보건 국립 연구소, 국립 과학 재단 (National Science Foundation), 미국 심장 협회, 근육질 영양 장애 협회, 도널드 B.와 델리아 E. 백스터 재단 Klingenstein 기금과 신경 과학에 대한 맥나이트 엔다우먼트 : 우리는 지원을위한 다음과 같은 자금 지원 기관 및 재단 감사드립니다.

풀링 pipettes 풀러 등 서터 P - 97 플라밍 / 브라운으로 microelectrode를 사용하여 약 10-20 pipettes 세트를 가져옵니다. <ol><li> 당신의 모세관 유리를 선택합니다. 우리는 borosilicate 모세관 (서터 BF150 - 110 - 10, 1.5mm 외경, 내경 1.1mm, 길이 10cm)을 사용합니다. 그것이 깨끗하고 먼지가 무료로 남아 있도록 신중하게 유리를 저장합니다. </li><li> 당기는 프로그램을 디자인합니다. 우리는 내림차순 열과 속도의 각 단계에서, 그리고 마지막 단계에있는 작은 당겨와 5 단계 프로그램을 사용합니다. 서터의 피펫 요리책은 적합한 프로그램을 개발하기위한 훌륭한 레퍼런스입니다. </li><li> 개방 직경과 매끄러운를 결정하는 현미경 피펫 팁을 검사합니다. 거친 고르지 또는 불규칙 조언을​​ 폐기하십시오. 좋은 당기는 프로토콜이가 드문 있도록해야합니다. 표준 패치 - 클램프 녹음, 팁 구멍은 직경 1-3 미크론해야합니다. 가파른 테이퍼 (뭉툭한 끝)은 같은 오프닝 직경에 대한 저항을 낮은 발생합니다. 모양과 크기가 압력 연마에 의해 수정할 수 있습니다. </li></ol> 소방 연마 pipettes <ol><li> 발 페달에 의해 통제 플래티넘 가열 필라멘트와 연마 장비 (microforge)를 설정합니다. 유용한 키트 에이러 과학 (CPM - 2) 또는 이와 유사한기구가 조금씩 조립하실 수 있습니다. </li><li> 옵션 : 코트 용량을 감소 및 소음 특성을 개선하기 위해 절연체로 피펫. 우리는 치과 왁스를 사용합니다. Sylgard 184 ™은 (또한 PDMS로 알려진 폴리 디메틸 실록산 - 엘라스토머)도 옵션입니다. </li><li> 왁스를 사용하는 경우, 작은 용융 공급이 근처에 보관. 피펫을 입력에서 왁스를 유지하기 위해 피펫의 뒷면에 공기 압력으로 액체 왁스에 팁 잠시을 찍어 제거합니다. Sylgard으로 저장, 준비 냉동 aliquots의 탄성과 현미경 피펫 팁을 페인트. 탄성체를 치료하기 위해 열. </li><li> 연마 장치에 피펫을 (코팅 여부) 장소 및 팁에게 필라멘트에서 ~ 50 미크론 가져. 가열하면 필라멘트가 확장됩니다 유의하십시오. </li><li> 추천 : 최적의 저항과 팁 직경에 대한 피펫의 모양을 변경하려면 &quot;압력 연마&quot;에 대한 별도의 프로토콜을 수행합니다. </li><li> 간단한 열 펄스 (1~2초)는 피펫의 팁을에서 왁스를 제거하고 유리를 부드럽게하기 위해 충분하다. </li><li> , 먼지로부터 보호하기 10 성공 pipettes을 반복하는 닫힌 상​​자에 완성된 피펫를 놓습니다. </li></ol>

프로토콜 그림 여기에 전기 생리학 실험실에서 일상 사용되고 또한 세포 및 동물 주입 바늘을하는 데 사용됩니다. 프로그램 풀러와 함께, 그것은 사용의 다양한 pipettes를 만들기 위해 쉽습니다. 관심과 관리를 통해 귀하의 풀러의 필라멘트는 1 년 이상 지속됩니다. 실험과 함께 행운을 빕니다.

