This work was funded by grants from the NIH and ACS, including T32CA009135, CA133027, CA070723, and CA022443. Bill Sugden is an American Cancer Society Research Professor.

1. Engineering of Cells with Visible Plasmids
<ol>
	<li>Use standard restriction digestion or homologous recombination techniques to introduce a DNA fragment containing approximately 250 copies of the lactose operator sequence (LacO) into the plasmid to be visualized. Our plasmid was a BACmid of approximately 170 kbp and contained the entire genome of Kaposi's Sarcoma-associated Herpes Virus (KSHV) and encoded resistance to hygromycin 12.</li>
	<li>Introduce LacO-containing plasmid DNA into mammalian cells by transfection with Lipofectamine 2000. We transfected HeLa and SLK cells in 60 mm dishes for 4 hr with 10 &mu;g purified plasmid DNA with 25 &mu;l Lipofectamine 2000, 0.5 ml OptiMEM, and 3.5 ml DMEM.</li>
	<li>Change the culture medium to 5 ml DMEM with 10% fetal bovine serum and incubate cells for 48 hr.</li>
	<li>Plate the cells at low density in large dishes and culture cells in medium containing an antibiotic selection for the introduced plasmid; we selected cells in DMEM with 10% fetal bovine serum with 300 &mu;g/ml hygromycin for 3 weeks.</li>
	<li>Use pieces of sterile, trypsin-soaked filter paper to transfer hygromycin-resistant colonies to new culture dishes.</li>
	<li>When transfected clones have grown to fill the dish, isolate DNA for restriction digestion and Southern blotting to ensure that the polymer of the LacO plasmid is homogenous and of the expected size.</li>
	<li>If the plasmid has not been engineered to express a fusion of LacI, infect suitable clones with a retrovirus encoding the lactose repressor (LacI) fused to a red fluorescent protein, such as LacI-tdTomato. We use tdTomato rather than proteins that fluoresce closer to green to minimize photoxicity that would occur from multiple exposures over long times13.</li>
	<li>Once the LacI fusion protein is introduced, culture the cells in the presence of 200 &mu;g/ml isopropyl-b-D-thiogalactoside (IPTG) to block the fusion proteins from binding DNA.</li>
	<li>Culture the cells for at least 3 days to allow expression of the LacI-tdTomato protein.</li>
	<li>Screen clones (by washing away the IPTG and viewing cells as described below) for those with optimal levels of LacI-tdTomato that reveal distinct signals with minimal background levels of unbound fusion protein.</li>
</ol>
2. Development of the Imaging System
Our imaging system consists of a Zeiss Axiovert 200M equipped with a motorized stage and Plan-Aochromat 63x/1.4 oil objective. Fluorescence excitation light is provided by the "neutral white" LED of a Colibri system. Emission is detected by a CascadeII:1024 EMCCD camera. This is a cooled, back-thinned camera with a gain register for amplifying signals; the dark current is 0.061 electrons per pixel per second, and the quantum efficiency is greater than 0.90. The system is controlled by Axiovision 4.8 software, including the SmartExperiments module. For detection of tdTomato we use a Zeiss filter set 75HE for which 85% of the passed light can excite the fluor and 95% of the emitted light will pass the emission filter.
<ol>
	<li>Assemble an imaging system with an inverted microscope and an EMCCD camera for sensitive detection of signals, a LED light source for specific excitation and fast shuttering, a motorized stage and focus arm, and software control for automated acquisitions of images over long time intervals with multiple z-stacks. The system should sit on an air table to be isolated from vibration.</li>
	<li>Assemble a stage-top incubator to maintain cells at 37 &deg;C, 5-10% CO2 in a humidified chamber. Allow sufficient time for the system to come to a stable temperature to minimize movement artifacts during the experiment.</li>
	<li>Use a heated collar for the objective to prevent the objective from siphoning heat from the sample.</li>
</ol>
3. Visualization of Labeled DNA
<ol>
	<li>Plate cells in a 35 mm glass-bottom dish at a density of less than 10% confluence, which will allow pairs of cells to divide into colonies of 8-16 cells without overlapping. Do this at least 24-48 hr before imaging so that the viable cells have time to divide.</li>
	<li>One to three hr before the start of imaging, rinse the plate 3 times with culture medium which lacks both selective drugs and IPTG. This procedure washes away non-viable cells and allows LacI fusion proteins to bind LacO sites in the plasmids.</li>
	<li>Replace the culture medium with 2 ml DMEM with 10% fetal bovine serum and 25 mM HEPES.</li>
	<li>Replace the top of the culture dish with CultFoil stretched in a mounting ring for a 35 mm dish. This cover will inhibit evaporation of the culture medium, which would increase salt concentration over time and kill the cells.</li>
	<li>Use differential interference contrast (DIC) imaging to view the cells and determine the focal plane of the top and bottom of the cells in multiple fields of view. In each field, move to a focal plane approximately 1/3 of the way down from the top of the cell. Save the x, y, z coordinates in the list of saved stage positions. 
	Note: Plasmid signals may not be visible through the eyepieces when cells are illuminated by LEDs of the wavelength appropriate for the LacI fusion protein. Weak signals may only be visible with the use of the EMCCD camera, which allows integration of the signal over time and thus effectively amplifies weak signals.</li>
	<li>In the microscope control software, define a z-stack of images to be performed at each saved stage position. Choose a z step size of 0.35-0.50 &mu;m, which will allow you to approximate Nyquist sampling in the z dimension14. Include "extra" slices above and below the planes in which the cells are located; these slices allow detection following cell movements during the cell cycle, and are also used in post-acquisition processing (deconvolution) of the image stacks. This inclusion will result in a stack of 40-60 images, depending on the thickness of the cells.</li>
	<li>In the microscope control software, define a time series experiment to collect a bright field image of the z-stacks at 30 min intervals (for tracking cell divisions and migrations over long times) and fluorescence images at 10-60 min intervals (for visualizing plasmids within cells). Do not use DIC imaging during the experiment, as it requires more light than standard bright field imaging, which can expose the cells unnecessarily to phototoxicity over the course of long experiments.</li>
	<li>Start the software-controlled experiment. Ensure that the system is not disturbed (bumped, exposed to excess room light, etc.) during the experiment.</li>
	<li>If necessary, replenish the water in the humidification system during the experiment.</li>
</ol>
4. Post-acquisition Processing of Images
<ol>
	<li>Review the images for each z-stack and time point. Crop out any unneeded areas of the images to reduce the computer processing time in the next step.</li>
	<li>Use a deconvolution algorithm to re-assign signal intensity to the pixel of origin in each z-stack15. While this deconvolution can be based on a theoretical point spread function for your imaging system, it is preferable to measure the point spread function of your particular system by imaging fluorescent microspheres, and use this measurement in the deconvolution algorithm. We measured the point spread function of our imaging system and applied a constrained iterative maximum likelihood algorithm in AxioVision software (AxioVision software description, Zeiss).</li>
	<li>Examine the images to track the intensity changes and movements of plasmids during the cell cycle and the partitioning of plasmids to daughter cells.</li>
</ol>

