This work was supported by the National Institutes of Health (NIH/NHLBI) [Grant no. HL114869] and the support from UTHSC to SM.

Ethics statement: All animal procedures mentioned in this video-article were reviewed and approved by the Animal Care and Use Committee (IACUC)&#160;at the University of Tennessee Health Science Center.
Note: Use sterile microsurgery instruments throughout the surgical procedure. Surgical instruments may be sterilized utilizing an infrared sterilizer at an optimum sterilizing temperature of 1,500 &#176;F (815.6&#160;&#176;C). Telemeters can be reused, however, before inserting it in the animal make sure to sterilize the entire transmitter according to the manufacturer instructions and that the tip of the catheter has been refilled with a special gel provided by the manufacturer.
1. Experimental Animals
<ol>
	<li>Obtain an IACUC approval for all experiments that will be performed. Consultation with institutional IACUC is highly recommended regarding requirements for post-operative analgesic use after this procedure.</li>
	<li>Keep the animals in climate-controlled rooms having an ambient temperature of 21&#160;&#176;C, 60% relative humidity, and a 12-hr light-dark cycle with free access to food and water. 
	Note: Because the mouse body has to accommodate the telemetry probe (approximatively 1 cm diameter x&#160;2 cm long), it is preferable to use mice that weigh 20 g or more.</li>
</ol>
2. Anesthesia and Operative Preparation
<ol>
	<li>Weigh the mouse using a precision scale and&#160;record its&#160;weight. Make sure to handle the animal gently and quietly.</li>
	<li>Place the animal into the induction chamber and close it. Set oxygen flow rate at 0.5 L/min and set isoflurane concentration at 4 - 5% .</li>
	<li>When the mouse loses consciousness, place it on a body warming plate maintained at 36 - 37 &#176;C. Maintain the anesthesia via nosecone with isoflurane set at 2% (keep flow rate at&#160;0.5 L/min).</li>
	<li>Prepare the operation regions by removing the hair from the back of the neck and the ventral aspect of the neck by applying hair removal cream.</li>
	<li>Treat the incision site with 3 applications of surgical scrub (betadine solution) alternating with 70% isopropyl alcohol.</li>
</ol>
3. Surgery&#160;
<ol>
	<li>Place the mouse in a supine position. Check for reflexes by pinching the foot, and adjust anesthesia&#160;until&#160;there is no response. Make an approximately 1-cm midline incision below the neck of the mouse with a scalpel. In the left side of the cut create a subcutaneous space by carefully separating the skin from the underlying connective tissue.</li>
	<li>Flip the mouse, and use a scalpel to make a skin incision of approximately 1.5 cm in the dorsal left side behind the scapula. Create a subcutaneous pocket along the animal&#39;s flank large enough to accommodate the device. Insert the transmitter into the pocket.</li>
	<li>Insert a small hemostat clamp in the back incision and maneuver it subcutaneously towards the anterior neck opening. Using the inserted-hemostat clamp, gently grasp a non-toxic polyethylene tube (4 cm length x&#160;1 mm I.D.).
	<ol>
		<li>Pull the hemostat back through the tunnel out the lateral incision in the back until the tubing protrudes from both abdominal and dorsal incision, release the tube from the hemostat. From the back, insert the catheter sensor into the tube to tunnel the tip of the pressure sensing catheter through the neck. From the neck anterior, pull and remove the polyethylene tube and close the dorsal incision using metal clips.</li>
	</ol>
	</li>
	<li>Carefully separate the mandibular glands using sterile cotton tip applicators and&#160;retract the left mandibular gland using an elastic stay hook. Using fine-tipped curved forceps, locate the carotid artery along the left side of the trachea.&#160;Keep the surgical site sterile&#160; by placing and securing a sterile drape.
	<ol>
		<li>Carefully isolate the vessel from the surrounding tissue and gently separate the vagus nerve (whitish in color) that is along the carotid artery away from the artery. Be careful not to cut or damage the nerve or the artery.</li>
	</ol>
	</li>
	<li>Pass three pieces of non-absorbable 7-0 suture underneath the isolated carotid artery section. Tie the cranial suture to close off blood flow. Pull the suture that is closest to the sternum to temporarily occlude blood flow from the aorta.</li>
	<li>Make a loose knot using the middle suture.&#160;This will be used to secure the catheter in the vessel. Cut a small incision into the artery between the cranial and the sternal sutures using micro-scissors.</li>
	<li>Grab the catheter with special vessel cannulation forceps, being careful not to squeeze the catheter to prevent gel loss from the probe. Gently, grab the artery with a fine tip curved tweezers forceps, retrieve the catheter, and insert it in the vessel through the small incision.</li>
	<li>Tighten the middle suture node around the artery and gently advance the catheter. Gently release the suture that is proximal to the sternum and continue to advance the catheter towards the transverse aorta.
	<ol>
		<li>Observe the mark on the catheter that gives an approximate index of how far the catheter needs to be inserted. Once the point is reached, gently tighten both the lower and the upper suture around the catheter. The catheter is secured to the carotid artery by suture knots.</li>
	</ol>
	</li>
	<li>Close the skin incision with non-absorbable 5-0 suture. Once closed, seal the incision with tissue adhesive.</li>
</ol>
4. Surgical Recovery and BP Measurements
<ol>
	<li>Monitor animal closely for the return of normal postures and behaviors. During the 24 hr post-surgery period administer analgesia as directed by a staff veterinarian.</li>
	<li>Once the animals have recovered (5 - 7 days post-surgery), house them individually in a regular mouse cage placed on top of the telemetry receiver plate.</li>
	<li>Turn the implanted transmitters &#34;on&#34; and &#34;off&#34; by using a magnetic device briefly positioned close to the animal from the outside of the cage.</li>
</ol>

