This work was supported by the University of Colorado School of Medicine Dean&#39;s Strategic Research Infrastructure award, R01 HL086680-09 and 1R35HL139726-01, to E.N.G. and UCD CFReT fellowship award (HE). The authors thank Dr. Sandra Eaton and Dr. Gareth Eaton (University of Denver), Dr. Gerald Rosen and Dr. Joseph P. Kao (University of Maryland), and Dr. Sujatha Venkataraman (University of Colorado Denver) for helpful discussions, and Joanne Maltzahn,&#160;Ashley Trumpie and Ivy McDermott (University of Colorado Denver) for technical support.

All animal studies were approved by the University of Colorado Denver Institutional Animal Care and Use Committee.
1. Preparation of Reagents
<ol>
	<li>Diethylenetriaminepentaacetic acid (DTPA) stock (150 mM)
	<ol>
		<li>Add 2.95 g of DTPA (393.35 g/mol) to 10 mL of deionized water.</li>
		<li>To dissolve DTPA, add 1 M NaOH dropwise and bring to a pH of 7.0.</li>
		<li>Bring the volume to 50 mL with water for a final DTPA concentration of 150 mM, and store at 4 &#176;C.</li>
	</ol>
	</li>
	<li>Phosphate buffer saline (PBS) (50 mM, pH 7.4)
	<ol>
		<li>Prepare 5 M of sodium chloride (NaCl) (58.44 g/mol; 29.22 g/100 mL).</li>
		<li>Prepare 1 M of potassium phosphate dibasic HK2PO4 (174.18 g/mol; 17.42 g/100 mL)</li>
		<li>Prepare 1 M of potassium phosphate monobasic KH2PO4 (136.1 g/mol; 13.61 g/100 mL). Mix 3 mL of 5 M NaCl with 4.24 mL of 1 M potassium phosphate dibasic and 0.760 mL of 1 M potassium phosphate monobasic. Check the pH.</li>
		<li>Bring the volume to 100 mL with deionized water.</li>
		<li>Store at room temperature (RT) for short-term (days) and at 4 &#176;C for long-term (weeks) storage.</li>
	</ol>
	</li>
	<li>Krebs-Henseleit buffer (KHB) containing 100 &#181;M DTPA 
	<ol>
		<li>In 50 mL conical centrifuge tube, add 33.3 &#181;L of 150 mM DTPA stock solution.</li>
		<li>Bring to a 50 mL volume with Krebs-Henseleit buffer (KHB).</li>
		<li>Prepare fresh buffer with DTPA each day and keep it at RT.</li>
	</ol>
	</li>
	<li>Tris-EDTA buffer containing sucrose
	<ol>
		<li>Prepare 0.5 M Tris stock: dissolve 15.14 g of Tris base (121.14 g/mol) in 150 mL of deionized water. Using HCl, adjust the pH to 7.8 and bring to a final volume of 250 mL.</li>
		<li>Dissolve 21.4 g of sucrose (342.29 g/mol; final concentration = 0.25 mM) in 150 mL of deionized water.</li>
		<li>Add 5 mL of Tris stock to sucrose to achieve a 10 mM final Tris concentration.</li>
		<li>Add 0.5 mL of 0.5 M EDTA stock to Tris-sucrose to achieve a 1 mM final concentration.</li>
		<li>Check the pH and adjust it to 7.4.</li>
		<li>Bring to a final volume of 250 mL with deionized water and store at 4 &#176;C.</li>
	</ol>
	</li>
	<li>Bovine erythrocyte Cu/Zn superoxide dismutase (SOD) stock (30,000 U/mL) 
	<ol>
		<li>Reconstitute 30,000 U of SOD in 1 mL of PBS (approximately 5.7 mg, depending on activity of SOD lot).</li>
		<li>Mix well, aliquot, and store at -20&#176; C for short-term (6-12 months) and at -80 &#176;C for long-term storage.</li>
	</ol>
	</li>
	<li>SOD working solution (1000 U/mL)
	<ol>
		<li>Transfer a 30 &#181;L aliquot of 30,000 U/mL SOD stock into a &#160;870 &#181;L of sterile PBS.</li>
		<li>Keep the solution on ice, and use it fresh.</li>
	</ol>
	</li>
	<li>Phorbol 12-myristate 13-acetate (PMA) stock (5 mM)
	<ol>
		<li>Dissolve 1 mg of PMA (616.83 g/mol) in 325 &#181;L of DMSO (final concentration = 5 mM).</li>
		<li>Aliquot a 5 mM PMA solution and store it at -20 &#176;C.</li>
	</ol>
	</li>
	<li>PMA working solution (125 &#181;M)
	<ol>
		<li>Dilute a 10 &#181;L aliquot of 5 mM PMA stock into 390 &#181;L of sterile PBS.</li>
		<li>Keep the solution on ice and use it fresh.</li>
		<li>For a vehicle control for PMA, use 10 &#181;L of DMSO in 390 &#181;L of PBS.</li>
	</ol>
	</li>
	<li>Diphenyliodonium chloride (DIP) (2.5 mM)
	<ol>
		<li>Dissolve 3.2 mg of DIP (316.57 g/mol) in 4 mL to obtain a 2.5 mM stock.</li>
		<li>Prepare the solution and use it fresh.</li>
	</ol>
	</li>
	<li>Deferoxamine mesylate salt (DFO) (20 mM)
	<ol>
		<li>Dissolve 4.5 mg of DFO (656.79 g/ mol) in 340 &#181;L to obtain a 20 mM stock.</li>
		<li>Prepare the solution and use it fresh.</li>
	</ol>
	</li>
	<li>Preparation of antimycin A (AA) stock (5 mM)
	<ol>
		<li>Dissolve 5.4 mg of AA (532 g/mol) in 2 mL of ethanol (final concentration = 5 mM).</li>
		<li>Aliquot the stock in glass vials and store at -20 &#176;C.</li>
	</ol>
	</li>
	<li>Preparation of spin probes 
	<ol>
		<li>Bubble 50 mM phosphate buffer containing 100 &#181;M DTPA with nitrogen for 30 min to remove dissolved oxygen from the buffer.</li>
		<li>Remove the spin probe from the -20 &#176;C freezer and allow the container to come to RT (10-15 min).</li>
		<li>Weigh out 2.4 mg of 1-hydroxy-3-methoxycarbonyl-2,2,5,5-tetramethylpyrrolidine&#183;HCl (CMH) (237.8 g/mol)</li>
		<li>Dissolve CMH into 1 mL of the deoxygenated phosphate buffer for a final concentration of 10 mM.</li>
		<li>Weigh out 5 mg of 1-hydroxy-4-[2-triphenylphosphonio)-acetamido]-2,2,6,6-tetramethylpiperidine,1-hydroxy-2,2,6,6-tetramethyl-4-[2-(triphenylphosphonio)acetamido]piperidinium dichloride (mito-TEMPO-H) (529.1 g/mol).</li>
		<li>Dissolve mito-TEMPO-H into 1 mL of the deoxygenated phosphate buffer for a final concentration of 9.5 mM.</li>
		<li>Weigh out 4.9 mg of 1-hydroxy-3-carboxy-2,2,5,5-tetramethylpyrrolidine&#183;HCl (CPH) (223.7 g/mol).</li>
		<li>Dissolve CPH into 1 mL of the deoxygenated phosphate buffer for a final concentration of 22 mM.</li>
		<li>Aliquot and store at -80 &#176;C (freeze-thaw is not recommended).</li>
	</ol>
	</li>
</ol>
2. Detection of Superoxide in vitro
<ol>
	<li>Detection of total, extracellular, and intracellular superoxide in PMA-stimulated RAW 264.7 cells at RT
	<ol>
		<li>Following proper aseptic technique, thaw RAW 264.7 cells and passage them in DMEM media supplemented with 10% FBS (low endotoxin-free) and 1% antimycotic/ampicillin at 37 &#176;C in CO2 incubator.</li>
