This study was supported by Netherlands Heart Foundation grant 2000T038 (to D.J. Duncker) grant 2000T042 (to D. Merkus), European Commission FP7-Health-2010 grant MEDIA-261409 (to D.J. Duncker and D. Merkus), Netherlands CardioVascular Research Initiative: the Dutch Heart Foundation, the Dutch Federation for University Medical Centers, the Netherlands Organisation for Health Research and Development and the Royal Netherlands Academy of Sciences &#160;CVON- ARENA CVON 2011-11 (to D.J. Duncker), CVON-PHAEDRA CVON2012-08 ( to D.&#160;Merkus) &#160;and CVON-RECONNECT CVON 2014-11 (to D.J. Duncker and D. Merkus), Sophia Foundation (to D. de Wijs-Meijler, D. Merkus and I.K.M. Reiss).

Procedures involving animal subjects have been approved by the Animal Care Committee at Erasmus Medical Center Rotterdam (NL). Swine with weights between 6 and 80 kg have been successfully instrumented using this protocol.
1. Adaptation of the Animals to Human Handling
<ol>
	<li>After arrival in the facility, house the animals solitarily but enable them to interact with each other.</li>
	<li>Accustomize swine to human handling and transportation from the animal facility to the experimental laboratory, by handling the animal at least once a day for one week.</li>
	<li>Train the animals appropriately for exercise experiments on a motor-driven treadmill by exercising them on the treadmill for a minimum of three times before surgery.</li>
	<li>Animals should be fasted O/N&#160;before surgery to prevent nausea, vomiting and thereby potential aspiration of stomach fluids.</li>
</ol>
2. Preparation for Surgery
<ol>
	<li>Sedation
	<ol>
		<li>Prepare medication for sedation in a 10 ml&#160;syringe. Premedication consists of tiletamine/zolazepam (5 mg/kg), xylazine, (2.25 mg/kg) and atropine (1 mg).</li>
		<li>Inject the medication intramuscularly in the trapezius muscle with a 19 G 1.5&#39;&#39; needle to sedate the pig.</li>
		<li>Wait for approximately 10 min and check for muscle relaxation and unconsciousness to confirm appropriate and stable level of sedation.</li>
		<li>Place a 20 G peripheral safety catheter in an ear vein for subsequent intravenous administration of anesthesia and/or fluids.</li>
	</ol>
	</li>
	<li>Intubation and Ventilation
	<ol>
		<li>Place the animal on a table and/or trolley in supine position.</li>
		<li>Open the mouth of the animal with an oral spreader.</li>
		<li>In case of insufficient relaxation of the jaws or presence of swallowing reflexes, which hinder intubation, administer thiopental (10 mg/kg) intravenously via the ear vein catheter.&#160;Alternatively, the pig could be masked with isoflurane to induce sedation.</li>
		<li>Use a conventional laryngoscope with a light and a Miller blade to allow the laryngoscopist to directly view the larynx.&#160;If there is laryngospasm, apply 2% lidocaine to the cords and larynx to reduce the spasm and allow intubation.</li>
		<li>Insert an intubating stylet into the endotracheal tube to make the tube conform better to the upper airway anatomy and pass the tube through the mouth and between the vocal cords into the trachea.</li>
		<li>Inflate the balloon cuff with a 10 ml syringe to help secure it in place, to prevent leakage of respiratory gases, and to protect the airways from possible aspiration of stomach fluid.</li>
		<li>Connect the tube to a breathing filter (heat and moisture exchanger) and to the mechanical ventilator.</li>
		<li>Place the animal on its right side on the surgical table.</li>
		<li>To achieve pO2 levels of 100-120 mmHg,&#160;ventilate the animal with a mixture of oxygen and nitrogen (1:2 v/v), using the following ventilator settings: Pressure control mode: positive end-expiratory pressure (PEEP) 4 cmH2O; peak inspiratory pressure 16 - 18 cmH2O; breathing frequency depending on the size of the animal (for a 20 kg animal, decrease frequency with increasing body weight)&#160;this should result in a tidal volume of ~10 ml/kg,&#160;monitor ventilation with capnography.</li>
		<li>Monitor temperature using a rectal thermometer and maintain temperature between 37 - 39 &#186;C using a heat lamp or heat mat.&#160;Moreover, monitor heart rate with electrocardiography.&#160;</li>
	</ol>
	</li>
	<li>Anesthesia
	<ol>
		<li>Induce and maintain anesthesia preferably by adding 2.0% of isoflurane (v/v) to the ventilation gas-mixture or alternatively by intravenous administration of fentanyl (10 &#956;g/kg/h) via the ear vein catheter.</li>
		<li>Check adequate depth of anesthesia by testing pain reflexes with a hind leg toe pinch before starting surgery. When necessary, add additional anesthesia or wait for a few minutes. Check pain reflexes regularly throughout the surgery.</li>
	</ol>
	</li>
	<li>Fluids and Antibiotics
	<ol>
		<li>Administer the first dose of amoxicillin&#160;(25 mg/kg) intravenously via the ear vein catheter.</li>
