This work was supported by the Federal Ministry of Education and Research Germany (FKZ 0312138A and FKZ 316159), the State Mecklenburg-Western Pomerania with EU Structural Funds (ESF/IVWM-B34-0030/10 and ESF/IVBM-B35-0010/12), the DFG (DA1296-1), and the German Heart Foundation (F/01/12). In addition, R.D. is supported by the FORUN Program of Rostock University Medical Centre (889001), the DAMP Foundation, and the BMBF (VIP+ 00240).

All steps in this protocol involving neonatal mice were performed per the ethical guidelines for animal care of the Rostock University Medical Centre.
1. Preparation of Cell Culture Dishes and the Medium for Cardiomyocyte Culture
<ol>
	<li>Coat a cell culture plate with 0.1% gelatin in PBS and incubate at 37 &#176;C for 4 h or at 4 &#176;C overnight. Remove the gelatin and allow it to dry under sterile laminar air flow.</li>
	<li>Prepare cell culture medium composed of 50 mL of DMEM supplemented with 10% fetal bovine serum (FBS) and 1% penicillin/streptomycin (P/S). Pre-warm it to 37 &#176;C.</li>
</ol>
2. Isolation of Neonatal Cardiomyocytes
<ol>
	<li>Sacrifice neonatal mice (1-2 days old) by decapitation with sterile scissors and open the chest along the sternum. Remove the heart using forceps while slightly pressing the thorax together and transfer the heart into a 24-well plate containing ice-cold HBSS (without Ca2+ and Mg2+).</li>
	<li>Remove non-cardiac tissue and larger vessels using sterile forceps. Transfer cleaned hearts into a 1.5 mL tube containing ice-cold HBSS. Use a maximum of 5 hearts per tube for enzymatic digestion.</li>
	<li>Mince the hearts with small scissors into &#60;0.5 to 1 mm&#179; pieces.</li>
	<li>Wash the minced hearts with HBSS two times by aspiration/addition of HBSS using a 1-mL microliter pipette. Remove the HBSS completely before adding the enzymes.</li>
	<li>For the enzymatic digestion of neonatal hearts, use a commercially available kit and follow the manufacturer&#39;s protocol (see the Table of Materials). Shake the tube containing the minced hearts every 5 min while incubating with enzymes at 37 &#176;C for 35 min.</li>
	<li>Seed suspended cells on non-coated cell culture dishes containing cell culture medium for 1.5-2 h to allow the adherence of the non-cardiomyocyte fraction. Repeat this step if a high purity of cardiomyocytes is needed. If desired, further culture the non-cardiomyocyte fraction. 
	NOTE: For a cell suspension of 15 hearts, the pre-plating step can be performed on 75 cm&#178; cell culture flasks. Usually, ~5 x 105 cells are obtained from one neonatal heart.</li>
	<li>Collect the supernatant in a 15 mL conical tube and centrifuge for 10 min at 300 x g. Resuspend the cells in 1 mL of cell culture medium.</li>
	<li>Count the cells with a Neubauer chamber hemocytometer, or use an equivalent method.</li>
	<li>Plate isolated cardiomyocytes on 6-well plates with a density of 3 x 105 cells/cm&#178;. Incubate the cells overnight at 37 &#176;C and 5% CO2. 
	NOTE: Optionally, cells can also be transfected at this point. However, increased viability was observed when the cells were cultured for one day before being subjected to electroporation.</li>
</ol>
3. Transfection with Fluorescently Labeled miRNA
<ol>
	<li>Pre-warm cell culture medium (see step 1.2) to 37 &#176;C in a water bath or an equivalent device.</li>
	<li>Prepare a 20 &#181;M stock solution of fluorescent miRNA in RNase-free sterile water.</li>
	<li>Prepare electroporation buffer containing 90 mM Na2HPO4, 90 mM NaH2PO4, 5 mM KCl, 10 mM MgCl2, and 10 mM sodium succinate and adjust the pH to 7.2; electroporation buffer can be stored at -20 &#176;C for several months.</li>
	<li>Detach the cells from the culture dish using 0.05% trypsin for 5 min. Inactivate the trypsin by adding cell culture medium.</li>
	<li>Count the cells with a Neubauer chamber hemocytometer, or use an equivalent method. Centrifuge the cells at 300 x g for 10 min.</li>
	<li>Resuspend the cells in electroporation buffer to obtain a concentration of 4 x 105 cells per 100 &#181;L.</li>
	<li>Mix 100 &#181;L of cell suspension with fluorescent miRNA (final concentration: 0.25 &#181;M) in a tube and load the mixture into an electroporation cuvette.
	<ol>
		<li>Empirically determine the appropriate amount of miRNA, depending upon cell type.</li>
	</ol>
	</li>
	<li>Perform electroporation using an electroporation device (see the Table of Materials), using the &#34;G-009&#34; program.</li>
	<li>Add 500 &#181;L of pre-warmed cell culture medium and transfer the whole cell suspension (4 x 105 cells) into a well of a 4-well glass-bottom chamber slide. Culture the cells for 1 day at 37 &#176;C in a 5% CO2 atmosphere. 
	NOTE: For FRAP analysis, a cell density of ~80% is optimal. The transfection of labeled miRNA should exclusively be performed using electroporation. Since the homogenous distribution of labeled miRNA molecules is beneficial for FRAP measurement, reagent-based transfection is not recommended.</li>
</ol>
4. Applying 3D-FRAP Microscopy (Day 3)
<ol>
	<li>Warm up the microscope incubator to 37 &#176;C and switch on the confocal microscope system at least 2 h before FRAP measurement to establish a thermal equilibrium and to decrease the possibility of drift. If possible, maintain a 5% CO2 atmosphere.</li>
	<li>Insert the chamber slide into the stage sample holder.</li>
	<li>Find a cluster of transfected cardiomyocytes using a 1.4 N.A. oil objective (400X magnification) and 561-nm laser excitation light at low laser power, with a detection range of 570-680 nm. 
	NOTE: The definition of FRAP settings, the image acquisition, and the analysis were done using microscope-specific software (see the Table of Materials).</li>
	<li>Activate the &#34;z-Stack,&#34; &#34;Time Series,&#34; &#34;Bleaching,&#34; and &#34;Regions&#34; buttons in the &#34;Setup Manager.&#34;</li>
	<li>Define the FRAP parameters.
	<ol>
		<li>Define the image acquisition settings in the &#34;Acquisition Mode&#34; menu by setting the &#34;frame size&#34; to 512 x 512, the &#34;line step&#34; to 1, and the &#34;scan time&#34; to &#60;1 s.</li>
		<li>In the &#34;Channels&#34; menu, adjust the laser power, offset, and gain settings to obtain maximal fluorescence from minimal laser excitation (e.g., laser power: 1-5%); adjust to avoid intensity saturation. Set the &#34;pinhole size&#34; to 2 &#181;m.</li>
		<li>Next, in the &#34;Regions&#34; menu, select the &#34;ROI drawing tool&#34; and use the cursor to mark the target cell, the reference cell, and the background area. If required, select several target cells for photobleaching.</li>
		<li>Adjust the photobleaching settings in the &#34;Bleaching&#34; menu (e.g., iterations: 9-14, laser power for photobleaching: 100%, interval of image acquisition: 60 s, cycles: 15). Define the start of bleaching after 3 initial scans.</li>
		<li>In the &#34;z-Stack&#34; menu, define the limits for z-stack acquisition, depending on the thickness of the cells. Adjust the &#34;number of z-layers&#34; to 12 - 15.</li>
		<li>Start the FRAP experiment and record the fluorescence recovery. 
		NOTE: Since the settings of a FRAP experiment depend upon the cell type, the fluorescent dye, and the microscope system, it is best to perform pilot experiments to determine the optimal parameters for FRAP. Bleaching should be sufficient to reduce the initial fluorescence intensity by at least 50%. Typically, fluorescence recovery should be recorded every 30-60 s until the plateau phase is reached.</li>
	</ol>
	</li>
</ol>
5. Data Analysis
<ol>
	<li>Create maximum projections of the acquired z-stacks and obtain fluorescence intensity values of the bleached target cells, the reference cell, and the background. Using microscope-specific software, click &#34;Processing&#34;&#160;&#8594; &#34;Maximum intensity projection&#34; &#8594; select file &#8594; &#34;Apply.&#34;
	<ol>
		<li>Alternatively, use an equivalent image analyzing tool (e.g., ImageJ).</li>
	</ol>
	</li>
	<li>Copy the fluorescence intensity data at all time points from the target cell, the background, and the reference cell into a spreadsheet.
	<ol>
		<li>Subtract the background intensity and the intensity of the reference cell from the intensity values of the target cell to correct for photobleaching caused during the acquisition process. Perform background and reference cell corrections for each time point.</li>
		<li>Normalize the corrected FRAP data to the initial fluorescence intensity before bleaching by dividing each value by the initial fluorescence.</li>
		<li>To obtain FRAP curves, set the baseline to the fluorescence intensity immediately after the bleach by subtracting from the intensity values of each time point.</li>
	</ol>
	</li>
</ol>

