Name: detecting protein subcellular localization by green fluorescence protein tagging and 4',6-diamidino-2-phenylindole staining in caenorhabditis elegans
Uploaded: 2018-07-30T13:00:00.000+00:00
Duration: 9 min 36 s
Description: 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.

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&amp;nbsp; 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&amp;nbsp;</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 &amp;micro;L pipettor</td><td></td><td></td><td></td></tr><tr><td>20-200 &amp;micro;L pipettor</td><td></td><td></td><td></td></tr><tr><td>200-1000 &amp;micro;L pipettor</td><td></td><td></td><td></td></tr><tr><td>1-200 &amp;micro;L pipet tips</td><td></td><td></td><td></td></tr><tr><td>200-1000 &amp;micro;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

The C. elegans Intestine As a Model for Intercellular Lumen Morphogenesis and In Vivo Polarized Membrane Biogenesis at the Single-cell Level: Labeling by Antibody Staining, RNAi Loss-of-function Analysis and Imaging

Multicellular tubes, fundamental units of all internal organs, are composed of polarized epithelial or endothelial cells, with apical membranes lining the lumen and basolateral membranes contacting each other and/or the extracellular matrix. How this distinctive membrane asymmetry is established and maintained during organ morphogenesis is still an unresolved question of cell biology. This protocol describes the C. elegans intestine as a&#160;model for the analysis of polarized membrane biogenesis during tube morphogenesis, with emphasis on apical membrane and lumen biogenesis. The C. elegans twenty-cell single-layered intestinal epithelium is arranged into a simple bilaterally symmetrical tube, permitting analysis on a single-cell level. Membrane polarization occurs concomitantly with polarized cell division and migration during early embryogenesis, but de novo polarized membrane biogenesis continues throughout larval growth, when cells no longer proliferate and move. The latter setting allows one to separate subcellular changes that simultaneously mediate these different polarizing processes, difficult to distinguish in most polarity models. Apical-, basolateral membrane-, junctional-, cytoskeletal- and endomembrane components can be labeled and tracked throughout development by GFP fusion proteins, or assessed by in situ antibody staining. Together with the organism&#39;s genetic versatility, the C. elegans intestine thus provides a unique in vivo model for the visual, developmental, and molecular genetic analysis of polarized membrane and tube biogenesis. The specific methods (all standard) described here include how to: label intestinal subcellular components by antibody staining; analyze genes involved in polarized membrane biogenesis by loss-of-function studies adapted to the typically essential tubulogenesis genes; assess polarity defects during different developmental stages; interpret phenotypes by epifluorescence, differential interference contrast (DIC) and confocal microscopy; quantify visual defects. This protocol can be adapted to analyze any of the often highly conserved molecules involved in epithelial polarity, membrane biogenesis, tube and lumen morphogenesis.

The C. elegans Intestine As a Model for Intercellular Lumen Morphogenesis and In Vivo Polarized Membrane Biogenesis at the Single-cell Level: Labeling by Antibody Staining, RNAi Loss-of-function Analy

The transparent C. elegans intestine can serve as an "in vivo tissue chamber" for studying apicobasal membrane and lumen biogenesis at the single-cell and subcellular level during multicellular tubulogenesis. This protocol describes how to combine standard labeling, loss-of-function genetic/RNAi and microscopic approaches to dissect these processes on a molecular level.

Multicellular tubes, fundamental units of all internal organs, are composed of polarized epithelial or endothelial cells, with apical membranes lining the ...

Developmental Biology

The C. elegans Excretory Canal as a Model for Intracellular Lumen Morphogenesis and In Vivo Polarized Membrane Biogenesis in a Single Cell: labeling by GFP-fusions, RNAi Interaction Screen and Imaging

