We are thankful to Manuela Ritzal, Hans-Peter Gensheimer, Christin Richter and the team from the Interfakult&#228;re Biomedizinische Forschungseinrichtung (IBF) from the Heidelberg University for expert technical assistance.
This work was supported by the DZHK (German Centre for Cardiovascular Research), the BMBF (German Ministry of Education and Research), a Baden-W&#252;rttemberg federal state Innovation fonds and the Deutsche Forschungsgemeinschaft (DFG, German Research Foundation) Project-ID 239283807 - TRR 152, FOR 2289 and the Collaborative Research Center (SFB) 1118.

All animal experiments were approved and performed according to the regulations of the Regional Council of Karlsruhe and the University of Heidelberg (AZ 35-9185.82/A-2/15, AZ 35-9185.82/A-18/15, AZ 35-9185.81/G131/15, AZ 35-9185.81/G121/17) conform to the guidelines from Directive 2010/63/EU of the European Parliament on the protection of animals used for scientific purposes. Data shown in this protocol are derived from wild type C57Bl6/N male mice (17 &#177; 1.4 weeks of age).&#160;Mice were maintained under specified ...

<ol>
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Here, we provide a protocol to analyze the in vivo cardiac function in mice under increasing &#946;-adrenergic stimulation. The procedure can be used to address both, baseline parameters of cardiac function and the adrenergic reserve (e.g., inotropy and chronotropy) in genetically modified mice or upon interventions. The most prominent advantage of pressure-volume loop (PVL) measurements as compared to other means of determining cardiac function is the analysis of intrinsic, load-independent cardiac function. All other m...

