Name: Video: Use of Enzymatic Biosensors to Quantify Endogenous ATP or H2O2 in the Kidney
Uploaded: 2015-10-12T15:00:00.000+00:00
Duration: 10 min
Description: 11.8K Views. Medical College of Wisconsin. Enzymatic microelectrode biosensors enable real-time measurements of extracellular cell signaling in biologically-relevant concentrations. The following protocols extend the applications of biosensors to the ex vivo and in vivo detection of ATP and H2O2 in the kidney.

We appreciate Sarissa Biomedical for their work in developing the sensors used in the present manuscript. This research was supported by the National Heart, Lung, and Blood Institute grants HL108880 (A. Staruschenko),&#160;HL 116264 (A. Cowley) and HL 122662 (A. Staruschenko and A. Cowley),&#160;a project funded by the Medical College of Wisconsin Research Affairs Committee #9306830 (O. Palygin) and &#160;Advancing a Healthier Wisconsin Research and Education Program&#160;#9520217, and the Young Investigator Grant of the National Kidney Foundation (O. Palygin).

The following animal&#160;procedures &#160;adhered to the NIH Guide for the Care and Use of Laboratory Animals. Prior approval was obtained from the Institutional Animal Care and Use Committee (IACUC). 
NOTE: Review of the sensor manufacturer instructions should be done during the experiment design&#160;and prior to their use. Following these instructions will produce optimal results when using the&#160;sensors.
1. Sensor Calibration
<ol>
	<li>Prepare fresh stock solutions prior the start of the experiment.</li>
	<li>Create Buffer A containing 10 mM NaPi buffer, 100 mM NaCl, 1 mM MgCl2, and 2 mM glycerol. Adjust the pH to 7.4 using NaOH. Store at 2-8 &#176;C.</li>
	<li>Using a stock ATP concentration (100 mM) stored at -20 &#176;C, create a fresh 10 mM ATP calibration solution by adding 10 &#181;l stock to 90 &#181;l of Buffer A.</li>
	<li>Rehydrate the ATP sensor by placing its tip (Figure 2) into the rehydration chamber containing Buffer A for at least 10 min at 2-8 &#176;C. 
	NOTE: After&#160;rehydration, the sensors should not be exposed to air for more than 20 sec&#160;or the sensor sensitivity may be reduced. If long exposures to air are anticipated, dip the sensor briefly into a solution of glycerol. The sensors may be used for multiple experiments but these must occur on the same day as sensor rehydration. The sensors can be stored in the rehydration chamber with Buffer A for up to 24 hr.</li>
	<li>Turn on the dual channel potentiostat (Figure 3) and start the recording system software.</li>
	<li>Prepare a calibration chamber with 3 ml of Buffer A. Place the reference electrode into the calibration chamber. Take each sensor from the rehydration chamber, then attach it to the manipulator and insert it into the calibration chamber solution. 
	NOTE: Use a standard silicone coated petri dish as a calibration chamber. Carry out all calibrations and studies in a Faraday cage on a high-performance lab air table to reduce signal noise during the amperometry recordings (Figure 4). Calibrations are best done as close to the start of data collection as possible. For in vivo applications the optimal time for calibration is during the animal post-surgery recovery period.</li>
	<li>Ex vivo calibration
	<ol>
		<li>Perform cyclic voltammetry in the calibration chamber by cycling the sensors from -500 mV to +500 mV at a rate of 100 mV/sec for 10 cycles. This greatly improves the sensitivity of the sensors. See Figure 5 for the traces observed from the 10 cycles.</li>
		<li>Polarize the sensors to +600 mV after the last cycle. The sensor current will decay to an asymptote. A steady reading is achieved after a minimum of 5 min. Record the zero reading.</li>
	</ol>
	</li>
	<li>In vivo sensor calibration
	<ol>
		<li>Do not perform the cyclic voltammetry on the&#160;in vivo sensors. Instead, polarize the sensor in the calibration chamber for 30 sec to +500 mV. Then set the potentiostat to 0 mV and allow the sensor current to rise to an asymptote. The sensor current will take at least 2 min to asymptote. Record the zero reading.</li>
	</ol>
	</li>
	<li>Consecutively add set amounts of ATP solution into the chamber to produce&#160;a calibration line encompassing a desired detection range. The ATP solution produces a sharp peak in the sensor signal initially, followed by a decay as the ATP diffuses evenly throughout the chamber. Record signal values once the signal level has stabilized after each ATP addition. Figures 6A and 7 show traces and suggested ATP concentrations for both ex vivo and in vivo studies, respectively. 
	NOTE: It is important to confirm the selectivity of the electrodes by using scavengers of the analytes. The present protocol used apyrase to test the specificity of the ATP sensor and catalase for the H2O2 signal (Figures 6B). If drugs are to be administered, measurements of their reactivity with the sensors should be determined before the study.</li>
	<li>Add 3 &#181;l of apyrase from a stock of 2 mg/ml (89 UN/mg) to test specificity of the ATP sensor (the current produced by ATP application should reduce to the zero level (Figure 7).</li>
	<li>Add 3 &#181;l of catalase from a stock of 2mg/ml (100 UN/mg) to test the specificity of the null sensor (the current produced by H2O2 application should reduce to the zero reading).</li>
</ol>
2. Animal Surgery For Sensor Studies
<ol>
	<li>Surgery for ex vivo
	<ol>
		<li>Anesthetize the experimental animal with isoflurane (5% induction, 1.5 to 2.5% maintenance)/medical grade O2 or another approved method. 15&#8217; 12 Animals must be continually monitored to ensure an adequate level of anesthesia. Stable respiratory rate and toe pinch reaction are used to confirm proper anesthesia. 
		Note: Euthanize the animal according to approved IACUC protocols. At the completion of all non-survival procedures euthanize deeply anaesthetized animals by thoracotomy inducing pneumothorax to ensure the humane demise of the animal12.</li>
