We would like to thank the Dorigo laboratory and the Neuroscience Imaging Service at Stanford University School of Medicine for providing equipment and space for this project.

1. Plasmid Construction
<ol>
	<li>Generate two-color fusion probes. Use a mammalian cell expression vector (see Table of Materials) in which mCherry112 and PA-mCherry113 have been inserted with the restriction sites AgeI and BsrGI.</li>
	<li>Order custom oligo-nucleotides to amplify the monomeric variants of eGFP and PA-eGFP containing the A206K mutation, i.e., mEGFP and PA-mEGFP14 without a stop codon as a SalI-BamHI fragment. Use the N-terminal primer 5&#8217;-AAT TAA CAG TCG ACG ATG GTG AGC AAG GGC GAG G 3&#8217; and the C-terminal primer 5&#8217;-AAT ATA TGG ATC CCG CTT GTA CAG CTC GTC CAT GC 3&#8217; and insert this SalI-BamHI fragment into the multiple cloning site of the expression vector. This will create the five amino acid linker RNPPV between the green and red fluorescent protein. We will refer to the fluorophore chimeras as GFP&#8212;Cherry, PA-GFP&#8212;Cherry, and GFP&#8212;PA-Cherry for the remainder of this article. 
	Note: The coupling of fluorophores and creation of internal rulers to assess photoactivation efficiency can be done with any spectrally distinct fluorophore pair, e.g. superfolder PA-GFP 15/ PA-TagRFP 16, etc. Many of the photoactivatable fluorescent proteins are derived from GFP. Hence, the primer sequence provided above can be used for many fluorescent proteins to construct a fluorophore chimera in a mammalian cell expression vector.</li>
</ol>
2. Cell Culture and Transfection
<ol>
	<li>Use either any standard cell line such as HeLa, NRK, Cos-7 or a specific cell line to be used for specific photoactivation experiments. Use DMEM (supplemented with 10% fetal bovine serum, and 2 mM glutamine) and trypsin without phenol red (see Table of Materials) to reduce background fluorescence.</li>
	<li>Detach the cells of a confluent culture with trypsin, count the number of cells in cell suspension using a Neubauer chamber, and seed 5,000 &#8211; 10,000 cells per well. Alternatively, use 1 drop from a 2-mL pipette of a 10-mL cell suspension from a confluent cell culture grown in a T 25-cell culture flask or 3 drops from a 5-mL cell suspension from a confluent culture grown in a T 12.5-cell culture flask.</li>
	<li>Grow cells in 8-well chambers with #1.0 cover glass (see Table of Materials) for fluorescence live-cell microscopy.</li>
	<li>Transfect cells 24 h after plating using commercial reagents (see Table of Materials) per distributer&#8217;s protocol with the GFP&#8212;Cherry, PA-GFP&#8212;Cherry, and GFP&#8212;PA-Cherry chimeras.</li>
	<li>Image cells after a total of 20 h post transfection to allow for protein expression, folding and maturation.</li>
</ol>
3. Imaging and Photoactivation
<ol>
	<li>Image cells in a humidified and heated environmental chamber at 37 degrees Celsius. To buffer the cell media at physiological pH and render it CO2-independent, add 20 mM HEPES, or use CO2 gas set to 5% flow.</li>
	<li>First, image cells expressing the GFP&#8212;Cherry construct. Set parameters that define time-integrated laser intensity per pixel in a confocal image, i.e., pixel dwell time in microseconds, acousto-optical tunable filter (AOTF) transmission in percent, and digital zoom. 