Cardiovascular diseases typically affect cardiac function. This issue points out the importance in assessing in vivo detailed cardiac function in animal disease models. Animal experimentation is surrounded by a frame of the three Rs (3Rs) guiding principles (Reduce/Refine/Replace). In case of understanding complex pathologies involving systemic responses (i.e., cardiovascular diseases) at the current developmental level, the main option is to refine the available methods. Refining will also lead to a reduction of the required animal numbers due to less variability, which improves the power of the analysis and conclusions. In addition, combination of cardiac contractility measurements with animal models of heart disease including those induced by neurohumoral stimulation or by pressure overload like aortic banding, which mimics for example altered catecholamine/&#946;-adrenergic levels1,2,3,4, provides a powerful method for pre-clinical studies. Taking into account that the catheter-based method remains the most widely used approach for in depth assessment of cardiac contractility5, we aimed to present here a refined measurement of in vivo cardiac function in mice by pressure-volume loop (PVL) measurements during &#946;-adrenergic stimulation based on previous experience including the evaluation of specific parameters of this approach6,7.
To determine cardiac hemodynamic parameters approaches that include imaging or catheter-based techniques are available. Both options are accompanied by advantages and disadvantages that carefully need to be considered for the respective scientific question. Imaging approaches include echocardiography and magnetic resonance imaging (MRI); both have been successfully used in mice. Echocardiographic measurements involve high initial costs from a high-speed probe required for the high heart rate of the mice; it is a relatively straightforward non-invasive approach, but it is variable among operators who ideally should be experienced recognizing and visualizing&#160;cardiac structures. In addition, no pressure measurements can be performed directly and calculations are obtained from combination of size magnitudes and flow measurements. On the other hand, it has the advantage that several measurements can be performed on the same animal and cardiac function can be monitored for example during disease progression. Regarding the volume measurement, the MRI is the gold standard procedure, but similar to echocardiography, no direct pressure measurements are possible and only preload dependent parameters can be obtained8. Limiting factors are also the availability, analysis effort and operating costs. Here catheter-based methods to measure cardiac function are a good alternative that additionally allow for the direct monitoring of intracardiac pressure and the determination of load-independent contractility parameters like preload recruitable stroke work (PRSW)9. However, ventricular volumes measured by a pressure-conductance catheter (through conductivity determination) are smaller than those from the MRI but group differences are maintained in the same range10. In order to determine reliable volume values the corresponding calibration is required, which is a critical step during the PVL measurements. It combines ex vivo measurements of blood conductivity in volume-calibrated cuvettes (conversion of conductance to volume) with the in vivo analysis for the parallel conductance of the myocardium during the bolus injection of the hypertonic saline11,12. Beyond that, the positioning of the catheter inside the ventricle and the correct orientation of the electrodes along the longitudinal axis of the ventricle are critical for the detection capability of the surrounding electrical field produced by them. Still with the reduced size of the mouse heart it is possible to avoid artifacts produced by changes in the intraventricular orientation of the catheter, even in dilated ventricles5,10, but artifacts can evolve under &#946;-adrenergic stimulation6,13. Additional to the conductance methods the development of admittance based method appeared to avoid the calibration steps, but here the volume values are rather overestimated14,15.
Since the mouse is one of the most important pre-clinical models in cardiovascular research and the &#946;-adrenergic reserve of the heart is of central interest in cardiac physiology and pathology, we here present a refined protocol to determine in vivo cardiac function in mice by PVL measurements during &#946;-adrenergic stimulation.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>1.4F SPR-839 catheter</td>
			<td>Millar Instruments, USA</td>
			<td>840-8111</td>
			<td></td>
		</tr>
		<tr>
			<td>1 ml syringes</td>
			<td>Beckton Dickinson, USA</td>
			<td>REF303172</td>
			<td></td>
		</tr>
		<tr>
			<td>Bio Amplifier</td>
			<td>ADInstruments, USA</td>
			<td>FE231</td>
			<td></td>
		</tr>
		<tr>
			<td>Bridge-Amplifier</td>
			<td>ADInstruments, USA</td>
			<td>FE221</td>
			<td></td>
		</tr>
		<tr>
			<td>Bovine Serum Albumin</td>
			<td>Roth, Germany</td>
			<td>8076.2</td>
			<td></td>
		</tr>
		<tr>
			<td>Buprenorphine hydrochloride</td>
			<td>Bayer, Germany</td>
			<td>4007221026402</td>
			<td></td>
		</tr>
		<tr>
			<td>Calibration cuvette</td>
			<td>Millar, USA</td>
			<td>910-1049</td>
			<td></td>
		</tr>
		<tr>
			<td>Differential pressure transducer MPX</td>
			<td>Hugo Sachs Elektronik- Harvard Apparatus, Germany</td>
			<td>Type 39912</td>
			<td></td>
		</tr>
		<tr>
			<td>Dumont Forceps #5/45</td>
			<td>Fine Science tools Inc.</td>
			<td>11251-35</td>
			<td></td>
		</tr>
		<tr>
			<td>Dumont Forceps #7B</td>
			<td>Fine Science tools Inc.</td>
			<td>11270-20</td>
			<td></td>
		</tr>
		<tr>
			<td>Graefe Forceps</td>
			<td>Fine Science tools Inc.</td>
			<td>11051-10</td>
			<td></td>
		</tr>
		<tr>
			<td>GraphPad Prism</td>
			<td>GraphPad Software</td>
			<td>Ver. 8.3.0</td>
			<td></td>
		</tr>
		<tr>
			<td>EcoLab-PE-Micotube</td>
			<td>Smiths, USA</td>
			<td>004/310/168-1</td>
			<td></td>
		</tr>
		<tr>
			<td>Etomidate Lipuro</td>
			<td>Braun, Germany</td>
			<td>2064006</td>
			<td></td>
		</tr>
		<tr>
			<td>Excel</td>
			<td>Microsoft</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Heparin</td>
			<td>Ratiopharm, Germany</td>
			<td>R26881</td>
			<td></td>
		</tr>
		<tr>
			<td>Hot plate and control unit</td>
			<td>Labotec, Germany</td>
			<td>Hot Plate 062</td>
			<td></td>
		</tr>
		<tr>
			<td>Isofluran</td>
			<td>Baxter, Germany</td>
			<td>HDG9623</td>
			<td></td>
		</tr>
		<tr>
			<td>Isofluran Vaporizer</td>
			<td>Abbot</td>
			<td>Vapor 19.3</td>
			<td></td>
		</tr>
		<tr>
			<td>Isoprenalinhydrochloride</td>
			<td>Sigma-Aldrich, USA</td>
			<td>I5627</td>
			<td></td>
		</tr>
		<tr>
			<td>Fine Bore Polythene tubing 0.61 mm OD, 0.28 mm ID</td>
			<td>Smiths Medical International Ltd, UK</td>
			<td>Ref. 800/100/100</td>
			<td></td>
		</tr>
		<tr>
			<td>MiniVent ventilator for mice</td>
			<td>Hugo Sachs Elektronik- Harvard Apparatus, Germany</td>
			<td>Type 845</td>
			<td></td>
		</tr>
		<tr>
			<td>MPVS Ultra PVL System</td>
			<td>Millar Instruments, USA</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>NaCl</td>
			<td>AppliChem, Germany</td>
			<td>A3597</td>
			<td></td>
		</tr>
		<tr>
			<td>NaCl 0.9% isotonic</td>
			<td>Braun, Germany</td>
			<td>2350748</td>
			<td></td>
		</tr>
		<tr>
			<td>Pancuronium-bromide</td>
			<td>Sigma-Aldrich, USA</td>
			<td>BCBQ8230V</td>
			<td></td>
		</tr>
		<tr>
			<td>Perfusor 11 Plus</td>
			<td>Harvard Apparatus</td>
			<td>Nr. 70-2209</td>
			<td></td>
		</tr>
		<tr>
			<td>Powerlab 4/35 control unit</td>
			<td>ADInstruments, USA</td>
			<td>PL3504</td>
			<td></td>
		</tr>
		<tr>
			<td>Rechargeable cautery-Set</td>
			<td>Faromed, Germany</td>
			<td>09-605</td>
			<td></td>
		</tr>
		<tr>
			<td>Scissors</td>
			<td>Fine Science tools Inc.</td>
			<td>140094-11</td>
			<td></td>
		</tr>
		<tr>
			<td>Software LabChart 7 Pro</td>
			<td>ADInstruments, USA</td>
			<td>LabChart 7.3 Pro</td>
			<td></td>
		</tr>
		<tr>
			<td>Standard mouse food</td>
			<td>LASvendi GmbH, Germany</td>
			<td>Rod18</td>
			<td></td>
		</tr>
		<tr>
			<td>Stereo microscope</td>
			<td>Zeiss, Germany</td>
			<td>Stemi 508</td>
			<td></td>
		</tr>
		<tr>
			<td>Surgical suture 8/0</td>
			<td>Suprama, Germany</td>
			<td>Ch.B.03120X</td>
			<td></td>
		</tr>
		<tr>
			<td>Venipuncture-cannula Venflon Pro Safty 20-gauge</td>
			<td>Beckton Dickinson, USA</td>
			<td>393224</td>
			<td></td>
		</tr>
		<tr>
			<td>Vessel Cannulation Forceps</td>
			<td>Fine Science tools Inc.</td>
			<td>00574-11</td>
			<td></td>
		</tr>
		<tr>
			<td>Water bath</td>
			<td>Thermo Fisher Scientific, USA</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Syringe filter (Filtropur S 0.45)</td>
			<td>Sarstedt, Germany</td>
			<td>Ref. 83.1826</td>
			<td></td>
		</tr>
	</tbody>
</table>

