The C. elegans strains used in this study were obtained from the Caenorhabditis Genetics Center, which is supported by the National Institutes of Health - Office of Research Infrastructure Programs (P40 OD010440). This work was supported by NIH: 1R03AG048578-01 to Jun Liang, CUNY-CCRG 1501 to Jun Liang, and PSC-CUNY 66184-00 44 and 67248-00 45 to Jun Liang. We thank Cathy Savage-Dunn for kindly sharing her laboratory space. All fluorescence images were collected at Queens College Core Facility. We thank William J. Rice for his comments on the manuscript.

1. Solutions
<ol>
	<li>NGM plates
	<ol>
		<li>In a 2 L Erlenmeyer flask, add 3 g of NaCl, 17 g of agar, 2.5 g of Peptone, and 975 mL of dH2O. Cover the mouth of the flask with aluminum foil. Autoclave the flask for 50 min. Cool it for 20&#8211;30 min.</li>
		<li>Then add sterilized solutions: 1 mL of 1 M CaCl2, 1 mL of 5 mg/mL cholesterol in ethanol, 1 mL of 1 M MgSO4, and 25 mL of 1 M KPO4 (pH 6.0) buffer. Swirl the solutions to mix them well.</li>
		<li>Using a liquid dispenser, dispense the NGM solution to 60 mm Petri plates. Fill the plates 2/3 full of agar. Store them in an air-tight container at 4 &#176;C for future use.</li>
	</ol>
	</li>
	<li>1 M KPO4 buffer (pH 6.0)
	<ol>
		<li>Dissolve 10.83 g of KH2PO4 and 3.56 g of K2HPO4 in dH2O, then adjust the total volume to 100 mL. Autoclave the solution for 15 min.</li>
	</ol>
	</li>
	<li>M9
	<ol>
		<li>Dissolve 3 g of KH2PO4, 6 g of Na2HPO4, and 5 g of NaCl in dH2O, add 1 mL of 1 M MgSO4, and adjust the total volume to 1 L.</li>
	</ol>
	</li>
	<li>10 &#181;g/mL DAPI
	<ol>
		<li>Add 1 &#181;L of 10 mg/mL DAPI into 999 &#181;L of dH2O.</li>
	</ol>
	</li>
	<li>95% alcohol (v/v)
	<ol>
		<li>Measure 95 mL of 100% alcohol and add distilled water to adjust the volume to 100 mL.</li>
	</ol>
	</li>
	<li>30% acetone (v/v)
	<ol>
		<li>Add 30 mL of acetone to dH2O and adjust the total volume to 100 mL.</li>
	</ol>
	</li>
</ol>
2. Heat Stress
<ol>
	<li>Generate an extrachromosomal array line with a translational construct of target genes9,10. Alternatively, obtain such lines from the Caenorhabditis Genetics Center (CGC), which is a public resource for C. elegans research, or from a previously published research laboratory.</li>
	<li>Integrate the extrachromosomal lines into the genome by exposing worms to 3,800 Rad &#947;-radiation9. Then, select the stably expressed lines so that each animal expresses a marker protein such as a GFP or roller.</li>
	<li>To remove mutations generated during the radiation, outcross the lines at least 2x. Use this transgenic line to detect protein expression pattern changes under stress.</li>
	<li>Prepare NGM worm plates as described in step 1.1. Streak an OP50 clone and culture the bacteria in liquid LB broth overnight. Seed the NGM plates with liquid OP50 culture and incubate the plates in a 37 &#176;C incubator overnight to grow a bacteria lawn as a food source for the worms. Cool down the plates at room temperature for at least 30 min before use.</li>
	<li>Grow the transgenic line at 20 &#176;C without starvation for at least 2 generations prior to the heat stress assay.</li>
	<li>Pick 8&#8211;10 young gravid adults (uterus filled with eggs) to a new worm plate, then let them lay eggs for about 3&#8211;5 h and remove all the adults from the plates.</li>
	<li>To synchronize the worms, let the eggs hatch and grow the larvae for approximately 48 h to the fourth larval (L4) stage. Examine their vulva structure (a half-moon structure) to identify the L4 stage (Figure 1, arrow)11.</li>
	<li>Place the worm plates with the L4 larvae in a 35 &#176;C incubator for 2&#8211;5 h to achieve heat stress. Place a thermometer in the incubator to accurately measure the temperature.</li>
</ol>
3. DAPI Staining
<ol>
	<li>Ethanol fixation
	<ol>
		<li>Add 10 &#181;L of the M9 buffer in the center of a glass slide.</li>
		<li>Pick the heat-shocked worms from step 2.8 and put them into the M9 buffer on the glass slide.
		<ol>
			<li>Alternatively, wash off the plate with M9, centrifuge it at 1,000 x g for 1 min at room temperature, and discard the supernatant. Add the worms to the glass slide.</li>
		</ol>
		</li>
		<li>Use a soft tissue to drain any excess liquid. Watch this step under a dissecting microscope.</li>
		<li>Add 10 &#181;L of 95% alcohol onto the worms and let them air dry. Watch the process under a dissecting microscope, and do not let the animals get too dry. Immediately after the ethanol has evaporated from the animals, proceed to the next step. 