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The authors declare no conflict of interest.

miRNAs are key players in cellular physiology and were shown to act as signaling molecules by using&#8212;among others&#8212;GJs as a pathway for intercellular exchange11,12,22. The current protocol presents an in vitro live-cell imaging technique to characterize this GJ-dependent shuttling using fluorescent miRNAs within cell clusters.
The protocol was developed on cardiomyocytes as a cell model system. However, this approach can be applied to several cell types if electroporation is a suitable method for miRNA transfection. In addition to cell-to-cell exchange, the presented technique is also suitable to investigate the intracellular mobility of miRNA molecules (e.g., for the identification of possible transport mechanisms within the cell)23,24.
The investigation of miRNA transfer by FRAP microscopy requires fluorescently tagged miRNA molecules (Figure 2). As the labeling of specific cellular miRNAs is not feasible, only exogenous, introduced labeled miRNA can be used for 3D-FRAP to analyze its intercellular mobility. In our experiments, cells were transfected with 250-nM miRNA. This concentration was much higher compared to physiological conditions, where the miRNA level ranges from 10 to 50,000 molecules per cell27,28. Such low concentrations cannot be used in FRAP experiments, as a certain level of fluorescent miRNA molecules is required to obtain a sufficient fluorescence signal and to ensure the proper discrimination between background fluorescence and fluorescently labeled miRNA. However, depending upon the confocal imaging system and the dye properties, the use of miRNA concentrations at low-nM range should be feasible for FRAP assays.
Different techniques are commonly applied to study intercellular miRNA transfer, including flow cytometry, PCR, and luciferase reporter assays12,13. These methods detect miRNA shuttling with low temporal resolution (i.e., hours to days). In contrast, 3D-FRAP microscopy allows for the visualization and quantification of miRNA transfer and mobility in real time. In addition to gap junctional cell-cell contacts, the intercellular shuttling of miRNA is also mediated by other mechanisms, such as exosomes18,29,30. Only 3D-FRAP provides sufficiently high temporal resolution to discriminate between exosomal and gap junctional transfer. Thus, it allows for specific investigation into the direct effect of GJ permeability on miRNA signaling under different pathological or physiological conditions (Figure 2).
We recommend the use of 3D-FRAP, which is superior to conventional 2D-FRAP, as it provides more accurate data on GJ-dependent miRNA transfer. 3D-FRAP measurements include the entire cell volume, which is important when using large-bulk cell types. In addition, it can compensate focus changes or cell movements, which were shown to significantly impair fluorescence recovery in 2D-FRAP (Figure 3). Finally, the recording of z-stacks in 3D-FRAP experiments provides the possibility to obtain spatial information, which is not feasible when just a single 2D layer is used for the analysis of fluorescence intensities (Figure 3B).
Since fluorescence recovery is dependent upon the influx of miRNA from adjacent cells, the number of cells connected to the bleached cell is critical and must be similar across different measurements to obtain reliable data. Therefore, a high cell density is required to ensure the formation of cellular clusters most suitable for FRAP analysis. Furthermore, preliminary experiments must be performed to select mild bleaching conditions (i.e., laser power, bleaching time, and scan settings) to avoid phototoxic effects caused by a high laser intensity25,26. This will also help to reduce the photobleaching that occurs during the acquisition process.
Despite the limitations, our data show that 3D-FRAP is a powerful tool to evaluate the role of miRNAs as signaling molecules within a cellular network. The impact of connexin composition on miRNA shuttling or the selective permeability of GJs for specific miRNAs can be addressed with the direct quantification of transport efficiency. 3D-FRAP can provide important information to improve the development of new therapies featuring miRNA and siRNA, such as by defining proper conditions that facilitate the distribution of small RNAs via the GJ network31,32.

