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 measurements of analytes for numerous experimental conditions. This will lead to greater insights into kidney diseases and the development of more effective pharmacological treatments. Here we used the sensors to analyze ATP and H2O2. However, sensors for other substances are also available and can be used. The protocols described here were applied to study ATP and H2O2 in rat kidneys, but the same method can easily be adjusted for application in mice. Thus, this approach has a great potential considering the abundance of genetic rodent models.
The basic design of enzymatic microelectrode biosensors was developed in the 1960&#8217;s by Clark and Lyons2. Following the initial development of biosensors, advances have been made in both design17-19 and applications20. Llaudet et al.21,22 developed the principle design of the ATP sensors used in the present protocols while using methods from Cosneir et al.23 for the enzyme coating. These sensors have detected ATP signaling in a number of physiological processes including ATP release from brain astrocytes8, regulation of breathing4,24, and skeletal muscle arteriole regulation25. Recently the ex vivo protocol has been used to measure ATP signaling in kidneys12. The goal of this manuscript is to provide effective directions and insights for the detection of endogenous substances such as ATP in kidneys.
Enzymatic microelectrode biosensors have many advantages over existing means of measuring analytes in vivo. However, special precautions should be used for this approach as for any other new method. Validation with an established approach provides confidence in the obtained data. For instance, microdialysis measurements were made as described previously26,27. No significant difference was observed between the peak concentrations determined by the&#160;sensors and those&#160;obtained from the dialysis samples12. The limitation of microdialysis is that it only measures steady-state levels of analyte. As such, the assessment of dynamic changes in cell signaling can only be achieved with the use of the enzymatic microelectrode biosensors. The manufactory specifications of the sensor response times differ between sensor types. Ex vivo sensors have a response time of 5-10 sec and in vivo sensors of 30-35 sec for the signal rise from 10 to 90%. The calibration traces (Figures 6 and 7) demonstrate the time resolution of analyte addition/removal. For both the sensor and microdialysis approaches, the use of specific enzymes to block the signal produced by analytes is required to ensure specificity.
The described protocols have several challenges. As with any sensor insertion into tissue, tissue damage does occur. This could&#160;activate signaling pathways which may influence the measurements28. To minimize the cell damage, thinner sensors would be needed. However, it is difficult to penetrate the kidney capsule with the sensors so needles are used to make a small entry hole. Thin sensors are more likely to bend and disrupt their coating on insertion. Larger diameter sensors are more resistant bending and the resultant damage to their coating. Figure 3 shows a thin and thick sensor. In addition to the thick sensor&#8217;s resistance to bending, they contain a thicker layer of coating, providing increased protection for the enzymatic layer. The concern of tissue damage caused by using thick sensors was considerably less in kidneys than it would be in brain tissue. The insertion depth is approximated and final placement should&#160;be verified after measurements on the kidney are completed. Estimation between the cortex and medulla layers using the sensor tip length as an insertion guide and&#160;is sufficient as the sensor tip is 0.5 mm and the kidney cortex is 2-3 mm thick. Fouling of the sensor also occurs at a greater rate in blood, thus limiting the sensor use in long in vivo protocols to one experiment. Recording times of&#160;1 to 1 &#189; hr resulted in only a small sensor drift. The main criteria when assessing the reduced sensor performance is the low sensitivity to the analyte during calibration process. Increased electrical noise and instability of the baseline current can also indicate that the sensor tip is damaged. For accurate measurements of ATP, both sensors (ATP and Null) should have similar noise and stability characteristics for further subtraction of the signal. Otherwise, one of the sensors should be replaced. For low noise and detection of small concentrations of analyte, the use of new sensors for each animal is suggested.
Several steps are crucial for successful experimental outcomes. The sensors are very fragile and should be handled carefully to avoid damaging the sensor tip. Also, after rehydration, the sensors should not be exposed to the air for more than 20 sec. Low noise recordings are required for the successful detection of biological signals in tissues. For that purpose, a high-performance lab air table and proper electrical grounding is required. The addition of exact amounts of analyte during the calibration steps is necessary for accurate concentration determination in experiments. Achieving good clearing of the kidney in ex vivo kidney surgeries will result in successful recordings. The use of enzymes apyrase and catalase in calibrations and in experiments allows for the assessment of sensor nonspecific sensitivity and confirms the measurements.
These protocols provide greatly enhanced detail of&#160;extracellular signaling in kidneys. The improved sensitivity and temporal resolution afforded by the sensors may allow us to resolve changes in ATP signaling in disease states and following pharmacological manipulation that were previously undetectable.&#160;The in vivo protocol is well suited for complex physiological studies while ex vivo is optimal for the study of pharmacological applications on kidneys. The design of the in vivo sensor used here enables unprecedented resistance against interference in blood-perfused tissue. Taken together, these sensors offer a large amount of applications that were previously not possible with existing protocols and techniques.