		Note: Ethanol evaporates very quickly.</li>
		<li>Repeat step 3.1.4 twice so that the worms are fixed. 
		Note: It is normal for the animals to look darker and distorted.</li>
		<li>Add 10 &#181;L of DAPI (10 &#181;g/mL) to the worms. Immediately cover them with a coverslip and seal the slide with transparent nail polish gel.</li>
		<li>View the animals after around 10 min on a fluorescence microscope.</li>
	</ol>
	</li>
	<li>Acetone fixation (an alternative approach)
	<ol>
		<li>Wash off the heat-shocked plate from step 2.8 with M9, centrifuge it at 1,000 x g for 1 min at room temperature, and discard the supernatant. Wash the pellet with 1 mL of dH2O, collect the pellet, and discard the supernatant.</li>
		<li>Add 400 &#181;L of 30% acetone for 15 min. Centrifuge at 1,000 x g for 1 min at room temperature and discard the supernatant. Use 500 &#181;L of dH2O to wash the animals 2x. Discard the supernatant.</li>
		<li>Add 200 &#181;L of DAPI (10 &#181;g/mL), or 4x the amount of the total worms&#8217; volume, in the tube for 15 min.</li>
		<li>Centrifuge at 1,000 x g for 1 min and discard the supernatant. Wash the pellet 2x with 500 &#181;L of dH2O. Place the worms on glass slides, cover them with coverslips, seal the slide with transparent nail polish gel, and observe the animals under a fluorescence microscope.</li>
	</ol>
	</li>
</ol>
4. Microscopic Imaging
<ol>
	<li>Turn on the UV light source and a fluorescence microscope. Turn on the computer connected to the microscope.</li>
	<li>Load the slide on the fluorescence microscope. Use a low power objective lens (10X) to locate the worms, increase the magnification power to 400X, and focus on the specimen.</li>
	<li>Open the software provided with the microscope. Click on &#34;Acquisition&#34; in the menu; click on &#34;Camera&#34; and select the camera connected to the microscope; this allows the selected camera to communicate with the software. 
	Note: Different microscopes may vary in operation.</li>
	<li>Under the same &#34;Acquisition&#34;, click on &#34;Multidimensional Acquisition&#34;. This opens a new &#34;Multidimensional Acquisition&#34; navigation window. Click the &#34;Multichannel&#34; button, and all available channels will appear.</li>
	<li>Choose blue, green, and DIC (differential interference contrast) to observe the specimen. In the &#34;Multidimensional acquisition&#34; navigation window, leave the &#34;Blue-DAPI&#34;, &#34;Green-GFP&#34;, and &#34;Grey-DIC&#34;, &#34;Nomarski&#34; channels on; right-click on other channels, such as &#34;Red-rhodamine&#34; and &#34;White-bright field&#34;, to close them.</li>
	<li>First, measure the exposure time for each channel:
	<ol>
		<li>Left-click the &#34;Blue-DAPI&#34; channel to select it and then click on &#34;Measure&#34; in order to take a live image under the DAPI channel with the automated exposure time. A new window with a live image will appear.</li>
		<li>On the left side of the live image window, manually adjust the exposure time if needed to make the image as desired.</li>
		<li>Finally, click on &#34;OK&#34; in the live image window. This sets the exposure time under the DAPI channel.</li>
	</ol>
	</li>
	<li>Repeat step 4.6 for the other channels, &#34;Green-GFP&#34; and &#34;Grey-DIC&#34;, &#34;Nomarski&#34;, to set up the exposure times for those channels.</li>
	<li>In the &#34;Multidimensional acquisition&#34; navigation window, click on &#34;Start&#34; to automatically collect 3 images under the 3 different channels in order (see above) with specific exposure times as set.</li>
	<li>To save the file, go to the main menu, click on &#34;File&#34; then on &#34;Save as&#34;. Input file name and the folder name. Save the file as the default file format to preserve the detailed imaging information for future reference.</li>
	<li>To export the file in other formats:
	<ol>
		<li>Go to &#34;File&#34; and click on &#34;Export&#34;. This will open up an export setting window.</li>
		<li>Set file name and folder. Check &#34;Create a project folder&#34; to better organize the data. The software also allows the user to specify 4 other options: &#34;Generate merged images&#34;, &#34;Use color for channel images&#34;, &#34;Use channel names&#34;, and &#34;Merged image only&#34;.</li>
		<li>Choose a desired file type from the drop-down menu, such as &#34;TIF&#34;, &#34;BMP&#34;, &#34;PSD&#34;, or &#34;JPG&#34; and other types. Then click on &#34;Start&#34; to export.</li>
	</ol>
	</li>
</ol>

