Wir danken folgenden Förderorganisationen und Stiftungen für die Unterstützung: National Institutes of Health, National Science Foundation, der American Heart Association, Gesellschaft für Muskeldystrophie, die Donald B. und Delia E. Baxter-Stiftung, die Klingenstein Fonds und der McKnight Endowment for Neuroscience.

Pulling Pipetten Mit einer Mikroelektrode puller wie Sutter P-97 Flaming / Brown, ziehen eine Reihe von etwa 10-20 Pipetten. <ol><li> Wählen Sie Ihre Kapillare aus Glas. Wir verwenden Borosilikat Kapillaren (Sutter BF150-110 bis 10 mm, 1,5 mm Außendurchmesser, 1,1 mm Innendurchmesser, 10 cm lang). Bewahren Sie das Glas vorsichtig damit es sauber und staubfrei bleibt. </li><li> Entwerfen Sie ein Ziehen Programm. Wir verwenden einen 5-Schritte-Programm, mit absteigender Hitze und Geschwindigkeit bei jedem Schritt, und ein kleiner Zug auf den letzten Schritt. Sutter ist Pipette Kochbuch ist eine hervorragende Referenz für die Entwicklung geeigneter Programme. </li><li> Untersuchen Sie die Pipettenspitzen unter dem Mikroskop auf Öffnungsdurchmesser und Glätte zu bestimmen. Discard rau, uneben oder unregelmäßig Tipps. Ein guter Zug-Protokoll sollten dafür sorgen, diese sind selten. Für Standard-Patch-Clamp-Aufzeichnung, sollte Bahnöffnungen 1-3 Mikrometer im Durchmesser sein. Ein Steilkegel (stumpfe Spitze) führt zu Widerstand für den gleichen Öffnungsdurchmesser geringer. Form und Größe können durch Druck Polieren modifiziert werden. </li></ol> Feuerpolieren Pipetten <ol><li> Richten Sie ein Polieren rig (microforge) mit einem Platin-Heizdraht durch ein Fußpedal gesteuert. Ein nützliches Kit ist ab ALA Scientific (CPM-2) oder eine ähnliche Vorrichtung kann stückweise zusammengesetzt werden. </li><li> Optional: Bestreichen Sie die Pipette mit einem Isolator, um die Kapazität zu verringern und verbessern das Geräuschverhalten. Wir verwenden Dentalwachs. Sylgard 184 ™ (Poly-dimethyl Siloxanelastomer auch als PDMS) ist auch eine Option. </li><li> Bei der Verwendung von Wachs, halten einen kleinen geschmolzenen Versorgung in der Nähe. Mit Luftdruck auf der Rückseite der Pipette auf Wachs von der Eingabe der Pipette zu halten, tauchen die Spitze kurz in das flüssige Wachs und zu entfernen. Mit Sylgard, speichern vorbereitet Elastomer in gefrorenen Aliquots und malen Sie die Pipettenspitze unter dem Mikroskop. Wärme an das Elastomer zu heilen. </li><li> Platzieren Sie die Pipette (beschichtet oder nicht) in der Poliervorrichtung und bringen die Spitze ~ 50 Mikron aus dem Filament. Denken Sie daran, dass der Faden dehnt sich bei Erwärmung. </li><li> Empfohlen: Folgen Sie der separaten Protokoll für &quot;pressure polishing&quot;, um die Form der Pipette für optimale Festigkeit und Spitzendurchmesser ändern. </li><li> Eine kurze Hitze-Impuls (1-2 Sekunden) ist ausreichend, um Wachs, das aus der Spitze der Pipette abnehmen und glatte Glas. </li><li> Legen Sie die fertigen Pipette in einem geschlossenen Kasten, um vor Staub zu schützen, wiederholen Sie für 10 erfolgreiche Pipetten. </li></ol>

Das Protokoll dargestellt ist hier im täglichen Einsatz in der Elektrophysiologie Labor und wird auch zur Injektionsnadeln für Zellen und Tieren zu machen. Mit einer programmierbaren Abzieher, ist es leicht, Pipetten für eine Vielzahl von Anwendungen zu machen. Mit Aufmerksamkeit und Pflege, wird das Filament auf Ihren Abzieher für ein Jahr oder länger dauern. Viel Glück mit Ihrer Experimente.

