Mark McCauley is supported by an NIH mentored-training grant 5T32HL066991-07
Xander Wehrens is supported by NIH/NHLBI grants 1R01HL091947-01A209 and 3R01HL089598-03S109

1. Preparation of Telemeter for Surgical Implantation
<ol>
 <li>Before inserting an ECG telemeter into the mouse, it is important to make sure the ECG telemeter is sterile and in good working order. New telemetry devices are typically provided in a sterile condition by the manufacturer. ECG telemeters can be reused provided the device is cleaned using Tergazyme 1% solution for at least 4 hours. You may rinse the telemeter with sterile water after cleaning with Tergazyme. Additionally, use Wavicide-01 disinfectant overnight to sterilize the ECG telemeter. Be sure to wash disinfectant off with sterile water 48 hours before implantation into mice. Store in a sterile container.</li>
 <li>Check the telemeter leads for integrity of both the conducting wires and the insulating sheath. Turn on the telemetry transmitter using a magnet waved within 5 cm of the telemeter, and test the signal with an AM radio, frequency 530. The signal should be strong and clear and should vary in intensity and pitch based on manipulation of the wires. Record the model number, the telemeter serial number, and the ECG calibration value. Manufacturers often recommend that the telemeter should be on for 24 hours prior to implantation, see the instruction manual for your specific telemeter for details. In this example, we will use a Data Sciences International (DSI) telemeter for implantation.</li>
 <li>Telemeter lead preparation may be achieved by cutting the negative (white) lead to approximately 3.5 cm and the positive (red) lead to 2.5 cm. These are the optimal lengths for implantation of the negative lead in the right chest, and positive lead in the left abdomen (see figure 2). Next, remove the insulating sheath to expose 7 mm of wire. Paint the end of the metal lead with sterile superglue, such as Vet-Bond, and then attach lead caps to the metal tips. These caps will avoid skin erosion due to lead placement. Approximately 2-3 mm metal wire should be exposed for electrical sensing of native heart rhythm.</li>
</ol>
2. Surgical Implantation of the Telemeter
 <ol>
 <li>In order to use sterile technique in murine surgery, you will need sterile gloves, a sterile drape, and 6-0 Prolene sutures. You will also need to 2 pairs of sterilized forceps, 1 pair of blunt-ended scissors, a scalpel, and a needle driver. Surgical instruments may be sterilized in a glass bead sterilizer heated to 250 degrees Celsius. Be sure to sterilize and wash the telemeter as previously described before use. </li>
 <li>Prepare the mouse for surgery by first anesthetizing the mouse in an induction box using 3% isoflurane in 0.5 L/min 100% O2. Use a heating pad, such as a T-pump, which circulates warm water, to maintain the anesthetized mouse's temperature. Temperature may be monitored by rectal probe, if desired. When the mouse is adequately sedated, completely shave the abdomen and chest of the mouse with electric clippers. Re-anesthetize the mouse with isoflurane before transferring the mouse to a constant-flow tube, supine, with head facing away and tail towards you. Tape down the mouse's paws to the heated surgical table. Wipe the abdomen with an alcohol swab to remove shaved hair and to clean the operative field. Apply three separate coats of betadine with swabs to disinfect the abdomen and chest. Verify correct level of anesthesia by applying pressure on the mouse nail bed. After the mouse has been prepared for surgery, use sterile gloves and apply the sterile drape to the surgical field. </li>
 <li>Intraperitoneal telemeter implantation offers the advantage of exercise physiology experimentation. For intraperitoneal insertion, begin surgery by using a scalpel to create a vertical midline incision in the skin overlying the abdomen roughly 2.5 cm in length. Carefully separate skin from underlying connective tissues using blunt-ended scissors. Then, create a vertical midline incision in the linea alba overlying the peritoneum, roughly 1.5 cm in length. Additionally, create a small hole in the peritoneum just above (cranial) to the peritoneal incision, which will serve as an outlet for subcutaneous leads. While performing surgery, keep the surgical site moist by occasionally dripping sterile saline onto the operating field. </li>
 <li>Insert the telemeter into the right peritoneal cavity. Insert forceps into the hole superior to the telemeter and pull both leads up through the lead hole so they are protruding from the peritoneum. Use continuous 6-0 Prolene sutures to close the 1.5 cm peritoneal vertical midline incision. </li>
 </ol>
3. Lead Implantation and Abdominal Closure
 <ol>