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 border="0" cellpadding="0" cellspacing="0">
	<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.&#160; Also included with the kit is the user&#39;s choice of sensors.</td>
		</tr>
		<tr>
			<td>Sarissaprobe ATP Biosensor 125 &#956;m</td>
			<td>Sarissa Biomedical</td>
			<td>SBS-ATP-05-125</td>
			<td>store at 2-8 C before use</td>
		</tr>
		<tr>
			<td>Sarissagold ATP Biosensor 50 &#956;m</td>
			<td>Sarissa Biomedical</td>
			<td>SGS-ATP-10-50</td>
			<td>store at 2-8 C before use</td>
		</tr>
		<tr>
			<td>Sarissaprobe null sensor 125&#956;m</td>
			<td>Sarissa Biomedical</td>
			<td>SBS-NUL-20-125</td>
			<td>store at 2-8 C before use</td>
		</tr>
		<tr>
			<td>Sarissagold null sensor 50 &#956;m</td>
			<td>Sarissa Biomedical</td>
			<td>SGS-NUL-10-50</td>
			<td>store at 2-8 C before use</td>
		</tr>
		<tr>
			<td>Sarissaprobe ATP Manual</td>
			<td>Sarissa Biomedical</td>
			<td></td>
			<td><a href="http://www.sarissa-biomedical.com/media/31563/instructions-atp.pdf">http://www.sarissa-biomedical.com/media/31563/instructions-atp.pdf</a></td>
		</tr>
		<tr>
			<td>Faraday cage&#160;</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 gauge</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&#160;</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&#160;</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 representative trace of the signal seen from the ex vivo ATP sensor during calibrations. Note that apyrase quickly eliminated the ATP signal. Catalase had no effect on the ATP signal and demonstrates its specificity to the H2O2 it creates (Figure 6B). The calibration procedure produces a linear fit that is used to calculate the dynamic changes of ATP (Figure 6A, right panel). In vivo sensor calibration produces a similar trace to that of the ex vivo sensor. However, this sensor detects reductive instead of oxidative currents and as such the current produced is negative. Calibration of these sensors also produces a linear fit in a 0.3 to 80 &#181;M range (Figure 7A right panel). Specificity of in vivo sensor to ATP over other purine products is shown in Figure 7B.
The approach described here allows us to measure both basal levels of endogenous substances and acute changes in response to drug infusion12. Shown in Figure 11 are Ang II-induced changes of interstitial endogenous concentration of H2O2 in salt-resistant and salt-sensitive rats. For these experiments, freshly isolated kidneys from Sprague Dawley (SD) or Dahl salt-sensitive (SS) rats were perfused with 1 &#181;M Ang II under constant laminar flow (650 &#181;l/min). As shown in Figure 11, Ang II induces the acute release of H2O2 in kidneys from both SD and SS rats. However, the maximum amplitude of each response was significantly elevated in SS rats, especially when the animals were fed with a high salt diet12. Shown in Figure 12 is a representative application of the biosensors in vivo. The infusion of catalase (5 &#181;g/ml) completely blocks the H2O2 signal detected by the biosensor. These experiments illustrate the use of specific enzymatic biosensors combined with the amperometric technique for the detection of basal levels of endogenous substances and real-time measurements of their release in response to pharmacological intervention. Further applications of this approach to study the role of purinergic signaling and hydrogen peroxide will improve&#160;our knowledge and&#160;understanding of renal pathologies like salt-sensitive hypertension, renal oxidative stress and chronic kidney disease.
<img alt="Figure 1" src="/files/ftp_upload/53059/53059fig1.jpg" /> 
Figure&#160;1. Schematic of the Protocols. Calibration with analyte of interest and testing its specificity is done immediately&#160;prior to the start of the experiments. The in vivo protocol should be followed for complex physiological experiments and ex vivo for pharmacological applications. Post experiment calibration should be done and taken into account during the data analysis. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/53059/53059fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" src="/files/ftp_upload/53059/53059fig2.jpg" /> 
Figure&#160;2.&#160;Enzymatic&#160;Microelectrode&#160;Sensor. The two sensor sizes used for ex vivo applications are shown, 50 &#181;m and 125 &#181;m diameters. Each sensor wire extends into a capillary tube to connect to a gold end connector (not shown). Inset shows a&#160;sensor tip coated with&#160;enzymes for ATP detection. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/53059/53059fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" src="/files/ftp_upload/53059/53059fig3.jpg" /> 
Figure&#160;3. Dual&#160;Channel&#160;Potentiostat. The potentiostat channels labeled CH1 and CH2. 1) CH1 the ATP sensor connection and 2) CH2 the Null sensor connection 3) connections that are electrically coupled to the reference electrode. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/53059/53059fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" src="/files/ftp_upload/53059/53059fig4.jpg" /> 
Figure&#160;4. Sensor&#160;Setup. The data acquisition is performed in a Faraday cage to reduce electrical noise and on a high-performance lab table for a&#160;vibration-free working surface. 1) A temperature-controlled surgical table is used to maintain&#160;&#160;animal body temperature during physiological experiments 2) &#160;the micromanipulators are attached to magnetic mounting adaptors for flexible positioning of the sensors during experiment 3) A light source is needed for surgical procedures and sensor insertion 4) dual channel potentiostat 5) holder for the sensors. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/53059/53059fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" src="/files/ftp_upload/53059/53059fig5.jpg" /> 
