We thank Dr. Seth Robia at Loyola University Chicago for his valuable comments on this manuscript. This work was supported by NIH grant 1R01NS073967-01A1 to Joanna C. Bakowska.

<ol>
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(60), e3747 (2012).</li><li>Nissim-Rafinia, M., Meshorer, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Photobleaching+assays+(FRAP+&amp;+FLIP)+to+measure+chromatin+protein+dynamics+in+living+embryonic+stem+cells.">Photobleaching assays (FRAP &amp; FLIP) to measure chromatin protein dynamics in living embryonic stem cells.</a> Journal of Visualized Experiments. (52), e2696 (2011).</li><li>Zheng, C., Petralia, R. S., Wang, Y., Kachar, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fluorescence+recovery+after+photobleaching+(FRAP)+of+fluorescence+tagged+proteins+in+dendritic+spines+of+cultured+hippocampal+neurons.">Fluorescence recovery after photobleaching (FRAP) of fluorescence tagged proteins in dendritic spines of cultured hippocampal neurons.</a> Journal of Visualized Experiments. 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C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=PB1+and+UBA+domains+of+p62+are+essential+for+aggresome-like+induced+structure+formation.">PB1 and UBA domains of p62 are essential for aggresome-like induced structure formation.</a> Biochemical and Biophysical Research Communications. 503 (4), 2306-2311 (2018).</li><li>Brown, E. B., Wu, E. S., Zipfel, W., Webb, W. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Measurement+of+molecular+diffusion+in+solution+by+multiphoton+fluorescence+photobleaching+recovery.">Measurement of molecular diffusion in solution by multiphoton fluorescence photobleaching recovery.</a> Biophysical Journal. 77 (5), 2837-2849 (1999).</li><li>Centonze, V., Pawley, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Tutorial+on+practical+confocal+microscopy+and+the+use+of+the+confocal+test+specimen.">Tutorial on practical confocal microscopy and the use of the confocal test specimen.</a> Handbook of biological confocal microscopy. , 549-570 (1995).</li><li>Chudakov, D. M., Matz, M. V., Lukyanov, S., Lukyanov, K. 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The authors have no conflict of interest to disclose.

We provide a comprehensive, practical, and simple step-by-step protocol for FRAP experiments with live cells. Herein, the protocol was used to measure the mobility of YFP-p62 in ALIS in RAW264.7 macrophages, but it can be applied to many of the laser scanning confocal microscope systems and genetically encoded fluorescent proteins that are now available. For any microscopy system, pilot experiments are critical for determining the optimal FRAP parameters, including acquisition, bleach, and control ROI sizes, laser intensity for photobleaching, and prebleach and postbleach image acquisition. It is reasonable to expect the optimal FRAP parameters to differ for each genetically encoded fluorescent protein and cell line.
Important factors to consider when conducting a FRAP experiment include (a) achieving suitable bleach depth, (b) the use of a brief bleaching step, (c) allowing sufficient time postbleach to observe the full recovery function, (d) photobleaching efficiency, (e) cytotoxicity with repeated FRAP, and (f) the inclusion of a control for fluorescence loss due to repeated imaging. We recommend that bleach depth, which can be calculated according to the equation provided in section 3.3.1, be &#8805;9013. When bleach depth is &#60;90, the degree of postbleach fluorescence recovery will be underestimated, and the values of If, Mf, and t1/2 will be incorrect. Although the duration and intensity of the bleach-inducing laser pulses may vary between FRAP experiments, it is important that the photobleaching step be brief and substantially faster than the fluorescence recovery function. If it is not, then a significant amount of fluorescence recovery could occur during the bleaching step. With a long bleach time, fluorescence recovery during the bleaching step would not be measured, and it would lead to incorrect measurements of If, Mf, and t1/2. In addition, to obtain correct values for If, Mf, and t1/2, the acquisition ROI should be observed postbleach until the fluorescence level in the bleach ROI has reached a plateau. For example, in our FRAP experiments, there was no difference between the If, Mf,&#160;and t1/2 values when we observed YFP-p62 fluorescence in the bleach ROI for 32.2 min postbleach versus when we observed YFP-p62 in the bleach ROI for 15.1 min postbleach; thus, we concluded that the recovery function reached a plateau at 15.1 min postbleach12. With regard to photobleaching efficiency, photobleaching increases with the square of the optical zoom factor14. Thus, the use of high optical zoom lenses is favorable for rapid photobleaching but can result in undesirable photobleaching during acquisition; the latter can be accounted for by imaging a control ROI. Repeated photobleaching is to be avoided as it can lead to the generation of cytotoxic reactive oxygen species (ROS). However, the degree of ROS generation due to exposure to a high-intensity laser is lower for genetically encoded fluorescent proteins than for chemical fluorophores (e.g., fluorescent antibodies)15, and the ROS generated are more likely to react within the genetically encoded fluorescent protein than with other molecules in the cell4. In addition to the increased probability of generating cytotoxic ROS, repeated photobleaching is to be avoided as it is difficult to control. Finally, although low laser transmission is used to acquire all non-bleach images, some photobleaching will invariably occur, which must be controlled. Possible controls for this include monitoring fluorescence in a control ROI within the acquisition ROI, obtaining control images in a neighboring unbleached cell, and performing control experiments with the identical settings for those used in the photobleaching experiments but without the photobleaching event.

