The present studies are supported by National Heart, Lung, and Blood Institute Grant R01-HL0921, R01-DK083385 and R01HL098294 to H. K. 
 Eltzschig, the 1K08HL102267-01 to T. Eckle, and Foundation for Anesthesia Education and Research Grants to T. Eckle and H. K. Eltzschig, and American Heart 
 Association Grant to T. Eckle and H. K. Eltzschig and Deutsche Forschungsgemeinschaft (DFG) research fellowship to M. Koeppen.

General remarks:
All operations should be performed under an upright dissecting microscope (Olympus, SZX10 with Z-Axis Crank Post with STU2 Stand Boom Stand) 
and by using a surgical coagulator (11). The experimental groups should be matched as best as possible in age and weight to ensure comparability of the results. 
Temperature, blood pressure, anesthesia and fluid administration should be stable throughout.

1. Anesthesia and trachea preparation
<ol>
	<li>Use C57BL/6 mice that are at least 10 weeks old and have a body of 22-25 g. Induce Anesthesia using sodium pentobarbital at a dose 70 mg/kg body weight i.p (6). 
	Maintain anesthesia approximately with 10 mg/kg/h sodium pentobarbital. Be cautious with overdosing since this might significantly lower the blood pressure. 
	Re-dosing of pentobarbital &#x2013; even after hrs- can lead to severe increases in plasma levels.</li>
	<li>After anesthesia induction, secure mice in a supine position with the upper and lower extremities attached to the table using a tape and a suture 
	fastened to the ankles. Do the same for the head by using the teeth. Sufficient restraining is important for a successful intubation and a well controlled 
	surgery. Prior to surgery, cover the mouse with mineral oil to reduce the risk of mouse hair allergy. In order to ensure that the body temperature remains 
	stable cover the mice with commercially available food wrap.</li>
	<li>Place mice on a temperature-controlled heated table (RT, Effenberg, Munich, Germany) with a rectal thermometer probe attached to thermal feedback controller
	to maintain body temperature at 37&deg;C. </li>
	<li>Expose the trachea surgically. Dissect lateral and dorsal sides of the trachea of the connective tissue and place two 3.0 silk surgical suture (Harvard apparatus,
	USA) each 10 cm long underneath the trachea. The sutures should be approximately 1 cm apart. </li>
	<li>Carefully incise the trachea 3 &#x2013; 4 mm below larynx between two circular cartilages using a McPherson-Vannas Scissors (8 cm, straight blade; World Precision Instruments, USA). Make sure not to cause a bleeding, since this might confound the outcome parameters. </li>
	<li>Perform a tracheal intubation using a blunt polyethylene cannulae (Insyte 22g, Beckton Dickinson, USA). Insert the tip of the polyethylene cannulae in an 85 degree angle into the trachea. Then tilt the cannulae so it is in line with the tracheal lumen. Slowly advance the tube further down the trachea until the tip of cannulae has disappeared in the thorax aperture. Fixate the tube in this position with the two surgical silk suture placed dorsal of the trachea (see 1.4).</li>
</ol>