Figure&#160;5. Cyclic&#160;Voltammetry. For ex vivo studies, sensor cyclic voltammetry is conducted for 10 cycles between -0.5 and 0.5 V prior to the calibration protocol. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/53059/53059fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" src="/files/ftp_upload/53059/53059fig6.jpg" /> 
Figure&#160;6. Amperometry Calibration&#160;for&#160;Ex&#160;Vivo.&#160;(A) Calibration uses&#160;the addition of known ATP concentrations to the bath solution. Corresponding amperometric values recorded at the asymptote level (black trace). The currents obtained create a calibration equation. An example is shown on the right panel. The red trace is the current of the Null sensor which&#160;responds to only the addition of H2O2. (B) The Null sensor is calibrated with the addition of known H2O2 concentrations (red arrows) to the bath solution and the asymptotes amperometric values are recorded (thin black arrows). Addition of catalase to the bath solution (thick black arrow) results in rapid current decay. The right panel shows the Null sensor calibration equation. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/53059/53059fig6large.jpg" target="_blank">Please click here to view a larger version of this figure.</a> <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/53059/53059fig6large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 7" src="/files/ftp_upload/53059/53059fig7.jpg" /> 
Figure&#160;7. Amperometry&#160;Calibration&#160;for&#160;In&#160;Vivo. Calibration of the&#160;in vivo electrodes is performed in a similar fashion to that&#160;detailed in the ex vivo studies except reduction reactions cause a reverse current (polarity). (A) The addition of known ATP concentrations produces an amperometric current on the ATP sensor (black trace) but has no effect on the Null sensor (red trace). Addition of apyrase extinguishes the current of the ATP sensor. (B) The specificity of the ATP sensor is confirmed by the addition of 10 &#181;M of different purinergic agents (UTP, UDP and adenosine). Further applications of ATP provide a stable detectable amperometric current. (C) Microphotograph of the in vivo sensors based on a mediator coated gold electrode. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/53059/53059fig7large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 8" src="/files/ftp_upload/53059/53059fig8.jpg" /> 
Figure&#160;8. Kidney&#160;Cups. In the in vivo&#160;studies, kidneys are held still using the stainless steel kidney cups shown. Two sizes of cups are used to accommodate kidney size variation. These cups reduce the movement artifacts which are generated by the animal&#8217;s respiration. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/53059/53059fig8large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 9" src="/files/ftp_upload/53059/53059fig9.jpg" /> 
Figure&#160;9. Ex&#160;Vivo&#160;Isolated&#160;and&#160;Perfused&#160;Kidney. The isolated kidney is placed into a 1) Petri dish coated with a thick silicone bottom for pin insertion&#160;2) the renal artery is cannulated and attached to a syringe pump for constant perfusion during experiment 3) the ATP sensor, 4) and Null sensor are inserted into the kidney 5) the reference electrode is placed near the kidney submerged into the bath solution. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/53059/53059fig9large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 10" src="/files/ftp_upload/53059/53059fig10.jpg" /> 
Figure&#160;10. In&#160;Vivo&#160;Blood&#160;Perfused&#160;Kidney.&#160;(A) The left kidney is exposed and placed in a kidney cup, the right kidney remains intact inside the animal. Both sensors are inserted into the kidney. BSA:NaCl is infused via the catheterized jugular vein to counteract the fluid loss caused by the large mid-line incision (B) Example of the in vivo experiment with an implanted interstitial catheter for direct pharmacological applications 1) the kidney is held by a kidney cup 2) ATP sensor 3) Null sensor and 4) reference electrode are inserted into the ventral surface of the kidney 5) catheter implanted into the kidney and attached to a peristaltic pump for laminar pharmacological infusions. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/53059/53059fig10large.jpg" target="_blank">Please click here to view a larger version of this figure.</a> <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/53059/53059fig10large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 11" src="/files/ftp_upload/53059/53059fig11.jpg" /> 
Figure&#160;11.&#160;Ex&#160;Vivo&#160;Analysis&#160;of&#160;H2O2&#160;in the kidney. Ang II perfusion causes H2O2 release in the rat kidney cortex. (A) Real time changes of the mean H2O2 concentration&#160;(gray bars show standard error) from a total of N = 8 applications (4 kidneys from 4 different rats). Bars on the top represent Ang II application. (B) The maximal H2O2 concentration amplitude values during Ang II perfusion for Sprague Dawley (SD) and Dahl salt-sensitive (SS) rats fed a low and high salt diets, respectively. * - P &#60; 0.05 versus SD rats. The figure is adapted from reference12 with permission. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/53059/53059fig11large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 12" src="/files/ftp_upload/53059/53059fig12.jpg" /> 
Figure&#160;12. In&#160;Vivo&#160;Analysis of&#160;H2O2. Example of the&#160;in vivo assessment of interstitial H2O2 concentration in the medulla of a SS rat fed a low salt diet (as shown in Figure 10B). The interstitial application of catalase via an implanted catheter for the 5 min interval produced a complete blockade of the H2O2 signal in the renal medulla. Reduction of catalase, from washout&#160;by renal blood flow, resulted in a partial recovery of the signal, which was blocked again by an additional catalase application (10 min). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/53059/53059fig12large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Use of Enzymatic Biosensors to Quantify Endogenous ATP or H2O2 in the Kidney at JoVE.com