		<li>Connect a transfusion system to the ear vein catheter to enable slow infusion of glucose 10% (500 ml) during surgery.</li>
	</ol>
	</li>
	<li>Sterilization of Surgical Site
	<ol>
		<li>Shave and clean the skin of the animal over an area of approximately 25 cm width from the vertebral column all the way to the left axilla.</li>
		<li>Scrub the moisturized skin with povidone-iodine scrub (75 mg/ml) for approximately 5 min.</li>
		<li>Remove the povidone-iodine soap from the skin with wet sterile gauzes, before sterilizing the skin with povidone-iodine lotion (100 mg/ml).</li>
		<li>Cover the animal with sterile surgical drapes to reduce bacterial transfer and subsequent contamination of the surgical site.</li>
	</ol>
	</li>
</ol>
3. Surgery
<ol>
	<li>Opening the Thorax (Thoracotomy)
	<ol>
		<li>Make an incision in the skin, starting 1 cm caudal to the left inferior angle of the scapula down to the left axilla (Figure 1). Use diathermy to cauterize blood vessels in the skin to prevent excessive bleeding.</li>
		<li>Cut through the serratus muscle and pectoralis major muscle, using the cutting modality of the diathermy. Also use diathermy to cauterize blood vessels in the muscle layer to prevent excessive bleeding.</li>
		<li>Use blunt dissection to carefully divide the intercostal muscle of the fourth left intercostal space with a mosquito clamp. Now the costal surface of the left lung covered with visceral and parietal pleura should be exposed.</li>
		<li>To enter the pleural cavity, carefully pierce both layers of the pleura and tear them open.</li>
		<li>Use a thoracic retractor to separate the edges of the wound and the ribs and to forcefully drive tissues apart to obtain good exposure of the pleural cavity.</li>
		<li>Push away the left lung in the caudal direction and keep it in place with a wet gauze. Now the heart and great vessels should be clearly exposed.</li>
	</ol>
	</li>
	<li>Placement of Catheters and Flow Probes (Figure 1)
	<ol>
		<li>Use blunt dissection to remove ~2 cm2 of the surrounding connective tissue of the descending thoracic aorta.</li>
		<li>Perform a purse-string suture, consisting of three stitches, in the aortic wall with a non-absorbable USP3-0 braided silk suture (&#216;0.2&#160;mm).</li>
		<li>Penetrate the aortic vessel wall with a stainless steel 16 G needle in the middle of the purse-string suture.</li>
		<li>Insert the tip of the fluid-filled catheter (until the ring) into the aorta, pull the purse-string suture firmly together and tie the two strings of the suture.</li>
		<li>To keep the catheter in place, wind the suture 3 times around the catheter above the ring and again tie the two strings of the suture. Further secure the catheter with a new stitch approximately 1 cm cranial from the insertion place.</li>
		<li>Connect the fluid-filled catheter to the calibrated pressure transducer, which is connected to the computer, to monitor the mean arterial pressure during the surgery.&#160;Obtain an arterial blood gas to verify or adjust for correct ventilation settings.</li>
		<li>Open the pericardium with a crossed cut. Be aware to keep the phrenic nerve that runs over the pericardium intact.</li>
		<li>Identify the pulmonary artery and pull it slightly in the caudal direction with a Farabeuf retractor. Now the ascending aorta and aortic arch should be exposed. Monitor mean arterial pressure while retracting the pulmonary artery.</li>
		<li>Make a small cut (~1 cm) in the connective tissue between the ascending aorta and the pulmonary artery using Metzenbaum scissors, to be able to dissect either the ascending aorta or the pulmonary artery with a large curved mosquito clamp to place the flow probe.</li>
		<li>Place the rubber band of the flow probe around the vessel. To make this easier, place a suture through one end of the rubber band, place this suture around the vessel and pull it until the rubber band surrounds the vessel.</li>
		<li>Fix the flow probe measurement device on the rubber band. Connect the flow probe to the computer and check the cardiac output signal on the computer to confirm a correct placement of the flow probe.</li>
		<li>Place fluid-filled catheters in the pulmonary artery, right ventricle, left ventricle and left atrium at the same manner as described for the aortic fluid-filled catheter (3.2.2 - 3.2.5). Note that it is not necessary to remove connective tissue before performing a purse-string suture in these structures.</li>
		<li>Expose and dissect the proximal part of the left anterior descending coronary artery by first lifting the tissue with a forceps and making a small (2 - 3 mm) cut with Metzenbaum scissors, followed by carefully teasing the tissue away from the artery with a cotton swab. Ensure complete dissection of the coronary artery by passing a small straight angled mosquito clamp underneath.</li>
		<li>Make a stitch parallel to the anterior interventricular coronary vein with a suture, which is connected to the coronary venous catheter.</li>