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A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Fields Virology 2 volume set. , (2006).</li><li>Adams, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Replication+of+latent+Epstein-Barr+virus+genomes+in+Raji+cells.">Replication of latent Epstein-Barr virus genomes in Raji cells.</a> J. Virol. 61, 1743-1746 (1987).</li><li>Humme, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+EBV+nuclear+antigen+1+(EBNA1)+enhances+B+cell+immortalization+several+thousandfold.">The EBV nuclear antigen 1 (EBNA1) enhances B cell immortalization several thousandfold.</a> Proc. Natl. Acad. Sci. U.S.A. 100, 10989-10994 (1073).</li><li>Verma, S. C., Choudhuri, T., Robertson, E. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+minimal+replicator+element+of+the+Kaposi's+sarcoma-associated+herpesvirus+terminal+repeat+supports+replication+in+a+semiconservative+and+cell-cycle-dependent+manner.">The minimal replicator element of the Kaposi's sarcoma-associated herpesvirus terminal repeat supports replication in a semiconservative and cell-cycle-dependent manner.</a> J. Virol. 81, 3402-3413 (2007).</li><li>Yates, J. L., Guan, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Epstein-Barr+virus-derived+plasmids+replicate+only+once+per+cell+cycle+and+are+not+amplified+after+entry+into+cells.">Epstein-Barr virus-derived plasmids replicate only once per cell cycle and are not amplified after entry into cells.</a> J. Virol. 65, 483-488 (1991).</li><li>Ye, F. 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No conflicts of interest declared.

