We thank Mike W. Davidson (Florida State University) for providing plasmid DNA encoding fluorescent proteins. This work was supported by National Institutes of Health grant GM72048 (to D.W.P.).

1. Preparation of fluorescent protein droplet samples
A fluorescent protein droplet sample consists of a 1-octanol/water emulsion with the fluorescent protein residing in the water phase. This emulsion is sandwiched between a microscope slide and a 22 mm square cover glass for microscopy applications.
<ol>
<li>Before making fluorescent protein droplet samples the microscope slides and cover glasses need to be cleaned and coated with a hydrophobic agent.</li>
<li>Clean glassware by washing 5 minutes with acetone and leave to dry by air. (Optionally, after cleaning the glassware can be treated for 30 seconds in a plasma cleaner to obtain optimal coating results).</li>
<li>Prepare a 2% methyltrimethoxysilane solution in acetone and coat the glassware during a 2 minutes incubation in this solution. After coating remove the glassware from the solution and leave to dry by air. Then rinse with 70% ethanol from a spray bottle and leave to dry again. (Optionally, at this point the glassware can be baked for 1 hour at 80&deg;C to covalently link the coating to the glassware). Coated glassware can be stored for at least one month.</li>
<li> Fluorescent proteins are purified as His6-tagged protein from E. Coli 1. Measure the absorbance spectrum of the purified protein and prepare a stock dilution with an optical density of ~0.1 in STE buffer (150 mM NaCl, 10 mM Tris-HCl pH 8, 1 mM EDTA), containing 0.1% bovine serum albumin (BSA). In addition prepare 10 ml of a 1:1 mixture of 1-octanol and STE buffer in a 15 ml conical tube and mix vigorously. After mixing leave until the phase separation is complete. The top phase is the 1-octanol. (Caution: Because 1-octanol has a strong smell it is important to use a closed waste container for everything that comes in contact with 1-octanol.)</li>
<li> To make the emulsion pipette 45 &mu;l 1-octanol and 5 &mu;l fluorescent protein in an microfuge tube. Tap the tube a few times with your finger to start formation of the emulsion and then sonicate the tube for 30 seconds in a sonication bath. In the meanwhile get a coated microscope slide and cover glass ready. After sonication the emulsion should be completely cloudy. Immediately after sonication pipette 4 &mu;l emulsion from the middle of the tube onto a coated microscope slide and cover with a coated cover glass.</li>
<li>If the procedure is done correctly the emulsion should spread evenly between the microscope slide and the object glass. Within minutes the sample should be stable, consisting of ~10 &mu;m thick fluorescent droplets with varying diameter. The largest droplets are close to the center of the sample and the smaller are located further towards the edges.</li>
</ol>
 2. Setting up a photoconversion experiment
 The following procedure is a general strategy for setting up a fluorescent protein photoconversion experiment. This procedure can be applied for purified proteins as well as for live cells.
 <ol>
 <li>The following parameters provide a general starting point to set up your photoconversion experiment: 
 40x 1.3NA oil immersion objective 
 Image size = 512 x 512 pixels 
 Scan zoom = 4 
 