The four C. elegans excretory canals are narrow tubes extended through the length of the animal from a single cell, with almost equally far extended intracellular endotubes that build and stabilize the lumen with a membrane and submembraneous cytoskeleton of apical character. The excretory cell expands its length approximately 2,000 times to generate these canals, making this model unique for the in vivo assessment of de novo polarized membrane biogenesis, intracellular lumen morphogenesis and unicellular tubulogenesis. The protocol presented here shows how to combine standard labeling, gain- and loss-of-function genetic or RNA interference (RNAi)-, and microscopic approaches to use this model to visually dissect and functionally analyze these processes on a molecular level. As an example of a labeling approach, the protocol outlines the generation of transgenic animals with fluorescent fusion proteins for live analysis of tubulogenesis. As an example of a genetic approach, it highlights key points of a visual RNAi-based interaction screen designed to modify a gain-of-function cystic canal phenotype. The specific methods described are how to: label and visualize the canals by expressing fluorescent proteins; construct a targeted RNAi library and strategize RNAi screening for the molecular analysis of canal morphogenesis; visually assess modifications of canal phenotypes; score them by dissecting fluorescence microscopy; characterize subcellular canal components at higher resolution by confocal microscopy; and quantify visual parameters. The approach is useful for the investigator who is interested in taking advantage of the C. elegans excretory canal for identifying and characterizing genes involved in the phylogenetically conserved processes of intracellular lumen and unicellular tube morphogenesis.

The C. elegans Excretory Canal as a Model for Intracellular Lumen Morphogenesis and In Vivo Polarized Membrane Biogenesis in a Single Cell: labeling by GFP-fusions, RNAi Interaction Screen and Imaging

The C. elegans excretory canal is a unique single-cell model for the visual in vivo analysis of de novo polarized membrane biogenesis. This protocol describes a combination of standard genetic/RNAi and imaging approaches, adaptable for the identification and characterization of molecules directing unicellular tubulogenesis, and apical membrane and lumen biogenesis.

The four C. elegans excretory canals are narrow tubes extended through the length of the animal from a single cell, with almost equally far extended ...

Immobilization of Caenorhabditis elegans to Analyze Intracellular Transport in Neurons

Axonal transport and intraflagellar transport (IFT) are essential for axon and cilia morphogenesis and function. Kinesin superfamily proteins and dynein are molecular motors that regulate anterograde and retrograde transport, respectively. These motors use microtubule networks as rails. Caenorhabditis elegans (C. elegans) is a powerful model organism to study axonal transport and IFT in vivo. Here, I describe a protocol to observe axonal transport and IFT in living C. elegans. Transported cargo can be visualized by tagging cargo proteins using fluorescent proteins such as green fluorescent protein (GFP). C. elegans is transparent and GFP-tagged cargo proteins can be expressed in specific cells under cell-specific promoters. Living worms can be fixed by microbeads on 10% agarose gel without killing or anesthetizing the worms. Under these conditions, cargo movement can be directly observed in the axons and cilia of living C. elegans without dissection. This method can be applied to the observation of any cargo molecule in any cells by modifying the target proteins and/or the cells they are expressed in. Most basic proteins such as molecular motors and adaptor proteins that are involved in axonal transport and IFT are conserved in C. elegans. Compared to other model organisms, mutants can be obtained and maintained more easily in C. elegans. Combining this method with various C. elegans mutants can clarify the molecular mechanisms of axonal transport and IFT.

Immobilization of Caenorhabditis elegans to Analyze Intracellular Transport in Neurons

Caenorhabditis elegans (C. elegans) is a good model to study axonal and intracellular transport. Here, I describe a protocol for in vivo recording and analysis of axonal and intraflagellar transport in C. elegans.

Axonal transport and intraflagellar transport (IFT) are essential for axon and cilia morphogenesis and function. Kinesin superfamily proteins and dynein ...

In vivo Neuronal Calcium Imaging in C. elegans

The nematode worm C. elegans is an ideal model organism for relatively simple, low cost neuronal imaging in vivo. Its small transparent body and simple, well-characterized nervous system allows identification and fluorescence imaging of any neuron within the intact animal. Simple immobilization techniques with minimal impact on the animal's physiology allow extended time-lapse imaging. The development of genetically-encoded calcium sensitive fluorophores such as cameleon 1 and GCaMP 2 allow in vivo imaging of neuronal calcium relating both cell physiology and neuronal activity. Numerous transgenic strains expressing these fluorophores in specific neurons are readily available or can be constructed using well-established techniques. Here, we describe detailed procedures for measuring calcium dynamics within a single neuron in vivo using both GCaMP and cameleon. We discuss advantages and disadvantages of both as well as various methods of sample preparation (animal immobilization) and image analysis. Finally, we present results from two experiments: 1) Using GCaMP to measure the sensory response of a specific neuron to an external electrical field and 2) Using cameleon to measure the physiological calcium response of a neuron to traumatic laser damage. Calcium imaging techniques such as these are used extensively in C. elegans and have been extended to measurements in freely moving animals, multiple neurons simultaneously and comparison across genetic backgrounds. C. elegans presents a robust and flexible system for in vivo neuronal imaging with advantages over other model systems in technical simplicity and cost.

