הצמד תיקון התקנה: אופטיקה: אופטיקה חשובים מאוד למצוא תא טובה, כדי לקבוע את המיקום של זה. נסה להשיג תמונה ברורה של התאים: קצוות חדים, רך למראה תא. סרוק את שדה שלם כדי לחפש טוב התאים העיקריים MNTB עם גביע המצורפת. סיבוב צלחת לחפש calyces מזוויות שונות. בהתחלה זה בסדר להתאמן על מסוף, בלון בצורת. פיק התא על פני השטח של פרוסה או שכבה 2 nd התא. היעד השלישי ראשון 1 / 3 תאים עם פיפטה זיהוי Calyxes: חפש קרום כפול (ראה איור 1), לסובב את המנה פרוסה לראות אותו מזוויות שונות כדי לעזור לאשר כי מדובר גביע. <img src="/files/ftp_upload/old/244_2008_8_22_2/244_Figure-01.png" alt="איור 1" width="298" height="159" /> איור 1 עומק Calyx: Calyx צריך להיות על פני השטח או קרוב לפני השטח ככל האפשר. שטח זמין עבור תיקון: השתמש עדין להתמקד על כפתור מיקרוסקופ כדי לקבוע כיצד &quot;רחב / עמוק&quot; גביע הוא. לדוגמה, ~ 10% סיבוב מלא על המוקד את הקנס ייתן לך יותר שטח מספיק על גבי טלאי. איטום על גבי תא presynaptic: כדי להגיע אל תא מצורף, לחפש עיוות &quot;קטן&quot; על פני התא עקב הלחץ pos (~ 0.15psi) ב פיפטה, להשתמש מניפולטור פיפטה לרוץ למעלה, למטה ולרוחב, ואפילו את פני השטח של הטרמינל presyn לקבל דפורמציה טוב, ואז לשחרר לחץ להחיל יניקה, אם על פי הצורך. PULSE, תוכנית רכישת נתונים: צריך להגדיל את גודל המאגר לפני השימוש הראשון, נדרש בכל פעם על ידי יכולת דגימה. צריך לטעון מאקרו שונה בכל פעם. צומת נוזלי אפשריים: הקלטה פתרון Presyn הוא-11mV (11mV שלילי). Presyn: Cslow = ~ 15 pF Rs comp עבור presyn: 10 μsec, 65% (פיצויים יותר עשוי אפשרי W / החוצה תנודות) לפני קיבול הקלטה: הרפיה הבדיקה הנוכחית (10 mV, 10 msec לקפוץ) כדי לראות אם היא הרפיה monoexponential, כלומר, אם המונח presyn יכול להיות המודל של תא יחיד. מסננים: הגדר בבית kHz 30 ו - 10 kHz (בסל מסנן 1 ו -2 בהתאמה) במהלך המדידה רגיעה, כדי לוודא מרכיב מהיר לא פיספס. במהלך ההקלטות מיני, המשמש Xinsheng 10 קילוהרץ ו 5kHz. LGW שאל אותי להשתמש רק להגדיר מסנן 1 ב 30 קילוהרץ כדי לוודא שאנחנו יכולים למדוד עליית פעמי של מיני. שמות קבצים: 7 ספרות 000mmxx, כאשר mm הוא החודש (או חודש, שנה) ו xx הוא מספר התא. ניקוי קווי הפתרון: קווים לנקות בסוף הניסוי עם 20 מ&quot;ל של HCl 0.1 M ואחריו 200 מ&quot;ל ddH2O. פתרונות חיצוניים הם ממוחזרים, מהירות המשאבה הוא 20-30 סל&quot;ד. (פתרון החוזרים קאמרית אמבטיה זורם חזרה לתוך צילינדר המדורג). הגדרת 3 קווי היוצא על מנת להבטיח יניקה מספיק. פיפטה פתרון הקלטה Presynaptic: ~ 310 מילי Osmolar הפתרון צריך להיות Presyn ~ 310 mOsm למנוע R-גישה להגדיל במהלך הניסוי. פתרון Presyn פיפטה שונה postsyn פיפטה פתרון (Presyn יש ATP GTP &amp; עבור לשעבר.). TTX-TEA פתרון (לבודד Ca זרמים): <ul style=";text-align:right;direction:rtl"><li style=";text-align:right;direction:rtl"> נסו ליישם לאחר איטום, או אפילו תיקון, על התא, כמו תה בלוקים הפתרון K-ערוצים depolarizes התא. זה לא טוב עבור התא. מאותה סיבה, זה לא יהיה ניתן לעשות שימוש חוזר פרוסה אחרי TTX מוחל. </li><li style=";text-align:right;direction:rtl"> השווה את התגובה הראשונה והאחרונה של התא, או פרוסה כדי לבדוק השפעה כלשהי של שלילת קוטביות ממושכת. </li><li style=";text-align:right;direction:rtl"> TTX משמש ב 1 מיקרומטר במקום 0.5 מיקרומטר ולכן ההשפעה היא מהירה יותר. צריך לחכות פחות זמן לאחר שינוי פתרונות. </li></ul> חיתוך Slices Vibratome הגדרות: <ul style=";text-align:right;direction:rtl"><li style=";text-align:right;direction:rtl"> 70Hz רעידות לרוחב </li><li style=";text-align:right;direction:rtl"> Amplitude: 1.0 מ&quot;מ </li><li style=";text-align:right;direction:rtl"> קדימה במהירות 0.01-0.02 מ&quot;מ / sec בזמן חיתוך פרוסות עם גביע, 0.1 אחרת עובי: 180 מיקרומטר (~ 150מיקרומטר בבעלי חיים מבוגרים) </li></ul> Slices לדגור על 37 º C למשך שעה 1 (כולל זמן חיתוך), כלומר, להשאיר באמבטיה 15-20 דקות לאחר חיתוך. Pipettes: Presyn: 3.5-4.5 MΩ בהתחלה, 3.0-3.5 MΩ כאשר מרגישים בטוחים יותר שימוש ידני חולץ להגדיר הגדרות. השתמש בתכונה עיכוב בגלל זכוכית עבה (עיכובים בעבודה 200 או 1) זכוכית סוג: בורוסיליקט, תקן הקיר, אין נימה קוטר חיצוני: 2.0 מ&quot;מ קוטר פנימי 1.16 מ&quot;מ אורך: 7.5 ס&quot;מ