Small noncoding RNAs are important players in cellular gene regulation. These molecules are composed of 20-25 nucleotides that bind to a specific target mRNA, leading to the blockage of translation or to mRNA degradation1,2. The gene regulatory process undertaken by small RNAs, such as miRNA and siRNA, is a highly conserved mechanism that has been found in many different species3. In particular, miRNA molecules are of crucial importance to a variety of physiological processes, including proliferation, differentiation, and regeneration4,5. In addition, the dysregulation of miRNA expression is attributed to many pathological disorders. Correspondingly, miRNAs have been demonstrated to be suitable as biomarkers for diagnosis and as therapeutic agents for gene therapy6,7.
Gap junctions (GJs) are specialized protein structures in the plasma membrane of two adjacent cells that allow the diffusional exchange of molecules with a molecular weight of up to 1 kD. They have been shown to be important to tissue development, differentiation, cell death, and pathological disorders such as cancer or cardiovascular disease8,9,10. Several molecules have been described as being capable of crossing GJ channels, including ions, metabolites, and nucleotides. Interestingly, GJs were also found to provide a pathway for the intercellular movement of small RNAs11,12. Thus, miRNAs can act not only within the cell in which they are produced, but also within recipient cells. This highlights the role of miRNAs in the intercellular signal transduction system. At the same time, the data demonstrate that gap junctional intercellular communication is closely linked to miRNA function. Because of the significant impact of miRNA and GJs on tissue homeostasis, pathology, and diagnosis, a comprehensive understanding of the function of GJs and the related intercellular dynamics of miRNA will help to clarify the mechanisms of miRNA-based diseases and to develop new strategies for miRNA therapies.
Depending on the extent of gap junctional coupling, the transfer of miRNA molecules between cells can be a very rapid process. Therefore, a methodology that allows the visualization and quantification of the fast intercellular movement of these regulatory signaling molecules is required. Commonly, flow cytometry and molecular biological techniques have been applied to demonstrate the shuttling of small RNAs11,12,13,14. However, as opposed to FRAP microscopy, these approaches lack high temporal resolution, which is mandatory when analyzing the exchange of miRNA via GJs. Moreover, FRAP microscopy is less invasive and therefore represents a powerful and novel live-cell imaging technique to evaluate the GJ-dependent exchange of molecules in several cell types15,16,17.
Here, we present a detailed protocol describing the application of 3D-FRAP to assess miRNA shuttling between cardiomyocytes. For this purpose, cardiomyocytes were transfected with fluorescently labeled miRNA. A cell marked with this miRNA was photobleached, and the gap junctional miRNA re-influx from adjacent cells was recorded in a time-dependent manner. The high temporal resolution of FRAP experiments offers the possibility to perform kinetic studies for the precise evaluation of the intercellular transfer of miRNA and siRNA between living cells. Moreover, as small RNAs can be exchanged via different mechanisms with highly different kinetics, FRAP microscopy can help to clarify the extent to which GJs are involved in the respective shuttling processes18. In addition, 3D-FRAP can be used to investigate physiological and pathological alterations in GJ permeability and its impact on small RNA transfer15,19.

<table><tbody><tr><td>neonatal NMRI mice</td><td>Charles River</td><td></td><td></td></tr><tr><td>Gelatin</td><td>Sigma Aldrich</td><td>G7041</td><td>0.1% solution in PBS, sterilized</td></tr><tr><td>PBS</td><td>Pan Biotech</td><td>P04-53500</td><td></td></tr><tr><td>Dulbecco&amp;acute;s modified medium</td><td>Pan Biotech</td><td>P04-03550</td><td></td></tr><tr><td>Penicillin/Streptomycin</td><td>Thermo Fisher Scientific</td><td>15140122</td><td>100 U/mL, 100&amp;micro;g/mL</td></tr><tr><td>Fetal bovine serum</td><td>Pan Biotech</td><td>P30-3306</td><td></td></tr><tr><td>Cell culture plastic</td><td>TPP</td><td></td><td></td></tr><tr><td>Primary Cardiomyocyte Isolation Kit</td><td>Thermo Fisher Scientific</td><td>88281</td><td></td></tr><tr><td>HBSS</td><td>Thermo Fisher Scientific</td><td>88281</td><td>included in primary cardiomyocyte isolation kit</td></tr><tr><td>Trypan blue</td><td>Thermo Fisher Scientific</td><td>15250061</td><td></td></tr><tr><td>miRIDIAN Dy547 labeled microRNA Mimic</td><td>Dharmacon</td><td>CP-004500-01-05</td><td>dissolve in RNAse free water, stock solution 20&amp;micro;M</td></tr><tr><td>Sodium phosphate monobasic</td><td>Sigma</td><td>S3139</td><td></td></tr><tr><td>Sodium phosphate dibasic</td><td>Carl Roth</td><td>T877.2</td><td></td></tr><tr><td>Potassium chloride</td><td>Sigma</td><td>P9333</td><td></td></tr><tr><td>Magnesium chloride</td><td>Serva</td><td>28305.01</td><td></td></tr><tr><td>Sodium succinate</td><td>Carl Roth</td><td>3195.1</td><td></td></tr><tr><td>0.05% Trypsin/EDTA solution</td><td>Merck</td><td>L2153</td><td></td></tr><tr><td>4-well- Glass bottom chamber slides</td><td>IBIDI</td><td>80827</td><td>coat with 0.1% gelatin solution</td></tr><tr><td>Amaxa Nucleofector II</td><td>Lonza</td><td></td><td>program G-009 was used for Electroporation&amp;nbsp;</td></tr><tr><td>LSM 780 ELYRA PS.1 system</td><td>Zeiss</td><td></td><td></td></tr><tr><td>Excel software</td><td>Microsoft</td><td></td><td></td></tr><tr><td>&amp;alpha;-actinin antibody</td><td>Abcam</td><td>ab9465</td><td>dilution 1:200</td></tr><tr><td>Connexin43 antibody</td><td>Santa Cruz</td><td>sc-9059</td><td>dilution 1:200</td></tr><tr><td>goat anti-mouse Alexa 594 antibody</td><td>Thermo Fisher Scientific</td><td>A-11005</td><td>dilution 1:300</td></tr><tr><td>goat anti-rabbit Alexa 488 antibody</td><td>Thermo Fisher Scientific</td><td>A-11034</td><td>dilution 1:300</td></tr><tr><td>Connexin43 siRNA</td><td>Thermo Fisher Scientific</td><td>AM16708 ID158724</td><td>final concentration 250 nM</td></tr><tr><td>Near-IR Live/Dead Cell Stain Kit</td><td>Thermo Fisher Scientific</td><td>L10119</td><td></td></tr><tr><td>CellTrace Calcein Red-Orange</td><td>Thermo Scientific</td><td>C34851</td><td></td></tr><tr><td>DAPI nuclear stain</td><td>Thermo Scientific</td><td>D1306</td><td></td></tr></tbody></table>