The method described here can be used to follow plasmids in live mammalian cells over time spanning multiple generations. By limiting exposure to excitation light, we have used these techniques to follow cells from newly-divided pairs through colonies of 16-32 cells, representing at least 72 hr and 3 cell divisions. These experiments provide information that cannot be obtained from static assays such as quantitative PCR, Southern blotting, and FISH.
It is essential to screen clones of cel...

<table border="1">
 <tbody>
 <tr>
 <td>Name of the reagent </td>
 <td>Company </td>
 <td>Catalogue number </td>
 <td>Comments </td>
 </tr>
 <tr>
 <td>DMEM</td>
 <td>GIBCO</td>
 <td>11965</td>
 <td></td>
 </tr>
 <tr>
 <td>OptiMEM</td>
 <td>GIBCO</td>
 <td>31985</td>
 <td></td>
 </tr>
 <tr>
 <td>Fetal bovine serum</td>
 <td>Hyclone</td>
 <td>SH30910.03</td>
 <td></td>
 </tr>
 <tr>
 <td>Penicillin</td>
 <td>Sigma</td>
 <td>P3032</td>
 <td></td>
 </tr>
 <tr>
 <td>Streptomycin sulfate</td>
 <td>Sigma</td>
 <td>S9137</td>
 <td></td>
 </tr>
 <tr>
 <td>Hygromycin</td>
 <td>Calbiochem</td>
 <td>400050</td>
 <td>400 &mu;g/ml for SLK; 300 &mu;g/ml for HeLa</td>
 </tr>
 <tr>
 <td>Isopropyl-b-D-thiogalactoside (IPTG)</td>
 <td>Roche</td>
 <td>10 724 815 001</td>
 <td>Final concentration 200 &mu;g/ml </td>
 </tr>
 <tr>
 <td>Lipofectamine 2000</td>
 <td>Invitrogen</td>
 <td>11668-027</td>
 <td></td>
 </tr>
 <tr>
 <td>HEPES</td>
 <td>GIBCO</td>
 <td>15630</td>
 <td></td>
 </tr>
 <tr>
 <td>Glass-bottom dishes</td>
 <td>MatTek</td>
 <td>P35G-1.5-10-C</td>
 <td></td>
 </tr>
 <tr>
 <td>CultFoil</td>
 <td>Pecon</td>
 <td>000000-1116-08</td>
 <td></td>
 </tr>
 <tr>
 <td>Axiovert 200M</td>
 <td>Zeiss</td>
 <td></td>
 <td></td>
 </tr>
 <tr>
 <td>Colibri</td>
 <td>Zeiss</td>
 <td>423052-9500-000</td>
 <td></td>
 </tr>
 <tr>
 <td>Neutral white LED</td>
 <td>Zeiss</td>
 <td>423052-9120-000</td>
 <td></td>
 </tr>
 <tr>
 <td>Filter Cube 75 HE</td>
 <td>Zeiss</td>
 <td>489075-0000-000</td>
 <td></td>
 </tr>
 <tr>
 <td>Plan-Apochromat 63x/1.4 objective</td>
 <td>Zeiss</td>
 <td>440762-9904-000</td>
 <td></td>
 </tr>
 <tr>
 <td>CascadeII:1024 EMCCD camera</td>
 <td>Photometrics</td>
 <td>B10C892007</td>
 <td></td>
 </tr>
 <tr>
 <td>Tempcontrol mini</td>
 <td>Zeiss/Pecon</td>
 <td>000000-1116-070</td>
 <td></td>
 </tr>
 <tr>
 <td>Tempcontrol 37-2 digital</td>
 <td>Zeiss/Pecon</td>
 <td>000000-1052-320</td>
 <td></td>
 </tr>
 <tr>
 <td>CTI Controller 3700 digital</td>
 <td>Zeiss/Pecon</td>
 <td>411856-9903</td>
 <td></td>
 </tr>
 <tr>
 <td>Objective heater</td>
 <td>Zeiss/Pecon</td>
 <td>440760-0000-000</td>
 <td></td>
 </tr>
 <tr>
 <td>Stage heating insert P</td>
 <td>Zeiss/Pecon</td>
 <td>411861-9901-000</td>
 <td></td>
 </tr>
 <tr>
 <td>Stage-top Incubator S</td>
 <td>Zeiss/Pecon</td>
 <td>411860-9902-000</td>
 <td></td>
 </tr>
 <tr>
 <td>Humidifier System</td>
 <td>Zeiss/Pecon</td>
 <td>000000-1116-065</td>
 <td></td>
 </tr>
 </tbody>
</table>

monitoring plasmid replication in live mammalian cells over multiple generations by fluorescence microscopy