 <li>The ECG leads are placed in the lead II configuration. The lead with the white/transparent sheath is negative, and is placed in the left upper abdomen. First, create a 0.5 cm skin incision in the mouse's upper right chest. Next, use the blunt scissors to create a tunnel back to the abdominal incision. Pull the lead through the tunnel and use a 6-0 Prolene suture to anchor the lead to the pectoral muscle. Make sure the suture is on top of the exposed part of the lead, and creates a good contact between lead and underlying muscle. Use a second suture proximal to the aforementioned one, to immobilize the lead to the muscle. Close the skin incision using a 6-0 Prolene suture.</li>
 <li>The positive lead (red sheath) is placed in the left abdomen below the left diaphragm and below the heart. The lead is anchored to the underlying peritoneal tissue by 6-0 Prolene suture and should have good contact with the peritoneal tissue. An additional incision is not necessary in this step because the site of lead implantation is close to the vertical midline incision, thus may be easily accessed with the existing surgical field. Note that the peritoneal tissue must be lifted to avoid perforation of the underlying intestine.</li>
 <li>Close the abdominal fascia and skin sequentially with 6-0 Prolene in layers.</li>
 </ol>
4. Post-Operative Care
 <ol>
 <li>Give the mouse 0.1 mL of [1 mg/mL] buprenorphine for analgesia immediately after surgery. Allow the mouse to recover from surgery on a heated pad. If the mouse has had prolonged surgery &gt; 30 min, or if the mouse appears dry, you may inject 0.2-0.3 mL sterile saline into the peritoneum for rehydration. For continued post-operative analgesia it is standard practice to provide mice with buprenorphine twice daily for three days, and then as needed every eight hours.</li>
 <li>Clean off betadine with an alcohol swab to avoid post-surgical irritation. Remember to record the post-surgical weight for subsequent determination of post-surgical health. </li>
 <li>8-24 hours after surgery, give the mouse additional analgesia with buprenorphine when needed, as this will reduce pain and reduce lead placement failure from mouse's clawing at the post-surgical site.</li>
 </ol>
5. Representative Results
When performed correctly, the mouse should have closed abdominal incisions with subcutaneous leads under the skin. Usually, the mouse takes 10-30 minutes to recover from anesthesia. It takes about 7-10 days before the mice have completely recovered from surgery, as evidenced by recovery of post-operative weight loss and normalization of mobility.
The most common complication from the surgery is erosion of limb leads through the skin in the first 10 days after the surgery. This common complication may be avoided by firmly adhering subcutaneous leads to subcutaneous tissues or peritoneum. This technique avoids slack leads, which create pressure and friction on the skin when the mouse ambulates. Although less common, infection and sepsis may occur from errors in sterile technique, or from incomplete cleaning of used telemeters. 
ECG output may be recorded by a receiver matrix coupled with data acquisition software, such as Data Sciences International (DSI). Waveform results should include a clearly defined P wave, denoting atrial depolarization, and also a QRS wave which signifies ventricular depolarization, as shown in Figure 3 below. A good quality ECG trace should have a clearly defined P wave preceding a clearly defined QRS wave in a 1:1 ratio. There should be low background signal and R-R intervals should be regular in wild-type mice. A poor quality ECG trace may have indistinct P or QRS waves, or background signal, which may make subsequent computer analysis difficult. A poor ECG trace may be fixed by re-implanting telemeter leads, usually one at a time. To re-implant a telemeter lead, place the mouse under anesthesia, and using surgical technique, make an incision in the skin directly overlying the lead in question. When the lead is accessed, cut the suture anchoring the lead, and move the lead to the desired location before anchoring the lead again. Finally, close the surgical site with 6-0 Prolene. 
<img src="/files/ftp_upload/1739/1739fig1.jpg" alt="Figure 1" /> Figure 1: An example of a small animal telemeter with positive and negative leads. In general, the red lead is the positive lead, and the white lead is the negative lead. Each lead should have an insulating sheath, 5-7 mm of exposed wire, and a plastic capped tip, which prevents erosion of the lead through the skin.
<img src="/files/ftp_upload/1739/1739fig2.jpg" alt="Figure 2" /> Figure 2: Cartoon showing proper telemeter implantation sites on a mouse. The white (negative) lead is implanted in the mouse's right upper chest, and the red (positive) lead is placed in the left abdomen, such that the telemeter senses the "lead two" configuration.
<img src="/files/ftp_upload/1739/1739fig3.jpg" alt="Figure 3" /> Figure 3: Representative waveform of a mouse in normal sinus rhythm. P wave represents atrial depolarization and QRS complex represents ventricular depolarization. A high quality waveform should have a distinct P wave preceding each QRS complex in a 1:1 ratio.