analysis of the gap junction-dependent transfer of mirna with 3d-frap microscopy

Small antisense RNAs, like miRNA and siRNA, play an important role in cellular physiology and pathology and, moreover, can be used as therapeutic agents in the treatment of several diseases. The development of new, innovative strategies for miRNA/siRNA therapy is based on an extensive knowledge of the underlying mechanisms. Recent data suggest that small RNAs are exchanged between cells in a gap junction-dependent manner, thereby inducing gene regulatory effects in the recipient cell. Molecular biological techniques and flow cytometric analysis are commonly used to study the intercellular exchange of miRNA. However, these methods do not provide high temporal resolution, which is necessary when studying the gap junctional flux of molecules. Therefore, to investigate the impact of miRNA/siRNA as intercellular signaling molecules, novel tools are needed that will allow for the analysis of these small RNAs at the cellular level. The present protocol describes the application of three-dimensional fluorescence recovery after photobleaching (3D-FRAP) microscopy to elucidating the gap junction-dependent exchange of miRNA molecules between cardiac cells. Importantly, this straightforward and non-invasive live-cell imaging approach allows for the visualization and quantification of the gap junctional shuttling of fluorescently labeled small RNAs in real time, with high spatio-temporal resolution. The data obtained by 3D-FRAP confirm a novel pathway of intercellular gene regulation, where small RNAs act as signaling molecules within the intercellular network.

