We thank Dr. Rachid Mazroui (Universit&#233; Laval, Qu&#233;bec, Canada) for the critical reading of the manuscript. We thank the Cell Imaging Unit of the Research Center for their technical assistance. M.-&#201;. Huot is a Junior 1 Research Scholar of the Fonds de Recherche du Qu&#233;bec-Sant&#233; (FRQ-S). This work was supported by the Canadian Institutes of Health Research (grant number CIHR, MOP-286437 to M.-&#201;. Huot).

1. Determination of Puromycilation Conditions
NOTE: This technique describes the method used to assess localized translation during the MRC-5 cell adhesion process5. As puromycilation can be done in any cell, it is important to optimize the puromycilation conditions for the specific cell lines to be used, because the treatment conditions are not identical for each cell line in terms of the puromycin concentration and the desired incubation time. To show how these conditions are defined, three example cell lines (i.e., HeLa, MRC-5, and Huh-7) were treated with increasing concentrations of puromycin for different incubation times (Figure 1).
<ol>
	<li>Grow identical numbers of cells in a 24-well plate in 0.5 mL of the appropriate complete culture medium 16 h before the test. Seed 30,000 MRC-5 cells, 50,000 HeLa cells, or 50,000 HuH-7 cells per well. Make sure to avoid exceeding 60-70% confluence (Figure 1A).</li>
	<li>Prepare 6 tubes with 1.5 mL of complete cell culture medium with increasing concentrations of puromycin (0, 5, 10, 15, 20, and 30 &#181;g/mL). Pre-warm at 37 &#176;C.</li>
	<li>For each concentration of puromycin medium (prepared in step 1.2), transfer 0.5 mL from each tube into 6 new tubes and add cycloheximide at a final concentration of 50 &#181;g/mL to all 6 new tubes. Pre-warm at 37 &#176;C.</li>
	<li>Change the medium with 0.5 mL of complete culture medium (row 1 and 2). For the third row, add 0.5 mL of complete culture medium supplemented with 50 &#181;g/mL of cycloheximide (Figure 1B). 
	NOTE: Cycloheximide treatment has been established as the best negative control for protein puromycilation due to its ability to block translation elongation. Because puromycin incorporation requires translation elongation, cycloheximide treatment leads to a loss of puromycilated protein signals5,18.</li>
	<li>Incubate for 15 min at 37 &#176;C (5% CO2).</li>
	<li>Add 0.5 mL of the puromycin medium prepared in step 1.2 to the second row to obtain final concentrations of 0, 2.5, 5, 7.5, 10, and 15 &#181;g/mL, respectively, in columns A, B, C, D, E, and F. At the same time, add 0.5 mL of puromycin to the cycloheximide-supplemented medium (step 1.3) in the third row (Figure 1C).</li>
	<li>Incubate for 5 min at 37 &#176;C (5% CO2).</li>
	<li>Add 0.5 mL of the puromycin medium prepared in step 1.2 to the first row to obtain final concentrations of 0.0, 2.5, 5, 7.5, 10, and 15 &#181;g/mL (Figure 1C).</li>
	<li>Incubate for 5 min at 37 &#176;C (5% CO2).</li>
	<li>Wash each well from rows 1 to 3 with 1 mL of ice-cold 1X phosphate-buffered saline (PBS) 5 min after the addition of puromycin to wells of the first row (wash twice). Add 75 &#181;L of 1X Laemmli buffer (4% sodium dodecyl sulfate (SDS), 4% 2-mercaptoethanol, 0.120 M Tris HCl pH 6.8, 0.004% bromophenol blue, and 10% glycerol) to obtain whole-cell lysates for each condition.</li>
	<li>Run 10% SDS-polyacrylamide gel electrophoresis (SDS-PAGE) using 1/3 of the prepared samples to assess puromycin incorporation.
	<ol>
		<li>Determine the level of puromycin incorporation by Western blot analysis using an anti-puromycin antibody (12D10) diluted at a ratio of 1:25,000 in primary antibody incubation buffer (2% bovine serum albumin (BSA)), 430 mM NaCl, 10 mM Tris pH 7.4, and 0.01% sodium azide). 
		NOTE: Representative images corresponding to different cell line extracts made as described in step 1 are shown as examples in Figure 2.</li>
	</ol>
	</li>
</ol>
2. In Cellulo Localized Translation Visualization
NOTE: The technique described here was used to assess localized translation within SICs in MRC-5 cells during the adhesion process5. Although MRC-5 cells are used here, other cell lines can be used with the same methodology described in step 2.1.
<ol>
	<li>Cell preparation.
	<ol>
		<li>Detach MRC-5 cells using 0.25% trypsin/2.21 mM ethylenediaminetetraacetic acid (EDTA) in Hank&#39;s Balanced Salt Solution (HBSS). Pellet the cells by centrifugation (5 min at 200 x g) in a 15-mL conical centrifuge tube and suspend them in Dulbecco&#39;s Modified Eagle Medium (DMEM) supplemented with 10% fetal bovine serum (FBS) at 40,000 cells/mL after counting with a counting chamber.</li>
		<li>Incubate the suspended cells at 37 &#176;C under gentle rotation using a tube rotator for 20 min to maintain them in suspension. 