		<li>Place the rat on a temperature-controlled surgical table in a supine position. While maintaining a proper anesthetic depth, make a midline incision of approximately 5 cm in line with the left kidney and expose the distal abdominal aorta.</li>
		<li>Wrap a ligature around the celiac and superior mesenteric arteries, and the abdominal aorta above these arteries but do not ligate. Wrap two ligatures around the abdominal aorta below the renal arteries.</li>
		<li>Clamp the abdominal aorta above the ligatures. Tie the lower ligature. Catheterize the abdominal aorta with polyethylene tubing (PE50). Secure the catheter with the second aorta ligature.</li>
		<li>Remove the clamp and ligate the mesenteric and celiac arteries. Perfuse the kidney at 6 ml/min with Hanks Balanced Salt Solution at RT for 2-3 min until the kidney is completely blanched.</li>
		<li>Excise the kidney and the catheter, connected portion of the aorta. Place the kidney into a 3 ml Petri dish filled with bath solution. 
		Note: Experiment protocol can be performed at RT. The bath solution contains in mM: 145 NaCl, 4.5 KCl, 2 MgCl2, 1 CaCl2, 10 HEPES, pH 7.35 adjusted with NaOH.</li>
	</ol>
	</li>
	<li>Surgery for in vivo
	<ol>
		<li>Anesthetize the rat using approved IACUC protocols. For in vivo analysis anesthetize the rats with ketamine (20 mg/kg i.m.) and inactin (50 mg/kg i.p.). Animals must be continually monitored to ensure an adequate level of anesthesia. A stable respiratory rate and toe pinch reaction are used to confirm proper anesthesia.</li>
		<li>After proper anesthesia depth is obtained, place the rat in a supine position on a temperature-controlled grounded surface located on the air table. The surface should be preheated and maintained at 36 &#176;C.</li>
		<li>While maintaining proper anesthetic depth, make a midline incision approximately 5 cm in line with the kidney.</li>
		<li>Use a suture to deflect and anchor the cutaneous and subcutaneous tissue so that the entire kidney is visible. Place the kidney in a kidney cup to minimize any movement artifacts.</li>
		<li>Use an IV infusion of 2% BSA: 0.9% NaCl at 1 ml/100 g/hr via the jugular vein to maintain blood volume. Cannulate both ureters for urine collection. Place ties around the superior mesenteric and celiac arteries and the distal aortal for the manipulation of renal perfusion pressure (Figures 10A).</li>
		<li>If the application of pharmacological agents is required during the in vivo experiments, insertion of an interstitial catheter is recommended (Figures 10B).</li>
	</ol>
	</li>
</ol>
3. Data Acquisition Setup
<ol>
	<li>Open the data acquisition program and set its polarity for both ex vivo and in vivo&#160;experiments to Anodic Positive. Set the program to save data as ASCII code.</li>
	<li>Position the micromanipulators for quick insertion of the sensors into the kidneys. 
	NOTE: Alternatively, use a dummy probe attached to the micromanipulator to help achieve the desired placement of sensors.</li>
	<li>Ex vivo data acquisition
	<ol>
		<li>Perfuse the kidney with bath solution (from 2.1.6) via the cannulated aorta at a constant rate of 650 &#181;l/min. Using surgical scissors, carefully remove the kidney capsule, which is necessary for sensor insertion.</li>
		<li>Secure the kidney with rubber bands strapped over the kidney and attached to the silicone-coated dish with pins.</li>
		<li>Place the reference electrode close to the kidney in the petri dish with its tip submerged in the buffer solution.</li>
	</ol>
	</li>
	<li>In vivo data acquisition
	<ol>
		<li>Place the kidney in a kidney cup. Depending on the strain and age of the animal use a size of cup that holds the kidney loosely. Figure 8 shows two sizes of kidney cups. Similar cups are used for different physiological approaches focused on the analysis of kidney function such as micropuncture etc16. 
		Note: The position of the kidney cup is critical to remove the mechanical noise produced by the animal breathing but should not interfere with or&#160;block kidney perfusion or urine flow.</li>
		<li>Allow 45 min&#160;of recovery time before performing data collection.</li>
		<li>Using a 26-30G needle, make a puncture hole at the desired location and depth of the sensor in the kidney. Blot the surface hole to remove&#160;exuded blood. Add glycerol solution to the surface of the kidney. This will prevent the kidney surface from drying out during the experiment.</li>
		<li>Remove the first sensor from the rehydration chamber and attach it to the micromanipulator. Quickly, within 20 sec, insert the electrode into the freshly created hole in the kidney. Repeat steps 3.5-3.9 for the null sensor.</li>
		<li>Insert the reference electrode into the kidney, approximately 1 cm from from the sensors.</li>
	</ol>
	</li>
	<li>Turn on the potentiostat and activate the recording program on the computer. Figure 9 shows the final setup of the ex vivo kidney data acquisition. Figure 10 shows the final setup of the in vivo kidney data acquisition with an inserted catheter.</li>
</ol>
4. Data Analysis
<ol>
	<li>Import the ASCII data file into Origin or any other similar software.</li>
	<li>Concentration current relation
	<ol>
		<li>Use the linear fit/extrapolate function to build a linear concentration (x-axis) to current (y-axis) relationship for the ATP or H2O2 calibration points (Figures 6 and 7). 
		linear fit line: y = mx + b</li>
	</ol>
	</li>
	<li>Equate the&#160;current to concentration</li>
	<li>Subtract the measured traces of the null sensor from those of the ATP sensor to obtain the actual current produced by intracellular ATP.</li>
	<li>Convert the ATP values in pA obtained by amperometry to nM using the calibration equation detailed in 4.2.1. Determine the concentration of the subtracted data trace of the ATP sensor by importing each adjusted current into the &#8220;x&#8221; value and solving for y (the concentration of analyte).</li>
	<li>Similarly, calculate H2O2 concentration from its calibration equation if needed.</li>
</ol>

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Sensors for the video recording of this manuscript were provided by Sarissa Biomedical Limited (Coventry, UK).