<ol>
	<li>Cerrone, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Bidirectional+ventricular+tachycardia+and+fibrillation+elicited+in+a+knock-in+mouse+model+carrier+of+a+mutation+in+the+cardiac+ryanodine+receptor.">Bidirectional ventricular tachycardia and fibrillation elicited in a knock-in mouse model carrier of a mutation in the cardiac ryanodine receptor.</a> Circ Res. 96 (10), e77-e77 (2005).</li><li>Chelu, M. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Calmodulin+kinase+II-mediated+sarcoplasmic+reticulum+Ca2++leak+promotes+atrial+fibrillation+in+mice.">Calmodulin kinase II-mediated sarcoplasmic reticulum Ca2+ leak promotes atrial fibrillation in mice.</a> J Clin Invest. 119 (7), 1940-1940 (1940).</li><li>Sood, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Intracellular+calcium+leak+due+to+FKBP12.6+deficiency+in+mice+facilitates+the+inducibility+of+atrial+fibrillation.">Intracellular calcium leak due to FKBP12.6 deficiency in mice facilitates the inducibility of atrial fibrillation.</a> Heart Rhythm. 5 (7), 1047-1047 (2008).</li><li>Swoap, S. J., Gutilla, M. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cardiovascular+changes+during+daily+torpor+in+the+laboratory+mouse.">Cardiovascular changes during daily torpor in the laboratory mouse.</a> Am J Physiol Regul Integr Comp Physiol. 297 (3), R769-R769 (2009).</li><li>Costa-Goncalves, d. a., C, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Role+of+the+multidomain+protein+spinophilin+in+blood+pressure+and+cardiac+function+regulation.">Role of the multidomain protein spinophilin in blood pressure and cardiac function regulation.</a> Hypertension. 52 (4), 702-702 (2008).</li><li>Wehrens, X. H., Kirchhoff, S., Doevendans, P. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mouse+electrocardiography:+an+interval+of+thirty+years.">Mouse electrocardiography: an interval of thirty years.</a> Cardiovasc Res. 45 (1), 231-231 (2000).</li></ol>

The critical steps of this procedure are proper cleaning and preparation of the mouse prior to survival surgery. When a telemeter and the mouse operative site are properly cleaned, there is a much better post-surgical outcome. Additionally, close attention to lead placement reduces post-surgical skin erosion. Possible modifications to the technique include alternative configurations to evaluate lateral LV wall function (lead I configuration), or the use of the telemeter not only for ECG waveform analysis, but also for...