In vivo Neuronal Calcium Imaging in C. elegans

With its small transparent body, well-documented neuroanatomy and a host of amenable genetic techniques and reagents, C. elegans makes an ideal model organism for in vivo neuronal imaging using relatively simple, low-cost techniques. Here we describe single neuron imaging within intact adult animals using genetically encoded fluorescent calcium indicators.

The nematode worm C. elegans is an ideal model organism  for relatively simple, low cost neuronal imaging in vivo. Its small transparent body and simple, ...

Neuroscience

Antibody Staining in C. Elegans Using &quot;Freeze-Cracking&quot;

To stain C. elegans with antibodies, the relatively impermeable cuticle must be bypassed by chemical or mechanical methods. "Freeze-cracking" is one method used to physically pull the cuticle from nematodes by compressing nematodes between two adherent slides, freezing them, and pulling the slides apart. Freeze-cracking provides a simple and rapid way to gain access to the tissues without chemical treatment and can be used with a variety of fixatives. However, it leads to the loss of many of the specimens and the required compression mechanically distorts the sample. Practice is required to maximize recovery of samples with good morphology. Freeze-cracking can be optimized for specific fixation conditions, recovery of samples, or low non-specific staining, but not for all parameters at once. For antibodies that require very hard fixation conditions and tolerate the chemical treatments needed to chemically permeabilize the cuticle, treatment of intact nematodes in solution may be preferred. If the antibody requires a lighter fix or if the optimum fixation conditions are unknown, freeze-cracking provides a very useful way to rapidly assay the antibody and can yield specific subcellular and cellular localization information for the antigen of interest.

Antibody Staining in C. Elegans Using "Freeze-Cracking"

"Freeze-cracking," a method for exposing the inner tissues of the nematode C. elegans to antibodies for protein localization, is demonstrated.

To stain C.  elegans with antibodies, the relatively impermeable cuticle must be  bypassed by chemical or mechanical methods.  "Freeze-cracking" is one ...

RNA Fluorescence in situ Hybridization (FISH) to Visualize Microbial Colonization and Infection in Caenorhabditis elegans Intestines

The intestines of wild Caenorhabditis nematodes are inhabited by a variety of microorganisms, including gut microbiome bacteria and pathogens, such as microsporidia and viruses. Because of the similarities between Caenorhabditis elegans and mammalian intestinal cells, as well as the power of the C. elegans system, this host has emerged as a model system to study host intestine-microbe interactions in vivo. While it is possible to observe some aspects of these interactions with bright-field microscopy, it is difficult to accurately classify microbes and characterize the extent of colonization or infection without more precise tools. RNA fluorescence in situ hybridization (FISH) can be used as a tool to identify and visualize microbes in nematodes from the wild or to experimentally characterize and quantify infection in nematodes infected with microbes in the lab. FISH probes, labeling the highly abundant small subunit ribosomal RNA, produce a bright signal for bacteria and microsporidian cells. Probes designed to target conserved regions of ribosomal RNA common to many species can detect a broad range of microbes, whereas targeting divergent regions of the ribosomal RNA is useful for narrower detection. Similarly, probes can be designed to label viral RNA. A protocol for RNA FISH staining with either paraformaldehyde (PFA) or acetone fixation is presented. PFA fixation is ideal for nematodes associated with bacteria, microsporidia, and viruses, whereas acetone fixation is necessary for the visualization of microsporida spores. Animals were first washed and fixed in paraformaldehyde or acetone. After fixation, FISH probes were incubated with samples to allow for the hybridization of probes to the desired target. The animals were again washed and then examined on microscope slides or using automated approaches. Overall, this FISH protocol enables detection, identification, and quantification of the microbes that inhabit the C. elegans intestine, including microbes for which there are no genetic tools available.

