<html><ol style="padding:5px;;text-align:right;direction:rtl"><li style=";text-align:right;direction:rtl"> ציוד הקמה <ol style="padding:5px;;text-align:right;direction:rtl"><li style=";text-align:right;direction:rtl"> בחר אזור במעבדה שבה בטמפרטורת החדר הוא בבית או מעל 20 ° C. </li><li style=";text-align:right;direction:rtl"> הימנע מקומות ליד פתחי אוורור ומיזוג ו / או רעש חזק. </li><li style=";text-align:right;direction:rtl"> הימנע לחשוף את בעל החיים ריחות שעלולים לגרום לגירוי / או לחץ, כמו בושם, בושם, כימיקלים, וניקיון. </li><li style=";text-align:right;direction:rtl"> הפעל את הבקר CODA. </li><li style=";text-align:right;direction:rtl"> הפעל את פלטפורמת מחמם להגדיר רמה 3. אם טמפרטורת החדר היא מעל 25 מעלות צלזיוס, להתחיל רמות 1 או 2. </li><li style=";text-align:right;direction:rtl"> בקר אבחון מבחן <ol style="padding:5px;;text-align:right;direction:rtl"><li style=";text-align:right;direction:rtl"> פתח את התוכנה CODA. </li><li style=";text-align:right;direction:rtl"> בחר את ההתקנים CODA ידי לחיצה עליו. </li><li style=";text-align:right;direction:rtl"> לחץ על &quot;התקן מבחן נבחרים&quot;. </li><li style=";text-align:right;direction:rtl"> בחר את האזיקים ערוצי לבדוק ולחץ על &quot;מבחן&quot;. </li><li style=";text-align:right;direction:rtl"> לאחר השלמת הבדיקה, לסגור את החלון מבחן ההתקנים. </li></ol></li></ol></li><li style=";text-align:right;direction:rtl"> תוכנה הקמה <ol style="padding:5px;;text-align:right;direction:rtl"><li style=";text-align:right;direction:rtl"> ניהול כוח אדם ובעלי חיים <ol style="padding:5px;;text-align:right;direction:rtl"><li style=";text-align:right;direction:rtl"> ניהול כוח אדם <ol style="padding:5px;;text-align:right;direction:rtl"><li style=";text-align:right;direction:rtl"> בחר כלים&gt; ניהול כוח אדם. </li><li style=";text-align:right;direction:rtl"> לחץ על &quot;חוקר 1&quot; כדי לשנות את השם. </li><li style=";text-align:right;direction:rtl"> לחץ על השורות הבאות כדי להוסיף חוקרים נוספים. </li><li style=";text-align:right;direction:rtl"> לחץ על &quot;Tab טכנאים&quot; לשנות או להוסיף טכנאים. </li><li style=";text-align:right;direction:rtl"> לחץ על &quot;Tab דוגמאות&quot; על שם בעלי חיים לצורך זיהוי קל לבחור את סוג החיה, חולדה או עכבר. </li><li style=";text-align:right;direction:rtl"> לחץ על &quot;Tab בעלי חיים&quot; כדי לשנות את סוג החיה ורגישות. </li></ol></li></ol></li><li style=";text-align:right;direction:rtl"> ניסוי חדש <ol style="padding:5px;;text-align:right;direction:rtl"><li style=";text-align:right;direction:rtl"> בחר קובץ&gt; ניסוי חדש&gt; </li><li style=";text-align:right;direction:rtl"> בגין ניסוי חדש <ol style="padding:5px;;text-align:right;direction:rtl"><li style=";text-align:right;direction:rtl"> הזן שם ניסוי. </li><li style=";text-align:right;direction:rtl"> בחר את חוקר מפתח. </li><li style=";text-align:right;direction:rtl"> בחר תאריך בגין ולחץ על &quot;הבא&quot;. </li></ol></li><li style=";text-align:right;direction:rtl"> מושב בסיסי מידע <ol style="padding:5px;;text-align:right;direction:rtl"><li style=";text-align:right;direction:rtl"> הזן את שם מושב </li><li style=";text-align:right;direction:rtl"> הגדרת &quot;מחזורי הסתגלות&quot; עד 5. אם אתה מעדיף שלא לכלול מחזורים הסתגלות, לאפס. כדי לסייע עם הסתגלות החיה לבעל וחפתים, להוסיף מחזורים נוספים הניסוי שלך ולהפעיל את הניסוי במשך 5 ימים רצופים. </li><li style=";text-align:right;direction:rtl"> הגדרת &quot;מספר סטים&quot; 1. </li><li style=";text-align:right;direction:rtl"> הגדרת &quot;זמן בין הסטים&quot; עד 30 שניות עבור קבוצות מרובות </li><li style=";text-align:right;direction:rtl"> הגדרת &quot;מחזורים לכל הגדר&quot; עד 20 מחזורים. </li><li style=";text-align:right;direction:rtl"> הגדרת &quot;זמן בין מחזורי&quot; עד 5 שניות ולחץ על &quot;הבא&quot;. </li></ol></li></ol></li><li style=";text-align:right;direction:rtl"> הדגימה לבחירה <ol style="padding:5px;;text-align:right;direction:rtl"><li style="list-style-type:none;;text-align:right;direction:rtl"><ol style="padding:5px;;text-align:right;direction:rtl"><li style=";text-align:right;direction:rtl"> בחר את הדגימה ממאגר הדגימה להקצות אותו הערוץ הזמין הראשון ולחץ על &quot;הבא&quot;. </li></ol></li><li style=";text-align:right;direction:rtl"> מושב פרמטרים <ol style="padding:5px;;text-align:right;direction:rtl"><li style=";text-align:right;direction:rtl"> הגדרת &quot;לחץ ספיגה מירבית&quot; כדי 250mmHg. </li><li style=";text-align:right;direction:rtl"> הגדרת &quot;זמן דפלציה&quot; עד 20 שניות עבור עכברים, חולדות במשך 15 שניות. </li><li style=";text-align:right;direction:rtl"> הגדרת &quot;כרך מינימום&quot; כדי 15ul. </li><li style=";text-align:right;direction:rtl"> הגדרת &quot;סגנון תצוגה&quot; לערוץ אחת לכל גרף. </li><li style=";text-align:right;direction:rtl"> הוסף חוקרים נוספים טכנאים ולחץ על &quot;הבא&quot;. </li></ol></li><li style=";text-align:right;direction:rtl"> מושב סקריפט <ol style="padding:5px;;text-align:right;direction:rtl"><li style=";text-align:right;direction:rtl"> סקור את סקריפט מושב ולחץ על &quot;הבא&quot;. </li><li style=";text-align:right;direction:rtl"> אל תבחר סיום עד שתשלים את סעיף הכנת בעלי חיים. </li></ol></li></ol></li></ol></li><li style=";text-align:right;direction:rtl"> הכנת בעלי חיים <ol style="padding:5px;;text-align:right;direction:rtl"><li style=";text-align:right;direction:rtl"> בעלי חיים הקמה <ol style="padding:5px;;text-align:right;direction:rtl"><li style=";text-align:right;direction:rtl"> בעלי חיים המחזיק <ol style="padding:5px;;text-align:right;direction:rtl"><li style=";text-align:right;direction:rtl"> כדי לאמן ולשפר הסתגלות בעלי חיים, להוסיף מחזורים נוספים הניסוי שלך ולהפעיל את הניסוי במשך חמישה ימים רצופים. אתה גם יכול לסייע הסתגלות ידי הצבת בעלי חיים בעל במשך 15 דקות במשך 3 ימים ברציפות לפני המחקר בפועל. </li><li style=";text-align:right;direction:rtl"> המקום כל חיה בעל ידי להרים את החיה בזנב בעדינות הצבת חיה לתוך האחורי של המחזיק הפונה את הקצה הפתוח של חרוט האף. הערה: גודל של חיה ביחס לגודל של בעל שנבחר. בעלי חיים צריכים להיות מסוגלים להיכנס בחופשיות בעל. </li><li style=";text-align:right;direction:rtl"> בזהירות לאבטח את הפתח האחורי לבעל על ידי סיבוב הבורג כובע אדום על הפתח האחורי. יש להקפיד להימנע צובט את הזנב או חלקי גוף אחרים תוך הבטחת הפתח האחורי. </li><li style=";text-align:right;direction:rtl"> החלק את חרטומו כלפי הפתח האחורי, הגבלת תנועה של החיה. חרטומו צריך להיות בעמדה כדי להגביל את בעלי החיים מן להסתובב בתוך זמן מחזיק. </li><li style=";text-align:right;direction:rtl"> המקום בעל לבמה התחממות במצב הייעודי. אם בודקים את אותה קבוצה של בעלי חיים על פני תקופה של זמן, כדי להיות בטוח לסובב את עמדת המחזיק בפלטפורמת ההתחממות לשפר עקביות ודיוק של מדידות לחץ הדם שלך. אפשר את החיה לפחות 5 דקות להסתגל לבעל. </li><li style=";text-align:right;direction:rtl"> אין לגעת או לטפל בבעלי חיים תוך בתוך לבעל. הקשר גדל יכול להרגיז את בעלי החיים. </li><li style=";text-align:right;direction:rtl"> אין להשאיר את בעל החיים של מחזיקי ללא השגחה. </li></ol></li><li style=";text-align:right;direction:rtl"> קאף מיקום <ol s</html style=";text-align:right;direction:rtl">tyle = &quot;padding: 5px;&quot;&gt; <li style=";text-align:right;direction:rtl"> נושא הזנב באמצעות &quot;קאף ספיגה.