Research supported by NSF award MCB-1021384.

1. Preparation of Anaerobic Buffer
<ol>
 <li>Prepare 1 L of 100 mM potassium phosphate buffer, pH 7.5. Pour 250 mL of the buffer into a 500-mL B&uuml;chner flask with a stir bar.</li>
 <li>Seal the flask with a rubber stopper and place it on a stir plate. Connect the short tube of the flask to a Schlenk line.</li>
 <li>Degas the buffer under vacuum for 5 hours at room temperature with agitation. During this time period, perform 5 consecutive rounds of vacuum degassing and flushing with argon every hour.</li>
 <li>Flush the flask with argon for 10 seconds and disconnect it from the vacuum manifold. Place the flask inside the glove box.</li>
 <li>Leave the flask open in the glove box overnight with vigorous agitation.</li>
</ol>
2. Removing Oxygen from the Stopped-flow System
<ol>
 <li>Prepare 1 L of 0.1 M sodium acetate, pH 5.0. Pour 125 mL of this buffer into a 250-mL B&uuml;chner flask and perform steps 1.2-1.4.</li>
 <li>Weigh out 5 mg glucose oxidase from Aspergillus niger (181300 U/g) and 0.9 g glucose using a 1.5 mL Eppendorf tube and 50 mL Falcon conical tube, respectively. Place both containers into the glove box.</li>
 <li>Dissolve glucose oxidase and glucose in 50 mL of anaerobic 0.1 M sodium acetate, pH 5.0 (final concentrations of 18.13 U/mL and 100 mM, respectively). Fill the reservoir syringes with this solution (Figure 1).</li>
 <li>Ensure that the drive valves are in the "load" position and fill the drive syringes (Figure 1). Turn the F valve to the "drive" position. Empty the stop syringe by clicking on Empty in the Pro-Data control software. Push the buffer from drive syringe F through the flow circuit by manually raising the corresponding drive ram. Repeat this step ten times in total. Perform the same procedure with the other three drive syringes. Wait one hour.</li>
 <li>Replace the solution of the reservoir syringes with the same buffer containing glucose oxidase and glucose. Repeat step 2.4.</li>
 <li>Repeat step 2.5 and allow the solution to stand in the flow system overnight.</li>
 <li>After overnight incubation, repeat step 2.5.</li>
</ol>
3. Preparation of Oxygen Saturated Buffer
<ol>
 <li>Prepare 100 mM potassium phosphate buffer (pH 7.5) and pour 50 mL into a 50-mL vial with a stir bar. Cap the vial with a Wheaton stopper and a Wheaton aluminum seal.</li>
 <li>Place the vial on ice. Immerse a long needle connected to a tank containing 100% oxygen into the solution. Insert another short needle through the stopper of the vial as a vent.</li>
 <li>Bubble the solution with 100% oxygen for 1 hour with agitation.</li>
 <li>First remove the short needle from the vial and wait 10 seconds before removing the long needle. Place the closed vial in the glove box.</li>
</ol>
4. Preparation of NADPH Solution
<ol>
 <li>Weigh out 1 mg of NADPH in a 1.5 mL Eppendorf tube and place it into the glove box.</li>
 <li>Dissolve the NADPH in 300 &mu;L of anaerobic 100 mM potassium phosphate buffer, pH 7.5. Remove 30 &mu;L of this solution from the glove box to determine the NADPH concentration with a spectrophotometer (&#949;340 nm= 6,270 M-1cm-1).</li>
</ol>
5. Removal of Oxygen from the Enzyme Solution
<ol>
 <li>Take 400 &mu;L of enzyme stock solution from the freezer and thaw in the glove box. The enzyme stock solution (360 &mu;M) was previously prepared in 100 mM potassium phosphate buffer (pH 7.5) containing 100 mM NaCl, and frozen in liquid nitrogen.</li>
 <li>Transfer the enzyme solution into a 25-mL vial with a stir bar, cap the vial with a Wheaton stopper and a Wheaton aluminum seal and remove from the glove box.</li>
 <li>Place the vial on ice and connect it to a Schlenk line by inserting a needle through the stopper of the vial.</li>
 <li>Degas the enzyme solution by performing 5 consecutive rounds of vacuum degassing and flushing with argon every 20 minutes for 1 hour.</li>
 <li>Flush the vial with argon for 10 seconds and disconnect it from the vacuum manifold. Place the vial on ice inside the glove box.</li>
</ol>
6. Reductive Half-reaction: Monitoring Flavin Reduction
<ol>
 <li>Prepare the stopped-flow instrument and the Pro-Data control software for single mixing mode following the manufacturer's protocol.</li>
 <li>Turn on a circulating water bath (15 &deg;C). Remove oxygen from the water by bubbling nitrogen through it for 20 minutes. This prevents the oxygen contamination through the flow circuit.</li>
 <li>Maintain the temperature of the drive syringes and the observation cell at 15 &deg;C by connecting the water bath housing of the stopped-flow to the circulating bath.</li>
 <li>In the Control Panel window of the Pro-Data control software select Photodiode Array, External trigger (to activate stopped-flow operation), and Logarithmic scale.</li>
 <li>Launch the Pro-Data viewer software and define the directory where the data files will be stored.</li>
 <li>Replace the solution of reservoir syringes C and F with anaerobic 100 mM potassium phosphate buffer (pH 7.5). Turn the C and F valves to the "drive" position. Empty the stop syringe by clicking Empty in the Pro-Data control software. Push the buffer from both drive syringes through the flow circuit by manually raising the corresponding drive ram five times in total (click Empty before each movement of the ram).</li>
 <li>In order to ensure that glucose oxidase has been removed completely from the flow circuit, perform step 6.6 four times in total.</li>
 <li>Click on Baseline in the Pro-Data control software.</li>
 <li>Mix 100 &mu;L of enzyme stock solution with 1100 &mu;L of anaerobic 100 mM potassium phosphate buffer (final concentration after mixing in the stopped flow of 15 &mu;M).</li>
 <li>Replace the solution of reservoir syringe F with the enzyme solution. Turn the F valve to the "drive" position. Empty the stop syringe by clicking Empty in the Pro-Data control software. Push the enzyme solution from drive syringe F through the flow circuit by manually raising the corresponding drive ram. Perform this three times in total. In the Pro-Data control software, set the acquisition time to 60 s.</li>
 <li>Ensure that both drive syringes are filled with their corresponding solutions and the drive rams are in contact with the drive syringe pistons. Turn both valves to the "drive" position and click Acquire in the Pro-Data control software to perform the drive. Repeat this step two times more to obtain the data in triplicate. This yields spectra of the oxidized form of the enzyme.</li>
 <li>Using the A452 nm value obtained from the drives, determine the enzyme concentration before mixing (&#949;452 nm= 13,700 M-1cm-1) and prepare 4 mL of a NADPH solution 1.5-fold more concentrated.</li>
 <li>Replace the solution of reservoir syringe C with the NADPH solution and empty and refill the stop syringe three times as described in 6.10.</li>
 <li>Repeat step 6.11. This yields spectra during the reduction of the enzyme.</li>