RNA Fluorescence in situ Hybridization FISH to Visualize Microbial Colonization and Infection in Caenorhabditis elegans Intestines

Intestinal microbes, including extracellular bacteria and intracellular pathogens like the Orsay virus and microsporidia (fungi), are often associated with wild Caenorhabditis nematodes. This article presents a protocol for detecting and quantifying microbes that colonize and/or infect C. elegans nematodes, and for measuring pathogen load after controlled infections in the lab.

The intestines of wild Caenorhabditis nematodes are inhabited by a variety of microorganisms, including gut microbiome bacteria and pathogens, such as ...

Immunology and Infection

Methods to Assess Subcellular Compartments of Muscle in C. elegans

Muscle is a dynamic tissue that responds to changes in nutrition, exercise, and disease state. The loss of muscle mass and function with disease and age are significant public health burdens. We currently understand little about the genetic regulation of muscle health with disease or age. The nematode C. elegans is an established model for understanding the genomic regulation of biological processes of interest. This worm&#8217;s body wall muscles display a large degree of homology with the muscles of higher metazoan species. Since C. elegans is a transparent organism, the localization of GFP to mitochondria and sarcomeres allows visualization of these structures in vivo. Similarly, feeding animals cationic dyes, which accumulate based on the existence of a mitochondrial membrane potential, allows the assessment of mitochondrial function in vivo. These methods, as well as assessment of muscle protein homeostasis, are combined with assessment of whole animal muscle function, in the form of movement assays, to allow correlation of sub-cellular defects with functional measures of muscle performance. Thus, C. elegans provides a powerful platform with which to assess the impact of mutations, gene knockdown, and/or chemical compounds upon muscle structure and function. Lastly, as GFP, cationic dyes, and movement assays are assessed non-invasively, prospective studies of muscle structure and function can be conducted across the whole life course and this at present cannot be easily investigated in vivo in any other organism.

Methods to Assess Subcellular Compartments of Muscle in C. elegans

Skeletal muscle is essential for locomotion and is the bodies&#8217; main protein store. Muscle health measurements within C. elegans are described. Prospective changes to muscle structure and function are assessed using localized GFP and cationic dyes.

Muscle is a dynamic tissue that responds to changes in nutrition, exercise, and disease state. The loss of muscle mass and function with disease and age ...

Subcellular Imaging of Neuronal Calcium Handling In Vivo

Calcium (Ca2+) imaging has been largely used to examine neuronal activity, but it is becoming increasingly clear that subcellular Ca2+ handling is a crucial component of intracellular signaling. The visualization of subcellular Ca2+ dynamics in vivo,&#160;where neurons can be studied in their native, intact circuitry, has proven technically challenging in complex nervous systems. The transparency and relatively simple nervous system of the nematode Caenorhabditis elegans enable the cell-specific expression and in vivo visualization of fluorescent tags and indicators. Among these are fluorescent indicators that have been modified for use in the cytoplasm as well as various subcellular compartments, such as the mitochondria. This protocol enables non-ratiometric Ca2+ imaging in vivo with a subcellular resolution that permits the analysis of Ca2+ dynamics down to the level of individual dendritic spines and mitochondria. Here, two available genetically encoded indicators with different Ca2+ affinities are used to demonstrate the use of this protocol for measuring relative Ca2+ levels within the cytoplasm or mitochondrial matrix in a single pair of excitatory interneurons (AVA). Together with the genetic manipulations and longitudinal observations possible in C. elegans, this imaging protocol may be useful for answering questions regarding how Ca2+ handling regulates neuronal function and plasticity.

Subcellular Imaging of Neuronal Calcium Handling In Vivo

The current methods describe a non-ratiometric approach for high-resolution, sub-compartmental calcium imaging in vivo in Caenorhabditis elegans using readily available genetically encoded calcium indicators.

Calcium (Ca2+) imaging has been largely used to examine neuronal activity, but it is becoming increasingly clear that subcellular Ca2+ handling is a ...