		NOTE: This step is critical to allow for the complete dissociation of the focal adhesion complexes.</li>
		<li>Plate 2 mL of suspended MRC-5 cells (40,000 cells/mL) in two 35-mm glass-bottom dishes. Use the first 35-mm glass-bottom dish for the puromycilation assay (see step 2.1.5) and use the second as a negative control (see step 2.1.6).</li>
		<li>Keep the cells at 37 &#176;C (5% CO2) for 55 min to allow for cellular adhesion and SIC formation.</li>
		<li>Add puromycin to the medium in the 35-mm glass-bottom dishes (assay) using the concentration previously defined in section 1. To treat MRC-5 cells, use 10 &#181;g/mL puromycin for 5 min at 37 &#176;C (5% CO2).</li>
		<li>As a negative control, add cycloheximide to the second 35-mm glass-bottom dish 40 min after seeding the cells to pre-treat them with 50 &#181;g/mL cycloheximide (Figure 2B). Then, 15 min after cycloheximide addition, add puromycin to the medium using the same concentration and incubation time as in step 2.1.5 (e.g., use 10 &#181;g/mL of puromycin for 5 min at 37 &#176;C (5% CO2) for MRC-5).</li>
		<li>Wash each plate twice with 2 mL of ice-cold 1X PBS and fix the cells with 1 mL of 4% formaldehyde (diluted in 1X PBS) for 15 min at room temperature (RT). 
		Caution: Paraformaldehyde must be used strictly in a chemical hood.</li>
	</ol>
	</li>
	<li>Immunostaining.
	<ol>
		<li>Following paraformaldehyde fixation, wash three times with 2 mL of 1X PBS and incubate in 500 &#181;L of PBS-TritonX-100 (0.5%) for 20 min at RT to permeabilize the cells.</li>
		<li>To prevent nonspecific antibody binding, block the sample with 500 &#181;L PBS &#8211; BSA (1%) for 20 min at RT.</li>
		<li>Wash three times with 2 mL of PBS-Tween20 (0.1%) and incubate with 300 &#181;L of anti-puromycin antibody (12D10) diluted 1:12,500 in 1X PBS for 1 h at RT.</li>
		<li>Wash 3 times with 2 mL of PBS-Tween20 (0.1%) and incubate with 300 &#181;L of anti-mouse IgG conjugated to a fluorophore (here, 488 nm) diluted in PBS to visualize puromycilated polypeptides and with phalloidin-conjugated to another fluorophore (here, 555 nm) to visualize F-actin. Incubate for 1 h at RT.</li>
		<li>Wash three times with 2 mL of PBS-Tween20 (0.1%) and then incubate for 5 min at RT in 300 &#181;L of PBS-Tween20 (0.1%) supplemented with 1 &#181;g/mL 4&#39;,6-diamidino-2-phenylindole (DAPI).</li>
		<li>Wash three times with 2 mL of PBS-Tween20 (0.1%) and then wash with 2 mL of 1X PBS. Prevent the cells from drying by keeping the sample in 2 mL of 1X PBS during image acquisition.</li>
	</ol>
	</li>
</ol>
3. Immunofluorescent Image Acquisition
NOTE: The following methodology can be used with any commercially available confocal imaging system.
<ol>
	<li>Determine the appropriate settings for each fluorophore to maximize the dynamic range for quantification. 
	NOTE: Representative quantification can only be obtained if pixel saturation is avoided and the signal threshold is adjusted properly. These settings can be attained by carefully adjusting the laser power, high voltage (HV), gain, and offset.
	<ol>
		<li>Adjust the zoom factor (5X) and the number of pixels (512 x 512) to optimize the pixel size. 
		NOTE: This should be at least 2.3-times smaller than the optical resolution, according to the Nyquist theorem.</li>
		<li>Decrease the scan speed (12.5-20 &#181;s/pixel) and use averaging (2-3 times) to improve the signal-to-noise ratio according to the settings mentioned above.</li>
	</ol>
	</li>
	<li>Manually determine the appropriate top and bottom focal planes along the Z-axis with the optimal step size (0.4 &#181;m/slice). 
	NOTE: The step size plane is determined by the axial resolution divided by 2.3, according to the Nyquist theorem. Typically, 20-25 layers are necessary to cover the entire cell depth of adherent cells.</li>
	<li>Acquire a confocal image of an entire single cell with a 60X Plan Apo oil immersion objective (1.42 numerical aperture).</li>
</ol>
4. Immunofluorescent Image Quantification
<ol>
	<li>Determination of enrichment.
	<ol>
		<li>Open an image corresponding to the z-plane layer of interest from the acquired cell data file using the ImageJ software (freeware available at http://fiji.sc).</li>
		<li>Draw a line using the line drawing tool (Tool bar &#8594; Straight) through the regions of the cell where quantification is desired. Modify the line width by double-clicking on &#34;straight;&#34; the intensity values will be averaged through the width of the line (set at 8 in Figure 3). 
		NOTE: A thinner line is preferred to avoid signal overlap from different structures.</li>
		<li>After the line is properly set, profile the signal density along the determined axis (menu bar &#8594; Analyze &#8594; Plot Profile). 