	Note: Using a 60x objective and a digital zoom of 3x allows imaging of a cell in its entirety while providing sufficient magnification. Set pixel dwell time to 2-4 &#956;s and AOTF transmission for the 488-nm and 561-nm laser such that images show a good signal-to-noise ratio without any bleaching and no pixels indicating fluorescence intensity saturation.</li>
	<li>Image, using the set laser power, AOTF transmission, pixel dwell time and digital zoom, 15-20 cells expressing GFP&#8212;Cherry.</li>
	<li>Then, image with the same set laser power, pixel dwell time, AOTF transmission and digital zoom cells expressing GFP&#8212;PA-Cherry and PA-GFP&#8212;Cherry. Search for expressing cells in the green channel or red channel, respectively. Avoid long exposure of the cells during the search for expressing cells in order to not bleach the fluorescent proteins.</li>
	<li>Set up a mini-time series with one pre-activation image and three post-activation images. The post-activation images will help identify potential transient dark states due to the exposure to UV-light. 
	Note: In our hands, changes in detected fluorescence intensity due to transient dark states were &#60;1% and could be neglected, but those dark states should be assessed for every experimental set-up.</li>
	<li>To determine photoactivation efficiency, i.e., the fraction of PA-FPs that is switched on to be fluorescent, in the specific photoactivation experiments that have already been established, apply the same photoactivation settings to 15-20 cells that are expressing the internal rulers introduced here and proceed with image analysis (section 4). If beginning to set up photoactivation experiments, find here a few different settings based on our experimental experience with PA-GFP and PA-Cherry; modify as needed.
	<ol>
		<li>For instantaneous photoactivation of PA-GFP and PA-Cherry using a confocal laser scanning microscope (CLSM), apply 90 &#956;W of 405-nm laser light in 3 or 5 iterations, respectively. 
		Note: With our microscopic set-up, using a 38% AOTM transmission and a 2 &#956;s pixel dwell time, we measured this 405-nm laser power in line-scan mode at the objective lens. With these settings, about 8% and 16% of the PA-GFP and PA-Cherry expressed can be photoactivated to be fluorescent. The entire titration series using CLSM for instantaneous photoactivation has been published previously17.</li>
		<li>If a higher fraction of photoactivated PA-GFP and PA-Cherry fluorophores is advantageous e.g. to achieve a higher signal-to-noise ratio, and photoactivation does not have to be immediate, apply 40 &#956;W of 405-nm laser light with a 2 &#956;s pixel dwell time and 6% AOTF transmission for 450 iterations. Then, photoactivation will take up to 4 min as opposed to only 1-2 seconds, but photoactivation efficiency for PA-GFP will be 29% instead of 8%, allowing for a higher signal-to-noise ratio.</li>
	</ol>
	</li>
	<li>If a mosaic digital illumination system that contains micro mirror arrays in a spatial light modulator is available, 405-nm laser light can be used for widefield-photoactivation. This allows for efficient photoactivation within milliseconds. 
	Note: With 1.6 mW laser power as measured at the objective lens and an exposure time of 250 ms, 29% of PA-GFP can be photoactivated to be fluorescent.</li>
	<li>Image with any set photoactivation parameters 15-20 cells expressing PA-GFP&#8212;Cherry and GFP&#8212;PA-Cherry, respectively.</li>