</ol>
7. Oxidative Half-reaction: Monitoring Flavin Oxidation
<ol>
 <li>Prepare the stopped-flow instrument and the Pro-Data control software for double mixing mode following the manufacturer's protocol.</li>
 <li>Repeat steps 6.2-6.5.</li>
 <li>Replace the solution in the four reservoir syringes with anaerobic 100 mM potassium phosphate buffer (pH 7.5). Turn the F valve to the "drive" position. Empty the stop syringe by clicking Empty in the Pro-Data control software. Push the buffer from the drive syringe F through the flow circuit by manually raising the corresponding drive ram. Perform this step five times in total with each drive syringe.</li>
 <li>Perform step 7.3 four times in total to completely remove the contents from the flow circuit.</li>
 <li>Click Baseline in the Pro-Data control software.</li>
 <li>Mix 200 &mu;L of enzyme stock solution with 1000 &mu;L of anaerobic 100 mM potassium phosphate buffer (final concentration after mixing in the stopped flow of 15 &mu;M).</li>
 <li>Replace the solution of reservoir syringe A with the enzyme solution. Turn the A valve to the "drive" position. Empty the stop syringe by clicking Empty in the Pro-Data control software. Push the enzyme solution from drive syringe A through the flow circuit by manually raising the corresponding drive ram. Perform this step three times in total. In the Pro-Data control software, set the delay time to 0.5 s and acquisition time to 60 s.</li>
 <li>Ensure that all drive syringes are filled with their corresponding solutions and the drive rams are in contact with the drive syringe pistons. Turn all valves to the "drive" position and click Acquire in the Pro-Data control software to perform the drive. Repeat this step two more times to obtain the data in triplicate. This yields spectra of the oxidized form of the enzyme.</li>
 <li>Replace the solution of reservoir syringe B with an NADPH solution 1.5-fold more concentrated than the enzyme solution in syringe A. Turn the B valve to the "drive" position and empty and refill the stop syringe three times as described in 7.7.</li>
 <li>Replace the solution of reservoir syringe C with oxygenated buffer. Turn the C valve to the "drive" position and empty and refill the stop syringe three times as described in 7.7. In the Pro-Data control software, set the delay and acquisition time at 15 and 900 s, respectively.</li>
 <li>Repeat step 7.8. This yields spectra during the reoxidation of the fully reduced enzyme.</li>
</ol>
8. Data Analysis
<ol>
 <li>Launch the Pro-Data converter software. Click on the Options icon and select Save in ProDataCSV.</li>
 <li>Open the directory where the data files were stored. Drag the files to the APL Pro-Data converter window. The data will be saved automatically as CSV format files in the same directory.</li>
 <li>Open the CSV format files using Microsoft Excel (Microsoft, Redmond, WA, USA). Normalize the baseline of the stopped-flow traces by subtracting the corresponding absorption at 700 nm.</li>
 <li>Analyze the corrected traces using KaleidaGraph (Synergy Software, Reading, PA) by fitting the plot of the absorbance at the corresponding maximum versus time to the appropriate exponential function. As shown in Figure 2B, the absorbance at 452 nm was plotted as a function of time in order to determine the rate of flavin reduction after mixing the enzyme and NADPH in the stopped-flow instrument. In this case, the best fit of the data was obtained using a double exponential equation and rates of 0.65 and 0.23 s-1 were calculated for the first and second phase of the reduction, respectively. To determine the rate of formation of the C4a-hydroperoxyflavin intermediate and the reoxidation after reacting the reduced enzyme with oxygen, we analyzed the stopped-flow traces at 380 and 452 nm, respectively (Figure 3B). Using a single exponential equation, we calculated rates of 1.4 and 0.006 s-1 for the first and second step of the oxidative half-reaction, respectively.</li>
</ol>
9. Representative Results
The results from the experiments described in the previous sections show how the reductive half-reaction of Af SidA can be monitored by measuring the changes in the absorbance at 452 nm, which correspond to the changes in the redox state of the flavin. The rate of this step can be determined by fitting the data to the appropriate equation (Figure 2; Step 8.4). The reduction rate obtained (0.65 s-1) is similar to the kcat value calculated with steady-state experiments2, suggesting that the reduction is the rate-determining step of the reaction. Taking advantage of the double mixing mode of the stopped-flow, the rate of oxidation and the intermediates in this half-reaction can be determined (Figure 3; Step 8.4). In the reaction catalyzed by Af SidA, the C4a-hydroperoxyflavin is clearly detected (&lambda;max of 380 nm), and the rate of formation and decay can be calculated. The slow rate of the reoxidation obtained (0.006 s-1) indicates that the C4a-hydroperoxyflavin is very stable in absence the ornithine.
<img alt="Scheme 1" src="/files/ftp_upload/3803/3803scheme1.jpg" /> 
Scheme 1. Mechanism of Af SidA. The isoallozaxine ring of the FAD cofactor is shown. The oxidized flavin (A) binds to NADPH (B) and reacts to form reduced flavin and NADP+ (C). After reaction with molecular oxygen and binding of ornithine, the C4a-hydroperoxyflavin is formed (D). This is the hydroxylating species. After hydroxylation of ornithine, the hydroxyflavin (E) must be dehydrated to form the oxidized enzyme. NADP+ remains bound throughout the catalytic cycle and is the last product to be released (F).
<img alt="Figure 1" src="/files/ftp_upload/3803/3803fig1.jpg" /> 
Figure 1. The stopped-flow instrument. A) Picture of the components of the Applied Photophysics SX20 stopped-flow spectrophotometer. B) Picture of the sample handling unit. C) Scheme of the flow circuit in double mixing mode.
<img alt="Figure 2" src="/files/ftp_upload/3803/3803fig2.jpg" /> 
Figure 2. Anaerobic reduction of SidA with NADPH in the stopped-flow instrument. A) Spectral changes recorded after mixing equal volumes of 30 &mu;M SidA and 45 &mu;M NADPH. The first spectrum (oxidized SidA) and last spectrum (fully reduced SidA) is highlighted in blue and red, respectively. B) Absorbance trace at 452 nm recorded as a function of time.
<img alt="Figure 3" src="/files/ftp_upload/3803/3803fig3.jpg" /> 
Figure 3. Oxidation of SidA in the stopped-flow instrument. A) Spectral changes recorded after mixing equal volumes of the fully reduced SidA and oxygenated buffer. The final concentrations were 15 &mu;M SidA, 22.5 &mu;M NADPH, and 0.95 mM oxygen. The spectrum recorded at 0.034, 4.268, and 727.494 s corresponds to the fully reduced enzyme, the C4a-hydroperoxyflavin intermediate (&lambda;max of 380 nm), and the oxidized enzyme (&lambda;max of 450 nm), respectively. B) Absorbance trace at 382 and 452 nm recorded as a function of time.