<!DOCTYPE html PUBLIC "-//W3C//DTD XHTML 1.0 Transitional//EN" "http://www.w3.org/TR/xhtml1/DTD/xhtml1-transitional.dtd">
<html xmlns="http://www.w3.org/1999/xhtml" xml:lang="en" lang="en">	
		
		<div>
		<table border="1">
		<thead>
		<tr>
		<th>Material Name</th>
		<th>Type</th>
		<th>Company</th>
		<th>Catalogue Number</th>
		<th>Comment</th>
		</tr>
		</thead>
		<tbody>
		
<tr>
<td>Micropipette Puller</td>
<td>Instrument</td>
<td>Sutter Instruments</td>
<td>P-97</td>
<td>Or similar instrument (e.g. Sutter P-87 or P-2000)</td>
</tr>
<tr>
<td>Glass Capillaries</td>
<td>Reagent</td>
<td>Sutter Instruments</td>
<td>BF150-86-10</td>
<td>Or, similar capillary glass. To make filling the pipette easier, use a capillary with a glass filament.</td>
</tr>		</tbody>
		</table>
		</div>

making patch-pipettes and sharp electrodes with a programmable puller

Glas-Mikroelektroden (auch als Pipetten) haben ein Arbeitspferd der Elektrophysiologie seit Jahrzehnten. Heute sind solche Pipetten von Glaskapillaren erfolgt über eine programmierbare Abzieher. Solche Instrumente Wärme der Kapillare entweder mit einem Metallfaden oder einem Laser und zeichnen sich aus dem Glas mit der Schwerkraft, einen Motor oder beides. Pipetten für Patch-Clamp-Aufzeichnung werden unter Verwendung nur Hitze und Schwerkraft, während scharfe Elektroden für die intrazelluläre Aufnahme verwenden eine Kombination von Wärme, Schwerkraft und einem Motor. Das Verfahren zur intrazellulären Aufnahme Pipetten machen, ist ähnlich, die zur Injektionsnadeln für eine Vielzahl von Anwendungen, einschließlich cRNA Injektion in Xenopus Oozyten zu machen. Im Allgemeinen, Kapillar-Glas &lt;1,2 mm im Durchmesser wird verwendet, um Pipetten für Patch-Clamp-Aufnahme zu machen, während engeren Glas für die intrazelluläre Aufnahme (Außendurchmesser = 1,0 mm) verwendet wird. Für jedes Werkzeug ist der Abzieher programmiert etwas anders. Dieses Video zeigt, wie man beide Arten von Aufnahmen Pipetten mit vorher festgelegten puller Programme machen.

Watch this Scientific Journal Video about Making Patch-pipettes and Sharp Electrodes with a Programmable Puller at JoVE.com

Making Patch-pipettes and Sharp Electrodes with a Programmable Puller

Dieses Video zeigt, wie eine programmierbare Abzieher verwenden, um Patch-Pipetten und scharfen Elektroden für die Elektrophysiologie zu machen. Das gleiche Verfahren kann verwendet werden, um eine Vielzahl von Glas-Tools, einschließlich Injektionsnadeln zu machen.

Machen Patch-Pipetten und Sharp Elektroden mit einem programmierbaren Puller

Glass microelectrodes (also called pipettes) have been a workhorse of electrophysiology for decades.  Today, such pipettes are made from glass capillaries ...

making-patch-pipettes-and-sharp-electrodes-with-a-programmable-puller

Universidad de Cartagena UDEC

Research

JoVE Journal

Biology

25.3K Aufrufe.  Stanford University. Dieses Video zeigt, wie eine programmierbare Abzieher verwenden, um Patch-Pipetten und scharfen Elektroden für die Elektrophysiologie zu machen. Das gleiche Verfahren kann verwendet werden, um eine Vielzahl von Glas-Tools, einschließlich Injektionsnadeln zu machen.

