Name: in vivo hydroxyl radical protein footprinting for the study of protein interactions in caenorhabditis elegans
Uploaded: 2020-04-01T14:00:00.000+00:00
Duration: 8 min
Description: Watch this Scientific Journal Video about In Vivo Hydroxyl Radical Protein Footprinting for the Study of Protein Interactions in Caenorhabditis elegans at JoVE.com

This work was supported by start-up funds from the University of Maryland, Baltimore and the NIH 1R01 GM 127595 awarded to LMJ. The authors thank Dr. Daniel Deredge for his help in editing the manuscript.

1. C. elegans maintenance and culture
<ol>
	<li>Grow and synchronize worms colonies to their fourth larvae (L4) stage following standard laboratory procedures8.</li>
	<li>The day of IV-FPOP experiment, wash L4 worms from bacterial lawns (OP50 E. coli) with M9 buffer (0.02 M KH2PO4, 0.08 M Na2HPO4, 0.08 M NaCl, 1 mM MgSO4).</li>
	<li>Obtain 500 &#181;L aliquots of ~10,000 worms per IV-FPOP sample.</li>
</ol>
2. Microfluidic flow system assembly
<ol>
	<li>Start the flow system assembly by cutting a 2 cm piece of fluorinated ethylene propylene (FEP) tubing (1/16 in. outer diameter (o.d.) x 0.020 in. inner diameter (i.d.)).</li>
	<li>Using a clean dissecting needle, widen the i.d. of the FEP tubing in order to make a small crater at one end only, ~50 mm in length. The crater should be big enough to fit two 360 &#181;m o.d. pieces of fused silica. 
	NOTE: Do not widen the opposite end as this end will only be used to fit the outlet capillary.</li>
	<li>With a ceramic cutter, cut two 15 cm pieces of 250 &#181;m i.d. fused silica. These two pieces will become the infusing lines of the flow system.</li>
	<li>Using self-adhesive tape, tape the two 250 &#181;m i.d. capillaries together ensuring they are parallel to each other and their ends are 100% flushed. 
	NOTE: To ensure the fused silica ends are not crushed after cutting, and are straight and 100% flushed against each other, it is recommended to examine them using a magnifying glass or under a stereo microscope.</li>
	<li>Insert the two taped capillaries into the handmade crater of the FEP tubing. Push the capillaries up to the very edge of the handmade crater. 
	CAUTION: To maintain the efficacy of the flow system, do not push past the handmade crater (Figure 1a).</li>
	<li>Place a small dot of epoxy resin on a clean surface and mix with a dissecting needle.</li>
	<li>Quickly, using the same needle, place a small drop of resin at the end of the infusing capillaries where they connect with the FEP tubing and allow the resin to dry, hanging outlet-side up, for a few minutes (Figure 1a).</li>
	<li>While the resin dries, binding the FEP tubing and two capillaries together, cut a new 250 &#181;m i.d. capillary. The new capillary will become the outlet capillary of the flow system. The desired length of the capillary can be calculated using the equation below: 
	<img alt="Equation 1" src="/files/ftp_upload/60910/60910eq1.jpg" /> 
	Where &#8467; is length of the capillary to be cut in centimeters, t is the desired reaction time in minutes, f is the flow rate in mL/min, and i.d. is the inner diameter of the capillary in centimeters. 
	NOTE: After cutting the outlet capillary, examine the ends ensuring they are straight and not crushed.</li>
	<li>Once the resin has dried, insert the new capillary through the FEP tubing outlet end. The inside ends of the outlet capillary and the two infusing capillaries should be flush against each other inside the FEP tubing, this creates the mixing-T (Figure 1b).</li>
	<li>Bind the outlet capillary and FEP tubing with fresh epoxy resin as described in steps 2.6 and 2.7.</li>
	<li>Allow the resin to dry overnight binding the flow system together. Set up the microfluidic flow system the next day (Figure 1c).</li>
</ol>
3. Microfluidic flow system for in vivo FPOP
<ol>
	<li>Insert four magnetic stirrers inside one 5 mL syringe. This syringe becomes the sample syringe. The magnetic stirrers prevent the worms from settling inside the syringe during IV-FPOP (Figure 2A).</li>
	<li>Fill the two 5 mL syringes with M9. Make sure there are no air bubbles inside each syringe as they can affect the flow rate and mixing efficiency.</li>
	<li>Connect a Luer adapter to each 5 mL syringe ensuring they are finger tight and secured in place.</li>
	<li>Attach each syringe to a single 3-2 valve. The syringe should be attached to the middle port of the valve (Figure 2B).</li>
	<li>Secure each 5 mL syringe to the dual syringe pump and adjust the mechanical stop collar to prevent over pressure to the syringe from the pusher block. Keep in mind the addition of the magnetic stirrers when setting the stop collar.</li>
