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="http://cloudflare.jove.com/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="http://cloudflare.jove.com/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="http://cloudflare.jove.com/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="http://cloudflare.jove.com/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="http://cloudflare.jove.com/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 Científica del Sur

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

High Throughput MicroRNA Profiling: Optimized Multiplex qRT-PCR at Nanoliter Scale on the Fluidigm Dynamic ArrayTM IFCs

The broad involvement of miRNAs in critical processes underlying development, tissue homoeostasis and disease has led to a surging interest among the research and pharmaceutical communities. To study miRNAs, it is essential that the quantification of microRNA levels is accurate and robust. By comparing wild-type to small RNA deficient mouse embryonic stem cells (mESC), we revealed a lack of accuracy and robustness in previous published multiplex qRT-PCR techniques. Here, we describe an optimized method, including purifying away excessive primers from previous multiplex steps before singleplex real time detection, which dramatically increases the accuracy and robustness of the technique. Furthermore, we explain how performing the technique on a microfluidic chip at nanoliter volumes significantly reduces reagent costs and permits time effective high throughput miRNA expression profiling.

Here we describe an optimized multiplex reverse transcriptase quantitative PCR (qRT-PCR) protocol in combination with a microfluidic platform as a cost and time effective high-throughput screening tool for microRNA (miRNA) expression levels, especially when working with limited amounts of sample.

The broad involvement of miRNAs in critical processes underlying development, tissue homoeostasis and disease has led to a surging interest among the ...

Bioengineering

Biotin-based Pulldown Assay to Validate mRNA Targets of Cellular miRNAs

MicroRNAs (miRNAs) are a class of small noncoding RNAs that post-transcriptionally regulate cellular gene expression. MiRNAs bind to the 3&#39; untranslated region (UTR) of target mRNA to inhibit protein translation or in some instances cause mRNA degradation. The binding of the miRNA to the 3&#39; UTR of the target mRNA is mediated by a 2&#8211;8 nucleotide seed sequence at the 5&#39; end of miRNA. While the role of miRNAs as cellular regulatory molecules is well established, identification of the target mRNAs with functional relevance remains a challenge. Bioinformatic tools have been employed to predict sequences within the 3&#39; UTR of mRNAs as potential targets for miRNA binding. These tools have also been utilized to determine the evolutionary conservation of such sequences among related species in an attempt to predict functional role. However, these computational methods often generate false positive results and are limited to predicting canonical interaction between miRNA and mRNA. Therefore, experimental procedures that measure direct binding of miRNA to its mRNA target are necessary to establish functional interaction. In this report, we describe a sensitive method for validating direct interaction between the cellular miRNA miR-125b and the 3&#39; UTR of PARP-1 mRNA. We elaborate a protocol in which synthetic biotinylated-miRNA mimics were transfected into mammalian cells and the miRNA-mRNA complex in the cellular lysate was pulled down with streptavidin-coated magnetic beads. Finally, the target mRNA in the pulled-down nucleic acid complex was quantified using a qPCR-based strategy.

This report describes a fast and reliable method for validating mRNA targets of cellular miRNAs. The method uses synthetic biotinylated Locked Nucleic Acid (LNA)-based miRNA mimics to capture target mRNA. Subsequently, streptavidin-coated magnetic beads are employed to pulldown the target mRNA for quantification by qPCR polymerase chain reaction.

MicroRNAs (miRNAs) are a class of small noncoding RNAs that post-transcriptionally regulate cellular gene expression. MiRNAs bind to the 3&#39; ...

Genetics

Characterization of Functionally Associated miRNAs in Glioblastoma and their Engineering into Artificial Clusters for Gene Therapy

The biological relevance of microRNAs (miRNAs) in health and disease significantly relies on specific combinations of many simultaneously deregulated miRNAs rather than the action of a single miRNA. The characterization of these specific miRNAs modules is a fundamental step in maximizing their use in therapy. This is extremely relevant because their combinatorial attributes can be practically exploited. Described here is a method to define a specific miRNA signature relevant to the control of oncogenic chromatin repressors in glioblastoma. The approach first defines a general group of miRNAs that are deregulated in tumors in comparison to normal tissue. The analysis is further refined by differential culture conditions, underscoring a subgroup of miRNAs that are co-expressed simultaneously during specific cellular states. Finally, the miRNAs that satisfy these filters are combined into an artificial polycistronic transgenes, which is based on a scaffold of naturally existing miRNA clusters genes, then used for overexpression of these miRNA modules into receiving cells.

Described here is a protocol for characterizing modules of biologically synergistic miRNAs and their assembly into short transgenes, which allows simultaneous overexpression for gene therapy applications.

