We thank Stilla Frede and Susanne Schulz for their technical assistance. This study received no funding.

This protocol was approved by the Ethics Committee for Animal Experimentation LANUV Recklinghausen (#84-02.04.2016.A374). Generally, this procedure is performed on adult mice &#62;10 weeks of age. However, it is possible to perform this surgery on younger animals as well. Surgical tools must be sterilized before use and all steps are to be performed under aseptic conditions.
1. Induction of Anesthesia and Intubation
<ol>
	<li>Inject buprenorphine 0.1 &#181;g/g body weight intraperitoneally for pain relief. Repeat the intraperitoneal injections of 0.1 mg/kg buprenorphine every 8 h for the next three days after surgery.</li>
	<li>For induction, place the mouse into an anesthesia induction box which is connected to the vaporizer set to 3.0 Vol% of isoflurane with an oxygen flow of 1 L/min.</li>
	<li>Ensure deep narcosis by inducing a tactile stimulus. 
	NOTE: Increase Vol% of isoflurane up to 5%, if anesthesia induction fails or narcosis is not deep enough.</li>
	<li>Pinch the tail of the mouse to ensure reflex absence. In case of complete absence of reflexes, weigh the mouse for optimal ventilator setup (see 1.12).</li>
	<li>Move the mouse to a temperature-controlled operating table to maintain a body temperature of 37 &#176;C throughout the procedure.</li>
	<li>Place the nose of the mouse in a plastic cone which is connected to the anesthesia induction box to maintain narcosis.</li>
	<li>Fix the upper incisors of the mouse with a nylon suture. Fixate the extremities with an adhesive tape.</li>
	<li>Apply pressure on the hind paw with the tip of the forceps to ensure an adequate narcosis again. In the absence of a withdrawal reflex proceed with the following steps. 
	NOTE: Increase volume % of isoflurane if anesthesia induction fails or narcosis is not deep enough and wait for the absence of the withdrawal reflex.</li>
	<li>Place sterile ophthalmic lubricant on the corneas to prevent desiccation under anesthesia.&#160;</li>
	<li>Lubricate the rectal probe to avoid rectal trauma. Insert a rectal temperature probe to ensure a core temperature of 37 &#176;C.</li>
	<li>Depilate the throat and the upper chest with depilatory cream according to the manufacturer&#39;s instructions. Wipe off the cream after 1 min. If necessary, repeat this step until successful. 
	NOTE: Use cotton-tipped swabs&#160;in case of bleeding.</li>
	<li>Clean the depilated area with 70% ethanol. Then apply povidone-iodine for local skin disinfection 3 times and for at least 3 min.</li>
	<li>Adjust ventilator settings to physiological parameters. Set the respiratory rate to 150/min and tidal volumes to 8-10 &#181;L/g body weight (BW).</li>
	<li>Put on a new pair of sterile gloves. Place the mouse under a surgical microscope and place a sterile fenestrated drape over the mouse.</li>
	<li>Incise the skin at the midline about 3 mm under the mandibular down to the 2nd rib. Identify midline and connective tissues of the submandibular gland. Then use angled intracapsular forceps to gently divide the gland at the midline bluntly with two forceps and explore the tracheal muscle.</li>
	<li>Prepare the trachea gently by pulling the para-tracheal muscles apart bluntly with intracapsular forceps.</li>
	<li>Pull on the tongue with forceps to straighten the throat for easier intubation conditions and gently insert an intubation cannula (OD 1.2 mm) inside the trachea. Confirm the intubation by direct visualization of the tube inside the trachea and by checking for proper chest movement.</li>
	<li>Adjust the isoflurane concentration after intubation to 2% isoflurane with a flow of 1.0 L/min and 100% O2. 
	NOTE: If breathing movements do not stop or mice start moving, first increase the respiratory rate up to 180/min. If necessary, increase the isoflurane concentration up to 3.5% secondarily until the mouse stops breathing on its own. Evaluate leakages or inadequate filling of the vaporizer as the most common problems.</li>
	<li>Alternatively, perform intubation as suggested in the following sub steps.
	<ol>
		<li>Position the mouse on a table at a 60&#176; angle.</li>
		<li>Fixate the extremities of the mouse with adhesive tape and recline the head.</li>
		<li>Place a cold-light source directly on the skin above the larynx.</li>
		<li>Pull the tongue gently with forceps to visualize the vocal chords.</li>
		<li>Insert a plastic tube of an IV-cannula (24 G) through the vocal chords and connect the plastic tube to the ventilator settings.</li>
		<li>Connect the ventilator to the cannula to confirm intubation by synchrone chest movements. 
		NOTE: If breathing movements do not stop or mice start moving, first increase the respiratory rate up to 180 per min. If necessary, increase secondarily the isoflurane concentration up to 3.5% until the mouse stops breathing on its own. Evaluate leakages or inadequate filling of the vaporizer as the most likely problems.</li>
	</ol>
	</li>
</ol>
2. Preoperative Doppler Measurement
<ol>
	<li>Prepare both carotid arteries which lay adjunct to the trachea by gently pulling the connective tissue apart with forceps.</li>
	<li>Place the tip of the 20 MHz Doppler probe with some sterile&#160;ultrasound gel on the right and the left carotid artery at an angle less than 45&#176;.</li>
	<li>Slowly rotate the probe to move it lateral and medial to find a Doppler signal and tilt the probe to optimize the signal.</li>
	<li>Use a Doppler software to display and store flow velocities in the right and left common carotid arteries on a computer.</li>
</ol>
3. Thoracotomy
<ol>
	<li>Use a set of sterile gloves for each individual mouse to prevent surgical site infections.</li>
	<li>Expand the skin incision down to the 2nd intercostal space with a scissor.</li>
	<li>Identify the 2nd intercostal space visually by counting the ribs and then bluntly penetrate this space with intracapsular forceps. 
	NOTE: The 1st rib is located under the clavicle and is therefore not visible because the 2nd intercostal space is found between first visible rib (i.e. 2nd rib) and the 3rd rib.</li>
	<li>Open the 2nd intercostal space with the help of forceps tips and insert the retractors.</li>
	<li>Adjust the retractors with a rubber band attached to the operating table to have a clear view of the thymus.</li>
	<li>In case of bleeding use a cotton-tip and press on the superficial vessels for 2 min.</li>
</ol>
4. Banding of the Transverse Aorta
<ol>
	<li>Adjust the magnification to 200% to identify midline and connective tissues. Then use angled forceps to gently divide the thymus. Remove fatty tissue until the aortic arch can be seen clearly.</li>
	<li>Prepare a tunnel with angled tying forceps under the transverse aorta between the brachiocephalic artery and left common carotid artery (see Figure 1). Hold the end of a 6.0 suture with the help of fine forceps and pass the thread under the aortic arch.</li>
	<li>Take the thread with the second forceps from the other side of the arch.</li>
	<li>Cut off a 3-mm long piece length of a 27 G to use the needle as a the spacer for TAC-ligation for mice weighing between 19 - 25 g body weight and a 26 G needle as a spacer for mice &#62;25 g BW.</li>
	<li>Carefully place the spacer parallel to the transverse aorta.</li>
	<li>Prepare a loose double knot on the spacer and ensure optimal placement of the spacer in parallel to the aorta. Then tie the first throw and quickly perform a second contrary throw. Remove the spacer promptly.</li>
	<li>To perform sham mice for control, follow the same protocol omitting the ligation of the aorta.</li>
	<li>Close the 2nd intercostal space with a 6.0 polypropylene suture. Pay special attention to the subclavian vessels when ligating.</li>
	<li>Suture&#160;the skin using a 6.0 polypropylene suture in a continuous suture pattern.</li>
</ol>
5. Confirmation of Successful Ligation of the Transverse Aorta
<ol>
	<li>Place a 20 MHz Doppler probe on both sides of the neck at a 45&#176; angle as in section 2.</li>
	<li>Document the flow velocities on each side. 
	NOTE: A successful TAC can be validated by Doppler flow velocity as shown in Figure 2. A flow velocity ratio of 4 - 10 between the right and left carotid artery usually guarantees adequate TAC (see Figure 2).</li>
</ol>
6. Heart harvest
<ol>
	<li>Induce narcosis according to steps 1.2. and 1.3.</li>
	<li>Place the mouse in an euthanasia chamber and adjust the carbon dioxide flow rate to displace 10 - 30% of the volume/minute.</li>
	<li>Fixate the mouse on a surgery table. Open the abdomen with scissors and harvest blood from the inferior vena cava with a cannula for further analysis.</li>
	<li>Cut the diaphragm and the sternal bone with strong scissors and remove the heart.</li>
	<li>Remove all arterial and connective tissue before weighing the heart.</li>
	<li>Separate the right ventricle and the septum from the left ventricle and weigh both samples.</li>
	<li>Freeze both tissue samples in liquid nitrogen.</li>
</ol>