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	<li>Hissen, A. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+Aspergillus+fumigatus+siderophore+biosynthetic+gene+sidA,+encoding+L&#45;ornithine+N5&#45;oxygenase,+is+required+for+virulence.">The Aspergillus fumigatus siderophore biosynthetic gene sidA, encoding L&#45;ornithine N5&#45;oxygenase, is required for virulence.</a> Infect. Immun. 73 (9), 5493-5503 (2005).</li><li>Chocklett, S. W., Sobrado, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Aspergillus+fumigatus+SidA+is+a+highly+specific+ornithine+hydroxylase+with+bound+flavin+cofactor.">Aspergillus fumigatus SidA is a highly specific ornithine hydroxylase with bound flavin cofactor.</a> Biochemistry. 49 (31), 6777-6783 (2010).</li><li>Mayfield, J. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Comprehensive+spectroscopic,+steady+state,+and+transient+kinetic+studies+of+a+representative+siderophore&#45;associated+flavin+monooxygenase.">Comprehensive spectroscopic, steady state, and transient kinetic studies of a representative siderophore&#45;associated flavin monooxygenase.</a> J. Biol. Chem. 285 (40), 30375-30388 (2010).</li><li>Fierke, C. A., Hammes, G. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Transient+kinetic+approaches+to+enzyme+mechanisms.">Transient kinetic approaches to enzyme mechanisms.</a> Contemporary enzyme kinetics and mechanism. , (2009).</li><li>Palfey, B. A., McDonald, C. 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J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=(&#956;-1,2-Peroxo)diiron(III/III)+complex+as+a+precursor+to+the+diiron(III/IV)+intermediate+X+in+the+assembly+of+the+iron&#45;radical+cofactor+of+ribonucleotide+reductase+from+mouse.">(&#956;-1,2-Peroxo)diiron(III/III) complex as a precursor to the diiron(III/IV) intermediate X in the assembly of the iron&#45;radical cofactor of ribonucleotide reductase from mouse.</a> Biochemistry. 46 (7), 1925-1932 (2007).</li><li>Pennati, A., Zanetti, G., Aliverti, A., Gadda, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Effect+of+salt+and+pH+on+the+reductive+half-reaction+of+Mycobacterium+tuberculosis+FprA+with+NADPH+Biochemistry.">Effect of salt and pH on the reductive half-reaction of Mycobacterium tuberculosis FprA with NADPH Biochemistry.</a> Biochemistry. 47 (11), 3418-3425 (2008).</li><li>Patil, P. 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We have nothing to disclose.