<ol>
	<li>Danaei, G., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+preventable+causes+of+death+in+the+United+States:+comparative+risk+assessment+of+dietary,+lifestyle,+and+metabolic+risk+factors.">The preventable causes of death in the United States: comparative risk assessment of dietary, lifestyle, and metabolic risk factors.</a> PLoS Med. 6 (4), e1000058 (2009).</li><li>Dornas, W. C., Silva, M. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Animal+models+for+the+study+of+arterial+hypertension.">Animal models for the study of arterial hypertension.</a> J Biosci. 36 (4), 731-737 (2011).</li><li>Henze, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Persistent+alterations+in+heart+rate+variability,+baroreflex+sensitivity,+and+anxiety-like+behaviors+during+development+of+heart+failure+in+the+rat.">Persistent alterations in heart rate variability, baroreflex sensitivity, and anxiety-like behaviors during development of heart failure in the rat.</a> Am J Physiol Heart Circ Physiol. 295 (1), H29-H38 (2008).</li><li>Hoffmann, D. S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Chronic+tempol+prevents+hypertension,+proteinuria,+and+poor+feto-placental+outcomes+in+BPH/5+mouse+model+of+preeclampsia.">Chronic tempol prevents hypertension, proteinuria, and poor feto-placental outcomes in BPH/5 mouse model of preeclampsia.</a> Hypertension. 51 (4), 1058-1065 (2008).</li><li>Lerman, L. O., Chade, A. R., Sica, V., Napoli, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Animal+models+of+hypertension:+an+overview.">Animal models of hypertension: an overview.</a> J Lab Clin Med. 146 (3), 160-173 (2005).</li><li>Johns, C., Gavras, I., Handy, D. E., Salomao, A., Gavras, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Models+of+experimental+hypertension+in+mice.">Models of experimental hypertension in mice.</a> Hypertension. 28 (6), 1064-1069 (1996).</li><li>Vatner, S. F., Braunwald, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cardiovascular+control+mechanisms+in+the+conscious+state.">Cardiovascular control mechanisms in the conscious state.</a> N Engl J Med. 293 (19), 970-976 (1975).</li><li>Zhao, X., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Arterial+Pressure+Monitoring+in+Mice.">Arterial Pressure Monitoring in Mice.</a> Curr Protoc Mouse Biol. 1, 105-122 (2011).</li><li>Whitesall, S. E., Hoff, J. B., Vollmer, A. P., D'Alecy, L. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Comparison+of+simultaneous+measurement+of+mouse+systolic+arterial+blood+pressure+by+radiotelemetry+and+tail-cuff+methods.">Comparison of simultaneous measurement of mouse systolic arterial blood pressure by radiotelemetry and tail-cuff methods.</a> Am J Physiol Heart Circ Physiol. 286 (6), H2408-H2415 (2004).</li><li>Kurtz, T. W., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Recommendations+for+blood+pressure+measurement+in+humans+and+experimental+animals.+Part+2:+Blood+pressure+measurement+in+experimental+animals:+a+statement+for+professionals+from+the+subcommittee+of+professional+and+public+education+of+the+American+Heart+Association+council+on+high+blood+pressure+research.">Recommendations for blood pressure measurement in humans and experimental animals. Part 2: Blood pressure measurement in experimental animals: a statement for professionals from the subcommittee of professional and public education of the American Heart Association council on high blood pressure research.</a> Hypertension. 45 (2), 299-310 (2005).</li><li>Kurtz, T. W., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Recommendations+for+blood+pressure+measurement+in+animals:+summary+of+an+AHA+scientific+statement+from+the+Council+on+High+Blood+Pressure+Research,+Professional+and+Public+Education+Subcommittee.">Recommendations for blood pressure measurement in animals: summary of an AHA scientific statement from the Council on High Blood Pressure Research, Professional and Public Education Subcommittee.</a> Arterioscler Thromb Vasc Biol. 25 (3), 478-479 (2005).</li></ol>

The authors have nothing to disclose.