		NOTE: A new window will open that shows a profile corresponding to the signal intensity for each pixel of the selected channel (specific fluorophore) along the line for the selected z-plane section.
		<ol>
			<li>Repeat the process for each channel corresponding to the fluorophore used (i.e., one quantification for 488, one for 568, and one for 405). 
			NOTE: The signal intensity value will always be ordered following the orientation of the drawn line.</li>
		</ol>
		</li>
		<li>To obtain the numerical gray-scaled values for each pixel of the profile corresponding to the quantifications of the selected z-plane layer, copy the profile data (menu bar in image window &#8594; Copy) and paste it in any spreadsheet software.</li>
		<li>Repeat steps 4.1.1-4.1.4 for each z-plane section of the acquired confocal image and each channel where quantification is desired. 
		NOTE: The quantification of different z-plane layers can be pooled to obtain a general assessment of the special enrichment of the signal by calculating the sum value of each pixel along the line for each z-plane layer. This type of pooling can only be achieved if the line drawn is identical for each z-plane layer included in the mean values.</li>
		<li>As an alternative method for the whole-cell quantification of different channels with a large number of layers, use the macro called StackprofileData (see the Supplemental File). 
		NOTE: This macro is accessible freely at https://imagej.nih.gov/ij/macros/StackProfileData.txt and can be used as follows:
		<ol>
			<li>Paste the macro into a text file and save it.</li>
			<li>Open the image file that includes all of the confocal layers of the cell.</li>
			<li>Draw a line, as described in step 4.1.2, open a new window to import the macro (menu bar &#8594; Plugins &#8594; Macros&#8594; Run), choose the previously saved text file corresponding to the StackprofileData macro, and click &#34;Open.&#34; 
			NOTE: Following macro importation, a new window named &#34;Results&#34; will open and all of the quantification along the determined axis will be listed for each z-plane layer and channel present in the image files.</li>
			<li>Copy the data for each channel (menu bar &#8594; Edit &#8594; Copy) to obtain the graphical profile representation corresponding to the signal quantifications. Paste it into any spreadsheet software. Calculate the sum value of each channel for each pixel along the line for each z-plane layer. Use these values to create a graphical representation similar to the one presented in Figure 3.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Quantification of signal in a designated cellular area or volume.
	<ol>
		<li>Open the image file with ImageJ software.</li>
		<li>Draw an area to denote the area of interest using the geometrical form function (tool bar &#8594;&#34;rectangle,&#34; &#34;oval,&#34; or &#34;polygon&#34;). 
		NOTE: The quantification value will be determined for the area corresponding to the drawn form.</li>
		<li>Adjust the threshold level to avoid any background signal (menu bar &#8594; Image Adjust &#8594; Threshold); a new window will appear to represent the pixel intensities in histogram form. Change the values to include/exclude pixels in the quantification using the slider. Select a threshold that eliminates red pixels outside the cell, as pixels highlighted in red will be included in the quantification.</li>
		<li>Define the measurement parameters to quantify the pixel signal within the selected area, without background signal (menu bar &#8594; Analyze &#8594; Set Measurements), and make sure to check the &#34;Limit to Threshold&#34; and the &#34;Integrated Density&#34; boxes.</li>
		<li>Measure the quantitative values for the selected area (menu bar &#8594; Analyze &#8594; Measure). 
		NOTE: Signal intensity quantification will be presented in a new window under the &#34;IntDen&#34; identification, which represents the product of the mean gray values and the number of pixels within the selected area.</li>
		<li>Draw an area that includes the entire cell using a geometrical form (tool bar &#8594;&#34;rectangle,&#34; &#34;oval,&#34; or &#34;polygon&#34;) and proceed with steps 4.2.3-4.2.5 to obtain a value corresponding to the total signal. Calculate the ratio of the signal within the selected area over the whole signal to obtain the percent of the signal within the area of interest.</li>
		<li>Repeat steps 4.2.2-4.2.6 to obtain the quantification of cell volume for each z-plane. Adapt the geometrical form as needed.</li>
		<li>Copy/paste the data obtained from the &#34;Results&#34; windows for each z-plane layer into a spreadsheet for graphical depiction and to find a mean value.</li>
	</ol>
	</li>
</ol>

