<ol style="padding:5px;"><li> Ausrüstung Set-up <ol style="padding:5px;"><li> Wählen Sie einen Bereich in dem Labor, in dem die Raumtemperatur bei oder über 20 ° C. </li><li> Vermeiden Sie Orte in der Nähe Luftdüsen und / oder laute Geräusche. </li><li> Vermeiden Sie es, das Tier, um Gerüche, die Hautreizungen und / oder Stress verursachen, wie Parfüm, Eau de Cologne, Chemikalien und Reinigungsmittel können. </li><li> Schalten Sie den CODA-Controller. </li><li> Schalten Sie den Warming-Plattform und auf Level 3. Wenn die Raumtemperatur über 25 ° C, auf Stufe 1 oder 2 zu beginnen. </li><li> Controller Diagnostics-Test <ol style="padding:5px;"><li> Öffnen Sie die CODA-Software. </li><li> Wählen Sie die CODA-Gerät, indem Sie auf sie. </li><li> Klicken Sie auf &quot;Test Selected Device&quot;. </li><li> Wählen Sie die Manschetten und Kanäle zu testen, und klicken Sie auf &quot;Test&quot;. </li><li> Nach der Test abgeschlossen ist, schließen Sie das Device Test-Fenster. </li></ol></li></ol></li><li> Software Set-up <ol style="padding:5px;"><li> Verwalten Personal und Tiere <ol style="padding:5px;"><li> Verwalten Personal <ol style="padding:5px;"><li> Wählen Sie Extras&gt; Personal. </li><li> Klicken Sie auf &quot;Researcher 1&quot;, um den Namen zu ändern. </li><li> Klicken Sie auf nachfolgenden Zeilen, um weitere Forscher hinzu. </li><li> Klicken Sie auf &quot;Techniker Tab&quot; zu ändern oder zu ergänzen Techniker. </li><li> Klicken Sie auf &quot;Proben Tab&quot;, um das Tier für eine einfache Identifizierung Name und das Tier auszuwählen; Ratte oder Maus. </li><li> Klicken Sie auf &quot;Animals Tab&quot;, um die Tierart und die Empfindlichkeit zu verändern. </li></ol></li></ol></li><li> New Experiment <ol style="padding:5px;"><li> Wählen Sie Datei&gt; Neu&gt; Experiment </li><li> Beginnen Sie ein neues Experiment <ol style="padding:5px;"><li> Geben Sie den Namen Experiment. </li><li> Wählen Sie den Key Researcher. </li><li> Wählen Sie ein Startdatum und klicken Sie auf &quot;Weiter&quot;. </li></ol></li><li> Grundlegende Session Info <ol style="padding:5px;"><li> Geben Sie die Session Name </li><li> Set &quot;Akklimatisierung Cycles&quot; bis 5. Wenn Sie nicht mit Akklimatisierung Zyklen, auf Null gesetzt sind bevorzugen. Um mit acclimating das Tier an den Halter und Manschetten zu unterstützen, zusätzliche Zyklen zu Ihrem Experiment und führen Sie das Experiment an 5 aufeinander folgenden Tagen. </li><li> Set &quot;Anzahl der Sätze&quot; auf 1. </li><li> Set &quot;Time Between Sets&quot; bis 30 Sekunden für mehrere Sätze </li><li> Set &quot;Zyklen pro Set&quot; bis 20 Zyklen. </li><li> Set &quot;Zeit zwischen den Zyklen&quot; auf 5 Sekunden und klicken Sie auf &quot;Weiter&quot;. </li></ol></li></ol></li><li> Specimen Selection <ol style="padding:5px;"><li style="list-style-type:none;"><ol style="padding:5px;"><li> Wählen Sie die Probe aus der Probe Pool und weisen es der erste verfügbare Kanal und klicken Sie auf &quot;Weiter&quot;. </li></ol></li><li> Session-Parameter <ol style="padding:5px;"><li> Set &quot;Maximum Occlusion Pressure&quot; auf 250mmHg. </li><li> Set &quot;Deflation Time&quot; bis 20 Sekunden für Mäuse, 15 Sekunden für Ratten. </li><li> Set &quot;Minimum Volume&quot; auf 15ul. </li><li> Set &quot;Display Style&quot; zu einem Kanal pro Grafik. </li><li> Fügen Sie zusätzliche Forscher und Techniker aus und klicken Sie auf &quot;Weiter&quot;. </li></ol></li><li> Session Script <ol style="padding:5px;"><li> Überprüfen Sie die Session Script und klicken Sie auf &quot;Weiter&quot;. </li><li> Wählen Sie nicht Fertig, bis Sie das Tier Vorbereitung Abschnitt abzuschließen. </li></ol></li></ol></li></ol></li><li> Tierische Vorbereitung <ol style="padding:5px;"><li> Tierische Set-up <ol style="padding:5px;"><li> Tierhalter <ol style="padding:5px;"><li> Zu trainieren und zu verbessern Tier Akklimatisierung, fügen Sie weitere Zyklen, Ihr Experiment und führen Sie das Experiment an fünf aufeinanderfolgenden Tagen. Sie können