我们感谢迈克W.戴维森（佛罗里达州立大学）提供荧光蛋白的质粒DNA编码。这项工作是由国家卫生授予GM72048（DWP）研究院的支持。

1。荧光蛋白滴样品制备一种荧光蛋白液滴样本包括与居住在水相中的荧光蛋白质1-octanol/water乳液。这个乳液是夹在显微镜幻灯片和一个22平方毫米的玻璃盖显微镜应用。 <ol><li>需要清理，并具有疏水性剂涂在荧光蛋白液滴样本的显微镜载玻片和盖玻片。 </li><li> 5分钟，用丙酮洗涤清洁的玻璃器皿和离开干燥空气。 （可选，清洗玻璃器皿后，可以被视为在等离子体清洁30秒，以获得最佳的涂层结果）。 </li><li> 2分钟，在这个解决方案孵化期间，准备在2％的丙酮和大衣的玻璃器皿methyltrimethoxysilane解决方案。涂后，从溶液中取出玻璃器皿，离开干燥空气。然后用70％乙醇喷雾瓶，再次留下来干。 （或者，在这一点上的玻璃器皿可烤1小时在80 ° C共价键链接涂层玻璃器皿）。镀膜玻璃制品可以保存至少一个月。 </li><li>他的6标记的大肠杆菌的蛋白质纯化荧光蛋白大肠杆菌 1。测量纯化的蛋白质的吸收光谱，并准备在STE缓冲液（150 mM氯化钠，10毫米的Tris - HCl pH值8，1毫米EDTA）〜0.1光密度的股票稀释，含0.1％牛血清白蛋白（BSA） 。另外准备10毫升1:1混合的1 - 辛醇和STE缓冲液15 ml锥形管，并大力混合。相分离混合后离开，直到完成。最高阶段是1 - 辛醇。 （注意：由于1 - 辛醇，具有很强的气味是很重要的一个封闭的废弃物容器，1 - 辛醇接触的一切。） </li><li>为了使1 - 辛醇和5μL离心管中的荧光蛋白的乳化吸液管45μL。用手指轻按管几次，开始形成乳液，然后管超声在超声浴30秒。在同时得到涂层的显微镜载玻片和盖玻片的准备。超声乳化液后应完全阴天。超声吸管从管中的4μL乳液到涂显微镜幻灯片后立即覆盖涂层的玻璃盖。 </li><li>如果过程是正确的乳液应均匀地分布之间的显微镜幻灯片和对象的玻璃。样品在几分钟之内应该是稳定的，不同直径〜10微米厚的荧光液滴组成。最大的水滴接近试样的中心，位于较小对边缘进一步。 </li></ol> 2。建立一个光转化实验下面的过程是一个总体战略，设立一种荧光蛋白的光转化实验。此过程可对活细胞中的蛋白质纯化以及应用。 <ol><li>以下参数设置你的光转化实验提供了一个一般的起点： 40X 1.3NA油浸物镜 图像尺寸为512 × 512像素 扫描变焦= 4 像素停留时间= 6微秒。 Z轴分辨率（针孔大小）= 3微米</li><li>配置初始photoconverted荧光检测通道，以及“光转化通道”。在这个例子中，我们将使用，纯化的mOrange2蛋白质，这是一个橙红色photoconvertible的荧光蛋白。橙色的物种是检测用561 nm激发和荧光收集的570纳米和630纳米之间。 photoconverted红色物种检测用633 nm激发和荧光收集的640纳​​米和700纳米之间。对于“光转化通道”，选择488 nm激发，并收集490 nm和540 nm处的荧光。 （注：不是绝对必要的成像光转化渠道。） </li><li>使用连续扫描调整激光功率和探测器获得最佳的图像质量与初始荧光成像通道。 </li><li>激活光转化渠道，并选择一个较低的激光功率。开始成像时间的推移系列，并逐步提高激光的光转化，直到观察到了显着的初始荧光漂白。继续扫描，直到初始荧光漂白约75％。 </li><li>停用的光转化渠道和激活的photoconverted荧光检测通道。开始与高探测器的增益和低激光功率成像，并逐步提高激光功率，直到photoconverted荧光检测。