This work was supported by grants from the National Science and Engineering Research Council of Canada to X.A.C. and A.F.P.

1. Preparation of Materials for Extraction
<ol>
	<li>Preparation of Cells
	<ol>
		<li>Seed tissue culture cells on acid-treated coverslips for at least one day prior to the experiment. We use COS-7 or U2OS, as these two cell lines exhibit robust production of secreted protein and have a well defined ER. Note that to achieve high extraction efficiency, cells should not exceed 80% confluency on the coverslip on the day of the experiment.</li>
		<li>If a particular exogenous mRNA is being investigated, cells are transfected one day after seeding with plasmids containing the gene of interest using standard transfection protocols. The cells are then allowed to express the mRNA for at least 18 hr prior to extraction.</li>
	</ol>
	</li>
	<li>Treating the Cells with Translation Inhibitors
	<ol>
		<li>To test the effect of intact ribosomes on the maintenance of mRNAs association with the ER, cells can be treated with translation inhibitors that disrupt the association of ribosomes with the mRNA11.</li>
		<li>Inhibitors of translation initiation, such as homoharringtonin (HHT), pactamycin, or 2-(4-methyl-2,6-dinitroanilino)-N-methylpropionamide (MDMP), prevent new ribosomes from associating with the transcript while simultaneously allowing engaged ribosomes to complete translation12-14. Since the translation rate for most proteins in mammalian cells is 5 amino acids per second, regardless of codon usage or mRNA length15, a treatment of 30 min should clear ribosomes off of messages with open reading frames as long as 27,000 nucleotides (coding for proteins as long as 9,000 amino acids).</li>
		<li>Inhibitors such as puromycin promote the premature ejection of the nascent chain and thus disrupt polysomes12. However to completely disrupt the association of puromycin-treated ribosomes with the mRNA, magnesium chelators such as EDTA must be added to the digitonin extraction buffer11.</li>
		<li>In this experiment, cells are treated with 200 &mu;M puromycin, 5 &mu;M HHT, or control medium for 30 min prior to extraction.</li>
	</ol>
	</li>
	<li>Preparation of Digitonin Extraction Buffer. To visualize ER-localized mRNAs without the obstruction of free (i.e. cytoplasmic, non-ER-associated) transcripts, we selectively permeabilize the cell with digitonin, a steroid glycoside that interacts preferentially with 3&beta;-hydroxysterols. At low concentrations, digitonin selectively permeabilizes portions of the plasma membrane, causing 'leakiness'16. In contrast, the ER membrane and nuclear envelope, which are poor in cholesterol, are left intact16,17.
	<ol>
		<li>Digitonin (Sigma Aldrich) powder is dissolved in distilled water at 5% weight/volume and this stock solution is aliquoted into small volumes and stored in -20 &deg;C. In our experience, multiple freeze-thaw cycles decreases the efficiency of dissolved digitonin.</li>
		<li>A stock solution of 10x CHO buffer (1x working solution concentration: 115 mM KAc, 25 mM HEPES pH 7.4, 2.5 mM MgCl2, 2 mM EGTA and 150 mM Sucrose) is prepared with RNase-free reagents and stored at -20 &deg;C. A working solution (1x CHO buffer) is prepared by diluting the stock solution with RNase-free water and can be stored at 4 &deg;C.</li>
	</ol>
	</li>
</ol>
2. Digitonin Extraction
<ol>
	<li>Preparing Extraction Solutions
	<ol>
		<li>A heating block with a flat surface is pre-heated to 40 &deg;C and maintained at this temperature during to the extraction.</li>
		<li>To form a flat working surface that is RNase free, the heating block is moistened with some water and overlaid with a fresh piece of parafilm M (Bemis Company, Inc). Air bubbles between the parafilm sheet and heating block are smoothed out using RNase-free gloves and kimwipes (VWR).</li>
		<li>1x CHO buffer is warmed to 37 &deg;C by incubating in a water bath.</li>
		<li>6-well plates are used to wash and fix the cells. Each row of 3 wells is used for one cover-slip. To the first two wells in the row, 2 ml of warmed CHO buffer is added. To the last well, 2 ml of the fixation solution (PBS + 4% paraformaldehyde (Electron Microscope Sciences)) is added. This tray can be stored in the 37 &deg;C incubator until the cells are ready for extraction.</li>