auch mit Akklimatisierung, indem das Tier in die Halterung für 15 Minuten an 3 aufeinanderfolgenden Tagen vor dem eigentlichen Studium zu unterstützen. </li><li> Legen Sie für jedes Tier in einer Halterung, indem Sie den Tier am Schwanz und sanftes Auflegen das Tier in den hinteren Teil der Halterung, die das offene Ende des Nasenkonus Gesichter. Hinweis: die Größe des Tieres in Bezug auf die Größe der ausgewählten Halter. Das Tier sollte in der Lage, frei zu geben den Halter. </li><li> Sorgfältig sichern die Heckklappe an den Inhaber durch Drehen des roten Schraubverschluss auf der Heckklappe. Es ist darauf zu vermeiden, Einklemmen des Schwanzes oder andere Körperteile bei gleichzeitiger Sicherstellung der Heckklappe werden. </li><li> Schieben Sie die Bugspitze auf der Heckklappe, die Begrenzung der Bewegung des Tieres. Die Nase Kegel sollten in der Lage, das Tier von sich umzudrehen, während im Inneren des Halters zu begrenzen. </li><li> Platzieren Sie die Halterung auf die Erwärmung Plattform in der dafür vorgesehenen Stelle. Wenn der Test die gleiche Gruppe von Tieren über einen Zeitraum von Zeit, sicher sein, auf den Inhaber Position auf die Erwärmung Plattform drehen, um die Konsistenz und Genauigkeit der BP-Messungen zu verbessern. Lassen Sie das Tier mindestens 5-Minuten, um die Halterung zu akklimatisieren zu. </li><li> Nicht berühren oder Griff des Tieres, während im Inneren des Halters. Die vermehrte Kontakte konnte reizen das Tier. </li><li> Lassen Sie das Tier in den Inhabern unbeaufsichtigt. </li></ol></li><li> Cuff Placement <ol style = &quot;padding: 5px;&quot;&gt; <li> Thema der Schwanz durch die &quot;Occlusion Cuff.&quot; Place dieser Manschette so nah an der Basis des Schwanzes wie möglich, ohne Kraft. </li><li> Sichern Sie die &quot;Occlusion Manschettenschlauch&quot; in die Kerbe am oberen Rückseite des Halters. </li><li> Thema der Schwanz durch die &quot;VPR Cuff&quot;, indem es innerhalb von 2 mm von der &quot;Occlusion Cuff&quot;. </li><li> Sichern Sie die &quot;VPR Manschettenschlauch&quot; in die Kerbe am oberen Rückseite des Halters. </li><li> Befestigen Sie die Manschette auf die CODA-Controller. </li></ol></li><li> Tierische Temperatur <ol style="padding:5px;"><li> Lassen Sie das Tier mindestens 5-Minuten thermoregulate. </li><li> Notieren Sie die Temperatur des Tieres häufig mit einem Infrarot-Thermometer. Das Tier Temperatur sollte 32 bis 35 ° C werden </li></ol></li></ol></li></ol></li><li> Experiment <ol style="padding:5px;"><li> Begin Experiment <ol style="padding:5px;"><li> Klicken Sie auf &quot;Fertig stellen&quot; und das Experiment beginnt sofort. </li></ol></li><li> End Experiment <ol style="padding:5px;"><li> Tier <ol style="padding:5px;"><li> Entfernen Sie unverzüglich die Tiere aus den Manschetten und Halter. </li><li> Zurück das Tier zu ihrem Käfig nach der &quot;Simple Session Summary Report&quot; angezeigt wird. </li></ol></li><li> Data Collection <ol style="padding:5px;"><li> Nach Abschluss des letzten Zyklus der akzeptierten Zyklen automatisch in Spreadsheet-Format innerhalb der CODA-Anwendung angezeigt werden, auf der &quot;Export to Excel&quot; klicken. </li><li> Wählen Sie den Ordner, um die Datei vorhanden, benennen Sie die Datei und klicken Sie auf &quot;Speichern&quot;. </li><li> Öffnen Sie die Excel-Anwendung und klicken Sie auf &quot;Öffnen&quot;. </li><li> Alle Dateitypen, die &quot;. Csv-Datei gespeichert&quot;-Display, auf die Datei. </li></ol></li><li> Data Processing <ol style="padding:5px;"><li> Es gibt mehrere Möglichkeiten, um die Daten in der Excel-Datei zu verarbeiten. Eine gängige Praxis ist, die durchschnittliche und die Standardabweichung zu erhalten. </li><li> Löschen Messungen, wenn die Standardabweichung größer als 30 ist. </li><li> Darüber hinaus kann abgelehnt Zyklen angesehen werden und die gesamte Datenbank kann einfach nach Excel exportiert werden. Wenden Sie sich an die Bedienungsanleitung für weitere Informationen. </li></ol></li></ol></li></ol></li></ol>