 Pixel dwell time = 6 &mu;sec. 
 Z-resolution (pinhole size) = 3 &mu;m</li>
 <li>Configure two detection channels for the initial and photoconverted fluorescence, as well as a "photoconversion channel". In this example we will use purified mOrange2 protein, which is a orange-to-red photoconvertible fluorescent protein. The orange species is detected using 561 nm excitation and the fluorescence is collected between 570 nm and 630 nm. The photoconverted red species is detected using 633 nm excitation and the fluorescence is collected between 640 nm and 700 nm. For the "photoconversion channel" select 488 nm excitation and collect fluorescence between 490 nm and 540 nm. (Note: imaging the photoconversion channel is not strictly necessary.)</li>
 <li> Use the channel for imaging the initial fluorescence with continuous scanning to adjust the laser power and detector gain for optimal image quality. </li>
 <li> Activate the photoconversion channel and select a low laser power. Start imaging a time lapse series and gradually increase the photoconversion laser until significant bleaching of the initial fluorescence is observed. Continue scanning until the initial fluorescence is approximately 75% bleached.</li>
 <li> Deactivate the photoconversion channel and activate the detection channel for the photoconverted fluorescence. Start imaging with a high detector gain and low laser power and gradually increase the laser power until the photoconverted fluorescence is detected. Once you detect the photoconverted fluorescence you can adjust laser power and detector gain for optimal image quality. </li>
 <li> Finally, the laser power used for photoconversion as well as the duration of photoconversion need to be optimized. Increasing the photoconversion laser power will accelerate the rate of photoconversion, however too much laser power will photobleach the protein.</li>
 <li> Once the optimal photoconversion laser power and duration have been determined, these parameters can be used to configure a standard photobleaching or FRAP module and the "photoconversion channel" is no longer required.</li>
 </ol>
 3. Dual-probe optical highlighting with mOrange2 and Dronpa
 Because of the red-shifted spectral properties, mOrange2 can be used in combination with the green photoswitchable fluorescent protein Dronpa for dual-probe optical highlighting to allow selective highlighting of 4 individual cell(organelle) populations.
 <ol>
 <li>Cells are grown in glass bottom MatTek dishes and transfected 24 hours prior to imaging using standard Lipofectamine2000 transfection1.</li>
 <li> Set up the microscope for mOrange2 photoconversion as described in section 2.</li>
 <li> Configure the microscope for Dronpa photoswitching. Dronpa fluorescence can be imaged using the mOrange2 "photoconversion channel" (see step 2.2). (Note: Minimize the laser power used for imaging Dronpa, because too much laser power will cause inactivation of Dronpa.) Add a channel for Dronpa photoactivation. We use 800 nm two-photon excitation for photoactivation, but alternatively this can be achieved using 405 nm excitation. Determine the laser power required for imaging, photoactivation, and photoinactivation of Dronpa fluorescence.</li>
 <li> Caution: Photoconversion of mOrange2 and inactivation of Dronpa both occur upon 488 nm excitation. Because of the high laser power required for mOrange2 photoconversion this will also inactivate Dronpa fluorescence. On the other hand, Dronpa inactivation occurs already at much lower laser power and can be performed without significant mOrange2 photoconversion.</li>
 <li> Once the parameters for mOrange2 photoconversion and Dronpa photoswitching are set, dual probe optical highlighting is achieved through the following steps. First, inactivate Dronpa fluorescence in the whole field of view with low power 488 nm excitation. Second, select a region of interest and photoconvert mOrange2 with high power 488 nm excitation. Finally, select a region of interest to activate Dronpa fluorescence.</li>
 </ol>
4. Representative Results
<img src="/files/ftp_upload/1995/1995fig1.jpg" alt="Figure 1" /> Figure 1. Droplet sample preparation. A) Correctly prepared droplet sample. B) Sample prepared without coating the microscope slide and cover glass. C) Sample prepared without adding 0.1% BSA.
<img src="/files/ftp_upload/1995/1995fig2.jpg" alt="Figure 2" /> Figure 2. Effect of photoconversion laser power and duration on mOrange2 photoconversion. Single droplets containing mOrange2 protein were continuously photoconverted using different amounts of 488 nm laser power. Relative laser power used for photoconversion was 10% (solid), 25% (dashed), and 100% (dotted). A) Orange fluorescent species. B) Photoconverted red fluorescent species.
<img src="/files/ftp_upload/1995/1995fig3.jpg" alt="Figure 3" /> Figure 3. Dual-probe optical highlighting with mOrange2 and Dronpa. A) Cell expressing mOrange2-Histone H2B and Dronpa-Mito before photoconversion, showing orange fluorescence in the nucleus and green fluorescence in the mitochondria. B) Dronpa fluorescence was switched off with low power 488 nm excitation, causing minimal photoconversion of mOrange2. C) mOrange2 was photoconverted to red with high power 488 nm excitation. D) Dronpa fluorescence was switched on again using 800 nm 2-photon excitation. The panels are overlays of the fluorescence images together with the differential interference contrast image.

<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>

No conflicts of interest declared.

The purified fluorescent protein droplet sample is a very convenient sample format for the photophysical characterization of fluorescent proteins, for example to study photobleaching kinetics and photoconversion kinetics. The extremely small droplet volume (~20 picoliter) facilitates photobleaching and photoconversion experiments, which can be difficult to perform in cuvette based systems. In addition, as shown here the droplet sample is ideally suited for setting up a confocal microscope for photoconversion applicatio...

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		<div>
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		<thead>
		<tr>
		<th>Material Name</th>
		<th>Type</th>
		<th>Company</th>
		<th>Catalogue Number</th>
		<th>Comment</th>
		</tr>
		</thead>
		<tbody>
		
<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 fluorescent proteins (pc-FPs) are a class of fluorescent proteins with "optical highlighter" capability, meaning that the color of fluorescence can be changed by exposure to light of a specific wavelength. Optical highlighting allows noninvasive marking of a subpopulation of fluorescent molecules, and is therefore ideal for tracking single cells or organelles.
Critical parameters for efficient photoconversion are the intensity and the exposure time of the photoconversion light. If the intensity is too low, photoconversion will be slow or not occur at all. On the other hand, too much intensity or too long exposure can photobleach the protein and thereby reduce the efficiency of photoconversion.
This protocol describes a general approach how to set up a confocal laser scanning microscope for pc-FP photoconversion applications. First, we describe a procedure for preparing purified protein droplet samples. This sample format is very convenient for studying the photophysical behavior of fluorescent proteins under the microscope. Second, we will use the protein droplet sample to show how to configure the microscope for photoconversion. And finally, we will show how to perform optical highlighting in live cells, including dual-probe optical highlighting with mOrange2 and Dronpa.

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

This protocol describes a general approach to perform photoconversion of fluorescent proteins on a confocal laser scanning microscope. We describe procedures for the photoconversion of puried protein samples, as well as for dual-probe optical highlighting in live cells with mOrange2 and Dronpa.

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

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12.1K Views.  Vanderbilt University. Этот протокол описывает общий подход для выполнения фотопреобразования флуоресцентных белков по конфокальной лазерной сканирующей микроскопии. Мы описываем процедуры фотопревращения puried образцов белков, а также для двойного зонда оптических подсветка в ж...

Video: 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.

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 ...

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.

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.

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.

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.

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.

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.

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.

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.

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.

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