2. Technique of ventilator-induced lung injury
<ol>
	<li>Connect the tube to a ventilator. To induce lung injury we use a pressure controlled ventilation technique by using a Servo 900 C from Siemens (DRE Veterinary,
	USA). Animals will be ventilated using peak inspiratory pressure of 45 mbar, frequency of 80 breaths/min and a positive end-expiratory pressure of 0-3 mbar with
	a FiO2 = 1.0. The inspiration to expiration ratio should be 1:1. Despite of the fact that the Servo 900 C is built as ventilator for humans, its use in a 
	pressure controlled ventilator setting works excellent for the ventilation of mice.</li>
	<li>Monitor heart rate with an ECG (e.g. Hewlett Packard, B blingen, Germany). Make sure that the heart rate does not drop below 400. One should see a shift of the 
	heart axis to the right, when mechanical ventilation is instituted as a sign of increased pulmonary artery pressures subsequent to an increase in intrathoracic 
	pressure. If the mouse develops bradycardia, check the temperature and the anesthetic dose/concentration. Xylacin/ Ketamin anesthesia induces a heart heart of 
	250/ min and is therefore not recommended.</li>
	<li>Apply a proper fluid replacement. An infusion with normal saline 0.1 ml/hour via an arterial or venous catheter should be performed prior to the ventilation to 
	ensure sufficient venous filing. Due to the high ventilation pressures the venous return to the heart is impaired, which might lead to a critical drop of the mean 
	arterial pressure. Also, a saline bolus of 500 &mu;l could be given i.p. prior to surgery.</li>
	<li>Place a catheter in the carotid artery for continuous recording of blood pressure (27). Attach the arm to the body before you start dissecting the artery. 
	The carotid artery is exposed via blunt dissection of the paratracheal muscles. Following further exposure and careful avoidance of any tissue trauma (particular 
	of the vagal nerve), a catheter is inserted into the vessel using two sutures and a small clamp (37). This will expose a longer segment of the artery. Place a 
	ligature at the very end of the carotid artery. Attach a larger clamp to the end of the suture to obtain tension or fixate the suture to the table using tape. 
	Place another suture around the artery and dissect the artery to the very distal end. Here, place a small clamp. Use micro scissors to cut a small diagonal 
	opening into the artery. Hold the opening with a fine forceps (Dumont, WPI) and advance the proper sized catheter with your hands/forceps. Make a knot with your	second suture and secure the artery. Loosen the clamp and advance the catheter further. Secure the catheter with several knots and tape. Alternatively the carotid artery catheter can be place at the end of the experiment to collect arterial blood samples for blood gas analysis.</li>
</ol>

3. Recovery of tissue samples
After 3 hours of mechanical ventilation, samples are collected to assess the extent of lung injury. We recommend collecting brochnoalveolar lavage fluid (BAL), arterial blood and lung tissue.
<ol>
	<li>Obtain BAL fluid at the end of the experiment. After deepening of anesthesia, flush the tracheal tube with 1 ml phosphate buffered saline (PBS). The fluid should remain in the trachea/lung for three seconds before it is recovery via the connected syringe. The BAL is snap-frozen in liquid nitrogen and stored at -80&deg;C for further analysis. Be aware that the recovered volume could be significantly less than 1 ml. </li>
	<li>Perform blood gas analysis at the end of the experiment. In order to do that, an incision should be made right below the sternum. Hold the sternum with forceps and extent the incision along the rib cage. Next, the diaphragm is incised at the edges and is cut out from the ribs. Now there is an open view into the lower aperture of the thorax. Lift the sternum up with forceps and open the thorax by long cuts at the right and left side (as lateral as possible) so that the complete anterior thorax wall is turned up-wards. This should be done, while the experimental animal is still mechanically ventilated. The left ventricle is punctured using a 27 &frac12;G needle and arterial blood analysis is performed using the i-STAT System (Abbott, USA). If the arterial blood gas analysis should be performed, a BAL cannot be obtained, since this will be a significant confounder to the results.</li>
	<li>Alternatively to method above, arterial blood samples can be collected via the carotid artery catheter. However this should be done before the BAL is obtained.</li>
	<li>Excise the lungs en-bloc by pulling up the heart and cut the trachea. Have a piece of tissue ready to absorb blood, so the surgical site is visible. Pull the heart in the direction of the abdomen and carefully cut along the spine to mobilize all thoracic organs. Cut the aorta, remove the thoracic organs and place them on a clean surgical table.</li>
	<li>Cut away the heart and the major vessel of the tissue sample. Make sure that there is no thymus tissue still attached to the lung. Separate the lungs with a scissor and place in individual tubes and snap-freeze. Store at -80&deg;C for further analysis.</li>
</ol>
4. Measurement of lung injury
We recommend using the following outcome parameters to assess the extent of lung injury: Perform an albumin ELISA (Bethyl Laboratories, USA) and a myeloperoxidase (MPO) ELISA (Hycult Biotechnology, USA) to assess the extent of barrier dysfunction and the amount of inflammatory cells in the BAL fluid. Perform an MPO ELISA also form the lung tissue. If wet-to-dry ratio is to be measured we do not obtain BAL fluid and the pulmonary circulation is not flushed (see 3.3). Measure the weight of the lungs after excision. Then lungs are lyophilized for 48 h and lung tissue again is measured. Then the wet-to-dry ratio is measured as mg of water per mg of dry tissue (5).
<img src="/files/ftp_upload/2525/2525fig1.jpg" alt="Figure 1" /> 
Figure 1. Protein content in the BAL in response to VILI. Mice were anesthetized with pentobarbital, mechanical ventilation
 was instituted and mice were ventilated using pressure-controlled settings (inspiratory pressure of 45 mbar, positive end-exspiratory pressure
 3 mbar, 100% inspired oxygen concentration). After 0, 1, 2 and 3 hours of ventilation BAL was harvested and the protein content was quantified using a 
 bicinchoninic acid assay (BCA assay). The relative change of the protein content is shown normalized to 0 hours of ventilation 
 (n=4 per group, * indicates p &lt; 0.05 compared to control, mean &plusmn;SEM)