The present protocols were developed to provide enhanced temporal and spatial resolution of ATP and H2O2 signaling for ex vivo isolated, perfused and in vivo blood-perfused kidneys. The differences between the protocols and the sensors used here provide optimal data acquisition for either pharmacological agents or physiological studies. The protocols consist of 1) sensor calibration, 2) surgical procedure, 3) data acquisition setup, and 4) data analysis. They enable the real-time m...

Enzymatic microelectrode biosensors (also referenced as sensors in the present manuscript) have been a valuable tool for studying dynamic signaling processes in living cells and tissues. The sensors provide increased temporal and spatial resolution of cell signaling molecules in biologically relevant concentrations. Instead of sampling and analyzing extracellular fluids taken at intervals over long periods of time, these sensors respond as fast as their enzymes react to the analyte, thereby producing real-time measurements1,2. Fast detection of interstitial concentrations of autocrine and paracrine factors, like purines or hydrogen peroxide, and the dynamics of their release can be used to establish a profile for the effects of drugs in normal and pathological conditions 3. Currently, the majority of applications using sensors have been in brain tissue slices and cell cultures4-10. The protocols detailed in this manuscript aim to establish the means to accurately measure real-time concentrations of analytes in whole kidneys.
The following protocols were developed to study interstitial ATP and H2O2 signaling in kidneys. In the native environment of the kidney, extracellular ATP is rapidly catabolized by endogenous ectonucleotidases into its derivatives (ADP, AMP and adenosine). The sensors used here are highly selective to ATP over other purines or ATP degradation products11. This offers a great advantage as it allows accurate monitoring of the constant and dynamic concentrations of ATP release and its signaling function. Interstitial ATP concentration is measured using the combination of two microelectrodes, an ATP sensor and a Null sensor. The Null sensor in&#160;combination with&#160;catalase applications is able to detect interstitial H2O2 concentrations12. The following protocols use two different designs of sensors that have characteristics optimal for either ex vivo or in vivo applications.
Both designs are based on the sequential catalytic reaction of glycerol kinase and glycerol-3-phosphate oxidase contained in a sensor enzymatic layer and is&#160;driven by the presence of ATP. In the set of sensors used in the&#160;ex vivo studies, H2O2, the final enzymatic reaction product, is detected by&#160;oxidation on a platinum/iridium (Pt-Ir) wire electrode. Sensors for in vivo studies are instead based on H2O2 reduction on a mediator coated gold electrode designed for blood-perfused tissue. Shown in Figure 1 is a scheme of both protocols described in this manuscript. The Null sensor is identical to its corresponding ATP sensor except it lacks the bound enzymes. Therefore, in addition to the detection of H2O2 with the catalase enzyme, the Null sensor measures nonspecific interferences. ATP concentrations are calculated by subtracting the Null detected nonspecific interferences and background H2O2 from the ATP sensor signal. Several sensors are also commercially available to detect other analytes including adenosine, ionosine, hypoxanthine, acetylcholine, choline, glutamate, glucose, lactate, d-serine for ex vivo applications or adenosine, ionosine, and hypoxanthine for in vivo&#160;when paired with the corresponding Null sensor.
The ability&#160;of the sensor to accurately detect analytes depends on the proper pre- and post- calibrations13. This ensures that the analysis accounts for the drift in sensor sensitivity that occurs during use in biological tissues. The sensor holds a depot of glycerol that is used as a reagent in the sensor enzymatic reactions. If the sensor is not used in bath solutions containing glycerol, it will wash out over time. Shorter recording times are then necessary to minimize the sensor drift. Additionally sensor fouling by endogenous proteases and protein fragments can greatly diminish the sensitivity of the sensors14.
The present manuscript establishes the use of enzymatic microelectrode biosensors for ex vivo and in vivo kidney preparations. Real-time analyte quantification provides unprecedented detail of cellular signaling that may reveal novel insights into the mechanisms of kidney diseases and pharmacological agents.