cardiac response to &#946;-adrenergic stimulation determined by pressure-volume loop analysis

Determination of the cardiac function is a robust endpoint analysis in animal models of cardiovascular diseases in order to characterize effects of specific treatments on the heart. Due to the feasibility of genetic manipulations the mouse has become the most common mammalian animal model to study cardiac function and to search for new potential therapeutic targets. Here we describe a protocol to determine cardiac function&#160;in vivo using pressure-volume loop measurements and analysis during basal conditions and under &#946;-adrenergic stimulation by intravenous infusion of increasing concentrations of isoproterenol. We provide a refined protocol including ventilation support taking into account the positive end-expiratory pressure to ameliorate negative effects during open-chest measurements, and potent analgesia (Buprenorphine) to avoid uncontrollable myocardial stress evoked by pain during the procedure. All together the detailed description of the procedure and discussion about possible pitfalls enables highly standardized and reproducible pressure-volume loop analysis, reducing the exclusion of animals from the experimental cohort by preventing possible methodological bias.

The pressure volume-loop (PVL) measurement is a powerful tool to analyze cardiac pharmacodynamics of drugs and to investigate the cardiac phenotype of genetically modified mouse models under normal and pathological conditions. The protocol allows the assessment of cardiac &#946;-adrenergic reserve in the adult mouse model. Here we describe an open-chest method under isoflurane anesthesia combined with buprenorphine (analgesic) and pancuronium (muscle relaxant), which focuses on the cardiac response to &#946;-adrenergic s...