<ol>
	<li>Tarnavski, O. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mouse+surgical+models+in+cardiovascular+research.">Mouse surgical models in cardiovascular research.</a> Methods Mol Biol Clifton NJ. 573, 115-137 (2009).</li><li>Tarnavski, O., McMullen, J. R., Schinke, M., Nie, Q., Kong, S., Izumo, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mouse+cardiac+surgery:+comprehensive+techniques+for+the+generation+of+mouse+models+of+human+diseases+and+their+application+for+genomic+studies.">Mouse cardiac surgery: comprehensive techniques for the generation of mouse models of human diseases and their application for genomic studies.</a> Physiol Genomics. 16 (3), 349-360 (2004).</li><li>Rockman, H. A., 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=Segregation+of+atrial-specific+and+inducible+expression+of+an+atrial+natriuretic+factor+transgene+in+an+in+vivo+murine+model+of+cardiac+hypertrophy.">Segregation of atrial-specific and inducible expression of an atrial natriuretic factor transgene in an in vivo murine model of cardiac hypertrophy.</a> Proc Natl Acad Sci U S A. 88 (18), 8277-8281 (1991).</li><li>deAlmeida, A. C., van Oort, R. J., Wehrens, X. H. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Transverse+Aortic+Constriction+in+Mice.">Transverse Aortic Constriction in Mice.</a> J Vis Exp JoVE. (38), (2010).</li><li>Grossman, W., Jones, D., McLaurin, L. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Wall+stress+and+patterns+of+hypertrophy+in+the+human+left+ventricle.">Wall stress and patterns of hypertrophy in the human left ventricle.</a> J Clin Invest. 56 (1), 56-64 (1975).</li><li>Hartley, C. J., Reddy, A. K., Madala, S., Michael, L. H., Entman, M. L., Taffet, G. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Doppler+estimation+of+reduced+coronary+flow+reserve+in+mice+with+pressure+overload+cardiac+hypertrophy.">Doppler estimation of reduced coronary flow reserve in mice with pressure overload cardiac hypertrophy.</a> Ultrasound Med Biol. 34 (6), 892-901 (2008).</li><li>Reddy, A. K., 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=Pulsed+Doppler+signal+processing+for+use+in+mice:+applications.">Pulsed Doppler signal processing for use in mice: applications.</a> IEEE Trans Biomed Eng. 52 (10), 1771-1783 (2005).</li><li>Shioura, K. M., Geenen, D. L., Goldspink, P. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Assessment+of+cardiac+function+with+the+pressure-volume+conductance+system+following+myocardial+infarction+in+mice.">Assessment of cardiac function with the pressure-volume conductance system following myocardial infarction in mice.</a> Am J Physiol - Heart Circ Physiol. 293 (5), H2870-H2877 (2007).</li><li>Zhang, B., Davis, J. P., Ziolo, M. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cardiac+Catheterization+in+Mice+to+Measure+the+Pressure+Volume+Relationship:+Investigating+the+Bowditch+Effect.">Cardiac Catheterization in Mice to Measure the Pressure Volume Relationship: Investigating the Bowditch Effect.</a> JoVE J Vis Exp. (100), e52618 (2015).</li><li>Barrick, C. J., Rojas, M., Schoonhoven, R., Smyth, S. S., Threadgill, D. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cardiac+response+to+pressure+overload+in+129S1/SvImJ+and+C57BL/6J+mice:+temporal-+and+background-dependent+development+of+concentric+left+ventricular+hypertrophy.">Cardiac response to pressure overload in 129S1/SvImJ and C57BL/6J mice: temporal- and background-dependent development of concentric left ventricular hypertrophy.</a> Am J Physiol Heart Circ Physiol. 292 (5), H2119-H2130 (2007).</li><li>Patten, R. D., Hall-Porter, M. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Small+animal+models+of+heart+failure:+development+of+novel+therapies,+past+and+present.">Small animal models of heart failure: development of novel therapies, past and present.</a> Circ Heart Fail. 2 (2), 138-144 (2009).</li><li>Zaw, A. M., Williams, C. M., Law, H. K. W., Chow, B. K. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Minimally+Invasive+Transverse+Aortic+Constriction+in+Mice.">Minimally Invasive Transverse Aortic Constriction in Mice.</a> J Vis Exp JoVE. (121), (2017).</li><li>Tavakoli, R., Nemska, S., Jamshidi, P., Gassmann, M., Frossard, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Technique+of+Minimally+Invasive+Transverse+Aortic+Constriction+in+Mice+for+Induction+of+Left+Ventricular+Hypertrophy.">Technique of Minimally Invasive Transverse Aortic Constriction in Mice for Induction of Left Ventricular Hypertrophy.</a> J Vis Exp. (127), (2017).</li><li>Kim, S. -. C., Boehm, O., Meyer, R., Hoeft, A., Kn&#252;fermann, P., Baumgarten, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+Murine+Closed-chest+Model+of+Myocardial+Ischemia+and+Reperfusion.">A Murine Closed-chest Model of Myocardial Ischemia and Reperfusion.</a> J Vis Exp JoVE. (65), (2012).</li><li>Veldhuizen, R. A., Slutsky, A. S., Joseph, M., McCaig, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Effects+of+mechanical+ventilation+of+isolated+mouse+lungs+on+surfactant+and+inflammatory+cytokines.">Effects of mechanical ventilation of isolated mouse lungs on surfactant and inflammatory cytokines.</a> Eur Respir J. 17 (3), 488-494 (2001).</li></ol>