一旦你发现photoconverted的荧光，您可以调整激光功率和探测器获得最佳的图像质量。 </li><li>最后，激光功率使用以及光转化的光转化的时间需要优化。提高光转化激光功率将加速光转化率，但是过多的激光功率，光漂白的蛋白质。 </li><li>一旦最佳的光转化的激光功率和持续时间已经确定，这些参数可用于配置一个标准的漂白或FRAP模块和不再需要的“光转化通道”。 </li></ol> 3。 mOrange2和Dronpa双探头光学突出由于红移光谱特性，mOrange2可以用来photoswitchable双探头光学突出绿色荧光蛋白Dronpa，允许4个独立的细胞（细胞器）人口的选择性突出的组合。 <ol><li>细胞是生长在玻璃底部MatTek菜和转染24小时前成像标准Lipofectamine2000转染 1 。 </li><li>设置第2条所述mOrange2光转化的显微镜。 </li><li>配置Dronpa photoswitching显微镜。 Dronpa荧光成像使用mOrange2“光转化通道”（见第2.2步）。 （注：用于成像Dronpa的激光功率最小化，因为太多的激光功率会导致Dronpa灭活）。Dronpa photoactivati​​on添加频道。我们用800 nm的双光子激发photoactivati​​on，但另外可以做到这一点使用405 nm激发。确定的成像所需的激光功率，photoactivati​​on，Dronpa荧光photoinactivati​​on。 </li><li>注意：光转化的mOrange2 Dronpa灭活发生后，488 nm激发。由于mOrange2光转化所需的激光功率高，这也将灭活Dronpa荧光。另一方面，Dronpa灭活发生已经在激光功率要低得多，可以不执行重大mOrange2光转化。 </li><li>一旦mOrange2光转化和Dronpa photoswitching的参数设置，双探头光学突出的是通过以下步骤。首先，在整场与低功耗488 nm激发荧光Dronpa灭活。其次，选择一个地区的利益，并具有高功率488 nm激发photoconvert mOrange2。最后，选择一个感兴趣的激活Dronpa荧光的区域。 </li></ol> 4。代表性的成果<img src="/files/ftp_upload/1995/1995fig1.jpg" alt="图1" /> 图1。液滴的样品制备。一个正确的准备）液滴样本。 b）样品准备，无涂层的显微镜载玻片和盖玻片。 c）样品准备而增加0.1％BSA。 <img src="/files/ftp_upload/1995/1995fig2.jpg" alt="图2" /> 图2。光转化mOrange2光转化的激光功率和持续时间的影响，含有mOrange2蛋白质的单液滴不断photoconverted使用不同金额的488 nm激光功率。相对激光功率为光转化为为10％（固体），25％（虚线），和100％（虚线）。一）橙色荧光物种。二）Photoconverted红色荧光物种。 <img src="/files/ftp_upload/1995/1995fig3.jpg" alt="图3" /> 图3。双探头光学突出mOrange2和Dronpa）前光转化细胞表达mOrange2组蛋白H2B和Dronpa三刀，在线粒体在细胞核和绿色荧光的橙色荧光。二）Dronpa荧光切断与低功耗488 nm激发，造成的mOrange2最小光转化。三）mOrange2 photoconverted以红色高功率488 nm激发。四）Dronpa接通再次使用800 nm的双光子激发荧光。该小组的荧光图像叠加在一起的微分干涉对比图像。