		<li>Digitonin extraction solution is prepared by diluting the stock digitonin solution to 0.025% with warm CHO buffer just prior to use. Again note that for puromycin-treated cells, the extraction buffer must additionally contain 20 mM EDTA to efficiently disrupt ribosomes11. Drops of 100 &mu;l of the extraction solution are placed on the parafilm-covered heating block immediately before extraction.</li>
	</ol>
	</li>
	<li>Digitonin Extraction and Cell Fixation
	<ol>
		<li>Remove the cells from the 37 &deg;C incubator.</li>
		<li>During the extraction process, coverslips are manipulated with the help of jewler's forceps. With the forceps pick up the coverslip and dip it into the first and then the second well containing CHO buffer to wash of the growth medium.</li>
		<li>Quickly blot off excess buffer off the back side of the coverslip with a kimwipe and immediately place the coverslip, cell side facing down, on the extraction solution.</li>
		<li>After 10 sec, pick up the coverslip with the forceps and immediately transfer it, cell side facing up, to the well containing the 4% paraformaldehyde fixing buffer.</li>
		<li>Let the sample incubate in fixative for at least 15 min at room temperature.</li>
	</ol>
	</li>
	<li>Preparation of Unextracted Control Cells. To determine the total amount of mRNA in the cytoplasm it is a good idea to prepare an unextracted control sample.
	<ol>
		<li>Cells grown on coverslips are prepared as above (see section 2.2), except that the digitonin extraction step (2.2.3) is skipped.</li>
		<li>After fixation, the un-extracted cells need to be permeabilized in PBS + 0.1% Triton X-100 for 15 min at room temperature.</li>
	</ol>
	</li>
</ol>
3. FISH Staining
<ol>
	<li>Designing probes for fluorescence in situ hybridization
	<ol>
		<li>The primary sequence of the mRNA of interested is folded using RNA secondary structure prediction software, such as RNAstructure 5.018. The mRNA folds are visually assessed to identify a region of about 50 nucleotides that is free of major secondary structures.</li>
		<li>The reverse complement of this region is synthesized with an Alexa546 fluorophore attached to the 5' end of the probe (purchased from Integrated DNA Technologies).</li>
		<li>The probe is diluted with RNase-free water to a stock concentration of 100 &mu;M which can be stored at -20 &deg;C.</li>
	</ol>
	</li>
	<li>Staining with FISH probes
	<ol>
		<li>Note that although the digitonin-extracted cells do not require further permeabilization, treating these fixed samples with 2 ml of 0.1% Triton X-100 diluted in PBS for 15 min at room temperature prior to FISH staining, helps to reduce the background signal.</li>
		<li>The slides are then washed twice with PBS to remove any residual paraformaldehyde or Triton X-100.</li>
		<li>The cells are then washed twice with 25% to 60% formamide in 1X Sodium Citrate Buffer (150 mM NaCl and 15 mM sodium citrate). Generally, for specific FISH probes that are 50-60 nucleotides in length, 60% formamide is used, but for poly(A) mRNA staining with oligo(dT) FISH probe, the level of formamide is reduced down to 25%.</li>
		<li>To prepare the staining chamber, the bottom of a 150 mm Petri dish is covered with water and a piece of parafilm is floated on the water. The water is removed by turning the dish over, thus allowing the parafilm to adhere to the bottom of the dish. Air bubbles are manually removed with kimwipes.</li>
		<li>For each coverslip to be stained, pipette a 100 &mu;l drop of the hybridization solution containing FISH probe of interest onto the parafilm in the staining chamber. The stock FISH probe is diluted with 25% to 60% formamide in hybridization buffer (1x SSC, 100 mg/ml dextran sulphate, 1 mg/ml yeast tRNA, 5mM vanadyl riboside complex, 25-60% formamide) to a working concentration of 0.2 &mu;M. Note that the amount of formamide should be optimized for the particular probe being used and is present at the same concentration as in the SSC wash buffer (see step 3.3.3).</li>
		<li>The coverslip is placed cell side face down onto the FISH solution in the staining chamber and incubated for 5-24 hr at 37 &deg;C in a humidified chamber such as the tissue culture incubator.</li>
	</ol>
	</li>
	<li>Washing and mounting the stained samples
	<ol>