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

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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

Use of the Protease Fluorescent Detection Kit to Determine Protease Activity

The Protease Fluorescent Detection Kit provides ready-to-use reagents for detecting the presence of protease activity. This simple assay to detect protease activity uses casein labeled with fluorescein isothiocyanate (FITC) as the substrate.
Protease activity results in the cleavage of the FITC-labeled casein substrate into smaller fragments, which do not precipitate under acidic conditions. After incubation of the protease sample and substrate, the reaction is acidified with the addition of trichloroacetic acid (TCA). The mixture is then centrifuged with the undigested substrate forming a pellet and the smaller, acid soluble fragments remaining in solution. The supernatant is neutralized and the fluorescence of the FITC-labeled fragments is measured.
The described kit procedure detects the trypsin protease control at a concentration of approximately 0.5 &mu;g/ml (5 ng of trypsin added to the assay). This sensitivity can be increased with a longer incubation time, up to 24 hours. The assay is performed in microcentrifuge tubes and procedures are provided for fluorescence detection using either cuvettes or multiwell plates.

The Protease Fluorescent Detection Kit is designed for the measurement of protease activity using fluorometry. It is also suitable for detection of trace amounts of protease contamination. The method is based on the proteolytic hydroysis of a proprietary formulation of a FITC-labeled casein substrate.