Fluorescence recovery after photobleaching (FRAP) is a microscopy technique that can be used to quantify protein dynamics in live cells1,2. The popularity of FRAP has increased because of the widespread commercial availability of laser scanning confocal microscopes with high resolution, speed, and sensitivity, and a &#34;rainbow&#34; of genetically encoded fluorescent proteins, such as green fluorescent protein (GFP) and yellow fluorescent protein (YFP)3. Genetically encoded fluorescent proteins are fused to a protein of interest to allow for the subcellular localization of the protein of interest. In a typical FRAP experiment, the steady-state fluorescence in an acquisition region of interest (ROI) within a cell is observed via repeated imaging of that ROI with low-intensity laser light. Subsequently, the fluorescent molecules are rapidly and irreversibly impaired in a predefined subset of the acquisition ROI, hereafter referred to as the bleach ROI, by brief exposure to high-intensity laser light. As new unbleached proteins replenish bleached proteins over time, the speed and intensity of fluorescence recovery in the bleach ROI provides information about protein mobility (Figure 1)4.
Our interest in determining the mobility of the ubiquitin binding protein p62 (also known as sequestosome-1) in aggresome-like induced structures (ALIS) in murine RAW264.7 macrophages after stimulation with lipopolysaccharide (LPS) led us to review the FRAP literature. Unfortunately, many of the existing FRAP protocols are incomplete or inordinately complex5,6,7,8,9. Some do not provide detailed information about the laser settings, beam path configuration, and image acquisition parameters5,6,7,8,9. Others omitted key details regarding data analysis, such as how to address the issue of bleach ROI drift6,9 or how to calculate important recovery parameters, including the mobile fraction (Mf), immobile fraction (If), and half-time of recovery (t&#189;)5,7. Conversely, others placed too much emphasis on complex mathematical formulas used to calculate Mf, If, and t1/25,6,8,9. Thus, our purpose is to provide a comprehensive, practical, and straightforward step-by-step protocol for FRAP experiments with live cells.

<table border="0" cellpadding="0" cellspacing="0" style="width:835px;" width="835">
	<colgroup>
		<col />
		<col />
		<col />
		<col />
	</colgroup>
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:320px;">Dulbecco&#39;s Modified Eagle Medium</td>
			<td style="width:211px;">Hyclone</td>
			<td style="width:137px;">SH30022.01</td>
			<td style="width:167px;">High Glucose (4.5 g/L)</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:320px;">Fetal bovine serum</td>
			<td style="width:211px;">Gemini Bio-Products</td>
			<td style="width:137px;">100-106</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:320px;">Glass bottom dishes</td>
			<td style="width:211px;">MatTek</td>
			<td style="width:137px;">P35G-1.5-14-C</td>
			<td>35-mm</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:320px;">Image J software</td>
			<td style="width:211px;">National Institutes of Health</td>
			<td style="width:137px;">N.A.</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:320px;">Lipopolysaccharide (LPS)</td>
			<td style="width:211px;">Sigma</td>
			<td style="width:137px;">L4391</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:320px;">Nocodazole</td>
			<td style="width:211px;">Sigma</td>
			<td style="width:137px;">M1404</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:320px;">Penicillin-streptomycin solution</td>
			<td style="width:211px;">Corning</td>
			<td style="width:137px;">30-002-Cl</td>
			<td>100&#215;</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:320px;">Plan-Apochromat 63&#215;/1.40 oil objective</td>
			<td style="width:211px;">Carl Zeiss MicroImaging, Inc</td>
			<td style="width:137px;">4407629904000000</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:320px;">Polyethylenimine (PEI)</td>
			<td style="width:211px;">Polysciences</td>
			<td style="width:137px;">23966-1</td>
			<td>linear (MW 25,000)</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:320px;">RAW 264.7 murine macrophages</td>
			<td style="width:211px;">ATCC</td>
			<td style="width:137px;">TIB-71</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:320px;">Zeiss confocal microscope</td>
			<td style="width:211px;">Carl Zeiss MicroImaging, Inc</td>
			<td style="width:137px;">LSM 510</td>
			<td></td>
		</tr>
	</tbody>
</table>