		<li>To prepare an RNase free washing area, lay down a strip of parafilm on a level flat surface, such as a lab bench. To ensure that the parafilm is flat, wet the level surface, place the parafilm on top and manually remove any air bubbles with kimwipes.</li>
		<li>The staining chamber is removed from the incubator and placed on a flat level surface. The coverslips are then floated by pipetting approximately 500 &mu;l of FISH wash buffer near the edge of each coverslip. The solution will be drawn in-between the coverslip and the parafilm via capillary action.</li>
		<li>For each coverslip, 1 ml of wash buffer (1x SCC with 25-60% formamide, see step 3.3.3) is pipetted onto the washing area parafilm (see step 3.4.1).</li>
		<li>Using forceps, the coverslips are removed from the staining chamber and each is placed onto the drop of wash buffer and incubated at room temperature for 5 min. The washing process is repeated 3 times.</li>
		<li>Glass slides are washed with 70% ethanol, and dried with kimwipes.</li>
		<li>For each coverslip, 25 &mu;l of mounting solution (DAPI Fluoromount G, Southern Biotech) is pipetted onto the slide. Note that this solution contains 4',6-diamidino-2-phenylindole (DAPI), which stains DNA and allows for the visualization of nuclei.</li>
		<li>Using forceps and kimwipes, the back of the coverslip is dried and excess FISH wash buffer is blotted off without drying the sample. Each coverslip is then placed cell side facing down, onto the drop of mounting solution.</li>
		<li>Mounted samples are then labeled and can be store at 4 &deg;C.</li>
	</ol>
	</li>
</ol>
4. Imaging and Quantification
<ol>
	<li>Imaging
	<ol>
		<li>An epifluorescence microscope is used to image the cells following FISH staining. The transfected cell can be differentiated from untransfected cells by bright fluorescence signal in the appropriate channel. Ideally, each acquired field will also contain an untransfected cell to be used to determine the background fluorescence for quantification.</li>
		<li>For each field images of the FISH and DAPI channel are acquired. In addition, if the cells are co-stained with protein markers, an image of the immunofluorescence channel is also acquired. Note that digitonin extraction can also be used to visualize ER-bound ribosomes and proteins11.</li>
		<li>To ensure that the fluorescence intensities between fields can be compared, the exposure time for each channel should remain constant for each set of experiment.</li>
		<li>Again to ensure that the fluorescent intensities between cells on different coverslips can be compared, the day-to-day changes in epiluminescence intensity or decay in the FISH signal should be minimized by imaging all of the coverslips in a single sitting.</li>
	</ol>
	</li>
	<li>Quantification. To quantify the amount of mRNA that is localized on the ER, we use Nikon NIS Element software. However other image analysis software such as ImageJ can be used.
	<ol>
		<li>Using the Nikon NIS ELEMENT image analysis tool, the cell periphery and the nucleus are outlined as separate Region of Interests (ROIs).</li>
		<li>For each ROI, the fluorescent intensity (IT for the total cell intensity and IN for the nuclear intensity) and the area (AT and AN) are recorded and exported to an excel spreadsheet.</li>
		<li>For each image, the background fluorescent intensity (IB) is determined by recording the intensity of a non-transfected cell. If poly(A) mRNA levels are being analyzed, a cell-free area is selected.</li>
		<li>The total amount of ER (in extracted cells) or cytoplasmic (in unextracted cells) fluorescence is calculated by the formula: 
		[AT][ IT - IB]- [AN][ IN - IB]</li>
		<li>The nuclear staining intensity can also be calculated and used as internal control between various treatments. Since nuclei are not affected by digitonin treatment17 or ribosome disruption11, the amount of nuclear mRNA should remain constant between each of these treatments. To calculate nuclear fluorescence the following formula is used: 
		[AN][ IN - IB]</li>
		<li>The percent of cytoplasmic mRNA that is ER-targeted can be calculated by taking the average ER fluorescence of 30-40 extracted cells (see 4.2.4) divided by the average cytoplasmic fluorescence from 30-40 unextracted cells (again see 4.2.4). It is critical that the extracted and unextracted cells are prepared, stained and imaged in parallel.</li>
	</ol>
	</li>
</ol>