Here, we present 3D-FRAP microscopy as a non-invasive technique to study the gap junctional shuttling of fluorescent miRNA within neonatal cardiomyocytes. The isolated cardiomyocytes revealed the typical striated &#945;-actinin pattern and contained large plaques of Cx43 along the cell-cell borders (Figure 1A, white arrowheads), which allowed for the high intercellular flux of molecules. The purity of isolated cardiomyocytes was assessed by the microscopic quantification of &#945;-actinin-positive cells. Although some non-cardiomyocytes (&#945;-actinin-negative cells) were present in the culture, neonatal cardiomyocytes represented the major cell type after isolation (79.62 &#177; 3.68%; Figure 1B).
To investigate the gap junctional exchange of miRNA, cells were transfected with fluorescently labeled miRNA. Electroporation is the method of choice to deliver miRNA molecules to cells, as it ensures a high transfection efficiency and the homogenous distribution of transfected compounds, which is mandatory for FRAP analysis. In our experimental setup, we achieved a transfection efficiency of ~ 45%, as determined by flow cytometry (Figure 2A). The preparation of cardiomyocytes for FRAP includes detachment from the culture plate, electroporation, and re-attachment onto chamber slides. A flow cytometric live/dead assay revealed that the majority of cells demonstrated high viability, which is important when analyzing gap junctional communication (Figure 2B). Moreover, cell density is a crucial parameter that affects FRAP results. For optimal FRAP measurements, cells should be seeded with a density of ~80% to allow for the formation of cell clusters and the establishment of functional gap junctions between cells (Figure 2C).
For FRAP analysis, target cells are selected within a cell cluster and are photobleached with 100% laser power, leading to reduced miRNA fluorescence in the selected cells (Figure 3A, bleach). Subsequently, fluorescence recovery is recorded to visualize the transfer of miRNA from adjacent cells into the bleached area. Since the plateau phase of the fluorescence recovery was reached after 13 min, this time span was used for image acquisition in the FRAP experiments. Quantitative analysis and normalization of the acquired data showed an average recovery of 20%. The data also demonstrated that FRAP microscopy allows for the rapid recording of miRNA shuttling with high temporal resolution. Depending on the dye properties, acquisition parameters can be changed to further increase temporal resolution.
The use of 3D-FRAP enables the comparison of GJ permeability to miRNA under different conditions. To demonstrate this, we induced an siRNA-mediated knockdown of Cx43, leading to reduced protein expression (Figure 3C) and the inhibition of gap junctional communication. As a result, fluorescence recovery was reduced by more than 50%, indicating that the efficiency of miRNA transfer is strongly dependent upon the extent of gap junctional coupling between cells (Figure 3B, Cx43 knockdown). To confirm that the fluorescence recovery reflects the passage of the fluorescently labeled miRNA, rather than the detached Dy547-tag, we also acquired FRAP data for calcein dye (Figure 3D). Calcein has a molecular weight comparable to that of Dy547 and thus possess similar transfer dynamics. In contrast to the fluorescence recovery of Dy547-labeled miRNA, calcein demonstrated increased gap junctional exchange, indicating that the Dy547 tag is not detached from the miRNA molecule (Figure 3D).
Stable focus conditions are mandatory throughout a FRAP experiment, especially when long-term measurements are performed. Compared to 2D applications, 3D-FRAP can compensate for potential focus drift. As shown in Figure 4, even slight focus changes affect the proper detection of the fluorescence signal of miRNA when just a single 2D layer is used for the FRAP experiment (Figure 4B). In contrast, for 3D-FRAP, a whole z-stack of the target cell is recorded, and maximum projections are subjected to data analysis (Figure 4A). The impact of focus drift on a normalized fluorescence-intensity curves is depicted in Figure 4C. While fluorescence recovery steadily increases over time in 3D-FRAP, 2D-FRAP acquisition results in a decline of the fluorescence signal.
Beside the compensation of focus changes, 3D-FRAP also provides spatial data of the gap junctional exchange of miRNA. Figure 5 shows a 3D surface rendering of a FRAP experiment. Compared to maximum projections, the acquisition of z-stacks can be used to create 3D reconstructions to investigate the localization and mobility of miRNA within cells (Figure 5). Co-labeling with organelles will allow for investigation into the involvement of other cellular components in miRNA transfer.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/55870/55870fig1.jpg" /> 
Figure 1: Representative Microscopic Images of Isolated Neonatal Cardiomyocytes. (A) Structured illumination microscopy shows the high expression of Cx43, which accumulates in large plaques at cell-cell borders (arrowheads). The pronounced formation of GJs with adjacent cells enables an extensive exchange of small molecules, including miRNA. Scale bar = 20 &#181;m (B) Purity of isolated cardiomyocytes. Labeling of &#945;-actinin demonstrates that cardiomyocytes represent the major cell type after isolation. Scale bar = 50 &#181;m. Cells were stained with anti-Cx43 (green) and anti-&#945;-actinin antibodies (red). Nuclei were visualized using DAPI (blue). A description of structured illumination microscopy and immunological labeling is provided in a previous publications20. <a href="https://cloudflare-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/55870/55870fig1large.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/55870/55870fig2.jpg" /> 
Figure 2: Transfection Efficiency, Viability, and Cell Density of Neonatal Cardiomyocyte culture. (A) Flow cytometry revealed that miR547 electroporation resulted in a transfection efficiency of ~45%. (B) Cell viability of cardiomyocytes used for FRAP. After isolation, electroporation, and reattachment, cardiomyocytes were subjected to live and dead cell staining and demonstrated high cell viability. (C) Microscopic image of mir547-transfected cardiomyocytes prepared for FRAP microscopy. A cell density of 80-90% ensures the pronounced formation of gap junctional cell-cell contacts. Scale bar = 50 &#181;m. A method for the flow cytometric analysis of transfection efficiency and viability staining is mentioned in previous publications20,21. <a href="https://cloudflare-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/55870/55870fig2large.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/55870/55870fig3.jpg" /> 
Figure 3: GJ-dependent Exchange of miRNA in Neonatal Cardiomyocytes. (A) Representative images of a FRAP experiment. After transfection with labeled miRNA, target cells within a cell cluster are photobleached by a strong laser pulse (pre-bleach versus bleach). Depending upon the extent of gap junctional coupling, the gap junctional influx of miRNA from adjacent cells results in increased fluorescence intensity in the bleached target cells (post-bleach t = 3, 6, 9, and 13 min). Scale bar = 20 &#181;m (B) Quantitative analysis of acquired FRAP data shows a fluorescence recovery of 20% after 13 min. To confirm the involvement GJs in the intercellular transfer of miRNA, a cx43 knockdown was performed, leading to a strong decrease of fluorescence recovery (50%). (C) Immunostaining with anti-Cx43 antibody and subsequent confocal microscopy demonstrate the efficiency of the Cx43 knockdown at the protein level. Scale bar = 50 &#181;m. (D) Comparison of the FRAP data of miR547 and calcein dye show the different transfer dynamics. The smaller molecular weight of calcein resulted in an increased fluorescence recovery compared to dy547-tagged miRNA molecules. FRAP curves summarize the data of n &#8805;56 cells, shown as the mean &#177; SEM. Statistical analysis was performed using two-way ANOVA followed by Bonferroni&#39;s post-hoc test (*P &#60;0.05, **P &#60;0.01, ***P &#60;0.001). <a href="https://cloudflare-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/55870/55870fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/55870/55870fig4.jpg" /> 
Figure 4: Compensation of Focus Drift using 3D-FRAP Microscopy. (A and B) Representative images of a photobleached cell (red line) demonstrate the impact of focus drift on the fluorescence intensity during 3D- and 2D-FRAP experiments. (A) In 3D-FRAP, z-stacks are recorded for each time-point, and maximum projections are used for data analysis, which corrects for focus drift. Scale bar = 20 &#181;m. (B) In contrast, for 2D-FRAP, just a single 2D-layer is subjected to data analysis. Thus, the overall fluorescence intensity is profoundly reduced by changes in the focus. Scale bar = 20 &#181;m. (C) Representative normalized fluorescence-intensity curves of 3D- and 2D-FRAP experiments indicate the strong effect of focus drift on fluorescence recovery and show the benefits of 3D acquisition when studying intercellular miRNA shuttling. <a href="https://cloudflare-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/55870/55870fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/55870/55870fig5.jpg" /> 
Figure 5: 3D Surface Rendering of a 3D-FRAP Experiment. (Left) Maximum projections of a FRAP measurement. The photobleached cell (red frame) demonstrates an increase in fluorescence intensity due to miRNA transfer from adjacent cells. (Right) Acquired z-stacks of the same photobleached cell (red) were subjected to 3D surface rendering to visualize the spatial distribution of the miRNA. Scale bar = 20 &#181;m. <a href="https://cloudflare-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/55870/55870fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Analysis of the Gap Junction-dependent Transfer of miRNA with 3D-FRAP Microscopy at JoVE.com

Analysis of the Gap Junction-dependent Transfer of miRNA with 3D-FRAP Microscopy

Here, we describe the application of three-dimensional fluorescence recovery after photobleaching (3D-FRAP) for the analysis of the gap junction-dependent shuttling of miRNA. In contrast to commonly applied methods, 3D-FRAP allows for the quantification of the intercellular transfer of small RNAs in real time, with high spatio-temporal resolution.

Small antisense RNAs, like miRNA and siRNA, play an important role in cellular physiology and pathology and, moreover, can be used as therapeutic agents ...

analysis-gap-junction-dependent-transfer-mirna-with-3d-frap

Universidad San Sebastian

Research

JoVE Journal

Biology

8.8K Views.  Reference and Translation Center for Cardiac Stem Cell Therapy (RTC). Here, we describe the application of three-dimensional fluorescence recovery after photobleaching (3D-FRAP) for the analysis of the gap junction-dependent shuttling of miRNA. In contrast to commonly applied methods, 3D-FRAP allows for the quantification of the intercellular transfer of small RNAs in real time, with high spatio-temporal resolution.