methods for patch clamp capacitance recordings from the calyx

אנו מדגימים את הטכניקות הבסיסיות הקלטה תיקון presynaptic מהדק את גביע של הלד, מסוף יונקים העצבים המרכזית מערכת העצבים. הקלטות חשמל ממסוף presynaptic לאפשר מדידה של פוטנציאל פעולה, ערוץ זרמי סידן, איחוי שלפוחית ​​(exocytosis) ספיגת הקרום הבאים (אנדוציטוזה). שילוב של שלפוחית ​​המכילה הנוירוטרנסמיטר גורם קרום שלפוחית ​​שיתווספו קרום התא של גביע. גידול זה בהיקף של קרום התא הוא, כפי שהיא נמדדת עלייה קיבול. הירידה עוקבת קיבול מציין אנדוציטוזה, תהליך של ספיגת קרום או הסרה של קרום גביע. אנדוציטוזה, נחוצה כדי לשמור על המבנה של גביע והוא נחוץ גם כדי ליצור שלפוחית ​​כי יתמלאו הנוירוטרנסמיטר לאירועים exocytosis בעתיד. הקלטות קיבול על גביע של הלד אפשרו ישירות ובמהירות למדוד לשחרר שלפוחי ו אנדוציטוזה מאוחרות מסוף יונקים עצב במערכת העצבים המרכזית. בנוסף, הפעילות postsynaptic המקביל ניתן למדוד בו זמנית באמצעות הקלטות לזווג. לכן תמונה מלאה של הפעילות החשמלית presynaptic ו postsynaptic בסינפסה מערכת העצבים המרכזית הוא בר השגה באמצעות הכנה. כאן, שיטות להכנת פרוסה, תכונות מורפולוגיות לזיהוי calyces, בטכניקות נערך תיקון בסיסי clamping, ודוגמאות של הקלטות קיבול למדוד exocytosis ו אנדוציטוזה מוצגים.