<!DOCTYPE html PUBLIC "-//W3C//DTD XHTML 1.0 Transitional//EN" "http://www.w3.org/TR/xhtml1/DTD/xhtml1-transitional.dtd">
<html xmlns="http://www.w3.org/1999/xhtml" xml:lang="en" lang="en">	
		
		<div>
		<table border="1">
		<thead>
		<tr>
		<th>Material Name</th>
		<th>Type</th>
		<th>Company</th>
		<th>Catalogue Number</th>
		<th>Comment</th>
		</tr>
		</thead>
		<tbody>
		
<tr>
<td>Steri 250 Bead Sterilizer</td>
<td>&nbsp;</td>
<td>Inotech</td>
<td>IS-250</td>
<td>&nbsp;</td>
<tr>
<td>Blunt-Ended Scissors</td>
<td>&nbsp;</td>
<td>Roboz</td>
<td>RS-5980</td>
<td>&nbsp;</td>
<tr>
<td>Blunt-Ended Forceps (x2)</td>
<td>&nbsp;</td>
<td>Roboz</td>
<td>RS-5138</td>
<td>&nbsp;</td>
<tr>
<td>Veterinary Anesthesia Vaporizer</td>
<td>&nbsp;</td>
<td>Surgivet</td>
<td>Isotec 4</td>
<td>&nbsp;</td>
<tr>
<td>T/Pump</td>
<td>&nbsp;</td>
<td>Gaymar</td>
<td>P/N11184-000</td>
<td>&nbsp;</td>
<tr>
<td>Propylene Blue Monofilament Suture</td>
<td>&nbsp;</td>
<td>Ethicon</td>
<td>8718</td>
<td>&nbsp;</td>
<tr>
<td>Povodone Iodine, 10%</td>
<td>&nbsp;</td>
<td>Equate</td>
<td>N/A</td>
<td>&nbsp;</td>
<tr>
<td>Alcohol Swabs</td>
<td>&nbsp;</td>
<td>PDI</td>
<td>B339</td>
<td>&nbsp;</td>
<tr>
<td>Tergazyme</td>
<td>&nbsp;</td>
<td>Alconox</td>
<td>21837-118</td>
<td>&nbsp;</td>
<tr>
<td>Wavidicde-01</td>
<td>&nbsp;</td>
<td>Medical Chemical Corporation</td>
<td>Wavicide-01</td>
<td>&nbsp;</td>
<tr>
<td>ECG Telemeter</td>
<td>&nbsp;</td>
<td>DataSciences International</td>
<td>EA-F20</td>
<td>&nbsp;</td>		</tbody>
		</table>
		</div>

ambulatory ecg recording in mice

Telemetric ECG recording in mice is essential to understanding the mechanisms behind arrhythmias, conduction disorders, and sudden cardiac death. Although the surface ECG is utilized for short-term measurements of waveform intervals, it is not practical for long-term studies of heart rate variability or the capture of rare episodes of arrhythmias. Implantable ECG telemeters offer the advantages of simple surgical implantation, long-term recording of electrograms in ambulatory mice, and scalability with simultaneous recordings of multiple animals. Here, we present a step-by-step guide to the implantation of telemeters for ambulatory ECG recording in mice. Careful adherence to aseptic technique is required for favorable survival results with the possibility of implantation and recording from weeks to months. Thus, implantable ECG telemetry is a valuable tool for detection of critical information on cardiac electrophysiology in ambulatory animal models such as the mouse. 

Watch this Scientific Journal Video about Ambulatory ECG Recording in Mice at JoVE.com

Ambulatory ECG Recording in Mice

Telemetric ECG has emerged as an essential tool in evaluating animal models for cardiac arrhythmias and sudden cardiac death. Here, we present a stepwise guide to telemetric ECG recordings for application in long-term ambulatory ECG monitoring in mice.

Telemetric ECG recording in mice is essential to understanding the mechanisms behind arrhythmias, conduction disorders, and sudden cardiac death. Although ...