Video: Analysis of the Gap Junction-dependent Transfer of miRNA with 3D-FRAP Microscopy

Method to Visualize and Analyze Membrane Interacting Proteins by Transmission Electron Microscopy

Monotopic proteins exert their function when attached to a membrane surface, and such interactions depend on the specific lipid composition and on the availability of enough area to perform the function. Nanodiscs are used to provide a membrane surface of controlled size and lipid content. In the absence of bound extrinsic proteins, sodium phosphotungstate-stained nanodiscs appear as stacks of coins when viewed from the side by transmission electron microscopy (TEM). This protocol is therefore designed to intentionally promote stacking; consequently, the prevention of stacking can be interpreted as the binding of the membrane-binding protein to the nanodisc. In a further step, the TEM images of the protein-nanodisc complexes can be processed with standard single-particle methods to yield low-resolution structures as a basis for higher resolution cryoEM work. Furthermore, the nanodiscs provide samples suitable for either TEM or non-denaturing gel electrophoresis. To illustrate the method, Ca2+-induced binding of 5-lipoxygenase on nanodiscs is presented.

Many proteins perform their function when attached to membrane surfaces. The binding of extrinsic proteins on nanodisc membranes can be indirectly imaged by transmission electron microscopy. We show that the characteristic stacking (rouleau) of nanodiscs induced by the negative stain sodium phosphotungstate is prevented by the binding of extrinsic protein.

Monotopic proteins exert their function when attached to a membrane surface, and such interactions depend on the specific lipid composition and on the ...

Biochemistry

Live-cell Imaging of Migrating Cells Expressing Fluorescently-tagged Proteins in a Three-dimensional Matrix

Traditionally, cell migration has been studied on two-dimensional, stiff plastic surfaces. However, during important biological processes such as wound healing, tissue regeneration, and cancer metastasis, cells must navigate through complex, three-dimensional extracellular tissue. To better understand the mechanisms behind these biological processes, it is important to examine the roles of the proteins responsible for driving cell migration. Here, we outline a protocol to study the mechanisms of cell migration using the epithelial cell line (MDCK), and a three-dimensional, fibrous, self-polymerizing matrix as a model system. This optically clear extracellular matrix is easily amenable to live-cell imaging studies and better mimics the physiological, soft tissue environment. This report demonstrates a technique for directly visualizing protein localization and dynamics, and deformation of the surrounding three-dimensional matrix. 
Examination of protein localization and dynamics during cellular processes provides key insight into protein functions. Genetically encoded fluorescent tags provide a unique method for observing protein localization and dynamics. Using this technique, we can analyze the subcellular accumulation of key, force-generating cytoskeletal components in real-time as the cell maneuvers through the matrix. In addition, using multiple fluorescent tags with different wavelengths, we can examine the localization of multiple proteins simultaneously, thus allowing us to test, for example, whether different proteins have similar or divergent roles. Furthermore, the dynamics of fluorescently tagged proteins can be quantified using Fluorescent Recovery After Photobleaching (FRAP) analysis. This measurement assays the protein mobility and how stably bound the proteins are to the cytoskeletal network.
By combining live-cell imaging with the treatment of protein function inhibitors, we can examine in real-time the changes in the distribution of proteins and morphology of migrating cells. Furthermore, we also combine live-cell imaging with the use of fluorescent tracer particles embedded within the matrix to visualize the matrix deformation during cell migration. Thus, we can visualize how a migrating cell distributes force-generating proteins, and where the traction forces are exerted to the surrounding matrix. Through these techniques, we can gain valuable insight into the roles of specific proteins and their contributions to the mechanisms of cell migration.

Cellular processes such as cell migration have traditionally been studied on two-dimensional, stiff plastic surfaces. This report describes a technique for directly visualizing protein localization and analyzing protein dynamics in cells migrating in a more physiologically relevant, three-dimensional matrix.

Traditionally, cell migration has been studied on two-dimensional, stiff plastic surfaces. However, during important biological processes such as wound ...

Bioengineering

Visualization of Cortex Organization and Dynamics in Microorganisms, using Total Internal Reflection Fluorescence Microscopy

TIRF microscopy has emerged as a powerful imaging technology to study spatio-temporal dynamics of fluorescent molecules in vitro and in living cells1. The optical phenomenon of total internal reflection occurs when light passes from a medium with high refractive index into a medium with low refractive index at an angle larger than a characteristic critical angle (i.e. closer to being parallel with the boundary). Although all light is reflected back under such conditions, an evanescent wave is created that propagates across and along the boundary, which decays exponentially with distance, and only penetrates sample areas that are 100-200 nm near the interface2. In addition to providing superior axial resolution, the reduced excitation of out of focus fluorophores creates a very high signal to noise ratios and minimizes damaging effects of photobleaching2,3. Being a widefield technique, TIRF also allows faster image acquisition than most scanning based confocal setups.
At first glance, the low penetration depth of TIRF seems to be incompatible with imaging of bacterial and fungal cells, which are often surrounded by thick cell walls. On the contrary, we have found that the cell walls of yeast and bacterial cells actually improve the usability of TIRF and increase the range of observable structures4-8. Many cellular processes can therefore be directly accessed by TIRF in small, single-cell microorganisms, which often offer powerful genetic manipulation techniques. This allows us to perform in vivo biochemistry experiments, where kinetics of protein interactions and activities can be directly assessed in living cells.
We describe here the individual steps required to obtain high quality TIRF images for Saccharomyces cerevisiae or Bacillus subtilis cells. We point out various problems that can affect TIRF visualization of fluorescent probes in cells and illustrate the procedure with several application examples. Finally, we demonstrate how TIRF images can be further improved using established image restoration techniques.

Total Internal Reflection Fluorescence (TIRF) microscopy is a powerful approach to observe structures close to the cell surface at high contrast and temporal resolution. We demonstrate how TIRF can be employed to study protein dynamics at the cortex of cell wall-enclosed bacterial and fungal cells.