Enzymes that catalyze redox reactions usually contain cofactors such as hemes and flavins that undergo significant absorbance changes during the catalytic cycle. The oxidized form of the flavin presents absorbance maxima at ~ 360 and 450 nm, and its reduction is typically monitored by following the absorbance decrease at 450 nm7. In general, some transient intermediates are present but form and decay too fast to be measured in regular spectrophotometers. Using the Applied Photophysics SX20 stopped-flow spectrophotometer (or similar instruments), it is possible to measure absorbance changes in the millisecond time scale (dead-time, 2 ms). Here we studied the reductive and oxidative half-reactions of the flavin-dependent monooxygenase Af SidA, serving as a model. The rate of hydride transfer was determined by measuring the change of absorbance at 452 nm after mixing the enzyme with NADPH under anaerobic conditions. Subsequently, taking advantage of the double mixing mode of the stopped-flow instrument, the enzyme was first reacted with NADPH, until full reduction was achieved, then the reduced enzyme-NADP+ complex was mixed with oxygen. Following this procedure, it is possible to detect transient oxygenated flavin intermediates and to measure rates of formation and decay. The identification of these intermediates provides experimental data about the nature of the reacting species in catalysis. In the case of Af SidA, the formation of the C4a-hydroperoxyflavin (typically monitored at 370-380 nm), which is the hydroxylating species. In addition, measuring the rate constant of each step allows one to obtain information about the rate-determining step of the reaction and help to elucidate the kinetic and chemical mechanisms of the enzyme. 
In general, similar approaches can be used for other flavoenzymes, or proteins for which absorbance changes occurred, such as proteins that contain heme, pyridoxal-phosphate, or non-heme iron8-10. A limitation to this method is that large amounts of purified enzyme are required, but this can be overcome by using an expression system with high yields. One determines the optimal protein concentration for recording spectra by using enough protein so that a strong enough signal can be observed, but not too much so that enzyme is not wasted. Typically, the lowest enzyme concentration for flavin-containing enzymes used in stopped-flow experiments is 6-10 &mu;M (after mixing) and is determined using the corresponding molar extinction coefficient of the enzyme. In the case of Af SidA, the percentage of the enzyme bound FAD is 50-65%2. Apo-protein is regarded as inactive in these experiments because a bound FAD cofactor is necessary for catalysis. Another possible limitation to this method is if processes in an enzyme occur faster than 2 ms (dead time of the stopped-flow) they will not be observed, but there are reported strategies where rates can be decreased to overcome this issue. One example for this includes using a high NaCl concentration in the reaction of a ferredoxin-NADP+ reductase11. The scrubbing of oxygen from the flow circuit of the stopped-flow is often a tricky step in this experiment and requires special attention. The glucose oxidase-glucose system described here is used successfully in most laboratories as it is an effective and inexpensive method. However, there are some drawbacks which include the production of H2O2 and for some applications other alternatives as the protocatechuate dioxygenase-protocatechuate system should be considered12. The utilization of an anaerobic glove box makes it easier to ensure anaerobic conditions, but is not essential. Oxygen must be removed from the flow circuit of the stopped-flow as we want the enzyme to be reduced in the absence of oxygen or react with oxygen at concentrations that we specify. Although the stopped-flow is in the glove box, there is oxygen in the flow circuit if we used aerobic buffers in previous experiments. In addition to absorbance measurements, fluorescence and circular dichroism assays can be performed in the Applied Photophysics SX20 stopped-flow spectrophotometer with the corresponding accessories.