		<li>Seed RAW 264.7 cells at 1 x 106 cells/well into 6-well plates one day prior to treatment.</li>
		<li>Gently remove media and wash the cells once with 1 mL of KHB buffer.</li>
		<li>Add KHB containing 100 &#181;M DTPA to each well, and treat in a total volume of 500 &#181;L with the following:</li>
		<li>For wells pretreated with SOD, add 15 &#181;L/well of SOD working solution (1000 U/mL; final concentration of SOD = 30 U/mL) and incubate for 10 min at 37 &#176;C prior to addition of CMH and PMA.</li>
		<li>Add 12.5 &#181;L/well of 10 mM CMH stock (final concentration = 0.25 mM).</li>
		<li>Add 40 &#181;L/well of 125 &#181;M PMA working solution (final concentration = 10 &#181;M) or 40 &#181;L vehicle (stock 10 &#181;L of DMSO in 390 &#181;L of PBS).</li>
		<li>Incubate for 50 min at 37 &#176;C in a CO2 incubator.</li>
		<li>Remove the plates from the incubator and place them immediately on ice.</li>
		<li>Collect buffer from each well in separate, 1.5 mL, labeled tubes. Keep on ice throughout.</li>
		<li>Add 100 &#181;L of fresh KHB buffer containing 100 &#181;M DTPA, gently scrape the cells, and resuspend by pipetting up and down several times. Keep on ice throughout cell resuspension.</li>
		<li>Load the sample collected in steps 2.1.10 and 2.1.11 (50 &#181;L) in each of the capillary tubes. Seal both ends and run the EPR. 
		NOTE: Always test a tube or well (without cells) containing the probe in buffer (same concentration = 0.25 mM), treated under the same conditions as the cells (same incubation time and temperature) as a control, since background intensity of the probe is temperature- and time-dependent.</li>
		<li>Set the EPR acquisition parameters to the following: microwave frequency = 9.65 GHz; center field = 3432 G; modulation amplitude = 2.0 G; sweep width = 80 G; microwave power = 19.9 mW; total number of scans = 10; sweep time = 12.11 s; and time constant = 20.48 ms.</li>
	</ol>
	</li>
	<li>Detection of mitochondrial superoxide in RAW 264.7 cells
	<ol>
		<li>Follow steps 2.1.1 and 2.1.2 to seed RAW 264.7 cells one day prior to the experiment.</li>
		<li>Remove media and wash the cells once with 1 mL of KHB buffer.</li>
		<li>Add 200 &#181;L of KHB containing 100 &#181;M DTPA to each well.</li>
		<li>Add 5.3 &#181;L/well of 9.5 mM mito-TEMPO-H stock (final concentration = 0.25 mM)</li>
		<li>Incubate for 10 min at RT.</li>
		<li>Add 1 &#181;L/well of antimycin A (AA), 5 mM stock solution in ethanol (final concentration = 25 &#181;M).</li>
		<li>Incubate for 50 min at 37 &#176;C in a CO2 incubator.</li>
		<li>Remove the plates from the incubator and place them immediately on ice.</li>
		<li>Gently scrape the cells and resuspend by pipetting up and down. Keep on ice.</li>
		<li>Load the sample in a capillary tube. Seal both ends.</li>
		<li>See the previous section for EPR setting.</li>
	</ol>
	</li>
	<li>Detection of superoxide in RAW 264.7 cells at 77 K
	<ol>
		<li>Place the buffer collected in step 1.1.10 in pre-prepared PTFE tubing 1-2 inches in length (3/16&#34; OD x 1/8&#34; ID). Make sure the PTFE tubing is straight so it can be easily inserted and removed from the finger dewar. Use a rubber stopper to close one end of the PTFE tubing, pipette the buffer or cell suspension (100 to 150 &#181;L) into the PTFE tubing, and seal the tubing with a second stopper.</li>
		<li>Flash freeze the sample in liquid nitrogen. The sample can be transferred to a labeled cryopreservation tube for storage at -80 &#176;C or run immediately.</li>
		<li>Fill the finger dewar with liquid nitrogen and insert the PTFE tubing containing the sample into the finger dewar. Make sure the sample is centered in the active space of the resonator and run EPR at 77 K. 
		NOTE: Start the nitrogen gas flow to your spectrometer 15-30 min before the measurements, and continue this flow throughout the measurements to prevent water condensation in the resonator.</li>
		<li>Set EPR acquisition parameters to the following: microwave frequency = 9.65 GHz; center field = 3438 G; modulation amplitude = 4.0 G; sweep width = 150 G; microwave power = 0.316 mW; total number of scans = 10; sweep time = 60 s; and time constant = 1.28 ms.</li>
	</ol>
	</li>
</ol>
3. EPR Measurements in Fluids
<ol>
	<li>Whole blood 
	<ol>
		<li>Treat mice (8-12 weeks old) with a single dose of intratracheal bleomycin (Bleo; 100 &#181;L at 1 U/mL) dissolved in PBS or PBS alone as previously described16,17.</li>
		<li>Euthanize mice by administering inhaled isoflurane (1.5-4%) followed by exsanguination and cervical dislocation. Aspirate blood through the right ventricle into a syringe coated with heparin (1000 USP/mL) containing 100 &#181;M DTPA and transfer to a 1.5 mL tube.</li>
		<li>In a separate 1.5 mL tube, add 15 &#181;L of PBS containing 100 &#181;M DTPA and 3 &#181;L of CMH (10 mM) to 132 &#181;L of blood for a total volume of 150 &#181;L and final CMH concentration of 0.2 mM.</li>
		<li>Incubate blood for 10 min at 37 &#176;C in a water bath.</li>
		<li>Remove the tubes from water bath.</li>
		<li>Take an aliquot by loading blood in a capillary tube and run EPR at RT with the following EPR acquisition parameters: microwave frequency = 9.65 GHz; center field = 3432 G; modulation amplitude = 1.0 G; sweep width = 80 G; microwave power = 19.9 mW; total number of scans = 3; sweep time = 12.11 s; and time constant = 20.48 ms. Alternatively, samples can be flash frozen as described in step 2.3 for measurements at 77 K. EPR acquisition parameters are the following: microwave frequency = 9.65 GHz; center field = 3438 G; modulation amplitude = 4.0 G; sweep width = 150 G; microwave power = 0.316 mW; total number of scans = 2; sweep time = 60 s; and time constant = 1.28 ms.</li>
	</ol>
	</li>
	<li>Bronchoalveolar lavage fluid (BALF)
	<ol>
		<li>After euthanasia (see step 3.1.2), collect BALF by slowly instilling and withdrawing 1 mL of PBS containing 100 &#181;M DTPA three times in a syringe via a cannula placed in the trachea.</li>