		<li>Puncture the coronary vein with the 20 G needle of the coronary venous catheter and insert the cannula of the catheter intravenously.</li>
		<li>Remove the needle and secure the catheter with the already performed stitch (3.2.14). Further secure the catheter with a new stitch approximately 1cm from the place of initial puncture.</li>
		<li>Place the coronary flow probe around the previously dissected left anterior descending coronary artery. When the artery is constricted and is hardly visible, use lidocaine 10% spray to relax the vessel to get a better exposure of the vessel. Check the signal of the coronary flow on the computer to confirm a correct placement of the flow probe (Figure 2).</li>
	</ol>
	</li>
	<li>Tunneling
	<ol>
		<li>Tunnel the flow probes individually through the third left intercostal space beneath the muscle and above the rib by using a large curved mosquito clamp.</li>
		<li>Tunnel the fluid-filled catheters through either the third or the fifth left intercostal space by piercing the intercostal muscle. Clamp off the fluid-filled catheters and remove the three-way stopcock to minimize the piercing area and prevent leakage of the fluid-filled catheters during the tunneling.</li>
		<li>Fix the flow probes and the fluid-filled catheters with non-absorbable USP2-0 braided silk (&#216;0.3&#160;mm) by means of a purse string suture on the intercostal muscle. This suture also serves to prevent air leakage after re-instating negative intrathoracic pressure.</li>
		<li>Make three incisions in the skin approximately 2 cm sinister and parallel to the vertebral column, approximately 3 cm in length 3 cm apart of each other.</li>
		<li>Pierce a trochar beneath the left latissimus dorsi muscle from rostral incision site to the incisions on the back. Tunnel the flow probes and fluid catheters to the back within this trochar.</li>
		<li>Place the stopcocks on the fluid-filled catheters and remove the clamp. Withdraw blood to remove clots and air bubbles and fill the fluid-filled catheters with 1,000 IU/ml heparin. Coronary venous catheters should be filled with 5,000 IU/ml heparin.</li>
	</ol>
	</li>
	<li>Closing the Thorax
	<ol>
		<li>Make an incision with a length of approximately 1.5 cm, 8 cm caudal and parallel to the first incision.</li>
		<li>Lead the drain from the pleural cavity through the sixth intercostal muscles subcutaneously to this incision with a large curved mosquito clamp. Connect the drain to the suction device to remove any remaining fluid and reinstate negative pressure in the pleural cavity during the closing of the thorax.</li>
		<li>Relieve and inflate the lung with an end-inspiratory hold. Ensure adequate filling of the lung by visual monitoring.</li>
		<li>Close the thorax by pulling the ribs of the fourth intercostal space together at two separate sites with non-absorbable USP6 braided polyester (&#216;0.8&#160;mm).</li>
		<li>Close the serratus muscle and pectoralis major muscle with a running stitch and the skin with a running subcuticular suture using non-absorbable USP2-0 braided silk (&#216;0.3 mm)</li>
		<li>Suture the incisions on the dorsal side with non-absorbable USP2-0 braided polyester (&#216;0.3&#160;mm) between the catheters. First tie a knot directly onto the skin to close the incision, then fixate the catheters to the suture with a knot 1 cm from the skin. For the flow probes, use an absorbable USP2-0 braided polyglactin (&#216;0.3&#160;mm) suture to prevent cutting of the suture in the flow probe wire (Figure 1).</li>
		<li>Carefully remove the drain while applying pressure on the cranial side of the incision to maintain negative pressure in the pleural cavity. Close the incision with a purse string suture using non-absorbable USP2-0 braided polyester (&#216;0.3&#160;mm) and seal the wound with petroleum jelly.</li>
	</ol>
	</li>
</ol>
<img alt="Figure 1" src="/files/ftp_upload/53772/53772fig1.jpg" />
Figure 1. Overview of the Surgery. Top left panel: The sterile area of the animal, which should be shaved and sterilized lies between the bleu lines. The incision site is depicted as the red dotted line. Bottom left panel: Picture of catheters and flow probes: fluid-filled catheter (A), aorta/ pulmonary flow probe including rubber band (B), coronary venous catheter including 20 G needle (C) and the coronary flow probe (D). Top right panel: Schematic overview of placement of the catheters and flow probes. MAP, mean arterial pressure; Cor venous, coronary venous catheter; LAP, left atrial pressure; LVP left ventricular pressure; RVP, right ventricular pressure; PAP, pulmonary artery pressure; CO, cardiac output; CBF, coronary blood flow. Bottom right panel: Tunneled catheters exiting the back secured with a stitch and a knot at approximately 1 cm distance along the suture. <a href="https://www.jove.com/files/ftp_upload/53772/53772fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