The authors have nothing to disclose.

measuring blood pressure in mice using volume pressure recording, a tail-cuff method

Die CODA 8-Kanal High-Throughput-Non-Invasive Blood Pressure System misst den Blutdruck in bis zu 8 Mäuse oder Ratten gleichzeitig. Die CODA tail-cuff-System verwendet Volume Druck Recording (VPR), um den Blutdruck durch die Bestimmung der Schwanz Blutvolumen messen. Eine speziell entwickelte Differenzdruck-Messumformer und eine Okklusion tail-cuff messen die gesamte Blutvolumen in den Schwanz, ohne die Notwendigkeit, die einzelnen Pulse Signal zu erhalten. Besonderes Augenmerk wird auf die Länge der Okklusion Manschette gewährt, um daraus die genauesten Blutdruckwerte. VPR kann leicht zu erhalten Lesungen auf dunkelhäutige Nagetiere wie Mäuse C57BL6 und MRI-kompatibel. Die CODA-System bietet Ihnen mit Messungen von sechs (6) verschiedene Parameter Blutdruck, systolischer und diastolischer Blutdruck, Herzfrequenz, der mittlere Blutdruck, Schwanz Durchblutung und Schwanz Blutvolumen. Die Messungen können entweder wach oder narkotisierten Mäusen oder Ratten durchgeführt werden. Die CODA-System enthält eine Steuereinheit, Laptop-Computer, Software, Manschetten, Tierhaltung, Infrarot-Wärmekabine Pads und einem Infrarot-Thermometer. Es gibt sieben verschiedene Halter Größen für Mäuse so klein wie 8 Gramm an Ratten so groß wie 900 Gramm.

Watch this Scientific Journal Video about Measuring Blood Pressure in Mice using Volume Pressure Recording, a Tail-cuff Method at JoVE.com

Measuring Blood Pressure in Mice using Volume Pressure Recording, a Tail-cuff Method

Die CODA 8-Kanal High-Throughput-Non-Invasive Blood Pressure System misst den Blutdruck in bis zu 8 Mäuse oder Ratten gleichzeitig. Dieser Schwanz-Manschette System nutzt Volume Druck Recording (VPR), um den Blutdruck durch die Bestimmung der Schwanz Blutvolumen messen.

Blutdruckmessung in Mäuse mit Volume Druck Recording, ein Tail-Manschette Methode

The CODA 8-Channel High Throughput Non-Invasive Blood Pressure system measures the blood pressure in up to 8 mice or rats simultaneously. The CODA ...

measuring-blood-pressure-mice-using-volume-pressure-recording-tail

Research

JoVE Journal

Biology

51.6K Aufrufe.  University of Kentucky. Die CODA 8-Kanal High-Throughput-Non-Invasive Blood Pressure System misst den Blutdruck in bis zu 8 Mäuse oder Ratten gleichzeitig. Dieser Schwanz-Manschette System nutzt Volume Druck Recording (VPR), um den Blutdruck durch die Bestimmung der Schwanz Blutvolumen messen.