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No conflicts of interest declared.

The localization of mRNAs to various subcellular sites, through the interaction of transcripts with mRNA localization proteins, is a widespread phenomenon important for the proper sorting of proteins to their final destination, and for the fine tuning of gene expression to the local requirements of a subcellular region19,20.
Using the assay described here, we revisited the question of whether mRNAs that encode secretory proteins can be maintained on the ER in the presence or absence...

In eukaryotes, mRNA encoding secreted and membrane proteins can be targeted to the ER co-translationally by the signal recognition particle1,2 and can be maintained on the ER via the direct interactions between ribosomes and translocons during the translation3,4. However, whether mRNAs can be targeted and maintained on the ER independent of either ribosomes or translation was unclear until very recently. Previous studies attempted to address whether there is translation-independent mRNA association with the ER using cellular fractionation techniques. Because harsh chemical conditions were required to disassociate ribosomes from ER derived vesicles, which might disrupt the potentially delicate mRNA-ER association, these studies were inconclusive, providing evidence for5-8 and against9,10 ribosomal-independent anchoring of mRNA to the ER.
To circumvent these problems we have developed a protocol to isolate and image ER-bound mRNAs. This procedure involves a mild extraction treatment, which effectively removes all the cytoplasmic content of the cell (including non ER-bound mRNAs) while simultaneously preserving the ER morphology and all of its associated molecules. Using this protocol we have demonstrated that a subset of mRNAs are targeted and then maintained on the ER independently of ribosomes or translation11.

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.

To determine the percentage of mRNA that is ER-associated, COS-7 cells that were transfected with plasmids that contained the placental alkaline phosphatase (ALPP) or cytochrome P450-8B1 (CYP8B1) were either extracted with digitonin and then fixed, or directly fixed (see Figure 1, compare "Unextracted Ctrl" to "Extracted Ctrl"). The non-nuclear fluorescence was quantified in both samples and the fraction of ER-associated mRNA was calculated (Figure 2)....

Watch this Scientific Journal Video about Visualization of Endoplasmic Reticulum Localized mRNAs in Mammalian Cells at JoVE.com

Visualization of Endoplasmic Reticulum Localized mRNAs in Mammalian Cells

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 ...

visualization-endoplasmic-reticulum-localized-mrnas-mammalian

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Biology

14.2K 조회수.  University of Toronto. 여기 포유류의 조직 배양 세포에서 endoplasmic 소포체 - 관련 mRNAs을 시각화하는 방법을 설명합니다. 이 기술은 형광등에 이어 세포질 내용을 제거 할 digitonin과 플라즈마 막의 선택적 permeabilization을 포함 현장에서 하이브리드.

비디오: Visualization of Endoplasmic Reticulum Localized mRNAs in Mammalian Cells

Trypsinizing and Subculturing Mammalian Cells

As cells reach confluency, they must be subcultured or passaged. Failure to subculture confluent cells results in reduced mitotic index and eventually in cell death. The first step in subculturing is to detach cells from the surface of the primary culture vessel by trypsinization or mechanical means. The resultant cell suspension is then subdivided, or reseeded, into fresh cultures. Secondary cultures are checked for growth and fed periodically, and may be subsequently subcultured to produce tertiary cultures. The time between passaging of cells varies with the cell line and depends on the growth rate.

As cells reach confluency, they must be subcultured or passaged. This video will demonstrate a procedure for subculturing both adherent and suspension cells.

As cells reach confluency, they must be subcultured or passaged. Failure to subculture confluent cells results in reduced mitotic index and eventually in ...

In situ Subcellular Fractionation of Adherent and Non-adherent Mammalian Cells

Protein function is intimately coupled to protein localization. Although some proteins are restricted to a specific location or subcellular compartment, many proteins are present as a freely diffusing population in free exchange with a sub-population that is tightly associated with a particular subcellular domain or structure. In situ subcellular fractionation allows the visualization of protein compartmentalization and can also reveal protein sub-populations that localize to specific structures. For example, removal of soluble cytoplasmic proteins and loosely held nuclear proteins can reveal the stable association of some transcription factors with chromatin. Subsequent digestion of DNA can in some cases reveal association with the network of proteins and RNAs that is collectively termed the nuclear scaffold or nuclear matrix. 
Here we describe the steps required during the in situ fractionation of adherent and non-adherent mammalian cells on microscope coverslips. Protein visualization can be achieved using specific antibodies or fluorescent fusion proteins and fluorescence microscopy. Antibodies and/or fluorescent dyes that act as markers for specific compartments or structures allow protein localization to be mapped in detail. In situ fractionation can also be combined with western blotting to compare the amounts of protein present in each fraction. This simple biochemical approach can reveal associations that would otherwise remain undetected.