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The authors have nothing to disclose.

Recent technological advances have allowed for a better understanding of the mechanisms involved in translational upregulation or the repression of specific mRNAs in biological processes, such as neuronal development, drug response, and metastasis. The cost-effective methodology described here allows translation events to be visualized in cells to study how RNA-binding proteins regulate metastatic processes, such as cellular adhesion, migration, and invasion.
Although numerous methods to asses...

The regulation of translation by different cellular functions has prompted many research teams to develop new tools and methods to determine the subcellular localization of mRNA translation and regulated protein synthesis1,2,3,4. These recent technological advances allow for an improved understanding of the mechanisms involving translation upregulation or the repression of specific mRNAs during biological processes, such as neuronal development, drug response, and metastasis5,6,7,8. However, most of these methods require expensive or hazardous reagents and specific equipment that might not be available to most laboratories. As such, a cost-effective method to allow for the rapid assessment of translation events was developed to specifically circumvent these potential issues. This method detects acute translational modulations that occur during specific cellular processes and also allows for the localization of translation using confocal microscopy.
The methods described here were used to monitor localized translation within subcellular compartments called spreading initiation centers (SIC)5. SICs are transient structures found in seeded cells that are localized on top of nascent adhesion complexes. Although SICs and adhesion complexes are distinct, their fates are closely linked. Indeed, SICs are known to gradually disappear upon focal adhesion complex maturation into an adhesion site during the initial phase of adhesion. We found that RNA-binding proteins known to specifically control mRNA translation (e.g., Sam68, FMRP, and G3BP1) and polyadenylated RNAs were enriched within these structures5. Using the methods described here, we showed that the regulation of SIC-associated mRNA translation acts as a checkpoint allowing seeded cells to consolidate cell adhesion. This method, based on puromycin incorporation, could be considered an adapted version of the surface sensing of translation assay (SunSET). Originally developed to measure global protein synthesis rates using a non-radioactively labeled amino acid, protein puromycilation offers an efficient way to visualize de novo protein synthesis9. This method relies on the intrinsic behavior of puromycin, an antibiotic that blocks translation through premature chain termination in the ribosome10. Indeed, puromycin is structurally analogous to tyrosyl-tRNA, which allows for incorporation into elongating peptide chains via the formation of a peptide bond. However, puromycin binding to a growing peptide chain prevents a new peptide bond from being formed with the next aminoacyl-tRNA, since puromycin has a non-hydrolysable amide bond instead of the hydrolysable ester bond found in tRNAs. Thus, the incorporation of puromycin into elongating polypeptides results in the premature release of numerous truncated puromycilated polypeptides corresponding to actively translated mRNA9,11,12.
Using this method, it was possible to assess active translation within a short time widow (e.g., 5 min) during cellular adhesion using a specific antibody directed against puromycin on cells that were supplemented with the antibiotic for 5-min periods at different time points during the cell adhesion process5. The precision of this assay relies on highly specific antibodies directed against the puromycilated moiety. Immunofluorescent detection of the puromycilated polypeptide provides a general subcellular repartition of the newly translated mRNA, which can also be quantified with great accuracy using confocal imaging systems.
Hence, this method offers a relevant option for a large number of laboratories studying translational regulatory mechanisms involved in processes such as neuronal granulation6,13,14,15, morphogen mRNA localization, and translation during development16,17. It is also well suited to studying localized or compartmentalized translation during rapid biological events, such as cell migration, adhesion, or invasion, or to simply assess drug treatments that might induce translational changes5,7,18. Overall, this method allows for the visualization of localized or controlled translation events in a rapid, precise, and cost-effective way.

<table><tbody><tr><td>DMEM</td><td>wisent</td><td>319-005-CL</td><td></td></tr><tr><td>Trypsine</td><td>wisent</td><td>325-043 EL</td><td></td></tr><tr><td>FBS</td><td>Thermo Fisher Scientific</td><td>12483020</td><td></td></tr><tr><td>Puroycin antibody 12D10</td><td>EMD millipore</td><td>MABE343</td><td>western blot dilution 1:25,000&lt;br/&gt; Immunofluoresence dilution 1:10,000</td></tr><tr><td>Anti-mouse IgG, HRP-linked Antibody</td><td>cell signaling technology</td><td>7076</td><td>western blot dilution 1:8,000</td></tr><tr><td>Western Lightning Plus-ECL</td><td>Perkin Elmer</td><td>NEL104001EA</td><td></td></tr><tr><td>Anti-mouse IgG (H+L), F(ab&#039;)&lt;sub&gt;2&lt;/sub&gt; Fragment (Alexa Fluor 488 Conjugate)</td><td>cell signaling technology</td><td>4408</td><td>immunofluoresecence dilution 1:400</td></tr><tr><td>CF568 Phalloidin</td><td>biotium</td><td>00044</td><td>immunofluoresecence dilution 1:400</td></tr><tr><td>Cyclohexmide</td><td>Sigma</td><td>C1988-1G</td><td>50&amp;micro;g/ml final concentration</td></tr><tr><td>DAPI (4&#039;,6-Diamidino-2-Phenylindole, Dihydrochloride)</td><td>Invitrogen</td><td>D1306</td><td>final concentration 1&amp;micro;g/ml</td></tr><tr><td>Puromycin</td><td>bio-Basic</td><td>PJ593</td><td>2.5&amp;micro;g/ml to 10&amp;micro;g/ml</td></tr><tr><td>Ibidi &amp;micro;-Dish 35 mm, high, ibiTreat</td><td>Ibidi</td><td>81156</td><td></td></tr><tr><td>MRC-5 cells</td><td>ATCC</td><td>CCL-171</td><td></td></tr><tr><td>HeLa cells</td><td>ATCC</td><td>CCL-2</td><td></td></tr><tr><td>Huh-7 cells</td><td>from Dr. Mazroui (Universit&amp;eacute; Laval)</td><td></td><td></td></tr><tr><td>Fv1000</td><td>olympus</td><td></td><td>confocal imaging system</td></tr><tr><td>Fiji software</td><td></td><td></td><td>http://fiji.sc</td></tr><tr><td>PBS (Phosphate BuffeRed Saline)</td><td>bio-Basic</td><td>PD8117</td><td></td></tr><tr><td>Formaldehyde 37% Solution</td><td>bio-Basic</td><td>C5300-1</td><td></td></tr><tr><td>Triton X-100</td><td>bio-Basic</td><td>TB0198</td><td></td></tr><tr><td>BSA</td><td>Fisher Bioreagents</td><td>BP9702-100</td><td></td></tr><tr><td>Tween20</td><td>Fisher Bioreagents</td><td>BP337-500</td><td></td></tr></tbody></table>