The Protease Fluorescent Detection Kit provides ready-to-use reagents for detecting the presence of protease activity. This simple assay to detect ...

The use of SC1 (Pluripotin) to Support mESC Self-renewal in the Absence of LIF

Mouse embryonic stem (ES) cells are conventionally cultured with Leukemia Inhibitory Factor (LIF) to maintain self-renewal.1 However, LIF is expensive and activation of the LIF/JAK/STAT3 pathway is not absolutely required to maintain the self-renewal state.2 The SC1 small molecule may be an economical alternative to LIF. SC1 functions through dual inhibition of Ras-GAP and ERK1.3 Illustration of its mechanism of action makes it a useful tool to study the fundamental molecular mechanism of self-renewal. Here we demonstrate the procedure for culturing mouse ES cells in the presence of SC1 and show that they are able to maintain self-renewal in the absence of LIF. Cells cultured with SC1 showed similar morphology compared to cells maintained with LIF. Both exhibited typical mouse ES morphology after five passages. Expression of typical pluripotency markers (Oct4, Sox2, Nanog, and SSEA1) was observed after five passages in the presence of SC1. Furthermore, SC1 caused no overt toxicity on mouse ES cells.

The use of SC1 Pluripotin to Support mESC Self-renewal in the Absence of LIF

SC1 functions through dual inhibition of Ras- GAP and ERK1. We tested the function of SC1 in supporting mouse ES cell self-renewal in the absence of LIF and showed that SC1 is able to maintain self-renewal of mouse ES cell cultures.

Mouse embryonic stem (ES) cells are conventionally cultured with Leukemia Inhibitory Factor (LIF) to maintain self-renewal.1 However, LIF is expensive and ...

Dissection of the Adult Zebrafish Kidney

Researchers working in the burgeoning field of adult stem cell biology seek to understand the signals that regulate the behavior and function of stem cells during normal homeostasis and disease states. The understanding of adult stem cells has broad reaching implications for the future of regenerative medicine1. For example, better knowledge about adult stem cell biology can facilitate the design of therapeutic strategies in which organs are triggered to heal themselves or even the creation of methods for growing organs in vitro that can be transplanted into humans1. The zebrafish has become a powerful animal model for the study of vertebrate cell biology2. There has been extensive documentation and analysis of embryonic development in the zebrafish3. Only recently have scientists sought to document adult anatomy and surgical dissection techniques4, as there has been a progressive movement within the zebrafish community to broaden the applications of this research organism to adult studies. For example, there are expanding interests in using zebrafish to investigate the biology of adult stem cell populations and make sophisticated adult models of diseases such as cancer5. Historically, isolation of the zebrafish adult kidney has been instrumental for studying hematopoiesis, as the kidney is the anatomical location of blood cell production in fish6,7. The kidney is composed of nephron functional units found in arborized arrangements, surrounded by hematopoietic tissue that is dispersed throughout the intervening spaces. The hematopoietic component consists of hematopoietic stem cells (HSCs) and their progeny that inhabit the kidney until they terminally differentiate8. In addition, it is now appreciated that a group of renal stem/progenitor cells (RPCs) also inhabit the zebrafish kidney organ and enable both kidney regeneration and growth, as observed in other fish species9-11. In light of this new discovery, the zebrafish kidney is one organ that houses the location of two exciting opportunities for adult stem cell biology studies. It is clear that many outstanding questions could be well served with this experimental system. To encourage expansion of this field, it is beneficial to document detailed methods of visualizing and then isolating the adult zebrafish kidney organ. This protocol details our procedure for dissection of the adult kidney from both unfixed and fixed animals.
Dissection of the kidney organ can be used to isolate and characterize hematopoietic and renal stem cells and their offspring using established techniques such as histology, fluorescence activated cell sorting (FACS)11,12, expression profiling13,14, and transplantation11,15. 
We hope that dissemination of this protocol will provide researchers with the knowledge to implement broader use of zebrafish studies that ultimately can be translated for human application.

The zebrafish kidney is home to both renal and hematopoietic adult stem/progenitor cells, and represents an outstanding opportunity to study these cell types and their progeny in a vertebrate model organism. Here, we demonstrate a detailed dissection procedure that enables the researcher to identify and surgically remove the adult zebrafish kidney, which can be used for applications such as cell isolation, transplantation, and expression studies of kidney and/or blood cell populations.