In situ Subcellular Fractionation of Adherent and Non-adherent Mammalian Cells

In situ subcellular fractionation of mammalian cells on microscope coverslips allows the visualisation of protein localisation.

Protein function is intimately coupled to protein localization. Although some proteins are restricted to a specific location or subcellular compartment, ...

MISSION esiRNA for RNAi Screening in Mammalian Cells

RNA interference (RNAi) is a basic cellular mechanism for the control of gene expression. RNAi is induced by short double-stranded RNAs also known as small interfering RNAs (siRNAs). The short double-stranded RNAs originate from longer double stranded precursors by the activity of Dicer, a protein of the RNase III family of endonucleases. The resulting fragments are components of the RNA-induced silencing complex (RISC), directing it to the cognate target mRNA. RISC cleaves the target mRNA thereby reducing the expression of the encoded protein1,2,3. RNAi has become a powerful and widely used experimental method for loss of gene function studies in mammalian cells utilizing small interfering RNAs.
Currently two main methods are available for the production of small interfering RNAs. One method involves chemical synthesis, whereas an alternative method employs endonucleolytic cleavage of target specific long double-stranded RNAs by RNase III in vitro. Thereby, a diverse pool of siRNA-like oligonucleotides is produced which is also known as endoribonuclease-prepared siRNA or esiRNA. A comparison of efficacy of chemically derived siRNAs and esiRNAs shows that both triggers are potent in target-gene silencing. Differences can, however, be seen when comparing specificity. Many single chemically synthesized siRNAs produce prominent off-target effects, whereas the complex mixture inherent in esiRNAs leads to a more specific knockdown10.
In this study, we present the design of genome-scale MISSION esiRNA libraries and its utilization for RNAi screening exemplified by a DNA-content screen for the identification of genes involved in cell cycle progression. We show how to optimize the transfection protocol and the assay for screening in high throughput. We also demonstrate how large data-sets can be evaluated statistically and present methods to validate primary hits. Finally, we give potential starting points for further functional characterizations of validated hits.

Here we use a human esiRNA library in a high-throughput screen for genes involved in cell division. We demonstrate how to set up and conduct an esiRNA screens, as well as how to analyze and validate the results.

RNA interference (RNAi) is a basic cellular mechanism for the control of gene expression. RNAi is induced by short double-stranded RNAs also known as ...

Analysis of mRNA Nuclear Export Kinetics in Mammalian Cells by Microinjection

In eukaryotes, messenger RNA (mRNA) is transcribed in the nucleus and must be exported into the cytoplasm to access the translation machinery. Although the nuclear export of mRNA has been studied extensively in Xenopus oocytes1 and genetically tractable organisms such as yeast2 and the Drosophila derived S2 cell line3, few studies had been conducted in mammalian cells. Furthermore the kinetics of mRNA export in mammalian somatic cells could only be inferred indirectly4,5. In order to measure the nuclear export kinetics of mRNA in mammalian tissue culture cells, we have developed an assay that employs the power of microinjection coupled with fluorescent in situ hybridization (FISH). These assays have been used to demonstrate that in mammalian cells, the majority of mRNAs are exported in a splicing dependent manner6,7, or in manner that requires specific RNA sequences such as the signal sequence coding region (SSCR) 6. In this assay, cells are microinjected with either in vitro synthesized mRNA or plasmid DNA containing the gene of interest. The microinjected cells are incubated for various time points then fixed and the sub-cellular localization of RNA is assessed using FISH. In contrast to transfection, where transcription occurs several hours after the addition of nucleic acids, microinjection of DNA or mRNA allows for rapid expression and allows for the generation of precise kinetic data.

Here we describe an assay that employs the power of microinjection coupled with fluorescent in situ hybridization in order to accurately measure the nuclear export kinetics of mRNA in mammalian somatic cells.

In eukaryotes, messenger RNA (mRNA) is transcribed in the nucleus and must be exported into the cytoplasm to access the translation machinery. Although ...