		<li>In a 1.5 mL tube, treat 200 &#181;L of BALF with 4 &#181;L of CMH (10 mM) to obtain a final concentration of 0.2 mM.</li>
		<li>Incubate BALF for 50 min at 37 &#176;C in a water bath.</li>
		<li>Take tubes out of the water bath and place them on ice.</li>
		<li>Load BALF in a capillary tube and run EPR at RT with the same EPR settings as used in step 1.1.13, or flash freeze in liquid nitrogen as described in step 2.3.</li>
	</ol>
	</li>
	<li>EPR measurements in blood and BALF at 77 K
	<ol>
		<li>Follow the protocol above to collect blood (steps 3.1.1. to 3.1.4) and BALF (steps 3.2.1 to 3.2.4).</li>
		<li>Place 150 &#181;L of the treated blood or BALF in PTFE tubing (1-2 in). Use a rubber stopper to close one end of the PTFE tubing prior to adding the sample and another stopper to seal the tubing.</li>
		<li>Flash freeze the sample in liquid nitrogen.</li>
		<li>See section 2.3 for details on running EPR in frozen samples in PTFE tubing using the finger dewar at 77 K. Run Frozen CMH treated Blood samples with in a week.</li>
	</ol>
	</li>
</ol>
4. EPR Measurements on Lung Tissue
<ol>
	<li>Flash frozen lung tissue 
	<ol>
		<li>After collecting the BALF in step 3.2.1, the chest is opened and lungs flushed with 10 mL of cold PBS via the right ventricle to remove blood. Flash freeze the lung tissue in liquid nitrogen. Frozen lung tissue can be stored at -80 &#176;C for up to 6 months until use for EPR measurements.</li>
		<li>Stabilize the lung tissue on dry ice with tweezers and cut multiple small pieces (5-15 mg) of lung tissue using a single-edge blade.</li>
		<li>Weigh the tissue in a 1.5 mL tube, place the tube on the scale and tare the scale, then add the tissue pieces and record the weight.</li>
		<li>To the tissue in the 1.5 mL tube, add 196 &#181;L of KHB containing DTPA and 4 &#181;L of CMH (0.2 mM) to achieve a 200 &#181;L total volume.</li>
		<li>Incubate for 1 h at 37 &#176;C in a water bath.</li>
		<li>Spin down (for a few seconds) in a microcentrifuge at 3,884 x g.</li>
		<li>Place on ice and pipette 150 &#181;L of the supernatant into the PTFE tubing and freeze for the 77 K measurements as described in section 2.3. 
		NOTE: For this method, the heterogeneity of the injury needs to be considered. For a bleomycin-induced lung injury, given that it is a highly heterogeneous injury, it is recommended to cut several tissue pieces from different parts of the lung from each mouse. Alternatively, a larger piece of tissue can be homogenized in KHB buffer containing 100 &#181;M DTPA at a 1:6 weight-to-volume ratio (mg/&#181;L) as described below.</li>
	</ol>
	</li>
	<li>Fresh lung tissue preserved in sucrose buffer 
	<ol>
		<li>Flush the lavaged lungs with cold PBS to remove blood as done in step 3.1.2.</li>
		<li>Homogenize the fresh lung tissue in Tris-EDTA buffer containing 0.25 M sucrose with a 1:6 lung/buffer (mg/&#181;L) ratio using Dounce tissue grinder with a glass or PTFE pestle.</li>
		<li>Add 50 &#181;L of the lung homogenate to 450 &#181;L of KHB containing 100 &#181;M DTPA.</li>
		<li>In a 1.5 mL tube (in a total volume of 100 &#181;L), to 98 &#181;L of lung homogenate in KHB, add 2 &#181;L of CMH of 10 mM stock to obtain a final concentration of 0.2 mM.</li>
		<li>Incubate for 20 min 37 &#176;C in a water bath.</li>
		<li>Place the samples on ice and load them in a capillary tube. Run EPR at RT (settings used in step 2.1.13).</li>
		<li>To test the contribution of specific species and sources using different inhibitors, pre-treat 88 &#181;L of lung homogenate +/- inhibitor, adjusting with KHB to achieve a final volume of 98 &#181;L. In this experiment, the inhibitors included 10 &#181;L of SOD (100 U/mL), 4 &#181;L of deferoxamine (DFO; final concentration = 800 &#181;M), or 4 &#181;L of diphenyliodonium chloride (DIP; final concentration = 100 &#181;M). Incubate for 20 min at 37 &#176;C in a water bath.</li>
		<li>Add 2 &#181;L of CMH and incubate for another 20 min at 37 &#176;C, followed by EPR measurements as described above. Include a one-time matched blank sample with CMH KHB containing sucrose buffer. Alternatively, store aliquots of the remaining lung homogenates (step 3.1.2) at -80 &#176;C for future measurements. 
		NOTE: The total volume can be scaled as needed.</li>
	</ol>
	</li>
	<li>EPR measurements on lung tissue from mice injected with spin probes in vivo (at RT using tissue cell)
	<ol>
		<li>Prepare CPH stock solution by dissolving 4.9 mg of CPH in 1 mL of filtered and deoxygenated 50 mM phosphate buffer.</li>
		<li>Anesthetize mice with inhaled isoflurane (1.5-4%) for 20-30 seconds until unresponsive to toe pinch. Inject mice via retroorbital route with 100 &#181;L of CPH spin probe for a 25 g mouse body weight (final dose = 20 mg/kg), and allow the probe to circulate for 1 h. Immediately after retroorbital injection, add one drop of 0.5% proparacaine HCl on the eye area to prevent eye pain and dryness. Monitor mice for 1 h and proceed to tissue harvesting.</li>
		<li>Harvest the lung tissue as described above and flash freeze the lungs.</li>
		<li>Cut 20-30 mg of frozen tissue on dry ice and record the exact weight.</li>
		<li>Gently wipe the tissue with cleaning wipes to absorb any surface water.</li>
		<li>Place the tissue within the window of the tissue cell (an accessory allows EPR measurements for tissue samples) and run EPR to determine total spins. The data can be expressed as total spins per mg of tissue.</li>
	</ol>
	</li>
</ol>
5. Data Analysis
<ol>
	<li>Simulate the EPR spectra using SpinFit module incorporated in the Xenon software of the bench-top EMXnano EPR spectrometer. Determine the nitroxide concentration by the SpinCount module. Alternatively, a calibration curve of a stable nitroxide such as 4-hydroxy-TEMPO or TEMPOL can be made, and the concentration can be obtained by comparing the intensity of the signal with the sample and standard.</li>
	<li>For the data collected at 77 K, use double integration followed by SpinCount.</li>
</ol>

