Kai Johnsson 
Ivan Correa 
Andreas Brecht 
Katrin M&uuml;entener 
Chris Noren 
Luo Sun 
Ming Xu 
Theodore Davis 
Salvatore Russello 
Aihua Zhang 
Inca Ghosh 
Elissa Maunus 
Nicholas Labarthe 
James MacFarland 
John Buswell 
Brenda Desmond 
James Ellard 
Don Comb

Plating CHO-K1 Cells:
<ol>
 <li>Add 50-100&mu;l of CHO-K1 cells that have been trypsinized according to standard protocols to 6ml of complete F-12K.</li>
 <li>&nbsp;Mix cell suspension by pipetting up and down several times, then aliquot 300 &mu;l of trypsinized cell suspension to each well of a sterile 8-chamber Lab-Tek II Chambered Coverglass with #1.5 borosilicate coverglass chambers (Nalgene Nunc Int.; Product #155409; Lot: 100808-8-0).</li>
 <li>Incubate samples at 37&deg;C, 5% CO2 overnight.</li>
</ol>
Transient Transfection of CHO-K1 Cells with pSNAP-ADRB2 and pSNAP-H2B:
<ol>
	<li>Check the cell culture chambers seeded on the previous day to look for cell density and health.</li>
	<li>Label the appropriate number of microfuge tubes.</li>
	<li>In a laminar flow hood, set up transfection complex mixtures according to the following sample chart, using either 0.3 &mu;g pSNAP-ADBR2 Control Plasmid DNA or 0.3 &mu;g of pSNAP-H2B Control Plasmid DNA, 1 &mu;l of TransPass D2 and 1 &mu;l of TransPass V per reaction.
<table border="1" cellspacing="0" cellpadding="0">
	<tbody>
		<tr>
		 <td>plasmid</td>
		 <td>number of reactions</td>
		 <td>&mu;l of serum-free medium</td>
		 <td>&mu;l of TransPass D2</td>
		 <td>&mu;l of TransPass V</td>
		 <td>&mu;l of plasmid DNA</td>
 		 <td>&mu;l of TransPass D2/reaction</td>
 	 	 <td>&mu;l of TransPass V/reaction</td>
		 <td>ng of DNA/ reaction</td>
		 <td>DNA conc. (ng/&mu;l)</td>
		 <td>&mu;l of plasmid DNA/ reaction</td>
 		 </tr>
 		 <tr>
		 <td>SNAP-ADRb2</td>
		 <td>5</td>
		 <td>236.5</td>
		 <td>5</td>
		 <td>5</td>
		 <td>3.5</td>
		 <td>1</td>
		 <td>1</td>
		 <td>300</td>
		 <td>472.34</td>
		 <td>0.7</td>
	 </tr>
 	 <tr>
		 <td>H2B-SNAP</td>
		 <td>5</td>
 		 <td>237</td>
 		 <td>5</td>
 		 <td>5</td>
 		 <td>3</td>
 		 <td>1</td>
		 <td>1</td>
		 <td>300</td>
		 <td>515.89</td>
		 <td>0.6</td>
	 </tr>
	 <tr>
		 <td>Mock</td>
		 <td>5</td>
		 <td>240</td>
		 <td>5</td>
		 <td>5</td>
		 <td>0</td>
		 <td>1</td>
		 <td>1</td>
		 <td>0</td>
		 <td>0</td>
		 <td>0</td>
 	 </tr>
 	 <tr>
 		 <td>Untransfected</td>
 		 <td>5</td>
 		 <td>250</td>
 		 <td>0</td>
 		 <td>0</td>
 		 <td>0</td>
 		 <td>0</td>
 		 <td>0</td>
 		 <td>0</td>
 		 <td>0</td>
		 <td>0</td>
	 </tr>
	</tbody>
</table>
</li>
	<li>First, mix the serum free F-12K with the appropriate plasmid DNA, then add the TransPass D2 Transfection Reagent, then add the TransPass V Transfection Reagent. Mix the components well by pipetting up and down gently several times after the addition of each reagent.</li>
	<li>Incubate the reaction at room temperature for 30 minutes.</li>
	<li>During the incubation, wash cells once with 600 &mu;l of complete DMEM and add 450 &mu;l of complete DMEM to each sample.</li>
	<li>Add 50 &mu;l of the transfection complex mixture to each sample, slowly and in a drop-wise fashion.</li>