The authors have nothing to disclose.

The rapid onset of hypertension due to TAC differs from clinically relevant hypertrophy caused by aortic stenosis or hypertension. Nevertheless, the use of small animal models to induce heart failure has many advantages and is, therefore, chosen by many investigators11. This closed chest-model improves the already existing models of the surgical technique to induce transverse aortic constriction in mice4.
The most critical step is the passage und...

Mouse models are often used to mimic human diseases1. Transverse aortic constriction (TAC) is used to induce pressure overload and left ventricular hypertrophy2. The open-chest TAC model in mice was validated by Rockman et al.3 and the surgical procedure is described in detail by DeAlmeida et al.4. Banding of the transverse aorta is more favorable in comparison to abdominal aortic constriction because a larger portion of the circulation can compensate negative effects of this latter procedure2.
The banding of the transverse aorta leads to an increased arterial pressure in the ascending aorta and brachiocephalic artery but leaves sufficient perfusion of the organs via the distal vessels (i.e. the left common carotid artery, the left subclavian artery, and descending aorta). This leads to an increased cardiac afterload and an elevated cardiac wall stress. The wall stress subsequently decreases due to fiber thickening5. The chronic change in cardiac hemodynamics results in maladaptation and dilatation of the left ventricle. This way the TAC creates a reproducible model of cardiac hypertrophy eventually leading to heart failure.
The standard procedure for TAC as described by DeAlmeide et al.4 approaches the aortic arch via a partial upper thoracotomy via dissection of the ribs or the sternum and entering the mediastinum as well as the pleural cavity. This allows for a good view of the aortic arch and its side branches. Unfortunately, the dissected ribs cannot be reattached, which leaves them floating freely and thereby altering the breathing dynamics.
We, therefore, established a minimally invasive closed-chest approach to the aortic arch using a lateral surgical approach via the 2nd intercostal space. The greatest advantage of this model is the ability to perform TAC without even cutting through the ribs. The surgical trauma is limited to the incision of the skin and the dissection of the intercostal muscles. This procedure minimizes the trauma itself and helps to maintain adequate chest stability.
Here we describe a detailed step-by-step procedure to perform TAC surgery in mice without performing the total or the upper thoracotomy. High frequency Doppler was used to ensure the success of TAC as previously described 6,7.