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E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Therapeutic+Targeting+of+Mitochondrial+Superoxide+in+Hypertension.">Therapeutic Targeting of Mitochondrial Superoxide in Hypertension.</a> Circulation Research. 107 (1), 106-116 (2010).</li><li>Dikalov, S. I., Polienko, Y. F., Kirilyuk, I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Electron+Paramagnetic+Resonance+Measurements+of+Reactive+Oxygen+Species+by+Cyclic+Hydroxylamine+Spin+Probes.">Electron Paramagnetic Resonance Measurements of Reactive Oxygen Species by Cyclic Hydroxylamine Spin Probes.</a> Antioxidants &amp; Redox Signaling. , (2017).</li><li>Sharma, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=L-Carnitine+preserves+endothelial+function+in+a+lamb+model+of+increased+pulmonary+blood+flow.">L-Carnitine preserves endothelial function in a lamb model of increased pulmonary blood flow.</a> Pediatric Research. 74 (1), 39-47 (2013).</li><li>Berg, K., Ericsson, M., Lindgren, M., Gustafsson, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+High+Precision+Method+for+Quantitative+Measurements+of+Reactive+Oxygen+Species+in+Frozen+Biopsies.">A High Precision Method for Quantitative Measurements of Reactive Oxygen Species in Frozen Biopsies.</a> PloS One. 9 (3), (2014).</li><li>Kozlov, A. V., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=EPR+analysis+reveals+three+tissues+responding+to+endotoxin+by+increased+formation+of+reactive+oxygen+and+nitrogen+species.">EPR analysis reveals three tissues responding to endotoxin by increased formation of reactive oxygen and nitrogen species.</a> Free Radical Biology and Medicine. 34 (12), 1555-1562 (2003).</li><li>Van Rheen, Z., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Lung+Extracellular+Superoxide+Dismutase+Overexpression+Lessens+Bleomycin-Induced+Pulmonary+Hypertension+and+Vascular+Remodeling.">Lung Extracellular Superoxide Dismutase Overexpression Lessens Bleomycin-Induced Pulmonary Hypertension and Vascular Remodeling.</a> American Journal of Respiratory Cell and Molecular Biology. 44 (4), 500-508 (2011).</li><li>Mouradian, G. C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Superoxide+Dismutase+3+R213G+Single-Nucleotide+Polymorphism+Blocks+Murine+Bleomycin-Induced+Fibrosis+and+Promotes+Resolution+of+Inflammation.">Superoxide Dismutase 3 R213G Single-Nucleotide Polymorphism Blocks Murine Bleomycin-Induced Fibrosis and Promotes Resolution of Inflammation.</a> American Journal of Respiratory Cell and Molecular Biology. 56 (3), 362-371 (2017).</li><li>Dikalov, S. I., Li, W., Mehranpour, P., Wang, S. S., Zafari, A. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Production+of+extracellular+superoxide+by+human+lymphoblast+cell+lines:+comparison+of+electron+spin+resonance+techniques+and+cytochrome+C+reduction+assay.">Production of extracellular superoxide by human lymphoblast cell lines: comparison of electron spin resonance techniques and cytochrome C reduction assay.</a> Biochem Pharmacol. 73 (7), 972-980 (2007).</li><li>Kozuleva, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Quantification+of+superoxide+radical+production+in+thylakoid+membrane+using+cyclic+hydroxylamines.">Quantification of superoxide radical production in thylakoid membrane using cyclic hydroxylamines.</a> Free Radical Biology and Medicine. 89, 1014-1023 (2015).</li><li>Chen, K., Swartz, H. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Oxidation+of+Hydroxylamines+to+Nitroxide+Spin+Labels+in+Living+Cells.">Oxidation of Hydroxylamines to Nitroxide Spin Labels in Living Cells.</a> Biochimica Et Biophysica Acta. 970 (3), 270-277 (1988).</li></ol>