&quot; המקום הזה השרוול קרוב לבסיס של הזנב ככל האפשר בלי כוח. </li><li style=";text-align:right;direction:rtl"> אבטח את &quot;קאף Tubing ספיגה&quot; ב החריץ על החלק האחורי העליון של המחזיק. </li><li style=";text-align:right;direction:rtl"> נושא הזנב דרך &quot;קאף הציבורי של ורמונט&quot;, הצבתו בתוך 2mm של &quot;קאף ספיגה&quot;. </li><li style=";text-align:right;direction:rtl"> אבטח את &quot;Tubing הציבורי של ורמונט קאף&quot; בתוך החריץ על החלק האחורי העליון של המחזיק. </li><li style=";text-align:right;direction:rtl"> צרף את האזיקים לבקר CODA. </li></ol></li><li style=";text-align:right;direction:rtl"> בעלי חיים טמפרטורה <ol style="padding:5px;;text-align:right;direction:rtl"><li style=";text-align:right;direction:rtl"> אפשר את החיה לפחות 5 דקות thermoregulate. </li><li style=";text-align:right;direction:rtl"> רשום את הטמפרטורה של החיה לעתים קרובות עם מדחום אינפרא אדום. הטמפרטורה צריכה להיות חיה 32-35 ° C. </li></ol></li></ol></li></ol></li><li style=";text-align:right;direction:rtl"> ניסוי <ol style="padding:5px;;text-align:right;direction:rtl"><li style=";text-align:right;direction:rtl"> בגין הניסוי <ol style="padding:5px;;text-align:right;direction:rtl"><li style=";text-align:right;direction:rtl"> לחץ על &quot;סיום&quot; ו הניסוי יחל באופן מיידי. </li></ol></li><li style=";text-align:right;direction:rtl"> סיום הניסוי <ol style="padding:5px;;text-align:right;direction:rtl"><li style=";text-align:right;direction:rtl"> בעל חיים <ol style="padding:5px;;text-align:right;direction:rtl"><li style=";text-align:right;direction:rtl"> מיד להסיר את החיה מן האזיקים מחזיק. </li><li style=";text-align:right;direction:rtl"> מחזירים את החיה לכלוב שלהם לאחר &quot;מושב פשוט דוח סיכום&quot; מוצג. </li></ol></li><li style=";text-align:right;direction:rtl"> איסוף נתונים <ol style="padding:5px;;text-align:right;direction:rtl"><li style=";text-align:right;direction:rtl"> לאחר השלמת המחזור האחרון מחזורי קיבל יוצג באופן אוטומטי בפורמט אלקטרוני בתוך היישום CODA, ללחוץ על כפתור &quot;ייצוא ל-Excel&quot;. </li><li style=";text-align:right;direction:rtl"> בחר את התיקייה כדי למקם את הקובץ, שם הקובץ ולאחר מכן לחץ על &quot;שמור&quot;. </li><li style=";text-align:right;direction:rtl"> פתח את היישום Excel ולחץ על &quot;פתח&quot;. </li><li style=";text-align:right;direction:rtl"> הצג את כל סוגי הקבצים כדי להציג את &quot;. Csv הקובץ שנשמר&quot;, לחץ על הקובץ. </li></ol></li><li style=";text-align:right;direction:rtl"> עיבוד נתונים <ol style="padding:5px;;text-align:right;direction:rtl"><li style=";text-align:right;direction:rtl"> ישנן כמה דרכים לעבד את הנתונים בתוך הקובץ Excel. מנהג נפוץ הוא לקבל את סטיית תקן ממוצעת. </li><li style=";text-align:right;direction:rtl"> מחיקת מדידות אם סטיית התקן גדולה יותר 30. </li><li style=";text-align:right;direction:rtl"> בנוסף, מחזורי דחה ניתן לראות את כל הנתונים ניתן לייצא בקלות ל-Excel. היוועץ במדריך למשתמש לקבלת מידע נוסף. </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