Implantable radio-telemetry has improved significantly over the last decade; smaller probe size makes the implant less traumatic for the animal, prolonged battery life helps to reduce the costs, and independent telemeter frequencies eliminate crosstalk between receivers. Telemetry is considered the state-of-the-art method for collecting a wide variety of physiological parameters from freely moving animals without the artifacts associated with the use of restraint, human interaction, or anesthesia that are required by oth...

Hypertension is one of the major risk factors for cardiovascular diseases, arguably it is a major public health issue both in developed and developing countries1. Several animal models of experimental hypertension have been developed to mimic hypertensive responses like those observed in humans2. Among others, the ambulatory mouse represents an excellent model to study the genesis and the progression of hypertension allowing in&#160;vivo analysis of the consequences of chronic exposure to hypertension.
Blood pressure (BP) monitoring in mice has helped researchers to unravel several mechanisms involved in the physiology and pathophysiology of diseases such as hypertension and heart failure3,4. Indeed, manipulation of the mouse genome allowed generation of transgenic or gene-targeted models suitable for studying hypertension5,6. However, even gentle manipulation of conscious mice induces excitements that can potentially introduce artifacts during data acquisition, while use of sedation or tranquilizers profoundly affect blood pressure7. These aspects are particularly important and must be taken in consideration when attempting long-term BP monitoring.
There are several ways to record BP in mice, and a comparison between the most common techniques currently available has been discussed elsewhere8,9. However, the AHA recommendations for BP measurement concluded that intra-arterial measurement of BP are generally preferred because of their ability to directly measure BP over an extended period of time10. Radio-telemetry coupled with direct measurement of arterial pressure is the state-of-the-art method for monitoring physiological functions in awake and freely moving laboratory animals while minimizing stress and environmentally-associated artifacts9,11. Radio-telemetry offers the ability to automatically collect blood pressures, heart rate, body temperature and animal activity from multiple conscious animals.
Although this methodology is becoming very popular in many laboratories, radio-telemetry implantation in mice can be technically challenging. Here we show a step-by-step protocol that illustrates how to implant a pressure transducer in mice. The technique involves subcutaneous insertion of the probe in the mouse body, channel the catheter to the neck and forward near the aortic arch via the left carotid artery. Remotely captured data are shown live on the computer monitor.&#160;Data are also stored for &#34;off-line&#34; analysis.

