Animal experiments were performed in compliance with the Principles of Laboratory Animal Care formulated by the National Society for Medical Research and the Guide for the Care and Use of Laboratory Animals prepared by the National Academy of Science and published by the National Institutes of Health (NIH Publication No. 86-23, revised 1985). All experiments were approved by the Austrian Ministry of Education, Science and Culture (BMWF-66.011/0071-II/3b/2012).
1. Arterial catheterization
<ol>
	<li>Anesthetize the mouse with an intraperitoneal injection of ketamine and xylazine10&#160;and an analgesic according to local practice (e.g. buprenorphine).&#160;&#160;Pinch the hind limbs with forceps to confirm the correct depth of anesthesia. 
	NOTE: For this study, male C57BL/6N mice at the age of 8&#8722;12 weeks were used (weight 20&#8722;25 g). The authors have tested male BALB/c of the same age unsuccessfully but have not tried other strains (see Discussion).</li>
	<li>Remove the hair in the regions of interest (head, neck) using an electric razor. Ensure that no loose hair remains to prevent wound contamination. Subsequently disinfect the surgical field with 70% ethanol/chlorhexidine/ betadine and place the mouse supine with the head facing the surgeon.</li>
	<li>Make a cervical midline incision with dissecting scissors.</li>
	<li>Dissect the submandibular glands and neck muscle tissue and separate them in order to expose the common carotid artery. Use mostly blunt dissection via forceps.</li>
	<li>Place three 8-0 silk ligatures beneath the right common carotid artery.</li>
	<li>Place a clamp on the proximal ligature and bring tension in the artery so that the flow is suspended.</li>
	<li>Close the most distal ligature.</li>
	<li>Insert the arterial catheter through a small, preformed skin hole on the cranial aspect of the incision. Squeeze and deform the lumen of the catheter if it appears too large to reduce blood backflow. Fixate with all three ligatures.</li>
	<li>Fixate the catheter to the skin to avoid dislocation. Do this by using a suture (e.g., 5-0 monofilament, non-absorbable) which connects the catheter to the skin in the area of the preformed skin hole.</li>
</ol>
2. Tracheostomy
<ol>
	<li>Dissect the pre-tracheal musculature bluntly using forceps.</li>
	<li>Place two 8-0 silk ligatures beneath the trachea.</li>
	<li>Tracheotomize using micro scissors as proximally as possible to avoid unilateral ventilation. Use a horizontal cutting line between two tracheal cartilages.</li>
	<li>Insert the ventilation tube and fixate with both prepared ligatures. 
	NOTE: The proximal end of the trachea does not need to be ligated.</li>
	<li>Close the skin with a running suture (e.g., 6-0 monofilament, non-absorbable).</li>
	<li>Ventilate the mouse with a frequency of 150/min and a tidal volume of 200 &#181;L.</li>
</ol>
3. Brain death induction
<ol>
	<li>Arrange the mouse to the prone position.</li>
	<li>Remove the skin from the skull using surgical scissors and forceps to hold the skin.</li>
	<li>Drill a 1 mm caliber borehole paramedially above the left parietal cortex. Stop drilling before breaching the inner compact bone and the dura mater.</li>
	<li>Penetrate the final tissue bridge of the skull using blunt forceps, removing sharp edges.</li>
	<li>Insert the balloon catheter, so that it is entirely within the cranial cavity. Ensure that the balloon is prefilled with saline and all air is evacuated.</li>
	<li>Begin inflation at ~0.1 mL/min over a period of 10&#8722;15 min (total volume of 0.8&#8722;1.2 mL) with the help of a syringe pump. 
	NOTE: The mouse will exhibit myoclonus, mydriasis, further seizure activity and agonal gasps.</li>
	<li>Pronounce the mouse brain dead once the tail of the mouse has gone stiff and erected. 
	NOTE: BD is confirmed by a characteristic initial blood pressure peak (Cushing reflex), the absence of brain stem reflexes and spontaneous breathing. Regular apnea testing should be avoided during experiments, as mice may become circulatory instable due to a lack of oxygen.</li>
	<li>Stop the inflation of the balloon catheter.</li>
	<li>Put a heating blanket over the mouse to avoid hypothermia.</li>
</ol>
4. During BD period
<ol>
	<li>Monitor and document blood pressure regularly. Exclude mice with a prolonged hypotensive phase (mean arterial pressure [MAP] &#60; 50 mmHg for &#62;30 min).</li>
	<li>Infuse 0.1 mL of saline every 30 min to stabilize the blood pressure of the mouse. 
	NOTE: In total 0.8 mL of saline was administered to each mouse in this study.</li>
	<li>After 4 h of BD duration, harvest mouse organs/tissues. Exclude the mouse from the experiment if the heart of the mouse is not beating at the end of the experiment.</li>
</ol>
5. Sham procedure
<ol>
	<li>Perform steps 1.1&#8722;3.3. 
	NOTE: Do not open the inner compact bone. Do not insert or inflate a balloon catheter.</li>
	<li>Stay close to the mouse and continuously observe the animal. Repeatedly pinch the hind limbs with forceps to confirm the correct depth of anesthesia. Apply additional anesthesia subcutaneously (approximately 1/2 of the starting dose) if the mouse shows signs of awakening, which, to our experience, happens approximately after 2&#8722;3 h post-anesthesia.&#160;&#160;</li>
	<li>Apply the same amount of intravenous saline as in the BD mice. 
	NOTE: The sham mouse will regain spontaneous breathing after cessation of ventilation. The BD mouse will not. Apnea testing should be done while establishing the model, but during experiments it should be avoided due to unnecessary physiological stress.</li>
</ol>