The authors have nothing to disclose.

The assessment of free radical production in biological settings is important in understanding redox regulated signaling in health and disease, but the measure of these species is highly challenging due to the short half-life of free radical species and technical limitations with commonly used methods. EPR is a valuable and powerful tool in redox biology, as it is the only unambiguous method for detecting free radicals. In this project, we demonstrate practical EPR methods for designing experiments and preparing samples ...

While measures of oxidative stress and reactive oxygen species are important to the study of diverse diseases across all organ systems, the detection of reactive oxygen species (ROS) is challenging due to a short half-life and high reactivity. An electron paramagnetic resonance (EPR) technique is the most unambiguous method for detecting free radicals. Spin probes have advantages over the more commonly used fluorescent probes. Though fluorescent probes are relatively inexpensive and easy to use and provide rapid, sensitive detection of ROS, they do have serious limitations due to artifactual signals, an inability to calculate ROS concentrations, and a general lack of specificity1.
To facilitate the use of EPR for biological studies, a variety of spin probes have been synthesized that can measure a range of biologically relevant free radical species as well as pO2, pH, and redox states2,3,4,5,6,7. Spin traps have also been developed to capture short-lived radicals and form long-living adducts, which facilitates detection by EPR8. Both classes (spin probes and spin traps) have advantages and limitations. One commonly used class of spin probes are cyclic hydroxylamines, which are EPR-silent and react with short-lived radicals to form a stable nitroxide. Cyclic hydroxylamines react with superoxide 100 times faster than spin traps, enabling them to compete with cellular antioxidants, but they lack specificity and require the use of appropriate controls and inhibitors to identify the radical species or source responsible for the nitroxide signal. While spin traps exhibit specificity, with distinct spectral patterns depending on the trapped species, they have slow kinetics for superoxide spin trapping and are prone to biodegradation of the radical adducts. Applications for spin trapping have been well-documented in biomedical research9,10,11,12,13.
The goal of this project is to demonstrate practical EPR methods for designing experiments and preparing samples to detect superoxide using spin probes in different cellular compartments in vitro and in different tissue compartments in vivo. Several manuscripts have published protocols relevant to these goals, using cell-permeable, cell-impermeable, and mitochondrial targeted spin probes to target different cellular compartments in vitro and process tissue for analysis in mouse models14,15. We build upon this body of literature by validating an approach to measure superoxide using a 1-hydroxy-3-methoxycarbonyl-2,2,5,5-tetramethylpyrrolidine (CMH) spin probe in different cellular compartments in vitro to ensure accurate measurements, highlighting potential technical problems that may skew results. We also provide methods to perform EPR measurements in blood, bronchoalveolar lavage fluid, and lung tissue using the CMH spin probe. These studies compare different methods to process the tissues as well as present a method to inject another spin probe, CPH, into mice prior to harvesting tissue. Finally, we develop a practical method to store samples in polytetrafluoroethylene (PTFE) tubing to allow for the storage and transfer of samples before EPR measurements at 77 K.

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	<tbody>
		<tr height="20">
			<td height="20" style="height:20px;width:931px;">DMEM</td>
			<td style="width:196px;">LifeTech</td>
			<td style="width:224px;">10566-016</td>
			<td style="width:643px;">cell culture media</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Diethylenetriaminepentaacetic acid (DTPA)</td>
			<td>Sigma Aldrich</td>
			<td>D6518-5G</td>
			<td></td>
		</tr>
		<tr height="23">
			<td height="23" style="height:23px;">sodium chloride (NaCl)&#160;</td>
			<td>Fisher Scientific&#160;</td>
			<td>&#160;BP358-212</td>
			<td>used to prepare 50 mM phosphate saline buffer&#160; according to Sigma aldrish&#160;&#160;</td>
		</tr>
		<tr height="24">
			<td height="24" style="height:24px;">potassium phosphate dibasic (HK2PO4 )</td>
			<td>Fisher Scientific&#160;</td>
			<td>&#160;BP363-500</td>
			<td>used to prepare 50 mM phosphate saline buffer&#160; according to Sigma aldrish&#160;&#160;</td>
		</tr>
		<tr height="24">
			<td height="24" style="height:24px;">potassium phosphate monobasic (KH2PO4 )</td>
			<td>Sigma Aldrich</td>
			<td>P-5379</td>
			<td>used to prepare 50 mM phosphate saline buffer&#160; according to Sigma aldrish&#160;&#160;</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Krebs-Henseleit buffer (KHB)&#160;</td>
			<td>(Alfa Aesar, Hill)</td>
			<td>J67820</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Bovine erythrocyte superoxide dismutase (SOD)</td>
			<td>Sigma Aldrich</td>
			<td>&#160;S7571-30KU</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Phorbol 12-myristate 13-acetate (PMA)&#160;</td>
			<td>Sigma Aldrich</td>
			<td>P1585-1MG</td>
			<td>Dissolve in DMSO</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Antimycin A (AA)</td>
			<td>Sigma Aldrich</td>
			<td>A8674-25MG</td>
			<td>Dissolve in Ethanol and store in glass vials(MW used is the averaged molecular weights for four lots)</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">1-Hydroxy-3-methoxycarbonyl-2,2,5,5-tetramethylpyrrolidine . HCl (CMH)</td>
			<td>Enzo Life Sciences</td>
			<td>ALX-430-117-M050</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">1-Hydroxy-3-carboxy-2,2,5,5-tetramethylpyrrolidine . HCl (CPH)</td>
			<td>Enzo Life Sciences</td>
			<td>ALX-430-078-M250</td>
			<td></td>
		</tr>
		<tr height="49">
			<td height="49" style="height:49px;width:931px;">1-Hydroxy-4-[2-triphenylphosphonio)-acetamido]-2,2,6,6-tetramethylpiperidine, 1-Hydroxy-2,2,6,6-tetramethyl-4-[2-(triphenylphosphonio)acetamido]piperidinium dichloride ( mito-TEMPO-H)</td>
			<td style="width:196px;">Enzo Life Sciences</td>
			<td style="width:224px;">ALX-430-171-M005</td>
			<td style="width:643px;"></td>
		</tr>
		<tr height="49">
			<td height="49" style="height:49px;width:931px;">1-Hydroxy-2,2,6,6-tetramethylpiperidin-4-yl-trimethylammonium chloride . HCl (CAT1H)</td>
			<td style="width:196px;">Enzo Life Sciences</td>
			<td style="width:224px;">ALX-430-131-M250</td>
			<td style="width:643px;"></td>
		</tr>
		<tr height="32">
			<td height="32" style="height:32px;width:931px;">Heparin&#160;</td>
			<td style="width:196px;">Sagent Pharmaceuticals</td>
			<td style="width:224px;">NDC 25021-400-10</td>
			<td style="width:643px;"></td>
		</tr>
		<tr height="30">
			<td height="30" style="height:30px;">Diphenyliodonium chloride&#160;</td>
			<td style="width:196px;">Sigma Aldrich</td>
			<td>43088</td>
			<td style="width:643px;"></td>
		</tr>
		<tr height="26">
			<td height="26" style="height:26px;width:931px;">Deferoxamin mesylate salt</td>
			<td>Sigma Aldrich</td>
			<td style="width:224px;">D9533-1G</td>
			<td style="width:643px;"></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Critoseal</td>
			<td>Leica</td>
			<td>39215003</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">BRAND disposable BLAUBRAND&#160;micropipettes, intraMark</td>
			<td>Sigma Aldrich</td>
			<td>708733</td>
			<td>Capillaries</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:931px;">PTFE FRACTIONAL FLUOROPOLYMER TUBING 
			3/16&#8221; OD x 1/8&#8221; ID</td>
			<td>NORELL</td>
			<td>1598774A</td>
			<td>Teflon tubing&#160;</td>
		</tr>
		<tr height="35">
			<td height="35" style="height:35px;">SILICONE RUBBER STOPPERS FOR NMR SAMPLE TUBES&#160; FOR THIN WALL TUBES HAVING AN OD OF 4mm-5mm (3.2mm TO 4.2mm ID) TS-4-5-SR</td>
			<td>NORELL</td>
			<td>94987</td>
			<td></td>
		</tr>
		<tr height="35">
			<td height="35" style="height:35px;">EMXnano Bench-Top EPR spectrometer&#160;</td>
			<td>Bruker BioSpin GmbH</td>
			<td>E7004002</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">EMX NANO TISSUE CELL</td>
			<td>Bruker BioSpin GmbH</td>
			<td>E7004542</td>
			<td></td>
		</tr>
	</tbody>
</table>

use of electron paramagnetic resonance in biological samples at ambient temperature and 77 k