Watch this Scientific Journal Video about Methods for Patch Clamp Capacitance Recordings from the Calyx at JoVE.com

Methods for Patch Clamp Capacitance Recordings from the Calyx

שיטות הצמד תיקון הקלטות קיבול מן Calyx

We demonstrate the basic techniques for presynaptic patch clamp recording at the calyx of Held, a mammalian central nervous system nerve terminal. ...

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12.6K Views.  National Institute of Health. 

Video: Methods for Patch Clamp Capacitance Recordings from the Calyx

Patch Clamp Recording of Ion Channels Expressed in  Xenopus Oocytes

Since its development by Sakmann and Neher 1, 2, the patch clamp has become established as an extremely useful technique for electrophysiological measurement of single or multiple ion channels in cells. This technique can be applied to ion channels in both their native environment and expressed in heterologous cells, such as oocytes harvested from the African clawed frog, Xenopus laevis. Here, we describe the well-established technique of patch clamp recording from Xenopus oocytes. This technique is used to measure the properties of expressed ion channels either in populations (macropatch) or individually (single-channel recording). We focus on techniques to maximize the quality of oocyte preparation and seal generation. With all factors optimized, this technique gives a probability of successful seal generation over 90 percent. The process may be optimized differently by every researcher based on the factors he or she finds most important, and we present the approach that have lead to the greatest success in our hands.

This is intended as an introduction to patch clamp recording from Xenopus laevis oocytes. It covers vitelline membrane removal, formation of a gigaohm seal (gigaseal), and the optional conversion of the patch to the outside-out topology.

הצמד תיקון הקלטה של ​​תעלות יונים מתבטא ביציות Xenopus

Since its development by Sakmann and Neher 1, 2, the patch clamp has become established as an extremely useful technique for electrophysiological ...

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.

לחץ לליטוש pipettes עבור Patch-clamp הקלטה משופרים

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

Electrophysiological Methods for Recording Synaptic Potentials from the NMJ of Drosophila Larvae

In this video, we describe the electrophysiological methods for recording synaptic transmission at the neuromuscular junction (NMJ) of Drosophila larva. The larval neuromuscular system is a model synapse for the study of synaptic physiology and neurotransmission, and is a valuable research tool that has defined genetics and is accessible to experimental manipulation. Larvae can be dissected to expose the body wall musculature, central nervous system, and peripheral nerves. The muscles of Drosophila and their innervation pattern are well characterized and muscles are easy to access for intracellular recording. Individual muscles can be identified by their location and orientation within the 8 abdominal segments, each with 30 muscles arranged in a pattern that is repeated in segments A2 - A7. Dissected drosophila larvae are thin and individual muscles and bundles of motor neuron axons can be visualized by transillumination1. Transgenic constructs can be used to label target cells for visual identification or for manipulating gene products in specific tissues. In larvae, excitatory junction potentials (EJP&#8217;s) are generated in response to vesicular release of glutamate from the motoneurons at the synapse. In dissected larvae, the EJP can be recorded in the muscle with an intracellular electrode. Action potentials can be artificially evoked in motor neurons that have been cut posterior to the ventral ganglion, drawn into a glass pipette by gentle suction and stimulated with an electrode. These motor neurons have distinct firing thresholds when stimulated, and when they fire simultaneously, they generate a response in the muscle. Signals transmitted across the NMJ synapse can be recorded in the muscles that the motor neurons innervate. The EJP&#8217;s and minature excitatory junction potentials (mEJP&#8217;s) are seen as changes in membrane potential. Electrophysiological responses are recorded at room temperature in modified minimal hemolymph-like solution2 (HL3) that contains 5 mM Mg2+ and 1.5 mM Ca2+. Changes in the amplitude of evoked EJP&#8217;s can indicate differences in synaptic function and structure. Digitized recordings are analyzed for EJP amplitude, mEJP frequency and amplitude, and quantal content.