Research

JoVE Journal

Biology

23.5K Views.  Baylor College of Medicine (BCM). Телеметрическая ЭКГ стала важным инструментом в оценке животных моделей для сердечной аритмии и внезапной сердечной смерти. Здесь мы представляем ступенчато руководство к телеметрической записи ЭКГ для применения в долгосрочной суточного мониторирования ...

Video: Ambulatory ECG Recording in Mice

Small Bowel Transplantation In Mice

Since 1990, the development of tacrolimus-based immunosuppression and improved surgical techniques, the increased array of potent immunosuppressive medications, infection prophylaxis, and suitable patient selection helped improve actuarial graft and patient survival rates for all types of intestine transplantation. Patients with irreversible intestinal failure and complications of parenteral nutrition should now be routinely considered for small intestine transplantation. However, Survival rates for small intestinal transplantation have been slow to improve compares increasingly favorably with renal, liver, heart and lung. The small bowel transplantation is still unsatisfactory compared with other organs. Further progress may depend on better understanding of immunology and physiology of the graft and can be greatly facilitated by animal models. A wider use of mouse small bowel transplantation model is needed in the study of immunology and physiology of the transplantation gut as well as efficient methods in diagnosing early rejection. However, this model is limited to use because the techniques involved is an extremely technically challenging. We have developed a modified technique. When making anastomosis of portal vein and inferior vena cava, two stay sutures are made at the proximal apex and distal apex of the recipient s inferior vena cava with the donor s portal vein. The left wall of the inferior vena cava and donor s portal vein is closed with continuing sutures in the inside of the inferior vena cava after, after one knot with the proximal apex stay suture the right wall of the inferior vena cava and the donor s portal vein are closed with continuing sutures outside the inferior vena cave with 10-0 sutures. This method is easier to perform because anastomosis is made just on the one side of the inferior vena cava and 10-0 sutures is the right size to avoid bleeding and thrombosis.  In this article, we provide details of the technique to supplement the video.

The mouse small bowel transplantation model has been recognized as an important tool to study mechanismes of immune rejection and screen new immunosuppressive drugs. However, this model is limited to use because the techniques involved is an extremely technically challenge. Now we introduce the modified technique.

Since 1990, the development of tacrolimus-based immunosuppression and improved surgical techniques, the increased array of potent immunosuppressive ...

Patch Clamp Recording of Ion Channels Expressed in  Xenopus Oocytes

Since its development by Sakmann and Neher 1, 2, the patch clamp has become established as an extremely useful technique for electrophysiological measurement of single or multiple ion channels in cells. This technique can be applied to ion channels in both their native environment and expressed in heterologous cells, such as oocytes harvested from the African clawed frog, Xenopus laevis. Here, we describe the well-established technique of patch clamp recording from Xenopus oocytes. This technique is used to measure the properties of expressed ion channels either in populations (macropatch) or individually (single-channel recording). We focus on techniques to maximize the quality of oocyte preparation and seal generation. With all factors optimized, this technique gives a probability of successful seal generation over 90 percent. The process may be optimized differently by every researcher based on the factors he or she finds most important, and we present the approach that have lead to the greatest success in our hands.

This is intended as an introduction to patch clamp recording from Xenopus laevis oocytes. It covers vitelline membrane removal, formation of a gigaohm seal (gigaseal), and the optional conversion of the patch to the outside-out topology.

Since its development by Sakmann and Neher 1, 2, the patch clamp has become established as an extremely useful technique for electrophysiological ...

Micro-drive Array for Chronic in vivo Recording: Drive Fabrication

Chronic recording of large populations of neurons is a valuable technique for studying the function of neuronal circuits in awake behaving rats. Lightweight recording devices carrying a high density array of tetrodes allow for the simultaneous monitoring of the activity of tens to hundreds of individual neurons. Here we describe a protocol for the fabrication of a micro-drive array with twenty one independently movable micro-drives. This device has been used successfully to record from hippocampal and cortical neurons in our lab. We show how to prepare a custom designed, 3-D printed plastic base that will hold the micro-drives. We demonstrate how to construct the individual micro-drives and how to assemble the complete micro-drive array. Further preparation of the drive array for surgical implantation, such as the fabrication of tetrodes, loading of tetrodes into the drive array and gold-plating, is covered in a subsequent video article.