	<li>Since pH, reactive oxygen species and other environmental factors may influence photoactivation efficiency, it may be important to assess photoactivation efficiencies of the PA-FPs in the respective subcellular micromilieu where the protein of interest is located. By coupling the fluorophore chimeras to the protein of interest, as the authors have done with the plasma membrane protein VSVG 18, photoactivation efficiency can be assessed in the specific subcellular compartment of interest.</li>
</ol>
4. Image Analysis and Algorithm for Ratiometric Intensity-based Quantification of Photoactivation Efficiency
<ol>
	<li>Image analysis can be done with the open source image processing platforms ImageJ or Fiji. Determine the background fluorescence intensity in non-transfected cells in the green (B1) and red (B2) channel. Avoid perinuclear or any areas showing increased auto-fluorescence.</li>
	<li>To determine the fluorescence intensity in a transfected cell, outline the cell body with the freehand selections tool. Again, avoid perinuclear or any other areas showing auto-fluorescence.</li>
	<li>Subtract the background from the measured fluorescence intensity in each channel. 
	IG = IGreen_measured &#8211; B1 
	IR = IRed_measured &#8211; B2</li>
	<li>Use the GFP&#8212;Cherry construct to calculate the red-to-green ratio (RtoGr) and correct for donor-quenching due to fluorescence resonance energy transfer (FRET). The FRET efficiency E was determined to be 0.3 in previous experiments for the GFP&#8212;Cherry construct using the same amino acid linker between the two fluorophores18. 
	RtoGr = (IRed_measured &#8211; B2) / (IGreen_measured &#8211; B1) 
	RtoGrcorr = RtoGr * (1 &#8211; E) 
	Note: In this intensity- based approach, donor quenching for mEGFP and PA-mGFP may be different given possible distinct spectral properties which have not been characterized. The rate of FRET (kET) and the Foerster distance (R0) depend upon the quantum yield of the donor which has not been determined for mEGFP and PA-mGFP.</li>
	<li>Use the GFP&#8212;PA-Cherry construct to assess the fraction of photoactivated PA-Cherry. Determine the expected fluorescence intensity of PA-Cherry by multiplying the measured unquenched green fluorescence intensity of the GFP&#8212;PA-Cherry construct prior to photoactivation with the corrected red-to-green-ratio (RtoGrcorr). 
	IRed_expected = (IGreen_measured &#8211; B1) * RtoGrcorr 
	Note: As indicated above, the molecular brightness of the always-on FP and the photoactivatable FP may be different. See Discussion for more information on how to account for these differences.</li>
	<li>Calculate the PA-Cherry photoactivation efficiency as a fraction of the measured red fluorescence intensity after photoactivation and the expected fluorescence intensity in the red channel. 
	(FPA-Cherry) = (IRed_measured &#8211; B2) / IRed_expected</li>
	<li>Use the PA-GFP&#8212;Cherry construct to assess the fraction of photoactivated PA-GFP. Determine the expected fluorescence intensity of PA-GFP by dividing the measured red fluorescence intensity of the PA-GFP&#8212;Cherry construct prior to photoactivation by the red-to-green-ratio (RtoGr). Here, the RtoGr does not need to be corrected for donor quenching, because GFP and PA-GFP are subject to donor quenching to the same amount. 
	IGreen_expected = (IRed_measured &#8211; B2) / RtoGr</li>
	<li>Calculate the PA-GFP photoactivation efficiency as a fraction of the measured green fluorescence intensity after photoactivation and the expected fluorescence intensity in the green channel. 
	(FPA-GFP) = (IGreen_measured &#8211; B1) / IGreen_expected</li>
</ol>