	<li>Attach each infusing capillary end of the homemade microfluidic flow system to the top port of each 3-2 valve using a super flangeless nut, FEP sleeve, and super flangeless ferrule (Figure 2B).</li>
	<li>To the remaining opened port of the 3-2 valve (bottom port), attach a 10 cm 450 &#181;m i.d. capillary using a super flangeless nut, FEP sleeve, and super flangeless ferrule (Figure 2B). These capillaries become the withdrawing sample capillaries.</li>
	<li>Start the pump flow and check all connections of the microfluidic flow system for visual leaks. Flow at least three syringe volumes using the experimental flow rate. The final flow rate is dependent on the laser irradiation window, laser frequency, and a zero-exclusion fraction volume. 
	NOTE: For this protocol, a final flow rate of 375.52 &#181;L/min was calculated from a laser irradiation window of 2.55 mm, 250 &#181;m i.d. fused silica, zero exclusion fraction, and 50 Hz frequency.
	<ol>
		<li>The flow path is marked by the arrows on the 3-2 valve handle. Each syringe can be refilled manually by moving the valve handle from the expelling position to the withdrawing position. 
		NOTE: Leaks at the 3-2 valve can be fixed by re-screwing the super flangeless nut or replacing any of the valve connections (super flangeless nut, FEP sleeve, or super flangeless ferrule). Leaks at the mixing-T require a reassembly of the microfluidic flow system (steps 2.1 to 2.11).</li>
	</ol>
	</li>
	<li>Move the microfluidic flow system to the experimental bench and secure the outlet capillary to the radiating stage using a 360 &#181;m o.d. stainless steel union (Figure 2C,D).</li>
	<li>Using a long-reach lighter, burn the coating of the fused silica at the laser irradiation window. Clean the burned coating using lint-free tissue and methanol. 
	NOTE: Alternate between burning cycles and methanol cleaning cycles to avoid over-heating the capillary as excess heat can melt and/or break the capillary.</li>
	<li>Position the magnetic stirrer block above the 5 mL syringe containing the magnetic stirrers. Adjust the speed and position of the magnetic stirrer block so that the magnetic stirrers inside the 5 mL syringe are rotating slowly and constantly.</li>
</ol>
4. In vivo FPOP
<ol>
	<li>Turn on the KrF excimer laser and allow the thyratron to warm-up. 
	CAUTION: The laser emits visible and invisible radiation at 248 nm wavelength that can damage eyes. Proper eye protection should be worn before turning on the laser and opening the beam exit window.</li>
	<li>Measure the laser energy at a frequency of 50 Hz for at least 100 pulses by placing the optical sensor at the beam exit window. 
	NOTE: For this protocol, a 2.55 mm irradiation window was used with a laser energy of 150 &#177; 2.32 mJ.</li>
	<li>Obtain approximatively 10,000 worms (500 &#181;L) and manually withdraw the sample into the sample syringe.</li>
	<li>Fill the sample syringe with 2.5 mL of M9 buffer for a final sample volume of 3 mL. Make sure that no air bubbles are introduced into the sample syringe.</li>
	<li>Fill the second syringe with 3 mL of 200 mM H2O2, again ensuring no air bubbles are introduced into the syringe.</li>
	<li>At the end of the outlet capillary, place a 15 mL conical tube wrapped in aluminum foil containing 6 mL of 40 mM N&#8217;-Dimethylthiourea (DMTU), 40 mM N-tert-Butyl-&#945;-phenylnitrone (PBN), and 1% dimethyl sulfoxide to quench excess H2O2, hydroxyl radicals, and inhibit methionine sulfoxide reductase activity, respectively.</li>
	<li>Start the excimer laser from the software window, wait for the first pulse, and start the sample flow from the dual syringe. 
	NOTE: Samples should be run in biological duplicates in technical triplicates under 3 samples conditions: laser irradiation with H2O2 (sample), no laser irradiation with H2O2 and no laser irradiation no H2O2 (controls), totaling to 9 samples per biological set.</li>
	<li>Collect the entire sample in the 15 mL tube, while actively monitoring the sample flow for any visual leaks as some back pressure from the sample syringe can sometimes build-up.</li>
	<li>Following IV-FPOP, pellet worms by centrifugation at 805 x g for 2 min. Remove quench solution, add 250 &#181;L of lysis buffer (8 M urea, 0.5% SDS, 50 mM HEPES, 50 mM NaCl, 1 mM EDTA, 1 mM phenylmethylsulfonyl fluoride [PMSF]). Transfer the sample to a clean microcentrifuge tube, flash freeze, and store at -80 &#176;C until sample digestion.</li>
</ol>
5. Protein extraction, purification, and proteolysis
<ol>
	<li>Thaw frozen samples on ice and homogenize by sonication for 10 s, followed by a 60 s ice incubation. Multiple rounds of sonication may be required, homogenate can be observed under a stereomicroscope by placing a 2 &#181;L sample aliquot on a microscope slide. Sample homogenization is complete when small to no pieces of worms are seen.</li>