<ol start="5">
	<li>Termination of anesthesia and recovery from surgery
	<ol>
		<li>Stop anesthesia when all incision sites are closed.</li>
		<li>Provide analgesia by administering buprenorphine (0.015 mg/kg) i.m. in the gracilis muscle.</li>
		<li>Stop the ventilation when the animal is breathing independently and disconnect the tracheal tube from the ventilator. Check regularly if the animal is breathing sufficiently.</li>
		<li>Place gauze pads between exteriorization sites of the catheters to absorb wound fluid.</li>
		<li>To protect the external segments of the catheters, give the animal an elastic vest and package the catheters between two pieces of artificial sheepskin.</li>
		<li>Deflate the balloon of the tracheal tube and extubate when the animal regains its swallowing reflex.</li>
		<li>Provide long-term analgesia by means of a Fentanyl slow-release patch (12 &#181;g/hr for a 20 kg pig; adjust strength according to bodyweight).&#160;Place the patch on a thin part of the skin (such as the lower abdomen) to ensure adequate delivery of analgesia.</li>
		<li>House the animal separately for the entire post-operative period. Provide a heating lamp for the first week after surgery to keep the animal warm.</li>
		<li>Supply enough fluid i.v. if the animal is not drinking independently.</li>
		<li>Flush the fluid-filled catheters daily, by first withdrawing blood to remove clots, then refilling with saline and finally with heparinized saline (1,000 - 5,000 IU/ml) to prevent blood clot formation. Take care not to infuse any air bubbles while flushing the catheters.</li>
		<li>Administer amoxicillin (25 mg/kg) i.v. daily for 6 days after surgery to prevent post-surgical infections.</li>
		<li>Allow the animal to recover for one week before starting the treadmill experiments.</li>
	</ol>
	</li>
</ol>
4. Treadmill Experiment (Figure 2)
<ol>
	<li>Flush the fluid-filled catheters as described (3.5.10) and attach the flushed catheters to the pressure transducers. Measure the rectal temperature to be able to obtain temperature corrected blood gas values.</li>
	<li>Flush the pressure transducers with saline to prevent damping of the signals due to air bubbles. Attach the pressure transducers to the elastic vest on the dorsal side.</li>
	<li>Connect the pressure transducers and flow probes to the amplifier. Start measuring in the computer program and calibrate the pressure transducers and flow probes with 0 mmHg being open to the air (and closed to animal) and 100 mmHg using a manometer.</li>
	<li>Switch the three-way stopcock in a way that the fluid catheters have an open connection with the pressure transducers. Note that the blood pressures can now be obtained. Check signals for shape and amplitude (Figure 2).</li>
	<li>If required, connect an extension line to either of the fluid catheters for sampling of mixed venous and arterial blood.</li>
	<li>Measure hemodynamics when the animal is lying as well as standing quietly on the treadmill. Average blood pressures are measured over a timeframe of 10 sec.</li>
	<li>Obtain arterial and mixed venous blood samples by first withdrawing 5 ml of blood using a 10 ml syringe so that 1 ml of pure blood can be obtained using a heparinized 1 ml syringe. For the coronary venous blood samples, a 2 ml syringe is used instead of the 10 ml syringe and withdrawal of 1 ml is sufficient to obtain pure blood.</li>
	<li>Keep the sealed 1 ml syringes on ice before processing the blood samples with a blood gas analyzer to determine the metabolic and ventilatory condition of the animal.</li>
	<li>Subject the swine to a five-stage exercise protocol on the treadmill, 3 min per speed, 1 - 5 km/hr (~85% of maximal heart rate). Obtain hemodynamics and blood gases after 1.5 - 2 min per speed on each speed as in the resting position.</li>
	<li>After the exercise protocol close the stopcocks and check if drift has occurred in the 0 mmHg calibration, make a note of this calibration. Remove the pressure transducers of the fluid-filled catheters and disconnect the flow probes.</li>
	<li>Flush the fluid-filled catheters with saline and heparin (1,000 - 5,000 IU/ml). Protect the catheters and flow probes by putting them beneath the elastic vest between two pieces of artificial sheepskin. The animal can now be returned to its cage.</li>
</ol>
<img alt="Figure 2" src="/files/ftp_upload/53772/53772fig2.jpg" /> 
Figure 2. Treadmill Experiment. Left panels: Instrumented swine on the treadmill. Fluid-filled catheters are connected to the pressure transducers, placed on the back of the swine. Top right panel: Overview of the total experimental set-up, including treadmill, amplifier and recording computer. Bottom right panel: Typical example of recorded hemodynamic data. From top to bottom; aortic pressure (AoP, blue) and left ventricular pressure (LVP, red); left atrial pressure (LAP,blue) and left ventricular pressure (red); pulmonary artery pressure (PAP, blue) and right ventricular pressure (RVP, red); aortic flow/cardiac output (AoF, blue); coronary blood flow (CBF, red). <a href="https://www.jove.com/files/ftp_upload/53772/53772fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