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P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Crucial+role+for+ecto-5'-nucleotidase+(CD73)+in+vascular+leakage+during+hypoxia.">Crucial role for ecto-5'-nucleotidase (CD73) in vascular leakage during hypoxia.</a> J. Exp. Med. 200, 1395-1405 (2004).</li><li>Aherne, C. M., Kewley, E. M., Eltzschig, H. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+resurgence+of+A2B+adenosine+receptor+signaling.">The resurgence of A2B adenosine receptor signaling.</a> Biochim Biophys Acta. , (2010).</li><li>Eckle, T., Koeppen, M., Eltzschig, H. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Role+of+extracellular+adenosine+in+acute+lung+injury.">Role of extracellular adenosine in acute lung injury.</a> Physiology (Bethesda). 24, 298-306 (2009).</li><li>Koeppen, M., Feil, R., Siegl, D., Feil, S., Hofmann, F., Pohl, U., de Wit, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=cGMP-dependent+protein+kinase+mediates+NO-+but+not+acetylcholine-induced+dilations+in+resistance+vessels+in+vivo.">cGMP-dependent protein kinase mediates NO- but not acetylcholine-induced dilations in resistance vessels in vivo.</a> Hypertension. 44, 952-955 (2004).</li></ol>

We have nothing to disclose.

The present study describes a technique of performing ventilator-induced lung injury in mice. This model demonstrates highly reproducible injury due to high pressure ventilation. Investigators who consider studying acute-lung injury in mice may benefit from this model.

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			<td>Sodium Pentobarbital (Fatal Plus)</td>
			<td>Vortech Pharmaceutical Ls, Ltd,</td>
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pressure controlled ventilation to induce acute lung injury in mice

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

Watch this Scientific Journal Video about Pressure Controlled Ventilation to Induce Acute Lung Injury in Mice at JoVE.com

Pressure Controlled Ventilation to Induce Acute Lung Injury in Mice

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

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

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18.2K Görüntüleme.  University of Colorado. Ventilatöre bağlı akciğer hasarı için bir fare modeli akut akciğer hasarı incelemek için önemli bir araçtır In vivo. Burada, kolay uygulanabilir rapor In situ Yüksek basınçlı mekanik ventilasyon akut akciğer yetmezliğine yol kullanarak akut akciğer hasarı modeli.

Video: Pressure Controlled Ventilation to Induce Acute Lung Injury 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 ...

Controlled Cervical Laceration Injury in Mice

Use of genetically modified mice enhances our understanding of molecular mechanisms underlying several neurological disorders such as a spinal cord injury (SCI). Freehand manual control used to produce a laceration model of SCI creates inconsistent injuries often associated with a crush or contusion component and, therefore, a novel technique was developed. Our model of cervical laceration SCI has resolved inherent difficulties with the freehand method by incorporating 1) cervical vertebral stabilization by vertebral facet fixation, 2) enhanced spinal cord exposure, and 3) creation of a reproducible laceration of the spinal cord using an oscillating blade with an accuracy of &plusmn;0.01 mm in depth without associated contusion. Compared to the standard methods of creating a SCI laceration such as freehand use of a scalpel or scissors, our method has produced a consistent lesion. This method is useful for studies on axonal regeneration of corticospinal, rubrospinal, and dorsal ascending tracts.