Few naturally-occurring plasmids are maintained in mammalian cells. Among these are genomes of gamma-herpesviruses, including Epstein-Barr virus (EBV) and Kaposi's Sarcoma-associated herpesvirus (KSHV), which cause multiple human malignancies 1-3. These two genomes are replicated in a licensed manner, each using a single viral protein and cellular replication machinery, and are passed to daughter cells during cell division despite their lacking traditional centromeres 4-8.
Much work has been done to characterize the replications of these plasmid genomes using methods such as Southern blotting and fluorescence in situ hybridization (FISH). These methods are limited, though. Quantitative PCR and Southern blots provide information about the average number of plasmids per cell in a population of cells. FISH is a single-cell assay that reveals both the average number and the distribution of plasmids per cell in the population of cells but is static, allowing no information about the parent or progeny of the examined cell.
Here, we describe a method for visualizing plasmids in live cells. This method is based on the binding of a fluorescently tagged lactose repressor protein to multiple sites in the plasmid of interest 9. The DNA of interest is engineered to include approximately 250 tandem repeats of the lactose operator (LacO) sequence. LacO is specifically bound by the lactose repressor protein (LacI), which can be fused to a fluorescent protein. The fusion protein can either be expressed from the engineered plasmid or introduced by a retroviral vector. In this way, the DNA molecules are fluorescently tagged and therefore become visible via fluorescence microscopy. The fusion protein is blocked from binding the plasmid DNA by culturing cells in the presence of IPTG until the plasmids are ready to be viewed.
This system allows the plasmids to be monitored in living cells through several generations, revealing properties of their synthesis and partitioning to daughter cells. Ideal cells are adherent, easily transfected, and have large nuclei. This technique has been used to determine that 84% of EBV-derived plasmids are synthesized each generation and 88% of the newly synthesized plasmids partition faithfully to daughter cells in HeLa cells. Pairs of these EBV plasmids were seen to be tethered to or associated with sister chromatids after their synthesis in S-phase until they were seen to separate as the sister chromatids separated in Anaphase10. The method is currently being used to study replication of KSHV genomes in HeLa cells and SLK cells. HeLa cells are immortalized human epithelial cells, and SLK cells are immortalized human endothelial cells. Though SLK cells were originally derived from a KSHV lesion, neither the HeLa nor SLK cell line naturally harbors KSHV genomes11. In addition to studying viral replication, this visualization technique can be used to investigate the effects of the addition, removal, or mutation of various DNA sequence elements on synthesis, localization, and partitioning of other recombinant plasmid DNAs.

Maximum intensity projections of z-stacks acquired in a representative experiment are shown in Figure 3. Typical plasmid signals are present in 3-8 slices of the z-stack, depending on whether they represent single or clusters of plasmids. In the case of EBV- and KSHV-derived plasmids, the signals move slowly within the cell until the cell reaches mitosis. The cells themselves migrate over the course of the experiment. They also become spherical and detach from the dish during mitosis. The cell cycl...

Watch this Scientific Journal Video about Monitoring Plasmid Replication in Live Mammalian Cells over Multiple Generations by Fluorescence Microscopy at JoVE.com

Monitoring Plasmid Replication in Live Mammalian Cells over Multiple Generations by Fluorescence Microscopy

A method of observing individual DNA molecules in live cells is described. The technique is based on the binding of a fluorescently tagged lac repressor protein to binding sites engineered into the DNA of interest. This method can be adapted to follow many recombinant DNAs in live cells over time.

Few naturally-occurring plasmids are maintained in mammalian cells.  Among these are genomes of gamma-herpesviruses, including Epstein-Barr virus  (EBV) ...