<ol>
	<li>Hart, 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=OPTN/SRTR+2017+Annual+Data+Report:+Kidney.">OPTN/SRTR 2017 Annual Data Report: Kidney.</a> American Journal of Transplantation. 19 (Suppl 2), 19 (2019).</li><li>The Quality Standards Subcommittee of the American Academy of Neurology. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Practice+parameters+for+determining+brain+death+in+adults+(summary+statement).">Practice parameters for determining brain death in adults (summary statement).</a> Neurology. 45 (5), 1012-1014 (1995).</li><li>Terasaki, P. I., Cecka, J. M., Gjertson, D. W., Takemoto, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=High+survival+rates+of+kidney+transplants+from+spousal+and+living+unrelated+donors.">High survival rates of kidney transplants from spousal and living unrelated donors.</a> New England Journal Medicine. 333 (6), 333-336 (1995).</li><li>Pratschke, J., Neuhaus, P., Tullius, S. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=What+can+be+learned+from+brain-death+models?.">What can be learned from brain-death models?.</a> Transplant International. 18 (1), 15-21 (2005).</li><li>Wilhelm, M. J., 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=Activation+of+the+heart+by+donor+brain+death+accelerates+acute+rejection+after+transplantation.">Activation of the heart by donor brain death accelerates acute rejection after transplantation.</a> Circulation. 102 (19), 2426-2433 (2000).</li><li>Pomper, G., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Introducing+a+mouse+model+of+brain+death.">Introducing a mouse model of brain death.</a> Journal of Neuroscience Methods. 192 (1), 70-74 (2010).</li><li>Ritschl, P. V., 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=Donor+brain+death+leads+to+differential+immune+activation+in+solid+organs+but+does+not+accelerate+ischaemia-reperfusion+injury.">Donor brain death leads to differential immune activation in solid organs but does not accelerate ischaemia-reperfusion injury.</a> Journal of Pathology. 239 (1), 84-96 (2016).</li><li>Atkinson, C., 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=Donor+brain+death+exacerbates+complement-dependent+ischemia/reperfusion+injury+in+transplanted+hearts.">Donor brain death exacerbates complement-dependent ischemia/reperfusion injury in transplanted hearts.</a> Circulation. 127 (12), 1290-1299 (2013).</li><li>Oberhuber, R., 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=Treatment+with+tetrahydrobiopterin+overcomes+brain+death-associated+injury+in+a+murine+model+of+pancreas+transplantation.">Treatment with tetrahydrobiopterin overcomes brain death-associated injury in a murine model of pancreas transplantation.</a> American Journal of Transplantation. 15 (11), 2865-2876 (2015).</li><li>Floerchinger, 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=Inflammatory+immune+responses+in+a+reproducible+mouse+brain+death+model.">Inflammatory immune responses in a reproducible mouse brain death model.</a> Transplant Immunology. 27 (1), 25-29 (2012).</li><li>Steen, P. A., Milde, J. H., Michenfelder, J. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=No+barbiturate+protection+in+a+dog+model+of+complete+cerebral+ischemia.">No barbiturate protection in a dog model of complete cerebral ischemia.</a> Annals of Neurology. 5 (4), 343-349 (1979).</li><li>Cooper, D. K., Novitzky, D., Wicomb, W. N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+pathophysiological+effects+of+brain+death+on+potential+donor+organs,+with+particular+reference+to+the+heart.">The pathophysiological effects of brain death on potential donor organs, with particular reference to the heart.</a> Annals of the Royal College of Surgeons of England. 71 (4), 261-266 (1989).</li><li>Herijgers, P., Leunens, V., Tjandra-Maga, T. B., Mubagwa, K., Flameng, W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Changes+in+organ+perfusion+after+brain+death+in+the+rat+and+its+relation+to+circulating+catecholamines.">Changes in organ perfusion after brain death in the rat and its relation to circulating catecholamines.</a> Transplantation. 62 (3), 330-335 (1996).</li></ol>