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H., Boomsma, F., Duncker, D. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Contribution+of+endothelin+to+coronary+vasomotor+tone+is+abolished+after+myocardial+infarction.">Contribution of endothelin to coronary vasomotor tone is abolished after myocardial infarction.</a> Am J Physiol Heart Circ Physiol. 288, H871-H880 (2005).</li><li>Haitsma, D. B., 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=Minimal+impairment+of+myocardial+blood+flow+responses+to+exercise+in+the+remodeled+left+ventricle+early+after+myocardial+infarction,+despite+significant+hemodynamic+and+neurohumoral+alterations.">Minimal impairment of myocardial blood flow responses to exercise in the remodeled left ventricle early after myocardial infarction, despite significant hemodynamic and neurohumoral alterations.</a> Cardiovasc Res. 52, 417-428 (2001).</li><li>Bender, S. B., van Houwelingen, M. J., Merkus, D., Duncker, D. J., Laughlin, M. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Quantitative+analysis+of+exercise-induced+enhancement+of+early-+and+late-systolic+retrograde+coronary+blood+flow.">Quantitative analysis of exercise-induced enhancement of early- and late-systolic retrograde coronary blood flow.</a> J Appl Physiol. 108 (3), 507-514 (2010).</li></ol>

The authors have nothing to disclose.

The present study describes the surgery for chronical instrumentation of swine as well as the protocol for exercising the instrumented swine on a motor-driven treadmill while measuring hemodynamics and taking blood samples for measurement of oxygen content in arterial, mixed venous and coronary venous blood.
Critical Steps within the Protocol 
There are several critical steps within the protocol that start already during the intubation procedure. Thiopental (2.1.5) is a respiratory depres...

Adequate cardiopulmonary function is essential to supply the body with oxygen and nutrients, particularly during conditions of increased metabolic demand such as during exercise1. The cardiopulmonary response to exercise is characterized by a number of adaptations in cardiac function, i.e., an increase in heart rate, contractility and stroke volume, and microvascular function, i.e., vasodilation in the vascular beds supplying exercising muscles as well as in the pulmonary vasculature, and vasoconstriction in the vascular beds supplying the gastrointestinal system as well as inactive muscles1. Impaired exercise capacity is an early hallmark of cardiopulmonary dysfunction, and cardiopulmonary exercise testing is used as an effective method to delineate between cardiac dysfunction, vascular dysfunction and/ or pulmonary dysfunction in patients with impaired exercise capacity2. Early cardiopulmonary dysfunction is difficult to diagnose, particularly in animal models, as cardiopulmonary function is often measured invasively, requiring anesthesia, with many anesthetic agents possessing cardiodepressive properties3.
Chronic instrumentation allows for measurement of cardiopulmonary function in the awake state, and when the animals are fully adjusted to the laboratory conditions measurements can be obtained under quiet resting conditions without the effects of anesthesia and acute surgical trauma. Furthermore, when the animals are appropriately trained, measurements can also be obtained during graded treadmill exercise4,5. More specifically, left and right ventricular function can be assessed and related to myocardial perfusion, while regulation of vasomotor tone in the coronary, systemic and pulmonary microcirculation can be determined. The use of fluid-filled catheters allows measurement of pressure as well as taking blood samples without imposing additional stress on the animals. Another advantage of studying chronically instrumented animals is that cardiopulmonary exercise testing can be repeated allowing the use of an animal as its own control, either during disease development or during chronic treatment, thereby increasing statistical power and hence limiting the number of animals required for a study.
Cardiopulmonary anatomy of swine closely resembles that of humans and it is possible to induce various forms of cardiopulmonary disease, such as diabetes 6, myocardial infarction 7, pulmonary hypertension 8,9 and pacing-induced heart failure10,11. Moreover, the size of swine allows chronic instrumentation, and repeated blood sampling of sufficient quantity to analyze not only blood gases, but also to perform neurohumoral measurements and/or to search for biomarkers of disease.
This protocol describes the surgery used to chronically instrument swine as well as the protocol for exercising the swine on a motor-driven treadmill.