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11.7K Görüntüleme.  University of Wisconsin - Madison. Canlı hücreler içinde ayrı DNA molekülleri gözlenmesi için bir yöntem tarif edilmektedir. Bu teknik ilgi DNA içine yerleştirilmiştir bağlanma yerleri için bir floresan etiketlendi lac represör proteininin bağlayıcı esas alınmaktadır. Bu yöntem, zaman içindeki çok sayıda canlı hücreler rekombinant DNA izlemek için adapte edilebilir.

Video: Monitoring Plasmid Replication in Live Mammalian Cells over Multiple Generations by Fluorescence Microscopy

Long-term Imaging Mammalian Cells using Wide-Field Microscopy

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.

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

Time-lapse Imaging of Mitosis After siRNA Transfection

Changes in cellular organization and chromosome dynamics that occur during mitosis are tightly coordinated to ensure accurate inheritance of genomic and cellular content. Hallmark events of mitosis, such as chromosome movement, can be readily tracked on an individual cell basis using time-lapse fluorescence microscopy of mammalian cell lines expressing specific GFP-tagged proteins. In combination with RNAi-based depletion, this can be a powerful method for pinpointing the stage(s) of mitosis where defects occur after levels of a particular protein have been lowered. In this protocol, we present a basic method for assessing the effect of depleting a potential mitotic regulatory protein on the timing of mitosis. Cells are transfected with siRNA, placed in a stage-top incubation chamber, and imaged using an automated fluorescence microscope. We describe how to use software to set up a time-lapse experiment, how to process the image sequences to make either still-image montages or movies, and how to quantify and analyze the timing of mitotic stages using a cell-line expressing mCherry-tagged histone H2B. Finally, we discuss important considerations for designing a time-lapse experiment. This strategy is complementary to other approaches and offers the advantages of 1) sensitivity to changes in kinetics that might not be observed when looking at cells as a population and 2) analysis of mitosis without the need to synchronize the cell cycle using drug treatments. The visual information from such imaging experiments not only allows the sub-stages of mitosis to be assessed, but can also provide unexpected insight that would not be apparent from cell cycle analysis by FACS.

Here we describe a basic protocol to image and quantify the mitotic timing of live mammalian tissue culture cells after siRNA transfection.

Changes in cellular organization and chromosome dynamics that occur during mitosis are tightly coordinated to ensure accurate inheritance of genomic and ...

A Fluorescence Microscopy Assay for Monitoring Mitophagy in the Yeast Saccharomyces cerevisiae

Autophagy is important for turnover of cellular components under a range of different conditions. It serves an essential homeostatic function as well as a quality control mechanism that can target and selectively degrade cellular material including organelles1-4. For example, damaged or redundant mitochondria (Fig. 1), not disposed of by autophagy, can represent a threat to cellular homeostasis and cell survival. In the yeast, Saccharomyces cerevisiae, nutrient deprivation (e.g., nitrogen starvation) or damage can promote selective turnover of mitochondria by autophagy in a process termed mitophagy 5-9.
We describe a simple fluorescence microscopy approach to assess autophagy. For clarity we restrict our description here to show how the approach can be used to monitor mitophagy in yeast cells. The assay makes use of a fluorescent reporter, Rosella, which is a dual-emission biosensor comprising a relatively pH-stable red fluorescent protein linked to a pH-sensitive green fluorescent protein. The operation of this reporter relies on differences in pH between the vacuole (pH ~ 5.0-5.5) and mitochondria (pH ~ 8.2) in living cells. Under growing conditions, wild type cells exhibit both red and green fluorescence distributed in a manner characteristic of the mitochondria. Fluorescence emission is not associated with the vacuole. When subjected to nitrogen starvation, a condition which induces mitophagy, in addition to red and green fluorescence labeling the mitochondria, cells exhibit the accumulation of red, but not green fluorescence, in the acidic vacuolar lumen representing the delivery of mitochondria to the vacuole. Scoring cells with red, but not green fluorescent vacuoles can be used as a measure of mitophagic activity in cells5,10-12.

A Fluorescence Microscopy Assay for Monitoring Mitophagy in the Yeast Saccharomyces cerevisiae

A robust approach to monitor the delivery of organelles to the acidic lumen of the yeast vacuole for degradation and recycling is described. The method relies on the specific labeling of target organelles with a genetically encoded dual-emission fluorescence pH-biosensor, and visualization of individual cells using fluorescence microscopy.