	<li>Incubate the samples overnight at 37&deg;C, 5% CO2.</li>
</ol>
SNAP-Cell TMR Star Labeling of CHO-K1 Cells Transiently Transfected with pSNAP-ADRB2 and pSNAP-H2B :
<ol>
	<li>Carefully remove the transfection complex media from each sample by vacuum suction. Wash cells twice with 600 &mu;l of complete F-12K and replace media in each well with 600 &mu;l of complete F-12K.</li>
	<li>Mix dye labeling media by adding 995 &mu;l of complete F-12K and 51 &mu;l 0.6 mM SNAP-Cell TMR-Star&nbsp; substrate. The final concentration of SNAP-Cell TMR-Star should be 3mM.&nbsp; Pipette up and down several times to mix after the addition of substrate.</li>
	<li>Carefully remove growth media from samples and add 200 &mu;l of dye labeling media to each well. Incubate samples at 37&deg;C, 5% CO2 for 30 minutes.</li>
	<li>While samples are incubating, prepare Hoechst 33342 solution for nuclear staining of samples by mixing 1 &mu;l of Hoechst 33342, trihydrochloride, trihydrate (10 mg/ml solution in water, 16.2 mM solution; Invitrogen Molecular Probes) in 10 ml of complete F-12K.</li>
	<li>Remove the media from all samples and add 600 &mu;l of the dilute Hoechst solution to each sample. Incubate samples at 37&deg;C, 5% CO2 for 5 minutes.</li>
	<li>Wash samples three times with 600 &mu;l of complete F-12K.</li>
	<li>Incubate the samples at 37&deg;C, 5% CO2 for an additional 30 minutes to allow unincorporated fluorophore to diffuse out of the cells.</li>
	<li>After 30 minutes, change the media on the SNAP-Cell samples one last time and proceed to imaging.</li>
	<li>Image the cells using a Zeiss Apotome Axiovert 200M fluorescent microscope.</li>
</ol>
Representative Results
<img src="/files/ftp_upload/1876/1876fig1.jpg" alt="Figure 1" /> Figure 1. Here are some representative results of fluorescent imaging using standard fluorescent microscopy of live COS-7 cells that are expressing SNAP-tag fusions. This first image shows live COS-7 cells expressing pSNAP-H2B (nuclear histone) labeled with SNAP-Cell TMR-Star. The pSNAP-H2B construct was generated using the pSNAP-tag(m) vector. The cells were labeled with SNAP-Cell TMR-Star for 30 minutes and counterstained with Hoechst 33342 (blue) for nuclei. The pink fluorescence demonstrates the overlay of red fluorescence where histone protein is present in addition to the blue fluorescence indicating the nucleus. The expression of the H2B protein is clearly and easily identified.
<img src="/files/ftp_upload/1876/1876fig2.jpg" alt="Figure 2" /> <img src="/files/ftp_upload/1876/1876fig3.jpg" alt="Figure 3" /> Figures 2 and 3. These next two images show live COS-7 cells transiently transfected with pSNAP-ADR&beta;2. Cells were labeled with SNAP-Cell TMR-Star (red) and counterstained with Hoechst 33342 (blue). The pSNAP-ADR&beta;2 construct was generated using the pSNAP-tag(m) vector. The cells were labeled with SNAP-Cell TMR-Star for 30 minutes and counterstained with Hoechst 33342 (blue) for nuclei. The red fluorescence indicates the presence of cell surface receptor protein ADRB2 while blue fluorescence identifies the nucleus.