The accurate and specific detection of reactive oxygen species (ROS) in different cellular and tissue compartments is essential to the study of redox-regulated signaling in biological settings. Electron paramagnetic resonance spectroscopy (EPR) is the only direct method to assess free radicals unambiguously. Its advantage is that it detects physiologic levels of specific species with a high specificity, but it does require specialized technology, careful sample preparation, and appropriate controls to ensure accurate interpretation of the data. Cyclic hydroxylamine spin probes react selectively with superoxide or other radicals to generate a nitroxide signal that can be quantified by EPR spectroscopy. Cell-permeable spin probes and spin probes designed to accumulate rapidly in the mitochondria allow for the determination of superoxide concentration in different cellular compartments.
In cultured cells, the use of cell permeable 1-hydroxy-3-methoxycarbonyl-2,2,5,5-tetramethylpyrrolidine (CMH) along with and without cell-impermeable superoxide dismutase (SOD) pretreatment, or use of cell-permeable PEG-SOD,&#160;allows for the differentiation of extracellular from cytosolic superoxide. The mitochondrial 1-hydroxy-4-[2-triphenylphosphonio)-acetamido]-2,2,6,6-tetramethyl-piperidine,1-hydroxy-2,2,6,6-tetramethyl-4-[2-(triphenylphosphonio)acetamido] piperidinium dichloride (mito-TEMPO-H) allows for measurement of mitochondrial ROS (predominantly superoxide).
Spin probes and EPR spectroscopy can also be applied to in vivo models. Superoxide can be detected in extracellular fluids such as blood and alveolar fluid, as well as tissues such as lung tissue. Several methods are presented to process and store tissue for EPR measurements and deliver intravenous 1-hydroxy-3-carboxy-2,2,5,5-tetramethylpyrrolidine (CPH) spin probe in vivo. While measurements can be performed at room temperature, samples obtained from in vitro and in vivo models can also be stored at -80 &#176;C and analyzed by EPR at 77 K. The samples can be stored in specialized tubing stable at -80 &#176;C and run at 77 K to enable a practical, efficient, and reproducible method that facilitates storing and transferring samples.

Superoxide detection using CMH was validated using the X/XO superoxide generating system to demonstrate that the nitroxide (CM.) signal was fully inhibited by SOD, while catalase had no effect&#160;(Figure 1A). The total, extracellular superoxide was then evaluated in RAW 264.7 cells by incubating cells with the cell-permeable CMH spin probe +/- SOD pretreatment. The nitroxide concentration was measured in both the cell suspension and buffer, which...

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Use of Electron Paramagnetic Resonance in Biological Samples at Ambient Temperature and 77 K

Electron paramagnetic resonance (EPR) spectroscopy is an unambiguous method to measure free radicals. The use of selective spin probes allows for detection of free radicals in different cellular compartments. We present a practical, efficient method to collect biological samples that facilitate treating, storing, and transferring samples for EPR measurements.

The accurate and specific detection of reactive oxygen species (ROS) in different cellular and tissue compartments is essential to the study of ...

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9.0K Views.  University of Colorado Anschutz Medical Campus. Electron paramagnetic resonance (EPR) spectroscopy is an unambiguous method to measure free radicals. The use of selective spin probes allows for detection of free radicals in different cellular compartments. We present a practical, efficient method to collect biological samples that facilitate treating, storing, and transferring samples for EPR measurements.

Video: Use of Electron Paramagnetic Resonance in Biological Samples at Ambient Temperature and 77 K

Sheathless Capillary Electrophoresis&#8211;Mass Spectrometry for Metabolic Profiling of Biological Samples

In metabolomics, a wide range of analytical techniques is used for the global profiling of (endogenous) metabolites in complex samples. In this paper, a protocol is presented for the analysis of anionic and cationic metabolites in biological samples by capillary electrophoresis&#8211;mass spectrometry (CE-MS). CE is well-suited for the analysis of highly polar and charged metabolites as compounds are separated on the basis of their charge-to-size ratio. A recently developed sheathless interfacing design, i.e., a porous tip interface, is used for coupling CE to electrospray ionization (ESI) MS. This interfacing approach allows the effective use of the intrinsically low-flow property of CE in combination with MS, resulting in nanomolar detection limits for a broad range of polar metabolite classes. The protocol presented here is based on employing a bare fused-silica capillary with a porous tip emitter at low-pH separation conditions for the analysis of a broad array of metabolite classes in biological samples. It is demonstrated that the same sheathless CE-MS method can be used for the profiling of cationic metabolites, including amino acids, nucleosides and small peptides, or anionic metabolites, including sugar phosphates, nucleotides and organic acids, by only switching the MS detection and separation voltage polarity. Highly information-rich metabolic profiles in various biological samples, such as urine, cerebrospinal fluid and extracts of the glioblastoma cell line, can be obtained by this protocol in less than 1 hr of CE-MS analysis.

A protocol for metabolic profiling of biological samples by capillary electrophoresis&#8211;mass spectrometry using a sheathless porous tip interface design is presented.

In metabolomics, a wide range of analytical techniques is used for the global profiling of (endogenous) metabolites in complex samples. In this paper, a ...

Multimer-PAGE: A Method for Capturing and Resolving Protein Complexes in Biological Samples

There are many well-developed methods for purifying and studying single proteins and peptides. However, most cellular functions are carried out by networks of interacting protein complexes, which are often difficult to investigate because their binding is non-covalent and easily perturbed by purification techniques. This work describes a method of stabilizing and separating native protein complexes from unmodified tissue using two-dimensional polyacrylamide gel electrophoresis. Tissue lysate is loaded onto a non-denaturing blue-native polyacrylamide gel, then an electric current is applied until the protein migrates a short distance into the gel. The gel strip containing the migrated protein is then excised and incubated with the amine-reactive cross-linking reagent dithiobis(succinimidyl propionate), which covalently stabilizes protein complexes. The gel strip containing cross-linked complexes is then cast into a sodium dodecyl sulfate polyacrylamide gel, and the complexes are separated completely. The method relies on techniques and materials familiar to most molecular biologists, meaning it is inexpensive and easy to learn. While it is limited in its ability to adequately separate extremely large complexes, and has not been universally successful, the method was able to capture a wide variety of well-studied complexes, and is likely applicable to many systems of interest.