<ol>
	<li>Kremers, G. J., Hazelwood, K. L., Murphy, C. S., Davidson, M. W., Piston, D. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Photoconversion+in+orange+and+red+fluorescent+proteins.">Photoconversion in orange and red fluorescent proteins.</a> Nature Methods. 6, 355-358 (2009).</li></ol>

纯化的荧光蛋白液滴样本的荧光蛋白的光物理特性是非常方便的样本格式，例如漂白动力学和光转化动力学研究。液滴体积非常小（〜20皮升）促进漂白和光转化实验，它可以比色皿的系统难以执行。此外，如图所示，这里液滴样本非常适合设立了光转化应用共聚焦显微镜。疏水性涂层和BSA的存在是重要的，获得均匀的液滴。无涂层的液滴往往是对玻璃表面的挤压和缺乏对BSA的荧光蛋白容易聚积?...

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		<th>Material Name</th>
		<th>Type</th>
		<th>Company</th>
		<th>Catalogue Number</th>
		<th>Comment</th>
		</tr>
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<tr>
<td>Microsope slides</td>
<td>&nbsp;</td>
<td>VWR Scientific</td>
<td>48312-003</td>
<td>&nbsp;</td>
<tr>
<td>22 mm cover glass</td>
<td>&nbsp;</td>
<td>Corning</td>
<td>2940-245</td>
<td>&nbsp;</td>
<tr>
<td>1-octanol</td>
<td>&nbsp;</td>
<td>Sigma</td>
<td>O4500</td>
<td>&nbsp;</td>
<tr>
<td>methyltrimethoxysilane</td>
<td>&nbsp;</td>
<td>Sigma</td>
<td>M6420</td>
<td>&nbsp;</td>
<tr>
<td>MatTek dishes</td>
<td>&nbsp;</td>
<td>MatTek Corp.</td>
<td>P35G-1.5-14-C</td>
<td>&nbsp;</td>
<tr>
<td>Lipofectamine2000</td>
<td>&nbsp;</td>
<td>Invitrogen</td>
<td>11668-019</td>
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photoconversion of purified fluorescent proteins and dual-probe optical highlighting in live cells

Photoconvertible荧光蛋白（PC - FPS）是一种荧光蛋白类的“光学荧光笔”的能力，这意味着暴露于一个特定波长的光可以通过改变颜色的荧光。光突出允许非侵入性标记的荧光分子亚群，因此跟踪单个细胞或细胞器的理想选择。 高效光转化的关键参数的光转化光的强度和曝光时间。如果强度太低，将光转化缓慢或不发生在所有。另一方面，强度太大或太长时间曝光可以光漂白的蛋白质，从而减少了光转化效率。 本协议描述了如何设置PC - FP光转化应用激光共聚焦扫描显微镜的一般方法。首先，我们描述准备纯化蛋白液滴样本的一个过程。此样本格式是非常方便的在显微镜下研究荧光蛋白的光物理行为。其次，我们将使用蛋白质液滴示例显示如何配置的光转化显微镜。最后，我们将展示如何执行在活细胞，包括mOrange2和Dronpa双探头光学突出中的光学突出。

Watch this Scientific Journal Video about Photoconversion of Purified Fluorescent Proteins and Dual-probe Optical Highlighting in Live Cells at JoVE.com

Photoconversion of Purified Fluorescent Proteins and Dual-probe Optical Highlighting in Live Cells

这个协议描述的一般方法执行在激光共聚焦显微镜，荧光蛋白的光转化。我们描述puried蛋白质样品的光转化，以及与mOrange2和Dronpa活细胞的双探头光学突出的程序。

在活细胞荧光蛋白的双探头光学突出纯净的光转化的

Photoconvertible fluorescent proteins (pc-FPs) are a class of fluorescent proteins with "optical highlighter" capability, meaning that the color of ...

photoconversion-purified-fluorescent-proteins-dual-probe-optical

Research

JoVE Journal

Biology

12.1K 次观看.  Vanderbilt University. 这个协议描述的一般方法执行在激光共聚焦显微镜，荧光蛋白的光转化。我们描述puried蛋白质样品的光转化，以及与mOrange2和Dronpa活细胞的双探头光学突出的程序。

视频: Photoconversion of Purified Fluorescent Proteins and Dual-probe Optical Highlighting in Live Cells

Labeling Stem Cells with Fluorescent Dyes for non-invasive Detection with Optical Imaging