fluorescence recovery after photobleaching of yellow fluorescent protein tagged p62 in aggresome-like induced structures

Fluorescence recovery after photobleaching (FRAP)&#160;is a microscopy technique that can be used to quantify protein mobility in live cells. In a typical FRAP experiment, steady-state fluorescence is observed by repeated imaging with low-intensity laser light. Subsequently, the fluorescent molecules are rapidly and irreversibly impaired via brief exposure to high-intensity laser light. Information about protein mobility is obtained by monitoring the recovery of fluorescence. We used FRAP to determine the mobility of p62 in aggresome-like induced structures (ALIS) in murine macrophages after stimulation with lipopolysaccharide (LPS). Because many existing FRAP protocols are either incomplete or complex, our goal was to provide a comprehensive, practical, and straightforward step-by-step protocol for FRAP experiments with live cells. Here, we describe RAW264.7 macrophage transfection with yellow fluorescent protein-p62 (YFP-p62), induction of ALIS by exposing the cells to LPS, and a step-by-step method for collecting prebleach and postbleach FRAP images and data analysis. Finally, we discuss important factors to consider when conducting a FRAP experiment.

Shown here are the results of a typical experiment in which we used FRAP to examine the degree of mobility of p62 in ALIS in RAW264.7 cells treated with LPS for 3-5 h12. Figure 3A shows the raw data obtained from a bleach, control, and background ROI after the stack of images had been aligned to correct for small amounts (&#60; ~3 &#181;m) of image drift of the ALIS. YFP-p62 fluorescence in this ALIS at prebleach and postbleach is shown in Figure 3E and Supplementary Video 1. Figure 3B shows these data after background-correction. Figure 3C shows these data after correcting for fluorescence in the control ROI. Figure 3D shows these data normalized to the mean fluorescence of the 5 prebleach values in the bleach ROI. The degree of bleaching was sufficient in this experiment (bleach depth = 91.94). YFP-p62 fluorescence within this ALIS recovered slowly, as t1/2 = 128.27 s (2.14 min). YFP-p62 was not a very mobile protein in this experiment, as Mf = 21.97 and If = 78.03.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/59288/59288fig1.jpg" /> 
Figure 1: A diagram of a RAW264.7 cell and steps for image analysis after the FRAP images have been aligned to correct for image drift of the ALIS. (A) Diagram of a RAW264.7 cell depicting several ALIS and the bleach, control, and background ROIs within the acquisition ROI. (B) An idealized depiction of raw FRAP data obtained in the bleach, control, and background ROIs in panel A. (C) An idealized graph of raw FRAP data that have been background-corrected. (D) An idealized graph of background-corrected FRAP data that have been normalized to fluorescence in the control ROI. (E) An idealized graph of normalized and background-corrected FRAP data that have been normalized to the prebleach fluorescence in the bleach ROI. Abbreviations: Con = control; BG = background. This figure was modified from Rabut and Ellenberg4. <a href="https://www.jove.com/files/ftp_upload/59288/59288fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/59288/59288fig2.jpg" /> 