	<li>Centrifuge homogenized worms at 400 x g for 5 min at 4 &#176;C and collect the supernatant into a clean microcentrifuge tube.</li>
	<li>Determine sample protein concentrations using a bicinchoninic acid assay (BCA assay) using the manufacturer&#8217;s instructions. 
	NOTE: Sample concentration can be determined using any biochemical protein concentration assay of choice. Keep in mind reagent compatibility with lysis buffer (8 M urea, 0.5% SDS, 50 mM HEPES, 50 mM NaCl, 1 mM EDTA, 1 mM PMSF). Additionally, a buffer dilution may be necessary.</li>
	<li>Obtain 100 &#181;g of protein from each sample and place in clean microcentrifuge tubes.</li>
	<li>Add dithiothreitol (DTT) to a 10 mM final concentration to reduce disulfide bonds in all samples. Vortex, spin down, and incubate samples at 50 &#176;C for 45 min.</li>
	<li>Cool samples to room temperature for 10 min.</li>
	<li>Add iodoacetamide (IAA) to a 50 mM final concentration to alkylate reduced residues. Vortex, spin down, and incubate samples for 20 min at room temperature protected from light.</li>
	<li>Immediately after alkylation, precipitate the protein sample by adding 4 volumes of 100% pre-chilled acetone and incubate at -20 &#176;C overnight.</li>
	<li>The next day, pellet protein precipitate by centrifugation at 16,000 x g for 10 min. Wash sample pellet with 90% acetone, remove supernatant, and allow pellet to dry for 2&#8211;3 min.</li>
	<li>Re-suspend the protein pellet in 25 mM Tris-HCl (1 &#181;g/&#181;L). Add trypsin at a final protease to protein ratio of 1:50 (w/w) and digest samples at 37 &#176;C overnight.</li>
	<li>The next day quench the trypsin digestion reaction by adding 5% formic acid. 
	NOTE: A minimum of 0.5 &#181;g of peptides per sample is needed for LC-MS/MS analysis (steps 6.1-6.5). Determine sample peptide concentrations using a quantitative colorimetric peptide assay (see the Table of Materials) as described by the manufacturer&#8217;s protocol.</li>
</ol>
6. High performance liquid chromatography-tandem mass spectrometry (LC-MS/MS)
<ol>
	<li>Use the following LC mobile phases: water in 0.1% FA (A) and ACN in 0.1% FA (B).</li>
	<li>Load 0.5 &#181;g of peptides onto a trap column (180 &#181;m &#215; 20 mm) containing C18 (5 &#181;m, 100 &#197;) and wash sample with 99% solvent A and 1% B for 15 min at 15 &#181;L/min flow rate.</li>
	<li>Separate peptides using an in-house packed column (0.075 mm i.d. &#215; 20 mm) with C18 reverse phase material (5 &#956;m, 125 &#197;).</li>
	<li>Use the following analytical separation method: the gradient was pumped at 300 nL/min for 120 min: 0&#8722;1 min, 3% B; 2&#8722;90 min, 10&#8722;45% B; 100&#8722;105 min, 100% B; 106&#8722;120 min, 3% B.</li>
	<li>Perform data acquisition of peptides in positive ion mode nano-electrospray ionization (nESI) using a high-resolution mass spectrometer. 
	NOTE: For this protocol, an ultra-performance LC (UPLC) instrument couple to a high-resolution mass spectrometer was utilized. Data dependent acquisition was utilized. The m/z scan range for MS1 was 3750&#8211;1500 at 60,000 resolution and 60 s dynamic exclusion. Ions with charge states of +1 and &#62;6 were excluded. An automatic gain control (AGC) target of 5.5e5 was used with a maximum injection time of 50 ms and an intensity threshold of 5.5e4. Ions selected for MS2 were subjected to higher-energy collisional dissociation (HCD) fragmentation using 32% normalized collision energy. Fragment ions were detected in the spectrometer with 15,000 resolution and a 5.0e4 AGC target.</li>
</ol>
7. Data analysis
<ol>
	<li>Search tandem MS files with a bottom-up proteomics analysis software against the C. elegans database.</li>
	<li>Set the protein analysis search parameters as follow: one trypsin missed cleavage, 375&#8211;1500 m/z peptide mass range, fragment mass tolerance of 0.02, and precursor mass tolerance of 10 ppm. Carbamidomethylation is set as a static modification and all know hydroxyl radical side-chain modifications9,10 as dynamic. Peptide identification is established at 95% confidence (medium) and residue at 99% confidence (high). The false discovery rate (FDR) is set a 1%.</li>
	<li>Export data to an electronic database and summarize the extent of oxidation per peptide or residue using the equation below11: 
	<img alt="Equation 2" src="/files/ftp_upload/60910/60910eq2.jpg" /> 
	NOTE: Where EIC area modified is the extracted ion chromatographic area (EIC) of the peptide or residue with an oxidative modification, and EIC area is the total area of the same peptide or residue with and without the oxidative modification.</li>
</ol>