quantitative immunofluorescence to measure global localized translation

The mechanisms regulating mRNA translation are involved in various biological processes, such as germ line development, cell differentiation, and organogenesis, as well as in multiple diseases. Numerous publications have convincingly shown that specific mechanisms tightly regulate mRNA translation. Increased interest in the translation-induced regulation of protein expression has led to the development of novel methods to study and follow de novo protein synthesis in cellulo. However, most of these methods are complex, making them costly and often limiting the number of mRNA targets that can be studied. This manuscript proposes a method that requires only basic reagents and a confocal fluorescence imaging system to measure and visualize the changes in mRNA translation that occur in any cell line under various conditions. This method was recently used to show localized translation in the subcellular structures of adherent cells over a short period of time, thus offering the possibility of visualizing de novo translation for a short period during a variety of biological processes or of validating changes in translational activity in response to specific stimuli.

To accurately observe translation events using puromycin incorporation, it is critical to determine the optimal conditions for each cell line because each shows different puromycin incorporation kinetics (Figure 1)9,11,12,18. Hence, to validate puromycin incorporation, it is necessary to treat the desired cell line with a standardiz...

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Quantitative Immunofluorescence to Measure Global Localized Translation

This manuscript describes a method to visualize and quantify localized translation events in subcellular compartments. The approach proposed in this manuscript requires a basic confocal imaging system and reagents and is rapid and cost-effective.

The mechanisms regulating mRNA translation are involved in various biological processes, such as germ line development, cell differentiation, and ...

quantitative-immunofluorescence-to-measure-global-localized

Research

JoVE Journal

Biochemistry

9.8K Views.  Université Laval. This manuscript describes a method to visualize and quantify localized translation events in subcellular compartments. The approach proposed in this manuscript requires a basic confocal imaging system and reagents and is rapid and cost-effective.

Video: Quantitative Immunofluorescence to Measure Global Localized Translation

Isolation and Quantitative Immunocytochemical Characterization of Primary Myogenic Cells and Fibroblasts from Human Skeletal Muscle

The repair and regeneration of skeletal muscle requires the action of satellite cells, which are the resident muscle stem cells. These can be isolated from human muscle biopsy samples using enzymatic digestion and their myogenic properties studied in culture. Quantitatively, the two main adherent cell types obtained from enzymatic digestion are: (i) the satellite cells (termed myogenic cells or muscle precursor cells), identified initially as CD56+ and later as CD56+/desmin+ cells and (ii) muscle-derived fibroblasts, identified as CD56&#8211; and TE-7+. Fibroblasts proliferate very efficiently in culture and in mixed cell populations these cells may overrun myogenic cells to dominate the culture. The isolation and purification of different cell types from human muscle is thus an important methodological consideration when trying to investigate the innate behavior of either cell type in culture. Here we describe a system of sorting based on the gentle enzymatic digestion of cells using collagenase and dispase followed by magnetic activated cell sorting (MACS) which gives both a high purity (&#62;95% myogenic cells) and good yield (~2.8 x 106 &#177; 8.87 x 105 cells/g tissue after 7 days in vitro) for experiments in culture. This approach is based on incubating the mixed muscle-derived cell population with magnetic microbeads beads conjugated to an antibody against CD56 and then passing cells though a magnetic field. CD56+ cells bound to microbeads are retained by the field whereas CD56&#8211; cells pass unimpeded through the column. Cell suspensions from any stage of the sorting process can be plated and cultured. Following a given intervention, cell morphology, and the expression and localization of proteins including nuclear transcription factors can be quantified using immunofluorescent labeling with specific antibodies and an image processing and analysis package.

The main adherent cell types derived from human muscle are myogenic cells and fibroblasts. Here, cell populations are enriched using magnetic-activated cell sorting based on the CD56 antigen. Subsequent immunolabelling with specific antibodies and use of image analysis techniques allows quantification of cytoplasmic and nuclear characteristics in individual cells.

The repair and regeneration of skeletal muscle requires the action of satellite cells, which are the resident muscle stem cells. These can be isolated ...

Developmental Biology

Use of Single Molecule Fluorescent In Situ Hybridization (SM-FISH) to Quantify and Localize mRNAs in Murine Oocytes

Current methods routinely used to quantify mRNA in oocytes and embryos include digital reverse-transcription polymerase chain reaction (dPCR), quantitative, real-time RT-PCR (RT-qPCR) and RNA sequencing. When these techniques are performed using a single oocyte or embryo, low-copy mRNAs are not reliably detected. To overcome this problem, oocytes or embryos can be pooled together for analysis; however, this often leads to high variability amongst samples. In this protocol, we describe the use of fluorescence in situ hybridization (FISH) using branched DNA chemistry. This technique identifies the spatial pattern of mRNAs in individual cells. When the technique is coupled with Spot Finding and Tracking&#160;computer software, the abundance of mRNAs in the cell can also be quantified. Using this technique, there is reduced variability within an experimental group and fewer oocytes and embryos are required to detect significant differences between experimental groups. Commercially available branched-DNA SM-FISH kits have been optimized to detect mRNAs in sectioned tissues or adherent cells on slides. However, oocytes do not effectively adhere to slides and some reagents in the kit were too harsh resulting in oocyte lysis. To prevent this lysis, several modifications were made to the FISH kit. Specifically, oocyte permeabilization and wash&#160;buffers designed for the immunofluorescence of oocytes and embryos replaced the proprietary buffers. The permeabilization, washes, and incubations with probes and amplifier were performed in 6-well plates and oocytes were placed on slides at the end of the protocol using mounting media. These modifications were able to overcome the limitations of the commercially available kit, in particular, the oocyte lysis. To accurately and reproducibly count the number of mRNAs in individual oocytes, computer software was used. Together, this protocol represents an alternative to PCR and sequencing to compare the expression of specific transcripts in single cells.