The authors have nothing to disclose.

BD, a risk factor for allograft quality in multi-organ donors, entails a plethora of pathophysiological changes, which can only be sufficiently assessed using in vivo models. Hemodynamic changes, cytokine storm, hormonal changes and their ultimate impact on organ graft quality and survival cannot be analyzed in vitro4. The majority of basic transplantation as well as immunological research is dependent on sophisticated diagnostic tools, which are widely available only in mice models. Mice models h...

Transplantation is currently the only curative treatment for end-stage organ failure. Until now, brain death (BD) patients have been the main source for organ donations, although living donation and donation after circulatory death are valuable alternatives1. BD is defined by an irreversible coma (with a known cause), the absence of brain stem reflexes and apnea2. Unfortunately, BD organs demonstrate inferior results in long-term graft survival independent of human leukocyte antigen (HLA)-mismatch and cold ischemic time3. Meanwhile, intensive research on this antigen-independent risk factor has been performed resulting in three main aspects of pathophysiological changes mediated as a consequence of BD: hemodynamic, hormonal, and inflammatory4.
To date, experimental BD models in rodents have been mostly performed using rats. In order to gain greater insight into the immunological consequences on solid organs following BD, we aimed to establish a murine model of BD, as currently only mouse models allow for comprehensive investigations into genetic or immunological factors. In this context, the mouse system provides a larger variety of analytical tools.
The principle of BD induction as described here is based on an increase in intracranial pressure induced by the inflation of a balloon catheter inserted under the skull. Increased intracranial pressure mimics the physiological mechanism of BD by blocking the perfusion of the cerebrum, cerebellum, and brain stem5,6. To guarantee sufficient perfusion of peripheral organs, blood pressure measurement is obligatory during the procedure. The catheter used for this purpose at the same time serves for saline administration in order to stabilize the blood pressure by fluid substitution. As BD is accompanied by cessation of spontaneous breathing, sufficient ventilation must be ensured. An electric blanket maintains physiological core body temperature.
In summary, this model will enable in-depth studies into the influence of BD-induced injury, on leukocyte migration7, compliment activation8, ischemic reperfusion injury9, and other factors.