<table><tbody><tr><td>Sensor Kit</td><td>Sarissa Biomedical</td><td>SBK-ATP-05-125</td><td>The kit includes storage bottle, rehydration chamber, electrode leads, and reference electrodes.&amp;nbsp; Also included with the kit is the user&#039;s choice of sensors.</td></tr><tr><td>Sarissaprobe ATP Biosensor 125 &amp;mu;m</td><td>Sarissa Biomedical</td><td>SBS-ATP-05-125</td><td>store at 2-8 &lt;sup&gt;o&lt;/sup&gt;C before use</td></tr><tr><td>Sarissagold ATP Biosensor 50 &amp;mu;m</td><td>Sarissa Biomedical</td><td>SGS-ATP-10-50</td><td>store at 2-8 &lt;sup&gt;o&lt;/sup&gt;C before use</td></tr><tr><td>Sarissaprobe null sensor 125 &amp;mu;m</td><td>Sarissa Biomedical</td><td>SBS-NUL-20-125</td><td>store at 2-8 &lt;sup&gt;o&lt;/sup&gt;C before use</td></tr><tr><td>Sarissagold null sensor 50 &amp;mu;m</td><td>Sarissa Biomedical</td><td>SGS-NUL-10-50</td><td>store at 2-8 &lt;sup&gt;o&lt;/sup&gt;C before use</td></tr><tr><td>Sarissaprobe ATP Manual</td><td>Sarissa Biomedical</td><td></td><td>&lt;a href=&quot;http://www.sarissa-biomedical.com/media/31563/instructions-atp.pdf&quot;&gt;http://www.sarissa-biomedical.com/media/31563/instructions-atp.pdf&lt;/a&gt;</td></tr><tr><td>Faraday cage&amp;nbsp;</td><td>TMC</td><td></td><td></td></tr><tr><td>Dual channel potentiostat</td><td>Digi-Ivy</td><td>DY2021</td><td>Type II Faraday cage</td></tr><tr><td>Data acquisition program</td><td>Digi-Ivy</td><td>DY2000</td><td></td></tr><tr><td>Perfusion pump</td><td>Razel Scientific Instruments</td><td>Model R99E</td><td></td></tr><tr><td>Fiber optic illuminator</td><td>Schott</td><td>ACE 1</td><td></td></tr><tr><td>micromanipulator</td><td>Narishige</td><td>MM-3</td><td></td></tr><tr><td>micromanipulator magnetic stand</td><td>Narishige</td><td>GJ-8</td><td></td></tr><tr><td>air table</td><td>TMC</td><td>63-500</td><td></td></tr><tr><td>isoflurane ventilator</td><td>LEI Medical</td><td>M2000</td><td></td></tr><tr><td>3 ml petri dish</td><td>Fisher Scientific</td><td>S3358OA</td><td></td></tr><tr><td>needle</td><td>Santa Cruz</td><td></td><td>26-30 G</td></tr><tr><td>pins</td><td></td><td></td><td>Standard dissection pins</td></tr><tr><td>catheter</td><td></td><td></td><td>Polyethylene tubing (PE50)</td></tr><tr><td>catheter tissue glue</td><td>Vetbond</td><td>1469SB</td><td></td></tr><tr><td>suture</td><td>Look</td><td>SP117</td><td></td></tr><tr><td>rubber bands</td><td></td><td></td><td>any 2-4 mm wide rubber bands</td></tr><tr><td>silicone</td><td>Momentive</td><td>RTV-615 Clear 1#</td><td></td></tr><tr><td>clamp</td><td>Fine Science Tools</td><td>18052-03</td><td></td></tr><tr><td>standard dissection kit</td><td></td><td></td><td>Kit should include scalpel and dissection sissors&amp;nbsp;</td></tr><tr><td>Kidney Cup</td><td></td><td></td><td>Of own design</td></tr><tr><td>standard chemicals</td><td>Sigma-Aldrich</td><td></td><td></td></tr><tr><td>ATP</td><td>Sigma-Aldrich</td><td>A6559-25UMO</td><td>100 mM ATP solution</td></tr><tr><td>hydrogen peroxide</td><td>Sigma-Aldrich</td><td>216763</td><td></td></tr><tr><td>glycerol</td><td>Sigma-Aldrich</td><td>G9012</td><td></td></tr><tr><td>Apyrase</td><td>Sigma-Aldrich</td><td>A7646</td><td></td></tr><tr><td>Catalase</td><td>Sigma-Aldrich</td><td>C40</td><td></td></tr><tr><td>isoflurane</td><td>Clipper</td><td>10250</td><td></td></tr><tr><td>inactin</td><td>Sigma-Aldrich</td><td>T133</td><td></td></tr><tr><td>ketamine</td><td>Clipper</td><td>2010012</td><td></td></tr><tr><td>Hanks Balanced Salt Solution&amp;nbsp;</td><td>Gibco</td><td>14025092</td><td></td></tr></tbody></table>

use of enzymatic biosensors to quantify endogenous atp or h2o2 in the kidney

Enzymatic microelectrode biosensors have been widely used to measure extracellular signaling in real-time. Most of their use has been limited to brain slices and neuronal cell cultures. Recently, this technology has been applied to the whole organs. Advances in sensor design have made &#160;possible the measuring of&#160;cell signaling in blood-perfused in vivo kidneys. The present protocols list the steps needed to measure ATP and H2O2 signaling in the rat kidney interstitium. Two separate sensor designs are used for the ex vivo and in vivo protocols. Both types of sensor&#160;are coated with&#160;a thin enzymatic biolayer on top of a permselectivity layer to give fast responding, sensitive and selective biosensors. The permselectivity layer protects the signal from the interferents in biological tissue, and the enzymatic layer utilizes&#160;the sequential catalytic reaction of glycerol kinase and glycerol-3-phosphate oxidase in the presence of ATP to produce H2O2. The set of sensors used for the ex vivo studies further detected analyte by oxidation of H2O2 on a platinum/iridium (Pt-Ir) wire electrode. The sensors for the&#160;in vivo studies are instead based on the reduction of H2O2 on a mediator coated gold electrode designed for blood-perfused tissue. Final concentration changes&#160;are detected by real-time amperometry followed by calibration to known concentrations of analyte. Additionally, the specificity of the amperometric signal can be confirmed by the addition of enzymes such as catalase and apyrase that break down H2O2 and ATP correspondingly. These sensors also rely heavily on accurate calibrations before and after each experiment. The following two protocols establish the study of real-time detection of ATP and H2O2 in kidney tissues, and can be further modified to extend the described method for use in other biological preparations or whole organs.