A novel technique to create a reproducible in vivo model of cervical spinal cord laceration injury in the mouse is described. This technique is based on spine stabilization by fixation of the cervical facets and laceration of the spinal cord using an oscillating blade with an accuracy of &plusmn;0.01 mm.

Use of genetically modified mice enhances our  understanding of molecular mechanisms underlying several neurological disorders  such as a spinal cord ...

Ischemia-reperfusion Model of Acute Kidney Injury and Post Injury Fibrosis in Mice

Ischemia-reperfusion induced acute kidney injury (IR-AKI) is widely used as a model of AKI in mice, but results are often quite variable with high, often unreported mortality rates that may confound analyses. Bilateral renal pedicle clamping is commonly used to induce IR-AKI, but differences between effective clamp pressures and/or renal responses to ischemia between kidneys often lead to more variable results. In addition, shorter clamp times are known to induce more variable tubular injury, and while mice undergoing bilateral injury with longer clamp times develop more consistent tubular injury, they often die within the first 3 days after injury due to severe renal insufficiency. To improve post-injury survival and obtain more consistent and predictable results, we have developed two models of unilateral ischemia-reperfusion injury followed by contralateral nephrectomy. Both surgeries are performed using a dorsal approach, reducing surgical stress resulting from ventral laparotomy, commonly used for mouse IR-AKI surgeries. For induction of moderate injury BALB/c mice undergo unilateral clamping of the renal pedicle for 26 min and also undergo simultaneous contralateral nephrectomy. Using this approach, 50-60% of mice develop moderate AKI 24 hr after injury but 90-100% of mice survive. To induce more severe AKI, BALB/c mice undergo renal pedicle clamping for 30 min followed by contralateral nephrectomy 8 days after injury. This allows functional assessment of renal recovery after injury with 90-100% survival. Early post-injury tubular damage as well as post injury fibrosis are highly consistent using this model.

We describe models of moderate and severe ischemia-reperfusion-induced kidney injury in which the mice undergo unilateral renal pedicle clamping followed by simultaneous or delayed contralateral nephrectomy, respectively. These models consistently give rise to renal dysfunction and post-injury fibrosis, but injury severity and survival are dependent on mouse background, age and surgical equipment.

Ischemia-reperfusion induced acute kidney injury (IR-AKI) is widely used as a model of AKI in mice, but results are often quite variable with high, often ...

Controlled Cortical Impact Model for Traumatic Brain Injury

Every year over a million Americans suffer a traumatic brain injury (TBI). Combined with the incidence of TBIs worldwide, the physical, emotional, social, and economical effects are staggering. Therefore, further research into the effects of TBI and effective treatments is necessary. The controlled cortical impact (CCI) model induces traumatic brain injuries ranging from mild to severe. This method uses a rigid impactor to deliver mechanical energy to an intact dura exposed following a craniectomy. Impact is made under precise parameters at a set velocity to achieve a pre-determined deformation depth. Although other TBI models, such as weight drop and fluid percussion, exist, CCI is more accurate, easier to control, and most importantly, produces traumatic brain injuries similar to those seen in humans. However, no TBI model is currently able to reproduce pathological changes identical to those seen in human patients. The CCI model allows investigation into the short-term and long-term effects of TBI, such as neuronal death, memory deficits, and cerebral edema, as well as potential therapeutic treatments for TBI.

Traumatic brain injuries (TBIs) remain a serious health problem. Using the controlled cortical impact surgery model, research on the effects of TBI and possible treatment methods may be performed.

Every year over a million Americans suffer a traumatic brain injury (TBI).  Combined with the incidence of TBIs worldwide, the physical, emotional, ...