<table><tbody><tr><td>Arterial catheter (BD Neoflon 26G)</td><td>BD</td><td>391349</td><td></td></tr><tr><td>Blood Pressure Transducers (APT300)</td><td>Harvard Apparatus Inc.</td><td>73-3862</td><td></td></tr><tr><td>Fogarty Arterial Embolectomy Catheter N&amp;deg; 3</td><td>Edwards Lifesciences Corporation</td><td>120403F</td><td></td></tr><tr><td>Forceps</td><td>FST</td><td>11271-30</td><td></td></tr><tr><td>Homeothermic Blanket Systems with Flexible Probe</td><td>Harvard Apparatus Inc.</td><td>55-7020</td><td></td></tr><tr><td>Ketansol</td><td>Graeub</td><td>6680110</td><td></td></tr><tr><td>Micro scissor</td><td>FST</td><td>15018-10</td><td></td></tr><tr><td>Needle holder</td><td>FST</td><td>12060-02</td><td></td></tr><tr><td>Prolene 5-0</td><td>Ethicon</td><td>8698H</td><td></td></tr><tr><td>Pump 11 Elite Infusion Only Single</td><td>Harvard Apparatus Inc.</td><td>70-4500</td><td></td></tr><tr><td>Scissor</td><td>FST</td><td>14075-11</td><td></td></tr><tr><td>Stereotactic microscope</td><td>Olympus</td><td>SZX7</td><td></td></tr><tr><td>Transpore Tape</td><td>3M</td><td>1527-1</td><td></td></tr><tr><td>Underpads</td><td>Molinea.A</td><td>274301</td><td></td></tr><tr><td>Ventilator for mice (MiniVent Model 845)</td><td>Harvard Apparatus Inc.</td><td>73-0043</td><td></td></tr><tr><td>Xylasol</td><td>Graeub</td><td>7630109</td><td></td></tr></tbody></table>

brain death induction in mice using intra-arterial blood pressure monitoring and ventilation via tracheostomy

While both living donation and donation after circulatory death provide alternative opportunities for organ transplantation, donation after donor brain death (BD) still represents the major source for solid transplants. Unfortunately, the irreversible loss of brain function is known to induce multiple pathophysiological changes, including hemodynamic as well as hormonal modifications, finally leading to a systemic inflammatory response. Models that allow a systematic investigation of these effects in vivo are scarce. We present a murine model of BD induction, which could aid investigations into the devastating effects of BD on allograft quality. After implementing intra-arterial blood pressure measurement via the common carotid artery and reliable ventilation via a tracheostomy, BD is induced by steadily increasing intracranial pressure using a balloon catheter. Four hours after BD induction, organs may be harvested for analysis or for further transplantation procedures. Our strategy enables the comprehensive analysis of donor BD in a murine model, therefore allowing an in-depth understanding of BD-related effects in solid organ transplantation and potentially paving the way to optimized organ preconditioning.

The murine BD model was successfully performed more than 100 times with a success rate of over 90%. Additionally, post interventional organ transplantation of heart and kidney has been safely performed7.
BD induces a variety of pathophysiological changes that may be further investigated using this model. As shown in Figure 1, the blood pressure shows an initial hypertensive p...

Watch this Scientific Journal Video about Brain Death Induction in Mice Using Intra-Arterial Blood Pressure Monitoring and Ventilation via Tracheostomy at JoVE.com

Brain Death Induction in Mice Using Intra-Arterial Blood Pressure Monitoring and Ventilation via Tracheostomy

We present a murine model of brain death induction in order to evaluate the influence of its pathophysiological effects on organs as well as on consecutive grafts in the context of solid organ transplantation.

While both living donation and donation after circulatory death provide alternative opportunities for organ transplantation, donation after donor brain ...

brain-death-induction-mice-using-intra-arterial-blood-pressure

Research

JoVE Journal

Neuroscience

5.5K Views.  Charité - Universitätsmedizin Berlin. Hoewel donatie na donorherdenking de belangrijkste bron is voor vaste orgaantransplantatie, veroorzaakt het onomkeerbare verlies van hersenfunctie pathofysiologische veranderingen die leiden tot een systemische ontstekingsreactie. Dit murine model van brain-death inductie maakt het gebruik van een grote verscheidenheid aan analytische instrumenten en knock-out modellen om de paden van de hersenen-dood regelgeving te bestuderen. Voor arteriële katheterisatie na bevestiging van een gebrek aan ...