Micro-drive Array for Chronic in vivo Recording: Drive Fabrication

In this protocol we demonstrate how to fabricate a micro-drive array for chronic electrophysiological recordings in rats.

Chronic recording of large populations of neurons is a valuable technique for studying the function of neuronal circuits in awake behaving rats. ...

Micro-drive Array for Chronic in vivo Recording: Tetrode Assembly

The tetrode, a bundle of four electrodes, has proven to be a valuable tool for the simultaneous recording of multiple neurons in-vivo.  The differential amplitude of action potential signatures over the channels of a tetrode allows for the isolation of single-unit activity from multi-unit signals.  The ability to precisely control the stereotaxic location and depth of the tetrode is critical for studying coordinated neural activity across brain regions.  In combination with a micro-drive array, it is possible to achieve precise placement and stable control of many tetrodes over the course of days to weeks.  In this protocol, we demonstrate how to fabricate and condition tetrodes using basic tools and materials, install the tetrodes into a multi-drive tetrode array for chronic in-vivo recording in the rat, make ground wire connections to the micro-drive array, and attach a protective cone onto the micro-drive array in order to protect the tetrodes from physical contact with the environment.



Micro-drive Array for Chronic in vivo Recording: Tetrode Assembly

In this protocol we demonstrate how to fabricate and condition tetrodes for use with a micro-drive array, which was designed for chronic electrophysiological recordings in rats. In addition, we illustrate the final stages of micro-drive array construction, which includes installing ground wires and a protective cone.

The tetrode, a bundle of four electrodes, has proven to be a valuable tool for the simultaneous recording of multiple neurons in-vivo.  The differential ...

Measuring Blood Pressure in Mice using Volume Pressure Recording, a Tail-cuff Method

The CODA 8-Channel High Throughput Non-Invasive Blood Pressure system measures the blood pressure in up to 8 mice or rats simultaneously. The CODA tail-cuff system uses Volume Pressure Recording (VPR) to measure the blood pressure by determining the tail blood volume. A specially designed differential pressure transducer and an occlusion tail-cuff measure the total blood volume in the tail without the need to obtain the individual pulse signal. Special attention is afforded to the length of the occlusion cuff in order to derive the most accurate blood pressure readings. VPR can easily obtain readings on dark-skinned rodents, such as C57BL6 mice and is MRI compatible. The CODA system provides you with measurements of six (6) different blood pressure parameters; systolic and diastolic blood pressure, heart rate, mean blood pressure, tail blood flow, and tail blood volume. Measurements can be made on either awake or anesthetized mice or rats. The CODA system includes a controller, laptop computer, software, cuffs, animal holders, infrared warming pads, and an infrared thermometer. There are seven different holder sizes for mice as small as 8 grams to rats as large as 900 grams.

The CODA 8-Channel High Throughput Non-Invasive Blood Pressure system measures the blood pressure in up to 8 mice or rats simultaneously. This tail-cuff system uses Volume Pressure Recording (VPR) to measure the blood pressure by determining the tail blood volume.

The CODA 8-Channel High Throughput Non-Invasive Blood Pressure system measures the blood pressure in up to 8 mice or rats simultaneously. The CODA ...