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This publication was written in partial fulfillment of JAE Ph.D. thesis. The authors declare no conflict of interest.

The current benchmark for the study of in vivo protein-protein interactions (PPI) is fluorescence resonance energy transfer (FRET). In its most simple form, this technique studies PPI by energy transfer between two molecules when they are in close proximity to one another15. Unlike MS techniques, FRET does not have the resolution to characterize conformational changes and interaction sites at the amino-acid level. MS based techniques have been increasingly utilized for the study of PPI16. IV-FPOP is a HRPF method that allows for the in vivo protein structural analysis in C. elegans. In order to successfully label C. elegans by IV-FPOP, it is important to properly assemble the microfluidic flow system to reduce sample loss. The 250 &#181;m i.d. capillary has shown to maximize sample recovery when compared to smaller i.d. capillaries4. Larger i.d. capillaries have not been tested, however the microfluidic flow system is designed using a capillary with the same i.d. as a commercially available flow cytometry system for the sorting of C. elegans.17 The worm sample size is also important, a sample size of less than ~10,000 per sample prior to FPOP does not yield protein concentrations high enough for LC-MS/MS analysis. Higher samples sizes (&#62;10,000 worms) can also be used by adjusting the initial starting volume (step 4.5).
Proper assembly of the microfluidic flow system is important. Leaks in the sample pathway result in an inconsistent flow of the worms or H2O2. The ferrules, sleeves, and 3-2 valves can be reused from multiple IV-FPOP experiments if properly cleaned after each experiment. However, we recommend to assemble a new microfluidic flow system for every biological replicate. If the microfluidics is properly assembled, the worms and H2O2 will mix at the mixing-T with minimal back pressure. As quality control (QC) of the microfluidic flow system, we recommend testing the mixing efficiency by using colored dyes. It is important to monitor the motion of the magnetic stirrers inside the worm syringe during IV-FPOP, improper sample mixing at the worm syringe or the mixing-T can result in back pressure causing leaks. In addition, poor mixing conditions lead to large sample losses, poor laser exposure of worms at the laser window, and clogging.
C. elegans maintenance is important in order to decrease background oxidation. We recommend growing the worms at low temperatures under low stress conditions as high temperatures can affect total background oxidation. A control sample set of worms only, no H2O2 and no laser irradiation, is recommended for all IV-FPOP experiments to account for background oxidation due to laboratory maintenance. One of the current limitations of this technique is total number of oxidatively modified peptides identified and the total number of residues oxidized per peptide in order to gain higher protein structural information. Although not recommended, an increase in oxidative modifications could be achieve by using higher concentrations of hydrogen peroxide. An increase in hydrogen peroxide could significantly alter important biological pathways as well as lead to oxidation-induced unfolding. If the hydrogen peroxide concentration for IV-FPOP is increased, it is recommended to test worm viability and background oxidation as concentrations higher than 200 mM have not been tested.
The LC-MS/MS protocol described can be optimized and modified to meet the MS QC of other laboratories. The use of 2D-chromatography techniques has been previously shown to increase the identification of oxidatively modified peptides and proteins18. Nonetheless, protein enrichment techniques that target a specific protein of interest are not recommend, including but not limited to antibody precipitation or pull-down assays. These techniques can bias towards one protein conformer if the epitope/binding site of the protein has been oxidatively modified by IV-FPOP. New developments in footprinting radical reagents, such as sulfate radical anion10 or trifluoromethylation19 could increase the versatility of IV-FPOP. Although the only labeling reagent tested in vivo thus far is hydrogen peroxide, other laser-activated radicals could be optimized. The use of other radicals should prove to be compatible with worm viability, cell permeable, and the 248 nm laser wavelength. Owing to the use of C. elegans as a model system for many human diseases, IV-FPOP has the potential to have a strong impact in studying the role of protein structure in disease pathogenesis.

Protein footprinting coupled to mass spectrometry (MS) has been used in recent years to study protein interactions and conformational changes. Hydroxyl radical protein footprinting (HRPF) methods probe protein solvent accessibility by modifying protein amino acid side chains. The HRPF method, fast photochemical oxidation of proteins (FPOP)1, has been used to probe protein structure in vitro2, in-cell (IC-FPOP)3, and most recently in vivo (IV-FPOP)4. FPOP utilizes a 248 nm wavelength excimer laser in order to rapidly generate hydroxyl radicals by photolysis of hydrogen peroxide to form hydroxyl radicals1. In turn, these radicals can label 19 out of 20 amino acids on a microsecond time scale, faster than proteins can unfold. Although, the reactivity of each amino acid with hydroxyl radicals extends 1000-fold, it is possible to normalize side chain oxidation by calculating a protection factor (PF)5.
Since FPOP can oxidatively modify proteins regardless of their size or primary sequence, it proves to be advantageous for in-cell and in vivo protein studies. IV-FPOP probes protein structure in C. elegans similarly to in vitro and in-cell studies4. C. elegans are part of the nematode family and are widely used as a model to study human diseases. The ability of the worm to uptake hydrogen peroxide by both passive and active diffusion allows for the study of protein structure in different body systems. In addition, C. elegans are suited for IV-FPOP due to their transparency at the 248 nm laser wavelength needed for FPOP6. Coupling of this method to mass spectrometry allows for the identification of multiple modified proteins using traditional bottom-up proteomics approaches.
In this protocol, we describe how to perform IV-FPOP for the analysis of protein structure in C. elegans. The experimental protocol requires the assembly and set-up of microfluidic flow system for IV-FPOP adapted from Konermann et al7. After IV-FPOP, samples are homogenized for protein extraction. Protein samples are proteolyzed and peptides are analyzed by liquid chromatograph (LC) tandem MS, followed by quantification.