Video: Brain Death Induction in Mice Using Intra-Arterial Blood Pressure Monitoring and Ventilation via Tracheostomy

Pressure Controlled Ventilation to Induce Acute Lung Injury in Mice

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

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

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

Medicine

Use of a Hanging Weight System for Coronary Artery Occlusion in Mice

Murine studies of acute injury are an area of intense investigation, as knockout mice for different genes are becoming increasingly available 1-38. Cardioprotection by ischemic preconditioning (IP) remains an area of intense investigation. To further elucidate its molecular basis, the use of knockout mouse studies is particularly important 7, 14, 30, 39. Despite the fact that previous studies have already successfully performed cardiac ischemia and reperfusion in mice, this model is technically very challenging. Particularly, visual identification of the coronary artery, placement of the suture around the vessel and coronary occlusion by tying off the vessel with a supported knot is technically difficult. In addition, re-opening the knot for intermittent reperfusion of the coronary artery during IP without causing surgical trauma adds additional challenge. Moreover, if the knot is not tied down strong enough, inadvertent reperfusion due to imperfect occlusion of the coronary may affect the results. In fact, this can easily occur due to the movement of the beating heart.
Based on potential problems associated with using a knotted coronary occlusion system, we adopted a previously published model of chronic cardiomyopathy based on a hanging weight system for intermittent coronary artery occlusion during IP 39. In fact, coronary artery occlusion can thus be achieved without having to occlude the coronary by a knot. Moreover, reperfusion of the vessel can be easily achieved by supporting the hanging weights which are in a remote localization from cardiac tissues.
We tested this system systematically, including variation of ischemia and reperfusion times, preconditioning regiments, body temperature and genetic backgrounds39. In addition to infarct staining, we tested cardiac troponin I (cTnI)
as a marker of myocardial infarction in this model. In fact, plasma levels of cTnI correlated with infarct sizes (R2=0.8). Finally, we could show in several studies that this technique yields highly reproducible infarct sizes during murine IP and myocardial infarction6, 8, 30, 40, 41. Therefore, this technique may be helpful for researchers who pursue molecular mechanisms involved in cardioprotection by IP using a genetic approach in mice with targeted gene deletion. Further studies on cardiac IP using transgenic mice may consider this technique.

A murine model for myocardial ischemia and ischemic preconditioning is an important tool study cardioprotective mechanisms in vivo. Here, we report an easy applicable in situ model for cardiac IP using a hanging-weight system for coronary artery occlusion.

Murine studies of acute injury are an area of intense investigation, as knockout mice for different genes are becoming increasingly available 1-38. ...

Normothermic Cardiac Arrest and Cardiopulmonary Resuscitation: A Mouse Model of Ischemia-Reperfusion Injury

Acute Kidney Injury (AKI) is a common, highly lethal, complication of critical illness which has a high mortality1-4 and which is most 
frequently caused by whole-body hypoperfusion.5,6 Successful reproduction of whole-body hypoperfusion in rodent models has been fraught with 
difficulty.7-9,9,10 Models which employ focal ischemia have repeatedly demonstrated results which do not translate to the clinical setting, and larger 
animal models which allow for whole body hypoperfusion lack access to the full toolset of genetic manipulation possible in the mouse.11,12 However, in recent 
years a mouse model of cardiac arrest and cardiopulmonary resuscitation has emerged which can be adapted to model AKI.13 This model reliably reproduces 
physiologic, functional, anatomic, and histologic outcomes seen in clinical AKI, is rapidly repeatable, and offers all of the significant advantages of a murine surgical 
model, including access to genetic manipulative techniques, low cost relative to large animals, and ease of use. Our group has developed extensive experience 
with use of this model to assess a number of organ-specific outcomes in AKI.14,15

A powerful model for perioperative and critical care related acute kidney injury is presented. Using whole body hypoperfusion induced by cardiac arrest it is possible to nearly replicate the histologic and functional changes of clinical AKI.

Acute Kidney Injury (AKI) is a common, highly lethal, complication of critical illness which has a high mortality1-4 and which is most 
frequently caused ...

A Murine Model of Subarachnoid Hemorrhage

In this video publication a standardized mouse model of subarachnoid hemorrhage (SAH) is presented. Bleeding is induced by endovascular Circle of Willis perforation (CWp) and proven by intracranial pressure (ICP) monitoring. Thereby a homogenous blood distribution in subarachnoid spaces surrounding the arterial circulation and cerebellar fissures is achieved. Animal physiology is maintained by intubation, mechanical ventilation, and continuous on-line monitoring of various physiological and cardiovascular parameters: body temperature, systemic blood pressure, heart rate, and hemoglobin saturation. Thereby the cerebral perfusion pressure can be tightly monitored resulting in a less variable volume of extravasated blood. This allows a better standardization of endovascular filament perforation in mice and makes the whole model highly reproducible. Thus it is readily available for pharmacological and pathophysiological studies in wild type and genetically altered mice.