Use of Single Molecule Fluorescent In Situ Hybridization SM-FISH to Quantify and Localize mRNAs in Murine Oocytes

To reproducibly count the numbers of mRNAs in individual oocytes, single molecule RNA fluorescence in situ hybridization (RNA-FISH) was optimized for non-adherent cells. Oocytes were collected, hybridized with the transcript specific probes, and quantified using an image quantification software.

Current methods routinely used to quantify mRNA in oocytes and embryos include digital reverse-transcription polymerase chain reaction (dPCR), ...

Genetics

An In Vitro Single-Molecule Imaging Assay for the Analysis of Cap-Dependent Translation Kinetics

Cap-dependent protein synthesis is the predominant translation pathway in eukaryotic cells. While various biochemical and genetic approaches have allowed extensive studies of cap-dependent translation and its regulation, high resolution kinetic characterization of this translation pathway is still lacking. Recently, we developed an in vitro assay to measure cap-dependent translation kinetics with single-molecule resolution. The assay is based on fluorescently labeled antibody binding to nascent epitope-tagged polypeptide. By imaging the binding and dissociation of antibodies to and from nascent peptide&#8211;ribosome&#8211;mRNA complexes, the translation progression on individual mRNAs can be tracked. Here, we present a protocol for establishing this assay, including mRNA and PEGylated slide preparations, real-time imaging of translation, and analysis of single molecule trajectories. This assay enables tracking of individual cap-dependent translation events and resolves key translation kinetics, such as initiation and elongation rates. The assay can be widely applied to distinct translation systems and should broadly benefit in vitro studies of cap-dependent translation kinetics and translational control mechanisms.

An In Vitro Single-Molecule Imaging Assay for the Analysis of Cap-Dependent Translation Kinetics

Tracking individual translation events allows for high-resolution kinetic studies of cap-dependent translation mechanisms. Here we demonstrate an in vitro single-molecule assay based on imaging interactions between fluorescently labeled antibodies and epitope-tagged nascent peptides. This method enables single-molecule characterization of initiation and peptide elongation kinetics during active in vitro cap-dependent translation.

Cap-dependent protein synthesis is the predominant translation pathway in eukaryotic cells. While various biochemical and genetic approaches have allowed ...

Quantitative Fluorescence In Situ Hybridization (FISH) and Immunofluorescence (IF) of Specific Gene Products in KSHV-Infected Cells

Mechanistic insight arrives from careful study and quantification of specific RNAs and proteins. The relative locations of these biomolecules throughout the cell at specific times can be captured with fluorescence in situ hybridization (FISH) and immunofluorescence (IF). During lytic herpesvirus infection, the virus hijacks the host cell to preferentially express viral genes, causing changes in cell morphology and behavior of biomolecules. Lytic activities are centered in nuclear factories, termed viral replication compartments, which are discernable only with FISH and IF. Here we describe an adaptable protocol of RNA FISH and IF techniques for Kaposi's sarcoma-associated herpesvirus (KSHV)-infected cells, both adherent and in suspension. The method includes steps for the development of specific anti-sense oligonucleotides, double RNA FISH, RNA FISH with IF, and quantitative calculations of fluorescence intensities. This protocol has been successfully applied to multiple cell types, uninfected cells, latent cells, lytic cells, time-courses, and cells treated with inhibitors to analyze the spatiotemporal activities of specific RNAs and proteins from both the human host and KSHV.

Quantitative Fluorescence In Situ Hybridization FISH and Immunofluorescence IF of Specific Gene Products in KSHV-Infected Cells

We describe a protocol utilizing fluorescence in situ hybridization (FISH) to visualize multiple herpesviral RNAs within lytically infected human cells, either in suspension or adherent. This protocol includes quantification of fluorescence producing a nucleocytoplasmic ratio and can be extended for simultaneous visualization of host and viral proteins with immunofluorescence (IF).

Mechanistic insight arrives from careful study and quantification of specific RNAs and proteins. The relative locations of these biomolecules throughout ...

Immunology and Infection

Visualization of Endoplasmic Reticulum Localized mRNAs in Mammalian Cells

In eukaryotes, most of the messenger RNAs (mRNAs) that encode secreted and membrane proteins are localized to the surface of the endoplasmic reticulum (ER). However, the visualization of these mRNAs can be challenging. This is especially true when only a fraction of the mRNA is ER-associated and their distribution to this organelle is obstructed by non-targeted (i.e. "free") transcripts. In order to monitor ER-associated mRNAs, we have developed a method in which cells are treated with a short exposure to a digitonin extraction solution that selectively permeabilizes the plasma membrane, and thus removes the cytoplasmic contents, while simultaneously maintaining the integrity of the ER. When this method is coupled with fluorescent in situ hybridization (FISH), one can clearly visualize ER-bound mRNAs by fluorescent microscopy. Using this protocol the degree of ER-association for either bulk poly(A) transcripts or specific mRNAs can be assessed and even quantified. In the process, one can use this assay to investigate the nature of mRNA-ER interactions.