The design of the enzymatic microelectrode biosensor allows the real-time detection of analytes in whole kidneys. The general experiment design for either ex vivo or in vivo studies is illustrated in Figure 1.The sensors used and the surgical procedures differ depending on whether the study is ex vivo or in vivo.
To obtain reproducible results, accurate pre- and post- calibrations are critical. Figure 6A shows a rep...

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Use of Enzymatic Biosensors to Quantify Endogenous ATP or H2O2 in the Kidney

Enzymatic microelectrode biosensors enable real-time measurements of extracellular cell signaling in biologically-relevant concentrations. The following protocols extend the applications of biosensors to the ex vivo and in vivo detection of ATP and H2O2 in the kidney.

Use of Enzymatic Biosensors to Quantify Endogenous ATP or H2O2 in the Kidney

Enzymatic microelectrode biosensors have been widely used to measure extracellular signaling in real-time. Most of their use has been limited to brain ...

use-enzymatic-biosensors-to-quantify-endogenous-atp-or-h2o2

Universidad de Cartagena UDEC

Research

JoVE Journal

Biology

11.8K Views.  Medical College of Wisconsin. Enzymatic microelectrode biosensors enable real-time measurements of extracellular cell signaling in biologically-relevant concentrations. The following protocols extend the applications of biosensors to the ex vivo and in vivo detection of ATP and H2O2 in the kidney. 

Video: Use of Enzymatic Biosensors to Quantify Endogenous ATP or H2O2 in the Kidney

Ratiometric Biosensors that Measure Mitochondrial Redox State and ATP in Living Yeast Cells

Mitochondria have roles in many cellular processes, from energy metabolism and calcium homeostasis to control of cellular lifespan and programmed cell death. These processes affect and are affected by the redox status of and ATP production by mitochondria. Here, we describe the use of two ratiometric, genetically encoded biosensors that can detect mitochondrial redox state and ATP levels at subcellular resolution in living yeast cells. Mitochondrial redox state is measured using redox-sensitive Green Fluorescent Protein (roGFP) that is targeted to the mitochondrial matrix. Mito-roGFP contains cysteines at positions 147 and 204 of GFP, which undergo reversible and environment-dependent oxidation and reduction, which in turn alter the excitation spectrum of the protein. MitGO-ATeam is a F&ouml;rster resonance energy transfer (FRET) probe in which the &epsilon; subunit of the FoF1-ATP synthase is sandwiched between FRET donor and acceptor fluorescent proteins. Binding of ATP to the &epsilon; subunit results in conformation changes in the protein that bring the FRET donor and acceptor in close proximity and allow for fluorescence resonance energy transfer from the donor to acceptor.

We describe the use of two ratiometric, genetically encoded biosensors, which are based on GFP, to monitor mitochondrial redox state and ATP levels at subcellular resolution in living yeast cells.

Mitochondria have roles in many cellular processes, from energy metabolism and calcium homeostasis to control of cellular lifespan and programmed cell ...

Bioengineering

A Semi-High-Throughput Adaptation of the NADH-Coupled ATPase Assay for Screening Small Molecule Inhibitors

ATPase enzymes utilize the free energy stored in adenosine triphosphate to catalyze a wide variety of endergonic biochemical processes in vivo that would not occur spontaneously. These proteins are crucial for essentially all aspects of cellular life, including metabolism, cell division, responses to environmental changes and movement. The protocol presented here describes a nicotinamide adenine dinucleotide (NADH)-coupled ATPase assay that has been adapted to semi-high throughput screening of small molecule ATPase inhibitors. The assay has been applied to cardiac and skeletal muscle myosin II's, two actin-based molecular motor ATPases, as a proof of principle. The hydrolysis of ATP is coupled to the oxidation of NADH by enzymatic reactions in the assay. First, the ADP generated by the ATPase is regenerated to ATP by pyruvate kinase (PK). PK catalyzes the transition of phosphoenolpyruvate (PEP) to pyruvate in parallel. Subsequently, pyruvate is reduced to lactate by lactate dehydrogenase (LDH), which catalyzes the oxidation of NADH in parallel. Thus, the decrease in ATP concentration is directly correlated to the decrease in NADH concentration, which is followed by change to the intrinsic fluorescence of NADH. As long as PEP is available in the reaction system, the ADP concentration remains very low, avoiding inhibition of the ATPase enzyme by its own product. Moreover, the ATP concentration remains nearly constant, yielding linear time courses. The fluorescence is monitored continuously, which allows for easy estimation of the quality of data and helps to filter out potential artifacts (e.g., arising from compound precipitation or thermal changes).

A nicotinamide adenine dinucleotide (NADH)-coupled ATPase assay has been adapted to semihigh throughput screening of small molecule myosin inhibitors. This kinetic assay is run in a 384-well microplate format with total reaction volumes of only 20 &#181;L per well. The platform should be applicable to virtually any ADP producing enzyme.

ATPase enzymes utilize the free energy stored in adenosine triphosphate to catalyze a wide variety of endergonic biochemical processes in vivo that would ...

Biochemistry

Nanosensors to Detect Protease Activity In Vivo for Noninvasive Diagnostics

Proteases are multi-functional enzymes that specialize in the hydrolysis of peptide-bonds and control broad biological processes including homeostasis and allostasis. Moreover, dysregulated protease activity drives pathogenesis and is a functional biomarker of diseases such as cancer; therefore, the ability to detect protease activity in vivo may provide clinically relevant information for biomedical diagnostics. The goal of this protocol is to create nanosensors that probe for protease activity in vivo by producing a quantifiable signal in urine. These protease nanosensors consist of two components: a nanoparticle and substrate. The nanoparticle functions to increase circulation half-life and substrate delivery to target disease sites. The substrate is a short peptide sequence (6-8 AA), which is designed to be specific to a target protease or group of proteases. The substrate is conjugated to the surface of the nanoparticle and is terminated by a reporter, such as a fluorescent marker, for detection. As dysregulated proteases cleave the peptide substrate, the reporter is filtered into urine for quantification as a biomarker of protease activity. Herein we describe construction of a nanosensor for matrix metalloproteinase 9 (MMP9), which is associated with tumor progression and metastasis, for detection of colorectal cancer in a mouse model.