Here we describe electrophysiological methods for measuring synaptic transmission at the neuromuscular junction of Drosophila larva. Evoked release is initiated artificially by stimulating the motor neuron axons, and transmission through the NMJ can be measured by the postsynaptic response evoked in the muscle.

שיטות אלקטרו עבור הקלטה פוטנציאלים Synaptic מן NMJ של הזחלים תסיסנית

In this video, we describe the electrophysiological methods for recording synaptic transmission at the neuromuscular junction (NMJ) of Drosophila larva. ...

Patch Clamp and Perfusion Techniques for Studying Ion Channels Expressed in Xenopus oocytes

The protocol presented here is designed to study the activation of the large conductance, voltage- and Ca2+-activated K+ (BK) channels. The protocol may also be used to study the structure-function relationship for other ion channels and neurotransmitter receptors1. BK channels are widely expressed in different tissues and have been implicated in many physiological functions, including regulation of smooth muscle contraction, frequency tuning of inner hair cells and regulation of neurotransmitter release2-6. BK channels are activated by membrane depolarization and by intracellular Ca2+ and Mg2+ 6-9. Therefore, the protocol is designed to control both the membrane voltage and the intracellular solution. In this protocol, messenger RNA of BK channels is injected into Xenopus laevis oocytes (stage V-VI) followed by 2-5 days of incubation at 18&deg;C10-13. Membrane patches that contain single or multiple BK channels are excised with the inside-out configuration using patch clamp techniques10-13. The intracellular side of the patch is perfused with desired solutions during recording so that the channel activation under different conditions can be examined. To summarize, the mRNA of BK channels is injected into Xenopus laevis oocytes to express channel proteins on the oocyte membrane; patch clamp techniques are used to record currents flowing through the channels under controlled voltage and intracellular solutions.

Patch Clamp and Perfusion Techniques for Studying Ion Channels Expressed in Xenopus oocytes

Ionic current of BK channels is recorded using patch clamp techniques. BK channels are expressed in Xenopus oocytes by injecting messenger RNA. The intracellular solution during patch clamp recordings is controlled by a perfusion system.

הצמד תיקון וטכניקות זלוף עבור לימוד תעלות יונים מתבטא Xenopus ביציות

The protocol presented here is designed to study the activation of the large conductance, voltage- and Ca2+-activated K+ (BK) channels. The protocol may ...

Exploring Arterial Smooth Muscle Kv7 Potassium Channel Function using Patch Clamp Electrophysiology and Pressure Myography

Contraction or relaxation of smooth muscle cells within the walls of resistance arteries determines the artery diameter and thereby controls flow of blood through the vessel and contributes to systemic blood pressure. The contraction process is regulated primarily by cytosolic calcium concentration ([Ca2+]cyt), which is in turn controlled by a variety of ion transporters and channels. Ion channels are common intermediates in signal transduction pathways activated by vasoactive hormones to effect vasoconstriction or vasodilation. And ion channels are often targeted by therapeutic agents either intentionally (e.g. calcium channel blockers used to induce vasodilation and lower blood pressure) or unintentionally (e.g. to induce unwanted cardiovascular side effects).
Kv7 (KCNQ) voltage-activated potassium channels have recently been implicated as important physiological and therapeutic targets for regulation of smooth muscle contraction. To elucidate the specific roles of Kv7 channels in both physiological signal transduction and in the actions of therapeutic agents, we need to study how their activity is modulated at the cellular level as well as evaluate their contribution in the context of the intact artery.
The rat mesenteric arteries provide a useful model system. The arteries can be easily dissected, cleaned of connective tissue, and used to prepare isolated arterial myocytes for patch clamp electrophysiology, or cannulated and pressurized for measurements of vasoconstrictor/vasodilator responses under relatively physiological conditions. Here we describe the methods used for both types of measurements and provide some examples of how the experimental design can be integrated to provide a clearer understanding of the roles of these ion channels in the regulation of vascular tone.