<ol>
	<li>Johnsson, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Visualizing+biochemical+activities+in+living+cells.">Visualizing biochemical activities in living cells.</a> Nat Chem Biol. 5, 63-65 (2009).</li><li>Johnsson, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=SNAP-tag+Technologies:+Novel+Tools+to+Study+Protein+Function.">SNAP-tag Technologies: Novel Tools to Study Protein Function.</a> NEB Expressions. 3.3, 1-3 (2008).</li></ol>

The authors are employed by New England Biolabs that produces the reagents and instruments used in this article.

There are several variables affecting the observed results such as focus, laser intensity, equipment capabilities, lens used, etc. The photostability of the fluorophore determines how quickly the fluorescence must be captured before fading.
This method of protein labeling is versatile and suitable for many applications. SNAP-tag and CLIP-tag technologies for the specific labeling of fusion proteins with synthetic probes enable various aspects of protein function, including a variety of dynamic processes in live cells and in cell lysates. SNAP-, CLIP-, MCP- and ACP-tag technologies provide simplicity and extraordinary versatility to the imaging of proteins in live and fixed cells and to the study of proteins in vitro. The creation of a single gene construct yields a tagged fusion protein capable of forming a covalent linkage to a variety of functional groups, including fluorophores, biotin, or beads. The protein of interest must be cloned and expressed only once and then the fusion can be used with a variety of fluorescent substrates for numerous downstream applications such as simultaneous protein labeling inside live cells, protein localization and translocation, pulse-chase experiments, receptor internalization studies, selective cell surface labeling, protein pull down assays, protein detection in SDS-PAGE, flow cytometry, high-throughput binding assays in microtiter plates, biosensor interaction experiments, and FRET-based binding assays.

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<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>CHO-K1 Cells</td>
<td>&nbsp;</td>
<td>ATCC</td>
<td>CCL-61</td>
<td>&nbsp;</td>
<tr>
<td>Serum free F-12K</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>Lab-Tek II Chambered Coverglass with #1.5 borosilicate coverglass chambers</td>
<td>&nbsp;</td>
<td>Nalgene Nunc Int.</td>
<td>155409</td>
<td>Lot: 100808-8-0</td>
</tr>
<tr>
<td>DMEM</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>Hoechst 33342, trihydrochloride, trihydrate (10 mg/ml solution in water, 16.2 mM solution)</td>
<td>&nbsp;</td>
<td>Invitrogen Molecular Probes</td>
<td>H3570</td>
<td>470519</td>
</tr>
<tr>
<td>Zeiss Apotome Axiovert 200M fluorescent microscope</td>
<td>&nbsp;</td>
<td>Zeiss</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>Transpass V</td>
<td>&nbsp;</td>
<td>New England Biolabs</td>
<td>M2561S</td>
<td>0030709</td>
</tr>
<tr>
<td>Transpass D2</td>
<td>&nbsp;</td>
<td>New England Biolabs</td>
<td>M2554S</td>
<td>0060802</td>
</tr>
<tr>
<td>SNAP-Cell TMR Star</td>
<td>&nbsp;</td>
<td>New England Biolabs</td>
<td>S9105S</td>
<td>0010811</td>
</tr>
<tr>
<td>pSNAP-ADBR2 Control Plasmid</td>
<td>&nbsp;</td>
<td>New England Biolabs</td>
<td>N9178S</td>
<td>0010811</td>
</tr>
<tr>
<td>pSNAP-H2B Control Plasmid </td>
<td>&nbsp;</td>
<td>New England Biolabs</td>
<td>N9179S</td>
<td>0010811</td>
</tr>		</tbody>
		</table>
		</div>

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.

Watch this Scientific Journal Video about Fluorescent Labeling of COS-7 Expressing SNAP-tag Fusion Proteins for Live Cell Imaging at JoVE.com

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. 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 ...

fluorescent-labeling-cos-7-expressing-snap-tag-fusion-proteins-for

Research

JoVE Journal

Biology

22.9K Views.  New England Biolabs. 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.

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

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 ...

Live Cell Imaging of F-actin Dynamics via Fluorescent Speckle Microscopy (FSM)

In this protocol we describe the use of Fluorescent Speckle Microscopy (FSM) to capture high-resolution images of actin dynamics in PtK1 cells. A unique advantage of FSM is its ability to capture the movement and turnover kinetics (assembly/disassembly) of the F-actin network within living cells. This technique is particularly useful in deriving quantitative measurements of F-actin dynamics when paired with computer vision software (qFSM).  We describe  the selection, microinjection and visualization of fluorescent actin probes in living cells. Importantly, similar procedures are applicable to visualizing other macomolecular assemblies. FSM has been demonstrated for microtubules, intermediate filaments, and adhesion complexes.  

Live Cell Imaging of F-actin Dynamics via Fluorescent Speckle Microscopy FSM

Selection, microinjection, and imaging of fluorescently-labeled F-actin via fluorescent speckle microscopy (FSM).

In this protocol we describe the use of Fluorescent Speckle Microscopy (FSM) to capture high-resolution images of actin dynamics in PtK1 cells. A unique ...