Optical imaging (OI) is an easy, fast and inexpensive tool for in vivo monitoring of new stem cell based therapies. The technique is based on ex vivo labeling of stem cells with a fluorescent dye, subsequent intravenous injection of the labeled cells and visualization of their accumulation in specific target organs or pathologies. The presented technique demonstrates how we label human mesenchymal stem cells (hMSC) by simple incubation with the lipophilic fluorescent dye DiD (C67H103CIN2O3S) and how we label human embryonic stem cells (hESC) with the FDA approved fluorescent dye Indocyanine Green (ICG). The uptake mechanism is via adherence and diffusion of the lypophilic dye across the phospholipid cell membrane bilayer. The labeling efficiency is usually improved if the cells are incubated with the dye in serum-free media as opposed to incubation in serum-containing media. Furthermore, the addition of the transfection agent Protamine Sulfate significantly improves contrast agent uptake.

This video shows techniques for labeling of human embryonic stem cells and mesenchymal stem cells with fluorescent dyes. This technique can be used for an in vivo tracking of transplanted stem cells with optical imaging and for histopathological correlations with fluorescence microscopy.

与非侵入性检测荧光染料标记的干细胞与光学成像

Optical imaging (OI) is an easy, fast and inexpensive tool for in vivo monitoring of new stem cell based therapies. The technique is based on ex vivo ...

科研

生物学

Proper Care and Cleaning of the Microscope

Keeping the microscope optics clean is important for high-quality imaging. Dust, fingerprints, excess immersion oil, or mounting medium on or in a microscope causes reduction in contrast and resolution. DIC is especially sensitive to contamination and scratches on the lens surfaces. This protocol details the procedure for keeping the microscope clean.

Keeping the microscope optics clean is important for high-quality imaging. Dust, fingerprints, excess immersion oil, or mounting medium on or in a microscope causes reduction in contrast and resolution. DIC is especially sensitive to contamination and scratches on the lens surfaces. This protocol details the procedure for keeping the microscope clean.

适当的照顾和清洁的显微镜

Keeping the microscope optics clean is important for high-quality imaging. Dust, fingerprints, excess immersion oil, or mounting medium on or in a ...

Fluorescent Labeling of COS-7 Expressing SNAP-tag Fusion Proteins for Live Cell Imaging

SNAP-tag and CLIP-tag protein labeling systems enable the specific, covalent attachment of molecules, including fluorescent dyes, to a protein of interest in live cells. These systems offer a broad selection of fluorescent substrates optimized for a range of imaging instrumentation. Once cloned and expressed, the tagged protein can be used with a variety of substrates for numerous downstream applications without having to clone again. There are two steps to using this system: cloning and expression of the protein of interest as a SNAP-tag fusion, and labeling of the fusion with the SNAP-tag substrate of choice. The SNAP-tag is a small protein based on human O6-alkylguanine-DNA-alkyltransferase (hAGT), a DNA repair protein. SNAP-tag labels are dyes conjugated to guanine or chloropyrimidine leaving groups via a benzyl linker. In the labeling reaction, the substituted benzyl group of the substrate is covalently attached to the SNAP-tag. CLIP-tag is a modified version of SNAP-tag, engineered to react with benzylcytosine rather than benzylguanine derivatives. When used in conjunction with SNAP-tag, CLIP-tag enables the orthogonal and complementary labeling of two proteins simultaneously in the same cells.

SNAP-tag and CLIP-tag protein labeling systems enable the specific, covalent attachment of molecules, including fluorescent dyes, to a protein of interest in live cells. Once cloned and expressed, the tagged protein can be used with a variety of substrates for numerous downstream applications without having to clone again.

对活细胞成像的SNAP - Tag融合蛋白的荧光标记的COS - 7

SNAP-tag and CLIP-tag protein labeling systems enable the specific, covalent attachment of molecules, including fluorescent dyes, to a protein of interest ...