<table>
	<tbody>
		<tr>
			<td>Small animal anesthesia system</td>
			<td>Kent Scientific Corp, Torrington, Connecticut, USA</td>
			<td></td>
			<td>low-Flow small animal anesthesia system</td>
		</tr>
		<tr>
			<td>Pad warmer and mouse termometer</td>
			<td>Kent Scientific Corp, Torrington, Connecticut, USA</td>
			<td></td>
			<td>allows monitoring body temperature,&#160; and homeothermic control in small animals.</td>
		</tr>
		<tr>
			<td>Binocular Microscope</td>
			<td>Kent Scientific Corp, Torrington, Connecticut, USA</td>
			<td>KSCXTS-1121</td>
			<td>binocular body with boom stand pole and top LED.</td>
		</tr>
		<tr>
			<td>Hemostat Forceps</td>
			<td>Kent Scientific Corp, Torrington, Connecticut, USA</td>
			<td>INS750451</td>
			<td>used to clamp blood vessels or tag sutures</td>
		</tr>
		<tr>
			<td>Small metal Clips, 7mm, Stainless Steel</td>
			<td>Kent Scientific Corp, Torrington, Connecticut, USA</td>
			<td>INS750344</td>
			<td>used for skin closure</td>
		</tr>
		<tr>
			<td>Betadine solution</td>
			<td>Purdue Products L.P., Stamford, CT, USA</td>
			<td>NDC-67618-150-01</td>
			<td>10% povidone iodine topical solution</td>
		</tr>
		<tr>
			<td>Normal saline solution</td>
			<td>Abott Laboratories</td>
			<td>04930-04-10</td>
			<td>Needed for preventing tissue from drying.</td>
		</tr>
		<tr>
			<td>Nair (Hair remover lotion)</td>
			<td></td>
			<td></td>
			<td>Needed for fur removal from the site of incision/surgery.</td>
		</tr>
		<tr>
			<td>Braide silk suture</td>
			<td>Teleflex Medical OEM, Coventry, Connecticut, USA</td>
			<td></td>
			<td>Size 5.0, 6.0, 7.0</td>
		</tr>
		<tr>
			<td>Ethanol</td>
			<td></td>
			<td>2716</td>
			<td>70% ethanol for disinfection</td>
		</tr>
		<tr>
			<td>Spring scissors&#160;</td>
			<td>Fine Science Tool</td>
			<td>15000-10</td>
			<td>for minor dissection</td>
		</tr>
		<tr>
			<td>Scissors (angled to side)</td>
			<td>Fine Science Tool</td>
			<td>14063-011</td>
			<td>No. 3 handle</td>
		</tr>
		<tr>
			<td>Scalpel blade</td>
			<td></td>
			<td>2976-0</td>
			<td>No. 10</td>
		</tr>
		<tr>
			<td>Forceps (curved)</td>
			<td>Fine Science Tool</td>
			<td>11150-10</td>
			<td>for holding tissue</td>
		</tr>
		<tr>
			<td>Forceps (straight)</td>
			<td>Fine Science Tool</td>
			<td>11151-10</td>
			<td>for holding tissue</td>
		</tr>
		<tr>
			<td>Needle holder&#160;</td>
			<td>Fine Science Tool</td>
			<td>12002-12</td>
			<td>for suturing</td>
		</tr>
		<tr>
			<td>Fine needle nose Forceps</td>
			<td>Fine Science Tool</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Isoflurane</td>
			<td>Henry Schein&#160;Animal Health, Melville, New York, USA</td>
			<td></td>
			<td>a general inhalation anesthetic agent</td>
		</tr>
		<tr>
			<td>Sterilizer</td>
			<td>Benchmark Scientific, 116 Corporate Blvd, South Plainfield, NJ, USA</td>
			<td>B1000</td>
			<td>sterilize surgical tools&#160;in 5-10 seconds using infrared heating</td>
		</tr>
		<tr>
			<td>Gauze Pads</td>
			<td>Johnson &#38; Johnson, New Brunswick, NJ, USA</td>
			<td>JJ8513</td>
			<td>to use for wound cleaning, prepping, scrubbin or dressing.</td>
		</tr>
		<tr>
			<td>Telemetry Device</td>
			<td>Data Sciences International, St. Paul, MN, USA</td>
			<td>DSI-PA-C10</td>
			<td>to record blood pressure in freely moving mice</td>
		</tr>
		<tr>
			<td>Telemetry receiver system&#160; coumpled with a PC</td>
			<td>Data Sciences International, St. Paul, MN, USA</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>small Tubing</td>
			<td>Instech Laboratories, Plymouth Meeting, PA USA</td>
			<td>BTPE-90</td>
			<td></td>
		</tr>
		<tr>
			<td>Vessel Cannulation Forceps, 13cm, 0.5mm OD</td>
			<td>World Precision Instruments</td>
			<td>503374</td>
			<td>special vessel cannulation forceps.</td>
		</tr>
		<tr>
			<td>Tissue adhesive</td>
			<td>3M Animal Care Products, St. Paul, MN, USA</td>
			<td>NAC No.:&#160;11380041</td>
			<td>&#160;use to close minor wounds, often eliminating the need for sutures and/or bandages</td>
		</tr>
		<tr>
			<td>Weighing scale&#160;</td>
			<td></td>
			<td>BB300</td>
			<td>&#160;Precision Analytical Laboratory Balance</td>
		</tr>
	</tbody>
</table>

long-term blood pressure measurement in freely moving mice using telemetry

During the development of new vasoactive agents, arterial blood pressure monitoring is crucial for evaluating the efficacy of the new proposed drugs. Indeed, research focusing on the discovery of new potential therapeutic targets using genetically altered mice requires a reliable, long-term assessment of the systemic arterial pressure variation. Currently, the gold standard for obtaining long-term measurements of blood pressure in ambulatory mice uses implantable radio-transmitters, which require&#160;artery cannulation. This technique eliminates the need for tethering, restraining, or anesthetizing the animals which introduce stress and artifacts during data sampling. However, arterial blood pressure monitoring in mice via catheterization can be rather challenging due to the small size of the arteries. Here we present a step-by-step guide to illustrate the crucial key passages for a successful subcutaneous implantation of radio-transmitters and carotid artery cannulation in mice. We also include examples of long-term blood pressure activity taken from freely moving mice after a period of post-surgery recovery. Following this procedure will allow reliable direct blood pressure recordings from multiple animals simultaneously.