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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.

Exercise up to 5 km/hr resulted in a doubling of cardiac output from 4.3 &#177; 0.3 to 8.5 &#177; 0.7 L/min which was principally accomplished by an increase in heart rate from 137 &#177; 7 to 256 &#177; 8 beats per min in combination with a small increase in stroke volume from 32 &#177; 2 to 36 &#177; 3 ml (Figure 3). The increase in stroke volume was facilitated by an increase in left ventricular contractility, as evidenced by an increase in the maximum of the first der...

Watch this Scientific Journal Video about Surgical Placement of Catheters for Long-term Cardiovascular Exercise Testing in Swine at JoVE.com

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

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 ...

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13.3K Views.  Erasmus MC. 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.

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

Minimal Invasive Surgical Procedure of Inducing Myocardial Infarction in Mice

Myocardial infarction still remains the main cause of death in western countries, despite considerable progress in the stent development area in the last decades. For clarification of the underlying mechanisms and the development of new therapeutic strategies, the availability of valid animal models are mandatory. Since we need new insights into pathomechanisms of cardiovascular diseases under in vivo conditions to combat myocardial infarction, the validity of the animal model is a crucial aspect. However, protection of animals are highly relevant in this context. Therefore, we establish a minimally invasive and simple model of myocardial infarction in mice, which assures a high reproducibility and survival rate of animals. Thus, this models fulfils the requirements of the 3R principle (Replacement, Refinement and Reduction) for animal experiments and assure the scientific information needed for further developing of therapeutical strategies for cardiovascular diseases.

A highly reproducible model for myocardial infarction in mice with minimal invasive manipulations is described. The model can be easily performed, resulting in a high reproducibility and survival rate. Thus, the described model will reduce the number of required animals as requested by the 3R principle (Replacement, Refinement and Reduction).

Myocardial infarction still remains the main cause of death in western countries, despite considerable progress in the stent development area in the last ...

Testing the Efficacy of Pharmacological Agents in a Pericardial Target Delivery Model in the Swine

To date, many pharmacological agents used to treat or prevent arrhythmias in open-heart cases create undesired systemic side effects. For example, antiarrhythmic drugs administered intravenously can produce drops in systemic pressure in the already compromised cardiac patient. While performing open-heart procedures, surgeons will often either create a small port or form a pericardial cradle to create suitable fields for operation. This access yields opportunities for target pharmacological delivery (antiarrhythmic or ischemic preconditioning agents) directly to the myocardial tissue without undesired side effects.
We have developed a swine model for testing pharmacological agents for target delivery within the pericardial fluid. While fully anesthetized, each animal was instrumented with a Swan-Ganz catheter as well as left and right ventricle pressure catheters, and pacing leads were placed in the right atrial appendage and the right ventricle. A medial sternotomy was then performed and a pericardial access cradle was created; a plunge pacing lead was placed in the left atrial appendage and a bipolar pacing lead was placed in the left ventricle. Utilizing a programmer and a cardiac mapping system, the refractory period of the atrioventricular node (AVN), atria and ventricles was determined. In addition, atrial fibrillation (AF) induction was produced utilizing a Grass stimulator and time in AF was observed. These measurements were performed prior to treatment, as well as 30 min&#160;and 60 min&#160;after pericardial treatment. Additional time points were added for selected studies. The heart was then cardiopleged and reanimated in a four chamber working mode. Pressure measurements and function were recorded for 1 hr after reanimation. This treatment strategy model allowed us to observe the effects of pharmacological agents that may decrease the incidence of cardiac arrhythmias and/or ischemic damage, during and after open-heart surgery.

We have developed a swine model for the target delivery of pharmacological agents within the pericardial space/fluid. Using this approach, the relative benefits of administered agents on induced atrial fibrillation, relative refractory periods and/or ischemic protection can be investigated.

To date, many pharmacological agents used to treat or prevent arrhythmias in open-heart cases create undesired systemic side effects. For example, ...

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.

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. ...

Tachycardia-Induced Cardiomyopathy As a Chronic Heart Failure Model in Swine

A stable and reliable model of chronic heart failure is required for many experiments to understand hemodynamics or to test effects of new treatment methods. Here, we present such a model by tachycardia-induced cardiomyopathy, which can be produced by rapid cardiac pacing in swine.
A single pacing lead is introduced transvenously into fully anaesthetized healthy swine, to the apex of the right ventricle, and fixated. Its other end is then tunneled dorsally to the paravertebral region. There, it is connected to an in-house modified heart pacemaker unit that is then implanted in a subcutaneous pocket. 
After 4 - 8 weeks of rapid ventricular pacing at rates of 200 - 240 beats/min, physical examination revealed signs of severe heart failure - tachypnea, spontaneous sinus tachycardia, and fatigue. Echocardiography and X-ray showed dilation of all heart chambers, effusions, and severe systolic dysfunction. These findings correspond well to decompensated dilated cardiomyopathy and are also preserved after the cessation of pacing.
This model of tachycardia-induced cardiomyopathy can be used for studying the pathophysiology of progressive chronic heart failure, especially hemodynamic changes caused by new treatment modalities like mechanical circulatory supports. This methodology is easy to perform and the results are robust and reproducible.

