upload/60911/60911fig7large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 8" class="xfigimg" src="/files/ftp_upload/60911/60911fig8.jpg" /> 
Figure 8: IC-FPOP is effective in probing the solvent accessibility of proteins. (A) Extent of modification for the 9 oxidatively modified peptides from actin. Values are shown as averages plus and minus standard deviation (n = 3). (B) Modification of actin peptides oxidized in vitro by synchrotron radiolysis from Guan et al.10 (C) Correlation of residue FPOP modifications with SASA in the tight (triangles, dashed trend line) and open (circles, solid trend line) states of actin. This figure has been modified from Espino et al.8 <a href="https://www.jove.com/files/ftp_upload/60911/60911fig8large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Characterizing Cellular Proteins with In-cell Fast Photochemical Oxidation of Proteins at JoVE.com

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

characterizing-cellular-proteins-with-cell-fast-photochemical

Research

JoVE Journal

Biochemistry

5.5K Views.  University of Maryland Baltimore. In-cell FPOP in combinatie met massaspectrometrie is een krachtige nieuwe techniek die wordt gebruikt om eiwiteiwitinteracties en eiwitstructurele veranderingen in levende cellen te bestuderen. Met in-cell FPOP hoeft u geen eiwitten van belang te isoleren, maar u ze eerder bestuderen in hun eigen omgevingen. Dit is vooral handig voor membraaneiwitten.Met deze methode u veranderingen in structuur- en eiwiteiwerkingen bestuderen in een verscheidenheid aan ziekten, waaronder kanker en gen...

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

Identification of Small Molecule-binding Proteins in a Native Cellular Environment by Live-cell Photoaffinity Labeling

Identifying the molecular target(s) of small molecules is a challenging but necessary step towards understanding their mechanism of action. While several target identification methods have been developed and used to successfully elucidate the binding proteins of a variety of small molecules, these techniques have drawbacks that make them unsuitable for detecting certain types of small molecule-target interactions. In particular, non-covalent interactions that depend on native cellular conditions, such as those of membrane proteins whose structures may be perturbed upon cell lysis, are often not amenable to affinity-based target identification methods. Here, we demonstrate a method wherein a probe containing a photolabile group is used to covalently crosslink to the small molecule binding protein within the environment of the live cell, allowing the detection and isolation of the target protein without the need for maintenance of the interaction after cell lysis. This technique is a valuable tool for studying biologically interesting small molecules with unknown mechanisms, both in the context of basic biology as well as drug discovery.

We describe here a method for identification of small molecule-binding proteins using photoaffinity labeling. The advantage of this technique is that binding and covalent labeling of the target proteins occurs within the live cellular environment, removing the risk of disrupting native protein structure and binding conditions upon cell lysis.

Identifying the molecular target(s) of small molecules is a challenging but necessary step towards understanding their mechanism of action. While several ...

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

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

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

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

Single-Cell Quantification of Protein Degradation Rates by Time-Lapse Fluorescence Microscopy in Adherent Cell Culture

Proteins are in a dynamic state of synthesis and degradation and their half-lives can be adjusted under various circumstances. However, most commonly used approaches to determine protein half-life are either limited to population averages from lysed cells or require the use of protein synthesis inhibitors. This protocol describes a method to measure protein half-lives in single living adherent cells, using SNAP-tag fusion proteins in combination with fluorescence time-lapse microscopy. Any protein of interest fused to a SNAP-tag can be covalently bound by a fluorescent, cell permeable dye that is coupled to a benzylguanine derivative, and the decay of the labeled protein population can be monitored after washout of the residual dye. Subsequent cell tracking and quantification of the integrated fluorescence intensity over time results in an exponential decay curve for each tracked cell, allowing for determining protein degradation rates in single cells by curve fitting. This method provides an estimate for the heterogeneity of half-lives in a population of cultured cells, which cannot easily be assessed by other methods. The approach presented here is applicable to any type of cultured adherent cells expressing a protein of interest fused to a SNAP-tag. Here we use mouse embryonic stem (ES) cells grown on E-cadherin-coated cell culture plates to illustrate how single cell degradation rates of proteins with a broad range of half-lives can be determined.

This protocol describes a method to determine protein half-lives in single living adherent cells, using pulse labeling and fluorescence time-lapse imaging of SNAP-tag fusion proteins.

Proteins are in a dynamic state of synthesis and degradation and their half-lives can be adjusted under various circumstances. However, most commonly used ...

Developmental Biology

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, ...