Data can be acquired remotely by a receiver; traces are visualized on a computer screen for quality control (Figure 1a). Details such as animal ID, diastolic blood pressure and&#160;systolic blood pressure&#160;are also shown (Figure 1b). Arterial BP can be recorded continuously (24/7), or for short programmed intervals (i.e., 60 sec&#160;acquisition every hour).&#160;Data can be automatically stored in a hard disk for later analysis. Averaged BP...

Watch this Scientific Journal Video about Long-term Blood Pressure Measurement in Freely Moving Mice Using Telemetry at JoVE.com

Long-term Blood Pressure Measurement in Freely Moving Mice Using Telemetry

The goal of this protocol is to assess systemic blood pressure in conscious freely moving mice using implantable radio-telemetry devices.

During the development of new vasoactive agents, arterial blood pressure monitoring is crucial for evaluating the efficacy of the new proposed drugs. ...

long-term-blood-pressure-measurement-freely-moving-mice-using

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JoVE Journal

Medicine

18.4K Views.  University of Tennessee Health Sciences Center. The goal of this protocol is to assess systemic blood pressure in conscious freely moving mice using implantable radio-telemetry devices.

Video: Long-term Blood Pressure Measurement in Freely Moving Mice Using Telemetry

Pressure Controlled Ventilation to Induce Acute Lung Injury in Mice

Murine models are extensively used to investigate acute injuries of different organs systems (1-34). Acute lung injury (ALI), which occurs with prolonged mechanical ventilation, contributes to morbidity and mortality of critical illness, and studies on novel genetic or pharmacological targets are areas of intense investigation (1-3, 5, 8, 26, 30, 33-36). ALI is defined by the acute onset of the disease, which leads to non-cardiac pulmonary edema and subsequent impairment of pulmonary gas exchange (36). We have developed a murine model of ALI by using a pressure-controlled ventilation to induce ventilator-induced lung injury (2). For this purpose, C57BL/6 mice are anesthetized and a tracheotomy is performed followed by induction of ALI via mechanical ventilation. Mice are ventilated in a pressure-controlled setting with an inspiratory peak pressure of 45 mbar over 1 - 3 hours. As outcome parameters, pulmonary edema (wet-to-dry ratio), bronchoalveolar fluid albumin content, bronchoalveolar fluid and pulmonary tissue myeloperoxidase content and pulmonary gas exchange are assessed (2). Using this technique we could show that it sufficiently induces acute lung inflammation and can distinguish between different treatment groups or genotypes (1-3, 5). Therefore this technique may be helpful for researchers who pursue molecular mechanisms involved in ALI using a genetic approach in mice with gene-targeted deletion.

A murine model for ventilator induced lung injury is an important tool to study an acute lung injury in vivo. Here, we report an easy applicable in situ model for acute lung injury using high-pressure mechanical ventilation to induce acute failure of the lung.

Murine models are extensively used to investigate acute injuries of different organs systems (1-34). Acute lung injury (ALI), which occurs with prolonged ...

Epidural Intracranial Pressure Measurement in Rats Using a Fiber-optic Pressure Transducer

Elevated intracranial pressure (ICP) is a significant problem in several forms of ischemic brain injury including stroke, traumatic brain injury and cardiac arrest. This elevation may result in further neurological injury, in the form of transtentorial herniation1,2,3,4, midbrain compression, neurological deficit or increased cerebral infarct2,4. Current therapies are often inadequate to control elevated ICP in the clinical setting5,6,7 . Thus there is a need for accurate methods of ICP measurement in animal models to further our understanding of the basic mechanisms and to develop new treatments for elevated ICP.
In both the clinical and experimental setting ICP cannot be estimated without direct measurement. Several methods of ICP catheter insertion currently exist. Of these the intraventricular catheter has become the clinical 'gold standard' of ICP measurement in humans8. This method involves the partial removal of skull and the instrumentation of the catheter through brain tissue. Consequently, intraventricular catheters have an infection rate of 6-11%9. For this reason, subdural and epidural cannulations have become the preferred methods in animal models of ischemic injury. 
Various ICP measurement techniques have been adapted for animal models, and of these, fluid-filled telemetry catheters10 and solid state catheters are the most frequently used11,12,13,14,15. The fluid-filled systems are prone to developing air bubbles in the line, resulting in false ICP readings. Solid state probes avoid this problem (Figure 1). An additional problem is fitting catheters under the skull or into the ventricles without causing any brain injury that might alter the experimental outcomes. Therefore, we have developed a method that places an ICP catheter contiguous with the epidural space, but avoids the need to insert it between skull and brain. 
An optic fibre pressure catheter (420LP, SAMBA Sensors, Sweden) was used to measure ICP at the epidural location because the location of the pressure sensor (at the very tip of the catheter) was found to produce a high fidelity ICP signal in this model. There are other manufacturers of similar optic fibre technologies13 that may be used with our methodology. Alternative solid state catheters, which have the pressure sensor located at the side of the catheter tip, would not be appropriate for this model as the signal would be dampened by the presence of the monitoring screw. 
Here, we present a relatively simple and accurate method to measure ICP. This method can be used across a wide range of ICP related animal models.