<table><tbody><tr><td>15mL 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>2 minimum</td></tr><tr><td>60 Sonic Dismembrator</td><td>Fisher Scientific</td><td>FM3279</td><td>This item is no longer available. Any low-volume sonicator will be sufficient</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></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></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>Dissecting Needle</td><td>Fisher Scientific</td><td>50-822-525</td><td>only a couple are needed</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>Epoxy instant mix 5 minute</td><td>Loctite</td><td>1365868</td><td></td></tr><tr><td>Ethylenediaminetetraacetic acid (EDTA)</td><td>Fisher Scientific</td><td>S311-100</td><td></td></tr><tr><td>EX350 excimer laser (248 nm wavelength)</td><td>GAM Laser</td><td></td><td></td></tr><tr><td>FEP Tubing 1/16&quot; OD x 0.020&quot; ID</td><td>IDEX Health &amp;amp; Sciene</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>HEPES</td><td>Fisher Scientific</td><td>BP310-500</td><td></td></tr><tr><td>HV3-2 VALVE</td><td>Hamilton</td><td>86728</td><td>2 minimum</td></tr><tr><td>Hydrochloric Acid</td><td>Fisher Scientific</td><td>A144S-500</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 101 syringe pump</td><td>KD Scientific</td><td>788101</td><td></td></tr><tr><td>Luer Adapter Female Luer to 1/4-28 Male Polypropylene</td><td>IDEX Health &amp;amp; Sciene</td><td>P-618L</td><td>2 minimum</td></tr><tr><td>Magnesium Sulfate</td><td>Fisher Scientific</td><td>M65-500</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&amp;prime;-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 x1.6&quot;&#039;</td><td>IDEX Health &amp;amp; Sciene</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; Sciene</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>OmniPur Phenylmethyl Sulfonyl Fluoride (PMSF)</td><td>Sigma-Aldrich</td><td>7110-OP</td><td>any protease inhibitor is sufficient</td></tr><tr><td>Orbitrap Fusion Lumos Tribrid Mass Spectrometer</td><td>Thermo Scientific</td><td></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>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>Polymicro Flexible Fused Silica Capillary Tubing, Inner Diameter 250&amp;micro;m, Outer Diameter 350&amp;micro;m, TSP250350</td><td>Polymicro Technologies</td><td>1068150026</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>1068150625</td><td></td></tr><tr><td>Polymicro Flexible Fused Silica Capillary Tubing, Inner Diameter 75&amp;micro;m, Outer Diameter 375&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 (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>Scissors</td><td>Fisher Scientific</td><td>50-111-1315</td><td>any scissors are sufficient</td></tr><tr><td>Self-Adhesive Label Tape</td><td>Fisher Scientific</td><td>15937</td><td>one roll is sufficient</td></tr><tr><td>Snap-Cap Microcentrifuge Flex-Tube Tubes</td><td>Fisher Scientific</td><td>05-402</td><td>any brand is sufficient</td></tr><tr><td>Sodium Chloride</td><td>Fisher Scientific</td><td>S271-500</td><td></td></tr><tr><td>Sodium Dodecyl Sulfate (SDS)</td><td>Fisher Scientific</td><td>15-525-017</td><td></td></tr><tr><td>Sodium Phosphate Dibasic Heptahydrate</td><td>Fisher Scientific</td><td>S373-500</td><td></td></tr><tr><td>Stereo Zoom Microscope</td><td>Fisher Scientific</td><td>03-000-014</td><td>a magnifying glass is sufficient</td></tr><tr><td>Super Flangeless Ferrule w/SST Ring, Tefzel (ETFE), 1/4-28 Flat-Bottom, for 1/16&quot; OD</td><td>IDEX Health &amp;amp; Sciene</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; Sciene</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>Tris Base</td><td>Fisher Scientific</td><td>BP152-500</td><td></td></tr><tr><td>Universal Base Plate, 2.5&quot; x 2.5&quot; x 3/8&quot;</td><td>Thorlabs Inc.</td><td>UBP2</td><td></td></tr><tr><td>Urea</td><td>Fisher Scientific</td><td>U5378</td><td></td></tr><tr><td>VHP MicroTight Union for 360&amp;micro;m OD</td><td>IDEX Health &amp;amp; Sciene</td><td>UH-436</td><td>2 minimum</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>