Electrophysiological Recording in the Drosophila Embryo

Drosophila is a premier genetic model for the study of both embryonic development and functional neuroscience. Traditionally, these fields are quite isolated from each other, with largely independent histories and scientific communities. However, the interface between these usually disparate fields is the developmental programs underlying acquisition of functional electrical signaling properties and differentiation of functional chemical synapses during the final phases of neural circuit formation. This interface is a critically important area for investigation. In Drosophila, these phases of functional development occur during a period of &lt;8 hours (at 25&deg;C) during the last third of embryogenesis. This late developmental period was long considered intractable to investigation owing to the deposition of a tough, impermeable epidermal cuticle. A breakthrough advance was the application of water-polymerizing surgical glue that can be locally applied to the cuticle to enable controlled dissection of late-stage embryos. With a dorsal longitudinal incision, the embryo can be laid flat, exposing the ventral nerve cord and body wall musculature to experimental investigation. Whole-cell patch-clamp techniques can then be employed to record from individually-identifiable neurons and somatic muscles. These recording configurations have been used to track the appearance and maturation of ionic currents and action potential propagation in both neurons and muscles. Genetic mutants affecting these electrical properties have been characterized to reveal the molecular composition of ion channels and associated signaling complexes, and to begin exploration of the molecular mechanisms of functional differentiation. A particular focus has been the assembly of synaptic connections, both in the central nervous system and periphery. The glutamatergic neuromuscular junction (NMJ) is most accessible to a combination of optical imaging and electrophysiological recording. A glass suction electrode is used to stimulate the peripheral nerve, with excitatory junction current (EJC) recordings made in the voltage-clamped muscle. This recording configuration has been used to chart the functional differentiation of the synapse, and track the appearance and maturation of presynaptic glutamate release properties. In addition, postsynaptic properties can be assayed independently via iontophoretic or pressure application of glutamate directly to the muscle surface, to measure the appearance and maturation of the glutamate receptor fields. Thus, both pre- and postsynaptic elements can be monitored separately or in combination during embryonic synaptogenesis. This system has been heavily used to isolate and characterize genetic mutants that impair embryonic synapse formation, and thus reveal the molecular mechanisms governing the specification and differentiation of synapse connections and functional synaptic signaling properties.

Electrophysiological Recording in the Drosophila Embryo

Electrophysiological recordings from Drosophila embryos allow analyses of developing muscle and neuron electrical properties, as well as characterization of functional synaptogenesis at the glutamatergic neuromuscular junction and central cholinergic and GABAergic synapses.

Drosophila is a premier genetic model for the study of both embryonic development and functional neuroscience. Traditionally, these fields are quite ...

Transverse Aortic Constriction in Mice

Transverse aortic constriction (TAC) in the mouse is a commonly used experimental model for pressure overload-induced cardiac hypertrophy and heart failure.1 TAC initially leads to compensated hypertrophy of the heart, which often is associated with a temporary enhancement of cardiac contractility. Over time, however, the response to the chronic hemodynamic overload becomes maladaptive, resulting in cardiac dilatation and heart failure.2 The murine TAC model was first validated by Rockman et al.1, and has since been extensively used as a valuable tool to mimic human cardiovascular diseases and elucidate fundamental signaling processes involved in the cardiac hypertrophic response and heart failure development. When compared to other experimental models of heart failure, such as complete occlusion of the left anterior descending (LAD) coronary artery, TAC provides a more reproducible model of cardiac hypertrophy and a more gradual time course in the development of heart failure. Here, we describe a step-by-step procedure to perform surgical TAC in mice. To determine the level of pressure overload produced by the aortic ligation, a high frequency Doppler probe is used to measure the ratio between blood flow velocities in the right and left carotid arteries.3, 4 With surgical survival rates of 80-90%, transverse aortic banding is an effective technique of inducing left ventricular hypertrophy and heart failure in mice.

Transverse aortic constriction (TAC) in the mouse is a commonly used experimental model to study mechanisms underlying cardiac hypertrophy and heart failure development. Here, we describe procedures to constrict the aorta to create a reproducible degree of cardiac hypertrophy in mice.

Transverse aortic constriction (TAC) in the mouse is a commonly used experimental model for pressure overload-induced cardiac hypertrophy and heart ...