TIRF microscopy has emerged as  a powerful imaging technology to study spatio-temporal dynamics of fluorescent molecules  in vitro and in living cells1. ...

Lateral Diffusion and Exocytosis of Membrane Proteins in Cultured Neurons Assessed using Fluorescence Recovery and Fluorescence-loss Photobleaching

Membrane proteins such as receptors and ion channels undergo active trafficking in neurons, which are highly polarised and morphologically complex. This directed trafficking is of fundamental importance to deliver, maintain or remove synaptic proteins.
Super-ecliptic pHluorin (SEP) is a pH-sensitive derivative of eGFP that has been extensively used for live cell imaging of plasma membrane proteins1-2. At low pH, protonation of SEP decreases photon absorption and eliminates fluorescence emission. As most intracellular trafficking events occur in compartments with low pH, where SEP fluorescence is eclipsed, the fluorescence signal from SEP-tagged proteins is predominantly from the plasma membrane where the SEP is exposed to a neutral pH extracellular environment. When illuminated at high intensity SEP, like every fluorescent dye, is irreversibly photodamaged (photobleached)3-5. Importantly, because low pH quenches photon absorption, only surface expressed SEP can be photobleached whereas intracellular SEP is unaffected by the high intensity illumination6-10. 
FRAP (fluorescence recovery after photobleaching) of SEP-tagged proteins is a convenient and powerful technique for assessing protein dynamics at the plasma membrane. When fluorescently tagged proteins are photobleached in a region of interest (ROI) the recovery in fluorescence occurs due to the movement of unbleached SEP-tagged proteins into the bleached region. This can occur via lateral diffusion and/or from exocytosis of non-photobleached receptors supplied either by de novo synthesis or recycling (see Fig. 1). The fraction of immobile and mobile protein can be determined and the mobility and kinetics of the diffusible fraction can be interrogated under basal and stimulated conditions such as agonist application or neuronal activation stimuli such as NMDA or KCl application8,10. 
We describe photobleaching techniques designed to selectively visualize the recovery of fluorescence attributable to exocytosis. Briefly, an ROI is photobleached once as with standard FRAP protocols, followed, after a brief recovery, by repetitive bleaching of the flanking regions. This 'FRAP-FLIP' protocol, developed in our lab, has been used to characterize AMPA receptor trafficking at dendritic spines10, and is applicable to a wide range of trafficking studies to evaluate the intracellular trafficking and exocytosis.

This report describes the use of live cell imaging and photobleach techniques to determine the surface expression, transport pathways and trafficking kinetics of exogenously expressed, pH-sensitive GFP-tagged proteins at the plasma membrane of neurons.

Membrane proteins such as receptors and ion channels undergo active trafficking in neurons, which are highly polarised and morphologically complex. This ...

Neuroscience

Measurement of Force-Sensitive Protein Dynamics in Living Cells Using a Combination of Fluorescent Techniques

Cells sense and respond to physical cues in their environment by converting mechanical stimuli into biochemically-detectable signals in a process called mechanotransduction. A crucial step in mechanotransduction is the transmission of forces between the external and internal environments. To transmit forces, there must be a sustained, unbroken physical linkage created by a series of protein-protein interactions. For a given protein-protein interaction, mechanical load can either have no effect on the interaction, lead to faster disassociation of the interaction, or even stabilize the interaction. Understanding how molecular load dictates protein turnover in living cells can provide valuable information about the mechanical state of a protein, in turn elucidating its role in mechanotransduction. Existing techniques for measuring force-sensitive protein dynamics either lack direct measurements of protein load or rely on the measurements performed outside of the cellular context. Here, we describe a protocol for the F&#246;rster resonance energy transfer-fluorescence recovery after photobleaching (FRET-FRAP) technique, which enables the measurement of force-sensitive protein dynamics within living cells. This technique is potentially applicable to any FRET-based tension sensor, facilitating the study of force-sensitive protein dynamics in variety of subcellular structures and in different cell types.

Here, we present a protocol for the simultaneous use of F&#246;rster resonance energy transfer-based tension sensors to measure protein load and fluorescence recovery after photobleaching to measure protein dynamics enabling the measurement of force-sensitive protein dynamics within living cells.

Cells sense and respond to physical cues in their environment by converting mechanical stimuli into biochemically-detectable signals in a process called ...

An Analytical Tool that Quantifies Cellular Morphology Changes from Three-dimensional Fluorescence Images

The most common software analysis tools available for measuring fluorescence images are for two-dimensional (2D) data that rely on manual settings for inclusion and exclusion of data points, and computer-aided pattern recognition to support the interpretation and findings of the analysis. It has become increasingly important to be able to measure fluorescence images constructed from three-dimensional (3D) datasets in order to be able to capture the complexity of cellular dynamics and understand the basis of cellular plasticity within biological systems. Sophisticated microscopy instruments have permitted the visualization of 3D fluorescence images through the acquisition of multispectral fluorescence images and powerful analytical software that reconstructs the images from confocal stacks that then provide a 3D representation of the collected 2D images. Advanced design-based stereology methods have progressed from the approximation and assumptions of the original model-based stereology1 even in complex tissue sections2. Despite these scientific advances in microscopy, a need remains for an automated analytic method that fully exploits the intrinsic 3D data to allow for the analysis and quantification of the complex changes in cell morphology, protein localization and receptor trafficking. 
Current techniques available to quantify fluorescence images include Meta-Morph (Molecular Devices, Sunnyvale, CA) and Image J (NIH) which provide manual analysis. Imaris (Andor Technology, Belfast, Northern Ireland) software provides the feature MeasurementPro, which allows the manual creation of measurement points that can be placed in a volume image or drawn on a series of 2D slices to create a 3D object. This method is useful for single-click point measurements to measure a line distance between two objects or to create a polygon that encloses a region of interest, but it is difficult to apply to complex cellular network structures. Filament Tracer (Andor) allows automatic detection of the 3D neuronal filament-like however, this module has been developed to measure defined structures such as neurons, which are comprised of dendrites, axons and spines (tree-like structure). This module has been ingeniously utilized to make morphological measurements to non-neuronal cells3, however, the output data provide information of an extended cellular network by using a software that depends on a defined cell shape rather than being an amorphous-shaped cellular model. To overcome the issue of analyzing amorphous-shaped cells and making the software more suitable to a biological application, Imaris developed Imaris Cell. This was a scientific project with the Eidgen&ouml;ssische Technische Hochschule, which has been developed to calculate the relationship between cells and organelles. While the software enables the detection of biological constraints, by forcing one nucleus per cell and using cell membranes to segment cells, it cannot be utilized to analyze fluorescence data that are not continuous because ideally it builds cell surface without void spaces. To our knowledge, at present no user-modifiable automated approach that provides morphometric information from 3D fluorescence images has been developed that achieves cellular spatial information of an undefined shape (Figure 1). 
We have developed an analytical platform using the Imaris core software module and Imaris XT interfaced to MATLAB (Mat Works, Inc.). These tools allow the 3D measurement of cells without a pre-defined shape and with inconsistent fluorescence network components. Furthermore, this method will allow researchers who have extended expertise in biological systems, but not familiarity to computer applications, to perform quantification of morphological changes in cell dynamics.