Here, we present a protocol to produce tachycardia-induced cardiomyopathy in swine. This model represents a potent way to study the hemodynamics of progressive chronic heart failure and the effects of applied treatment.

A stable and reliable model of chronic heart failure is required for many experiments to understand hemodynamics or to test effects of new treatment ...

Lentiviral Vector-mediated Gene Therapy of Hepatocytes Ex Vivo for Autologous Transplantation in Swine

Gene therapy is an ideal choice to cure many inborn errors of metabolism of the liver. Ex-vivo, lentiviral vectors have been used successfully in the treatment of many hematopoietic diseases in humans, as their use offers stable transgene expression due to the vector&#39;s ability to integrate into the host genome. This method demonstrates the application of ex vivo gene therapy of hepatocytes to a large animal model of hereditary tyrosinemia type I. This process consists of 1) isolation of primary hepatocytes from the autologous donor/recipient animal, 2) ex vivo gene delivery via hepatocyte transduction with a lentiviral vector, and 3) autologous transplant of corrected hepatocytes via portal vein injection. Success of the method generally relies upon efficient and sterile removal of the liver resection, careful handling of the excised specimen for isolation of viable hepatocytes sufficient for re-engrafting, high-percentage transduction of the isolated cells, and aseptic surgical procedures throughout to prevent infection. Technical failure at any of these steps will result in low yield of viable transduced hepatocytes for autologous transplant or infection of the donor/recipient animal. The pig model of human type 1 hereditary tyrosinemia (HT-1) chosen for this approach is uniquely amenable to such a method, as even a small percentage of engraftment of corrected cells will lead to repopulation of the liver with healthy cells based on a powerful selective advantage over native-diseased hepatocytes. Although this growth selection will not be true for all indications, this approach is a foundation for expansion into other indications and allows for manipulation of this environment to address additional diseases, both within the liver and beyond, while controlling for exposure to viral vector and opportunity for off-target toxicity and tumorigenicity.

Lentiviral Vector-mediated Gene Therapy of Hepatocytes Ex Vivo for Autologous Transplantation in Swine

This protocol is intended to describe porcine hepatocyte isolation and ex vivo gene delivery to cure models of metabolic diseases via autologous cell transplantation. Although this particular model enjoys unique advantages that favor successful therapy, the application is a relevant foundation to address additional diseases and indications.

Gene therapy is an ideal choice to cure many inborn errors of metabolism of the liver. Ex-vivo, lentiviral vectors have been used successfully in the ...

Conducting Maximal and Submaximal Endurance Exercise Testing to Measure Physiological and Biological Responses to Acute Exercise in Humans

Regular physical activity has a positive effect on human health, but the mechanisms controlling these effects remain unclear. The physiologic and biologic responses to acute exercise are predominantly influenced by the duration and intensity of the exercise regimen. As exercise is increasingly thought of as a therapeutic treatment and/or diagnostic tool, it is important that standardizable methodologies be utilized to understand the variability and to increase the reproducibility of exercise outputs and measurements of responses to such regimens. To that end, we describe two different cycling exercise regimens that yield different physiologic outputs. In a maximal exercise test, exercise intensity is continually increased with a greater workload resulting in an increasing cardiopulmonary and metabolic response (heart rate, stroke volume, ventilation, oxygen consumption and carbon dioxide production). In contrast, during endurance exercise tests, the demand is increased from that at rest, but is raised to a fixed submaximal exercise intensity resulting in a cardiopulmonary and metabolic response that typically plateaus. Along with the protocols, we provide suggestions on measuring physiologic outputs that include, but are not limited to, heart rate, slow and forced vital capacity, gas exchange metrics, and blood pressure to enable the comparison of exercise outputs between studies. Biospecimens can then be sampled to assess cellular, protein, and/or gene expression responses. Overall, this approach can be easily adapted into both short- and long-term effects of two distinct exercise regimens.

To assess the influence of exercise intensity on physiologic and biologic responses, two different exercise testing protocols were utilized. Methods outlining exercise testing on a cycle ergometer as an incremental maximal oxygen consumption test and endurance, steady state submaximal endurance test are described.