<table border="0" cellpadding="0" cellspacing="0" width="1303">
	<tbody>
		<tr height="20">
			<td height="20" style="height:20px;width:312px;">Pressure-volume catheter</td>
			<td style="width:243px;">Millar Instruments, USA</td>
			<td style="width:591px;">SPR-839</td>
			<td style="width:159px;"></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Mouse ventilator</td>
			<td>Harvard Apparatus GmbH, Germany</td>
			<td>Minivent - TYPE 845</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Mouse ventilator</td>
			<td>Harvard Apparatus GmbH, Germany</td>
			<td>Y-connection with intubation cannula OD 1.2mm 73-2844</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Vaporizer</td>
			<td>Dr&#228;ger Medical AG&#38;CO.KG, Germany</td>
			<td>19.3 Isofluran-Vaporizer (a newer version is available under catalog number&#160; D-877-2010)</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Microscope&#160;</td>
			<td>Leica Microsystems, Germany</td>
			<td>MZ 7.5</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Light source&#160;</td>
			<td>Schott AG, Germany</td>
			<td>KL 1500 LCD</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">6-0 Prolene</td>
			<td>Ethicon, USA</td>
			<td>Polypropylene suture BV-1 9.3 mm 3/8c</td>
			<td>suture for surgery</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Seraflex</td>
			<td>Serag Wiessner, Germany</td>
			<td>USP 5/0 schwarz;&#160; IC108000&#160;</td>
			<td>suture for constriction</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Homoeothermic Controlled Operating Tables</td>
			<td>Harvard Apparatus GmbH, Germany</td>
			<td>Typ 872/3 HT with tripod stand and homoeothermic controller Type 874; 73-4233</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Flexible Rectal Probe</td>
			<td>Harvard Apparatus GmbH, Germany</td>
			<td>1.6 mm OD; 55-7021</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Doppler Signal Visualisation Instrument</td>
			<td>Indus Instruments, USA</td>
			<td>Doppler Signal Processing Workstation (DSWP) with 20MHz Pulsed Doppler Module</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Doppler Probe</td>
			<td>Indus Instruments, USA</td>
			<td>20MHz Tubing-mounted Probe</td>
			<td></td>
		</tr>
	</tbody>
</table>

a closed-chest model to induce transverse aortic constriction in mice

Research on cardiac hypertrophy and heart failure is frequently based on pressure overload mouse models induced by TAC. The standard procedure is to perform a partial thoracotomy to visualize the transverse aortic arch. However, the surgical trauma caused by the thoracotomy in open-chest models changes the respiratory physiology as the ribs are dissected and left unattached after chest closure. To prevent this, we established a minimally invasive, closed chest approach via lateral thoracotomy. Herein we approach the aortic arch via the 2nd intercostal space without entering the chest cavities, leaving the mouse with a less traumatic injury to recover from. We perform this operation using standard laboratory settings for open chest TAC procedures with equal survival rates. Apart from maintaining physiological breathing patterns due to the closed chest approach, the mice seem to benefit by showing rapid recovery, as the less invasive technique appears to facilitate a fast healing process and to reduce immune response after trauma.

A successful TAC guarantees the induction of pressure overload and left ventricular hypertrophy. An ad hoc validation of pressure overload can be achieved using Doppler flow velocity measurement as shown in Figure 2. While preoperative blood flow velocity is equal in both carotid arteries, TAC causes an augmented blood velocity in the right carotid artery due to elevated pressure in the left ventricle and aorta while causing post-stenotic attenuated blood flo...

Watch this Scientific Journal Video about A Closed-chest Model to Induce Transverse Aortic Constriction in Mice at JoVE.com

A Closed-chest Model to Induce Transverse Aortic Constriction in Mice

Here, we present a protocol of Transverse Aortic constriction (TAC) via a lateral thoracotomy. This technique is a minimally invasive, closed chest surgical procedure aiming to simulate pressure overload and heart failure in mice utilizing standard TAC laboratory settings.

Research on cardiac hypertrophy and heart failure is frequently based on pressure overload mouse models induced by TAC. The standard procedure is to ...

a-closed-chest-model-to-induce-transverse-aortic-constriction-in-mice

Universidad de Cartagena UDEC

Research

JoVE Journal

Medicine

18.8K Views.  University Hospital Bonn. Here, we present a protocol of Transverse Aortic constriction (TAC) via a lateral thoracotomy. This technique is a minimally invasive, closed chest surgical procedure aiming to simulate pressure overload and heart failure in mice utilizing standard TAC laboratory settings.