A standardized mouse model of subarachnoid hemorrhage by intraluminal Circle of Willis perforation is described. Vessel perforation and subarachnoid bleeding are monitored by intracranial pressure monitoring. In addition various vital parameters are recorded and controlled to maintain physiologic conditions.

In this video publication a standardized mouse model of subarachnoid  hemorrhage (SAH) is presented. Bleeding is induced by endovascular Circle of  Willis ...

Cardiopulmonary Bypass in a Mouse Model: A Novel Approach

As prolonged cardiopulmonary bypass becomes more essential during cardiac interventions, an increasing clinical demand arises for procedure optimization and for minimizing organ damage resulting from prolonged extracorporal circulation. The goal of this paper was to demonstrate a fully functional and clinically relevant model of cardiopulmonary bypass in a mouse. We report on the device design, perfusion circuit optimization, and microsurgical techniques. This model is an acute model, which is not compatible with survival due to the need for multiple blood drawings. Because of the range of tools available for mice (e.g., markers, knockouts, etc.), this model will facilitate investigation into the molecular mechanisms of organ damage and the effect of cardiopulmonary bypass in relation to other comorbidities.

This paper describes how to perform cardiopulmonary bypass in mice. This novel model will facilitate the investigation of the molecular mechanisms involved in organ damage.

As prolonged cardiopulmonary bypass becomes more essential during cardiac interventions, an increasing clinical demand arises for procedure optimization ...

Biventricular Assessment of Cardiac Function and Pressure-Volume Loops by Closed-Chest Catheterization in Mice

Assessment of cardiac function is essential to conduct cardiovascular and pulmonary-vascular preclinical research. Pressure-volume loops (PV loops) generated by recording both pressure and volume during cardiac catheterization are vital when assessing both systolic and diastolic cardiac function. Left and right heart function are closely related, reflected in ventricular interdependence. Thus, recording biventricular function in the same animal is important to get a complete assessment of cardiac function. In this protocol, a closed chest approach to cardiac catheterization consistent with the way catheterization is performed in patients is adopted in mice. While challenging, the closed chest strategy is a more physiological approach, because opening the chest results in major changes in preload and afterload that create artifacts, most notably a fall in systemic blood pressure. While high-resolution echocardiography is used to assess rodents, cardiac catheterization is invaluable, particularly when assessing diastolic pressures in both ventricles.
Described here is a procedure to perform invasive, closed chest, sequential left and right ventricular pressure-volume (PV) loops in the same animal. PV loops are acquired using&#160;admittance technology with a mouse pressure-volume catheter and pressure-volume system acquisition. The procedure is described, beginning with the neck dissection, which is required to access the right jugular vein and the right carotid artery, to the insertion and positioning of the catheter, and finally the data acquisition. Then, the criteria required to ensure the acquisition of high-quality PV loops are discussed. Finally, the analysis of the left and right ventricular PV loops and the different hemodynamic parameters available to quantify systolic and diastolic ventricular function are briefly described.

Presented here is a protocol to assess biventricular heart function in mice by generating pressure-volume (PV) loops from the right and left ventricle in the same animal using closed chest catheterization. The focus is on the technical aspect of surgery and data acquisition.

Assessment of cardiac function is essential to conduct cardiovascular and pulmonary-vascular preclinical research. Pressure-volume loops (PV loops) ...

A Mouse Model to Study Brain Physiology During Cardiac Arrest

Mouse Cardiac Arrest Model for Brain Imaging and Brain Physiology Monitoring During Ischemia and Resuscitation

Most cardiac arrest (CA) survivors experience varying degrees of neurologic deficits. To understand the mechanisms that underpin CA-induced brain injury and, subsequently, develop effective treatments, experimental CA research is essential. To this end, a few mouse CA models have been established. In most of these models, the mice are placed in the supine position in order to perform chest compression for cardiopulmonary resuscitation (CPR). However, this resuscitation procedure makes the real-time imaging/monitoring of brain physiology during CA and resuscitation challenging. To obtain such critical knowledge, the present protocol presents a mouse asphyxia CA model that does not require the chest compression CPR step. This model allows for the study of dynamic changes in blood flow, vascular structure, electrical potentials, and brain tissue oxygen from the pre-CA baseline to early post-CA reperfusion. Importantly, this model applies to aged mice. Thus, this mouse CA model is expected to be a critical tool for deciphering the impact of CA on brain physiology.