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The authors have nothing to declare.

This article presented a fast and efficient method to verify protein subcellular changes from the cytoplasm to the nucleus. The protein expression was shown by a GFP fusion, while the nucleus structure was verified by DAPI staining (Figure 3). Since immunostaining C. elegans proteins is challenging, most C. elegans protein subcellular localizations are characterized by tagging them with marker proteins such as GFP, Laz, mCherry, and others18,</s...

A change of protein subcellular localization is a common mechanism in response to internal or external signals such as heat stress, starvation, oxidative stress, apoptosis, protein phosphorylation, and others. For example, heat stress induces a FOXO member DAF-16 nuclear translocation1,2, and the pro-apoptotic BCL-2 protein BID translocates to the mitochondria upon receiving death signaling3,4. Various techniques are available to detect these changes. A combination of western blotting and biochemically isolating subcellular structures (e.g., mitochondria or the nuclei) could well achieve the goal3. However, it requires a specific antibody against the protein of interest. Thus, a well-established antibody becomes the key to the success. An alternative approach is to label different subcellular structures or organelles with various markers such as green fluorescence protein (GFP), red fluorescence protein (RFP), yellow fluorescence protein (YFP), and mCherry, and meanwhile label the protein of interest with other markers. Then, observe them under a confocal microscope to localize the targets5,6. Radioactive isotopes are an alternative choice for labeling target proteins and then detecting their subcellular localization7. However, this method requires proper training and handling of radioactive wastes. Under circumstances such as the lack of a specific antibody, the absence of a proper marker, or the scarceness of equipment such as a confocal microscope, an alternative approach needs to be considered. To identify a protein nuclear translocation, it is attractive to only label the target proteins with a marker and to stain nuclei with the chemical reagent 4',6-diamidino-2-phenylindole (DAPI) since this only requires a regular fluorescence microscope. 
Immunolabeling C. elegans with antibodies is challenging due to the low permeability of either the eggshell or the collagenous cuticle surrounding the animal. Meanwhile, since C. elegans proteins are significantly divergent from their vertebrate orthologs, a few commercial companies provide C. elegans with specific products. It is difficult for a small laboratory to generate C. elegans antibodies for themselves. Researchers in the community often use tagged proteins as markers to demonstrate a protein localization or gene expression. This article uses EXL-1::GFP as an example to track a protein nuclear translocation under heat stress8. An integrated translational exl-1::gfp into the animal genome is used to stably express the gene with gfp fusion. Research showed that exl-1 is expressed in intestine, body wall muscle, and other subcellular structures8. This protocol demonstrates how to synchronize worms to the fourth larval (L4) stage, perform a heat stress experiment, and conduct DAPI staining, ethanol fixation, acetone fixation, and imaging under a regular fluorescence microscope.