Video: A Closed-chest Model to Induce Transverse Aortic Constriction in Mice

The CYP2D6 Animal Model: How to Induce Autoimmune Hepatitis in Mice

Autoimmune hepatitis is a rare but life threatening autoimmune disease of the liver of unknown etiology1,2. In the past many attempts have been made to generate an animal model that reflects the characteristics of the human disease 3-5. However, in various models the induction of disease was rather complex and often hepatitis was only transient3-5. Therefore, we have developed a straightforward mouse model that uses the major human autoantigen in type 2 autoimmune hepatitis (AIH-2), namely hCYP2D6, as a trigger6. Type 1 liver-kidney microsomal antibodies (LKM-1) antibodies recognizing hCYP2D6 are the hallmark of AIH-27,8. Delivery of hCYP2D6 into wildtype FVB or C57BL/6 mice was by an Adenovirus construct (Ad-2D6) that ensures a direct delivery of the triggering antigen to the liver. Thus, the ensuing local inflammation generates a fertile field9 for the subsequent development of autoimmunity. A combination of intravenous and intraperitoneal injection of Ad-2D6 is the most effective route to induce a long-lasting autoimmune damage to the liver (section 1). Here we provide a detailed protocol on how autoimmune liver disease is induced in the CYP2D6 model and how the different aspects of liver damage can be assessed. First, the serum levels of markers indicating hepatocyte destruction, such as aminotransferases, as well as the titers of hCYP2D6 antibodies are determined by sampling blood retroorbitaly (section 2). Second, the hCYP2D6-specific T cell response is characterized by collecting lymphocytes from the spleen and the liver. In order to obtain pure liver lymphocytes, the livers are perfused by PBS via the portal vein (section 3), digested in collagen and purified over a Percoll gradient (section 4). The frequency of hCYP2D6-specific T cells is analyzed by stimulation with hCYP2D6 peptides and identification of IFN&gamma;-producing cells by flow cytometry (section 5). Third, cellular infiltration and fibrosis is determined by immunohistochemistry of liver sections (section 6). Such analysis regimen has to be conducted at several times after initiation of the disease in order to prove the chronic nature of the model. The magnitude of the immune response characterized by the frequency and activity of hCYP2D6-specific T and/or B cells and the degree of the liver damage and fibrosis have to be assessed for a subsequent evaluation of possible treatments to prevent, delay or abrogate the autodestructive process of the liver.

Infection of mice with an Adenovirus expressing the major human autoantigen cytochrome P450 2D6 (hCYP2D6) recognized by sera of patients suffering from type 2 autoimmune hepatitis results in a persistent form of autoimmune-mediated liver disease characterized by extensive hepatitis, fibrosis and generation of a CYP2D6-specific immune response.

Autoimmune hepatitis is a rare but life threatening autoimmune disease of the liver of unknown etiology1,2. In the past many attempts have been made to ...

In vitro Measurements of Tracheal Constriction Using Mice

Transgenic and knockout mice have been powerful tools for the investigation of the physiology and pathophysiology of airways1,2. In vitro tensometry of isolated tracheal preparations has proven to be a useful assay of airway smooth muscle (ASM) contractile response in genetically modified mice. These in vitro tracheal preparations are relatively simple, provide a robust response, and retain both functional cholinergic nerve endings and muscle responses, even after long incubations.
Tracheal tensometry also provides a functional assay to study a variety of second messenger signaling pathways that affect contraction of smooth muscle. Contraction in trachea is primarily mediated by parasympathetic, cholinergic nerves that release acetylcholine onto ASM (Figure 1). The major ASM acetylcholine receptors are muscarinic M2 and M3 which are Gi/o and Gq coupled receptors, respectively3,4,5. M3 receptors evoke contraction by coupling to Gq to activate phospholipase C, increase IP3 production and IP3-mediated calcium release from the sarcoplasmic reticulum3,6,7. M2/Gi/o signaling is believed to enhance contractions by inhibition of adenylate cyclase leading to a decrease in cAMP levels5,8,9,10. These pathways constitute the so called "pharmaco-contraction coupling" of airway smooth muscle11. In addition, cholinergic signaling through M2 receptors (and modulated by M3 signaling) involves pathways that depolarize the ASM which in turn activate L-type, voltage-dependent calcium channels (Figure 1) and calcium influx (so called "excitation-contraction coupling")4,7. More detailed reviews on signaling pathways controlling airway constriction can be found4,12. The above pathways appear to be conserved between mice and other species. However, mouse tracheas differ from other species in some signaling pathways. Most prominent is their lack of contractile response to histamine and adenosine13,14, both well-known ASM modulators in humans and other species5,15.
Here we present protocols for the isolation of murine tracheal rings and the in vitro measurement of their contractile output. Included are descriptions of the equipment configuration, trachea ring isolation and contractile measurements. Examples are given for evoking contractions indirectly using high potassium stimulation of nerves and directly by depolarization of ASM muscle to activate voltage-dependent calcium influx (1. high K+, Figure 1). In addition, methods are presented for stimulations of nerves alone using electric field stimulation (2. EFS, Figure 1), or for direct stimulation of ASM muscle using exogenous neurotransmitter applied to the bath (3. exogenous ACH, Figure 1). This flexibility and ease of preparation renders the isolated trachea ring model a robust and functional assay for a number of signaling cascades involved in airway smooth muscle contraction.

In vitro Measurements of Tracheal Constriction Using Mice

Transgenic mice have been extremely useful in ascribing physiological function to genes. As such, research in general, and functional studies of airway, in particular, have undergone a remarkable shift toward murine models. Here we provide protocols for in vitro trachea constriction studies to evaluate smooth muscle function in murine airway.