Watch this Scientific Journal Video about Cardiac Response to Î²-Adrenergic Stimulation Determined by Pressure-Volume Loop Analysis at JoVE.com

Cardiac Response to &amp;#946;-Adrenergic Stimulation Determined by Pressure-Volume Loop Analysis

Here we describe a cardiac pressure-volume loop analysis under increasing doses of intravenously infused isoproterenol to determine the intrinsic cardiac function and the &#946;-adrenergic reserve in mice. We use a modified open-chest approach for the pressure-volume loop measurements, in which we include ventilation with positive end-expiratory pressure.

Cardiac Response to &#946;-Adrenergic Stimulation Determined by Pressure-Volume Loop Analysis

Determination of the cardiac function is a robust endpoint analysis in animal models of cardiovascular diseases in order to characterize effects of ...

Our open-chest approach, in which we include ventilation with positive end-expiratory pressure, is significant to assess and analyze cardiac function in vivo, and the increase in beta-adrenergic stimulation. The procedure can be used to address both baseline parameters of load-independent cardiac function, and the beta-adrenergic reserve in genetically modified mice or upon interventions. After confirming a lack of response to pedal reflex, disinfect the left hind limb of the anesthetized mouse with 70%ethanol and make an incision to expose the left femoral vein.Blast the epigastric artery and vein with a cautery. To better see the femoral vein, the field of vision is spanned by a suture through the abdomen. Then, a suture is placed distal to the catheter access, to ligate the femoral vein.Pass a suture under the femoral vein, and prepare a knot cranial to the puncture site. Puncture the femoral vein with a prepared micro tube attached to a 1-milliliter syringe and secure the tube inside the vessel with the knotted suture. To counteract any fluid loss, use an automatic syringe pump to infuse 0.9%sodium chloride, supplemented with 12.5%albumin, at a 15 microliter infusion rate, and hydrate the exposed tissue with pre-warmed 0.9%sodium chloride.To access the thorax, rinse the thorax with 70%ethanol, and incise the skin just beneath the xiphoid process. Bluntly separate the pectoral muscles from the chest wall, and use forceps to lift the xiphoid process to allow the chest wall to be cut, until the diaphragm is fully visible. Incise the diaphragm from below to expose the cardiac apex, and use forceps to carefully remove the pericardium.Perform a limited costotomy on the left side, and pass a suture beneath the inferior caval vein to allow downstream preload reduction. Using a 25 gauge cannula, gently puncture the cardiac apex, and replace the cannula with a pressure-volume catheter, until all the electrodes are within the ventricle. Then, gently adjust the position of the catheter until rectangular-shaped loops are obtained.To acquire pressure-volume loop measurements, perform an online analysis of the cardiac function parameters of interest, and wait until a steady-state cardiac function is obtained. Stop the respirator at the end-expiratory position, and record the baseline parameters. After 3 to 5 seconds, use forceps to lift the suture beneath the inferior caval vein to obtain pre-load independent parameters, and turn on the ventilator.Wait at least 30 seconds for the second occlusion until the hemodynamic parameters are stabilized. After obtaining the measurement under basal conditions, mount an isoproterenol-loaded syringe onto the syringe pump. Wait at least two minutes, until a new steady-state cardiac function is observed.Before stopping the respirator at the end-expiratory position, and recording the baseline parameters. After 3 to 5 seconds, lift the suture beneath the inferior caval vein to obtain preload independent parameters, and wait at least 30 seconds for the second occlusion. Then mount the syringe containing the next isoproterenol concentration, and repeat the baseline, and pre-load independent parameters recordings.To perform a parallel-conductance calibration, after the last isoproterenol dose-response measurement, connect a syringe loaded with the 15%sodium chloride solution to the femoral cannula, and carefully infuse 5 microliters of the hypertonic solution until the pressure-volume loop slightly shifts to the right. Wait until the loops return to the steady-state, before stopping the respirator at end-expiration, and immediately injecting a 10 microliter bolus of 15%sodium chloride. Then check whether the pressure-volume loop has largely broadened, and shifted to the right.To perform a conductance-to-volume ratio, wait 5 minutes until the hypertonic saline bolus is completely diluted before removing the catheter, and using a 1-milliliter syringe, equipped with a 21 gauge cannula to remove at least 600 microliters of blood from the left ventricle of the beating heart. Then transfer the blood into a pre-warmed calibration cuvette in a 37-degree celsius water bath, with cylinders of a known volume, and place the pressure-volume catheter centrally into each cylinder to allow recording of the conductance. Using this open chest pressure-volume-loop measurement method, if the catheter is correctly placed within the ventricle, a full cardiac cycle will be represented by a rectangular-shaped pressure-volume-loop.As illustrated, systole begins with a phase of isovolumetric contraction, during which both cardiac valves are closed. When the ventricular pressure, exceeds the aortic pressure, the aortic valve opens, and blood is pumped into the aorta during the ejection phase. Subsequently, when the aortic pressure exceeds the ventricular pressure, the aortic valve closes, and diastole begins.During the isovolumetric relaxation, the ventricular pressure falls, until the atrial pressure exceeds the ventricular pressure, and the mitral valve opens. Passive diastolic filling, characterized by the end-diastolic pressure-volume relationship then takes place until the next cardiac cycle begins. Since it is capable of determining cardiac function independent of preload, pressure-volume analysis can provide detailed insight into cardiac function, in cardiac contractility.For example, in this analysis isoproterenol induced a significant effect on each measured cardiac function, and cardiac contractility parameter. In contrast, a reduction in the diastolic parameters was observed in response to, increasing isoproterenol concentrations. Precise catheter placement within the left ventricle, is crucial to obtain reproducible loops without artifacts.To optimize catheter placement, we prefer the open chest approach. Following PV loop measurements, allometric endysis can be determined, and tissue can be isolated, to identify molecular and histopathological alterations, caused by treatments, and or genetic modifications. This technique paves the way to analyze beta-adrenergic reserve in mouse models under normal and pathological conditions and may be used as a template for testing other stimuli.