Regular physical activity has a positive effect on human health, but the mechanisms controlling these effects remain unclear. The physiologic and biologic ...

Integration of Brain Tissue Saturation Monitoring in Cardiopulmonary Exercise Testing in Patients with Heart Failure

Cerebral hypo-oxygenation during rest or exercise negatively impacts the exercise capacity of patients with heart failure with reduced ejection fraction (HF). However, in clinical cardiopulmonary exercise testing (CPET), cerebral hemodynamics is not assessed. NIRS is used to measure cerebral tissue oxygen saturation (SctO2) in the frontal lobe. This method is reliable and valid and has been utilized in several studies. SctO2 is lower during both rest and peak exercise in patients with HF than in healthy controls (66.3 &#177; 13.3% and 63.4 &#177; 13.8% vs. 73.1 &#177; 2.8% and 72 &#177; 3.2%). SctO2 at rest is significantly linearly correlated with peak VO2 (r = 0.602), oxygen uptake efficiency slope (r = 0.501), and brain natriuretic peptide (r = -0.492), all of which are recognized prognostic and disease severity markers, indicating its potential prognostic value. SctO2 is determined mainly by end-tidal CO2 pressure, mean arterial pressure, and hemoglobin in the HF population. This article demonstrates a protocol that integrates SctO2 using NIRS into incremental CPET on a calibrated bicycle ergometer.

This protocol integrated near-infrared spectroscopy into conventional cardiopulmonary exercise testing to identify the involvement of the cerebral hemodynamic response in exercise intolerance in patients with heart failure.

Cerebral hypo-oxygenation during rest or exercise negatively impacts the exercise capacity of patients with heart failure with reduced ejection fraction ...

Novel Percutaneous Approach for Deployment of 3D Printed Coronary Stenosis Implants in Swine Models of Ischemic Heart Disease

Minimally invasive methods for creating models of focal coronary narrowing in large animals are challenging. Rapid prototyping using three-dimensionally (3D) printed coronary implants can be employed to percutaneously create a focal coronary stenosis. However, reliable delivery of the implants can be difficult without the use of ancillary equipment. We describe the use of a mother-and-child coronary guide catheter for stabilization of the implant and for effective delivery of the 3D printed implant to any desired location along the length of the coronary vessel. The focal coronary narrowing was confirmed under coronary cineangiography and the functional significance of the coronary stenosis was assessed using gadolinium-enhanced first-pass cardiac perfusion MRI. We showed that reliable delivery of 3D printed coronary implants in swine models (n = 11) of ischemic heart disease can be achieved through repurposing mother-and-child coronary guide catheters. Our technique simplifies the percutaneous delivery of coronary implants to create closed-chest swine models of focal coronary artery stenosis and can be performed expeditiously, with a low procedural failure rate.

We describe a novel, cost-effective, and efficient technique for percutaneous delivery of three-dimensionally printed coronary implants to create closed-chest swine models of ischemic heart disease. The implants were fixed in place using a mother-and-child extension catheter with high success rate.

Minimally invasive methods for creating models of focal coronary narrowing in large animals are challenging. Rapid prototyping using three-dimensionally ...

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 Continuous Measurement of Renal Blood Flow in Conscious Rats

The kidneys play a crucial role in maintaining the homeostasis of body fluids. The regulation of renal blood flow (RBF) is essential to the vital functions of filtration and metabolism in kidney function. Many acute studies have been carried out in anesthetized animals to measure RBF under various conditions to determine mechanisms responsible for the regulation of kidney perfusion. However, for technical reasons, it has not been possible to measure RBF continuously (24 h/day) in unrestrained unanesthetized rats over prolonged periods. These methods allow the continuous determination of RBF over many weeks while also simultaneously recording blood pressure (BP) with implanted catheters (fluid-filled or by telemetry). RBF monitoring is carried out with rats placed in a circular servo-controlled rat cage that enables the unrestrained movement of the rat throughout the study. At the same time, the tangling of cables from the flow probe and arterial catheters is prevented. Rats are first instrumented with an ultrasonic flow probe placement on the left renal artery and an arterial catheter implanted in the right femoral artery. These are routed subcutaneously to the nape of the neck, and connected to the flowmeter and pressure transducer, respectively, to measure RBF and BP. Following surgical implantation, rats are immediately placed in the cage to recover for at least one week and stabilize the ultrasonic probe recordings. Urine collection is also feasible in this system. The surgical and post-surgical procedures for continuous monitoring are demonstrated in this protocol.

The present protocol describes a long-term continuous measurement of renal blood flow in conscious rats and simultaneously recording blood pressure with implanted catheters (fluid-filled or by telemetry).