A method for stabilizing and separating native protein complexes from unmodified tissue lysate using an amine-reactive protein cross-linker coupled to a novel two-dimensional polyacrylamide gel electrophoresis (PAGE) system is presented.

There are many well-developed methods for purifying and studying single proteins and peptides. However, most cellular functions are carried out by ...

Analysis of &#946;-Amyloid-induced Abnormalities on Fibrin Clot Structure by Spectroscopy and Scanning Electron Microscopy

This article presents methods for generating in vitro fibrin clots and analyzing the effect of beta-amyloid (A&#946;) protein on clot formation and structure by spectrometry and scanning electron microscopy (SEM). A&#946;, which forms neurotoxic amyloid aggregates in Alzheimer&#39;s disease (AD), has been shown to interact with fibrinogen. This A&#946;-fibrinogen interaction makes the fibrin clot structurally abnormal and resistant to fibrinolysis. A&#946;-induced abnormalities in fibrin clotting may also contribute to cerebrovascular aspects of the AD pathology such as microinfarcts, inflammation, as well as, cerebral amyloid angiopathy (CAA). Given the potentially critical role of neurovascular deficits in AD pathology, developing compounds which can inhibit or lessen the A&#946;-fibrinogen interaction has promising therapeutic value. In vitro methods by which fibrin clot formation can be easily and systematically assessed are potentially useful tools for developing therapeutic compounds. Presented here is an optimized protocol for in vitro generation of the fibrin clot, as well as analysis of the effect of A&#946; and A&#946;-fibrinogen interaction inhibitors. The clot turbidity assay is rapid, highly reproducible and can be used to test multiple conditions simultaneously, allowing for the screening of large numbers of A&#946;-fibrinogen inhibitors. Hit compounds from this screening can be further evaluated for their ability to ameliorate A&#946;-induced structural abnormalities of the fibrin clot architecture using SEM. The effectiveness of these optimized protocols is demonstrated here using TDI-2760, a recently identified A&#946;-fibrinogen interaction inhibitor.

Presented here are two methods that can be used individually or in combination to analyze the effect of beta-amyloid on fibrin clot structure. Included is a protocol for creating an in vitro fibrin clot, followed by clot turbidity and scanning electron microscopy methods.

This article presents methods for generating in vitro fibrin clots and analyzing the effect of beta-amyloid (A&#946;) protein on clot formation and ...

Using In Vitro Fluorescence Resonance Energy Transfer to Study the Dynamics Of Protein Complexes at a Millisecond Time Scale

Proteins are the primary operators of biological systems, and they usually interact with other macro- or small molecules to carry out their biological functions. Such interactions can be highly dynamic, meaning the interacting subunits are constantly associated and dissociated at certain rates. While measuring the binding affinity using techniques such as quantitative pull-down reveals the strength of the interaction, studying the binding kinetics provides insights on how fast the interaction occurs and how long each complex can exist. Furthermore, measuring the kinetics of an interaction in the presence of an additional factor, such as a protein exchange factor or a drug, helps reveal the mechanism by which the interaction is regulated by the other factor, providing important knowledge for the advancement of biological and medical research. Here, we describe a protocol for measuring the binding kinetics of a protein complex that has a high intrinsic association rate and can be dissociated quickly by another protein. The method uses fluorescence resonance energy transfer to report the formation of the protein complex in vitro, and it enables monitoring the fast association and dissociation of the complex in real time on a stopped-flow fluorimeter. Using this assay, the association and dissociation rate constants of the protein complex are quantified.

Protein-protein interactions are critical for biological systems, and studies of the binding kinetics provide insights into the dynamics and function of protein complexes. We describe a method that quantifies the kinetic parameters of a protein complex using fluorescence resonance energy transfer and the stopped-flow technique. &#160;

Proteins are the primary operators of biological systems, and they usually interact with other macro- or small molecules to carry out their biological ...

In Vivo Calcium Imaging in C. elegans Body Wall Muscles

The model organism C. elegans provides an excellent system to perform in vivo calcium imaging. Its transparent body and genetic manipulability allow for the targeted expression of genetically encoded calcium sensors. This protocol outlines the use of these sensors for the in vivo imaging of calcium dynamics in targeted cells, specifically the body wall muscles of the worms. By utilizing the co-expression of presynaptic channelrhodopsin, stimulation of acetylcholine release from excitatory motor neurons can be induced using blue light pulses, resulting in muscle depolarization and reproducible changes in cytoplasmic calcium levels. Two worm immobilization techniques are discussed with varying levels of difficulty. Comparison of these techniques demonstrates that both approaches preserve the physiology of the neuromuscular junction and allow for the reproducible quantification of calcium transients. By pairing optogenetics and functional calcium imaging, changes in postsynaptic calcium handling and homeostasis can be evaluated in a variety of mutant backgrounds. Data presented validates both immobilization techniques and specifically examines the roles of the C. elegans sarco(endo)plasmic reticular calcium ATPase and the calcium-activated BK potassium channel in the body wall muscle calcium regulation.

In Vivo Calcium Imaging in C. elegans Body Wall Muscles

This method provides a way to couple optogenetics and genetically encoded calcium sensors to image baseline cytosolic calcium levels and changes in evoked calcium transients in the body wall muscles of the model organism C. elegans.

The model organism C. elegans provides an excellent system to perform in vivo calcium imaging. Its transparent body and genetic manipulability allow for ...

Use of Microscale Thermophoresis to Measure Protein-Lipid Interactions

The ability to determine the binding affinity of lipids to proteins is an essential part of understanding protein-lipid interactions in membrane trafficking, signal transduction and cytoskeletal remodeling. Classic tools for measuring such interactions include surface plasmon resonance (SPR) and isothermal titration calorimetry (ITC). While powerful tools, these approaches have setbacks. ITC requires large amounts of purified protein as well as lipids, which can be costly and difficult to produce. Furthermore, ITC as well as SPR are very time consuming, which could add significantly to the cost of performing these experiments. One way to bypass these restrictions is to use the relatively new technique of microscale thermophoresis (MST). MST is fast and cost effective using small amounts of sample to obtain a saturation curve for a given binding event. There currently are two types of MST systems available. One type of MST requires labeling with a fluorophore in the blue or red spectrum. The second system relies on the intrinsic fluorescence of aromatic amino acids in the UV range. Both systems detect the movement of molecules in response to localized induction of heat from an infrared laser. Each approach has its advantages and disadvantages. Label-free MST can use untagged native proteins; however, many analytes, including pharmaceuticals, fluoresce in the UV range, which can interfere with determination of accurate KD values. In comparison, labeled MST allows for a greater diversity of measurable pairwise interactions utilizing fluorescently labeled probes attached to ligands with measurable absorbances in the visible range as opposed to UV, limiting the potential for interfering signals from analytes.