<table>
	<tbody>
		<tr>
			<td>DAPI</td>
			<td>Biotium</td>
			<td>40043</td>
			<td>both Hoechst 33258 and Hoechst 33342 also works well</td>
		</tr>
		<tr>
			<td>OP50</td>
			<td>CGC</td>
			<td>OP50</td>
			<td>https://cgc.umn.edu/</td>
		</tr>
		<tr>
			<td>Caenorhabditis Genetics Center (CGC)</td>
			<td></td>
			<td></td>
			<td>https://cgc.umn.edu/</td>
		</tr>
		<tr>
			<td>Kim Wipe</td>
			<td>Kimberly-Clark Corporation</td>
			<td></td>
			<td>soft tissue</td>
		</tr>
		<tr>
			<td>Zeiss&#160; AX10</td>
			<td>Zeiss</td>
			<td></td>
			<td>fluorescence microscope</td>
		</tr>
		<tr>
			<td>Axiovision Rel 4.8</td>
			<td>Zeiss</td>
			<td></td>
			<td>microscope software</td>
		</tr>
		<tr>
			<td>AxioCam MR Rev3</td>
			<td>Zeiss</td>
			<td></td>
			<td>digital camera</td>
		</tr>
		<tr>
			<td>Incubator&#160;</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Micro centrifuge</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Transparent nail polish gel</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>60 mm petri dishes</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Glass slides</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Glass coverslips</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>1-20 &#181;L pipettor</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>20-200 &#181;L pipettor</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>200-1000 &#181;L pipettor</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>1-200 &#181;L pipet tips</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>200-1000 &#181;L pipet tips</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>1.5 mL microcentrifuge tubes</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Platinum wire worm pick</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
	</tbody>
</table>

detecting protein subcellular localization by green fluorescence protein tagging and 4',6-diamidino-2-phenylindole staining in caenorhabditis elegans

In this protocol, a green fluorescence protein (GFP) fusion protein and 4',6-diamidino-2-phenylindole (DAPI) staining are used to track protein subcellular localization changes; in particular, a nuclear translocation under a heat stress condition. Proteins react correspondingly to external and internal signals. A common mechanism is to change its subcellular localization. This article describes a protocol to track protein localization that does not require an antibody, radioactive labeling, or a confocal microscope. In this article, GFP is used to tag the target protein EXL-1 in C. elegans, a member of the chloride intracellular channel proteins (CLICs) family, including mammalian CLIC4. An integrated translational exl-1::gfp transgenic line (with a promoter and a full gene sequence) was created by transformation and &#947;-radiation, and stably expresses the gene and gfp. Recent research showed that upon heat stress, not oxidative stress, EXL-1::GFP accumulates in the nucleus. Overlapping the GFP signal with both the nuclei structure and the DAPI signals confirms the EXL-1 subcellular localization changes under stress. This protocol presents two different fixation methods for DAPI staining: ethanol fixation and acetone fixation. The DAPI staining protocol presented in this article is fast and efficient and preserves both the GFP signal and the protein subcellular localization changes. This method only requires a fluorescence microscope with Nomarski, a FITC filter, and a DAPI filter. It is suitable for a small laboratory setting, undergraduate student research, high school student research, and biotechnology classrooms.

Chloride intracellular channel proteins (CLIC) are multifunctional proteins that are highly conserved across species12. Much research shows that CLICs regulate cellular stress, autophagy, apoptosis, carcinogenesis, angiogenesis, and the macrophage innate immune response in the mammalian system13,14,15,16,17</s...

Watch this Scientific Journal Video about Detecting Protein Subcellular Localization by Green Fluorescence Protein Tagging and 4',6-Diamidino-2-phenylindole Staining in Caenorhabditis elegans at JoVE.

Detecting Protein Subcellular Localization by Green Fluorescence Protein Tagging and 4',6-Diamidino-2-phenylindole Staining in Caenorhabditis elegans

This protocol demonstrates how to track a protein nuclear translocation under heat stress by using a green fluorescence protein (GFP) fusion protein as a marker and 4',6-diamidino-2-phenylindole (DAPI) staining. The DAPI staining protocol is fast and preserves the GFP and protein subcellular localization signals.

Detecting Protein Subcellular Localization by Green Fluorescence Protein Tagging and 4',6-Diamidino-2-phenylindole Staining in Caenorhabditis elegans

In this protocol, a green fluorescence protein (GFP) fusion protein and 4',6-diamidino-2-phenylindole (DAPI) staining are used to track protein ...

detecting-protein-subcellular-localization-green-fluorescence-protein

Research

JoVE Journal

Biology

9.5K Views.  Borough of Manhattan Community College/CUNY. This protocol demonstrates how to track a protein nuclear translocation under heat stress by using a green fluorescence protein (GFP) fusion protein as a marker and 4',6-diamidino-2-phenylindole (DAPI) staining. The DAPI staining protocol is fast and preserves the GFP and protein subcellular localization signals. 