ערוץ 8-CODA תפוקה גבוהה לא פולשני לחץ דם מערכת מודד את לחץ הדם עד 8 עכברים או חולדות בו זמנית. מערכת זנב השרוול CODA משתמשת בלחץ נפח הקלטה (הציבורי של ורמונט) כדי למדוד את לחץ הדם על ידי קביעת נפח הדם הזנב. לחץ שתוכנן במיוחד ההפרש מתמר ואת אמצעי זנב השרוול חסימה נפח הדם הכולל בזנב ללא צורך לקבל את האות הדופק הפרט. תשומת לב מיוחדת הניתנת אורך השרוול חסימה על מנת להפיק הלחץ קריאה מדויקת ביותר דם. הרדיו הציבורי של ורמונט יכול בקלות לקבל קריאות על כהי עור מכרסמים, כגון C57BL6 עכברים הוא תואם ה-MRI. מערכת CODA מספק לך מדידות של שישה (6) שונות הפרמטרים לחץ דם, לחץ דם סיסטולי ודיאסטולי, קצב הלב, כלומר לחץ הדם, זרימת הדם הזנב, ודם נפח הזנב. מדידות יכול להתבצע גם על עכברים או חולדות ער או מורדם. המערכת כוללת בקר CODA, מחשב נייד, תוכנה, חפתים, בעלי חיים, כריות חימום אינפרא אדום, מד חום אינפרא אדום. ישנם שבעה גדלים שונים עבור בעל עכברים קטנים כמו 8 גרם לחולדות גדולים כמו 900 גרם.