A novel technique to record the pressures within the skull is described. The minimally invasive method uses a fibre-optic pressure sensing system to accurately measure intracranial pressure (ICP) in anaesthetized rats without causing significant brain trauma. The technique may be used in a wide range of experimental models.

Elevated intracranial pressure (ICP) is a significant problem in several forms of ischemic brain injury including stroke, traumatic brain injury and ...

Measurement of the Pressure-volume Curve in Mouse Lungs

In recent decades the mouse has become the primary animal model of a variety of lung diseases. In models of emphysema or fibrosis, the essential phenotypic changes are best assessed by measurement of the changes in lung elasticity. To best understand specific mechanisms underlying such pathologies in mice, it is essential to make functional measurements that can reflect the developing pathology. Although there are many ways to measure elasticity, the classical method is that of the total lung pressure-volume (PV) curve done over the whole range of lung volumes. This measurement has been made on adult lungs from nearly all mammalian species dating back almost 100 years, and such PV curves also played a major role in the discovery and understanding of the function of pulmonary surfactant in fetal lung development. Unfortunately, such total PV curves have not been widely reported in the mouse, despite the fact that they can provide useful information on the macroscopic effects of structural changes in the lung. Although partial PV curves measuring just the changes in lung volume are sometimes reported, without a measure of absolute volume, the nonlinear nature of the total PV curve makes these partial ones very difficult to interpret. In the present study, we describe a standardized way to measure the total PV curve. We have then tested the ability of these curves to detect changes in mouse lung structure in two common lung pathologies, emphysema and fibrosis. Results showed significant changes in several variables consistent with expected structural changes with these pathologies. This measurement of the lung PV curve in mice thus provides a straightforward means to monitor the progression of the pathophysiologic changes over time and the potential effect of therapeutic procedures.

Here we present a protocol to simply and reliably measure the lung pressure-volume curve in mice, showing that it is sufficiently sensitive to detect phenotypic parenchymal changes in two common lung pathologies, pulmonary fibrosis and emphysema. This metric provides a means to quantify the lung&#8217;s structural changes with developing pathology.

In recent decades the mouse has become the primary animal model of a variety of lung diseases. In models of emphysema or fibrosis, the essential ...

Intrauterine Telemetry to Measure Mouse Contractile Pressure In Vivo

A complex integration of molecular and electrical signals is needed to transform a quiescent uterus into a contractile organ at the end of pregnancy. Despite the discovery of key regulators of uterine contractility, this process is still not fully understood. Transgenic mice provide an ideal model in which to study parturition. Previously, the only method to study uterine contractility in the mouse was ex vivo isometric tension recordings, which are suboptimal for several reasons. The uterus must be removed from its physiological environment, a limited time course of investigation is possible, and the mice must be sacrificed. The recent development of radiometric telemetry has allowed for longitudinal, real-time measurements of in vivo intrauterine pressure in mice. Here, the implantation of an intrauterine telemeter to measure pressure changes in the mouse uterus from mid-pregnancy until delivery is described. By comparing differences in pressures between wild type and transgenic mice, the physiological impact of a gene of interest can be elucidated. This technique should expedite the development of therapeutics used to treat myometrial disorders during pregnancy, including preterm labor.

Intrauterine Telemetry to Measure Mouse Contractile Pressure In Vivo

This manuscript provides a protocol for implanting telemeters in the mouse for the purpose of measuring intrauterine pressures during pregnancy.

A complex integration of molecular and electrical signals is needed to transform a quiescent uterus into a contractile organ at the end of pregnancy. ...

Surgical Placement of Catheters for Long-term Cardiovascular Exercise Testing in Swine