Video: Detecting Protein Subcellular Localization by Green Fluorescence Protein Tagging and 4',6-Diamidino-2-phenylindole Staining in Caenorhabditis elegans

Staining of Proteins in Gels with Coomassie G-250 without Organic Solvent and Acetic Acid

In classical protein staining protocols using Coomassie Brilliant Blue (CBB), solutions with high contents of toxic and flammable organic solvents (Methanol, Ethanol or 2-Propanol) and acetic acid are used for fixation, staining and destaining of proteins in a gel after SDS-PAGE. To speed up the procedure, heating the staining solution in the microwave oven for a short time is frequently used. This usually results in evaporation of toxic or hazardous Methanol, Ethanol or 2-Propanol and a strong smell of acetic acid in the lab which should be avoided due to safety considerations. In a protocol originally published in two patent applications by E.M. Wondrak (US2001046709 (A1), US6319720 (B1)), an alternative composition of the staining solution is described in which no organic solvent or acid is used. The CBB is dissolved in bidistilled water (60-80mg of CBB G-250 per liter) and 35 mM HCl is added as the only other compound in the staining solution. The CBB staning of the gel is done after SDS-PAGE and thorough washing of the gel in bidistilled water. By heating the gel during the washing and staining steps, the process can be finished faster and no toxic or hazardous compunds are evaporating. The staining of proteins occurs already within 1 minute after heating the gel in staining solution and is fully developed after 15-30 min with a slightly blue background that is destained completely by prolonged washing of the stained gel in bidistilled water, without affecting the stained protein bands.

A short protocol for protein staining with Coomassie Brilliant Blue (CBB) G-250 in polyacrylamide gels is described without using organic solvents or acetic acid as in the classical staining procedures with CBB.

In classical protein staining protocols using Coomassie Brilliant Blue (CBB), solutions with high contents of toxic and flammable organic solvents ...

Using the GELFREE 8100 Fractionation System for Molecular Weight-Based Fractionation with Liquid Phase Recovery

The GELFREE 8100 Fractionation System is a novel protein fractionation system designed to maximize protein recovery during molecular weight based fractionation. The system is comprised of single-use, 8-sample capacity cartridges and a benchtop GELFREE Fractionation Instrument. During separation, a constant voltage is applied between the anode and cathode reservoirs, and each protein mixture is electrophoretically driven from a loading chamber into a specially designed gel column gel. Proteins are concentrated into a tight band in a stacking gel, and separated based on their respective electrophoretic mobilities in a resolving gel. As proteins elute from the column, they are trapped and concentrated in liquid phase in the collection chamber, free of the gel. The instrument is then paused at specific time intervals, and fractions are collected using a pipette. This process is repeated until all desired fractions have been collected. If fewer than 8 samples are run on a cartridge, any unused chambers can be used in subsequent separations.
This novel technology facilitates the quick and simple separation of up to 8 complex protein mixtures simultaneously, and offers several advantages when compared to previously available fractionation methods. This system is capable of fractionating up to 1mg of total protein per channel, for a total of 8mg per cartridge. Intact proteins over a broad mass range are separated on the basis of molecular weight, retaining important physiochemical properties of the analyte. The liquid phase entrapment provides for high recovery while eliminating the need for band or spot cutting, making the fractionation process highly reproducible1.

The accompanying video describes the use of the GELFREE 8100 Fractionation System, which partitions complex protein samples on the basis of molecular weight and recovers the fractions in liquid phase. The video describes how the technology works, how it is used, and provides resultant data, with polyacrylamide gel electrophoresis analysis of fractionated bovine liver homogenate.

The GELFREE 8100 Fractionation System is a novel protein fractionation system designed to maximize protein recovery during molecular weight based ...

Basic Caenorhabditis elegans Methods: Synchronization and Observation

Research into the molecular and developmental biology of the nematode Caenorhabditis elegans was begun in the early seventies by Sydney Brenner and it has since been used extensively as a model organism 1. C. elegans possesses key attributes such as simplicity, transparency and short life cycle that have made it a suitable experimental system for fundamental biological studies for many years 2. Discoveries in this nematode have broad implications because many cellular and molecular processes that control animal development are evolutionary conserved 3.
C. elegans life cycle goes through an embryonic stage and four larval stages before animals reach adulthood. Development can take 2 to 4 days depending on the temperature. In each of the stages several characteristic traits can be observed. The knowledge of its complete cell lineage 4,5 together with the deep annotation of its genome turn this nematode into a great model in fields as diverse as the neurobiology 6, aging 7,8, stem cell biology 9 and germ line biology 10. 
An additional feature that makes C. elegans an attractive model to work with is the possibility of obtaining populations of worms synchronized at a specific stage through a relatively easy protocol. The ease of maintaining and propagating this nematode added to the possibility of synchronization provide a powerful tool to obtain large amounts of worms, which can be used for a wide variety of small or high-throughput experiments such as RNAi screens, microarrays, massive sequencing, immunoblot or in situ hybridization, among others. 
Because of its transparency, C. elegans structures can be distinguished under the microscope using Differential Interference Contrast microscopy, also known as Nomarski microscopy. The use of a fluorescent DNA binder, DAPI (4',6-diamidino-2-phenylindole), for instance, can lead to the specific identification and localization of individual cells, as well as subcellular structures/defects associated to them.