Polarized Translocation of Fluorescent Proteins in Xenopus Ectoderm in Response to Wnt Signaling

Cell polarity is a fundamental property of eukaryotic cells that is dynamically regulated by both intrinsic and extrinsic factors during embryonic development 1, 2. One of the signaling pathways involved in this regulation is the Wnt pathway, which is used many times during embryogenesis and critical for human disease3, 4, 5. Multiple molecular components of this pathway coordinately regulate signaling in a spatially-restricted manner, but the underlying mechanisms are not fully understood. Xenopus embryonic epithelial cells is an excellent system to study subcellular localization of various signaling proteins. Fluorescent fusion proteins are expressed in Xenopus embryos by RNA microinjection, ectodermal explants are prepared and protein localization is evaluated by epifluorescence. In this experimental protocol we describe how subcellular localization of Diversin, a cytoplasmic protein that has been implicated in signaling and cell polarity determination6, 7 is visualized in Xenopus ectodermal cells to study Wnt signal transduction8. Coexpression of a Wnt ligand or a Frizzled receptor alters the distribution of Diversin fused with red fluorescent protein, RFP, and recruits it to the cell membrane in a polarized fashion 8, 9. This ex vivo protocol should be a useful addition to in vitro studies of cultured mammalian cells, in which spatial control of signaling differs from that of the intact tissue and is much more difficult to analyze.

Polarized Translocation of Fluorescent Proteins in Xenopus Ectoderm in Response to Wnt Signaling

Xenopus embryonic ectoderm has become an attractive model for studies of cell polarity. An assay is described, in which subcellular distribution of fluorescent proteins is assessed in ectoderm cells. This protocol will help address questions related to spatial control of signaling.

偏光易位荧光蛋白爪蟾外胚层响应Wnt信号

Cell polarity is a fundamental property of eukaryotic cells that is dynamically regulated by both intrinsic and extrinsic factors during embryonic ...

Live Imaging of GFP-labeled Proteins in Drosophila Oocytes

The Drosophila oocyte has been established as a versatile system for investigating fundamental questions such as cytoskeletal function, cell organization, and organelle structure and function. The availability of various GFP-tagged proteins means that many cellular processes can be monitored in living cells over the course of minutes or hours, and using this technique, processes such as RNP transport, epithelial morphogenesis, and tissue remodeling have been described in great detail in Drosophila oocytes1,2.
The ability to perform video imaging combined with a rich repertoire of mutants allows an enormous variety of genes and processes to be examined in incredible detail. One such example is the process of ooplasmic streaming, which initiates at mid-oogenesis3,4. This vigorous movement of cytoplasmic vesicles is microtubule and kinesin-dependent5 and provides a useful system for investigating cytoskeleton function at these stages.
Here I present a protocol for time lapse imaging of living oocytes using virtually any confocal microscopy setup.

Live Imaging of GFP-labeled Proteins in Drosophila Oocytes

A protocol for live imaging of GFP-tagged proteins or autofluorescent structures in individual Drosophila oocytes is described.

GFP标记的蛋白质在实时成像<em&gt;果蝇</em&gt;卵母细胞

The  Drosophila oocyte has been established as a versatile system for investigating  fundamental questions such as cytoskeletal function, cell ...

Intravital Microscopy for Imaging Subcellular Structures in Live Mice Expressing Fluorescent Proteins

Here we describe a procedure to image subcellular structures in live rodents that is based on the use of confocal intravital microscopy. As a model organ, we use the salivary glands of live mice since they provide several advantages. First, they can be easily exposed to enable access to the optics, and stabilized to facilitate the reduction of the motion artifacts due to heartbeat and respiration. This significantly facilitates imaging and tracking small subcellular structures. Second, most of the cell populations of the salivary glands are accessible from the surface of the organ. This permits the use of confocal microscopy that has a higher spatial resolution than other techniques that have been used for in vivo imaging, such as two-photon microscopy. Finally, salivary glands can be easily manipulated pharmacologically and genetically, thus providing a robust system to investigate biological processes at a molecular level.
In this study we focus on a protocol designed to follow the kinetics of the exocytosis of secretory granules in acinar cells and the dynamics of the apical plasma membrane where the secretory granules fuse upon stimulation of the beta-adrenergic receptors. Specifically, we used a transgenic mouse that co-expresses cytosolic GFP and a membrane-targeted peptide fused with the fluorescent protein tandem-Tomato. However, the procedures that we used to stabilize and image the salivary glands can be extended to other mouse models and coupled to other approaches to label in vivo cellular components, enabling the visualization of various subcellular structures, such as endosomes, lysosomes, mitochondria, and the actin cytoskeleton.