Figure 2:&#160;Images of the user interface in the AIM software (Table of Materials) for laser selection, beam path configuration, and image acquisition parameters for the FRAP experiments. (A) Laser control screen showing the Argon/2 laser lines, output, and tube current used. (B) Configuration control screen showing the beam splitter and emission filter settings used. (C)&#160;Scan control screen showing the setting used for image acquisition.&#160;(D) Scan control screen showing the pinhole, detector gain, amplifier offset, amplifier gain, and transmission settings for the Argon/2 514 nm laser line. (E) Time series control screen showing the time series settings used. (F) Bleach control screen showing the prebleach and postbleach parameters used. Abbreviations: HFT = Haupt Farb Teiler (primary dichroic beam splitter); NT1 = Neben Farb Teiler 1 (first secondary dichroic beam splitter); NT2 = Neben Farb Teiler 2 (second secondary dichroic beam splitter); EF = emission filter. <a href="https://www.jove.com/files/ftp_upload/59288/59288fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/59288/59288fig3.jpg" /> 
Figure 3: A representative FRAP experiment to study YFP-p62 dynamics in ALIS in RAW264.7 cells. (A) Raw fluorescence intensity data for a bleach, control, and background ROI. (B) The data given in panel A after the bleach and control ROI had been corrected for fluorescence in the background ROI. (C) The data given in panels A and B after normalizing the fluorescence intensity in the bleach ROI to account for fluorescence in the background-corrected control ROI. (D) The data given in panels A-C after normalizing the fluorescence intensity in the normalized and background-corrected bleach ROI to the average of the first 5 prebleach values in the bleach ROI. Mf = 21.97, If = 78.03, t1/2 = 128.27 s (2.14 min), and bleach depth = 91.94. (E) An image of an ALIS at prebleach (1.5 min) and postbleach (0, 6, and 16 min). Scale bar = 5 &#181;m and is valid for all panels. Abbreviations: Con = control; BG = background; Corr. = corrected; Norm. = normalized; Avg. = average. This figure was modified from Cabe et al.12 <a href="https://www.jove.com/files/ftp_upload/59288/59288fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Supplementary Video 1" src="/files/ftp_upload/59288/59288supvideo1.jpg" /> 
Supplementary Video 1: A typical ALIS in a RAW264.7 macrophage before and after photobleaching. YFP-p62 fluorescence is slow to recover after the photobleach; hence, p62 is not a very mobile protein. For this experiment, Mf = 25.62, If = 74.38, t1/2 = 442.79 s (7.38 min), and bleach depth = 90.19. Scale bar = 5 &#181;m. This video was modified from Cabe et al.12 <a href="https://www.jove.com/files/ftp_upload/59288/Fig_4-Vid_FINAL.avi" target="_blank">Please click here to view this video. (Right-click to download.)</a>

Watch this Scientific Journal Video about Fluorescence Recovery after Photobleaching of Yellow Fluorescent Protein Tagged p62 in Aggresome-like Induced Structures at JoVE.com

Fluorescence Recovery after Photobleaching of Yellow Fluorescent Protein Tagged p62 in Aggresome-like Induced Structures

We describe a comprehensive and practical protocol for fluorescence recovery after photobleaching experiments with live cells. Although the protocol was used to measure the mobility of yellow fluorescent protein-tagged p62 in aggresome-like induced structures, it can be applied to a variety of microscopy systems and fluorescent proteins.

Fluorescence recovery after photobleaching (FRAP)&#160;is a microscopy technique that can be used to quantify protein mobility in live cells. In a typical ...

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14.7K Views.  Loyola University Chicago. We describe a comprehensive and practical protocol for fluorescence recovery after photobleaching experiments with live cells. Although the protocol was used to measure the mobility of yellow fluorescent protein-tagged p62 in aggresome-like induced structures, it can be applied to a variety of microscopy systems and fluorescent proteins.

Video: Fluorescence Recovery after Photobleaching of Yellow Fluorescent Protein Tagged p62 in Aggresome-like Induced Structures

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 ...