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 the microfluidic flow system used for IV-FPOP, H2O2 and the worms are kept separated until just prior to laser irradiation. This separation eliminates breakdown of H2O2 by endogenous catalase and other cellular mechanisms12. The use of a 250 &#181;m i.d. capillary shows a total sample recovery between 63&#8211;89% across two biological replicates, while the 150 &#181;m i.d. capillary only shows 21&#8211;31% recovery (Figure 3A). The use of a larger i.d. capillary (250 &#181;m) leads to better worm flow during IV-FPOP and single worm flow (Figure 3B) when compared to a smaller i.d. capillary (150 &#181;m) (Figure 3C). The 150 &#181;m i.d. capillary does not allow for single worm flow (Figure 3C) and multiple worms are seen flowing together at the laser irradiating window which decreases the amount of laser exposure per single worm.
IV-FPOP is a covalent labeling technique that probes solvent accessibility in C. elegans. Figure 4A shows a representative extracted ion chromatograms (EIC) of a FPOP modified and unmodified peptide. The hydroxyl radical label changes the chemistry of oxidatively modified peptides, thus making FPOP modified peptides more polar. In reverse phase chromatography, IV-FPOP modified peptides have earlier retention times than unmodified peptides. MS/MS fragmentation of isolated peptides allows for the identification of oxidatively modified residues (Figure 4B).
IV-FPOP has shown to oxidatively modified a total of 545 proteins across two biological replicates within C. elegans (Figure 5A,B). An advantage of IV-FPOP as a protein footprinting method relies on the technique&#8217;s ability to modify proteins in a variety of body systems within the worms (Figure 5C). This method would allow to probe protein structure and protein interactions regardless of body tissue or organ within the worm. Further, tandem MS analysis confirms IV-FPOP probes solvent accessibility in vivo. The oxidation pattern of the heat shock protein 90 (Hsp90) in complex with the myosin chaperon protein UNC-45 was analyzed (Figure 6). MS/MS analysis for Hsp90 shows four oxidatively modified residues (Figure 6C,D), the normalized extent of FPOP modification (ln(PF))5 indicates Hsp90&#8217;s residue M698 to be less solvent accessible than residues R697, E699, and E700 when bound to UNC-45 (Figure 6C). These differences in oxidation are validated by literature solvent accessible surface area (SASA) calculations (PDB 4I2Z13). Residue M698 has a SASA value of 0.03 which is consider to be a buried residue when compared to residues R697, E699, and E700 with higher SASA values (Figure 6C).14
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/60910/60910fig1.jpg" /> 
Figure 1. In vivo FPOP microfluidic flow system schematic. (A) The two infusing lines (orange) of the IV-FPOP flow system are shown inside the FEP tubing (yellow), the correct binding position of the epoxy resin is represented by the light blue circle. (B) Complete assembled mixing-T formed by the three 250 &#181;m i.d. capillaries. The correct resin binding position of the outlet capillary to the FEP tubing is represented by the light blue circle. (C) The complete assembled flow system for in vivo covalent labeling of C. elegans. Prior to FPOP, worms are kept separated from H2O2 until just prior to labeling; the laser irradiation window is shown in light blue and the laser beam is represented by the purple lightning bolt. Figures are not to scale. This figure has been modified from Espino et al.4. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/60910/60910fig1large.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/60910/60910fig2.jpg" /> 
Figure 2. Microfluidic system during IV-FPOP. (A) Representative picture of C. elegans inside the 5 mL syringe. Without stirring, the worms settle at the bottom of the syringe (left). The magnetic stirrers and stirrer block keep the worms in suspension during the IV-FPOP experiments (right). (B) Representative picture of a 5 mL syringe, infusing capillary, and withdrawing capillary connected to the 3-2 valve. The 3-2 valve handle is shown in the withdrawing position. (C) Microfluidic flow system during IV-FPOP, the magnetic stirrer block is position above the worms&#8217; 5 mL syringe. (D) Outlet capillary secured to the radiating stage. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/60910/60910fig2large.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/60910/60910fig3.jpg" /> 
Figure 3. Comparison of C. elegans flow and recovery using two i.d. capillaries. (A) Percent recovery of worms after IV-FPOP for two biological replicates (BR) with 250 (gray) and 150 (black) &#956;m i.d. capillaries. Error bars are calculated from the standard deviation across technical triplicates. C. elgans flowing through the laser irradiating window through a 250 &#181;m (B) and 150 &#181;m (C) i.d. capillaries. The worms are more tightly compacted in the smaller capillary. The 150 &#181;m i.d. capillary shows clumping of worms. This figure has been modified from Espino et al.4. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/60910/60910fig3large.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/60910/60910fig4.jpg" /> 
Figure 4. Representative LC-MS/MS results following IV-FPOP. (A) EIC of a FPOP modified peptide (red) and unmodified (blue). The selected peptide belongs to the actin-1 protein. (B) MS/MS spectrum of doubly charged unmodified actin-1 peptide 317-327. (C) MS/MS spectrum of doubly charged FPOP modified actin-1 peptide 317-327, in this example P323 was oxidatively modified (y5+ ion, red). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/60910/60910fig4large.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/60910/60910fig5.jpg" /> 
Figure 5. IV-FPOP oxidatively modifies proteins within C. elegans. (A) Venn diagram of oxidatively modified proteins in the presence of 200 mM hydrogen peroxide at 50 Hz across two biological replicates (BR), BR1 is in blue and BR2 is in yellow. (B) Venn diagram of oxidatively modified proteins identified in irradiated samples, hydrogen peroxide control, and worm-only control in BR2 across technical triplicates. (C) Pie chart of oxidatively modified proteins within different C. elegans body systems. This figure has been modified from Espino et al.4. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/60910/60910fig5large.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/60910/60910fig6.jpg" /> 
Figure 6. Correlating IV-FPOP modifications to solvent accessibility. (A) Myosin chaperon protein UNC-45 (gray) (PDB ID 4I2Z13) highlighting two modified peptides identified by LC/MS/MS analysis, 669&#8722;680 and 698&#8722;706 (green, left inset). UNC-45 is bound to the Hsp90 peptide fragment (blue). Oxidatively modified residues within this fragment are shown in sticks (red), and UNC-45 is rendered as a surface (right inset). (B) Tandem MS spectra of UNC-45 peptide 669&#8722;680 (top) and 698&#8722;706 (bottom) showing b- and y-ions for the loss of CO2, an FPOP modification. (C) The calculated ln(PF) for the Hsp90 oxidatively modified residues, R697, M698, E699, and E700. Calculated SASA values for Hsp90 are denoted above each residue. (D) Tandem MS spectra for R697, M698, E699, and E700 showing a +16 FPOP modification. This figure has been modified from Espino et al.4. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/60910/60910fig6large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about In Vivo Hydroxyl Radical Protein Footprinting for the Study of Protein Interactions in Caenorhabditis elegans at JoVE.com

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.

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, ...

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Nanyang Technological University

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JoVE Journal

Biochemistry

6.8K Views.  University of Maryland. 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.

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

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 ...

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 ...

Monitoring Protein Aggregation Kinetics In Vivo using Automated Inclusion Counting in Caenorhabditis elegans

Protein aggregation into insoluble inclusions is a hallmark of a variety of human diseases, many of which are age-related. The nematode Caenorhabditis elegans is a well-established model organism that has been widely used in the field to study protein aggregation and toxicity. Its optical transparency enables the direct visualization of protein aggregation by fluorescence microscopy. Moreover, the fast reproductive cycle and short lifespan make the nematode a suitable model to screen for genes and molecules that modulate this process.
However, the quantification of aggregate load in living animals is poorly standardized, typically performed by manual inclusion counting under a fluorescence dissection microscope at a single time point. This approach can result in high variability between observers and limits the understanding of the aggregation process. In contrast, amyloid-like protein aggregation in vitro is routinely monitored by thioflavin T fluorescence in a highly quantitative and time-resolved fashion.
Here, an analogous method is presented for the unbiased analysis of aggregation kinetics in living C. elegans, using a high-throughput confocal microscope combined with custom-made image analysis and data fitting. The applicability of this method is demonstrated by monitoring inclusion formation of a fluorescently labeled polyglutamine (polyQ) protein in the body wall muscle cells. The image analysis workflow allows the determination of the numbers of inclusions at different timepoints, which are fitted to a mathematical model based on independent nucleation events in individual muscle cells. The method described here may prove useful to assess the effects of proteostasis factors and potential therapeutics for protein aggregation diseases in a living animal in a robust and quantitative manner.