Serie: Making Patch-pipettes and Sharp Electrodes with a Programmable Puller

Patterning Cells on Optically Transparent Indium Tin Oxide Electrodes

The ability to exercise precise spatial and temporal control over cell-surface interactions is an important prerequisite to the assembly of multi-cellular constructs serving as in vitro mimics of native tissues.  In this study, photolithography and wet etching techniques were used to fabricate individually addressable indium tin oxide (ITO) electrodes on glass substrates.  The glass substrates containing ITO microelectrodes were modified with poly(ethylene glycol) (PEG) silane to make them protein and cell resistive.  Presence of insulating PEG molecules on the electrode surface was verified by cyclic voltammetry employing potassium ferricyanide as a redox reporter molecule.  Importantly, the application of reductive potential caused desorption of the PEG layer, resulting in regeneration of the conductive electrode surface and appearance of typical ferricyanide redox peaks.    Application of reductive potential also corresponded to switching of ITO electrode properties from cell non-adhesive to cell-adhesive. Electrochemical stripping of PEG-silane layer from ITO microelectrodes allowed for cell adhesion to take place in a spatially defined fashion, with cellular patterns corresponding closely to electrode patterns.  Micropatterning of several cell types was demonstrated on these substrates.   In the future, the control of the biointerfacial properties afforded by this method will allow to engineer cellular microenvironments through the assembly of three or more cell types into a precise geometric configuration on an optically transparent substrate.

Non-fouling PEG silane monolayer was desorbed from individually addressable ITO electrodes on glass by application of a reductive potential. Electrochemical stripping of PEG-silane layer from ITO microelectrodes allowed for cell adhesion to take place in a spatially defined fashion, with cellular patterns corresponding closely to electrode patterns.

Patterning Cells auf optisch transparenten Indium-Zinn-Oxid-Elektroden

The ability to exercise precise spatial and temporal control over cell-surface interactions is an important prerequisite to the assembly of multi-cellular ...

Forschung

Biologie

Fabrication of Amperometric Electrodes

Carbon fiber electrodes are crucial for the detection of catecholamine release from vesicles in single cells for amperometry measurements.  Here, we describe the techniques needed to generate low noise (&lt;0.5 pA) electrodes.  The techniques have been modified from published descriptions by previous researchers (1,2).  Electrodes are made by preparing carbon fibers and threading them individually into each capillary tube by using a vacuum with a filter to aspirate the fiber.  Next, the capillary tube with fiber is pulled by an electrode puller, creating two halves, each with a fine-pointed tip.  The electrodes are dipped in hot, liquid epoxy mixed with hardener to create an epoxy-glass seal.  Lastly, the electrodes are placed in an oven to cure the epoxy.  Careful handling of the electrodes is critical to ensure that they are made consistently and without damage.  This protocol shows how to fabricate and cut amperometric electrodes for recording from single cells.

This protocol describes how to generate carbon fiber electrodes. The electrodes are subsequently used to detect catecholamine release from vesicles with carbon fiber amperometry.

Herstellung von amperometrische Elektroden

Carbon fiber electrodes are crucial for the detection of catecholamine release from vesicles in single cells for amperometry measurements.  Here, we ...

Brain Banking: Making the Most of your Research Specimens

Unbiased stereology is a method for accurately and efficiently estimating the total neuron number (or other cell type) in a given area of interest1. To achieve this goal 6-10 systematic sections should be probed covering the entire structure. Typically this involves processing 1/5 sections which leaves a significant amount of material unprocessed. In order to maximize the material, we propose an inexpensive method for preserving fixed tissue as part of a long-term storage research plan. As tissue is sliced and processed for the desired stain or antibody, alternate sections should be systematically placed in antigen preserve at -20&deg;C for future processing. Using 24-well plates, sections can be placed in order for future retrieval. Using this method, tissue can be stored and processed for immunohistochemistry over the course of years.