Measuring Diffusion Coefficients via Two-photon Fluorescence Recovery After Photobleaching

Multi-fluorescence recovery after photobleaching is a microscopy technique used to measure the diffusion coefficient (or analogous transport parameters) of macromolecules, and can be applied to both in vitro and in vivo biological systems. Multi-fluorescence recovery after photobleaching is performed by photobleaching a region of interest within a fluorescent sample using an intense laser flash, then attenuating the beam and monitoring the fluorescence as still-fluorescent molecules from outside the region of interest diffuse in to replace the photobleached molecules. We will begin our demonstration by aligning the laser beam through the Pockels Cell (laser modulator) and along the optical path through the laser scan box and objective lens to the sample. For simplicity, we will use a sample of aqueous fluorescent dye. We will then determine the proper experimental parameters for our sample including, monitor and bleaching powers, bleach duration, bin widths (for photon counting), and fluorescence recovery time. Next, we will describe the procedure for taking recovery curves, a process that can be largely automated via LabVIEW (National Instruments, Austin, TX) for enhanced throughput. Finally, the diffusion coefficient is determined by fitting the recovery data to the appropriate mathematical model using a least-squares fitting algorithm, readily programmable using software such as MATLAB (The Mathworks, Natick, MA).

In this article we will describe the procedure for measuring diffusion coefficients using multi-photon fluorescence recovery after photobleaching. We will begin by aligning the laser along the optical path to the sample and determining the proper experimental parameters, then continue generating and finally fitting fluorescence recovery curves.

Multi-fluorescence recovery after photobleaching is a microscopy technique used to measure the diffusion coefficient (or analogous transport parameters) ...

Using the GELFREE 8100 Fractionation System for Molecular Weight-Based Fractionation with Liquid Phase Recovery

The GELFREE 8100 Fractionation System is a novel protein fractionation system designed to maximize protein recovery during molecular weight based fractionation. The system is comprised of single-use, 8-sample capacity cartridges and a benchtop GELFREE Fractionation Instrument. During separation, a constant voltage is applied between the anode and cathode reservoirs, and each protein mixture is electrophoretically driven from a loading chamber into a specially designed gel column gel. Proteins are concentrated into a tight band in a stacking gel, and separated based on their respective electrophoretic mobilities in a resolving gel. As proteins elute from the column, they are trapped and concentrated in liquid phase in the collection chamber, free of the gel. The instrument is then paused at specific time intervals, and fractions are collected using a pipette. This process is repeated until all desired fractions have been collected. If fewer than 8 samples are run on a cartridge, any unused chambers can be used in subsequent separations.
This novel technology facilitates the quick and simple separation of up to 8 complex protein mixtures simultaneously, and offers several advantages when compared to previously available fractionation methods. This system is capable of fractionating up to 1mg of total protein per channel, for a total of 8mg per cartridge. Intact proteins over a broad mass range are separated on the basis of molecular weight, retaining important physiochemical properties of the analyte. The liquid phase entrapment provides for high recovery while eliminating the need for band or spot cutting, making the fractionation process highly reproducible1.

The accompanying video describes the use of the GELFREE 8100 Fractionation System, which partitions complex protein samples on the basis of molecular weight and recovers the fractions in liquid phase. The video describes how the technology works, how it is used, and provides resultant data, with polyacrylamide gel electrophoresis analysis of fractionated bovine liver homogenate.

The GELFREE 8100 Fractionation System is a novel protein fractionation system designed to maximize protein recovery during molecular weight based ...

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 ...

Photobleaching Assays (FRAP &amp; FLIP) to Measure Chromatin Protein Dynamics in Living Embryonic Stem Cells