<table border="1">
<tbody>
 <tr>
 <td>General Laboratory Equipment</td>
 <td>Company</td>
 <td>Catalogue Number</td>
 </tr>
 <tr>
 <td>Vacuum pump</td>
 <td>Welch</td>
 <td>-</td>
 </tr>
 <tr>
 <td>B&uuml;chner flasks</td>
 <td>Fisher</td>
 <td>70340-500</td>
 </tr>
 <tr>
 <td>Stir bars</td>
 <td>Fisher</td>
 <td>14-512-129</td>
 </tr>
 <tr>
 <td>Stir plates</td>
 <td>Fisher</td>
 <td>11-100-49S</td>
 </tr>
 <tr>
 <td>Schlenk lines</td>
 <td>Kontes Glass</td>
 <td>-</td>
 </tr>
 <tr>
 <td>Argon tank </td>
 <td>Airgas</td>
 <td>AR UPC300</td>
 </tr>
 <tr>
 <td>Nitrogen tank</td>
 <td>Airgas</td>
 <td>NI200</td>
 </tr>
 <tr>
 <td>Nitrogen tank, ultra high purity grade</td>
 <td>Airgas</td>
 <td>NI UHP200</td>
 </tr>
 <tr>
 <td>Oxygen tank</td>
 <td>Airgas</td>
 <td>OX 40</td>
 </tr>
 <tr>
 <td>5% Hydrogen balance nitrogen tank</td>
 <td>Airgas</td>
 <td>X02NI95B200H998</td>
 </tr>
 <tr>
 <td>SX20 Stopped-flow spectrophotometer</td>
 <td>AppliedPhotophysics</td>
 <td>-</td>
 </tr>
 <tr>
 <td>Glove box</td>
 <td>Coy</td>
 <td>-</td>
 </tr>
 <tr>
 <td>Water bath</td>
 <td>Brinkmann Lauda</td>
 <td>-</td>
 </tr>
 <tr>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Supplies</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>50 mL BD Falcon tubes</td>
 <td>Fisher</td>
 <td>14-432-23</td>
 </tr>
 <tr>
 <td>15 mL BD Falcon conical tubes</td>
 <td>Fisher</td>
 <td>05-527-90</td>
 </tr>
 <tr>
 <td>1.5 mL Eppendorf microcentrifuge tubes</td>
 <td>Fisher</td>
 <td>05-402-18</td>
 </tr>
 <tr>
 <td>50 and 25 mL glass vials</td>
 <td>Fisher</td>
 <td>06-402</td>
 </tr>
 <tr>
 <td>Rubber stoppers</td>
 <td>Fisher</td>
 <td>06-447H</td>
 </tr>
 <tr>
 <td>Aluminum seals</td>
 <td>Fisher</td>
 <td>06-406-15</td>
 </tr>
 <tr>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Reagents</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Potassium phosphate, monobasic</td>
 <td>Fisher</td>
 <td>AC2714080025</td>
 </tr>
 <tr>
 <td>Potassium phosphate, dibasic</td>
 <td>Fisher</td>
 <td>P288-500</td>
 </tr>
 <tr>
 <td>Sodium acetate</td>
 <td>Sigma</td>
 <td>S-2889</td>
 </tr>
 <tr>
 <td>Glucose oxidase from A. niger</td>
 <td>Sigma</td>
 <td>G7141-250KU</td>
 </tr>
 <tr>
 <td>D-Glucose</td>
 <td>Fisher</td>
 <td>D16-500</td>
 </tr>
 <tr>
 <td>&beta;-NADPH</td>
 <td>Fisher</td>
 <td>ICN10116783</td>
 </tr>
 <tr>
 <td>L(+)-Ornithine hydrochloride</td>
 <td>Fisher</td>
 <td>ICN10116783</td>
 </tr>
 </tbody>
</table>

monitoring the reductive and oxidative half-reactions of a flavin-dependent monooxygenase using stopped-flow spectrophotometry

Aspergillus fumigatus siderophore A (SidA) is an FAD-containing monooxygenase that catalyzes the hydroxylation of ornithine in the biosynthesis of hydroxamate siderophores that are essential for virulence (e.g. ferricrocin or N',N",N'''-triacetylfusarinine C)1. The reaction catalyzed by SidA can be divided into reductive and oxidative half-reactions (Scheme 1). In the reductive half-reaction, the oxidized FAD bound to Af SidA, is reduced by NADPH2,3. In the oxidative half-reaction, the reduced cofactor reacts with molecular oxygen to form a C4a-hydroperoxyflavin intermediate, which transfers an oxygen atom to ornithine. Here, we describe a procedure to measure the rates and detect the different spectral forms of SidA using a stopped-flow instrument installed in an anaerobic glove box. In the stopped-flow instrument, small volumes of reactants are rapidly mixed, and after the flow is stopped by the stop syringe (Figure 1), the spectral changes of the solution placed in the observation cell are recorded over time. In the first part of the experiment, we show how we can use the stopped-flow instrument in single mode, where the anaerobic reduction of the flavin in Af SidA by NADPH is directly measured. We then use double mixing settings where Af SidA is first anaerobically reduced by NADPH for a designated period of time in an aging loop, and then reacted with molecular oxygen in the observation cell (Figure 1). In order to perform this experiment, anaerobic buffers are necessary because when only the reductive half-reaction is monitored, any oxygen in the solutions will react with the reduced flavin cofactor and form a C4a-hydroperoxyflavin intermediate that will ultimately decay back into the oxidized flavin. This would not allow the user to accurately measure rates of reduction since there would be complete turnover of the enzyme. When the oxidative half-reaction is being studied the enzyme must be reduced in the absence of oxygen so that just the steps between reduction and oxidation are observed. One of the buffers used in this experiment is oxygen saturated so that we can study the oxidative half-reaction at higher concentrations of oxygen. These are often the procedures carried out when studying either the reductive or oxidative half-reactions with flavin-containing monooxygenases. The time scale of the pre-steady-state experiments performed with the stopped-flow is milliseconds to seconds, which allow the determination of intrinsic rate constants and the detection and identification of intermediates in the reaction4. The procedures described here can be applied to other flavin-dependent monooxygenases.5,6

Watch this Scientific Journal Video about Monitoring the Reductive and Oxidative Half-Reactions of a Flavin-Dependent Monooxygenase using Stopped-Flow Spectrophotometry at JoVE.com

Monitoring the Reductive and Oxidative Half-Reactions of a Flavin-Dependent Monooxygenase using Stopped-Flow Spectrophotometry

We describe the use of a stopped-flow instrument to investigate both the reductive and oxidative half-reactions of Aspergillus fumigatus siderophore A (SidA), a flavin-dependent monooxygenase. We then show the spectra corresponding to the species in the reaction of SidA and we calculate the rate constants for their formation.