Nanosensors to Detect Protease Activity In Vivo for Noninvasive Diagnostics

Proteases are tightly regulated enzymes involved in fundamental biological processes, and dysregulated protease activity drives progression of complex diseases such as cancer. This method&#39;s goal is to create nanosensors that measure protease activity in vivo by producing a cleavage signal that is detectable from host urine and discriminates disease.

Proteases are multi-functional enzymes that specialize in the hydrolysis of peptide-bonds and control broad biological processes including homeostasis and ...

Assessment of Mitochondrial Functions and Cell Viability in Renal Cells Overexpressing Protein Kinase C Isozymes

The protein kinase C (PKC) family of isozymes is involved in numerous physiological and pathological processes. Our recent data demonstrate that PKC regulates mitochondrial function and cellular energy status. Numerous reports demonstrated that the activation of PKC-a and PKC-&epsilon; improves mitochondrial function in the ischemic heart and mediates cardioprotection. In contrast, we have demonstrated that PKC-&alpha; and PKC-&epsilon; are involved in nephrotoxicant-induced mitochondrial dysfunction and cell death in kidney cells. Therefore, the goal of this study was to develop an in vitro model of renal cells maintaining active mitochondrial functions in which PKC isozymes could be selectively activated or inhibited to determine their role in regulation of oxidative phosphorylation and cell survival. Primary cultures of renal proximal tubular cells (RPTC) were cultured in improved conditions resulting in mitochondrial respiration and activity of mitochondrial enzymes similar to those in RPTC in vivo. Because traditional transfection techniques (Lipofectamine, electroporation) are inefficient in primary cultures and have adverse effects on mitochondrial function, PKC-&epsilon; mutant cDNAs were delivered to RPTC through adenoviral vectors. This approach results in transfection of over 90% cultured RPTC.
Here, we present methods for assessing the role of PKC-&epsilon; in: 1. regulation of mitochondrial morphology and functions associated with ATP synthesis, and 2. survival of RPTC in primary culture. PKC-&epsilon; is activated by overexpressing the constitutively active PKC-&epsilon; mutant. PKC-&epsilon; is inhibited by overexpressing the inactive mutant of PKC-&epsilon;. Mitochondrial function is assessed by examining respiration, integrity of the respiratory chain, activities of respiratory complexes and F0F1-ATPase, ATP production rate, and ATP content. Respiration is assessed in digitonin-permeabilized RPTC as state 3 (maximum respiration in the presence of excess substrates and ADP) and uncoupled respirations. Integrity of the respiratory chain is assessed by measuring activities of all four complexes of the respiratory chain in isolated mitochondria. Capacity of oxidative phosphorylation is evaluated by measuring the mitochondrial membrane potential, ATP production rate, and activity of F0F1-ATPase. Energy status of RPTC is assessed by determining the intracellular ATP content. Mitochondrial morphology in live cells is visualized using MitoTracker Red 580, a fluorescent dye that specifically accumulates in mitochondria, and live monolayers are examined under a fluorescent microscope. RPTC viability is assessed using annexin V/propidium iodide staining followed by flow cytometry to determine apoptosis and oncosis.
These methods allow for a selective activation/inhibition of individual PKC isozymes to assess their role in cellular functions in a variety of physiological and pathological conditions that can be reproduced in in vitro.

The effects of activation of protein kinase C (PKC) isozymes on mitochondrial functions associated with respiration and oxidative phosphorylation and on cell viability are described. The approach adapts adenoviral technique to selectively overexpress PKC isozymes in primary cell culture and a variety of assays to determine mitochondrial functions and energy status of the cell.

The protein kinase C (PKC) family of isozymes is involved in numerous physiological and pathological processes. Our recent data demonstrate that PKC ...

Measuring Cation Transport by Na,K- and H,K-ATPase in Xenopus Oocytes by Atomic Absorption Spectrophotometry: An Alternative to Radioisotope Assays

Whereas cation transport by the electrogenic membrane transporter Na+,K+-ATPase can be measured by electrophysiology, the electroneutrally operating gastric H+,K+-ATPase is more difficult to investigate. Many transport assays utilize radioisotopes to achieve a sufficient signal-to-noise ratio, however, the necessary security measures impose severe restrictions regarding human exposure or assay design. Furthermore, ion transport across cell membranes is critically influenced by the membrane potential, which is not straightforwardly controlled in cell culture or in proteoliposome preparations. Here, we make use of the outstanding sensitivity of atomic absorption spectrophotometry (AAS) towards trace amounts of chemical elements to measure Rb+ or Li+ transport by Na+,K+- or gastric H+,K+-ATPase in single cells. Using Xenopus oocytes as expression system, we determine the amount of Rb+ (Li+) transported into the cells by measuring samples of single-oocyte homogenates in an AAS device equipped with a transversely heated graphite atomizer (THGA) furnace, which is loaded from an autosampler. Since the background of unspecific Rb+ uptake into control oocytes or during application of ATPase-specific inhibitors is very small, it is possible to implement complex kinetic assay schemes involving a large number of experimental conditions simultaneously, or to compare the transport capacity and kinetics of site-specifically mutated transporters with high precision. Furthermore, since cation uptake is determined on single cells, the flux experiments can be carried out in combination with two-electrode voltage-clamping (TEVC) to achieve accurate control of the membrane potential and current. This allowed e.g. to quantitatively determine the 3Na+/2K+ transport stoichiometry of the Na+,K+-ATPase and enabled for the first time to investigate the voltage dependence of cation transport by the electroneutrally operating gastric H+,K+-ATPase. In principle, the assay is not limited to K+-transporting membrane proteins, but it may work equally well to address the activity of heavy or transition metal transporters, or uptake of chemical elements by endocytotic processes.