This protocol describes the surgical procedure to chronically instrument swine and the procedure to exercise swine on a motor-driven treadmill. Early cardiopulmonary dysfunction is difficult to diagnose, particularly in animal models, as cardiopulmonary function is often measured invasively, requiring anesthesia. As many anesthetic agents are cardiodepressive, subtle changes in cardiovascular function may be masked. In contrast, chronic instrumentation allows for measurement of cardiopulmonary function in the awake state, so that measurements can be obtained under quiet resting conditions, without the effects of anesthesia and acute surgical trauma. Furthermore, when animals are properly trained, measurements can also be obtained during graded treadmill exercise.
Flow probes are placed around the aorta or pulmonary artery for measurement of cardiac output and around the left anterior descending coronary artery for measurement of coronary blood flow. Fluid-filled catheters are implanted in the aorta, pulmonary artery, left atrium, left ventricle and right ventricle for pressure measurement and blood sampling. In addition, a 20 G&#160;catheter is positioned in the anterior interventricular vein to allow coronary venous blood sampling.
After a week of recovery, swine are placed on a motor-driven treadmill, the catheters are connected to pressure and flow meters, and swine are subjected to a five-stage progressive exercise protocol, with each stage lasting 3 min. Hemodynamic signals are continuously recorded and blood samples are taken during the last 30 sec of each exercise stage.
The major advantage of studying chronically instrumented animals is that it allows serial assessment of cardiopulmonary function, not only at rest but also during physical stress such as exercise. Moreover, cardiopulmonary function can be assessed repeatedly during disease development and during chronic treatment, thereby increasing statistical power and hence limiting the number of animals required for a study.

Here we present a protocol to assess cardiopulmonary function in awake swine, at rest and during graded treadmill exercise. Chronic instrumentation allows for repeated hemodynamic measurements uninfluenced by cardiodepressive anesthetic agents.

This protocol describes the surgical procedure to chronically instrument swine and the procedure to exercise swine on a motor-driven treadmill. Early ...

Transdermal Measurement of Glomerular Filtration Rate in Mice

Transdermal analysis of glomerular filtration rate (GFR) is an established technique that is used to assess renal function in mouse and rat models of acute kidney injury and chronic kidney disease. The measurement system consists of a miniaturized fluorescence detector that is directly attached to the skin on the back of conscious, freely moving animals, and measures the excretion kinetics of the exogenous GFR tracer, fluorescein-isothiocyanate (FITC) conjugated sinistrin (an inulin analog). This system has been described in detail in rats. However, because of their smaller size, measurement of transcutaneous GFR in mice presents additional technical challenges. In this paper we therefore provide the first detailed practical guide to the use of transdermal GFR monitors in mice based on the combined experience of three different investigators who have been performing this assay in mice over a number of years.

Here we describe a protocol to measure glomerular filtration rate (GFR) in conscious, freely moving mice using a transdermal GFR monitor.

Transdermal analysis of glomerular filtration rate (GFR) is an established technique that is used to assess renal function in mouse and rat models of ...

Simultaneous Electrocardiography Recording and Invasive Blood Pressure Measurement in Rats

For studies related to cardiovascular physiology or pathophysiology, blood pressure (BP) and electrocardiography are basic observational parameters. Research focusing on cardiovascular disease models, potential cardiovascular therapeutic targets or pharmaceutical agents requires assessment of systemic arterial pressure and heart rhythm changes. In situations where radio telemetry systems are not available or affordable, the technique of femoral artery cannulation is an alternative way to obtain intra-arterial pressure waveform recordings and systemic BP measurements. This technique is economical and can be performed with standard equipment in animal facilities. However, invasive arterial pressure recording requires cannulation of small arteries, which can be a challenging surgical skill. Here, we present step-by-step protocols for femoral artery cannulation procedures. Key procedures include the calibration of the data acquisition system, tissue dissection and femoral artery cannulation, and setup of the arterial cannulation system for pressure recording. Surface electrocardiography recording procedures are also included. We also present examples of BP recordings from normotensive and hypertensive rats. This protocol allows reliable direct recordings of systemic BP with simultaneous electrocardiography.

Here, we describe a setup for simultaneous recording of electrocardiography and intra-arterial blood pressure (BP) in experimental rats, which can be done with standard equipment in animal facilities and can be applied to physiological or pharmacological studies to investigate pathogenic or therapeutic mechanisms in cardiovascular medicine.

For studies related to cardiovascular physiology or pathophysiology, blood pressure (BP) and electrocardiography are basic observational parameters. ...

Blood Collection Through Subclavian Vein Puncture in Mice

The mouse is the foremost mammalian model for studying human disease and human health. However, blood sample collection from mice is challenging in research work. Tail blood collection is a popular method when a small amount of blood sample is needed. Orbital artery could be considered if a large amount of blood is needed but this blood collection method has ethical issues. Formerly, we demonstrated the feasibility and safety of blood sample collection through subclavian vein puncture in rats, and here we investigate whether this method could be used in mice. We report that this method is safe and practical for blood collection in mice. Blood collection through the subclavian vein puncture in mice can be a convenient method in daily research works.

Here, we present a protocol to take blood samples from the subclavian vein of mice.

The mouse is the foremost mammalian model for studying human disease and human health. However, blood sample collection from mice is challenging in ...