Aspergillus fumigatus siderophore A (SidA) is an FAD-containing monooxygenase that catalyzes the hydroxylation of ornithine in the biosynthesis of ...

monitoring-reductive-oxidative-half-reactions-flavin-dependent

Research

JoVE Journal

Bioengineering

High-throughput Protein Expression Generator Using a Microfluidic Platform

Rapidly increasing fields, such as systems biology, require the development and implementation of new technologies, enabling high-throughput and high-fidelity measurements of large systems. Microfluidics promises to fulfill many of these requirements, such as performing high-throughput screening experiments on-chip, encompassing biochemical, biophysical, and cell-based assays1. Since the early days of microfluidics devices, this field has drastically evolved, leading to the development of microfluidic large-scale integration2,3. This technology allows for the integration of thousands of micromechanical valves on a single device with a postage-sized footprint (Figure 1). We have developed a high-throughput microfluidic platform for generating in vitro expression of protein arrays (Figure 2) named PING (Protein Interaction Network Generator). These arrays can serve as a template for many experiments such as protein-protein 4, protein-RNA5 or protein-DNA6 interactions.
The device consist of thousands of reaction chambers, which are individually programmed using a microarrayer. Aligning of these printed microarrays to microfluidics devices programs each chamber with a single spot eliminating potential contamination or cross-reactivity Moreover, generating microarrays using standard microarray spotting techniques is also very modular, allowing for the arraying of proteins7, DNA8, small molecules, and even colloidal suspensions. The potential impact of microfluidics on biological sciences is significant. A number of microfluidics based assays have already provided novel insights into the structure and function of biological systems, and the field of microfluidics will continue to impact biology.

We present a microfluidic approach for the expression of protein arrays. The device consists of thousands of reaction chambers controlled by micro-mechanical valves. The microfluidic device is mated to a microarray-printed gene library. These genes are then transcribed and translated on-chip, resulting in a protein array ready for experimental use.

Rapidly increasing fields, such as systems biology, require the development and implementation of new technologies, enabling high-throughput and ...

Monitoring the Wall Mechanics During Stent Deployment in a Vessel

Clinical trials have reported different restenosis rates for various stent designs1. It is speculated that stent-induced strain concentrations on the arterial wall lead to tissue injury, which initiates restenosis2-7. This hypothesis needs further investigations including better quantifications of non-uniform strain distribution on the artery following stent implantation. A non-contact surface strain measurement method for the stented artery is presented in this work. ARAMIS stereo optical surface strain measurement system uses two optical high speed cameras to capture the motion of each reference point, and resolve three dimensional strains over the deforming surface8,9. As a mesh stent is deployed into a latex vessel with a random contrasting pattern sprayed or drawn on its outer surface, the surface strain is recorded at every instant of the deformation. The calculated strain distributions can then be used to understand the local lesion response, validate the computational models, and formulate hypotheses for further in vivo study.

Stent-induced arterial strain distributions are characterized using an optical surface strain measurement system. This visualization technique is used to gain insights into the impact of stent implantation on the host vessel.

Clinical trials have reported different restenosis rates for various stent designs1. It is speculated that stent-induced strain concentrations on the ...

Non-contact, Label-free Monitoring of Cells and Extracellular Matrix using Raman Spectroscopy

Non-destructive, non-contact and label-free technologies to monitor cell and tissue cultures are needed in the field of biomedical research.1-5 However, currently available routine methods require processing steps and alter sample integrity. Raman spectroscopy is a fast method that enables the measurement of biological samples without the need for further processing steps. This laser-based technology detects the inelastic scattering of monochromatic light.6 As every chemical vibration is assigned to a specific Raman band (wavenumber in cm-1), each biological sample features a typical spectral pattern due to their inherent biochemical composition.7-9 Within Raman spectra, the peak intensities correlate with the amount of the present molecular bonds.1 Similarities and differences of the spectral data sets can be detected by employing a multivariate analysis (e.g. principal component analysis (PCA)).10
Here, we perform Raman spectroscopy of living cells and native tissues. Cells are either seeded on glass bottom dishes or kept in suspension under normal cell culture conditions (37 &deg;C, 5% CO2) before measurement. Native tissues are dissected and stored in phosphate buffered saline (PBS) at 4 &deg;C prior measurements. Depending on our experimental set up, we then either focused on the cell nucleus or extracellular matrix (ECM) proteins such as elastin and collagen. For all studies, a minimum of 30 cells or 30 random points of interest within the ECM are measured. Data processing steps included background subtraction and normalization.

Raman spectroscopy is a suitable technique for the non-contact, label-free analysis of living cells, tissue-engineered constructs and native tissues. Source-specific spectral fingerprints can be generated and analyzed using multivariate analysis.

Non-destructive, non-contact and label-free technologies to monitor cell and tissue cultures are needed in the field of biomedical research.1-5 However, ...