Transgenic and knockout mice have been powerful tools for the investigation of the physiology and pathophysiology of airways1,2. In vitro tensometry of ...

A Murine Closed-chest Model of Myocardial Ischemia and Reperfusion

Surgical trauma by thoracotomy in open-chest models of coronary ligation induces an immune response which modifies different mechanisms involved in ischemia and reperfusion. Immune response includes cytokine expression and release or secretion of endogenous ligands of innate immune receptors. Activation of innate immunity can potentially modulate infarct size. We have modified an existing murine closed-chest model using hanging weights which could be useful for studying myocardial pre- and postconditioning and the role of innate immunity in myocardial ischemia and reperfusion. This model allows animals to recover from surgical trauma before onset of myocardial ischemia.
Volatile anesthetics have been intensely studied and their preconditioning effect for the ischemic heart is well known. However, this protective effect precludes its use in open chest models of coronary artery ligation. Thus, another advantage could be the use of the well controllable volatile anesthetics for instrumentation in a chronic closed-chest model, since their preconditioning effect lasts up to 72 hours. Chronic heart diseases with intermittent ischemia and multiple hit models are other possible applications of this model.
For the chronic closed-chest model, intubated and ventilated mice undergo a lateral blunt thoracotomy via the 4th intercostal space. Following identification of the left anterior descending a ligature is passed underneath the vessel and both suture ends are threaded through an occluder. Then, both suture ends are passed through the chest wall, knotted to form a loop and left in the subcutaneous tissue. After chest closure and recovery for 5 days, mice are anesthetized again, chest skin is reopened and hanging weights are hooked up to the loop under ECG control.
At the end of the ischemia/reperfusion protocol, hearts can be stained with TTC for infarct size assessment or undergo perfusion fixation to allow morphometric studies in addition to histology and immunohistochemistry.

Surgical trauma induces an inflammatory response. Cytokines and endogenous ligands are known to modulate myocardial infarct size following ischemia and reperfusion. We present a modified closed-chest model of murine ischemia and reperfusion using hanging weights to minimize effects of thoracotomy.

Surgical trauma by thoracotomy in open-chest models of coronary ligation induces an immune response which modifies different mechanisms involved in ...

Ascending Aortic Constriction in Rats for Creation of Pressure Overload Cardiac Hypertrophy Model

Ascending aortic constriction is the most common and successful surgical model for creating pressure overload induced cardiac hypertrophy and heart failure. Here, we describe a detailed surgical procedure for creating pressure overload and cardiac hypertrophy in rats by constriction of the ascending aorta using a small metallic clip. After anesthesia, the trachea is intubated by inserting a cannula through a half way incision made between two cartilage rings of trachea. Then a skin incision is made at the level of the second intercostal space on the left chest wall and muscle layers are cleared to locate the ascending portion of aorta. The ascending aorta is constricted to 50&#8211;60% of its original diameter by application of a small sized titanium clip. Following aortic constriction, the second and third ribs are approximated with prolene sutures. The tracheal cannula is removed once spontaneous breathing was re-established. The animal is allowed to recover on the heating pad by gradually lowering anesthesia. The intensity of pressure overload created by constriction of the ascending aorta is determined by recording the pressure gradient using trans-thoracic two dimensional Doppler-echocardiography. Overall this protocol is useful to study the remodeling events and contractile properties of the heart during the gradual onset and progression from compensated cardiac hypertrophy to heart failure stage.

We describe a stepwise procedure for creating pressure overload and left ventricular hypertrophy in Wistar rats by constriction of the ascending aorta using a small metallic clip. This model is extensively used for studying remodeling changes during cardiac hypertrophy and for identifying and evaluating strategies for regression of such changes.

Ascending aortic constriction is the most common and successful surgical model for creating pressure overload induced cardiac hypertrophy and heart ...

A Rat Model of Ventricular Fibrillation and Resuscitation by Conventional Closed-chest Technique

A rat model of electrically-induced ventricular fibrillation followed by cardiac resuscitation using a closed chest technique that incorporates the basic components of cardiopulmonary resuscitation in humans is herein described. The model was developed in 1988 and has been used in approximately 70 peer-reviewed publications examining a myriad of resuscitation aspects including its physiology and pathophysiology, determinants of resuscitability, pharmacologic interventions, and even the effects of cell therapies. The model featured in this presentation includes: (1) vascular catheterization to measure aortic and right atrial pressures, to measure cardiac output by thermodilution, and to electrically induce ventricular fibrillation; and (2) tracheal intubation for positive pressure ventilation with oxygen enriched gas and assessment of the end-tidal CO2. A typical sequence of intervention entails: (1) electrical induction of ventricular fibrillation, (2) chest compression using a mechanical piston device concomitantly with positive pressure ventilation delivering oxygen-enriched gas, (3) electrical shocks to terminate ventricular fibrillation and reestablish cardiac activity, (4) assessment of post-resuscitation hemodynamic and metabolic function, and (5) assessment of survival and recovery of organ function. A robust inventory of measurements is available that includes &#8211; but is not limited to &#8211; hemodynamic, metabolic, and tissue measurements. The model has been highly effective in developing new resuscitation concepts and examining novel therapeutic interventions before their testing in larger and translationally more relevant animal models of cardiac arrest and resuscitation.