Cardiac Catheterization in Mice to Measure the Pressure Volume Relationship: Investigating the Bowditch Effect

Animal models that mimic human cardiac disorders have been created to test potential therapeutic strategies. A key component to evaluating these strategies is to examine their effects on heart function. There are several techniques to measure in vivo cardiac mechanics (e.g., echocardiography, pressure/volume relations, etc.). Compared to echocardiography, real-time left ventricular (LV) pressure/volume analysis via catheterization is more precise and insightful in assessing LV function. Additionally, LV pressure/volume analysis provides the ability to instantaneously record changes during manipulations of contractility (e.g., &#946;-adrenergic stimulation) and pathological insults (e.g., ischemia/reperfusion injury). In addition to the maximum (+dP/dt) and minimum (-dP/dt) rate of pressure change in the LV, an accurate assessment of LV function via several load-independent indexes (e.g., end systolic pressure volume relationship and preload recruitable stroke work) can be attained. Heart rate has a significant effect on LV contractility such that an increase in the heart rate is the primary mechanism to increase cardiac output (i.e., Bowditch effect). Thus, when comparing hemodynamics between experimental groups, it is necessary to have similar heart rates. Furthermore, a hallmark of many cardiomyopathy models is a decrease in contractile reserve (i.e., decreased Bowditch effect). Consequently, vital information can be obtained by determining the effects of increasing heart rate on contractility. Our and others data has demonstrated that the neuronal nitric oxide synthase (NOS1) knockout mouse has decreased contractility. Here we describe the procedure of measuring LV pressure/volume with increasing heart rates using the NOS1 knockout mouse model.

This article describes the measurement of murine left ventricular function via pressure/volume analysis at different heart rates.

Animal models that mimic human cardiac disorders have been created to test potential therapeutic strategies. A key component to evaluating these ...

Open Tracheostomy Gastric Acid Aspiration Murine Model of Acute Lung Injury Results in Maximal Acute Nonlethal Lung Injury

Acid pneumonitis is a major cause of sterile acute lung injury (ALI) in humans. Acid pneumonitis spans the clinical spectrum from asymptomatic to acute respiratory distress syndrome (ARDS), characterized by neutrophilic alveolitis, and injury to both alveolar epithelium and vascular endothelium. Clinically, ARDS is defined by acute onset of hypoxemia, bilateral patchy pulmonary infiltrates and non-cardiogenic pulmonary edema. Human studies have provided us with valuable information about the physiological and inflammatory changes in the lung caused by ARDS, which has led to various hypotheses about the underling mechanisms. Unfortunately, difficulties determining the etiology of ARDS, as well as a wide range of pathophysiology have resulted in a lack of critical information that could be useful in developing therapeutic strategies.
Translational animal models are valuable when their pathogenesis and pathophysiology accurately reproduce a concept proven in both in vitro and clinical settings. Although large animal models (e.g., sheep) share characteristics of the anatomy of human trachea-bronchial tree, murine models provide a host of other advantages including: low cost; short reproductive cycle lending itself to greater data acquisition; a well understood immunologic system; and a well characterized genome leading to the availability of a variety of gene deletion and transgenic strains. A robust model of low pH induced ARDS requires a murine ALI that targets mainly the alveolar epithelium, secondarily the vascular endothelium, as well as the small airways leading to the alveoli. Furthermore, a reproducible injury with wide differences between different injurious and non-injurious insults is important.
The murine gastric acid aspiration model presented here using hydrochloric acid employs an open tracheostomy and recreates a pathogenic scenario that reproduces the low pH pneumonitis injury in humans. Additionally, this model can be used to examine interaction of a low pH insult with other pulmonary injurious entities (e.g., food particles, pathogenic bacteria).

This protocol induces acute lung injury in a mouse that has close fidelity to the pathogenesis of acid pneumonitis observed in humans. We generate a maximal acute nonlethal low pH lung injury and account for differences in rodent-human anatomic respiratory structure using an open tracheostomy coupled with circumferential pressure release.

Acid pneumonitis is a major cause of sterile acute lung injury (ALI) in humans. Acid pneumonitis spans the clinical spectrum from asymptomatic to acute ...