Autophagy is important for turnover of cellular components under a range of different conditions. It serves an essential homeostatic function as well as a ...

Fluorescence-based Measurement of Store-operated Calcium Entry in Live Cells: from Cultured Cancer Cell to Skeletal Muscle Fiber

Store operated Ca2+ entry (SOCE), earlier termed capacitative Ca2+ entry, is a tightly regulated mechanism for influx of extracellular Ca2+ into cells to replenish depleted endoplasmic reticulum (ER) or sarcoplasmic reticulum (SR) Ca2+ stores1,2. Since Ca2+ is a ubiquitous second messenger, it is not surprising to see that SOCE plays important roles in a variety of cellular processes, including proliferation, apoptosis, gene transcription and motility. Due to its wide occurrence in nearly all cell types, including epithelial cells and skeletal muscles, this pathway has received great interest3,4. However, the heterogeneity of SOCE characteristics in different cell types and the physiological function are still not clear5-7.
The functional channel properties of SOCE can be revealed by patch-clamp studies, whereas a large body of knowledge about this pathway has been gained by fluorescence-based intracellular Ca2+ measurements because of its convenience and feasibility for high-throughput screening. The objective of this report is to summarize a few fluorescence-based methods to measure the activation of SOCE in monolayer cells, suspended cells and muscle fibers5,8-10. The most commonly used of these fluorescence methods is to directly monitor the dynamics of intracellular Ca2+ using the ratio of F340nm and F380nm (510 nm for emission wavelength) of the ratiometric Ca2+ indicator Fura-2. To isolate the activity of unidirectional SOCE from intracellular Ca2+ release and Ca2+ extrusion, a Mn2+ quenching assay is frequently used. Mn2+ is known to be able to permeate into cells via SOCE while it is impervious to the surface membrane extrusion processes or to ER uptake by Ca2+ pumps due to its very high affinity with Fura-2. As a result, the quenching of Fura-2 fluorescence induced by the entry of extracellular Mn2+ into the cells represents a measurement of activity of SOCE9. Ratiometric measurement and the Mn+2 quenching assays can be performed on a cuvette-based spectrofluorometer in a cell population mode or in a microscope-based system to visualize single cells. The advantage of single cell measurements is that individual cells subjected to gene manipulations can be selected using GFP or RFP reporters, allowing studies in genetically modified or mutated cells. The spatiotemporal characteristics of SOCE in structurally specialized skeletal muscle can be achieved in skinned muscle fibers by simultaneously monitoring the fluorescence of two low affinity Ca2+ indicators targeted to specific compartments of the muscle fiber, such as Fluo-5N in the SR and Rhod-5N in the transverse tubules9,11,12.

The extent of store-operated Ca2+ entry (SOCE) can be monitored using fluorescent Ca2+ indicators. Mn2+ quenching of such indicators assays SOCE in cultured cells and skeletal muscle fibers. A technique allowing spatial and temporal resolution of SOCE by confocal imaging of mechanically skinned muscle fibers is also described.

Store operated Ca2+ entry (SOCE), earlier termed capacitative Ca2+ entry, is a tightly regulated mechanism for influx of extracellular Ca2+ into cells to ...