Basic Caenorhabditis elegans Methods: Synchronization and Observation

The easiness of maintaining and propagating the nematode C. elegans make it a nice model organism to work with. The possibility of synchronizing worms allows the work with a significant amount of subjects at the same developmental stage, what facilitates the study of one particular process in many animals.

Research into the molecular and developmental biology of the nematode Caenorhabditis elegans was begun in the early seventies by Sydney Brenner and it has ...

Preparation of the Mgm101 Recombination Protein by MBP-based Tagging Strategy

The MGM101 gene was identified 20 years ago for its role in the maintenance of mitochondrial DNA. Studies from several groups have suggested that the Mgm101 protein is involved in the recombinational repair of mitochondrial DNA. Recent investigations have indicated that Mgm101 is related to the Rad52-type recombination protein family. These proteins form large oligomeric rings and promote the annealing of homologous single stranded DNA molecules. However, the characterization of Mgm101 has been hindered by the difficulty in producing the recombinant protein. Here, a reliable procedure for the preparation of recombinant Mgm101 is described. Maltose Binding Protein (MBP)-tagged Mgm101 is first expressed in Escherichia coli. The fusion protein is initially purified by amylose affinity chromatography. After being released by proteolytic cleavage, Mgm101 is separated from MBP by cationic exchange chromatography. Monodispersed Mgm101 is then obtained by size exclusion chromatography. A yield of ~0.87 mg of Mgm101 per liter of bacterial culture can be routinely obtained. The recombinant Mgm101 has minimal contamination of DNA. The prepared samples are successfully used for biochemical, structural and single particle image analyses of Mgm101. This protocol may also be used for the preparation of other large oligomeric DNA-binding proteins that may be misfolded and toxic to bacterial cells.

The yeast mitochondrial nucleoid protein, Mgm101, is a Rad52-type recombination protein that forms large oligomeric rings. A protocol is described to prepare soluble recombinant Mgm101 using the Maltose Binding Protein (MBP)-tagging strategy coupled with cation exchange and size exclusion chromatography.

The MGM101 gene was identified 20 years ago for its role in the maintenance of mitochondrial DNA. Studies from several groups have suggested that the ...

Using Caenorhabditis elegans as a Model System to Study Protein Homeostasis in a Multicellular Organism

The folding and assembly of proteins is essential for protein function, the long-term health of the cell, and longevity of the organism. Historically, the function and regulation of protein folding was studied in vitro, in isolated tissue culture cells and in unicellular organisms. Recent studies have uncovered links between protein homeostasis (proteostasis), metabolism, development, aging, and temperature-sensing. These findings have led to the development of new tools for monitoring protein folding in the model metazoan organism Caenorhabditis elegans. In our laboratory, we combine behavioral assays, imaging and biochemical approaches using temperature-sensitive or naturally occurring metastable proteins as sensors of the folding environment to monitor protein misfolding. Behavioral assays that are associated with the misfolding of a specific protein provide a simple and powerful readout for protein folding, allowing for the fast screening of genes and conditions that modulate folding. Likewise, such misfolding can be associated with protein mislocalization in the cell. Monitoring protein localization can, therefore, highlight changes in cellular folding capacity occurring in different tissues, at various stages of development and in the face of changing conditions. Finally, using biochemical tools ex vivo, we can directly monitor protein stability and conformation. Thus, by combining behavioral assays, imaging and biochemical techniques, we are able to monitor protein misfolding at the resolution of the organism, the cell, and the protein, respectively.

Using Caenorhabditis elegans as a Model System to Study Protein Homeostasis in a Multicellular Organism

To study the relationship between protein homeostasis, stress and aging, we monitored changes in protein folding by following protein dysfunction, protein localization in the cell and protein stability at the organismal, cellular and protein levels, using the genetically tractable metazoan Caenorhabditis elegans as a model system.

The folding and assembly of proteins is essential for protein function, the long-term health of the cell, and longevity of the organism. Historically, the ...