This article describes a rat model of electrically-induced ventricular fibrillation and resuscitation by chest compression, ventilation, and delivery of electrical shocks that simulates an episode of sudden cardiac arrest and conventional cardiopulmonary resuscitation. The model enables gathering insights on the pathophysiology of cardiac arrest and exploration of new resuscitation strategies.

A rat model of electrically-induced ventricular fibrillation followed by cardiac resuscitation using a closed chest technique that incorporates the basic ...

A Magnetic Microbead Occlusion Model to Induce Ocular Hypertension-Dependent Glaucoma in Mice

The use of rodent models of glaucoma has been essential to understand the molecular mechanisms that underlie the pathophysiology of this multifactorial neurodegenerative disease. With the advent of numerous transgenic mouse lines, there is increasing interest in inducible murine models of ocular hypertension. Here, we present an occlusion model of glaucoma based on the injection of magnetic microbeads into the anterior chamber of the eye using a modified microneedle with a facetted bevel. The magnetic microbeads are attracted to the iridocorneal angle using a handheld magnet to block the drainage of aqueous humour from the anterior chamber. This disruption in aqueous dynamics results in a steady elevation of intraocular pressure, which subsequently leads to the loss of retinal ganglion cells, as observed in human glaucoma patients. The microbead occlusion model presented in this manuscript is simple compared to other inducible models of glaucoma and also highly effective and reproducible. Importantly, the modifications presented here minimize common issues that often arise in occlusion models. First, the use of a bevelled glass microneedle prevents backflow of microbeads and ensures that minimal damage occurs to the cornea during the injection, thus reducing injury-related effects. Second, the use of magnetic microbeads ensures the ability to attract most beads to the iridocorneal angle, effectively reducing the number of beads floating in the anterior chamber avoiding contact with other structures (e.g., iris, lens). Lastly, the use of a handheld magnet allows flexibility when handling the small mouse eye to efficiently direct the magnetic microbeads and ensure that there is little reflux of the microbeads from the eye when the microneedle is withdrawn. In summary, the microbead occlusion mouse model presented here is a powerful investigative tool to study neurodegenerative changes that occur during the onset and progression of glaucoma.

Here, we present a protocol to induce ocular hypertension in the murine eye that results in the loss of retinal ganglion cells as observed in glaucoma. Magnetic microbeads are injected into the anterior chamber and attracted to the iridocorneal angle using a magnet to block the outflow of aqueous humour.

The use of rodent models of glaucoma has been essential to understand the molecular mechanisms that underlie the pathophysiology of this multifactorial ...

Assessment of Cardiac Morphological and Functional Changes in Mouse Model of Transverse Aortic Constriction by Echocardiographic Imaging

Transverse aortic constriction (TAC) in mice has been used as a valuable model to study mechanisms of cardiac hypertrophy and heart failure1. A reliable noninvasive method is essential to assess real-time cardiac morphological and functional changes in animal models of heart disease. Transthoracic echocardiography represents an important tool for noninvasive assessment of cardiac structure and function2. Here we used a high-resolution ultrasound imaging system to monitor myocardial remodeling and heart failure progression over time in a mouse model of TAC. B-mode, M-mode, and Doppler imaging were used to precisely assess cardiac hypertrophy, ventricular dilatation, and functional deterioration in mice following TAC. Color and pulse wave (PW) Doppler imaging was used to noninvasively measure pressure gradient across the aortic constriction created by TAC and to assess transmitral blood flow in mice. Thus transthoracic echocardiographic imaging provides comprehensive noninvasive measurements of cardiac dimensions and function in mouse models of heart disease.

The goal of this protocol is to noninvasively assess cardiac structural and functional changes in a mouse model of heart disease created by transverse aortic constriction, using B- and M-mode echocardiography and color/pulse wave Doppler imaging.

Transverse aortic constriction (TAC) in mice has been used as a valuable model to study mechanisms of cardiac hypertrophy and heart failure1. A reliable ...

Minimally Invasive Transverse Aortic Constriction in Mice

Minimally invasive transverse aortic constriction (MTAC) is a more desirable method for the constriction of the transverse aorta in mice than standard open-chest transverse aortic constriction (TAC). Although transverse aortic constriction is a highly functional method for the induction of high pressure in the left ventricle, it is a more difficult and lengthy procedure due to its use of artificial ventilation with tracheal intubation. TAC is oftentimes also less survivable, as the newer method, MTAC, neither requires the cutting of the ribs and intercostal muscles nor tracheal intubation with a ventilation setup. In MTAC, as opposed to a thoracotomy to access to the chest cavity, the aortic arch is reached through a midline incision in the anterior neck. The thyroid is pulled back to reveal the sternal notch. The sternum is subsequently cut down to the second rib level, and the aortic arch is reached simply by separating the connective tissues and thymus. From there, a suture can be wrapped around the arch and tied with a spacer, and then the sternal cut and skin can be closed. MTAC is a much faster and less invasive way to induce left ventricular hypertension and enables the possibility for high-throughput studies. The success of the constriction can be verified using high-frequency trans-thoracic echocardiography, particularly color Doppler and pulsed-wave Doppler, to determine the flow velocities of the aortic arch and left and right carotid arteries, the dimension of the blood vessels, and the left ventricular function and morphology. A successful constriction will also trigger significant histopathological changes, such as cardiac muscle cell hypertrophy with interstitial and perivascular fibrosis. Here, the procedure of MTAC is described, demonstrating how the resulting flow changes in the carotid arteries can be examined with echocardiography, gross morphology, and histopathological changes in the heart.