Serie: Measuring Blood Pressure in Mice using Volume Pressure Recording, a Tail-cuff Method

Pressure-polishing Pipettes for Improved Patch-clamp Recording

Pressure-polishing is a method for shaping glass pipettes for patch-clamp recording. We first developed this method for fabricating pipettes suitable for recording from small (&lt;3 m) neuronal cell bodies. The basic principal is similar to glass-blowing and combines air pressure and heat to modify the shape of patch pipettes prepared by a conventional micropipette puller. It can be applied to so-called soft (soda lime) and hard (borosilicate) glasses. Generally speaking, pressure polishing can reduce pipette resistance by 25% without decreasing the diameter of the tip opening (Goodman and Lockery, 2000). It can be applied to virtually any type of glass and requires only the addition of a high-pressure valve and fitting to a microforge. This technique is essential for recording from ultrasmall cells (&lt;5 m) and can also improve single-channel recording by minimizing pipette resistance. The blunt shape is also useful for perforated-patch clamp recording since this tip shape results in a larger membrane bleb available for perforation.

This is a guide to modifying the shape of glass micropipettes. Specifically, by using heat and air pressure the taper is widened without increasing the tip opening, leading to lower pipette resistance. This is critical to obtain low noise recordings of small cells but is useful in many applications.

Pressure-Polieren Pipetten für verbesserte Patch-Clamp Recording

Pressure-polishing is a method for shaping glass pipettes for patch-clamp recording.  We first developed this method for fabricating pipettes suitable for ...

Forschung

Biologie

Making Gynogenetic Diploid Zebrafish by Early Pressure

Heterozygosity in diploid eukaryotes often makes genetic studies cumbersome. Methods that produce viable homozygous diploid offspring directly from heterozygous females allow F1 mutagenized females to be screened directly for deleterious mutations in an accelerated forward genetic screen. Streisinger et al.1,2 described methods for making gynogenetic (homozygous) diploid zebrafish by activating zebrafish eggs with ultraviolet light-inactivated sperm and preventing either the second meiotic or the first zygotic cell division using physical treatments (heat or pressure) that deploymerize microtubules. The "early pressure" (EP) method blocks the meiosis II, which occurs shortly after fertilization. The EP method produces a high percentage of viable embryos that can develop to fertile adults of either sex. The method generates embryos that are homozygous at all loci except those that were separated from their centromere by recombination during meiosis I. Homozygous mutations are detected in EP clutches at between 50% for centromeric loci and less than 1% for telomeric loci. This method is reproduced verbatim from the Zebrafish Book3.

This is a method for generating gynogenetic diploid zebrafish embryos (embryos whose only genetic contribution comes from the mother) by blocking the second meiotic division immediately after fertilization with ultraviolet light-inactivated sperm. EP embryos are not fully homozygous due to recombination during the first meiotic division, however they are homozygous at all loci that have not been separated from their centromere by recombination.

Machen Gynogenetic Diploid Zebrafisch von Anfang Druck

Heterozygosity in diploid eukaryotes often makes genetic studies cumbersome. Methods that produce viable homozygous diploid offspring directly from ...

Ambulatory ECG Recording in Mice

Telemetric ECG recording in mice is essential to understanding the mechanisms behind arrhythmias, conduction disorders, and sudden cardiac death. Although the surface ECG is utilized for short-term measurements of waveform intervals, it is not practical for long-term studies of heart rate variability or the capture of rare episodes of arrhythmias. Implantable ECG telemeters offer the advantages of simple surgical implantation, long-term recording of electrograms in ambulatory mice, and scalability with simultaneous recordings of multiple animals. Here, we present a step-by-step guide to the implantation of telemeters for ambulatory ECG recording in mice. Careful adherence to aseptic technique is required for favorable survival results with the possibility of implantation and recording from weeks to months. Thus, implantable ECG telemetry is a valuable tool for detection of critical information on cardiac electrophysiology in ambulatory animal models such as the mouse. 