Measuring Cation Transport by Na,K- and H,K-ATPase in Xenopus Oocytes by Atomic Absorption Spectrophotometry: An Alternative to Radioisotope Assays

We describe a method to quantify the activity of K+-countertransporting P-type ATPases by heterologous expression of the enzymes in Xenopus oocytes and measuring Rb+ or Li+ uptake into individual cells by atomic absorption spectrophotometry. The method is a sensitive and safe alternative to radioisotope flux experiments facilitating complex kinetic studies.

Whereas cation transport by the electrogenic membrane transporter Na+,K+-ATPase can be measured by electrophysiology, the electroneutrally operating ...

Imaging of Intracellular ATP in Organotypic Tissue Slices of the Mouse Brain using the FRET-based Sensor ATeam1.03YEMK

Neuronal activity in the central nervous system (CNS) evokes a high demand on cellular energy provided by the breakdown of adenosine triphosphate (ATP). A large share of ATP is needed to re-install ion gradients across plasma membranes degraded by electrical signaling of neurons. There is evidence that astrocytes - while not generating fast electrical signals themselves - undergo increased production of ATP in response to neuronal activity and support active neurons by providing energy metabolites to them. The recent development of genetically encoded sensors for different metabolites now enables the study of such metabolic interactions between neurons and astrocytes. Here, we describe a protocol for cell-type specific expression of the ATP-sensitive Fluorescence Resonance Energy Transfer- (FRET-) sensor ATeam1.03YEMK in organotypic tissue slice cultures of the mouse hippocampus and cortex using adeno-associated viral vectors (AAV). Furthermore, we demonstrate how this sensor can be employed for dynamic measurement of changes in cellular ATP levels in neurons and astrocytes upon increases in extracellular potassium and following induction of chemical ischemia (i.e., an inhibition of cellular energy metabolism).

Imaging of Intracellular ATP in Organotypic Tissue Slices of the Mouse Brain using the FRET-based Sensor ATeam1.03YEMK

We describe a protocol for cell-type specific expression of the genetically encoded FRET-based sensor ATeam1.03YEMK in organotypic slice cultures of the mouse forebrain. Furthermore, we show how to use this sensor for dynamic imaging of cellular ATP levels in neurons and astrocytes.

Neuronal activity in the central nervous system (CNS) evokes a high demand on cellular energy provided by the breakdown of adenosine triphosphate (ATP). A ...

Neuroscience

Biochemical Assays for Analyzing Activities of ATP-dependent Chromatin Remodeling Enzymes

Members of the SNF2 family of ATPases often function as components of multi-subunit chromatin remodeling complexes that regulate nucleosome dynamics and DNA accessibility by catalyzing ATP-dependent nucleosome remodeling. Biochemically dissecting the contributions of individual subunits of such complexes to the multi-step ATP-dependent chromatin remodeling reaction requires the use of assays that monitor the production of reaction products and measure the formation of reaction intermediates. This JOVE protocol describes assays that allow one to measure the biochemical activities of chromatin remodeling complexes or subcomplexes containing various combinations of subunits. Chromatin remodeling is measured using an ATP-dependent nucleosome sliding assay, which monitors the movement of a nucleosome on a DNA molecule using an electrophoretic mobility shift assay (EMSA)-based method. Nucleosome binding activity is measured by monitoring the formation of remodeling complex-bound mononucleosomes using a similar EMSA-based method, and DNA- or nucleosome-dependent ATPase activity is assayed using thin layer chromatography (TLC) to measure the rate of conversion of ATP to ADP and phosphate in the presence of either DNA or nucleosomes. Using these assays, one can examine the functions of subunits of a chromatin remodeling complex by comparing the activities of the complete complex to those lacking one or more subunits. The human INO80 chromatin remodeling complex is used as an example; however, the methods described here can be adapted to the study of other chromatin remodeling complexes.

Here we describe biochemical assays that can be used to characterize ATP-dependent chromatin remodeling enzymes for their abilities to 1) catalyze ATP-dependent nucleosome sliding, 2) engage with nucleosome substrates, and 3) hydrolyze ATP in a nucleosome- or DNA-dependent manner.

Members of the SNF2 family of ATPases often function as components of multi-subunit chromatin remodeling complexes that regulate nucleosome dynamics and ...

A Simple Bioassay for the Evaluation of Vascular Endothelial Growth Factors

The analysis of receptor tyrosine kinases and their interacting ligands involved in vascular biology is often challenging due to the constitutive expression of families of related receptors, a broad range of related ligands and the difficulty of dealing with primary cultures of specialized endothelial cells. Here we describe a bioassay for the detection of ligands to the vascular endothelial growth factor receptor-2 (VEGFR-2), a key transducer of signals that promote angiogenesis and lymphangiogenesis. A cDNA encoding a fusion of the extracellular (ligand-binding) region of VEGFR-2 with the transmembrane and cytoplasmic regions of the erythropoietin receptor (EpoR) is expressed in the factor-dependent cell line Ba/F3. This cell line grows in the presence of interleukin-3 (IL-3) and withdrawal of this factor results in death of the cells within 24 hr. Expression of the VEGFR-2/EpoR receptor fusion provides an alternative mechanism to promote survival and potentially proliferation of stably transfected Ba/F3 cells in the presence of a ligand capable of binding and cross-linking the extracellular portion of the fusion protein (i.e., one that can cross-link the VEGFR-2 extracellular region). The assay can be performed in two ways: a semi-quantitative approach in which small volumes of ligand and cells permit a rapid result in 24 hr, and a quantitative approach involving surrogate markers of a viable cell number. The assay is relatively easy to perform, is highly responsive to known VEGFR-2 ligands and can accommodate extracellular inhibitors of VEGFR-2 signaling such as monoclonal antibodies to the receptor or ligands, and soluble ligand traps.