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A Murine Model of Muscle Training by Neuromuscular Electrical Stimulation

Neuromuscular electrical stimulation (NMES) is a common clinical modality that is widely used to restore1, maintain2 or enhance3-5 muscle functional capacity. Transcutaneous surface stimulation of skeletal muscle involves a current flow between a cathode and an anode, thereby inducing excitement of the motor unit and the surrounding muscle fibers. 
NMES is an attractive modality to evaluate skeletal muscle adaptive responses for several reasons. First, it provides a reproducible experimental model in which physiological adaptations, such as myofiber hypertophy and muscle strengthening6, angiogenesis7-9, growth factor secretion9-11, and muscle precursor cell activation12 are well documented. Such physiological responses may be carefully titrated using different parameters of stimulation (for Cochrane review, see 13). In addition, NMES recruits motor units non-selectively, and in a spatially fixed and temporally synchronous manner14, offering the advantage of exerting a treatment effect on all fibers, regardless of fiber type. Although there are specified contraindications to NMES in clinical populations, including peripheral venous disorders or malignancy, for example, NMES is safe and feasible, even for those who are ill and/or bedridden and for populations in which rigorous exercise may be challenging. 
Here, we demonstrate the protocol for adapting commercially available electrodes and performing a NMES protocol using a murine model. This animal model has the advantage of utilizing a clinically available device and providing instant feedback regarding positioning of the electrode to elicit the desired muscle contractile effect. For the purpose of this manuscript, we will describe the protocol for muscle stimulation of the anterior compartment muscles of a mouse hindlimb.

A murine model of neuromuscular electrical stimulation (NMES), a safe and inexpensive clinical modality, to the anterior compartment muscles is described. This model has the advantage of modifying a readily available clinical device for the purpose of eliciting targeted and specific muscle contractions in mice.

Neuromuscular electrical stimulation (NMES) is a common clinical modality that is widely used to restore1, maintain2 or enhance3-5 muscle functional ...

Cardiac Stress Test Induced by Dobutamine and Monitored by Cardiac Catheterization in Mice

Dobutamine is a &beta;-adrenergic agonist with an affinity higher for receptor expressed in the heart (&beta;1) than for receptors expressed in the arteries (&beta;2). When systemically administered, it increases cardiac demand. Thus, dobutamine unmasks abnormal rhythm or ischemic areas potentially at risk of infarction. 
Monitoring of heart function during a cardiac stress test can be performed by either ecocardiography or cardiac catheterization. The latter is an invasive but more accurate and informative technique that the former.
Cardiac stress test induced by dobutamine and monitored by cardiac catheterization accomplished as described here allows, in a single experiment, the measurement of the following hemodynamic parameters: heart rate (HR), systolic pressure, diastolic pressure, end-diastolic pressure, maximal positive pressure development (dP/dtmax) and maximal negative pressure development (dP/dtmin), at baseline conditions and under increasing doses of dobutamine.
As expected, in normal mice we observed a dobutamine dose-related increase in HR, dP/dtmax and dP/dtmin. Moreover, at the highest dose tested (12 ng/g/min) the cardiac decompensation of high fat diet-induced obese mice was unmasked.