Author Spotlight: A Unique Mouse Model of Asphyxia-Induced Cardiac Arrest 

This protocol demonstrates a unique mouse model of asphyxia cardiac arrest that does not require chest compression for resuscitation. This model is useful for monitoring and imaging the dynamics of brain physiology during cardiac arrest and resuscitation.

Most cardiac arrest (CA) survivors experience varying degrees of neurologic deficits. To understand the mechanisms that underpin CA-induced brain injury ...

Multiple Intravenous Bolus Dosing and Invasive Hemodynamic Assessment in a Hypoxia-Induced Mouse Pulmonary Artery Hypertension Model

Pulmonary arterial hypertension (PAH) is a progressive life-threatening disease, primarily affecting small pulmonary arterioles of the lung. Currently, there is no cure for PAH. It is important to discover new compounds that can be used to treat PAH. The mouse hypoxia-induced PAH model is a widely used model for PAH research. This model recapitulates human clinical manifestations of PAH Group 3 disease and is an important research tool to evaluate the effectiveness of new experimental therapies for PAH. Research using this model often requires the administration of compounds in mice. For a compound that needs to be given directly into the bloodstream, optimizing intravenous (IV) administration is a key part of the experimental procedures. Ideally, the IV injection system should permit multiple injections over a set time course. Although the mouse hypoxia-induced PAH model is very popular in many laboratories, it is technically challenging to perform multiple IV bolus dosing and invasive hemodynamic assessment in this model. In this protocol, we present step-by-step instructions on how to carry out multiple IV bolus dosing via mouse jugular vein and perform arterial and right ventricle catheterization for hemodynamic assessment in mouse hypoxia-induced PAH model.

This protocol provides a step-by-step procedure for executing multiple intravenous bolus dose administration and invasive hemodynamic monitoring in mice. Investigators can use this protocol for future therapeutic compound screening for pulmonary artery hypertension.

Pulmonary arterial hypertension (PAH) is a progressive life-threatening disease, primarily affecting small pulmonary arterioles of the lung. Currently, ...

Study of Experimental Organ Donation Models for Lung Transplantation

Experimental models are important tools for understanding the etiological phenomena involved in various pathophysiological events. In this context, different animal models are used to study the elements triggering the pathophysiology of primary graft dysfunction after transplantation to evaluate potential treatments. Currently, we can divide experimental donation models into two large groups: donation after brain death and donation after circulatory arrest. In addition, the deleterious effects associated with hemorrhagic shock should be considered when considering animal models of organ donation. Here, we describe the establishment of three different lung donation models (post-brain death donation, post-circulatory death donation, and post-hemorrhagic shock donation) and compare the inflammatory processes and pathological disorders associated with these events. The objective is to provide the scientific community with reliable animal models of lung donation for studying the associated pathological mechanisms and searching for new therapeutic targets to optimize the number of viable grafts for transplantation.

The present study shows the establishment of three different lung donation models (post-brain death donation, post-circulatory death donation, and post-hemorrhagic shock donation). It compares the inflammatory processes and pathological disorders associated with these events.

Experimental models are important tools for understanding the etiological phenomena involved in various pathophysiological events. In this context, ...

Inducing Brain Death in a Mouse Model Using a Balloon Catheter

Source: Ritschl, P. V., et al. &#160;<a href="https://app.jove.com/v/60831">Brain Death Induction in Mice Using Intra-Arterial Blood Pressure Monitoring and Ventilation via Tracheostomy.</a> J. Vis. Exp. (2020).
This video demonstrates the procedure for inducing brain death in a mouse model by increasing controlled intracranial pressure through a balloon catheter, mimicking natural brain death.

All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.
1. ...

Brain Death Induction in Mice Using Intra-Arterial Blood Pressure Monitoring and Ventilation via Tracheostomy

Samenvatting

Meer video's verkennen

Hoofdstukken in deze video