Assessment of Cellular Oxidation using a Subcellular Compartment-Specific Redox-Sensitive Green Fluorescent Protein

Measuring the intracellular oxidation/reduction balance provides an overview of the physiological and/or pathophysiological redox status of an organism. Thiols are especially important for illuminating the redox status of cells via their reduced dithiol and oxidized disulfide ratios. Engineered cysteine-containing fluorescent proteins open a new era for redox-sensitive biosensors. One of them, redox-sensitive green fluorescent protein (roGFP), can easily be introduced into cells with adenoviral transduction, allowing the redox status of subcellular compartments to be evaluated without disrupting cellular processes. Reduced cysteines and oxidized cystines of roGFP have excitation maxima at 488 nm and 405 nm, respectively, with emission at 525 nm. Assessing the ratios of these reduced and oxidized forms allows the convenient calculation of redox balance within the cell. In this method article, immortalized human triple-negative breast cancer cells (MDA-MB-231) were used to assess redox status within the living cell. The protocol steps include MDA-MB-231 cell line transduction with adenovirus to express cytosolic roGFP, treatment with H2O2, and assessment of cysteine and cystine ratio with both flow cytometry and fluorescence microscopy.

This protocol describes the assessment of subcellular compartment-specific redox status within the cell. A redox-sensitive fluorescent probe allows convenient ratiometric analysis in intact cells.

Measuring the intracellular oxidation/reduction balance provides an overview of the physiological and/or pathophysiological redox status of an organism. ...

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 ...

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

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

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

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

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

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

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

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

Photoconversion of Purified Fluorescent Proteins and Dual-probe Optical Highlighting in Live Cells

Photoconvertible fluorescent proteins (pc-FPs) are a class of fluorescent proteins with "optical highlighter" capability, meaning that the color of fluorescence can be changed by exposure to light of a specific wavelength. Optical highlighting allows noninvasive marking of a subpopulation of fluorescent molecules, and is therefore ideal for tracking single cells or organelles.
Critical parameters for efficient photoconversion are the intensity and the exposure time of the photoconversion light. If the intensity is too low, photoconversion will be slow or not occur at all. On the other hand, too much intensity or too long exposure can photobleach the protein and thereby reduce the efficiency of photoconversion.
This protocol describes a general approach how to set up a confocal laser scanning microscope for pc-FP photoconversion applications. First, we describe a procedure for preparing purified protein droplet samples. This sample format is very convenient for studying the photophysical behavior of fluorescent proteins under the microscope. Second, we will use the protein droplet sample to show how to configure the microscope for photoconversion. And finally, we will show how to perform optical highlighting in live cells, including dual-probe optical highlighting with mOrange2 and Dronpa.

This protocol describes a general approach to perform photoconversion of fluorescent proteins on a confocal laser scanning microscope. We describe procedures for the photoconversion of puried protein samples, as well as for dual-probe optical highlighting in live cells with mOrange2 and Dronpa.

Photoconvertible fluorescent proteins (pc-FPs) are a class of fluorescent proteins with "optical highlighter" capability, meaning that the color of ...

Biology

Micromanipulation Techniques Allowing Analysis of Morphogenetic Dynamics and Turnover of Cytoskeletal Regulators

Examining the spatiotemporal dynamics of proteins can reveal their functional importance in various contexts. In this article, it is discussed how fluorescent recovery after photobleaching (FRAP) and photoactivation techniques can be used to study the spatiotemporal dynamics of proteins in subcellular locations. We also show how these techniques enable straightforward determination of various parameters linked to actin cytoskeletal regulation and cell motility. Moreover, the microinjection of cells is additionally described as an alternative treatment (potentially preceding or complementing the aforementioned photomanipulation techniques) to trigger instantaneous effects of translocated proteins on cell morphology and function. Micromanipulation such as protein injection or local application of plasma membrane-permeable drugs or cytoskeletal inhibitors can serve as powerful tool to record immediate consequences of a given treatment on cell behavior at the single cell and subcellular level. This is exemplified here by immediate induction of lamellipodial cell edge protrusion by the injection of recombinant Rac1 protein, as established a quarter-century ago. In addition, we provide a protocol for determining the turnover of enhanced green fluorescent protein (EGFP)-VASP, an actin filament polymerase prominently accumulating at lamellipodial tips of B16-F1 cells, employing FRAP and including associated data analysis and curve fitting. We also present guidelines for estimating the rates of lamellipodial actin network polymerization, as exemplified by cells expressing EGFP-tagged &#946;-actin. Finally, instructions are given for how to investigate the rates of actin monomer mobility within the cell cytoplasm, followed by actin incorporation at sites of rapid filament assembly, such as the tips of protruding lamellipodia, using photoactivation approaches. None of these protocols is&#160;restricted to components or regulators of the actin cytoskeleton, but can easily be extended to explore in analogous fashion the spatiotemporal dynamics and function of proteins in various different subcellular structures or functional contexts.

We describe how micro- and photomanipulation techniques such as FRAP and photoactivation enable the determination of motility parameters and the spatiotemporal dynamics of proteins within migrating cells. Experimental readouts include subcellular dynamics and turnover of motility regulators or of the underlying actin cytoskeleton.

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

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