Telemetric ECG has emerged as an essential tool in evaluating animal models for cardiac arrhythmias and sudden cardiac death. Here, we present a stepwise guide to telemetric ECG recordings for application in long-term ambulatory ECG monitoring in mice.

Ambulante EKG-Aufzeichnung in Mäuse

Telemetric ECG recording in mice is essential to understanding the mechanisms behind arrhythmias, conduction disorders, and sudden cardiac death. Although ...

Isolation and Culture of Pulmonary Endothelial Cells from Neonatal Mice

Endothelial cells provide a useful research model in many areas of vascular biology. Since its first isolation 1, human umbilical vein endothelial cells (HUVECs) have shown to be convenient, easy to obtain and culture, and thus are the most widely studied endothelial cells. However, for research focused on processes like angiogenesis, permeability or many others, microvascular endothelial cells (ECs) are a much more physiologically relevant model to study 2. Furthermore, ECs isolated from knockout mice provide a useful tool for analysis of protein function ex vivo. Several approaches to isolate and culture microvascular ECs of different origin have been reported to date 3-7, but consistent isolation and culture of pure ECs is still a major technical problem in many laboratories. Here, we provide a step-by-step protocol on a reliable and relatively simple method of isolating and culturing mouse lung endothelial cells (MLECs). In this approach, lung tissue obtained from 6- to 8-day old pups is first cut into pieces, digested with collagenase/dispase (C/D) solution and dispersed mechanically into single-cell suspension. MLECS are purified from cell suspension using positive selection with anti-PECAM-1 antibody conjugated to Dynabeads using a Magnetic Particle Concentrator (MPC). Such purified cells are cultured on gelatin-coated tissue culture (TC) dishes until they become confluent. At that point, cells are further purified using Dynabeads coupled to anti-ICAM-2 antibody. MLECs obtained with this protocol exhibit a cobblestone phenotype, as visualized by phase-contrast light microscopy, and their endothelial phenotype has been confirmed using FACS analysis with anti-VE-cadherin 8 and anti-VEGFR2 9 antibodies and immunofluorescent staining of VE-cadherin. In our hands, this two-step isolation procedure consistently and reliably yields a pure population of MLECs, which can be further cultured. This method will enable researchers to take advantage of the growing number of knockout and transgenic mice to directly correlate in vivo studies with results of in vitro experiments performed on isolated MLECs and thus help to reveal molecular mechanisms of vascular phenotypes observed in vivo.

Here, we describe a protocol for isolation and culture of murine pulmonary endothelial cells. This method comprises mechanic and enzymatic lung tissue dissociation as well as a 2-step purification process using anti-PECAM-1 and anti-ICAM-2 antibodies conjugated to magnetic beads, which produces a pure endothelial cell population of mostly microvascular origin.

Isolierung und Kultivierung von pulmonaler Endothelzellen aus neugeborenen Mäusen

Endothelial cells provide a useful research model in many areas of vascular biology. Since its first isolation 1, human umbilical vein endothelial cells ...

Assessing Murine Resistance Artery Function Using Pressure Myography

Pressure myograph systems are exquisitely useful in the functional assessment of small arteries, pressurized to a suitable transmural pressure. The near physiological condition achieved in pressure myography permits in-depth characterization of intrinsic responses to pharmacological and physiological stimuli, which can be extrapolated to the in vivo behavior of the vascular bed. Pressure myograph has several advantages over conventional wire myographs. For example, smaller resistance vessels can be studied at tightly controlled and physiologically relevant intraluminal pressures. Here, we study the ability of 3rd order mesenteric arteries (3-4 mm long), preconstricted with phenylephrine, to vaso-relax in response to acetylcholine. Mesenteric arteries are mounted on two cannulas connected to a pressurized and sealed system that is maintained at constant pressure of 60 mmHg. The lumen and outer diameter of the vessel are continuously recorded using a video camera, allowing real time quantification of the vasoconstriction and vasorelaxation in response to phenylephrine and acetylcholine, respectively.
	To demonstrate the applicability of pressure myography to study the etiology of cardiovascular disease, we assessed endothelium-dependent vascular function in a murine model of systemic hypertension. Mice deficient in the &alpha;1 subunit of soluble guanylate cyclase (sGC&alpha;1-/-) are hypertensive when on a 129S6 (S6) background (sGC&alpha;1-/-S6) but not when on a C57BL/6 (B6) background (sGC&alpha;1-/-B6). Using pressure myography, we demonstrate that sGC&alpha;1-deficiency results in impaired endothelium-dependent vasorelaxation. The vascular dysfunction is more pronounced in sGC&alpha;1-/-S6 than in sGC&alpha;1-/-B6 mice, likely contributing to the higher blood pressure in sGC&alpha;1-/-S6 than in sGC&alpha;1-/-B6 mice. 
	Pressure myography is a relatively simple, but sensitive and mechanistically useful technique that can be used to assess the effect of various stimuli on vascular contraction and relaxation, thereby augmenting our insight into the mechanisms underlying cardiovascular disease.