Fluorescence Recovery After Photobleaching (FRAP) and Fluorescence Loss In Photobleaching (FLIP) enable the study of protein dynamics in living cells with good spatial and temporal resolution. Here we describe how to perform FRAP and FLIP assays of chromatin proteins, including H1 and HP1, in mouse embryonic stem (ES) cells. In a FRAP experiment, cells are transfected, either transiently or stably, with a protein of interest fused with the green fluorescent protein (GFP) or derivatives thereof (YFP, CFP, Cherry, etc.). In the transfected, fluorescing cells, an intense focused laser beam bleaches a relatively small region of interest (ROI). The laser wavelength is selected according to the fluorescent protein used for fusion. The laser light irreversibly bleaches the fluorescent signal of molecules in the ROI and, immediately following bleaching, the recovery of the fluorescent signal in the bleached area - mediated by the replacement of the bleached molecules with the unbleached molecules - is monitored using time lapse imaging. The generated fluorescence recovery curves provide information on the protein's mobility. If the fluorescent molecules are immobile, no fluorescence recovery will be observed. In a complementary approach, Fluorescence Loss in Photobleaching (FLIP), the laser beam bleaches the same spot repeatedly and the signal intensity is measured elsewhere in the fluorescing cell. FLIP experiments therefore measure signal decay rather than fluorescence recovery and are useful to determine protein mobility as well as protein shuttling between cellular compartments. Transient binding is a common property of chromatin-associated proteins. Although the major fraction of each chromatin protein is bound to chromatin at any given moment at steady state, the binding is transient and most chromatin proteins have a high turnover on chromatin, with a residence time in the order of seconds. These properties are crucial for generating high plasticity in genome expression1. Photobleaching experiments are therefore particularly useful to determine chromatin plasticity using GFP-fusion versions of chromatin structural proteins, especially in ES cells, where the dynamic exchange of chromatin proteins (including heterochromatin protein 1 (HP1), linker histone H1 and core histones) is higher than in differentiated cells2,3.

Photobleaching Assays FRAP &amp; FLIP to Measure Chromatin Protein Dynamics in Living Embryonic Stem Cells

We describe photobleaching methods including Fluorescence Recovery After Photobleaching (FRAP) and Fluorescence Loss In Photobleaching (FLIP) to monitor chromatin protein dynamics in embryonic stem (ES) cells. Chromatin protein dynamics, which is considered to be one of the means to study chromatin plasticity, is enhanced in pluripotent cells.

Fluorescence Recovery After Photobleaching (FRAP) and Fluorescence Loss In Photobleaching (FLIP) enable the study of protein dynamics in living cells with ...

Determination of Lipid Raft Partitioning of Fluorescently-tagged Probes in Living Cells by Fluorescence Correlation Spectroscopy (FCS)

In the past fifteen years the notion that cell membranes are not homogenous and rely on microdomains to exert their functions has become widely accepted. Lipid rafts are membrane microdomains enriched in cholesterol and sphingolipids. They play a role in cellular physiological processes such as signalling, and trafficking1,2 but are also thought to be key players in several diseases including viral or bacterial infections and neurodegenerative diseases3.
Yet their existence is still a matter of controversy4,5. Indeed, lipid raft size has been estimated to be around 20 nm6, far under the resolution limit of conventional microscopy (around 200 nm), thus precluding their direct imaging. Up to now, the main techniques used to assess the partition of proteins of interest inside lipid rafts were Detergent Resistant Membranes (DRMs) isolation and co-patching with antibodies. Though widely used because of their rather easy implementation, these techniques were prone to artefacts and thus criticized7,8. Technical improvements were therefore necessary to overcome these artefacts and to be able to probe lipid rafts partition in living cells.
Here we present a method for the sensitive analysis of lipid rafts partition of fluorescently-tagged proteins or lipids in the plasma membrane of living cells. This method, termed Fluorescence Correlation Spectroscopy (FCS), relies on the disparity in diffusion times of fluorescent probes located inside or outside of lipid rafts. In fact, as evidenced in both artificial membranes and cell cultures, probes would diffuse much faster outside than inside dense lipid rafts9,10. To determine diffusion times, minute fluorescence fluctuations are measured as a function of time in a focal volume (approximately 1 femtoliter), located at the plasma membrane of cells with a confocal microscope (Fig. 1). The auto-correlation curves can then be drawn from these fluctuations and fitted with appropriate mathematical diffusion models11.
FCS can be used to determine the lipid raft partitioning of various probes, as long as they are fluorescently tagged. Fluorescent tagging can be achieved by expression of fluorescent fusion proteins or by binding of fluorescent ligands. Moreover, FCS can be used not only in artificial membranes and cell lines but also in primary cultures, as described recently12. It can also be used to follow the dynamics of lipid raft partitioning after drug addition or membrane lipid composition change12.

Determination of Lipid Raft Partitioning of Fluorescently-tagged Probes in Living Cells by Fluorescence Correlation Spectroscopy FCS

A technique to probe the lipid raft partitioning of fluorescent proteins at the plasma membrane of living cells is described. It takes advantage of the disparity in diffusion times of proteins located inside or outside of lipid rafts. Acquisition can be performed dynamically in control conditions or after drug addition.