Researchers working in the burgeoning field of adult stem cell biology seek to understand the signals that regulate the behavior and function of stem ...

FRET Microscopy for Real-time Monitoring of Signaling Events in Live Cells Using Unimolecular Biosensors

F&ouml;rster resonance energy transfer (FRET) microscopy continues to gain increasing interest as a technique for real-time monitoring of biochemical and signaling events in live cells and tissues. Compared to classical biochemical methods, this novel technology is characterized by high temporal and spatial resolution. FRET experiments use various genetically-encoded biosensors which can be expressed and imaged over time in situ or in vivo1-2. Typical biosensors can either report protein-protein interactions by measuring FRET between a fluorophore-tagged pair of proteins or conformational changes in a single protein which harbors donor and acceptor fluorophores interconnected with a binding moiety for a molecule of interest3-4. Bimolecular biosensors for protein-protein interactions include, for example, constructs designed to monitor G-protein activation in cells5, while the unimolecular sensors measuring conformational changes are widely used to image second messengers such as calcium6, cAMP7-8, inositol phosphates9 and cGMP10-11. Here we describe how to build a customized epifluorescence FRET imaging system from single commercially available components and how to control the whole setup using the Micro-Manager freeware. This simple but powerful instrument is designed for routine or more sophisticated FRET measurements in live cells. Acquired images are processed using self-written plug-ins to visualize changes in FRET ratio in real-time during any experiments before being stored in a graphics format compatible with the build-in ImageJ freeware used for subsequent data analysis. This low-cost system is characterized by high flexibility and can be successfully used to monitor various biochemical events and signaling molecules by a plethora of available FRET biosensors in live cells and tissues. As an example, we demonstrate how to use this imaging system to perform real-time monitoring of cAMP in live 293A cells upon stimulation with a &beta;-adrenergic receptor agonist and blocker.

F&#246;rster resonance energy transfer (FRET) microscopy is a powerful technique for real-time monitoring of signaling events in live cells using various biosensors as reporters. Here we describe how to build a customized epifluorescence FRET imaging system from commercially available components and how to use it for FRET experiments.

F&ouml;rster resonance energy transfer (FRET) microscopy  continues to gain increasing interest as a technique for real-time monitoring  of biochemical ...

The Use of Chemostats in Microbial Systems Biology

Cells regulate their rate of growth in response to signals from the external world. As the cell grows, diverse cellular processes must be coordinated including macromolecular synthesis, metabolism and ultimately, commitment to the cell division cycle. The chemostat, a method of experimentally controlling cell growth rate, provides a powerful means of systematically studying how growth rate impacts cellular processes - including gene expression and metabolism - and the regulatory networks that control the rate of cell growth. When maintained for hundreds of generations chemostats can be used to study adaptive evolution of microbes in environmental conditions that limit cell growth. We describe the principle of chemostat cultures, demonstrate their operation and provide examples of their various applications. Following a period of disuse after their introduction in the middle of the twentieth century, the convergence of genome-scale methodologies with a renewed interest in the regulation of cell growth and the molecular basis of adaptive evolution is stimulating a renaissance in the use of chemostats in biological research.

Cell growth rate is a regulated process and a primary determinant of cell physiology. Continuous culturing using chemostats enables extrinsic control of cell growth rate by nutrient limitation facilitating the study of molecular networks that control cell growth and how those networks evolve to optimize cell growth.

Cells regulate their rate of growth in response to signals from the external world. As the cell grows, diverse cellular processes must be coordinated ...