Protein-protein Interactions Visualized by Bimolecular Fluorescence Complementation in Tobacco Protoplasts and Leaves

Many proteins interact transiently with other proteins or are integrated into multi-protein complexes to perform their biological function. Bimolecular fluorescence complementation (BiFC) is an in vivo method to monitor such interactions in plant cells. In the presented protocol the investigated candidate proteins are fused to complementary halves of fluorescent proteins and the respective constructs are introduced into plant cells via agrobacterium-mediated transformation. Subsequently, the proteins are transiently expressed in tobacco leaves and the restored fluorescent signals can be detected with a confocal laser scanning microscope in the intact cells. This allows not only visualization of the interaction itself, but also the subcellular localization of the protein complexes can be determined. For this purpose, marker genes containing a fluorescent tag can be coexpressed along with the BiFC constructs, thus visualizing cellular structures such as the endoplasmic reticulum, mitochondria, the Golgi apparatus or the plasma membrane. The fluorescent signal can be monitored either directly in epidermal leaf cells or in single protoplasts, which can be easily isolated from the transformed tobacco leaves. BiFC is ideally suited to study protein-protein interactions in their natural surroundings within the living cell. However, it has to be considered that the expression has to be driven by strong promoters and that the interaction partners are modified due to fusion of the relatively large fluorescence tags, which might interfere with the interaction mechanism. Nevertheless, BiFC is an excellent complementary approach to other commonly applied methods investigating protein-protein interactions, such as coimmunoprecipitation, in vitro pull-down assays or yeast-two-hybrid experiments.

Formation of protein complexes in vivo can be visualized by bimolecular fluorescence complementation. Interaction partners are fused to complementary parts of fluorescent tags and transiently expressed in tobacco leaves, resulting in a reconstituted fluorescent signal upon close proximity of the two proteins.

Many proteins interact transiently with other proteins or are integrated into multi-protein complexes to perform their biological function. Bimolecular ...

Green Fluorescent Protein-based Expression Screening of Membrane Proteins in Escherichia coli

The production of recombinant membrane proteins for structural and functional studies remains technically challenging due to low levels of expression and the inherent instability of many membrane proteins once solubilized in detergents. A protocol is described that combines ligation independent cloning of membrane proteins as GFP fusions with expression in Escherichia coli detected by GFP fluorescence. This enables the construction and expression screening of multiple membrane protein/variants to identify candidates suitable for further investment of time and effort. The GFP reporter is used in a primary screen of expression by visualizing GFP fluorescence following SDS polyacrylamide gel electrophoresis (SDS-PAGE). Membrane proteins that show both a high expression level with minimum degradation as indicated by the absence of free GFP, are selected for a secondary screen. These constructs are scaled and a total membrane fraction prepared and solubilized in four different detergents. Following ultracentrifugation to remove detergent-insoluble material, lysates are analyzed by fluorescence detection size exclusion chromatography (FSEC). Monitoring the size exclusion profile by GFP fluorescence provides information about the mono-dispersity and integrity of the membrane proteins in different detergents. Protein: detergent combinations that elute with a symmetrical peak with little or no free GFP and minimum aggregation are candidates for subsequent purification. Using the above methodology, the heterologous expression in E. coli of SED (shape, elongation, division, and sporulation) proteins from 47 different species of bacteria was analyzed. These proteins typically have ten transmembrane domains and are essential for cell division. The results show that the production of the SEDs orthologues in E. coli was highly variable with respect to the expression levels and integrity of the GFP fusion proteins. The experiment identified a subset for further investigation.

Green Fluorescent Protein-based Expression Screening of Membrane Proteins in Escherichia coli

A streamlined approach to screening for the expression of recombinant membrane proteins in Escherichia coli based on fusion to green fluorescent protein is presented.

The production of recombinant membrane proteins for structural and functional studies remains technically challenging due to low levels of expression and ...