Monitoring Protein Aggregation Kinetics In Vivo using Automated Inclusion Counting in Caenorhabditis elegans

Here, a method is presented for the analysis of protein aggregation kinetics in the nematode Caenorhabditis elegans. Animals from an age-synchronized population are imaged at different time points, followed by semiautomated inclusion counting in CellProfiler and fitting to a mathematical model in AmyloFit.

Protein aggregation into insoluble inclusions is a hallmark of a variety of human diseases, many of which are age-related. The nematode Caenorhabditis ...

Protein Extract Preparation and Co-immunoprecipitation from Caenorhabditis elegans

Co-immunoprecipitation methods are frequently used to study protein-protein interactions. Confirmation of hypothesized protein-protein interactions or identification of new ones can provide invaluable information about the function of a protein of interest. Some of the traditional methods for extract preparation frequently require labor-intensive and time-consuming techniques. Here, a modified extract preparation protocol using a bead mill homogenizer and metal beads is described as a rapid alternative to traditional protein preparation methods. This extract preparation method is compatible with downstream co-immunoprecipitation studies. As an example, the method was used to successfully co-immunoprecipitate&#160;C. elegans microRNA Argonaute ALG-1 and two known ALG-1 interactors: AIN-1, and HRPK-1. This protocol includes descriptions of animal sample collection, extract preparation, extract clarification, and protein immunoprecipitation. The described protocol can be adapted to test for interactions between any two or more endogenous, endogenously tagged, or overexpressed C. elegans proteins in a variety of genetic backgrounds.

Protein Extract Preparation and Co-immunoprecipitation from Caenorhabditis elegans

This method describes a protocol for high-throughput protein extract preparation from Caenorhabditis elegans samples and subsequent co-immunoprecipitation.

Co-immunoprecipitation methods are frequently used to study protein-protein interactions. Confirmation of hypothesized protein-protein interactions or ...

Developmental Biology

Stable Isotopic Profiling of Intermediary Metabolic Flux in Developing and Adult Stage Caenorhabditis elegans

Stable isotopic profiling has long permitted sensitive investigations of the metabolic consequences of genetic mutations and/or pharmacologic therapies in cellular and mammalian models. Here, we describe detailed methods to perform stable isotopic profiling of intermediary metabolism and metabolic flux in the nematode, Caenorhabditis elegans. Methods are described for profiling whole worm free amino acids, labeled carbon dioxide, labeled organic acids, and labeled amino acids in animals exposed to stable isotopes either from early development on nematode growth media agar plates or beginning as young adults while exposed to various pharmacologic treatments in liquid culture. Free amino acids are quantified by high performance liquid chromatography (HPLC) in whole worm aliquots extracted in 4% perchloric acid. Universally labeled 13C-glucose or 1,6-13C2-glucose is utilized as the stable isotopic precursor whose labeled carbon is traced by mass spectrometry in carbon dioxide (both atmospheric and dissolved) as well as in metabolites indicative of flux through glycolysis, pyruvate metabolism, and the tricarboxylic acid cycle. Representative results are included to demonstrate effects of isotope exposure time, various bacterial clearing protocols, and alternative worm disruption methods in wild-type nematodes, as well as the relative extent of isotopic incorporation in mitochondrial complex III mutant worms (isp-1(qm150)) relative to wild-type worms. Application of stable isotopic profiling in living nematodes provides a novel capacity to investigate at the whole animal level real-time metabolic alterations that are caused by individual genetic disorders and/or pharmacologic therapies.

Stable Isotopic Profiling of Intermediary Metabolic Flux in Developing and Adult Stage Caenorhabditis elegans

Stable isotopic profiling by gas chromatography mass spectrometric analysis of intermediary metabolic flux is described in the nematode, Caenorhabditis elegans. Methods are detailed for assessing isotopic enrichment in carbon dioxide, organic acids, and amino acids following isotope exposure either during development on agar plates or during adulthood in liquid culture.

Stable isotopic profiling has long permitted sensitive investigations of the metabolic consequences of genetic mutations and/or pharmacologic therapies in ...

Biology

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 ...

Using Caenorhabditis elegans as a Model System to Study Protein Homeostasis in a Multicellular Organism

The folding and assembly of proteins is essential for protein function, the long-term health of the cell, and longevity of the organism. Historically, the function and regulation of protein folding was studied in vitro, in isolated tissue culture cells and in unicellular organisms. Recent studies have uncovered links between protein homeostasis (proteostasis), metabolism, development, aging, and temperature-sensing. These findings have led to the development of new tools for monitoring protein folding in the model metazoan organism Caenorhabditis elegans. In our laboratory, we combine behavioral assays, imaging and biochemical approaches using temperature-sensitive or naturally occurring metastable proteins as sensors of the folding environment to monitor protein misfolding. Behavioral assays that are associated with the misfolding of a specific protein provide a simple and powerful readout for protein folding, allowing for the fast screening of genes and conditions that modulate folding. Likewise, such misfolding can be associated with protein mislocalization in the cell. Monitoring protein localization can, therefore, highlight changes in cellular folding capacity occurring in different tissues, at various stages of development and in the face of changing conditions. Finally, using biochemical tools ex vivo, we can directly monitor protein stability and conformation. Thus, by combining behavioral assays, imaging and biochemical techniques, we are able to monitor protein misfolding at the resolution of the organism, the cell, and the protein, respectively.