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

ערוץ 8-CODA תפוקה גבוהה לא פולשני לחץ דם מערכת מודד את לחץ הדם עד 8 עכברים או חולדות בו זמנית. מערכת זו זנב השרוול משתמשת בלחץ נפח הקלטה (הציבורי של ורמונט) כדי למדוד את לחץ הדם על ידי קביעת נפח הדם הזנב.

מדידת לחץ דם בעכברים באמצעות לחץ נפח הקלטה, שיטת זנב השרוול

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 Views.  University of Kentucky. ערוץ 8-CODA תפוקה גבוהה לא פולשני לחץ דם מערכת מודד את לחץ הדם עד 8 עכברים או חולדות בו זמנית. מערכת זו זנב השרוול משתמשת בלחץ נפח הקלטה (הציבורי של ורמונט) כדי למדוד את לחץ הדם על ידי קביעת נפח הדם הזנב.

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

Measuring Left Ventricular Pressure in Late Embryonic and Neonatal Mice

Blood pressure increases significantly during embryonic and postnatal development in vertebrate animals. In the mouse, blood flow is first detectable around embryonic day (E) 8.51. Systolic left ventricular (LV) pressure is 2 mmHg at E9.5 and 11 mmHg at E14.52. At these mid-embryonic stages, the LV is clearly visible through the chest wall for invasive pressure measurements because the ribs and skin are not fully developed. Between E14.5 and birth (approximately E21) imaging methods must be used to view the LV. After birth, mean arterial pressure increases from 30 - 70 mmHg from postnatal day (P) 2 - 353. Beyond P20, arterial pressure can be measured with solid-state catheters (i.e. Millar or Scisense). Before P20, these catheters are too big for developing mouse arteries and arterial pressure must be measured with custom pulled plastic catheters attached to fluid-filled pressure transducers3 or glass micropipettes attached to servo null pressure transducers4.
Our recent work has shown that the greatest increase in blood pressure occurs during the late embryonic to early postnatal period in mice5-7. This large increase in blood pressure may influence smooth muscle cell (SMC) phenotype in developing arteries and trigger important mechanotransduction events. In human disease, where the mechanical properties of developing arteries are compromised by defects in extracellular matrix proteins (i.e. Marfan's Syndrome8 and Supravalvular Aortic Stenosis9) the rapid changes in blood pressure during this period may contribute to disease phenotype and severity through alterations in mechanotransduction signals. Therefore, it is important to be able to measure blood pressure changes during late embryonic and neonatal periods in mouse models of human disease.
We describe a method for measuring LV pressure in late embryonic (E18) and early postnatal (P1 - 20) mice. A needle attached to a fluid-filled pressure transducer is inserted into the LV under ultrasound guidance. Care is taken to maintain normal cardiac function during the experimental protocol, especially for the embryonic mice. Representative data are presented and limitations of the protocol are discussed.

Measuring left ventricular pressure (LV) in embryonic and neonatal mice is described. Pressure is measured by inserting a needle connected to a fluid-filled transducer into the LV under ultrasound guidance. Care must be taken to maintain normal cardiac function during the experimental protocol.

מדידת לחץ חדרית השמאל עכברים עובריים ילודים המאוחרת

Blood pressure increases significantly during embryonic and postnatal development in vertebrate animals. In the mouse, blood flow is first detectable ...

Bioengineering

Right Ventricular Systolic Pressure Measurements in Combination with Harvest of Lung and Immune Tissue Samples in Mice

The function of the right heart is to pump blood through the lungs, thus linking right heart physiology and pulmonary vascular physiology. Inflammation is a common modifier of heart and lung function, by elaborating cellular infiltration, production of cytokines and growth factors, and by initiating remodeling processes 1.
Compared to the left ventricle, the right ventricle is a low-pressure pump that operates in a relatively narrow zone of pressure changes. Increased pulmonary artery pressures are associated with increased pressure in the lung vascular bed and pulmonary hypertension 2. Pulmonary hypertension is often associated with inflammatory lung diseases, for example chronic obstructive pulmonary disease, or autoimmune diseases 3. Because pulmonary hypertension confers a bad prognosis for quality of life and life expectancy, much research is directed towards understanding the mechanisms that might be targets for pharmaceutical intervention 4. The main challenge for the development of effective management tools for pulmonary hypertension remains the complexity of the simultaneous understanding of molecular and cellular changes in the right heart, the lungs and the immune system.
Here, we present a procedural workflow for the rapid and precise measurement of pressure changes in the right heart of mice and the simultaneous harvest of samples from heart, lungs and immune tissues. The method is based on the direct catheterization of the right ventricle via the jugular vein in close-chested mice, first developed in the late 1990s as surrogate measure of pressures in the pulmonary artery5-13. The organized team-approach facilitates a very rapid right heart catheterization technique. This makes it possible to perform the measurements in mice that spontaneously breathe room air. The organization of the work-flow in distinct work-areas reduces time delay and opens the possibility to simultaneously perform physiology experiments and harvest immune, heart and lung tissues.
The procedural workflow outlined here can be adapted for a wide variety of laboratory settings and study designs, from small, targeted experiments, to large drug screening assays. The simultaneous acquisition of cardiac physiology data that can be expanded to include echocardiography5,14-17 and harvest of heart, lung and immune tissues reduces the number of animals needed to obtain data that move the scientific knowledge basis forward. The procedural workflow presented here also provides an ideal basis for gaining knowledge of the networks that link immune, lung and heart function. The same principles outlined here can be adapted to study other or additional organs as needed.