Fluorescent Labeling of Drosophila Heart Structures

The Drosophila melanogaster dorsal vessel, or heart, is a tubular structure comprised of a single layer of contractile cardiomyocytes, pericardial cells that align along each side of the heart wall, supportive alary muscles and, in adults, a layer of ventral longitudinal muscle cells. The contractile fibers house conserved constituents of the muscle cytoarchitecture including densely packed bundles of myofibrils and cytoskeletal/submembranous protein complexes, which interact with homologous components of the extracellular matrix. Here we describe a protocol for the fixation and the fluorescent labeling of particular myocardial elements from the hearts of dissected larvae and semi-intact adult Drosophila. Specifically, we demonstrate the labeling of sarcomeric F-actin and of &alpha;-actinin in larval hearts. Additionally, we perform labeling of F-actin and &alpha;-actinin in myosin-GFP expressing adult flies and of &alpha;-actinin and pericardin, a type IV extracellular matrix collagen, in wild type adult hearts. Particular attention is given to a mounting strategy for semi-intact adult hearts that minimizes handling and optimizes the opportunity for maintaining the integrity of the cardiac tubes and the associated tissues. These preparations are suitable for imaging via fluorescent and confocal microscopy. Overall, this procedure allows for careful and detailed analysis of the structural characteristics of the heart from a powerful genetically tractable model system.

Fluorescent Labeling of Drosophila Heart Structures

Here we describe a basic protocol for fluorescent labeling of different elements of heart tubes from larva and adult Drosophila melanogaster. These specimens are well-suited for imaging via fluorescent or confocal microscopy. This technique permits detailed structural analysis of the features of the hearts from a powerful model organism.

The Drosophila melanogaster dorsal vessel, or heart, is a tubular structure comprised of a single layer of contractile cardiomyocytes, pericardial cells ...

Transplantation of GFP-expressing Blastomeres for Live Imaging of Retinal and Brain Development in Chimeric Zebrafish Embryos

Cells change extensively in their locations and property during embryogenesis. These changes are regulated by the interactions between the cells and their environment. Chimeric embryos, which are composed of cells of different genetic background, are great tools to study the cell-cell interactions mediated by genes of interest. The embryonic transparency of zebrafish at early developmental stages permits direct visualization of the morphogenesis of tissues and organs at the cellular level. Here, we demonstrate a protocol to generate chimeric retinas and brains in zebrafish embryos and to perform live imaging of the donor cells. The protocol covers the preparation of transplantation needles, the transplantation of GFP-expressing donor blastomeres to GFP-negative hosts, and the examination of donor cell behavior under live confocal microscopy. With slight modifications, this protocol can also be used to study the embryonic development of other tissues and organs in zebrafish. The advantages of using GFP to label donor cells are also discussed. 

We demonstrate a protocol to generate chimeric zebrafish embryos for live imaging cellular behavior during embryogenesis.

Cells change extensively in their locations and property during embryogenesis. These changes are regulated by the interactions between the cells and their ...

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.

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 ...

Two- and Three-Dimensional Live Cell Imaging of DNA Damage Response Proteins

Double-strand breaks (DSBs) are the most deleterious DNA lesions a cell can encounter. If left unrepaired, DSBs harbor great potential to generate mutations and chromosomal aberrations1. To prevent this trauma from catalyzing genomic instability, it is crucial for cells to detect DSBs, activate the DNA damage response (DDR), and repair the DNA. When stimulated, the DDR works to preserve genomic integrity by triggering cell cycle arrest to allow for repair to take place or force the cell to undergo apoptosis. The predominant mechanisms of DSB repair occur through nonhomologous end-joining (NHEJ) and homologous recombination repair (HRR) (reviewed in2). There are many proteins whose activities must be precisely orchestrated for the DDR to function properly. Herein, we describe a method for 2- and 3-dimensional (D) visualization of one of these proteins, 53BP1.
The p53-binding protein 1 (53BP1) localizes to areas of DSBs by binding to modified histones3,4, forming foci within 5-15 minutes5. The histone modifications and recruitment of 53BP1 and other DDR proteins to DSB sites are believed to facilitate the structural rearrangement of chromatin around areas of damage and contribute to DNA repair6. Beyond direct participation in repair, additional roles have been described for 53BP1 in the DDR, such as regulating an intra-S checkpoint, a G2/M checkpoint, and activating downstream DDR proteins7-9. Recently, it was discovered that 53BP1 does not form foci in response to DNA damage induced during mitosis, instead waiting for cells to enter G1 before localizing to the vicinity of DSBs6. DDR proteins such as 53BP1 have been found to associate with mitotic structures (such as kinetochores) during the progression through mitosis10.
In this protocol we describe the use of 2- and 3-D live cell imaging to visualize the formation of 53BP1 foci in response to the DNA damaging agent camptothecin (CPT), as well as 53BP1's behavior during mitosis. Camptothecin is a topoisomerase I inhibitor that primarily causes DSBs during DNA replication. To accomplish this, we used a previously described 53BP1-mCherry fluorescent fusion protein construct consisting of a 53BP1 protein domain able to bind DSBs11. In addition, we used a histone H2B-GFP fluorescent fusion protein construct able to monitor chromatin dynamics throughout the cell cycle but in particular during mitosis12. Live cell imaging in multiple dimensions is an excellent tool to deepen our understanding of the function of DDR proteins in eukaryotic cells.