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W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Ch.+9.">Ch. 9.</a> The Fluorescent Protein Revolution Series in Cellular and Clinical Imaging. , 201-228 (2014).</li><li>Shcherbakova, D. M., Sengupta, P., Lippincott-Schwartz, J., Verkhusha, V. 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The authors have nothing to disclose.

So far, no method existed to determine in bulk the fraction of PA-FPs expressed in live cells that is photoactivated to be fluorescent. The presented protocol can be used for any spectrally distinct fluorescent protein pair. While exemplified here for the irreversible PA-FPs PA-GFP and PA-Cherry, this approach is in principle applicable to photoconvertible proteins as well. The spectrally distinct fluorescent protein, however, must be selected carefully to minimize spectral overlap given that photoconvertible fluorescent...

In 2002, the first broadly applicable photoactivatable (PA-GFP1) and photoconvertible (Kaede2) fluorescent proteins were described. These optical highlighter fluorescent proteins change their spectral properties upon irradiation with UV-light, i.e., they become bright (photoactivatable fluorescent proteins, i.e., PA-FPs), or change their color (photoconvertible FPs). To date, several reversible and irreversible photoactivatable and photoconvertible fluorescent proteins have been developed3,4. In ensemble or bulk studies, optical highlighters have been used to study the dynamics of entire cells or proteins, and the connectivity of subcellular compartments. Furthermore, optical highlighters enabled single-molecule based superresolution imaging techniques such as PALM5 and FPALM6.
Although the photochemical processes during photoactivation or -conversion have been described for many optical highlighters and even crystallographic structures before and after photoactivation/ -conversion have been made available7,8, the underlying photophysical mechanism of photoactivation and -conversion is not completely understood. Furthermore, thus far only crude estimates exist of the efficiency of photoactivation and -conversion, that is, the fraction of fluorescent proteins expressed that is actually photoconverted or photoactivated to be fluorescent. In vitro ensemble studies have been reported quantifying the shift in absorption spectra and the amount of native and activated protein in a gel9,10,11.
Here, we present a protocol involving fluorescent protein chimeras to assess the fraction of photoactivated fluorescent proteins in bulk and in live cells. Whenever working with genetically encoded fluorescent proteins, the absolute amount of protein expressed varies from cell to cell and is unknown. If one cell expressing a PA-FP shows a brighter signal after photoactivation than another cell, it cannot be differentiated if this brighter signal is due to higher expression of the PA-FP or a more efficient photoactivation of the PA-FP. To standardize the expression level in cells, we introduce internal rulers of genetically coupled spectrally distinct fluorescent proteins. By coupling the genetic information of a photoactivatable fluorescent protein to a spectrally distinct always-on fluorescent proteins, internal rulers are created that will still be expressed to an unknown total amount but in a fixed and known relative amount of 1:1. This strategy allows the quantitative characterization of different UV-light photoactivation schemes, i.e., the assessment of the relative amount of PA-FPs that can be photoactivated with different modes of photoactivation, and thereby permits to define photoactivation schemes that are more effective than others. Furthermore, this strategy allows in principle the assessment of the absolute quantification of the photoactivated PA-FP fraction. To this end, it is important to realize that the presented ensemble studies are intensity-based which makes the analysis more complex as laid out in this protocol. Parameters determining the measured fluorescent intensity, i.e., different molecular brightness, absorbance and emission spectra and FRET effects, need to be considered when comparing fluorescence intensities of different fluorescent proteins.
The presented ratiometric intensity-based quantification of photoactivation efficiency is exemplified for PA-GFP and PA-Cherry in live cells, but is in principle broadly applicable and can be used for any photoactivatable fluorescent protein under any experimental condition.

<table><tbody><tr><td>pEGFP-N1 mammalian cell expression vector</td><td>Clontech</td><td></td><td></td></tr><tr><td>DMEM w/o phenol red</td><td>Thermo Fisher Scientific</td><td>11054020</td><td></td></tr><tr><td>Trypsin w/o phenol red</td><td>Thermo Fisher Scientific</td><td>15400054</td><td></td></tr><tr><td>L-Glutamine (200 mM)</td><td>Thermo Fisher Scientific</td><td>25030081</td><td></td></tr><tr><td>HEPES</td><td>Thermo Fisher Scientific</td><td>15630080</td><td></td></tr><tr><td>LabTek 8-well chambers #1.0</td><td>Thermo Fisher Scientific</td><td>12565470</td><td></td></tr><tr><td>Fugene 6</td><td>Promega</td><td>E2691</td><td></td></tr></tbody></table>

how to quantify the fraction of photoactivated fluorescent proteins in bulk and in live cells

Photoactivatable and -convertible fluorescent proteins (PA-FPs) have been used in fluorescence live-cell microscopy for analyzing the dynamics of cells and protein ensembles. Thus far, no method has been available to quantify in bulk and in live cells how many of the PA-FPs expressed are photoactivated to fluoresce.
Here, we present a protocol involving internal rulers, i.e., genetically coupled spectrally distinct (photoactivatable) fluorescent proteins, to ratiometrically quantify the fraction of all PA-FPs expressed in a cell that are switched on to be fluorescent. Using this protocol, we show that different modes of photoactivation yielded different photoactivation efficiencies. Short high-power photoactivation with a confocal laser scanning microscope (CLSM) resulted in up to four times lower photoactivation efficiency than hundreds of low-level exposures applied by CLSM or a short pulse applied by widefield illumination. While the protocol has been exemplified here for (PA-)GFP and (PA-)Cherry, it can in principle be applied to any spectrally distinct photoactivatable or photoconvertible fluorescent protein pair and any experimental set-up.

The protocol presented here shows the ratiometric quantification of the fraction of fluorescent proteins that are photoactivated to be fluorescent (Figure 1). This fraction differs depending upon the mode of photoactivation.
A typical result using short time high-power photoactivation with a confocal laser scanning microscope (CLSM) is shown in Figure 2c. After titratin...