Measurements of Kv7 (KCNQ) potassium channel activity in isolated arterial myocytes (using patch clamp electrophysiological techniques) in parallel with measurements of constrictor/dilator responses (using pressure myography) can reveal important information about the roles of Kv7 channels in vascular smooth muscle physiology and pharmacology.

היכרות עם פונקצית עורקי שריר חלק Kv7 אשלגן ערוץ באמצעות אלקטרופיזיולוגיה קלאמפ תיקון וMyography לחץ

Contraction  or relaxation of smooth muscle cells within the walls of resistance arteries  determines the artery diameter and thereby controls flow of ...

Giant Liposome Preparation for Imaging and Patch-Clamp Electrophysiology

The reconstitution of ion channels into chemically defined lipid membranes for electrophysiological recording has been a powerful technique to identify and explore the function of these important proteins. However, classical preparations, such as planar bilayers, limit the manipulations and experiments that can be performed on the reconstituted channel and its membrane environment. The more cell-like structure of giant liposomes permits traditional patch-clamp experiments without sacrificing control of the lipid environment.
Electroformation is an efficient mean to produce giant liposomes &gt;10 &mu;m in diameter which relies on the application of alternating voltage to a thin, ordered lipid film deposited on an electrode surface. However, since the classical protocol calls for the lipids to be deposited from organic solvents, it is not compatible with less robust membrane proteins like ion channels and must be modified. Recently, protocols have been developed to electroform giant liposomes from partially dehydrated small liposomes, which we have adapted to protein-containing liposomes in our laboratory.
We present here the background, equipment, techniques, and pitfalls of electroformation of giant liposomes from small liposome dispersions. We begin with the classic protocol, which should be mastered first before attempting the more challenging protocols that follow. We demonstrate the process of controlled partial dehydration of small liposomes using vapor equilibrium with saturated salt solutions. Finally, we demonstrate the process of electroformation itself. We will describe simple, inexpensive equipment that can be made in-house to produce high-quality liposomes, and describe visual inspection of the preparation at each stage to ensure the best results.

Reconstituting functional membrane proteins into giant liposomes of defined composition is a powerful approach when combined with patch-clamp electrophysiology. However, conventional giant liposome production may be incompatible with protein stability. We describe protocols for producing giant liposomes from pure lipids or small liposomes containing ion channels.

הכנת Liposome ענקית הדמיה ותיקון מהדק Electrophysiology

The reconstitution of ion channels into chemically defined lipid membranes for electrophysiological recording has been a powerful technique to identify ...

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.

הקלטת תיקון מהדק מצורף תא אחד ערוצים

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

Reconstitution of a Transmembrane Protein, the Voltage-gated Ion Channel, KvAP, into Giant Unilamellar Vesicles for Microscopy and Patch Clamp Studies

Giant Unilamellar Vesicles (GUVs) are a popular biomimetic system for studying membrane associated phenomena. However, commonly used protocols to grow GUVs must be modified in order to form GUVs containing functional transmembrane proteins. This article describes two dehydration-rehydration methods &#8212; electroformation and gel-assisted swelling &#8212; to form GUVs containing the voltage-gated potassium channel, KvAP. In both methods, a solution of protein-containing small unilamellar vesicles is partially dehydrated to form a stack of membranes, which is then allowed to swell in a rehydration buffer. For the electroformation method, the film is deposited on platinum electrodes so that an AC field can be applied during film rehydration. In contrast, the gel-assisted swelling method uses an agarose gel substrate to enhance film rehydration. Both methods can produce GUVs in low (e.g., 5 mM) and physiological (e.g., 100 mM) salt concentrations. The resulting GUVs are characterized via fluorescence microscopy, and the function of reconstituted channels measured using the inside-out patch-clamp configuration. While swelling in the presence of an alternating electric field (electroformation) gives a high yield of defect-free GUVs, the gel-assisted swelling method produces a more homogeneous protein distribution and requires no special equipment.