In pressure myography, an intact small segment of a vessel is mounted onto two small cannulas and pressurized to a suitable luminal pressure. Here, we describe the method to measure vasorelaxation response of the mouse 3rd order mesenteric arteries in c57 and sGC&alpha;1-/- mice using pressure myography.

Beurteilung Murine Widerstand Artery Funktion Mit Druck Myographie

Pressure myograph systems are  exquisitely useful in the functional assessment of small arteries, pressurized  to a suitable transmural pressure. The  ...

One-channel Cell-attached Patch-clamp Recording

Ion channel proteins are universal devices for fast communication across biological membranes. The temporal signature of the ionic flux they generate depends on properties intrinsic to each channel protein as well as the mechanism by which it is generated and controlled and represents an important area of current research. Information about the operational dynamics of ion channel proteins can be obtained by observing long stretches of current produced by a single molecule. Described here is a protocol for obtaining one-channel cell-attached patch-clamp current recordings for a ligand gated ion channel, the NMDA receptor, expressed heterologously in HEK293 cells or natively in cortical neurons. Also provided are instructions on how to adapt the method to other ion channels of interest by presenting the example of the mechano-sensitive channel PIEZO1. This method can provide data regarding the channel&#8217;s conductance properties and the temporal sequence of open-closed conformations that make up the channel&#8217;s activation mechanism, thus helping to understand their functions in health and disease.

Described here is a procedure for obtaining long stretches of current recording from one ion channel with the cell-attached patch-clamp technique. This method allows for observing, in real time, the pattern of open-close channel conformations that underlie the biological signal. These data inform about channel properties in undisturbed biological membranes.

Ein-Kanal-Cell-Attached-Patch-Clamp-Aufnahme

Ion channel proteins are universal devices for fast communication across biological membranes. The temporal signature of the ionic flux they generate ...

Measuring Respiratory Function in Mice Using Unrestrained Whole-body Plethysmography

Respiratory dysfunction is one of the leading causes of morbidity and mortality in the world and the rates of mortality continue to rise. Quantitative assessment of lung function in rodent models is an important tool in the development of future therapies. Commonly used techniques for assessing respiratory function including invasive plethysmography and forced oscillation. While these techniques provide valuable information, data collection can be fraught with artefacts and experimental variability due to the need for anesthesia and/or invasive instrumentation of the animal. In contrast, unrestrained whole-body plethysmography (UWBP) offers a precise, non-invasive, quantitative way by which to analyze respiratory parameters. This technique avoids the use of anesthesia and restraints, which is common to traditional plethysmography techniques. This video will demonstrate the UWBP procedure including the equipment set up, calibration and lung function recording. It will explain how to analyze the collected data, as well as identify experimental outliers and artefacts that results from animal movement. The respiratory parameters obtained using this technique include tidal volume, minute volume, inspiratory duty cycle, inspiratory flow rate and the ratio of inspiration time to expiration time. UWBP does not rely on specialized skills and is inexpensive to perform. A key feature of UWBP, and most appealing to potential users, is the ability to perform repeated measures of lung function on the same animal.

The assessment of respiratory physiology has traditionally relied upon techniques, which require restraint or sedation of the animal. Unrestrained whole-body plethysmography, however, provides precise, non-invasive, quantitative analysis of respiratory physiology in animal models. In addition, the technique allows repeated respiratory assessment of mice allowing for longitudinal studies.