Quantitation of Protein Expression and Co-localization Using Multiplexed Immuno-histochemical Staining and Multispectral Imaging

Immunohistochemistry is a commonly used clinical and research lab detection technique for investigating protein expression and localization within tissues. Many semi-quantitative systems have been developed for scoring expression using immunohistochemistry, but inherent subjectivity limits reproducibility and accuracy of results. Furthermore, the investigation of spatially overlapping biomarkers such as nuclear transcription factors is difficult with current immunohistochemistry techniques. We have developed and optimized a system for simultaneous investigation of multiple proteins using high throughput methods of multiplexed immunohistochemistry and multispectral imaging. Multiplexed immunohistochemistry is performed by sequential application of primary antibodies with secondary antibodies conjugated to horseradish peroxidase or alkaline phosphatase. Different chromogens are used to detect each protein of interest. Stained slides are loaded into an automated slide scanner and a protocol is created for automated image acquisition. A spectral library is created by staining a set of slides with a single chromogen on each. A subset of representative stained images are imported into multispectral imaging software and an algorithm for distinguishing tissue type is created by defining tissue compartments on images. Subcellular compartments are segmented by using hematoxylin counterstain and adjusting the intrinsic algorithm. Thresholding is applied to determine positivity and protein co-localization. The final algorithm is then applied to the entire set of tissues. Resulting data allows the user to evaluate protein expression based on tissue type (ex. epithelia vs. stroma) and subcellular compartment (nucleus vs. cytoplasm vs. plasma membrane). Co-localization analysis allows for investigation of double-positive, double-negative, and single-positive cell types. Combining multispectral imaging with multiplexed immunohistochemistry and automated image acquisition is an objective, high-throughput method for investigation of biomarkers within tissues.

Immunohistochemistry is a powerful lab technique for evaluating protein localization and expression within tissues. Current semi-automated methods for quantitation introduce subjectivity and often create irreproducible results. Herein, we describe methods for multiplexed immunohistochemistry and objective quantitation of protein expression and co-localization using multispectral imaging.

Immunohistochemistry is a commonly used clinical and research lab detection technique for investigating protein expression and localization within ...

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, ...

Methods to Study Changes in Inherent Protein Aggregation with Age in Caenorhabditis elegans

In the last decades, the prevalence of neurodegenerative disorders, such as Alzheimer's disease (AD) and Parkinson's disease (PD), has grown. These age-associated disorders are characterized by the appearance of protein aggregates with fibrillary structure in the brains of these patients. Exactly why normally soluble proteins undergo an aggregation process remains poorly understood. The discovery that protein aggregation is not limited to disease processes and instead part of the normal aging process enables the study of the molecular and cellular mechanisms that regulate protein aggregation, without using ectopically expressed human disease-associated proteins. Here we describe methodologies to examine inherent protein aggregation in Caenorhabditis elegans through complementary approaches. First, we examine how to grow large numbers of age-synchronized C. elegans to obtain aged animals and we present the biochemical procedures to isolate highly-insoluble-large aggregates. In combination with a targeted genetic knockdown, it is possible to dissect the role of a gene of interest in promoting or preventing age-dependent protein aggregation by using either a comprehensive analysis with quantitative mass spectrometry or a candidate-based analysis with antibodies. These findings are then confirmed by in vivo analysis with transgenic animals expressing fluorescent-tagged aggregation-prone proteins. These methods should help clarify why certain proteins are prone to aggregate with age and ultimately how to keep these proteins fully functional.

Methods to Study Changes in Inherent Protein Aggregation with Age in Caenorhabditis elegans

The goal of the method presented here is to explore protein aggregation during normal aging in the model organism C. elegans. The protocol represents a powerful tool to study the highly insoluble large aggregates that form with age and to determine how changes in proteostasis impact protein aggregation.

In the last decades, the prevalence of neurodegenerative disorders, such as Alzheimer's disease (AD) and Parkinson's disease (PD), has grown. These ...

Detecting Protein Subcellular Localization by Green Fluorescence Protein Tagging and 4',6-Diamidino-2-phenylindole Staining in Caenorhabditis elegans

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Chapters in this video

Staining of Proteins in Gels with Coomassie G-250 without Organic Solvent and Acetic Acid

Using the GELFREE 8100 Fractionation System for Molecular Weight-Based Fractionation with Liquid Phase Recovery

Basic Caenorhabditis elegans Methods: Synchronization and Observation

Preparation of the Mgm101 Recombination Protein by MBP-based Tagging Strategy

Using Caenorhabditis elegans as a Model System to Study Protein Homeostasis in a Multicellular Organism

Protein-protein Interactions Visualized by Bimolecular Fluorescence Complementation in Tobacco Protoplasts and Leaves

Green Fluorescent Protein-based Expression Screening of Membrane Proteins in Escherichia coli

Quantitation of Protein Expression and Co-localization Using Multiplexed Immuno-histochemical Staining and Multispectral Imaging

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

Methods to Study Changes in Inherent Protein Aggregation with Age in Caenorhabditis elegans