The reconstitution of the transmembrane protein, KvAP, into giant unilamellar vesicles (GUVs) is demonstrated for two dehydration-rehydration methods &#8212; electroformation, and gel-assisted swelling. In both methods, small unilamellar vesicles containing the protein are fused together to form GUVs that can then be studied by fluorescence microscopy and patch-clamp electrophysiology.

כינונה מחדש של חלבון טראנסממברנלי, יון ערוץ מתח מגודרת, KvAP, לענק Unilamellar שלפוחית ​​למיקרוסקופי וצמד תיקון ללימודים

Giant Unilamellar Vesicles (GUVs) are a popular biomimetic system for studying membrane associated phenomena. However, commonly used protocols to grow ...

Implementing Patch Clamp and Live Fluorescence Microscopy to Monitor Functional Properties of Freshly Isolated PKD Epithelium

Cyst initiation and expansion during polycystic kidney disease is a complex process characterized by abnormalities in tubular cell proliferation, luminal fluid accumulation and extracellular matrix formation. Activity of ion channels and intracellular calcium signaling are key physiologic parameters which determine functions of tubular epithelium. We developed a method suitable for real-time observation of ion channels activity with patch-clamp technique and registration of intracellular Ca2+ level in epithelial monolayers freshly isolated from renal cysts. PCK rats, a genetic model of autosomal recessive polycystic kidney disease (ARPKD), were used here for ex vivo analysis of ion channels and calcium flux. Described here is a detailed step-by-step procedure designed to isolate cystic monolayers and non-dilated tubules from PCK or normal Sprague Dawley (SD) rats, and monitor single channel activity and intracellular Ca2+ dynamics. This method does not require enzymatic processing and allows analysis in a native setting of freshly isolated epithelial monolayer. Moreover, this technique is very sensitive to intracellular calcium changes and generates high resolution images for precise measurements. Finally, isolated cystic epithelium can be further used for staining with antibodies or dyes, preparation of primary cultures and purification for various biochemical assays.

Ion channels expressed in renal tubular epithelium play a significant role in the pathology of polycystic kidney disease. Here we describe experimental protocols used to perform patch-clamp analysis and intracellular calcium level measurements in cystic epithelium freshly isolated from rodent kidneys.

יישום הצמד תיקון והחי מיקרוסקופ פלואורסצנטי לפקח על מאפיינים פונקציונליים של מבודד טרי PKD האפיתל

Cyst initiation and expansion during polycystic kidney disease is a complex process characterized by abnormalities in tubular cell proliferation, luminal ...

Methods for the Isolation, Culture, and Functional Characterization of Sinoatrial Node Myocytes from Adult Mice

Sinoatrial node myocytes (SAMs) act as the natural pacemakers of the heart, initiating each heart beat by generating spontaneous action potentials (APs). These pacemaker APs reflect the coordinated activity of numerous membrane currents and intracellular calcium cycling. However the precise mechanisms that drive spontaneous pacemaker activity in SAMs remain elusive. Acutely isolated SAMs are an essential preparation for experiments to dissect the molecular basis of cardiac pacemaking. However, the indistinct anatomy, complex microdissection, and finicky enzymatic digestion conditions have prevented widespread use of acutely isolated SAMs. In addition, methods were not available until recently to permit longer-term culture of SAMs for protein expression studies. Here we provide a step-by-step protocol and video demonstration for the isolation of SAMs from adult mice. A method is also demonstrated for maintaining adult mouse SAMs in vitro and for expression of exogenous proteins via adenoviral infection. Acutely isolated and cultured SAMs prepared via these methods are suitable for a variety of electrophysiological and imaging studies.

Methods are demonstrated for the isolation of sinoatrial node myocytes (SAMs) from adult mice for patch clamp electrophysiology or imaging studies. Isolated cells can be used directly or can be maintained in culture to permit expression of proteins of interest, such as genetically encoded reporters.

שיטות הבידוד, תרבות האפיון פונקציונלי של myocytes צומת Sinoatrial של עכברים בוגרים

Sinoatrial node myocytes (SAMs) act as the natural pacemakers of the heart, initiating each heart beat by generating spontaneous action potentials (APs). ...