We describe the protocol to perform a cardiac stress test induced by dobutamine and monitored by cardiac catheterization in normal mice. Also we show its application to unmask subclinical cardiac disease in high fat diet-induced obese mice.

Dobutamine is a &beta;-adrenergic agonist with an affinity higher for receptor expressed in the heart (&beta;1) than for receptors expressed in the ...

Measurement of the Pressure-volume Curve in Mouse Lungs

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

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

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

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 ...

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 ...

Hemocompatibility Testing of Blood-Contacting Implants in a Flow Loop Model Mimicking Human Blood Flow

The growing use of medical devices (e.g., vascular grafts, stents, and cardiac catheters) for temporary or permanent purposes that remain in the body&#39;s circulatory system demands a reliable and multiparametric approach that evaluates the possible hematologic complications caused by these devices (i.e., activation and destruction of blood components). Comprehensive in vitro hemocompatibility testing of blood-contacting implants is the first step towards successful in vivo implementation. Therefore, extensive analysis according to the International Organization for Standardization 10993-4 (ISO 10993-4) is mandatory prior to clinical application. The presented flow loop describes a sensitive model to analyze the hemostatic performance of stents (in this case, neurovascular) and reveal adverse effects. The use of fresh human whole blood and gentle blood sampling are essential to avoid the preactivation of blood. The blood is perfused through a heparinized tubing containing the test specimen by using a peristaltic pump at a rate of 150 mL/min at 37 &#176;C for 60 min. Before and after perfusion, hematologic markers (i.e., blood cell count, hemoglobin, hematocrit, and plasmatic markers) indicating the activation of leukocytes (polymorphonuclear [PMN]-elastase), platelets (&#946;-thromboglobulin [&#946;-TG]), the coagulation system (thombin-antithrombin III [TAT]), and the complement cascade (SC5b-9) are analyzed. In conclusion, we present an essential and reliable model for extensive hemocompatibility testing of stents and other blood-contacting devices prior to clinical application.

This protocol describes a comprehensive hemocompatibility evaluation of blood-contacting devices using laser-cut neurovascular implants. A flow loop model with fresh, heparinized human blood is applied to mimic blood flow. After perfusion, various hematologic markers are analyzed and compared to the values gained directly after blood collection for hemocompatibility evaluation of the tested devices.

The growing use of medical devices (e.g., vascular grafts, stents, and cardiac catheters) for temporary or permanent purposes that remain in the ...

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) ...

In vitro Assessment of Cardiac Reprogramming by Measuring Cardiac Specific Calcium Flux with a GCaMP3 Reporter

Cardiac reprogramming has become a potentially promising therapy to repair a damaged heart. By introducing multiple transcription factors, including Mef2c, Gata4, Tbx5 (MGT), fibroblasts can be reprogrammed into induced cardiomyocytes (iCMs). These iCMs, when generated in situ in an infarcted heart, integrate electrically and mechanically with the surrounding myocardium, leading to a reduction in scar size and an improvement in heart function. Because of the relatively low reprogramming efficiency, purity, and quality of the iCMs, characterization of iCMs remains a challenge. The currently used methods in this field, including flow cytometry, immunocytochemistry, and qPCR, mainly focus on cardiac-specific gene and protein expression but not on the functional maturation of iCMs. Triggered by action potentials, the opening of voltage-gated calcium channels in cardiomyocytes leads to a rapid influx of calcium into the cell. Therefore, quantifying the rate of calcium influx is a promising method to evaluate cardiomyocyte function. Here, the protocol introduces a method to evaluate iCM function by calcium (Ca2+) flux. An &#945;MHC-Cre/Rosa26A-Flox-Stop-Flox-GCaMP3 mouse strain was established by crossing Tg(Myh6-cre)1Jmk/J (referred to as Myh6-Cre below) with Gt(ROSA)26Sortm38(CAG-GCaMP3)Hze/J (referred to as Rosa26A-Flox-Stop-Flox-GCaMP3 below) mice. Neonatal cardiac fibroblasts (NCFs) from P0-P2 neonatal mice were isolated and cultured in vitro, and a polycistronic construction of MGT was introduced to NCFs, which led to their reprogramming to iCMs. Because only successfully reprogrammed iCMs will express GCaMP3 reporter, the functional maturation of iCMs can be visually assessed by Ca2+ flux with fluorescence microscopy. Compared with un-reprogrammed NCFs, NCF-iCMs showed significant calcium transient flux and spontaneous contraction, similar to CMs. This protocol describes in detail the mouse strain establishment, isolation and selection of neonatal mice hearts, NCF isolation, production of retrovirus for cardiac reprogramming, iCM induction, the evaluation of iCM Ca2+ flux using our reporter line, and related statistical analysis and data presentation. It is expected that the methods described here will provide a valuable platform to assess the functional maturation of iCMs for cardiac reprogramming studies.