The biological relevance of microRNAs (miRNAs) in health and disease significantly relies on specific combinations of many simultaneously deregulated ...

A Reporter Assay to Analyze Intronic microRNA Maturation in Mammalian Cells

MicroRNAs (miRNAs) are short RNA molecules that are widespread in eukaryotes. Most miRNAs are transcribed from introns, and their maturation involves different RNA-binding proteins in the nucleus. Mature miRNAs frequently mediate gene silencing, and this has become an important tool for comprehending post-transcriptional events. Besides that, it can be explored as a promising methodology for gene therapies. However, there is currently a lack of direct methods for assessing miRNA expression in mammalian cell cultures. Here, we describe an efficient and simple method that aids in determining miRNA biogenesis and maturation through confirmation of its interaction with target sequences. Also, this system allows the separation of exogenous miRNA maturation from its endogenous activity using a doxycycline-inducible promoter capable of controlling primary miRNA (pri-miRNA) transcription with high efficiency and low cost. This tool also allows modulation with RNA-binding proteins in a separate plasmid. In addition to its use with a variety of different miRNAs and their respective targets, it can be adapted to different cell lines, provided these are amenable to transfection.

We developed an intronic microRNA biogenesis reporter assay to be used in cells in vitro with four plasmids: one with intronic miRNA, one with the target, one to overexpress a regulatory protein, and one for Renilla luciferase. The miRNA was processed and could control luciferase expression by binding to the target sequence.

MicroRNAs (miRNAs) are short RNA molecules that are widespread in eukaryotes. Most miRNAs are transcribed from introns, and their maturation involves ...

Cancer Research

Genome-wide Screen for miRNA Targets Using the MISSION Target ID Library

The Target ID Library is designed to assist in discovery and identification of microRNA (miRNA) targets. The Target ID Library is a plasmid-based, genome-wide cDNA library cloned into the 3'UTR downstream from the dual-selection fusion protein, thymidine kinase-zeocin (TKzeo). The first round of selection is for stable transformants, followed with introduction of a miRNA of interest, and finally, selecting for cDNAs containing the miRNA's target. Selected cDNAs are identified by sequencing (see Figure 1-3 for Target ID Library Workflow and details).
To ensure broad coverage of the human transcriptome, Target ID Library cDNAs were generated via oligo-dT priming using a pool of total RNA prepared from multiple human tissues and cell lines. Resulting cDNA range from 0.5 to 4 kb, with an average size of 1.2 kb, and were cloned into the p3&#900;TKzeo dual-selection plasmid (see Figure 4 for plasmid map). The gene targets represented in the library can be found on the Sigma-Aldrich webpage. Results from Illumina sequencing (Table 3), show that the library includes 16,922 of the 21,518 unique genes in UCSC RefGene (79%), or 14,000 genes with 10 or more reads (66%).

The Target ID Library is a plasmid-based, genome-wide collection of cloned cDNA used to identify miRNA targets. Here we demonstrate its use and application.

The Target ID Library is designed to assist in discovery and identification of microRNA (miRNA) targets. The Target ID Library is a plasmid-based, ...

Identifying Targets of Human microRNAs with the LightSwitch Luciferase Assay System using 3'UTR-reporter Constructs and a microRNA Mimic in Adherent Cells

MicroRNAs (miRNAs) are important regulators of gene expression and play a role in many biological processes. More than 700 human miRNAs have been identified so far with each having up to hundreds of unique target mRNAs. Computational tools, expression and proteomics assays, and chromatin-immunoprecipitation-based techniques provide important clues for identifying mRNAs that are direct targets of a particular miRNA. In addition, 3'UTR-reporter assays have become an important component of thorough miRNA target studies because they provide functional evidence for and quantitate the effects of specific miRNA-3'UTR interactions in a cell-based system. To enable more researchers to leverage 3'UTR-reporter assays and to support the scale-up of such assays to high-throughput levels, we have created a genome-wide collection of human 3'UTR luciferase reporters in the highly-optimized LightSwitch Luciferase Assay System. The system also includes synthetic miRNA target reporter constructs for use as positive controls, various endogenous 3'UTR reporter constructs, and a series of standardized experimental protocols.
Here we describe a method for co-transfection of individual 3'UTR-reporter constructs along with a miRNA mimic that is efficient, reproducible, and amenable to high-throughput analysis.

MicroRNAs (miRNAs) are important regulators of gene expression and have been shown to play a role in numerous biological processes. To better understand miRNA-UTR interactions, we have created a genome-wide collection of 3 UTR luciferase reporters paired with a novel luciferase gene and assay reagent, the LightSwitch system.

MicroRNAs (miRNAs) are important regulators of gene expression and play a role in many biological processes.  More than 700 human miRNAs have been ...