In the past fifteen years the notion that cell membranes are not homogenous and rely on microdomains to exert their functions has become widely accepted. ...

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, ...

Simultaneous Multicolor Imaging of Biological Structures with Fluorescence Photoactivation Localization Microscopy

Localization-based super resolution microscopy can be applied to obtain a spatial map (image) of the distribution of individual fluorescently labeled single molecules within a sample with a spatial resolution of tens of nanometers. Using either photoactivatable (PAFP) or&#160;photoswitchable (PSFP) fluorescent proteins fused to proteins of interest, or organic dyes conjugated to antibodies or other molecules of interest, fluorescence photoactivation localization microscopy (FPALM) can simultaneously image multiple species of molecules within single cells. By using the following approach, populations of large numbers (thousands to hundreds of thousands) of individual molecules are imaged in single cells and localized with a precision of ~10-30 nm. Data obtained can be applied to understanding the nanoscale spatial distributions of multiple protein types within a cell. One primary advantage of this technique is the dramatic increase in spatial resolution: while diffraction limits resolution to ~200-250 nm in conventional light microscopy, FPALM can image length scales more than an order of magnitude smaller. As many biological hypotheses concern the spatial relationships among different biomolecules, the improved resolution of FPALM can provide insight into questions of cellular organization which have previously been inaccessible to conventional fluorescence microscopy. In addition to detailing the methods for sample preparation and data acquisition, we here describe the optical setup for FPALM. One additional consideration for researchers wishing to do super-resolution microscopy is cost: in-house setups are significantly cheaper than most commercially available imaging machines. Limitations of this technique include the need for optimizing the labeling of molecules of interest within cell samples, and the need for post-processing software to visualize results. We here describe the use of PAFP and PSFP expression to image two protein species in fixed cells. Extension of the technique to living cells is also described.

We demonstrate the use of fluorescence photo activation localization microscopy (FPALM) to simultaneously image multiple types of fluorescently labeled molecules&#160;within cells. The techniques described yield the localization of thousands to hundreds of thousands of individual fluorescent labeled proteins, with a precision of tens of nanometers&#160;within single cells.

Localization-based super resolution microscopy can be applied to obtain a spatial map (image) of the distribution of individual fluorescently labeled ...

Protein-protein Interactions Visualized by Bimolecular Fluorescence Complementation in Tobacco Protoplasts and Leaves

Many proteins interact transiently with other proteins or are integrated into multi-protein complexes to perform their biological function. Bimolecular fluorescence complementation (BiFC) is an in vivo method to monitor such interactions in plant cells. In the presented protocol the investigated candidate proteins are fused to complementary halves of fluorescent proteins and the respective constructs are introduced into plant cells via agrobacterium-mediated transformation. Subsequently, the proteins are transiently expressed in tobacco leaves and the restored fluorescent signals can be detected with a confocal laser scanning microscope in the intact cells. This allows not only visualization of the interaction itself, but also the subcellular localization of the protein complexes can be determined. For this purpose, marker genes containing a fluorescent tag can be coexpressed along with the BiFC constructs, thus visualizing cellular structures such as the endoplasmic reticulum, mitochondria, the Golgi apparatus or the plasma membrane. The fluorescent signal can be monitored either directly in epidermal leaf cells or in single protoplasts, which can be easily isolated from the transformed tobacco leaves. BiFC is ideally suited to study protein-protein interactions in their natural surroundings within the living cell. However, it has to be considered that the expression has to be driven by strong promoters and that the interaction partners are modified due to fusion of the relatively large fluorescence tags, which might interfere with the interaction mechanism. Nevertheless, BiFC is an excellent complementary approach to other commonly applied methods investigating protein-protein interactions, such as coimmunoprecipitation, in vitro pull-down assays or yeast-two-hybrid experiments.

Formation of protein complexes in vivo can be visualized by bimolecular fluorescence complementation. Interaction partners are fused to complementary parts of fluorescent tags and transiently expressed in tobacco leaves, resulting in a reconstituted fluorescent signal upon close proximity of the two proteins.

Many proteins interact transiently with other proteins or are integrated into multi-protein complexes to perform their biological function. Bimolecular ...

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 ...