We developed a software platform that utilizes Imaris Neuroscience, ImarisXT and MATLAB to measure the changes in morphology of an undefined shape taken from three-dimensional confocal fluorescence of single cells. This novel approach can be used to quantify changes in cell shape following receptor activation and therefore represents a possible additional tool for drug discovery.

The most common software analysis tools available for measuring fluorescence images are for two-dimensional (2D) data that rely on manual settings for ...

Single-cell Microinjection for Cell Communication Analysis

Gap junctions are intercellular channels that allow the communication of neighboring cells. This communication depends on the contribution of a hemichannel by each neighboring cell to form the gap junction. In mammalian cells, the hemichannel is formed by six connexins, monomers with four transmembrane domains and a C and N terminal within the cytoplasm. Gap junctions permit the exchange of ions, second messengers, and small metabolites. In addition, they have important roles in many forms of cellular communication within physiological processes such as synaptic transmission, heart contraction, cell growth and differentiation. We detail how to perform a single-cell microinjection of Lucifer Yellow to visualize cellular communication via gap-junctions in living cells. It is expected that in functional gap junctions, the dye will diffuse from the loaded cell to the connected cells. It is a very useful technique to study gap junctions since you can evaluate the diffusion of the fluorescence in real time. We discuss how to prepare the cells and the micropipette, how to use a micromanipulator and inject a low molecular weight fluorescent dye in an epithelial cell line.

We describe here how to perform a single-cell microinjection of Lucifer Yellow to visualize cellular communication via gap-junctions in living cells, and provide some useful tips. We expect that this paper will help everyone to evaluate the degree of cellular coupling due to functional gap junctions. Everything described here could be, in principle, adapted to other fluorescent dyes with molecular weight below 1,000 Daltons.

Gap junctions are intercellular channels that allow the communication of neighboring cells. This communication depends on the contribution of a ...

FRET Imaging in Three-dimensional Hydrogels

Imaging of F&#246;rster resonance energy transfer (FRET)&#160;is a powerful tool for examining cell biology in real-time. Studies utilizing FRET commonly employ two-dimensional (2D) culture, which does not mimic the three-dimensional (3D) cellular microenvironment. A method to perform quenched emission FRET imaging using conventional widefield epifluorescence microscopy of cells within a 3D hydrogel environment is presented. Here an analysis method for ratiometric FRET probes that yields linear ratios over the probe activation range is described. Measurement of intracellular cyclic adenosine monophosphate (cAMP) levels is demonstrated in chondrocytes under forskolin stimulation using a probe for EPAC1 activation (ICUE1) and the ability to detect differences in cAMP signaling dependent on hydrogel material type, herein a photocrosslinking hydrogel (PC-gel, polyethylene glycol dimethacrylate) and a thermoresponsive hydrogel (TR-gel). Compared with 2D FRET methods, this method requires little additional work. Laboratories already utilizing FRET imaging in 2D can easily adopt this method to perform cellular studies in a 3D microenvironment. It can further be applied to high throughput drug screening in engineered 3D microtissues. Additionally, it is compatible with other forms of FRET imaging, such as anisotropy measurement and fluorescence lifetime imaging (FLIM), and with advanced microscopy platforms using confocal, pulsed, or modulated illumination.

F&#246;rster resonance energy transfer (FRET) imaging is a powerful tool for real-time cell biology studies. Here a method for FRET imaging cells in physiologic three-dimensional (3D) hydrogel microenvironments using conventional epifluorescence microscopy is presented. An analysis for ratiometric FRET probes that yields linear ratios over the activation range is described.

Imaging of F&#246;rster resonance energy transfer (FRET)&#160;is a powerful tool for examining cell biology in real-time. Studies utilizing FRET commonly ...

Functional Imaging of Viral Transcription Factories Using 3D Fluorescence Microscopy

It is well known that spatial and temporal regulation of genes is an integral part of governing proper gene expression. Consequently, it is invaluable to understand where and when transcription is taking place within nuclear space and to visualize the relationship between episomes infected within the same cell's nucleus. Here, both immunofluorescence (IFA) and RNA-FISH have been combinedto identify actively transcribing Kaposi's sarcoma-associated herpesvirus (KSHV) episomes. By staining KSHV latency-associated nuclear antigen (LANA), it is possible to locate where viral episomes exist within the nucleus. In addition, by designing RNA-FISH probes to target the intron region of a viral gene, which is expressed only during productive infection, nascent RNA transcripts can be located. Using this combination of molecular probes, it is possible to visualize the assembly of large viral transcription factories and analyze the spatial regulation of viral gene expression during KSHV reactivation. By including anti-RNA polymerase II antibody staining, one can also visualize the association between RNA polymerase II (RNAPII) aggregation and KSHV transcription during reactivation.

Viral transcriptional factories are discrete structures that are enriched with cellular RNA polymerase II to increase viral gene transcription during reactivation. Here, a method to locate sites of actively transcribing viral chromatin in 3D nuclear space by a combination of immunofluorescence staining and in situ RNA hybridization is described.

It is well known that spatial and temporal regulation of genes is an integral part of governing proper gene expression. Consequently, it is invaluable to ...

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.