Mass Cytometry: Protocol for Daily Tuning and Running Cell Samples on a CyTOF Mass Cytometer

In recent years, the rapid analysis of single cells has commonly been performed using flow cytometry and fluorescently-labeled antibodies. However, the issue of spectral overlap of fluorophore emissions has limited the number of simultaneous probes. In contrast, the new CyTOF mass cytometer by DVS Sciences couples a liquid single-cell introduction system to an ICP-MS.1 Rather than fluorophores, chelating polymers containing highly-enriched metal isotopes are coupled to antibodies or other specific probes.2-5 Because of the metal purity and mass resolution of the mass cytometer, there is no "spectral overlap" from neighboring isotopes, and therefore no need for compensation matrices. Additionally, due to the use of lanthanide metals, there is no biological background and therefore no equivalent of autofluorescence. With a mass window spanning atomic mass 103-203, theoretically up to 100 labels could be distinguished simultaneously. Currently, more than 35 channels are available using the chelating reagents available from DVS Sciences, allowing unprecedented dissection of the immunological profile of samples.6-7
Disadvantages to mass cytometry include the strict requirement for a separate metal isotope per probe (no equivalent of forward or side scatter), and the fact that it is a destructive technique (no possibility of sorting recovery). The current configuration of the mass cytometer also has a cell transmission rate of only ~25%, thus requiring a higher input number of cells.
Optimal daily performance of the mass cytometer requires several steps. The basic goal of the optimization is to maximize the measured signal intensity of the desired metal isotopes (M) while minimizing the formation of oxides (M+16) that will decrease the M signal intensity and interfere with any desired signal at M+16. The first step is to warm up the machine so a hot, stable ICP plasma has been established. Second, the settings for current and make-up gas flow rate must be optimized on a daily basis. During sample collection, the maximum cell event rate is limited by detector efficiency and processing speed to 1000 cells/sec. However, depending on the sample quality, a slower cell event rate (300-500 cells/sec) is usually desirable to allow better resolution between cells events and thus maximize intact singlets over doublets and debris. Finally, adequate cleaning of the machine at the end of the day helps minimize background signal due to free metal.

The steps necessary for daily tuning and optimization of the performance of a CyTOF mass cytometer are described. Comments on optimal sample preparation and flow rate are discussed

In recent years, the rapid analysis of single cells has commonly been performed using flow cytometry and fluorescently-labeled antibodies. However, the ...

Surgical Technique for the Implantation of Tissue Engineered Vascular Grafts and Subsequent In Vivo Monitoring

The development of Tissue Engineered Vessels (TEVs) is advanced by the ability to routinely and effectively implant TEVs (4-5 mm in diameter) into a large animal model. A step by-step protocol for inter-positional placement of the TEV and real-time digital assessment of the TEV and native carotid arteries is described here. In vivo monitoring is made possible by the implantation of flow probes, catheters and ultrasonic crystals (capable of recording dynamic diameter changes of implanted TEVs and native carotid arteries) at the time of surgery. Once implanted, researchers can calculate arterial blood flow patterns, invasive blood pressure and artery diameter yielding parameters such as pulse wave velocity, augmentation index, pulse pressures and compliance. Data acquisition is accomplished using a single computer program for analysis throughout the duration of the experiment. Such invaluable data provides insight into TEV matrix remodeling, its resemblance to native/sham controls and overall TEV performance in vivo.

Surgical Technique for the Implantation of Tissue Engineered Vascular Grafts and Subsequent In Vivo Monitoring

A step-by-step protocol for the inter-positional placement of Tissue Engineered Vessels (TEVs) into the carotid artery of a sheep using end-to-end anastomosis and real-time digital assessment in vivo until animal sacrifice.

The development of Tissue Engineered Vessels (TEVs) is advanced by the ability to routinely and effectively implant TEVs (4-5 mm in diameter) into a large ...

Real Time Monitoring of Intracellular Bile Acid Dynamics Using a Genetically Encoded FRET-based Bile Acid Sensor

F&#246;rster Resonance Energy Transfer (FRET) has become a powerful tool for monitoring protein folding, interaction and localization in single cells. Biosensors relying on the principle of FRET have enabled real-time visualization of subcellular signaling events in live cells with high temporal and spatial resolution. Here, we describe the application of a genetically encoded Bile Acid Sensor (BAS) that consists of two fluorophores fused to the farnesoid X receptor ligand binding domain (FXR-LBD), thereby forming a bile acid sensor that can be activated by a large number of bile acids species and other (synthetic) FXR ligands. This sensor can be targeted to different cellular compartments including the nucleus (NucleoBAS) and cytosol (CytoBAS) to measure bile acid concentrations locally. It allows rapid and simple quantitation of cellular bile acid influx, efflux and subcellular distribution of endogenous bile acids without the need for labeling with fluorescent tags or radionuclei. Furthermore, the BAS FRET sensors can be useful for monitoring FXR ligand binding. Finally, we show that this FRET biosensor can be combined with imaging of other spectrally distinct fluorophores. This allows for combined analysis of intracellular bile acid dynamics and i) localization and/or abundance of proteins of interest, or ii) intracellular signaling in a single cell.

We provide a detailed protocol to study bile acid dynamics in living cells using a genetically encoded BAS FRET sensor. This Bile Acid Sensor represents a unique tool to study (regulation of) bile acid transport and FXR activation in a wide range of cell types.

F&#246;rster Resonance Energy Transfer (FRET) has become a powerful tool for monitoring protein folding, interaction and localization in single cells. ...