Analysis of Nephron Composition and Function in the Adult Zebrafish Kidney

The zebrafish model has emerged as a relevant system to study kidney development, regeneration and disease. Both the embryonic and adult zebrafish kidneys are composed of functional units known as nephrons, which are highly conserved with other vertebrates, including mammals. Research in zebrafish has recently demonstrated that two distinctive phenomena transpire after adult nephrons incur damage: first, there is robust regeneration within existing nephrons that replaces the destroyed tubule epithelial cells; second, entirely new nephrons are produced from renal progenitors in a process known as neonephrogenesis. In contrast, humans and other mammals seem to have only a limited ability for nephron epithelial regeneration. To date, the mechanisms responsible for these kidney regeneration phenomena remain poorly understood. Since adult zebrafish kidneys undergo both nephron epithelial regeneration and neonephrogenesis, they provide an outstanding experimental paradigm to study these events. Further, there is a wide range of genetic and pharmacological tools available in the zebrafish model that can be used to delineate the cellular and molecular mechanisms that regulate renal regeneration. One essential aspect of such research is the evaluation of nephron structure and function. This protocol describes a set of labeling techniques that can be used to gauge renal composition and test nephron functionality in the adult zebrafish kidney. Thus, these methods are widely applicable to the future phenotypic characterization of adult zebrafish kidney injury paradigms, which include but are not limited to, nephrotoxicant exposure regimes or genetic methods of targeted cell death such as the nitroreductase mediated cell ablation technique. Further, these methods could be used to study genetic perturbations in adult kidney formation and could also be applied to assess renal status during chronic disease modeling.

The zebrafish adult kidney is an excellent system for renal regeneration and disease studies. An essential aspect of such research is the assessment of nephron structure and function. This protocol describes several methodologies that can be implemented to assess nephron tubule composition and to evaluate renal reabsorption.

The zebrafish model has emerged as a relevant system to study kidney development, regeneration and disease. Both the embryonic and adult zebrafish kidneys ...

Nephrotoxin Microinjection in Zebrafish to Model Acute Kidney Injury

The kidneys are susceptible to harm from exposure to chemicals they filter from the bloodstream. This can lead to organ injury associated with a rapid decline in renal function and development of the clinical syndrome known as acute kidney injury (AKI). Pharmacological agents used to treat medical circumstances ranging from bacterial infection to cancer, when administered individually or in combination with other drugs, can initiate AKI. Zebrafish are a useful animal model to study the chemical effects on renal function in vivo, as they form an embryonic kidney comprised of nephron functional units that are conserved with higher vertebrates, including humans. Further, zebrafish can be utilized to perform genetic and chemical screens, which provide opportunities to elucidate the cellular and molecular facets of AKI and develop therapeutic strategies such as the identification of nephroprotective molecules. Here, we demonstrate how microinjection into the zebrafish embryo can be utilized as a paradigm for nephrotoxin studies.

Renal injuries incurred from nephrotoxins, which include drugs ranging from antibiotics to chemotherapeutics, can result in complex disorders whose pathogenesis remains incompletely understood. This protocol demonstrates how zebrafish can be used for disease modeling of these conditions, which can be applied to the identification of renoprotective measures.

The kidneys are susceptible to harm from exposure to chemicals they filter from the bloodstream. This can lead to organ injury associated with a rapid ...

Measuring Lactase Enzymatic Activity in the Teaching Lab

Understanding how enzymes work, and relating this to real life examples, is critical to a wide range of undergraduate degrees in the biological and biomedical sciences. This easy to follow protocol was developed for first year undergraduate pharmacy students and provides an entry-level introduction to enzyme reactions and analytical procedures for enzyme analysis. The enzyme of choice is lactase, as this represents an example of a commercially available enzyme relevant to human disease/pharmaceutical practice. Lactase is extracted from dietary supplement tablets, and assessed using a colorimetric assay based upon hydrolysis of an artificial substrate for lactase (ortho-nitrophenol-beta-D-galactopyranoside, ONPG). Release of ortho-nitrophenol following the hydrolytic cleavage of ONPG by lactase is measured by a change in absorbance at 420 nm, and the effect of the temperature on the enzymatic reaction is evaluated by carrying out the reaction on ice, at room temperature and at 37 &#176;C. More advanced analysis can be implemented using this protocol by assessing the enzyme activity under different conditions and using different reagents.

The enzymatic activity of lactase is essential for the catabolic processing of the disaccharide lactose. Here, the activity of lactase found in dietary supplements is assayed using a colorimetric assay. This provides students with an experimental platform for understanding the activity of lactase and enzyme kinetics.

Understanding how enzymes work, and relating this to real life examples, is critical to a wide range of undergraduate degrees in the biological and ...