We describe a simple cell-based bioassay for detecting, quantifying and monitoring the activity of members of the vascular endothelial growth factor family of ligands. The assay uses chimeric receptors expressed in a factor-dependent cell line to provide a semi-quantitative or quantitative assessment of receptor binding and cross-linking by the ligand.

The analysis of receptor tyrosine kinases and their interacting ligands involved in vascular biology is often challenging due to the constitutive ...

Measuring In Vitro ATPase Activity for Enzymatic Characterization

Adenosine triphosphate-hydrolyzing enzymes, or ATPases, play a critical role in a diverse array of cellular functions. These dynamic proteins can generate energy for mechanical work, such as protein trafficking and degradation, solute transport, and cellular movements. The protocol described here is a basic assay for measuring the in vitro activity of purified ATPases for functional characterization. Proteins hydrolyze ATP in a reaction that results in inorganic phosphate release, and the amount of phosphate liberated is then quantitated using a colorimetric assay. This highly adaptable protocol can be adjusted to measure ATPase activity in kinetic or endpoint assays. A representative protocol is provided here based on the activity and requirements of EpsE, the AAA+ ATPase involved in Type II Secretion in the bacterium Vibrio cholerae. The amount of purified protein needed to measure activity, length of the assay and the timing and number of sampling intervals, buffer and salt composition, temperature, co-factors, stimulants (if any), etc. may vary from those described here, and thus some optimization may be necessary. This protocol provides a basic framework for characterizing ATPases and can be performed quickly and easily adjusted as necessary.

Measuring In Vitro ATPase Activity for Enzymatic Characterization

We describe a basic protocol for quantitating in vitro ATPase activity. This protocol can be optimized based on the level of activity and requirements for a given purified ATPase.

Adenosine triphosphate-hydrolyzing enzymes, or ATPases, play a critical role in a diverse array of cellular functions. These dynamic proteins can generate ...

Using Real-Time Cell Metabolic Flux Analyzer to Monitor Osteoblast Bioenergetics

Bone formation by osteoblasts is an essential process for proper bone acquisition and bone turnover to maintain skeletal homeostasis, and ultimately, prevent fracture. In the interest to both optimize peak bone mass and combat various musculoskeletal diseases (i.e., post-menopausal osteoporosis, anorexia nervosa, type 1 and 2 diabetes mellitus), incredible efforts have been made in the field of bone biology to fully characterize osteoblasts throughout their differentiation process. Given the primary role of mature osteoblasts to secrete matrix proteins and mineralization vesicles, it has been noted that these processes take an incredible amount of cellular energy, or adenosine triphosphate (ATP). The overall cellular energy status is often referred to as cellular bioenergetics, and it includes a series of metabolic reactions that sense substrate availability to derive ATP to meet cellular needs. Therefore, the current method details the process of isolating primary, murine bone marrow stromal cells (BMSCs) and monitoring their bioenergetic status using the Real-time cell metabolic flux analyzer at various stages in osteoblast differentiation. Importantly, these data have demonstrated that the metabolic profile changes dramatically throughout osteoblast differentiation. Thus, using this physiologically relevant cell type is required to fully appreciate how a cell's bioenergetic status can regulate the overall function.

Real-time cell metabolic flux assay measures the oxygen consumption rate and extracellular acidification rate, which corresponds to mitochondrial and glycolytic adenosine triphosphate production, using pH and oxygen sensors. The manuscript explains a method to understand the energy status of osteoblasts and the characterization and interpretation of the cellular bioenergetic status.

Bone formation by osteoblasts is an essential process for proper bone acquisition and bone turnover to maintain skeletal homeostasis, and ultimately, ...

Use of Enzymatic Biosensors to Quantify Endogenous ATP or H₂O₂ in the Kidney

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Rozdziały w tym wideo

Ratiometric Biosensors that Measure Mitochondrial Redox State and ATP in Living Yeast Cells

A Semi-High-Throughput Adaptation of the NADH-Coupled ATPase Assay for Screening Small Molecule Inhibitors

Nanosensors to Detect Protease Activity In Vivo for Noninvasive Diagnostics

Assessment of Mitochondrial Functions and Cell Viability in Renal Cells Overexpressing Protein Kinase C Isozymes

Measuring Cation Transport by Na,K- and H,K-ATPase in Xenopus Oocytes by Atomic Absorption Spectrophotometry: An Alternative to Radioisotope Assays

Imaging of Intracellular ATP in Organotypic Tissue Slices of the Mouse Brain using the FRET-based Sensor ATeam1.03^YEMK

Biochemical Assays for Analyzing Activities of ATP-dependent Chromatin Remodeling Enzymes

A Simple Bioassay for the Evaluation of Vascular Endothelial Growth Factors

Measuring In Vitro ATPase Activity for Enzymatic Characterization

Using Real-Time Cell Metabolic Flux Analyzer to Monitor Osteoblast Bioenergetics