Intravital microscopy is a powerful tool that enables imaging various biological processes in live animals. In this article, we present a detailed method for imaging the dynamics of subcellular structures, such as the secretory granules, in the salivary glands of live mice.

活体显微镜成像的亚细胞结构的活体小鼠表达荧光蛋白

Here we describe a procedure to image subcellular structures in live rodents that is based on the use of confocal intravital microscopy. As a model organ, ...

In vivo Clonal Tracking of Hematopoietic Stem and Progenitor Cells Marked by Five Fluorescent Proteins using Confocal and Multiphoton Microscopy

We developed and validated a fluorescent marking methodology for clonal tracking of hematopoietic stem and progenitor cells (HSPCs) with high spatial and temporal resolution to study in vivo hematopoiesis using the murine bone marrow transplant experimental model. Genetic combinatorial marking using lentiviral vectors encoding fluorescent proteins (FPs) enabled cell fate mapping through advanced microscopy imaging. Vectors encoding five different FPs: Cerulean, EGFP, Venus, tdTomato, and mCherry were used to concurrently transduce HSPCs, creating a diverse palette of color marked cells. Imaging using confocal/two-photon hybrid microscopy enables simultaneous high resolution assessment of uniquely marked cells and their progeny in conjunction with structural components of the tissues. Volumetric analyses over large areas reveal that spectrally coded HSPC-derived cells can be detected non-invasively in various intact tissues, including the bone marrow (BM), for extensive periods of time following transplantation. Live studies combining video-rate multiphoton and confocal time-lapse imaging in 4D demonstrate the possibility of dynamic cellular and clonal tracking in a quantitative manner.

In vivo Clonal Tracking of Hematopoietic Stem and Progenitor Cells Marked by Five Fluorescent Proteins using Confocal and Multiphoton Microscopy

Combinatorial 5 fluorescent proteins marking of hematopoietic stem and progenitor cells allows in vivo clonal tracking via confocal and two-photon microscopy, providing insights into bone marrow hematopoietic architecture during regeneration. This method allows non-invasive fate mapping of spectrally-coded HSPCs-derived cells in intact tissues for extensive periods of time following transplantation.

在造血干细胞和祖细胞利用共聚焦和多光子显微镜五荧光蛋白标记的活体克隆跟踪

We developed and validated a fluorescent marking methodology for clonal tracking of hematopoietic stem and progenitor cells (HSPCs) with high spatial and ...

Green Fluorescent Protein-based Expression Screening of Membrane Proteins in Escherichia coli

The production of recombinant membrane proteins for structural and functional studies remains technically challenging due to low levels of expression and the inherent instability of many membrane proteins once solubilized in detergents. A protocol is described that combines ligation independent cloning of membrane proteins as GFP fusions with expression in Escherichia coli detected by GFP fluorescence. This enables the construction and expression screening of multiple membrane protein/variants to identify candidates suitable for further investment of time and effort. The GFP reporter is used in a primary screen of expression by visualizing GFP fluorescence following SDS polyacrylamide gel electrophoresis (SDS-PAGE). Membrane proteins that show both a high expression level with minimum degradation as indicated by the absence of free GFP, are selected for a secondary screen. These constructs are scaled and a total membrane fraction prepared and solubilized in four different detergents. Following ultracentrifugation to remove detergent-insoluble material, lysates are analyzed by fluorescence detection size exclusion chromatography (FSEC). Monitoring the size exclusion profile by GFP fluorescence provides information about the mono-dispersity and integrity of the membrane proteins in different detergents. Protein: detergent combinations that elute with a symmetrical peak with little or no free GFP and minimum aggregation are candidates for subsequent purification. Using the above methodology, the heterologous expression in E. coli of SED (shape, elongation, division, and sporulation) proteins from 47 different species of bacteria was analyzed. These proteins typically have ten transmembrane domains and are essential for cell division. The results show that the production of the SEDs orthologues in E. coli was highly variable with respect to the expression levels and integrity of the GFP fusion proteins. The experiment identified a subset for further investigation.

