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 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 rep...

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

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

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 ...

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 ...

Techniques to Induce and Quantify Cellular Senescence

In response to cellular stress or damage, proliferating cells can induce a specific program that initiates a state of long-term cell-cycle arrest, termed cellular senescence. Accumulation of senescent cells occurs with organismal aging and through continual culturing in vitro. Senescent cells influence many biological processes, including embryonic development, tissue repair and regeneration, tumor suppression, and aging. Hallmarks of senescent cells include, but are not limited to, increased senescence-associated &#946;-galactosidase activity (SA-&#946;-gal); p16INK4A, p53, and p21 levels; higher levels of DNA damage, including &#947;-H2AX; the formation of Senescence-associated Heterochromatin Foci (SAHF); and the acquisition of a Senescence-associated Secretory Phenotype (SASP), a phenomenon characterized by the secretion of a number of pro-inflammatory cytokines and signaling molecules. Here, we describe protocols for both replicative and DNA damage-induced senescence in cultured cells. In addition, we highlight techniques to monitor the senescent phenotype using several senescence-associated markers, including SA-&#946;-gal, &#947;-H2AX and SAHF staining, and to quantify protein and mRNA levels of cell cycle regulators and SASP factors. These methods can be applied to the assessment of senescence in various models and tissues.

Cellular senescence, the irreversible state of cell-cycle arrest, can be induced by various cellular stresses. Here, we describe protocols to induce cellular senescence and methods to assess markers of senescence.

In response to cellular stress or damage, proliferating cells can induce a specific program that initiates a state of long-term cell-cycle arrest, termed ...

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 ...

Applications of Spatio-temporal Mapping and Particle Analysis Techniques to Quantify Intracellular Ca2+ Signaling In Situ

Ca2+ imaging of isolated cells or specific types of cells within intact tissues often reveals complex patterns of Ca2+ signaling. This activity requires careful and in-depth analyses and quantification to capture as much information about the underlying events as possible. Spatial, temporal and intensity parameters intrinsic to Ca2+ signals such as frequency, duration, propagation, velocity and amplitude may provide some biological information required for intracellular signalling. High-resolution Ca2+ imaging typically results in the acquisition of large data files that are time consuming to process in terms of translating the imaging information into quantifiable data, and this process can be susceptible to human error and bias. Analysis of Ca2+ signals from cells in situ typically relies on simple intensity measurements from arbitrarily selected regions of interest (ROI) within a field of view (FOV). This approach ignores much of the important signaling information contained in the FOV. Thus, in order to maximize recovery of information from such high-resolution recordings obtained with Ca2+dyes or optogenetic Ca2+ imaging, appropriate spatial and temporal analysis of the Ca2+ signals is required. The protocols outlined in this paper will describe how a high volume of data can be obtained from Ca2+ imaging recordings to facilitate more complete analysis and quantification of Ca2+ signals recorded from cells using a combination of spatiotemporal map (STM)-based analysis and particle-based analysis. The protocols also describe how different patterns of Ca2+ signaling observed in different cell populations in situ can be analyzed appropriately. For illustration, the method will examine Ca2+ signaling in a specialized population of cells in the small intestine, interstitial cells of Cajal (ICC), using GECIs.

Applications of Spatio-temporal Mapping and Particle Analysis Techniques to Quantify Intracellular Ca2+ Signaling In Situ

Genetically encoded Ca2+ indicators (GECIs) have radically changed how in situ Ca2+ imaging is performed. To maximize data recovery from such recordings, appropriate analysis of Ca2+ signals is required. The protocols in this paper facilitate the quantification of Ca2+ signals recorded in situ using spatiotemporal mapping and particle-based analysis.

Ca2+ imaging of isolated cells or specific types of cells within intact tissues often reveals complex patterns of Ca2+ signaling. This activity requires ...