Brain banking and systematic sampling of biological material provides the basis for unbiased stereology and maximizes the potential data obtained from each specimen.

Brain-Banking: Making the Most Ihrer Forschung Proben

Unbiased stereology is a method for accurately and efficiently estimating the total neuron number (or other cell type) in a given area of interest1.  To ...

Making MR Imaging Child's Play - Pediatric Neuroimaging Protocol, Guidelines and Procedure

Within the last decade there has been an increase in the use of structural and functional magnetic resonance imaging (fMRI) to investigate the neural basis of human perception, cognition and behavior 1, 2. Moreover, this non-invasive imaging method has grown into a tool for clinicians and researchers to explore typical and atypical brain development. Although advances in neuroimaging tools and techniques are apparent, (f)MRI in young pediatric populations remains relatively infrequent 2. Practical as well as technical challenges when imaging children present clinicians and research teams with a unique set of problems 3, 2. To name just a few, the child participants are challenged by a need for motivation, alertness and cooperation. Anxiety may be an additional factor to be addressed. Researchers or clinicians need to consider time constraints, movement restriction, scanner background noise and unfamiliarity with the MR scanner environment2,4-10. A progressive use of functional and structural neuroimaging in younger age groups, however, could further add to our understanding of brain development. As an example, several research groups are currently working towards early detection of developmental disorders, potentially even before children present associated behavioral characteristics e.g.11. Various strategies and techniques have been reported as a means to ensure comfort and cooperation of young children during neuroimaging sessions. Play therapy 12, behavioral approaches 13, 14,15, 16-18 and simulation 19, the use of mock scanner areas 20,21, basic relaxation 22 and a combination of these techniques 23 have all been shown to improve the participant's compliance and thus MRI data quality. Even more importantly, these strategies have proven to increase the comfort of families and children involved 12. One of the main advances of such techniques for the clinical practice is the possibility of avoiding sedation or general anesthesia (GA) as a way to manage children's compliance during MR imaging sessions 19,20. In the current video report, we present a pediatric neuroimaging protocol with guidelines and procedures that have proven to be successful to date in young children.

Despite an increase in the use of structural and functional magnetic resonance imaging (fMRI) in humans, the study of young pediatric populations remains a challenge. We present a hands-on, step-by-step video protocol including guidelines for clinicians and researchers intending to perform (f)MRI in young children.

Erstellen MR Imaging Kinderspiel - Pediatric Neuroimaging-Protokoll, Richtlinien und Verfahren

Within the last decade there has been an increase in the use of structural and functional magnetic resonance imaging (fMRI) to investigate the neural ...

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 ...

Visualizing Single-molecule DNA Replication with Fluorescence Microscopy

We describe a simple fluorescence microscopy-based real-time method for observing DNA replication at the single-molecule level. A circular, forked DNA template is attached to a functionalized glass coverslip and replicated extensively after introduction of replication proteins and nucleotides (Figure 1). The growing product double-strand DNA (dsDNA) is extended with laminar flow and visualized by using an intercalating dye. Measuring the position of the growing DNA end in real time allows precise determination of replication rate (Figure 2). Furthermore, the length of completed DNA products reports on the processivity of replication. This experiment can be performed very easily and rapidly and requires only a fluorescence microscope with a reasonably sensitive camera.


This protocol demonstrates a simple single-molecule fluorescence microscopy technique for visualizing DNA replication by individual replisomes in real time.

Visualizing Einzel-Molekül-DNA-Replikation mit Fluoreszenz-Mikroskopie

We describe a simple fluorescence microscopy-based real-time method for observing DNA replication at the single-molecule level. A circular, forked DNA ...