Messatemfunktion im Maus Mit Hemmungslose Ganzkörper-Plethysmographie

Respiratory dysfunction is one of the leading causes of morbidity and mortality in the world and the rates of mortality continue to rise. Quantitative ...

A Method for Screening and Validation of Resistant Mutations Against Kinase Inhibitors

The discovery of BCR/ABL as a driver oncogene in chronic myeloid leukemia (CML) resulted in the development of Imatinib, which, in fact, demonstrated the potential of targeting the kinase in cancers by effectively treating the CML patients. This observation revolutionized drug development to target the oncogenic kinases implicated in various other malignancies, such as, EGFR, B-RAF, KIT and PDGFRs. However, one major drawback of anti-kinase therapies is the emergence of drug resistance mutations rendering the target to have reduced or lost affinity for the drug. Understanding the mechanisms employed by resistant variants not only helps in developing the next generation inhibitors but also gives impetus to clinical management using personalized medicine. We reported a retroviral vector based screening strategy to identify the spectrum of resistance conferring mutations in BCR/ABL, which has helped in developing the next generation BCR/ABL inhibitors. Using Ruxolitinib and JAK2 as a drug target pair, here we describe in vitro screening methods that utilizes the mouse BAF3 cells expressing the random mutation library of JAK2 kinase.

Emergence of genetic resistance against kinase inhibitor therapy poses significant challenge for effective cancer therapy. Identification and characterization of resistant mutations against a newly developed drug helps in better clinical management and next generation drug design. Here, we describe our protocol for in vitro screening and validation of resistant mutations.

Verfahren zum Screening und Validierung von Beständig Mutationen Vor-Inhibitoren

The discovery of BCR/ABL as a driver oncogene in chronic myeloid leukemia (CML) resulted in the development of Imatinib, which, in fact, demonstrated the ...

Cardiac Pressure-Volume Loop Analysis Using Conductance Catheters in Mice

Cardiac pressure-volume loop analysis is the &#8220;gold-standard&#8221; in the assessment of load-dependent and load-independent measures of ventricular systolic and diastolic function. Measures of ventricular contractility and compliance are obtained through examination of cardiac response to changes in afterload and preload. These techniques were originally developed nearly three decades ago to measure cardiac function in large mammals and humans. The application of these analyses to small mammals, such as mice, has been accomplished through the optimization of microsurgical techniques and creation of conductance catheters. Conductance catheters allow for estimation of the blood pool by exploiting the relationship between electrical conductance and volume. When properly performed, these techniques allow for testing of cardiac function in genetic mutant mouse models or in drug treatment studies. The accuracy and precision of these studies are dependent on careful attention to the calibration of instruments, systematic conduct of hemodynamic measurements and data analyses. We will review the methods of conducting pressure-volume loop experiments using a conductance catheter in mice.

Cardiac pressure-volume loop analysis is the most comprehensive way to measure cardiac function in the intact heart. We describe a technique to perform and analyze cardiac pressure volume loops, using conductance catheters.

Cardiac Druck-Volumen-Loop-Analyse mit Conductance-Katheter bei Mäusen

Cardiac pressure-volume loop analysis is the &#8220;gold-standard&#8221; in the assessment of load-dependent and load-independent measures of ventricular ...

Measuring Pressure Volume Loops in the Mouse

Understanding the causes and progression of heart disease presents a significant challenge to the biomedical community. The genetic flexibility of the mouse provides great potential to explore cardiac function at the molecular level. The mouse&#39;s small size does present some challenges in regards to performing detailed cardiac phenotyping. Miniaturization and other advancements in technology have made many methods of cardiac assessment possible in the mouse. Of these, the simultaneous collection of pressure and volume data provides a detailed picture of cardiac function that is not available through any other modality. Here a detailed procedure for the collection of pressure-volume loop data is described. Included is a discussion of the principles underlying the measurements and the potential sources of error. Anesthetic management and surgical approaches are discussed in great detail as they are both critical to obtaining high quality hemodynamic measurements. The principles of hemodynamic protocol development and relevant aspects of data analysis are also addressed.

This manuscript describes a detailed protocol for the collection of pressure-volume data from the mouse.

Mess Druck Volumen Loops in der Maus

Understanding the causes and progression of heart disease presents a significant challenge to the biomedical community. The genetic flexibility of the ...