<!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

유리 microelectrodes는 (또한 pipettes라고도 함) 수십 년 동안 전기 생리학의 일하는 말이되어 왔습니다. 오늘날과 같은 pipettes은 프로그램 풀러를 사용하여 유리 모세관에서 만들어집니다. 이러한 악기 열 금속 필라멘트 또는 레이저와 중력, 모터 또는 둘 모두를 사용하여 유리를 그릴 중 하나를 사용하여 모세관. 세포내 레코딩을위한 날카로운 전극은 열, 중력, 그리고 모터의 조합을 사용하는 동안 패치 - 클램프 녹음 Pipettes은 단 열과 중력을 이용하여 형성됩니다. 세포내 기록 pipettes를 만들기 위해 사용되는 절차는 Xenopus의 oocytes에 cRNA 분사를 포함한 다양한 어플 리케이션을위한 주사 바늘을 만들기 위해 사용하는 것과 비슷합니다. 좁은 유리 세포 기록 (외경 = 1.0 ㎜)에 사용되는 동안 일반적으로, 모세관 유리에 직경 &lt;1.2 mm은 패치 클램프 녹음 pipettes을하는 데 사용됩니다. 각 도구에 대한, 풀러가 약간 다르게 프로그램입니다. 이 비디오는 사전에 설립 풀러 프로그램을 사용 기록 pipettes 두 가지를하는 방법을 보여줍니다.

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

이 동영상은 전기 생리학에 대한 패치 pipettes과 날카로운 전극을 만들 수있는 프로그램 풀러를 사용하는 방법을 보여줍니다. 동일한 절차는 주사 바늘을 포함한 유리 다양한 도구를 만드는 데 사용할 수 있습니다.

프로그래밍 풀러와 패치 - pipettes와 샤프 전극을 만들기

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

Research

JoVE Journal

Biology

25.3K 조회수.  Stanford University. 이 동영상은 전기 생리학에 대한 패치 pipettes과 날카로운 전극을 만들 수있는 프로그램 풀러를 사용하는 방법을 보여줍니다. 동일한 절차는 주사 바늘을 포함한 유리 다양한 도구를 만드는 데 사용할 수 있습니다.

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

Application of Light-cured Dental Adhesive Resin for Mounting Electrodes or Microdialysis Probes in Chronic Experiments

In chronic recording experiments, self-curing dental acrylic resins have been used as a mounting base of electrodes or microdialysis-probes. Since these acrylics do not bond to the bone, screws have been used as anchors. However, in small experimental animals like finches or mouse, their craniums are very fragile and can not successfully hold the anchors. In this report, we propose a new application of light-curing dental resins for mounting base of electrodes or microdialysis probes in chronic experiments. This material allows direct bonding to the cranium. Therefore, anchor screws are not required and surgical field can be reduced considerably. Past experiences show that the bonding effect maintains more than 2 months. Conventional resin's window of time when the materials are pliable and workable is a few minutes. However, the window of working time for these dental adhesives is significantly wider and adjustable.

In this report, we propose a new application of light-curing dental resins for mounting base of electrodes or microdialysis probes in chronic experiments. This material allows direct bonding to the cranium.

만성 실험에 장착 전극 또는 Microdialysis 프로브를위한 라이트 - 치료 치과 접착제 수지의 응용

In chronic recording experiments, self-curing dental acrylic resins have been used as a mounting base of electrodes or microdialysis-probes. Since these ...

연구

생물학

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.

광학 투명 인듐 틴 산화물 전극에 Patterning 셀

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

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.

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.

브레인 은행 : 귀하의 연구 표본의 대부분을 만들기

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.

Making MR Imaging Child&#039;s Play - Pediatric Neuroimaging Protocol, Guidelines and Procedure

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.

소아 Neuroimaging 프로토콜, 지침 및 절차 - MR 이미징 아동의 놀이 만들기

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.

초기 압력에 의해 Gynogenetic의 Diploid의 Zebrafish 만들기

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.

형광 현미경과 함께 머릿속으로 단일 분자의 DNA 복제

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.

레이저 Uncagable 플루오레신 Dextran과 Zebrafish 세포의 리니지 라벨

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.

ECIS / 택시와 셀룰러 Chemotaxis 측정

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.

생산<em&gt; Xenopus의 tropicalis</em&gt; 달걀은 미세 소관 관련 RNA를 식별하기 위해 추출

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