Lineage Labeling of Zebrafish Cells with Laser Uncagable Fluorescein Dextran

A central problem in developmental biology is to deduce the origin of the myriad cell types present in vertebrates as they arise from undifferentiated precursors. Researchers have employed various methods of lineage labeling, such as DiI labeling1 and pressure injection of traceable enzymes2 to ascertain cell fate at later stages of development in model systems. The first fate maps in zebrafish (Danio rerio) were assembled by iontophoretic injection of fluorescent dyes, such as rhodamine dextran, into single cells in discrete regions of the embryo and tracing the labeled cell's fate over time3-5. While effective, these methods are technically demanding and require specialized equipment not commonly found in zebrafish labs. Recently, photoconvertable fluorescent proteins, such as Eos and Kaede, which irreversibly switch from green to red fluorescence when exposed to ultraviolet light, are seeing increased use in zebrafish6-8. The optical clarity of the zebrafish embryo and the relative ease of transgenesis have made these particularily attractive tools for lineage labeling and to observe the migration of cells in vivo7. Despite their utility, these proteins have some disadvantages compared to dye-mediated lineage labeling methods. The most crucial is the difficulty we have found in obtaining high 3-D resolution during photoconversion of these proteins. In this light, perhaps the best combination of resolution and ease of use for lineage labeling in zebrafish makes use of caged fluorescein dextran, a fluorescent dye that is bound to a quenching group that masks its fluorescence9. The dye can then be "uncaged" (released from the quenching group) within a specific cell using UV light from a laser or mercury lamp, allowing visualization of its fluorescence or immunodetection. Unlike iontophoretic methods, caged fluorescein can be injected with standard injection apparatuses and uncaged with an epifluorescence microscope equipped with a pinhole10. In addition, antibodies against fluorescein detect only the uncaged form, and the epitope survives fixation well11. Finally, caged fluorescein can be activated with very high 3-D resolution, especially if two-photon microscopy is employed 12,13. This protocol describes a method of lineage labeling by caged fluorescein and laser uncaging. Subsequenctly, uncaged fluorescein is detected simultaneously with other epitopes such as GFP by labeling with antibodies.

This protocol delineates a way to label and trace the fate of small groups of cells zebrafish embryos using UV-uncaging of caged fluorescein, followed by whole mount immunolabeling to amplify the signal from the uncaged fluorescein.

Lineage Kennzeichnung von Zebrafisch-Zellen mit Laser Uncagable Fluorescein Dextran

A central problem in developmental biology is to deduce the origin of the myriad cell types present in vertebrates as they arise from undifferentiated ...

Measurement of Cellular Chemotaxis with ECIS/Taxis

Cellular movement in response to external stimuli is fundamental to many cellular processes including wound healing, inflammation and the response to infection. A common method to measure chemotaxis is the Boyden chamber assay, in which cells and chemoattractant are separated by a porous membrane. As cells migrate through the membrane toward the chemoattractant, they adhere to the underside of the membrane, or fall into the underlying media, and are subsequently stained and visually counted 1. In this method, cells are exposed to a steep and transient chemoattractant gradient, which is thought to be a poor representation of gradients found in tissues 2.
Another assay system, the under-agarose chemotaxis assay, 3, 4 measures cell movement across a solid substrate in a thin aqueous film that forms under the agarose layer. The gradient that develops in the agarose is shallow and is thought to be an appropriate representation of naturally occurring gradients. Chemotaxis can be evaluated by microscopic imaging of the distance traveled. Both the Boyden chamber assay and the under-agarose assay are usually configured as endpoint assays.
The automated ECIS/Taxis system combines the under-agarose approach with Electric Cell-substrate Impedance Sensing (ECIS) 5, 6. In this assay, target electrodes are located in each of 8 chambers. A large counter-electrode runs through each of the 8 chambers (Figure 2). Each chamber is filled with agarose and two small wells are the cut in the agarose on either side of the target electrode. One well is filled with the test cell population, while the other holds the sources of diffusing chemoattractant (Figure 3). Current passed through the system can be used to determine the change in resistance that occurs as cells pass over the target electrode. Cells on the target electrode increase the resistance of the system 6. In addition, rapid fluctuations in the resistance represent changes in the interactions of cells with the electrode surface and are indicative of ongoing cellular shape changes. The ECIS/Taxis system can measure movement of the cell population in real-time over extended periods of time, but is also sensitive enough to detect the arrival of a single cell at the target electrode. 
Dictyostelium discoidium is known to migrate in the presence of a folate gradient 7, 8 and its chemotactic response can be accurately measured by ECIS/Taxis 9. Leukocyte chemotaxis, in response to SDF1&alpha; and to chemotaxis antagonists has also been measured with ECIS/Taxis 10, 11. An example of the leukocyte response to SDF1&alpha; is shown in Figure 1.