Intra-cardiac Side-Firing Light Catheter for Monitoring Cellular Metabolism using Transmural Absorbance Spectroscopy of Perfused Mammalian Hearts

Absorbance spectroscopy of cardiac muscle provides non-destructive assessment of cytosolic and mitochondrial oxygenation via myoglobin and cytochrome absorbance respectively. In addition, numerous aspects of the mitochondrial metabolic status such as membrane potential and substrate entry can also be estimated. To perform cardiac wall transmission optical spectroscopy, a commercially available side-firing optical fiber catheter is placed in the left ventricle of the isolated perfused heart as a light source. Light passing through the heart wall is collected with an external optical fiber to perform optical spectroscopy of the heart in near real- time. The transmission approach avoids numerous surface scattering interference occurring in widely used reflection approaches. Changes in transmural absorbance spectra were deconvolved using a library of chromophore reference spectra, providing quantitative measures of all the known cardiac chromophores simultaneously. This spectral deconvolution approach eliminated intrinsic errors that may result from using common dual wavelength methods applied to overlapping absorbance spectra, as well as provided a quantitative evaluation of the goodness of fit. A custom program was designed for data acquisition and analysis, which permitted the investigator to monitor the metabolic state of the preparation during the experiment. These relatively simple additions to the standard heart perfusion system provide a unique insight into the metabolic state of the heart wall in addition to conventional measures of contraction, perfusion, and substrate/oxygen extraction.

Here we introduce a method for using an intra-ventricle optical catheter in perfused hearts to perform absorbance spectroscopy across the heart wall. The data obtained provides robust information on tissue oxygen tension as well as substrate utilization and membrane potential simultaneously with cardiac performance measures in this ubiquitous preparation.

Absorbance spectroscopy of cardiac muscle provides non-destructive assessment of cytosolic and mitochondrial oxygenation via myoglobin and cytochrome ...

Preclinical Cardiac Electrophysiology Assessment by Dual Voltage and Calcium Optical Mapping of Human Organotypic Cardiac Slices

Human cardiac slice preparations have recently been developed as a platform for human physiology studies and therapy testing to bridge the gap between animal and clinical trials. Numerous animal and cell models have been used to examine the effects of drugs, yet these responses often differ in humans. Human cardiac slices offer an advantage for drug testing in that they are directly derived from viable human hearts. In addition to having preserved multicellular structures, cell-cell coupling, and extracellular matrix environments, human cardiac tissue slices can be used to directly test the effect of innumerable drugs on adult human cardiac physiology. What distinguishes this model from other heart preparations, such as whole hearts or wedges, is that slices can be subjected to longer-term culture. As such, cardiac slices allow for studying the acute as well as chronic effects of drugs. Furthermore, the ability to collect several hundred to a thousand slices from a single heart makes this a high-throughput model to test several drugs at varying concentrations and combinations with other drugs at the same time. Slices can be prepared from any given region of the heart. In this protocol, we describe the preparation of left ventricular slices by isolating tissue cubes from the left ventricular free wall and sectioning them into slices using a high precision vibrating microtome. These slices can then either be subjected to acute experiments to measure baseline cardiac electrophysiological function or cultured for chronic drug studies. This protocol also describes dual optical mapping of cardiac slices for simultaneous recordings of transmembrane potentials and intracellular calcium dynamics to determine the effects of the drugs being investigated.

This protocol describes the procedure for sectioning and culturing human cardiac slices for preclinical drug testing and details the use of optical mapping for recording transmembrane voltage and intracellular calcium signals simultaneously from these slices.

Human cardiac slice preparations have recently been developed as a platform for human physiology studies and therapy testing to bridge the gap between ...

Imaging and Quantification of the Area of Fast-Moving Microbubbles Using a High-Speed Camera and Image Analysis

An experimental and image analysis technique is presented for imaging cavitation bubbles and calculating their area. The high-speed imaging experimental technique and image analysis protocol presented here can also be applied for imaging microscopic bubbles in other fields of research; therefore, it has a wide range of applications. We apply this to image cavitation around dental ultrasonic scalers. It is important to image cavitation to characterize it and to understand how it can be exploited for various applications. Cavitation occurring around dental ultrasonic scalers can be used as a novel method of dental plaque removal, which would be more effective and cause less damage than current periodontal therapy techniques. We present a method for imaging the cavitation bubble clouds occurring around dental ultrasonic scaler tips using a high-speed camera and a zoom lens. We also calculate the area of cavitation using machine learning image analysis. Open source software is used for image analysis. The image analysis presented is easy to replicate, does not require programming experience, and can be modified easily to suit the application of the user.

Cavitation microbubbles are imaged using a high-speed camera attached to a zoom lens. The experimental setup is explained, and image analysis is used to calculate the area of the cavitation. Image analysis is done using ImageJ.

An experimental and image analysis technique is presented for imaging cavitation bubbles and calculating their area. The high-speed imaging experimental ...

Labeling of Extracellular Vesicles for Monitoring Migration and Uptake in Cartilage Explants

Extracellular vesicles (EVs) are used in different studies to prove their potential as a cell-free treatment due to their cargo derived from their cellular source, such as platelet lysate (PL). When used as treatment, EVs are expected to enter the target cells and effect a response from these. In this research, PL-derived EVs have been studied as a cell-free treatment for osteoarthritis (OA). Thus, a method was set up to label EVs and test their uptake on cartilage explants. PL-derived EVs are labeled with the lipophilic dye PKH26, washed twice through a column, and then tested in an in vitro inflammation-driven&#160;OA model for 5 h after particle quantification by nanoparticle tracking analysis (NTA). Hourly, cartilage explants are fixed, paraffined, cut into 6 &#181;m sections to mount on slides, and observed under a confocal microscope. This allows verification of whether EVs enter the target cells (chondrocytes) during this period and analyze their direct effect.

Here, we present a protocol to label platelet lysate-derived extracellular vesicles to monitor their migration and uptake in cartilage explants used as a model for osteoarthritis.