Quantitative Live Cell Fluorescence-microscopy Analysis of Fission Yeast

Several microscopy techniques are available today that can detect a specific protein within the cell. During the last decade live cell imaging using fluorochromes like Green Fluorescent Protein (GFP) directly attached to the protein of interest has become increasingly popular 1. Using GFP and similar fluorochromes the subcellular localisations and movements of proteins can be detected in a fluorescent microscope. Moreover, also the subnuclear localisation of a certain region of a chromosome can be studied using this technique. GFP is fused to the Lac Repressor protein (LacR) and ectopically expressed in the cell where tandem repeats of the lacO sequence has been inserted into the region of interest on the chromosome2. The LacR-GFP will bind to the lacO repeats and that area of the genome will be visible as a green dot in the fluorescence microscope. Yeast is especially suited for this type of manipulation since homologous recombination is very efficient and thereby enables targeted integration of the lacO repeats and engineered fusion proteins with GFP 3. Here we describe a quantitative method for live cell analysis of fission yeast. Additional protocols for live cell analysis of fission yeast can be found, for example on how to make a movie of the meiotic chromosomal behaviour 4. In this particular experiment we focus on subnuclear organisation and how it is affected during gene induction. We have labelled a gene cluster, named Chr1, by the introduction of lacO binding sites in the vicinity of the genes. The gene cluster is enriched for genes that are induced early during nitrogen starvation of fission yeast 5. In the strain the nuclear membrane (NM) is labelled by the attachment of mCherry to the NM protein Cut11 giving rise to a red fluorescent signal. The Spindle Pole body (SPB) compound Sid4 is fused to Red Fluorescent Protein (Sid4-mRFP) 6. In vegetatively growing yeast cells the centromeres are always attached to the SPB that is embedded in the NM 7. The SPB is identified as a large round structure in the NM. By imaging before and 20 minutes after depletion of the nitrogen source we can determine the distance between the gene cluster (GFP) and the NM/SPB. The mean or median distances before and after nitrogen depletion are compared and we can thus quantify whether or not there is a shift in subcellular localisation of the gene cluster after nitrogen depletion.

The fission yeast, Schizosaccharomyces pombe, is a good model system to study basic cellular processes. Here we describe a method to perform quantitative live cell analysis of fission yeast. In this particular experiment we focus on organisation of the genome within the cell nucleus, but the method can also be used to study cytosolic factors.

Several microscopy techniques are available today that can detect a specific protein within the cell. During the last decade live cell imaging using ...

FRET Microscopy for Real-time Monitoring of Signaling Events in Live Cells Using Unimolecular Biosensors

F&ouml;rster resonance energy transfer (FRET) microscopy continues to gain increasing interest as a technique for real-time monitoring of biochemical and signaling events in live cells and tissues. Compared to classical biochemical methods, this novel technology is characterized by high temporal and spatial resolution. FRET experiments use various genetically-encoded biosensors which can be expressed and imaged over time in situ or in vivo1-2. Typical biosensors can either report protein-protein interactions by measuring FRET between a fluorophore-tagged pair of proteins or conformational changes in a single protein which harbors donor and acceptor fluorophores interconnected with a binding moiety for a molecule of interest3-4. Bimolecular biosensors for protein-protein interactions include, for example, constructs designed to monitor G-protein activation in cells5, while the unimolecular sensors measuring conformational changes are widely used to image second messengers such as calcium6, cAMP7-8, inositol phosphates9 and cGMP10-11. Here we describe how to build a customized epifluorescence FRET imaging system from single commercially available components and how to control the whole setup using the Micro-Manager freeware. This simple but powerful instrument is designed for routine or more sophisticated FRET measurements in live cells. Acquired images are processed using self-written plug-ins to visualize changes in FRET ratio in real-time during any experiments before being stored in a graphics format compatible with the build-in ImageJ freeware used for subsequent data analysis. This low-cost system is characterized by high flexibility and can be successfully used to monitor various biochemical events and signaling molecules by a plethora of available FRET biosensors in live cells and tissues. As an example, we demonstrate how to use this imaging system to perform real-time monitoring of cAMP in live 293A cells upon stimulation with a &beta;-adrenergic receptor agonist and blocker.

F&#246;rster resonance energy transfer (FRET) microscopy is a powerful technique for real-time monitoring of signaling events in live cells using various biosensors as reporters. Here we describe how to build a customized epifluorescence FRET imaging system from commercially available components and how to use it for FRET experiments.

F&ouml;rster resonance energy transfer (FRET) microscopy  continues to gain increasing interest as a technique for real-time monitoring  of biochemical ...

In Vivo 4-Dimensional Tracking of Hematopoietic Stem and Progenitor Cells in Adult Mouse Calvarial Bone Marrow

Through a delicate balance between quiescence and proliferation, self renewal and production of differentiated progeny, hematopoietic stem cells (HSCs) maintain the turnover of all mature blood cell lineages. The coordination of the complex signals leading to specific HSC fates relies upon the interaction between HSCs and the intricate bone marrow microenvironment, which is still poorly understood[1-2].
We describe how by combining a newly developed specimen holder for stable animal positioning with multi-step confocal and two-photon in vivo imaging techniques, it is possible to obtain high-resolution 3D stacks containing HSPCs and their surrounding niches and to monitor them over time through multi-point time-lapse imaging. High definition imaging allows detecting ex vivo labeled hematopoietic stem and progenitor cells (HSPCs) residing within the bone marrow. Moreover, multi-point time-lapse 3D imaging, obtained with faster acquisition settings, provides accurate information about HSPC movement and the reciprocal interactions between HSPCs and stroma cells.
Tracking of HSPCs in relation to GFP positive osteoblastic cells is shown as an exemplary application of this method. This technique can be utilized to track any appropriately labeled hematopoietic or stromal cell of interest within the mouse calvarium bone marrow space.