Highly Efficient Ligation of Small RNA Molecules for MicroRNA Quantitation by High-Throughput Sequencing

MiRNA cloning and high-throughput sequencing, termed miR-Seq, stands alone as a transcriptome-wide approach to quantify miRNAs with single nucleotide resolution. This technique captures miRNAs by attaching 3&#8217; and 5&#8217; oligonucleotide adapters to miRNA molecules and allows de novo miRNA discovery. Coupling with powerful next-generation sequencing platforms, miR-Seq has been instrumental in the study of miRNA biology. However, significant biases introduced by oligonucleotide ligation steps have prevented miR-Seq from being employed as an accurate quantitation tool. Previous studies demonstrate that biases in current miR-Seq methods often lead to inaccurate miRNA quantification with errors up to 1,000-fold for some miRNAs1,2. To resolve these biases imparted by RNA ligation, we have developed a small RNA ligation method that results in ligation efficiencies of over 95% for both 3&#8217; and 5&#8242; ligation steps. Benchmarking this improved library construction method using equimolar or differentially mixed synthetic miRNAs, consistently yields reads numbers with less than two-fold deviation from the expected value. Furthermore, this high-efficiency miR-Seq method permits accurate genome-wide miRNA profiling from in vivo total RNA samples2.

MicroRNAs (miRNAs) are a widely conserved class of regulatory molecules. Here we describe a miRNA cloning method that relies upon two potent ligation steps followed by high-throughput sequencing. Our method permits accurate genome-wide quantitation of miRNAs.

MiRNA cloning and high-throughput sequencing, termed miR-Seq, stands alone as a transcriptome-wide approach to quantify miRNAs with single nucleotide ...

Probe-based Real-time PCR Approaches for Quantitative Measurement of microRNAs

Probe-based quantitative PCR (qPCR) is a favoured method for measuring transcript abundance, since it is one of the most sensitive detection methods that provides an accurate and reproducible analysis. Probe-based chemistry offers the least background fluorescence as compared to other (dye-based) chemistries. Presently, there are several platforms available that use probe-based chemistry to quantitate transcript abundance. qPCR in a 96 well plate is the most routinely used method, however only a maximum of 96 samples or miRNAs can be tested in a single run. This is time-consuming and tedious if a large number of samples/miRNAs are to be analyzed. High-throughput probe-based platforms such as microfluidics (e.g. TaqMan Array Card) and nanofluidics arrays (e.g. OpenArray) offer ease to reproducibly and efficiently detect the abundance of multiple microRNAs in a large number of samples in a short time. Here, we demonstrate the experimental setup and protocol for miRNA quantitation from serum or plasma-EDTA samples, using probe-based chemistry and three different platforms (96 well plate, microfluidics and nanofluidics arrays) offering increasing levels of throughput.

Circulating microRNAs have recently emerged as promising and novel biomarkers for various cancers and other diseases. The goal of this article is to discuss three different probe-based real-time PCR platforms and methods that are available to quantify and determine the abundance of circulating microRNAs.

Probe-based quantitative PCR (qPCR) is a favoured method for measuring transcript abundance, since it is one of the most sensitive detection methods that ...

Detection of miRNA Targets in High-throughput Using the 3'LIFE Assay

Luminescent Identification of Functional Elements in 3&#8217;UTRs (3&#8217;LIFE) allows the rapid identification of targets of specific miRNAs within an array of hundreds of queried 3&#8217;UTRs. Target identification is based on the dual-luciferase assay, which detects binding at the mRNA level by measuring translational output, giving a functional readout of miRNA targeting. 3&#8217;LIFE uses non-proprietary buffers and reagents, and publically available reporter libraries, making genome-wide screens feasible and cost-effective. 3&#8217;LIFE can be performed either in a standard lab setting or scaled up using liquid handling robots and other high-throughput instrumentation. We illustrate the approach using a dataset of human 3&#8217;UTRs cloned in 96-well plates, and two test miRNAs, let-7c and miR-10b. We demonstrate how to perform DNA preparation, transfection, cell culture and luciferase assays in 96-well format, and provide tools for data analysis. In conclusion 3&#39;LIFE is highly reproducible, rapid, systematic, and identifies high confidence targets.

Luminescent identification of functional elements in 3&#8217; untranslated regions (3&#8217;UTRs) (3&#8217;LIFE) is a technique to identify functional regulation in 3&#8217;UTRs by miRNAs or other regulatory factors. This protocol utilizes high-throughput methodology such as 96-well transfection and luciferase assays to screen hundreds of putative interactions for functional repression.

Luminescent Identification of Functional Elements in 3&#8217;UTRs (3&#8217;LIFE) allows the rapid identification of targets of specific miRNAs within an ...

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.