Green Fluorescent Protein-based Expression Screening of Membrane Proteins in Escherichia coli

A streamlined approach to screening for the expression of recombinant membrane proteins in Escherichia coli based on fusion to green fluorescent protein is presented.

膜蛋白的绿色荧光蛋白为基础的表达筛选<em&gt;大肠杆菌</em

The production of recombinant membrane proteins for structural and functional studies remains technically challenging due to low levels of expression and ...

Functional Reconstitution and Channel Activity Measurements of Purified Wildtype and Mutant CFTR Protein

The Cystic Fibrosis Transmembrane Conductance Regulator (CFTR) is a unique channel-forming member of the ATP Binding Cassette (ABC) superfamily of transporters. The phosphorylation and nucleotide dependent chloride channel activity of CFTR has been frequently studied in whole cell systems and as single channels in excised membrane patches. Many Cystic Fibrosis-causing mutations have been shown to alter this activity. While a small number of purification protocols have been published, a fast reconstitution method that retains channel activity and a suitable method for studying population channel activity in a purified system have been lacking. Here rapid methods are described for purification and functional reconstitution of the full-length CFTR protein into proteoliposomes of defined lipid composition that retains activity as a regulated halide channel. This reconstitution method together with a novel flux-based assay of channel activity is a suitable system for studying the population channel properties of wild type CFTR and the disease-causing mutants F508del- and G551D-CFTR. Specifically, the method has utility in studying the direct effects of phosphorylation, nucleotides and small molecules such as potentiators and inhibitors on CFTR channel activity. The methods are also amenable to the study of other membrane channels/transporters for anionic substrates.

Described here is a rapid and effective procedure for functional reconstitution of purified wild-type and mutant CFTR protein that preserves activity for this chloride channel, which is defective in Cystic Fibrosis. Iodide efflux from reconstituted proteoliposomes mediated by CFTR allows studies of channel activity and the effects of small molecules.

纯化野生型和突变CFTR蛋白的功能重建和通道活性测定

The Cystic Fibrosis Transmembrane Conductance Regulator (CFTR) is a unique channel-forming member of the ATP Binding Cassette (ABC) superfamily of ...

Characterization of Calcification Events Using Live Optical and Electron Microscopy Techniques in a Marine Tubeworm

Characterizing the first event of biological production of calcium carbonate requires a combination of microscopy approaches. First, intracellular pH distribution and calcium ions can be observed using live microscopy over time. This allows identification of the life stage and the tissue with the feature of interest for further electron microscopy studies. Life stage and tissues of interest are typically higher in pH and Ca signals. 
Here, using H. elegans, we present a protocol to characterize the presence of calcium carbonate structures in a biological specimen on the scanning electron microscope (SEM), using energy-dispersive X-ray spectroscopy (EDS) to visualize elemental composition, using electron backscatter diffraction (EBSD) to determine the presence of crystalline structures, and using transmission electron microscopy (TEM) to analyze the composition and structure of the material. In this protocol, a focused ion beam (FIB) is used to isolate samples with dimension suitable for TEM analysis. As FIB is a site specific technique, we demonstrate how information from the previous techniques can be used to identify the region of interest, where Ca signals are highest.

We demonstrate the use of various microscopy methods that are useful in observing the calcification of a tubeworm, Hydroides elegans, as well as locating and characterizing the first calcified material. Live microscopy and electron microscopy are used together to provide functional and material information that are important in studying biomineralization.

钙化事件的表征在海洋Tubeworm使用实时光学和电子显微镜技术

Characterizing the first event of biological production of calcium carbonate requires a combination of microscopy approaches. First, intracellular pH ...