Quantitative Approaches for Studying Cellular Structures and Organelle Morphology in Caenorhabditis elegans

Defining the cellular mechanisms underlying disease is essential for the development of novel therapeutics. A strategy frequently used to unravel these mechanisms is to introduce mutations in candidate genes and qualitatively describe changes in the morphology of tissues and cellular organelles. However, qualitative descriptions may not capture subtle phenotypic differences, might misrepresent phenotypic variations across individuals in a population, and are frequently assessed subjectively. Here, quantitative approaches are described to study the morphology of tissues and organelles in the nematode Caenorhabditis elegans using laser scanning confocal microscopy combined with commercially available bio-image processing software. A quantitative analysis of phenotypes affecting synapse integrity (size and integrated fluorescence levels), muscle development (muscle cell size and myosin filament length), and mitochondrial morphology (circularity and size) was performed to understand the effects of genetic mutations on these cellular structures. These quantitative approaches are not limited to the applications described here, as they could readily be used to quantitatively assess the morphology of other tissues and organelles in the nematode, as well as in other model organisms.

Quantitative Approaches for Studying Cellular Structures and Organelle Morphology in Caenorhabditis elegans

This study outlines quantitative measurements of synaptic size and localization, muscle morphology, and mitochondrial shape in C. elegans using freely available image processing tools. This approach allows future studies in&#160;C. elegans&#160;to quantitatively compare the extent of tissue and organelle structural changes as a result of genetic mutations.

Defining the cellular mechanisms underlying disease is essential for the development of novel therapeutics. A strategy frequently used to unravel these ...

Studying Membrane Protein Trafficking in Drosophila Photoreceptor Cells Using eGFP-Tagged Proteins

Membrane protein trafficking regulates the incorporation and removal of receptors and ion channels into the plasma membrane. This process is fundamentally important for cell function and cell integrity of neurons. Drosophila photoreceptor cells have become a model for studying membrane protein trafficking. Besides rhodopsin, which upon illumination becomes internalized from the photoreceptor membrane and is degraded, the transient receptor potential-like (TRPL) ion channel in Drosophila exhibits a light-dependent translocation between the rhabdomeral photoreceptor membrane (where it is located in the dark) and the photoreceptor cell body (to which it is transported upon illumination). This intracellular transport of TRPL can be studied in a simple and non-invasive way by expressing eGFP-tagged TRPL in photoreceptor cells. The eGFP fluorescence can then be observed either in the deep pseudopupil or by water immersion microscopy. These methods allow detection of fluorescence in the intact eye and are therefore useful for high-throughput assays and genetic screens for Drosophila mutants defective in TRPL translocation. Here, the preparation of flies, the microscopic techniques, as well as quantification methods used to study this light-triggered translocation of TRPL are explained in detail. These methods can be applied also for trafficking studies on other Drosophila photoreceptor proteins, for example, rhodopsin. In addition, by using eGFP-tagged rhabdomeral proteins, these methods can be used to assess the degeneration of photoreceptor cells.

Studying Membrane Protein Trafficking in Drosophila Photoreceptor Cells Using eGFP-Tagged Proteins

Here, non-invasive methods are described for localization of photoreceptor membrane proteins and assessment of retinal degeneration in the Drosophila compound eye using eGFP fluorescence.

Membrane protein trafficking regulates the incorporation and removal of receptors and ion channels into the plasma membrane. This process is fundamentally ...

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

The C. elegans Intestine As a Model for Intercellular Lumen Morphogenesis and In Vivo Polarized Membrane Biogenesis at the Single-cell Level: Labeling by Antibody Staining, RNAi Loss-of-function Analysis and Imaging

The C. elegans Excretory Canal as a Model for Intracellular Lumen Morphogenesis and In Vivo Polarized Membrane Biogenesis in a Single Cell: labeling by GFP-fusions, RNAi Interaction Screen and Imaging

Immobilization of Caenorhabditis elegans to Analyze Intracellular Transport in Neurons

In vivo Neuronal Calcium Imaging in C. elegans

Antibody Staining in C. Elegans Using "Freeze-Cracking"

RNA Fluorescence in situ Hybridization (FISH) to Visualize Microbial Colonization and Infection in Caenorhabditis elegans Intestines

Methods to Assess Subcellular Compartments of Muscle in C. elegans

Subcellular Imaging of Neuronal Calcium Handling In Vivo

Quantitative Approaches for Studying Cellular Structures and Organelle Morphology in Caenorhabditis elegans

Studying Membrane Protein Trafficking in Drosophila Photoreceptor Cells Using eGFP-Tagged Proteins