Analysis of Cell Cycle Position in Mammalian Cells

The regulation of cell proliferation is central to tissue morphogenesis during the development of multicellular organisms. Furthermore, loss of control of cell proliferation underlies the pathology of diseases like cancer. As such there is great need to be able to investigate cell proliferation and quantitate the proportion of cells in each phase of the cell cycle. It is also of vital importance to indistinguishably identify cells that are replicating their DNA within a larger population. Since a cell&prime;s decision to proliferate is made in the G1 phase immediately before initiating DNA synthesis and progressing through the rest of the cell cycle, detection of DNA synthesis at this stage allows for an unambiguous determination of the status of growth regulation in cell culture experiments.
DNA content in cells can be readily quantitated by flow cytometry of cells stained with propidium iodide, a fluorescent DNA intercalating dye. Similarly, active DNA synthesis can be quantitated by culturing cells in the presence of radioactive thymidine, harvesting the cells, and measuring the incorporation of radioactivity into an acid insoluble fraction. We have considerable expertise with cell cycle analysis and recommend a different approach. We Investigate cell proliferation using bromodeoxyuridine/fluorodeoxyuridine (abbreviated simply as BrdU) staining that detects the incorporation of these thymine analogs into recently synthesized DNA. Labeling and staining cells with BrdU, combined with total DNA staining by propidium iodide and analysis by flow cytometry1 offers the most accurate measure of cells in the various stages of the cell cycle. It is our preferred method because it combines the detection of active DNA synthesis, through antibody based staining of BrdU, with total DNA content from propidium iodide. This allows for the clear separation of cells in G1 from early S phase, or late S phase from G2/M. Furthermore, this approach can be utilized to investigate the effects of many different cell stimuli and pharmacologic agents on the regulation of progression through these different cell cycle phases.
In this report we describe methods for labeling and staining cultured cells, as well as their analysis by flow cytometry. We also include experimental examples of how this method can be used to measure the effects of growth inhibiting signals from cytokines such as TGF-&beta;1, and proliferative inhibitors such as the cyclin dependent kinase inhibitor, p27KIP1. We also include an alternate protocol that allows for the analysis of cell cycle position in a sub-population of cells within a larger culture5. In this case, we demonstrate how to detect a cell cycle arrest in cells transfected with the retinoblastoma gene even when greatly outnumbered by untransfected cells in the same culture. These examples illustrate the many ways that DNA staining and flow cytometry can be utilized and adapted to investigate fundamental questions of mammalian cell cycle control.

Determining the cell cycle position of a population of cells, or understanding how signals affect proliferation, can be readily measured by flow cytometry using this protocol. We report a simple experimental approach to staining cells and quantifying their position in the cell cycle.

The regulation of cell proliferation is central to tissue morphogenesis during the development of multicellular organisms. Furthermore, loss of control of ...

Visualization of Endoplasmic Reticulum Subdomains in Cultured Cells

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

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

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

Isolation of mRNAs Associated with Yeast Mitochondria to Study Mechanisms of Localized Translation

Most of mitochondrial proteins are encoded in the nucleus and need to be imported into the organelle. Import may occur while the protein is synthesized near the mitochondria. Support for this possibility is derived from recent studies, in which many mRNAs encoding mitochondrial proteins were shown to be localized to the mitochondria vicinity. Together with earlier demonstrations of ribosomes&#8217; association with the outer membrane, these results suggest a localized translation process. Such localized translation may improve import efficiency, provide unique regulation sites and minimize cases of ectopic expression. Diverse methods have been used to characterize the factors and elements that mediate localized translation. Standard among these is subcellular fractionation by differential centrifugation. This protocol has the advantage of isolation of mRNAs, ribosomes and proteins in a single procedure. These can then be characterized by various molecular and biochemical methods. Furthermore, transcriptomics and proteomics methods can be applied to the resulting material, thereby allow genome-wide insights. The utilization of yeast as a model organism for such studies has the advantages of speed, costs and simplicity. Furthermore, the advanced genetic tools and available deletion strains facilitate verification of candidate factors.

Many mRNAs encoding mitochondrial proteins are associated with the mitochondria outer membrane. We describe a subcellular fractionation procedure aimed at isolation of yeast mitochondria with its associated mRNAs and ribosomes. This protocol can be applied to cells grown under diverse conditions&#160;in order to reveal mechanisms of mRNA localization and localized translation near the mitochondria.

Most of mitochondrial proteins are encoded in the nucleus and need to be imported into the organelle. Import may occur while the protein is synthesized ...