This protocol describes a method for visualizing a DNA double-strand break signaling protein activated in response to DNA damage as well as its localization during mitosis.

Double-strand breaks (DSBs) are the most deleterious DNA lesions a cell can encounter. If left unrepaired, DSBs harbor great potential to generate ...

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 ...

Live Imaging of Apoptotic Cell Clearance during Drosophila Embryogenesis

The proper elimination of unwanted or aberrant cells through apoptosis and subsequent phagocytosis (apoptotic cell clearance) is crucial for normal development in all metazoan organisms. Apoptotic cell clearance is a highly dynamic process intimately associated with cell death; unengulfed apoptotic cells are barely seen in vivo under normal conditions. In order to understand the different steps of apoptotic cell clearance and to compare 'professional' phagocytes - macrophages and dendritic cells to 'non-professional' - tissue-resident neighboring cells, in vivo live imaging of the process is extremely valuable. Here we describe a protocol for studying apoptotic cell clearance in live Drosophila embryos. To follow the dynamics of different steps in phagocytosis we use specific markers for apoptotic cells and phagocytes. In addition, we can monitor two phagocyte systems in parallel: 'professional' macrophages and 'semi-professional' glia in the developing central nervous system (CNS). The method described here employs the Drosophila embryo as an excellent model for real time studies of apoptotic cell clearance.

Live Imaging of Apoptotic Cell Clearance during Drosophila Embryogenesis

Here we describe an effective method for studying dynamics of apoptotic cell clearance in vivo. This method employs live Drosophila embryos as a powerful model for monitoring phagocytosis of apoptotic cells using specific labeling of apoptotic cells and phagocytes.

The  proper elimination of unwanted or aberrant cells through apoptosis and  subsequent phagocytosis (apoptotic cell clearance) is crucial for normal  ...

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, ...

Expression of Fluorescent Fusion Proteins in Murine Bone Marrow-derived Dendritic Cells and Macrophages

Dendritic cells and macrophages are crucial cells that form the first line of defense against pathogens. They also play important roles in the initiation of an adaptive immune response. Experimental work with these cells is rather challenging. Their abundance in organs and tissues is relatively low. As a result, they cannot be isolated in large numbers. They are also difficult to transfect with cDNA constructs. In the murine model, these problems can be partially overcome by in vitro differentiation from bone marrow progenitors in the presence of M-CSF for macrophages or GM-CSF for dendritic cells. In this way, it is possible to obtain large amounts of these cells from very few animals. Moreover, bone marrow progenitors can be transduced with retroviral vectors carrying cDNA constructs during early stages of cultivation prior to their differentiation into bone marrow derived dendritic cells and macrophages. Thus, retroviral transduction followed by differentiation in vitro can be used to express various cDNA constructs in these cells. The ability to express ectopic proteins substantially extends the range of experiments that can be performed on these cells, including live cell imaging of fluorescent proteins, tandem purifications for interactome analyses, structure-function analyses, monitoring of cellular functions with biosensors and many others. In this article, we describe a detailed protocol for retroviral transduction of murine bone marrow derived dendritic cells and macrophages with vectors coding for fluorescently-tagged proteins. On the example of two adaptor proteins, OPAL1 and PSTPIP2, we demonstrate its practical application in flow cytometry and microscopy. We also discuss the advantages and limitations of this approach.

In this article, we provide a detailed protocol for the expression of fluorescent fusion proteins in murine bone marrow derived dendritic cells and macrophages. The method is based on the transduction of bone marrow progenitors with retroviral constructs followed by differentiation into macrophages and dendritic cells in vitro.

Dendritic cells and macrophages are crucial cells that form the first line of defense against pathogens. They also play important roles in the initiation ...