Watch this Scientific Journal Video about How to Quantify the Fraction of Photoactivated Fluorescent Proteins in Bulk and in Live Cells at JoVE.com

How to Quantify the Fraction of Photoactivated Fluorescent Proteins in Bulk and in Live Cells

Here, we present a protocol that involves genetically coupled spectrally distinct photoactivatable and fluorescent proteins. These fluorescent protein chimeras permit quantification of the PA-FP fraction that is photoactivated to be fluorescent, i.e., the photoactivation efficiency. The protocol reveals that different modes of photoactivation yield different photoactivation efficiencies.

Photoactivatable and -convertible fluorescent proteins (PA-FPs) have been used in fluorescence live-cell microscopy for analyzing the dynamics of cells ...

how-to-quantify-fraction-photoactivated-fluorescent-proteins-bulk

Research

JoVE Journal

Biochemistry

6.6K Views.  Stanford University School of Medicine. We presenteren hier een protocol dat genetisch gekoppelde spectrally verschillende foto-geactiveerde en fluorescerende eiwitten omvat. Deze interne linialen maken kwantificering van de PAFP-fractie mogelijk die wordt geactiveerd om fluorescerend te zijn. Tot nu toe is er geen methode beschikbaar om in bulk in levenscellen te kwantificeren hoeveel van de uitgedrukte PAFP's zijn geactiveerd om te fluoresten. De gepresenteerde kwantificering van foto-activering efficiëntie wordt geïllustreerd ...

Video: How to Quantify the Fraction of Photoactivated Fluorescent Proteins in Bulk and in Live Cells

A Fluorescence Fluctuation Spectroscopy Assay of Protein-Protein Interactions at Cell-Cell Contacts

A variety of biological processes involves cell-cell interactions, typically mediated by proteins that interact at the interface between neighboring cells. Of interest, only few assays are capable of specifically probing such interactions directly in living cells. Here, we present an assay to measure the binding of proteins expressed at the surfaces of neighboring cells, at cell-cell contacts. This assay consists of two steps: mixing of cells expressing the proteins of interest fused to different fluorescent proteins, followed by fluorescence fluctuation spectroscopy measurements at cell-cell contacts using a confocal laser scanning microscope. We demonstrate the feasibility of this assay in a biologically relevant context by measuring the interactions of the amyloid precursor-like protein 1 (APLP1) across cell-cell junctions. We provide detailed protocols on the data acquisition using fluorescence-based techniques (scanning fluorescence cross-correlation spectroscopy, cross-correlation number and brightness analysis) and the required instrument calibrations. Further, we discuss critical steps in the data analysis and how to identify and correct external, spurious signal variations, such as those due to photobleaching or cell movement.
In general, the presented assay is applicable to any homo- or heterotypic protein-protein interaction at cell-cell contacts, between cells of the same or different types and can be implemented on a commercial confocal laser scanning microscope. An important requirement is the stability of the system, which needs to be sufficient to probe diffusive dynamics of the proteins of interest over several minutes.

This protocol describes a fluorescence fluctuation spectroscopy-based approach to investigate interactions among proteins mediating cell-cell interactions, i.e. proteins localized in cell junctions, directly in living cells. We provide detailed guidelines on instrument calibration, data acquisition and analysis, including corrections to possible artefact sources.

A variety of biological processes involves cell-cell interactions, typically mediated by proteins that interact at the interface between neighboring ...

Visualizing Protein-DNA Interactions in Live Bacterial Cells Using Photoactivated Single-molecule Tracking

Protein-DNA interactions are at the heart of many fundamental cellular processes. For example, DNA replication, transcription, repair, and chromosome organization are governed by DNA-binding proteins that recognize specific DNA structures or sequences. In vitro experiments have helped to generate detailed models for the function of many types of DNA-binding proteins, yet, the exact mechanisms of these processes and their organization in the complex environment of the living cell remain far less understood. We recently introduced a method for quantifying DNA-repair activities in live Escherichia coli cells using Photoactivated Localization Microscopy (PALM) combined with single-molecule tracking. Our general approach identifies individual DNA-binding events by the change in the mobility of a single protein upon association with the chromosome. The fraction of bound molecules provides a direct quantitative measure for the protein activity and abundance of substrates or binding sites at the single-cell level. Here, we describe the concept of the method and demonstrate sample preparation, data acquisition, and data analysis procedures.