The ECIS/Taxis system is an automated, real-time assay that measures cellular chemotaxis. In this assay, cells move beneath a layer of agarose to arrive at a target electrode. Cellular movement is measured by the onset of resistance to AC current 0.

Messung zellulärer Chemotaxis mit ECIS / Taxis

Cellular movement in response to external stimuli is fundamental to many cellular processes including wound healing, inflammation and the response to ...

Production of Xenopus tropicalis Egg Extracts to Identify Microtubule-associated RNAs

Many organisms localize mRNAs to specific subcellular destinations to spatially and temporally control gene expression. Recent studies have demonstrated that the majority of the transcriptome is localized to a nonrandom position in cells and embryos. One approach to identify localized mRNAs is to biochemically purify a cellular structure of interest and to identify all associated transcripts. Using recently developed high-throughput sequencing technologies it is now straightforward to identify all RNAs associated with a subcellular structure. To facilitate transcript identification it is necessary to work with an organism with a fully sequenced genome. One attractive system for the biochemical purification of subcellular structures are egg extracts produced from the frog Xenopus laevis. However, X. laevis currently does not have a fully sequenced genome, which hampers transcript identification. In this article we describe a method to produce egg extracts from a related frog, X. tropicalis, that has a fully sequenced genome. We provide details for microtubule polymerization, purification and transcript isolation. While this article describes a specific method for identification of microtubule-associated transcripts, we believe that it will be easily applied to other subcellular structures and will provide a powerful method for identification of localized RNAs.

Production of Xenopus tropicalis Egg Extracts to Identify Microtubule-associated RNAs

We describe the collection of unfertilized Xenopus tropicalis eggs and production of a meiosis II-arrested egg extract. This egg extract can be used to purify microtubules and microtubule-associated RNAs.

Produktion von<em&gt; Xenopus tropicalis</em&gt; Egg Extrakte Mikrotubuli-assoziierten RNAs identifizieren

Many  organisms localize mRNAs to specific subcellular destinations to spatially and  temporally control gene expression. Recent studies have demonstrated ...

Transfecting RAW264.7 Cells with a Luciferase Reporter Gene

Transfection of desired genetic materials into cells is an inevitable procedure in biomedical research studies. While numerous methods have been described, certain types of cells are resistant to many of these methods and yield low transfection efficiency1, potentially hindering research in those cell types. In this protocol, we present an optimized transfection procedure to introduce luciferase reporter genes as a plasmid DNA into the RAW264.7 macrophage cell line. Two different types of transfection reagents (lipid-based and polyamine-based) are described, and important notes are given throughout the protocol to ensure that the RAW264.7 cells are minimally altered by the transfection procedure and any experimental data obtained are the direct results of the experimental treatment. While transfection efficiency may not be higher compared to other transfection methods, the described procedure is robust enough for detecting luciferase signal in RAW264.7 without changing the physiological response of the cells to stimuli.

Transfection into the macrophage cell line, RAW264.7, is difficult due to the cell&#8217;s natural response against foreign materials. We described here a gentle yet robust procedure for transfecting luciferase reporter genes into RAW264.7 cells.

Machen Patch-Pipetten und Sharp Elektroden mit einem programmierbaren Puller

Zusammenfassung

Weitere Videos entdecken

Kapitel in diesem Video