A specific and rapid protocol to simultaneously investigate right heart function, lung inflammation, and the immune response is described as a learning tool. Video and figures describe physiology and microdissection techniques in an organized team-approach that is adaptable to be used for small to large sized studies.

מדידות נכונות חדרית סיסטולי לחץ בשילוב עם קציר של ריאה ודגימות רקמה חיסוניות בעכברים

The function of the right heart is to pump blood through the lungs, thus linking right heart physiology and pulmonary vascular physiology. Inflammation is ...

Immunology and Infection

Monitoring of Systemic and Hepatic Hemodynamic Parameters in Mice

The use of mouse models in experimental research is of enormous importance for the study of hepatic physiology and pathophysiological disturbances. However, due to the small size of the mouse, technical details of the intraoperative monitoring procedure suitable for the mouse were rarely described. Previously we have reported a monitoring procedure to obtain hemodynamic parameters for rats. Now, we adapted the procedure to acquire systemic and hepatic hemodynamic parameters in mice, a species ten-fold smaller than rats. This film demonstrates the instrumentation of the animals as well as the data acquisition process needed to assess systemic and hepatic hemodynamics in mice. Vital parameters, including body temperature, respiratory&#160;rate&#160;and heart&#160;rate were recorded throughout the whole procedure. Systemic hemodynamic parameters consist of carotid artery pressure (CAP) and central venous pressure (CVP). Hepatic perfusion parameters include portal vein pressure (PVP), portal flow rate as well as the flow rate of the common hepatic artery (table 1). Instrumentation and data acquisition to record the normal values was completed within 1.5 h. Systemic and hepatic hemodynamic parameters remained within normal ranges during this procedure.
This procedure is challenging but feasible. We have already applied this procedure to assess hepatic hemodynamics in normal mice as well as during 70% partial hepatectomy and in liver lobe clamping experiments. Mean PVP after resection (n= 20), was 11.41&#177;2.94 cmH2O which was significantly higher (P&#60;0.05) than before resection (6.87&#177;2.39 cmH2O). The results of liver lobe clamping experiment indicated that this monitoring procedure is sensitive and suitable for detecting small changes in portal pressure and portal flow rate. In conclusion, this procedure is reliable in the hands of an experienced micro-surgeon but should be limited to experiments where mice are absolutely needed.

This film demonstrates how to acquire systemic and hepatic hemodynamics in mice. The whole monitoring includes acquisition of vital parameters, systemic blood pressure, central venous pressure, common hepatic artery flow rate, and portal vein pressure as well as the portal flow rate in mice.

ניטור של פרמטרים מערכתיים והבדיה המודינמית בעכברים

The use of mouse models in experimental research is of enormous importance for the study of hepatic physiology and pathophysiological disturbances. ...

Medicine

Measuring Ascending Aortic Stiffness In Vivo in Mice Using Ultrasound

We present a protocol for measuring in vivo aortic stiffness in mice using high-resolution ultrasound imaging. Aortic diameter is measured by ultrasound and aortic blood pressure is measured invasively with a solid-state pressure catheter. Blood pressure is raised then lowered incrementally by intravenous infusion of vasoactive drugs phenylephrine and sodium nitroprusside. Aortic diameter is measured for each pressure step to characterize the pressure-diameter relationship of the ascending aorta. Stiffness indices derived from the pressure-diameter relationship can be calculated from the data collected. Calculation of arterial compliance is described in this protocol.
This technique can be used to investigate mechanisms underlying increased aortic stiffness associated with cardiovascular disease and aging. The technique produces a physiologically relevant measure of stiffness compared to ex vivo approaches because physiological influences on aortic stiffness are incorporated in the measurement. The primary limitation of this technique is the measurement error introduced from the movement of the aorta during the cardiac cycle. This motion can be compensated by adjusting the location of the probe with the aortic movement as well as making multiple measurements of the aortic pressure-diameter relationship and expanding the experimental group size.