The Mouse Isolated Perfused Kidney Technique

The mouse isolated perfused kidney (MIPK) is a technique for keeping a mouse kidney under ex vivo conditions perfused and functional for 1 hr. This is a prerequisite for studying the physiology of the isolated organ and for many innovative applications that may be possible in the future, including perfusion decellularization for kidney bioengineering or the administration of anti-rejection or genome-editing drugs in high doses to prime the kidney for transplantation. During the time of the perfusion, the kidney can be manipulated, renal function can be assessed, and various pharmaceuticals administered. After the procedure, the kidney can be transplanted or processed for molecular biology, biochemical analysis, or microscopy.
This paper describes the perfusate and the surgical technique needed for the ex vivo perfusion of mouse kidneys. Details of the perfusion apparatus are given and data are presented showing the viability of the kidney&#39;s preparation: renal blood flow, vascular resistance, and urine data as functional, transmission electron micrographs of different nephron segments as morphological readouts, and western blots of transport proteins of different nephron segments as molecular readout.

The mouse isolated perfused kidney (MIPK) is a technique for keeping a mouse kidney under ex vivo conditions perfused and functional for 1 hr. The buffers and surgical technique are described in detail.

The mouse isolated perfused kidney (MIPK) is a technique for keeping a mouse kidney under ex vivo conditions perfused and functional for 1 hr. This is a ...

Visualizing and Tracking Endogenous mRNAs in Live Drosophila melanogaster Egg Chambers

Fluorescence-based imaging techniques, in combination with developments in light microscopy, have revolutionized how cell biologists conduct live cell imaging studies. Methods for detecting RNAs have expanded greatly since seminal studies linked site-specific mRNA localization to gene expression regulation. Dynamic mRNA processes can now be visualized via approaches that detect mRNAs, coupled with microscopy set-ups that are fast enough to capture the dynamic range of molecular behavior. The molecular beacon technology is a hybridization-based approach capable of direct detection of endogenous transcripts in living cells. Molecular beacons are hairpin-shaped, internally quenched, single-nucleotide discriminating nucleic acid probes, which fluoresce only upon hybridization to a unique target sequence. When coupled with advanced fluorescence microscopy and high-resolution imaging, they enable one to perform spatial and temporal tracking of intracellular movement of mRNAs. Although this technology is the only method capable of detecting endogenous transcripts, cell biologists have not yet fully embraced this technology due to difficulties in designing such probes for live cell imaging. A new software application, PinMol, allows for enhanced and rapid design of probes best suited to efficiently hybridize to mRNA target regions within a living cell. In addition, high-resolution, real-time image acquisition and current, open source image analysis software allow for a refined data output, leading to a finer evaluation of the complexity underlying the dynamic processes involved in the mRNA&#39;s life cycle.
Here we present a comprehensive protocol for designing and delivering molecular beacons into Drosophila melanogaster egg chambers. Direct and highly specific detection and visualization of endogenous maternal mRNAs is performed via spinning disc confocal microscopy. Imaging data is processed and analyzed using object detection and tracking in Icy software to obtain details about the dynamic movement of mRNAs, which are transported and localized to specialized regions within the oocyte.

Visualizing and Tracking Endogenous mRNAs in Live Drosophila melanogaster Egg Chambers

Here, we present a protocol for the visualization, detection, analysis and tracking of endogenous mRNA trafficking in live Drosophila melanogaster egg chamber using molecular beacons, spinning disc confocal microscopy, and open-source analysis software.

Fluorescence-based imaging techniques, in combination with developments in light microscopy, have revolutionized how cell biologists conduct live cell ...

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

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Use of the Protease Fluorescent Detection Kit to Determine Protease Activity

The use of SC1 (Pluripotin) to Support mESC Self-renewal in the Absence of LIF

Dissection of the Adult Zebrafish Kidney

FRET Microscopy for Real-time Monitoring of Signaling Events in Live Cells Using Unimolecular Biosensors

The Use of Chemostats in Microbial Systems Biology

Analysis of Nephron Composition and Function in the Adult Zebrafish Kidney

Nephrotoxin Microinjection in Zebrafish to Model Acute Kidney Injury

Measuring Lactase Enzymatic Activity in the Teaching Lab

The Mouse Isolated Perfused Kidney Technique

Visualizing and Tracking Endogenous mRNAs in Live Drosophila melanogaster Egg Chambers