Measurement of Heme Synthesis Levels in Mammalian Cells

Heme serves as the prosthetic group for a wide variety of proteins known as hemoproteins, such as hemoglobin, myoglobin and cytochromes. It is involved in various molecular and cellular processes such as gene transcription, translation, cell differentiation and cell proliferation. The biosynthesis levels of heme vary across different tissues and cell types and is altered in diseased conditions such as anemia, neuropathy and cancer. This technique uses [4-14C] 5-aminolevulinic acid ([14C] 5-ALA), one of the early precursors in the heme biosynthesis pathway to measure the levels of heme synthesis in mammalian cells. This assay involves incubation of cells with [14C] 5-ALA followed by extraction of heme and measurement of the radioactivity incorporated into heme. This procedure is accurate and quick. This method measures the relative levels of heme biosynthesis rather than the total heme content. To demonstrate the use of this technique the levels of heme biosynthesis were measured in several mammalian cell lines.

Altered intracellular heme levels are associated with common diseases such as cancer. Thus, there is a need to measure heme biosynthesis levels in diverse cells. The goal of this protocol is to provide a fast and sensitive method to measure and compare the levels of heme synthesis in different cells.

Heme serves as the prosthetic group for a wide variety of proteins known as hemoproteins, such as hemoglobin, myoglobin and cytochromes. It is involved in ...

Massively Parallel Reporter Assays in Cultured Mammalian Cells

The genetic reporter assay is a well-established and powerful tool for dissecting the relationship between DNA sequences and their gene regulatory activities. The potential throughput of this assay has, however, been limited by the need to individually clone and assay the activity of each sequence on interest using protein fluorescence or enzymatic activity as a proxy for regulatory activity. Advances in high-throughput DNA synthesis and sequencing technologies have recently made it possible to overcome these limitations by multiplexing the construction and interrogation of large libraries of reporter constructs. This protocol describes implementation of a Massively Parallel Reporter Assay (MPRA) that allows direct comparison of hundreds of thousands of putative regulatory sequences in a single cell culture dish.

The genetic reporter assay is a well-established and powerful tool for dissecting the relationship between DNA sequences and their gene regulatory activities. Coupling candidate regulatory elements to reporter genes that carry identifying sequence tags enables massive parallelization of these assays.

The genetic reporter assay is a well-established and powerful tool for dissecting the relationship between DNA sequences and their gene regulatory ...

Imaging Local Ca2+ Signals in Cultured Mammalian Cells

Cytosolic Ca2+ ions regulate numerous aspects of cellular activity in almost all cell types, controlling processes as wide-ranging as gene transcription, electrical excitability and cell proliferation. The diversity and specificity of Ca2+ signaling derives from mechanisms by which Ca2+ signals are generated to act over different time and spatial scales, ranging from cell-wide oscillations and waves occurring over the periods of minutes to local transient Ca2+ microdomains (Ca2+ puffs) lasting milliseconds. Recent advances in electron multiplied CCD (EMCCD) cameras now allow for imaging of local Ca2+ signals with a 128 x 128 pixel spatial resolution at rates of &#62;500 frames sec-1 (fps). This approach is highly parallel and enables the simultaneous monitoring of hundreds of channels or puff sites in a single experiment. However, the vast amounts of data generated (ca. 1 Gb per min) render visual identification and analysis of local Ca2+ events impracticable. Here we describe and demonstrate the procedures for the acquisition, detection, and analysis of local IP3-mediated Ca2+ signals in intact mammalian cells loaded with Ca2+ indicators using both wide-field epi-fluorescence (WF) and total internal reflection fluorescence (TIRF) microscopy. Furthermore, we describe an algorithm developed within the open-source software environment Python that automates the identification and analysis of these local Ca2+ signals. The algorithm localizes sites of Ca2+ release with sub-pixel resolution; allows user review of data; and outputs time sequences of fluorescence ratio signals together with amplitude and kinetic data in an Excel-compatible table.

Imaging Local Ca2+ Signals in Cultured Mammalian Cells

Here we present techniques for imaging local IP3-mediated Ca2+ events using fluorescence microscopy in intact mammalian cells loaded with Ca2+ indicators together with an algorithm that automates identification and analysis of these events.

Cytosolic Ca2+ ions regulate numerous aspects of cellular activity in almost all cell types, controlling processes as wide-ranging as gene transcription, ...