Here we describe a method to visualize endoplasmic reticulum-associated mRNAs in mammalian tissue culture cells. This technique involves the selective permeabilization of the plasma membrane with digitonin to remove cytoplasmic contents followed by fluorescent in situ hybridization to detect either bulk poly(A) mRNA or specific transcripts.

In eukaryotes, most of the messenger RNAs (mRNAs) that encode secreted and membrane proteins are localized to the surface of the endoplasmic reticulum ...

Biology

Quantifying the Heterogeneous Distribution of a Synaptic Protein in the Mouse Brain Using Immunofluorescence

The presence, absence, or levels of specific synaptic proteins can severely influence synaptic transmission. In addition to elucidating the function of a protein, it is vital to also determine its distribution. Here, we describe a protocol employing immunofluorescence, confocal microscopy, and computer-based analysis to determine the distribution of the synaptic protein Mover (also called TPRGL or SVAP30). We compare the distribution of Mover to that of the synaptic vesicle protein synaptophysin, thereby determining the distribution of Mover in a quantitative manner relative to the abundance of synaptic vesicles. Notably, this method could potentially be implemented to allow for comparison of the distribution of proteins using different antibodies or microscopes or across different studies. Our method circumvents the inherent variability of immunofluorescent stainings by yielding a ratio rather than absolute fluorescence levels. Additionally, the method we describe enables the researcher to analyze the distribution of a protein on different levels: from whole brain slices to brain regions to different subregions in one brain area, such as the different layers of the hippocampus or sensory cortices. Mover is a vertebrate-specific protein that is associated with synaptic vesicles. With this method, we show that Mover is heterogeneously distributed across brain areas, with high levels in the ventral pallidum, the septal nuclei, and the amygdala, and also within single brain areas, such as the different layers of the hippocampus.

Here, we describe a quantitative approach to determining the distribution of a synaptic protein relative to a marker protein using immunofluorescence staining, confocal microscopy, and computer-based analysis.

The presence, absence, or levels of specific synaptic proteins can severely influence synaptic transmission. In addition to elucidating the function of a ...

Neuroscience

Workflow for High-content, Individual Cell Quantification of Fluorescent Markers from Universal Microscope Data, Supported by Open Source Software

Advances in understanding the control mechanisms governing the behavior of cells in adherent mammalian tissue culture models are becoming increasingly dependent on modes of single-cell analysis. Methods which deliver composite data reflecting the mean values of biomarkers from cell populations risk losing subpopulation dynamics that reflect the heterogeneity of the studied biological system. In keeping with this, traditional approaches are being replaced by, or supported with, more sophisticated forms of cellular assay developed to allow assessment by high-content microscopy. These assays potentially generate large numbers of images of fluorescent biomarkers, which enabled by accompanying proprietary software packages, allows for multi-parametric measurements per cell. However, the relatively high capital costs and overspecialization of many of these devices have prevented their accessibility to many investigators.
Described here is a universally applicable workflow for the quantification of multiple fluorescent marker intensities from specific subcellular regions of individual cells suitable for use with images from most fluorescent microscopes. Key to this workflow is the implementation of the freely available Cell Profiler software1&#160;to distinguish individual cells in these images, segment them into defined subcellular regions and deliver fluorescence marker intensity values specific to these regions. The extraction of individual cell intensity values from image data is the central purpose of this workflow and will be illustrated with the analysis of control data from a siRNA screen for G1 checkpoint regulators in adherent human cells. However, the workflow presented here can be applied to analysis of data from other means of cell perturbation (e.g., compound screens) and other forms of fluorescence based cellular markers and thus should be useful for a wide range of laboratories.

Presented is a flexible informatics workflow enabling multiplexed image-based analysis of fluorescently labeled cells. The workflow quantifies nuclear and cytoplasmic markers and computes marker translocation between these compartments. Procedures are provided for perturbation of cells using siRNA and reliable methodology for marker detection by indirect immunofluorescence in 96-well formats.

Advances in understanding the control mechanisms governing the behavior of cells in adherent mammalian tissue culture models are becoming increasingly ...

Quantitative Immunofluorescence Assay to Measure the Variation in Protein Levels at Centrosomes

Centrosomes are small but important organelles that serve as the poles of mitotic spindle to maintain genomic integrity or assemble primary cilia to facilitate sensory functions in cells. The level of a protein may be regulated differently at centrosomes than at other .cellular locations, and the variation in the centrosomal level of several proteins at different points of the cell cycle appears to be crucial for the proper regulation of centriole assembly. We developed a quantitative fluorescence microscopy assay that measures relative changes in the level of a protein at centrosomes in fixed cells from different samples, such as at different phases of the cell cycle or after treatment with various reagents. The principle of this assay lies in measuring the background corrected fluorescent intensity corresponding to a protein at a small region, and normalize that measurement against the same for another protein that does not vary under the chosen experimental condition. Utilizing this assay in combination with BrdU pulse and chase strategy to study unperturbed cell cycles, we have quantitatively validated our recent observation that the centrosomal pool of VDAC3 is regulated at centrosomes during the cell cycle, likely by proteasome-mediated degradation specifically at centrosomes.