Balloon-based Injury to Induce Myointimal Hyperplasia in the Mouse Abdominal Aorta

The use of animal models is essential for a better understanding of MH, one major cause for arterial stenosis.In this article, we demonstrate a murine balloon denudation model, which is comparable with established vessel injury models in large animals. The aorta denudation model with balloon catheters mimics the clinical setting and leads to comparable pathobiological and physiological changes. Briefly, after performing a horizontal incision in the aorta abdominalis, a balloon catheter will be inserted into the vessel, inflated, and introduced retrogradely. Inflation of the balloon will lead to intima injury and overdistension of the vessel. After removing the catheter, the aortic incision will be closed with single stiches. The model shown in this article is reproducible, easy to perform, and can be established quickly and reliably. It is especially suitable for evaluating expensive experimental therapeutic agents, which can be applied in an economical fashion. By using different knockout-mouse strains, the impact of different genes on MH development can be assessed.

This article demonstrates a murine model to study the development of myointimal hyperplasia (MH) after aortic balloon injury.

The use of animal models is essential for a better understanding of MH, one major cause for arterial stenosis.In this article, we demonstrate a murine ...

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.

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 ...

Real-time Pressure-volume Analysis of Acute Myocardial Infarction in Mice

Acute myocardial infarction can lead to acute heart failure and cardiogenic shock. The evaluation of hemodynamics is critical for the evaluation of any potential therapeutic approach directed against acute left ventricular (LV) dysfunction. Current imaging modalities (e.g., echocardiography and magnetic resonance imaging) have several limitations since data on LV pressure cannot directly be measured. LV catheterization in mice undergoing coronary artery occlusion could serve as a novel method for a real-time evaluation of LV function.
At the beginning of the procedure, mice were anesthetized followed by endotracheal intubation. For LV catheterization, the right carotid artery was exposed via middle-neck incision. The catheter was introduced and placed into the LV cavity. Left thoracotomy was conducted and the left main coronary artery (LCA) was ligated. To induce reperfusion, the suture was released after 45 min. Pressure-volume data was recorded at all times.
Ligation of the LCA caused a decrease in LV systolic function as evidenced by a 30% reduction in stroke volume, LV ejection fraction (EF), and cardiac output. Maximum dP/dt as a parameter for LV contractility was also significantly reduced and diastolic function was severely impaired (minimum dP/dt -40%). Reperfusion over a period of 20 min did not lead to a complete recovery of LV function.
Real-time pressure-volume analysis served as a valid procedure for monitoring cardiac function during acute myocardial infarction in mice. Maintaining stable anesthesia and a standardized surgical approach was crucial to ensure valid results. As the early phase of acute myocardial infarction is critical for morbidity and mortality, the delineated method could be beneficial for preclinical evaluation of new strategies for cardioprotection.

Acute myocardial infarction in mice induces acute but incompletely characterized changes in left ventricular (LV) function. LV catheterization in mice undergoing coronary artery occlusion serves as a novel method for a real-time evaluation of LV function.

Acute myocardial infarction can lead to acute heart failure and cardiogenic shock. The evaluation of hemodynamics is critical for the evaluation of any ...

Lung Fixation under Constant Pressure for Evaluation of Emphysema in Mice

Emphysema is a significant feature of chronic obstructive pulmonary disease (COPD). Studies involving an emphysematous mouse model require optimal lung fixation to produce reliable histological specimens of the lung. Due to the nature of the lung&rsquo;s structural composition, which consists largely of air and tissue, there is a risk that it collapses or deflates during the fixation process. Various lung fixation methods exist, each of which has its own advantages and disadvantages. The lung fixation method presented here utilizes constant pressure to enable optimal tissue evaluation for studies using an emphysematous mouse lung model. The main advantage is that it can fix many lungs with the same condition at one time. Lung specimens are obtained from chronic cigarette smoke-exposed mice. Lung fixation is performed using specialized equipment that enables the production of constant pressure. This constant pressure maintains the lung in a reasonably inflated state. Thus, this method generates a histological specimen of the lung that is suitable to evaluate cigarette smoke-induced mild emphysema.

Presented here is a useful protocol for lung fixation that creates a stable condition for histological evaluate of lung specimens from a mouse model of emphysema. The main advantage of this model is that it can fix many lungs with the same constant pressure without lung collapse or deflation.

Emphysema is a significant feature of chronic obstructive pulmonary disease (COPD). Studies involving an emphysematous mouse model require optimal lung ...