In Vivo 4-Dimensional Tracking of Hematopoietic Stem and Progenitor Cells in Adult Mouse Calvarial Bone Marrow

The nature of the interactions between hematopoietic stem and progenitor cells (HSPCs) and bone marrow niches is poorly understood. Custom hardware modifications and a multi-step acquisition protocol allow the use of two-photon and confocal microscopy to image ex vivo labeled HSPCs homed within bone marrow areas, tracking interactions and movement.

Through a delicate balance between quiescence and proliferation, self renewal and production of differentiated progeny, hematopoietic stem cells (HSCs) ...

Detection of Intracellular Gene Expression in Live Cells of Murine, Human and Porcine Origin Using Fluorescence-labeled Nanoparticles

The reprogramming of somatic cells to induced pluripotent stem cells (iPS) has successfully been performed in different mammalian species including mouse, rat, human, pig and others. The verification of iPS clones mainly relies on the detection of the endogenous expression of different pluripotency genes. These genes mostly represent transcription factors which are located in the cell nucleus. Traditionally, the proof of their endogenous expression is supplied by immunohistochemical staining after fixation of the cells. This approach requires replicate cultures of each clone at this early stage to preserve validated clones for further experiments. The present protocol describes an approach with gene-specific nanoparticles which allows the evaluation of intracellular gene expression directly in live cells by fluorescence. The nanoparticles consist of a central gold particle coupled to a capture strand carrying a sequence complementary to the target mRNA as well as a quenched reporter strand. These nanoparticles are actively endocytosed and the target mRNA displaces the reporter strand which then start to fluoresce. Therefore, specific target gene expression can be detected directly under the microscope. In addition, the emitted fluorescence allows the identification, isolation and enrichment of cells expressing a specific gene by flow cytometry. This method can be applied directly to live cells in culture without any manipulation of the target cells.

This manuscript describes a novel technique which allows the detection of intracellular gene expression after endocytosis of fluorescence-labeled nanoparticles directly in live cells. The method does not require manipulation of the cells and is not restricted with respect to the target gene or species.

The reprogramming of somatic cells to induced pluripotent stem cells (iPS) has successfully been performed in different mammalian species including mouse, ...

Fecal Glucocorticoid Analysis: Non-invasive Adrenal Monitoring in Equids

Adrenal activity can be assessed in the equine species by analysis of feces for corticosterone metabolites. During a potentially aversive situation, corticotrophin releasing hormone (CRH) is released from the hypothalamus in the brain. This stimulates the release of adrenocorticotrophic hormone (ACTH) from the pituitary gland, which in turn stimulates release of glucocorticoids from the adrenal gland. In horses the glucocorticoid corticosterone is responsible for several adaptations needed to support equine flight behaviour and subsequent removal from the aversive situation. Corticosterone metabolites can be detected in the feces of horses and assessment offers a non-invasive option to evaluate long term patterns of adrenal activity. Fecal assessment offers advantages over other techniques that monitor adrenal activity including blood plasma and saliva analysis. The non-invasive nature of the method avoids sampling stress which can confound results. It also allows the opportunity for repeated sampling over time and is ideal for studies in free ranging horses. This protocol describes the enzyme linked immunoassay (EIA) used to assess feces for corticosterone, in addition to the associated biochemical validation.

Adrenal activity can be assessed in the equine species by analysis of feces for corticosterone metabolites. The method offers a non-invasive option to assess long term patterns in both domestic and free ranging horses. This protocol describes the enzyme linked immunoassay involved and the associated biochemical validation.

Adrenal activity can be assessed in the equine species by analysis of feces for corticosterone metabolites. During a potentially aversive situation, ...