Using Caenorhabditis elegans as a Model System to Study Protein Homeostasis in a Multicellular Organism

To study the relationship between protein homeostasis, stress and aging, we monitored changes in protein folding by following protein dysfunction, protein localization in the cell and protein stability at the organismal, cellular and protein levels, using the genetically tractable metazoan Caenorhabditis elegans as a model system.

The folding and assembly of proteins is essential for protein function, the long-term health of the cell, and longevity of the organism. Historically, the ...

Detecting Protein Subcellular Localization by Green Fluorescence Protein Tagging and 4',6-Diamidino-2-phenylindole Staining in Caenorhabditis elegans

In this protocol, a green fluorescence protein (GFP) fusion protein and 4',6-diamidino-2-phenylindole (DAPI) staining are used to track protein subcellular localization changes; in particular, a nuclear translocation under a heat stress condition. Proteins react correspondingly to external and internal signals. A common mechanism is to change its subcellular localization. This article describes a protocol to track protein localization that does not require an antibody, radioactive labeling, or a confocal microscope. In this article, GFP is used to tag the target protein EXL-1 in C. elegans, a member of the chloride intracellular channel proteins (CLICs) family, including mammalian CLIC4. An integrated translational exl-1::gfp transgenic line (with a promoter and a full gene sequence) was created by transformation and &#947;-radiation, and stably expresses the gene and gfp. Recent research showed that upon heat stress, not oxidative stress, EXL-1::GFP accumulates in the nucleus. Overlapping the GFP signal with both the nuclei structure and the DAPI signals confirms the EXL-1 subcellular localization changes under stress. This protocol presents two different fixation methods for DAPI staining: ethanol fixation and acetone fixation. The DAPI staining protocol presented in this article is fast and efficient and preserves both the GFP signal and the protein subcellular localization changes. This method only requires a fluorescence microscope with Nomarski, a FITC filter, and a DAPI filter. It is suitable for a small laboratory setting, undergraduate student research, high school student research, and biotechnology classrooms.

Detecting Protein Subcellular Localization by Green Fluorescence Protein Tagging and 4',6-Diamidino-2-phenylindole Staining in Caenorhabditis elegans

This protocol demonstrates how to track a protein nuclear translocation under heat stress by using a green fluorescence protein (GFP) fusion protein as a marker and 4',6-diamidino-2-phenylindole (DAPI) staining. The DAPI staining protocol is fast and preserves the GFP and protein subcellular localization signals.

In this protocol, a green fluorescence protein (GFP) fusion protein and 4',6-diamidino-2-phenylindole (DAPI) staining are used to track protein ...

Combined Nucleotide and Protein Extractions in Caenorhabditis elegans

A single biological sample holds a plethora of information, and it is now common practice to simultaneously investigate several macromolecules to capture a full picture of the multiple levels of molecular processing and changes between different conditions. This protocol presents the method of isolating DNA, RNA, and protein from the same sample of the nematode Caenorhabditis elegans to remove the variation introduced when these biomolecules are isolated from replicates of similarly treated but ultimately different samples. Nucleic acids and protein are extracted from the nematode using the acid guanidinium thiocyanate-phenol-chloroform extraction method, with subsequent precipitation, washing, and solubilization of each. We show the successful isolation of RNA, DNA, and protein from a single sample from three strains of nematode and HeLa cells, with better protein isolation results in adult animals. Additionally, guanidinium thiocyanate-phenol-chloroform-extracted protein from nematodes improves the resolution of larger proteins, with enhanced detectable levels as observed by immunoblotting, compared to the traditional RIPA extraction of protein.
The method presented here is useful when investigating samples using a multiomic approach, specifically for the exploration of the proteome and transcriptome. Techniques that simultaneously assess multiomics are appealing because molecular signaling underlying complex biological phenomena is thought to occur at complementary levels; however, it has become increasingly common to see that changes in mRNA levels do not always reflect the same change in protein levels and that the time of collection is relevant in the context of circadian regulations. This method removes any intersample variation when assaying different contents within the same sample (intrasample.)

Combined Nucleotide and Protein Extractions in Caenorhabditis elegans

Here, we present a protocol for the isolation of RNA, DNA, and protein from the same sample, in an effort to reduce variation, improve reproducibility, and facilitate interpretations.

A single biological sample holds a plethora of information, and it is now common practice to simultaneously investigate several macromolecules to capture ...

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

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

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

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

Monitoring Protein Aggregation Kinetics In Vivo using Automated Inclusion Counting in Caenorhabditis elegans

Protein Extract Preparation and Co-immunoprecipitation from Caenorhabditis elegans

Stable Isotopic Profiling of Intermediary Metabolic Flux in Developing and Adult Stage Caenorhabditis elegans

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

Using Caenorhabditis elegans as a Model System to Study Protein Homeostasis in a Multicellular Organism

Detecting Protein Subcellular Localization by Green Fluorescence Protein Tagging and 4',6-Diamidino-2-phenylindole Staining in Caenorhabditis elegans

Combined Nucleotide and Protein Extractions in Caenorhabditis elegans