Recording Multicellular Behavior in Myxococcus xanthus Biofilms using Time-lapse Microcinematography

A swarm of the &delta;-proteobacterium Myxococcus xanthus contains millions of cells that act as a collective, coordinating movement through a series of signals to create complex, dynamic patterns as a response to environmental cues. These patterns are self-organizing and emergent; they cannot be predicted by observing the behavior of the individual cells. Using a time-lapse microcinematography tracking assay, we identified a distinct emergent pattern in M. xanthus called chemotaxis, defined as the directed movement of a swarm up a nutrient gradient toward its source 1.
In order to efficiently characterize chemotaxis via time-lapse microcinematography, we developed a highly modifiable plate complex (Figure 1) and constructed a cluster of 8 microscopes (Figure 2), each capable of capturing time-lapse videos. The assay is rigorous enough to allow consistent replication of quantifiable data, and the resulting videos allow us to observe and track subtle changes in swarm behavior. Once captured, the videos are transferred to an analysis/storage computer with enough memory to process and store thousands of videos. The flexibility of this setup has proven useful to several members of the M. xanthus community.

Recording Multicellular Behavior in Myxococcus xanthus Biofilms using Time-lapse Microcinematography

To study Myxococcus xanthus swarm behavior, we have designed a time-lapse microcinematography protocol that can be modified for different assays. It employs standard growth conditions adapted for microscopy, and yields reproducible results by the use of inexpensive, reusable silicone gaskets. We have used this method to quantify multicellular chemotaxis.

A swarm of the &delta;-proteobacterium Myxococcus xanthus contains millions of cells that act as a collective, coordinating movement through a series of ...

One-channel Cell-attached Patch-clamp Recording

Ion channel proteins are universal devices for fast communication across biological membranes. The temporal signature of the ionic flux they generate depends on properties intrinsic to each channel protein as well as the mechanism by which it is generated and controlled and represents an important area of current research. Information about the operational dynamics of ion channel proteins can be obtained by observing long stretches of current produced by a single molecule. Described here is a protocol for obtaining one-channel cell-attached patch-clamp current recordings for a ligand gated ion channel, the NMDA receptor, expressed heterologously in HEK293 cells or natively in cortical neurons. Also provided are instructions on how to adapt the method to other ion channels of interest by presenting the example of the mechano-sensitive channel PIEZO1. This method can provide data regarding the channel&#8217;s conductance properties and the temporal sequence of open-closed conformations that make up the channel&#8217;s activation mechanism, thus helping to understand their functions in health and disease.

Described here is a procedure for obtaining long stretches of current recording from one ion channel with the cell-attached patch-clamp technique. This method allows for observing, in real time, the pattern of open-close channel conformations that underlie the biological signal. These data inform about channel properties in undisturbed biological membranes.

Ion channel proteins are universal devices for fast communication across biological membranes. The temporal signature of the ionic flux they generate ...

Intravenous Injections in Neonatal Mice

Intravenous injection is a clinically applicable manner to deliver therapeutics. For adult rodents and larger animals, intravenous injections are technically feasible and routine. However, some mouse models can have early onset of disease with a rapid progression that makes administration of potential therapies difficult. The temporal (or facial) vein is just anterior to the ear bud in mice and is clearly visible for the first two days after birth on either side of the head using a dissecting microscope. During this window, the temporal vein can be injected with volumes up to 50 &#956;l. The injection is safe and well tolerated by both the pups and the dams. A typical injection procedure is completed within 1-2 min, after which the pup is returned to the home cage. By the third postnatal day the vein is difficult to visualize and the injection procedure becomes technically unreliable. This technique has been used for delivery of adeno-associated virus (AAV) vectors, which in turn can provide almost body-wide, stable transgene expression for the life of the animal depending on the viral serotype chosen.

Animal models of pediatric disease can experience early onset and aggressive disease progression. Clinically relevant therapy delivery to young mouse models can be technically difficult. This protocol describes a non-invasive intravenous injection method for newborn mice within the first two postnatal days of life.