Minimally invasive transverse aortic constriction (MTAC) conserves the essentials of regular transverse aortic constriction (TAC) while eliminating the use of a ventilator with tracheal intubation. It proves to be a highly desirable method for high-throughput studies on left ventricular overload, particularly in translational studies.

Minimally invasive transverse aortic constriction (MTAC) is a more desirable method for the constriction of the transverse aorta in mice than standard ...

Technique of Minimally Invasive Transverse Aortic Constriction in Mice for Induction of Left Ventricular Hypertrophy

Transverse aortic constriction (TAC) in mice is one of the most commonly used surgical techniques for experimental investigation of pressure overload-induced left ventricular hypertrophy (LVH) and its progression to heart failure. In the majority of the reported investigations, this procedure is performed with intubation and ventilation of the animal which renders it demanding and time-consuming and adds to the surgical burden to the animal. The aim of this protocol is to describe a simplified technique of minimally invasive TAC without intubation and ventilation of mice. Critical steps of the technique are emphasized in order to achieve low mortality and high efficiency in inducing LVH.
Male C57BL/6 mice (10-week-old, 25-30 g, n=60) were anesthetized with a single intraperitoneal injection of a mixture of ketamine and xylazine. In a spontaneously breathing animal following a 3-4 mm upper partial sternotomy, a segment of 6/0 silk suture threaded through the eye of a ligation aid was passed under the aortic arch and tied over a blunted 27-gauge needle. Sham-operated animals underwent the same surgical preparation but without aortic constriction. The efficacy of the procedure in inducing LVH is attested by a significant increase in the heart/body weight ratio. This ratio is obtained at days 3, 7, 14 and 28 after surgery (n = 6 - 10 in each group and each time point). Using our technique, LVH is observed in TAC compared to sham animals from day 7 through day 28. Operative and late (over 28 days) mortalities are both very low at 1.7%.
In conclusion, our cost-effective technique of minimally invasive TAC in mice carries very low operative and post-operative mortalities and is highly efficient in inducing LVH. It simplifies the operative procedure and reduces the strain put on the animal. It can be easily performed by following the critical steps described in this protocol.

The aim of this protocol is to describe step-by-step the technique of minimally invasive transverse aortic constriction (TAC) in mice. By elimination of intubation and ventilation which are mandatory for the commonly used standard procedure, minimally invasive TAC simplifies the operative procedure and reduces the strain put on the animal.

Transverse aortic constriction (TAC) in mice is one of the most commonly used surgical techniques for experimental investigation of pressure ...

Improvement of a Closed Chest Porcine Myocardial Infarction Model by Standardization of Tissue and Blood Sampling Procedures

Myocardial ischemia reperfusion (I/R) injury contributes to almost half of the necrotic area after myocardial infarction. To date there is no approved drug to prevent or reduce myocardial I/R injury. The study and understanding of the pathophysiological mechanisms of myocardial I/R injury is essential to develop successful treatments. Large animal experiments are an important step in translational methods. The porcine model of acute myocardial infarction has been established and described by ourselves and others. We aimed to further improve the value of the model by focusing in detail on the sampling techniques for use in future experiments. Furthermore, we emphasize small but important steps that can affect the quality of the final results. To mimic the clinical situation of myocardial I/R injury, a percutaneous coronary intervention (PCI) catheter was inserted into the left anterior descending coronary artery (LAD) of an anesthetized pig. &#176;&#176;&#176;This model mimics acute myocardial infarction and PCI treatment in humans with the possibility of accurately determining the area at risk as well as the necrotic- and viable ischemic tissue. Here the model was used to investigate the effect of a bicyclic peptide inhibitor of FXIIa. The model can also be modified to allow longer reperfusion times to study later effects of myocardial infarction.

Here we demonstrate a protocol to standardize sampling procedures of an established porcine model of acute myocardial infarction in order to increase its translational value in the understanding of the pathophysiology of myocardial ischemia/reperfusion injury and to test novel drug candidates.

Myocardial ischemia reperfusion (I/R) injury contributes to almost half of the necrotic area after myocardial infarction. To date there is no approved ...

A Closed-chest Model to Induce Transverse Aortic Constriction in Mice

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Chapters in this video

The CYP2D6 Animal Model: How to Induce Autoimmune Hepatitis in Mice

In vitro Measurements of Tracheal Constriction Using Mice

A Murine Closed-chest Model of Myocardial Ischemia and Reperfusion

Ascending Aortic Constriction in Rats for Creation of Pressure Overload Cardiac Hypertrophy Model

A Rat Model of Ventricular Fibrillation and Resuscitation by Conventional Closed-chest Technique

A Magnetic Microbead Occlusion Model to Induce Ocular Hypertension-Dependent Glaucoma in Mice

Assessment of Cardiac Morphological and Functional Changes in Mouse Model of Transverse Aortic Constriction by Echocardiographic Imaging

Minimally Invasive Transverse Aortic Constriction in Mice

Technique of Minimally Invasive Transverse Aortic Constriction in Mice for Induction of Left Ventricular Hypertrophy

Improvement of a Closed Chest Porcine Myocardial Infarction Model by Standardization of Tissue and Blood Sampling Procedures