We describe here, the establishment and application of an Tg(Myh6-cre)1Jmk/J /Gt(ROSA)26Sortm38(CAG-GCaMP3)Hze/J (referred to as &#945;MHC-Cre/Rosa26A-Flox-Stop-Flox-GCaMP3 below) mouse reporter line for cardiac reprogramming assessment. Neonatal cardiac fibroblasts (NCFs) isolated from the mouse strain are converted into induced cardiomyocytes (iCMs), allowing for convenient and efficient evaluation of reprogramming efficiency and functional maturation of iCMs via calcium (Ca2+) flux.

Cardiac reprogramming has become a potentially promising therapy to repair a damaged heart. By introducing multiple transcription factors, including ...

Closed Chest Biventricular Pressure-Volume Loop Recordings with Admittance Catheters in a Porcine Model

Pressure-volume (PV) loop recording enables the state-of-the-art investigation of load-independent variables of ventricular performance. Uni-ventricular evaluation is often performed in preclinical research. However, the right and left ventricles exert functional interdependence due to their parallel and serial connections, encouraging simultaneous evaluation of both ventricles. Furthermore, various pharmacological interventions may affect the ventricles and their preloads and afterloads differently. 
We describe our closed chest approach to admittance-based bi-ventricular PV loop recordings in a porcine model of acute right ventricular (RV) overload. We utilize minimally invasive techniques with all vascular accesses guided by ultrasound. PV catheters are positioned, under fluoroscopic guidance, to avoid thoracotomy in animals, as the closed chest approach maintains the relevant cardiopulmonary physiology. The admittance technology provides real-time PV loop recordings without the need for post-hoc processing. Furthermore, we explain some essential troubleshooting steps during critical timepoints of the presented procedure. 
The presented protocol is a reproducible and physiologically relevant approach to obtain a bi-ventricular cardiac PV loop recording in a large animal model. This can be applied to a large variety of cardiovascular animal research.

Here we present a closed chest approach to admittance-based bi-ventricular pressure-volume loop recordings in pigs with acute right ventricular dysfunction.

Pressure-volume (PV) loop recording enables the state-of-the-art investigation of load-independent variables of ventricular performance. Uni-ventricular ...

Improved Renal Denervation Mitigated Hypertension Induced by Angiotensin II Infusion

The benefits of renal sympathetic denervation (RDN) on blood pressure have been proved in a large number of clinical trials in recent years. However, the regulatory mechanism of RDN on hypertension remains elusive. Thus, it&#39;s essential to establish a simpler RDN model in mice. In this study, osmotic mini pumps filled with Angiotensin II were implanted in 14-week-old C57BL/6 mice. One week after the implantation of the mini-osmotic pump, a modified RDN procedure was performed on bilateral renal arteries of the mice using phenol. Age-sex-matched mice were given saline and served as sham group. Blood pressure was measured at baseline and every week subsequently for 21 days. Then, renal artery, abdominal aorta and heart were collected for histological examination using H&#38;E and Masson staining. In this study, we present a simple, practical, repeatable, and standardized RDN model, which can control hypertension and alleviate cardiac hypertrophy. The technique can denervate peripheral renal sympathetic nerves without renal artery damage. Compared to previous models, the modified RDN facilitates the study of the pathobiology and pathophysiology of hypertension.

Here, we present a protocol for renal sympathetic denervation (RDN) in mice with hypertension induced by Angiotensin II infusion. The procedure is repeatable, convenient and allows to study the regulatory mechanisms of RDN on hypertension and cardiac hypertrophy.

The benefits of renal sympathetic denervation (RDN) on blood pressure have been proved in a large number of clinical trials in recent years. However, the ...