Here, a novel quantitative fluorescence assay is developed to measure changes in the level of a protein specifically at centrosomes by normalizing that protein&#8217;s fluorescence intensity to that of an appropriate internal standard.

Centrosomes are small but important organelles that serve as the poles of mitotic spindle to maintain genomic integrity or assemble primary cilia to ...

Quantitative Localization of a Golgi Protein by Imaging Its Center of Fluorescence Mass

The Golgi complex consists of serially stacked membrane cisternae which can be further categorized into sub-Golgi regions, including the cis-Golgi, medial-Golgi, trans-Golgi and trans-Golgi network. Cellular functions of the Golgi are determined by the characteristic distribution of its resident proteins. The spatial resolution of conventional light microscopy is too low to resolve sub-Golgi structure or cisternae. Thus, the immuno-gold electron microscopy is a method of choice to localize a protein at the sub-Golgi level. However, the technique and instrument are beyond the capability of most cell biology labs. We describe here our recently developed super-resolution method called Golgi protein localization by imaging centers of mass (GLIM) to systematically and quantitatively localize a Golgi protein. GLIM is based on standard fluorescence labeling protocols and conventional wide-field or confocal microscopes. It involves the calibration of chromatic-shift aberration of the microscopic system, the image acquisition and the post-acquisition analysis. The sub-Golgi localization of a test protein is quantitatively expressed as the localization quotient. There are four main advantages of GLIM; it is rapid, based on conventional methods and tools, the localization result is quantitative, and it affords ~ 30 nm practical resolution along the Golgi axis. Here we describe the detailed protocol of GLIM to localize a test Golgi protein.

The precise localization of Golgi residents is essential for understanding the cellular functions of the Golgi. However, conventional optical microscopy is unable to resolve the sub-Golgi structure. Here we describe the protocol for a conventional microscopy based super-resolution method to quantitatively determine the sub-Golgi localization of a protein.

The Golgi complex consists of serially stacked membrane cisternae which can be further categorized into sub-Golgi regions, including the cis-Golgi, ...

Quantitative Immunoblotting of Cell Lines as a Standard to Validate Immunofluorescence for Quantifying Biomarker Proteins in Routine Tissue Samples

Quantification of proteins of interest in formalin-fixed, paraffin-embedded (FFPE) tissue samples is important in clinical and research applications. An optimal method of quantification is accurate, has a broad linear dynamic range and maintains the structural integrity of the sample to allow for identification of individual cell types. Current methods such as immunohistochemistry (IHC), mass spectrometry, and immunoblotting each fail to meet these stipulations due to their categorical nature or need to homogenize the sample. As an alternative method, we propose the use of immunofluorescence (IF) and image analysis to determine the relative abundance of a protein of interest in FFPE tissues. Herein we demonstrate that this method is easily optimized, yields a wide dynamic range, and is linearly quantifiable as compared to the gold standard of quantitative immunoblotting. Furthermore, this method permits the maintenance of the structural integrity of the sample and allows for the distinction of various cell types, which may be crucial in diagnostic applications. Overall, this is a robust method for the relative quantification of proteins in FFPE samples and can be easily adapted to suit clinical or research needs.

We describe the use of quantitative immunoblotting to validate immunofluorescence histology coupled with image analysis as a means of quantifying a protein of interest in formalin-fixed, paraffin-embedded (FFPE) tissue samples. Our results demonstrate the utility of immunofluorescence histology for ascertaining the relative quantity of biomarker proteins in routine biopsy samples.

Quantification of proteins of interest in formalin-fixed, paraffin-embedded (FFPE) tissue samples is important in clinical and research applications. An ...

Quantitative Immunofluorescence to Measure Global Localized Translation

Summary

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Chapters in this video

Isolation and Quantitative Immunocytochemical Characterization of Primary Myogenic Cells and Fibroblasts from Human Skeletal Muscle

Use of Single Molecule Fluorescent In Situ Hybridization (SM-FISH) to Quantify and Localize mRNAs in Murine Oocytes

An In Vitro Single-Molecule Imaging Assay for the Analysis of Cap-Dependent Translation Kinetics

Quantitative Fluorescence In Situ Hybridization (FISH) and Immunofluorescence (IF) of Specific Gene Products in KSHV-Infected Cells

Visualization of Endoplasmic Reticulum Localized mRNAs in Mammalian Cells

Quantifying the Heterogeneous Distribution of a Synaptic Protein in the Mouse Brain Using Immunofluorescence

Workflow for High-content, Individual Cell Quantification of Fluorescent Markers from Universal Microscope Data, Supported by Open Source Software

Quantitative Immunofluorescence Assay to Measure the Variation in Protein Levels at Centrosomes

Quantitative Localization of a Golgi Protein by Imaging Its Center of Fluorescence Mass

Quantitative Immunoblotting of Cell Lines as a Standard to Validate Immunofluorescence for Quantifying Biomarker Proteins in Routine Tissue Samples