Microscale thermophoresis obtains binding constants quickly at low material cost. Either labeled or label free microscale thermophoresis is commercially available; however, label free thermophoresis is not capable of the diversity of interaction measurements that can be performed using fluorescent labels. We provide a protocol for labeled thermophoresis measurements.

The ability to determine the binding affinity of lipids to proteins is an essential part of understanding protein-lipid interactions in membrane ...

Removal and Replacement of Endogenous Ligands from Lipid-Bound Proteins and Allergens

Many major allergens bind to hydrophobic lipid-like molecules, including Mus m 1, Bet v 1, Der p 2, and Fel d 1. These ligands are strongly retained and have the potential to influence the sensitization process either through directly stimulating the immune system or altering the biophysical properties of the allergenic protein. In order to control for these variables, techniques are required for the removal of endogenously bound ligands and, if necessary, replacement with lipids of known composition. The cockroach allergen Bla g 1 encloses a large hydrophobic cavity which binds a heterogeneous mixture of endogenous lipids when purified using traditional techniques. Here, we describe a method through which these lipids are removed using reverse-phase HPLC followed by thermal annealing to yield Bla g 1 in either its Apo-form or reloaded with a user-defined mixture of fatty acid or phospholipid cargoes. Coupling this protocol with biochemical assays reveal that fatty acid cargoes significantly alter the thermostability and proteolytic resistance of Bla g 1, with downstream implications for the rate of T-cell epitope generation and allergenicity. These results highlight the importance of lipid removal/reloading protocols such as the one described herein when studying allergens from both recombinant and natural sources. The protocol is generalizable to other allergen families including lipocalins (Mus m 1), PR-10 (Bet v 1), MD-2 (Der p 2) and Uteroglobin (Fel d 1), providing a valuable tool to study the role of lipids in the allergic response.

This protocol describes the removal of endogenous lipids from allergens, and their replacement with user-specified ligands through reverse-phase HPLC coupled with thermal annealing. 31P-NMR and circular dichroism allow for the rapid confirmation of ligand removal/loading, and the recovery of native allergen structure.

Many major allergens bind to hydrophobic lipid-like molecules, including Mus m 1, Bet v 1, Der p 2, and Fel d 1. These ligands are strongly retained and ...

Fast Grid Preparation for Time-Resolved Cryo-Electron Microscopy

The field of cryo-electron microscopy (cryo-EM) is rapidly developing with new hardware and processing algorithms, producing higher resolution structures and information on more challenging systems. Sample preparation for cryo-EM is undergoing a similar revolution with new approaches being developed to supersede the traditional blotting systems. These include the use of piezo-electric dispensers, pin printing and direct spraying. As a result of these developments, the speed of grid preparation is going from seconds to milliseconds, providing new opportunities, especially in the field of time-resolved cryo-EM where proteins and substrates can be rapidly mixed before plunge freezing, trapping short lived intermediate states. Here we describe, in detail, a standard protocol for making grids on our in-house time-resolved EM device both for standard fast grid preparation and also for time-resolved experiments. The protocol requires a minimum of about 50 &#181;L sample at concentrations of &#8805; 2 mg/mL for the preparation of 4 grids. The delay between sample application and freezing can be as low as 10 ms. One limitation is increased ice thickness at faster speeds and compared to the blotting method. We hope this protocol will aid others in designing their own grid making devices and those interested in designing time-resolved experiments.

Here, we provide a detailed protocol for the use of a rapid grid making device for both fast grid-making and for rapid mixing and freezing to conduct time-resolved experiments.

The field of cryo-electron microscopy (cryo-EM) is rapidly developing with new hardware and processing algorithms, producing higher resolution structures ...

Microcrystal Electron Diffraction of Small Molecules

A detailed protocol for preparing small molecule samples for microcrystal electron diffraction (MicroED) experiments is described. MicroED has been developed to solve structures of proteins and small molecules using standard electron cryo-microscopy (cryo-EM) equipment. In this way, small molecules, peptides, soluble proteins, and membrane proteins have recently been determined to high resolutions. Protocols are presented here for preparing grids of small-molecule pharmaceuticals using the drug carbamazepine as an example. Protocols for screening and collecting data are presented. Additional steps in the overall process, such as data integration, structure determination, and refinement are presented elsewhere. The time required to prepare the small-molecule grids is estimated to be less than 30 min.

Here, we describe the procedures developed in our laboratory for preparing powders of small molecule crystals for microcrystal electron diffraction (MicroED) experiments.

A detailed protocol for preparing small molecule samples for microcrystal electron diffraction (MicroED) experiments is described. MicroED has been ...

Preparing Lamellae from Vitreous Biological Samples Using a Dual-Beam Scanning Electron Microscope for Cryo-Electron Tomography

Presented here is a protocol for preparing cryo-lamellae from plunge-frozen grids of Plasmodium falciparum-infected&#160;human erythrocytes, which could easily be adapted for other biological samples. The basic principles for preparing samples, milling, and viewing lamellae are common to all instruments and the protocol can be followed as a general guide to on-grid cryo-lamella preparation for cryo-electron microscopy (cryoEM) and cryo-electron tomography (cryoET). Electron microscopy grids supporting the cells are plunge-frozen into liquid nitrogen-cooled liquid ethane using a manual or automated plunge freezer, then screened on a light microscope equipped with a cryo-stage. Frozen grids are transferred into a cryo-scanning electron microscope equipped with a focused ion beam (cryoFIB-SEM). Grids are routinely sputter coated prior to milling, which aids dispersal of charge build-up during milling. Alternatively, an e-beam rotary coater can be used to apply a layer of carbon-platinum to the grids, the exact thickness of which can be more precisely controlled. Once inside the cryoFIB-SEM an additional coating of an organoplatinum compound is applied to the surface of the grid via a gas injection system (GIS). This layer protects the front edge of the lamella as it is milled, the integrity of which is critical for achieving uniformly thin lamellae. Regions of interest are identified via SEM and milling is carried out in a step-wise fashion, reducing the current of the ion beam as the lamella reaches electron transparency, in order to avoid excessive heat generation. A grid with multiple lamellae is then transferred to a transmission electron microscope (TEM) under cryogenic conditions for tilt-series acquisition. A robust and contamination-free workflow for lamella preparation is an essential step for downstream techniques, including cellular cryoEM, cryoET, and sub-tomogram averaging. Development of these techniques, especially for lift-out and milling of high-pressure frozen samples, is of high-priority in the field.

Using focused ion beam milling to produce vitreous on-grid lamellae from plunge frozen biological samples for cryo-electron tomography.

Presented here is a protocol for preparing cryo-lamellae from plunge-frozen grids of Plasmodium falciparum-infected&#160;human erythrocytes, which could ...