Photoactivated localization microscopy (PALM) combined with single-molecule tracking allows direct observation and quantification of protein-DNA interactions in live Escherichia coli cells.

Protein-DNA interactions are at the heart of many fundamental cellular processes. For example, DNA replication, transcription, repair, and chromosome ...

Immunology and Infection

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 ...

Biology

Imaging Protein-protein Interactions in vivo

Protein-protein interactions are a hallmark of all essential cellular processes. However, many of these interactions are transient, or energetically weak, preventing their identification and analysis through traditional biochemical methods such as co-immunoprecipitation. In this regard, the genetically encodable fluorescent proteins (GFP, RFP, etc.) and their associated overlapping fluorescence spectrum have revolutionized our ability to monitor weak interactions in vivo using F&ouml;rster resonance energy transfer (FRET)1-3. Here, we detail our use of a FRET-based proximity assay for monitoring receptor-receptor interactions on the endothelial cell surface.

Imaging Protein-protein Interactions in vivo

This protocol describes how to image protein-protein interactions using a FRET-based proximity assay.

Protein-protein interactions are a hallmark of all essential cellular processes. However, many of these interactions are transient, or energetically weak, ...

In vivo Quantification of G Protein Coupled Receptor Interactions using Spectrally Resolved Two-photon Microscopy

The study of protein interactions in living cells is an important area of research because the information accumulated both benefits industrial applications as well as increases basic fundamental biological knowledge. F&ouml;rster (Fluorescence) Resonance Energy Transfer (FRET) between a donor molecule in an electronically excited state and a nearby acceptor molecule has been frequently utilized for studies of protein-protein interactions in living cells. The proteins of interest are tagged with two different types of fluorescent probes and expressed in biological cells. The fluorescent probes are then excited, typically using laser light, and the spectral properties of the fluorescence emission emanating from the fluorescent probes is collected and analyzed. Information regarding the degree of the protein interactions is embedded in the spectral emission data. Typically, the cell must be scanned a number of times in order to accumulate enough spectral information to accurately quantify the extent of the protein interactions for each region of interest within the cell. However, the molecular composition of these regions may change during the course of the acquisition process, limiting the spatial determination of the quantitative values of the apparent FRET efficiencies to an average over entire cells. By means of a spectrally resolved two-photon microscope, we are able to obtain a full set of spectrally resolved images after only one complete excitation scan of the sample of interest. From this pixel-level spectral data, a map of FRET efficiencies throughout the cell is calculated. By applying a simple theory of FRET in oligomeric complexes to the experimentally obtained distribution of FRET efficiencies throughout the cell, a single spectrally resolved scan reveals stoichiometric and structural information about the oligomer complex under study. Here we describe the procedure of preparing biological cells (the yeast Saccharomyces cerevisiae) expressing membrane receptors (sterile 2 &alpha;-factor receptors) tagged with two different types of fluorescent probes. Furthermore, we illustrate critical factors involved in collecting fluorescence data using the spectrally resolved two-photon microscopy imaging system. The use of this protocol may be extended to study any type of protein which can be expressed in a living cell with a fluorescent marker attached to it.

In vivo Quantification of G Protein Coupled Receptor Interactions using Spectrally Resolved Two-photon Microscopy

By employing a spectrally resolved two-photon microscopy imaging system, pixel-level maps of F&ouml;rster Resonance Energy Transfer (FRET) efficiencies are obtained for cells expressing membrane receptors hypothesized to form homo-oligomeric complexes. From the FRET efficiency maps, we are able to estimate stoichiometric information about the oligomer complex under study.

The study of protein interactions in living cells is an important area of research because the information accumulated both benefits industrial ...