Implantation of Combined Telemetric ECG and Blood Pressure Transmitters to Determine Spontaneous Baroreflex Sensitivity in Conscious Mice

Blood pressure (BP) and heart rate (HR) are both controlled by the autonomic nervous system (ANS) and are closely intertwined due to reflex mechanisms. The baroreflex is a key homeostatic mechanism to counteract acute, short-term changes in arterial BP and to maintain BP in a relatively narrow physiological range. BP is sensed by baroreceptors located in the aortic arch and carotid sinus. When BP changes, signals are transmitted to the central nervous system and are then communicated to the parasympathetic and sympathetic branches of the autonomic nervous system to adjust HR. A rise in BP causes a reflex decrease in HR, a drop in BP causes a reflex increase in HR.
Baroreflex sensitivity (BRS) is the quantitative relationship between changes in arterial BP and corresponding changes in HR. Cardiovascular diseases are often associated with impaired baroreflex function. In various studies reduced BRS has been reported in e.g., heart failure, myocardial infarction, or coronary artery disease. 
Determination of BRS requires information from both BP and HR, which can be recorded simultaneously using telemetric devices. The surgical procedure is described beginning with the insertion of the pressure sensor into the left carotid artery and positioning of its tip in the aortic arch to monitor arterial pressure followed by the subcutaneous placement of the transmitter and ECG electrodes. We also describe postoperative intensive care and analgesic management. After a two-week period of post-surgery recovery long-term ECG and BP recordings are performed in conscious and unrestrained mice. Finally, we include examples of high-quality recordings and the analysis of spontaneous baroreceptor sensitivity using the sequence method.

The baroreflex is a heart-rate regulation mechanism by the autonomic nervous system in response to blood-pressure changes. We describe a surgical technique to implant telemetry transmitters for continuous and simultaneous measurement of electrocardiogram and blood pressure in mice. This can determine spontaneous baroreflex sensitivity, an important prognostic marker for cardiovascular disease.

Blood pressure (BP) and heart rate (HR) are both controlled by the autonomic nervous system (ANS) and are closely intertwined due to reflex mechanisms. ...

Analyzing Long-Term Electrocardiography Recordings to Detect Arrhythmias in Mice

Arrhythmias are common, affecting millions of patients worldwide. Current treatment strategies are associated with significant side effects and remain ineffective in many patients. To improve patient care, novel and innovative therapeutic concepts causally targeting arrhythmia mechanisms are needed. To study the complex pathophysiology of arrhythmias, suitable animal models are necessary, and mice have been proven to be ideal model species to evaluate the genetic impact on arrhythmias, to investigate fundamental molecular and cellular mechanisms, and to identify potential therapeutic targets.
Implantable telemetry devices are among the most powerful tools available to study electrophysiology in mice, allowing continuous ECG recording over a period of several months in freely moving, awake mice. However, due to the huge number of data points (&gt;1 million QRS complexes per day), analysis of telemetry data remains challenging. This article describes a step-by-step approach to analyze ECGs and to detect arrhythmias in long-term telemetry recordings using the software, Ponemah, with its analysis modules, ECG Pro and Data Insights, developed by Data Sciences International (DSI). To analyze basic ECG parameters, such as heart rate, P wave duration, PR interval, QRS interval, or QT duration, an automated attribute analysis was performed using Ponemah to identify P, Q, and T waves within individually adjusted windows around detected R waves. 
Results were then manually reviewed, allowing adjustment of individual annotations. The output from the attribute-based analysis and the pattern recognition analysis was then used by the Data Insights module to detect arrhythmias. This module allows an automatic screening for individually defined arrhythmias within the recording, followed by a manual review of suspected arrhythmia episodes. The article briefly discusses challenges in recording and detecting ECG signals, suggests strategies to improve data quality, and provides representative recordings of arrhythmias detected in mice using the approach described above.

Here we present a step-by-step protocol for a semiautomated approach to analyze murine long-term electrocardiography (ECG) data for basic ECG parameters and common arrhythmias. Data are obtained by implantable telemetry transmitters in living and awake mice and analyzed using Ponemah and its analysis modules.

Arrhythmias are common, affecting millions of patients worldwide. Current treatment strategies are associated with significant side effects and remain ...

Long-term Blood Pressure Measurement in Freely Moving Mice Using Telemetry

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Surgical Placement of Catheters for Long-term Cardiovascular Exercise Testing in Swine

Transdermal Measurement of Glomerular Filtration Rate in Mice

Simultaneous Electrocardiography Recording and Invasive Blood Pressure Measurement in Rats

Blood Collection Through Subclavian Vein Puncture in Mice

Implantation of Combined Telemetric ECG and Blood Pressure Transmitters to Determine Spontaneous Baroreflex Sensitivity in Conscious Mice

Analyzing Long-Term Electrocardiography Recordings to Detect Arrhythmias in Mice