Measuring Ascending Aortic Stiffness In Vivo in Mice Using Ultrasound

We describe a technique for measuring aortic stiffness from its pressure-diameter relationship in vivo in mice. Aortic diameter is recorded by ultrasound and aortic pressure is measured invasively with a solid-state pressure catheter. Blood pressure is changed incrementally and the resulting diameter is measured.

מדידה עולה אבי העורקים נוקשים<em&gt; In vivo</em&gt; בעכברים באמצעות אולטראסאונד

We present a protocol for measuring in vivo aortic stiffness in mice using high-resolution ultrasound imaging. Aortic diameter is measured by ultrasound ...

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.

מדידה של עקומת לחץ-נפח ריאות עכבר

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.

צנתור לב בעכברים כדי למדוד את יחסי נפח לחץ: חקירת אפקט Bowditch

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

Long-term Blood Pressure Measurement in Freely Moving Mice Using Telemetry

During the development of new vasoactive agents, arterial blood pressure monitoring is crucial for evaluating the efficacy of the new proposed drugs. Indeed, research focusing on the discovery of new potential therapeutic targets using genetically altered mice requires a reliable, long-term assessment of the systemic arterial pressure variation. Currently, the gold standard for obtaining long-term measurements of blood pressure in ambulatory mice uses implantable radio-transmitters, which require&#160;artery cannulation. This technique eliminates the need for tethering, restraining, or anesthetizing the animals which introduce stress and artifacts during data sampling. However, arterial blood pressure monitoring in mice via catheterization can be rather challenging due to the small size of the arteries. Here we present a step-by-step guide to illustrate the crucial key passages for a successful subcutaneous implantation of radio-transmitters and carotid artery cannulation in mice. We also include examples of long-term blood pressure activity taken from freely moving mice after a period of post-surgery recovery. Following this procedure will allow reliable direct blood pressure recordings from multiple animals simultaneously.

The goal of this protocol is to assess systemic blood pressure in conscious freely moving mice using implantable radio-telemetry devices.

מדידת לחץ דם לטווח ארוך בעכברים לנוע בחופשיות באמצעות טלמטריה

During the development of new vasoactive agents, arterial blood pressure monitoring is crucial for evaluating the efficacy of the new proposed drugs. ...

Simultaneous Electrocardiography Recording and Invasive Blood Pressure Measurement in Rats

For studies related to cardiovascular physiology or pathophysiology, blood pressure (BP) and electrocardiography are basic observational parameters. Research focusing on cardiovascular disease models, potential cardiovascular therapeutic targets or pharmaceutical agents requires assessment of systemic arterial pressure and heart rhythm changes. In situations where radio telemetry systems are not available or affordable, the technique of femoral artery cannulation is an alternative way to obtain intra-arterial pressure waveform recordings and systemic BP measurements. This technique is economical and can be performed with standard equipment in animal facilities. However, invasive arterial pressure recording requires cannulation of small arteries, which can be a challenging surgical skill. Here, we present step-by-step protocols for femoral artery cannulation procedures. Key procedures include the calibration of the data acquisition system, tissue dissection and femoral artery cannulation, and setup of the arterial cannulation system for pressure recording. Surface electrocardiography recording procedures are also included. We also present examples of BP recordings from normotensive and hypertensive rats. This protocol allows reliable direct recordings of systemic BP with simultaneous electrocardiography.

Here, we describe a setup for simultaneous recording of electrocardiography and intra-arterial blood pressure (BP) in experimental rats, which can be done with standard equipment in animal facilities and can be applied to physiological or pharmacological studies to investigate pathogenic or therapeutic mechanisms in cardiovascular medicine.

ההקלטה סימולטני אלקטרוקרדיוגרם ודם פולשנית בלחץ מדידה בחולדות

For studies related to cardiovascular physiology or pathophysiology, blood pressure (BP) and electrocardiography are basic observational parameters. ...

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.

מדידת לולאות נפח לחץ העכבר

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

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.

מינון בולוס תוך ורידי מרובים והערכה המודינמית פולשנית במודל יתר לחץ דם של עורק ריאתי בעכבר הנגרם על ידי היפוקסיה

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