Quantification of Efferocytosis by Single-cell Fluorescence Microscopy

Studying the regulation of efferocytosis requires methods that are able to accurately quantify the uptake of apoptotic cells and to probe the signaling and cellular processes that control efferocytosis. This quantification can be difficult to perform as apoptotic cells are often efferocytosed piecemeal, thus necessitating methods which can accurately delineate between the efferocytosed portion of an apoptotic target versus residual unengulfed cellular fragments. The approach outlined herein utilizes dual-labeling approaches to accurately quantify the dynamics of efferocytosis and efferocytic capacity of efferocytes such as macrophages. The cytosol of the apoptotic cell is labeled with a cell-tracking dye to enable monitoring of all apoptotic cell-derived materials, while surface biotinylation of the apoptotic cell allows for differentiation between internalized and non-internalized apoptotic cell fractions. The efferocytic capacity of efferocytes is determined by taking fluorescent images of live or fixed cells and quantifying the amount of bound versus internalized targets, as differentiated by streptavidin staining. This approach offers several advantages over methods such as flow cytometry, namely the accurate delineation of non-efferocytosed versus efferocytosed apoptotic cell fractions, the ability to measure efferocytic dynamics by live-cell microscopy, and the capacity to perform studies of cellular signaling in cells expressing fluorescently-labeled transgenes. Combined, the methods outlined in this protocol serve as the basis for a flexible experimental approach that can be used to accurately quantify efferocytic activity and interrogate cellular signaling pathways active during efferocytosis.

Efferocytosis, the phagocytic removal of apoptotic cells, is required to maintain homeostasis and is facilitated by receptors and signaling pathways that allow for the recognition, engulfment, and internalization of apoptotic cells. Herein, we present a fluorescence microscopy protocol for the quantification of efferocytosis and the activity of efferocytic signaling pathways.

Studying the regulation of efferocytosis requires methods that are able to accurately quantify the uptake of apoptotic cells and to probe the signaling ...

Temporal Quantification of MAPK Induced Expression in Single Yeast Cells

The quantification of gene expression at the single cell level uncovers novel regulatory mechanisms obscured in measurements performed at the population level. Two methods based on microscopy and flow cytometry are presented to demonstrate how such data can be acquired. The expression of a fluorescent reporter induced upon activation of the high osmolarity glycerol MAPK pathway in yeast is used as an example. The specific advantages of each method are highlighted. Flow cytometry measures a large number of cells (10,000) and provides a direct measure of the dynamics of protein expression independent of the slow maturation kinetics of the fluorescent protein. Imaging of living cells by microscopy is by contrast limited to the measurement of the matured form of the reporter in fewer cells. However, the data sets generated by this technique can be extremely rich thanks to the combinations of multiple reporters and to the spatial and temporal information obtained from individual cells. The combination of these two measurement methods can deliver new insights on the regulation of protein expression by signaling pathways.

Two complementary methods based on flow cytometry and microscopy are presented which enable the quantification, at the single cell level, of the dynamics of gene expression induced by the activation of a MAPK pathway in yeast.

The quantification of gene expression at the  single cell level uncovers novel regulatory mechanisms obscured in measurements  performed at the population ...

Visualization of Endoplasmic Reticulum Subdomains in Cultured Cells

The lipids and proteins in eukaryotic cells are continuously exchanged between cell compartments, although these retain their distinctive composition and functions despite the intense interorganelle molecular traffic. The techniques described in this paper are powerful means of studying protein and lipid mobility and trafficking in vivo and in their physiological environment. Fluorescence recovery after photobleaching (FRAP) and fluorescence loss in photobleaching (FLIP) are widely used live-cell imaging techniques for studying intracellular trafficking through the exo-endocytic pathway, the continuity between organelles or subcompartments, the formation of protein complexes, and protein localization in lipid microdomains, all of which can be observed under physiological and pathological conditions. The limitations of these approaches are mainly due to the use of fluorescent fusion proteins, and their potential drawbacks include artifactual over-expression in cells and the possibility of differences in the folding and localization of tagged and native proteins. Finally, as the limit of resolution of optical microscopy (about 200 nm) does not allow investigation of the fine structure of the ER or the specific subcompartments that can originate in cells under stress (i.e. hypoxia, drug administration, the over-expression of transmembrane ER resident proteins) or under pathological conditions, we combine live-cell imaging of cultured transfected cells with ultrastructural analyses based on transmission electron microscopy.

We describe the imaging approaches we use to investigate the distribution and mobility of the transfected fluorescent proteins resident in the endoplasmic reticulum (ER) by means of the confocal imaging of living cells. We also ultrastructurally analyze the effect of their expression on the architecture of this subcellular compartment.

The lipids and proteins in eukaryotic cells are continuously exchanged between cell compartments, although these retain their distinctive composition and ...

Internalization and Observation of Fluorescent Biomolecules in Living Microorganisms via Electroporation

The ability to study biomolecules in vivo is crucial for understanding their function in a biological context. One powerful approach involves fusing molecules of interest to fluorescent proteins such as GFP to study their expression, localization and function. However, GFP and its derivatives are significantly larger and less photostable than organic fluorophores generally used for in vitro experiments, and this can limit the scope of investigation. 
We recently introduced a straightforward, versatile and high-throughput method based on electroporation, allowing the internalization of biomolecules labeled with organic fluorophores into living microorganisms. Here we describe how to use electroporation to internalize labeled DNA fragments or proteins into Escherichia coli and Saccharomyces cerevisi&#230;, how to quantify the number of internalized molecules using fluorescence microscopy, and how to quantify the viability of electroporated cells. Data can be acquired at the single-cell or single-molecule level using fluorescence or FRET. The possibility of internalizing non-labeled molecules that trigger a physiological observable response in vivo is also presented. Finally, strategies of optimization of the protocol for specific biological systems are discussed.

Studies of biomolecules in vivo are crucial for understanding molecular function in a biological context. Here we describe a novel method allowing the internalization of fluorescent biomolecules, such as DNA or proteins, into living microorganisms. Analysis of in vivo data recorded by fluorescence microscopy is also presented and discussed.

The ability to study biomolecules in vivo is crucial for understanding their function in a biological context. One powerful approach involves fusing ...

Micromanipulation Techniques Allowing Analysis of Morphogenetic Dynamics and Turnover of Cytoskeletal Regulators

Examining the spatiotemporal dynamics of proteins can reveal their functional importance in various contexts. In this article, it is discussed how fluorescent recovery after photobleaching (FRAP) and photoactivation techniques can be used to study the spatiotemporal dynamics of proteins in subcellular locations. We also show how these techniques enable straightforward determination of various parameters linked to actin cytoskeletal regulation and cell motility. Moreover, the microinjection of cells is additionally described as an alternative treatment (potentially preceding or complementing the aforementioned photomanipulation techniques) to trigger instantaneous effects of translocated proteins on cell morphology and function. Micromanipulation such as protein injection or local application of plasma membrane-permeable drugs or cytoskeletal inhibitors can serve as powerful tool to record immediate consequences of a given treatment on cell behavior at the single cell and subcellular level. This is exemplified here by immediate induction of lamellipodial cell edge protrusion by the injection of recombinant Rac1 protein, as established a quarter-century ago. In addition, we provide a protocol for determining the turnover of enhanced green fluorescent protein (EGFP)-VASP, an actin filament polymerase prominently accumulating at lamellipodial tips of B16-F1 cells, employing FRAP and including associated data analysis and curve fitting. We also present guidelines for estimating the rates of lamellipodial actin network polymerization, as exemplified by cells expressing EGFP-tagged &#946;-actin. Finally, instructions are given for how to investigate the rates of actin monomer mobility within the cell cytoplasm, followed by actin incorporation at sites of rapid filament assembly, such as the tips of protruding lamellipodia, using photoactivation approaches. None of these protocols is&#160;restricted to components or regulators of the actin cytoskeleton, but can easily be extended to explore in analogous fashion the spatiotemporal dynamics and function of proteins in various different subcellular structures or functional contexts.

We describe how micro- and photomanipulation techniques such as FRAP and photoactivation enable the determination of motility parameters and the spatiotemporal dynamics of proteins within migrating cells. Experimental readouts include subcellular dynamics and turnover of motility regulators or of the underlying actin cytoskeleton.

How to Quantify the Fraction of Photoactivated Fluorescent Proteins in Bulk and in Live Cells

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