D.S.A acknowledges funding from Conselho Nacional de Pesquisa e Desenvolvimento (CNPq/Brazil), grant number #301808/2018-0, #313117/2019-5, Funda&#231;&#227;o de Amparo &#224; Pesquisa do Estado do Rio Grande do Sul (FAPERGS/Brazil), grant number, 21/2551-0001963-8, Coordena&#231;&#227;o de Aperfei&#231;oamento de Pessoal de N&#237;vel Superior (CAPES, Finance Code 001 for N.S.J and A.C.S)

NOTE: Escherichia coli OP50 (E. coli OP50) bacteria and Bristol N2 wild type strains can be obtained from the CGC, University of Minnesota, USA or from donation from a C. elegans laboratory. For researchers&#39; safety, it is imperative to use Personal Protective Equipment. Although the concentrations of reagents like hypochlorite and sodium hydroxide are low, it is essential to wear the recommended PPE, as highlighted in the manuscript, to minimize any potential risks associated with these chemicals.
1. Glycogen content
<ol>
	<li>Preparation of test plates
	<ol>
		<li>Prepare six, 60 mm x 15 mm NGM agar plates20&#160;(10 mL of nematode growth media agar, NGM, as described in Supplementary Table 1) per worm strain and allow them to dry for 1 day at room temperature. Keep the plates closed to prevent contamination.</li>
		<li>Add 200 &#181;L of E. coli OP50 liquid culture (with an optical density = 0.600, approximately at a wavelength of 600 nm) per plate to inoculate a total of four, 60 mm x 15 mm NGM plates per strain in the flow hood. Allow the closed plates to incubate at 37 &#176;C for 1 day before use. 
		NOTE: Prepare six plates (step 1.1) but inoculate only 4 with E. coli OP50 bacteria. Reserve the remaining two plates at 4 &#176;C for later use when placing the worms under starvation conditions. This analysis may be performed in duplicate.</li>
		<li>On the test day, add 200 &#181;L of 0.025 M D-Glucose to two (6 mm x 15 mm) NGM agar plates per strain that were previously seeded with E. coli OP50 bacteria. Allow the plates to dry at room temperature in the flow hood. 
		NOTE: The NGM agar plates seeded with E. coli OP50 bacteria can also be dried near to alcohol lamps as an alternative to using a flow hood.</li>
	</ol>
	</li>
	<li>Synchronization
	<ol>
		<li>Approximately 3 days before the synchronization, transfer 3-4 chunks (3 cm x 3cm) from a maintenance NGM agar with C. elegans Bristol N2 (wild type) at different stages (around 500 worms per chunk) to a new plate (150 mm x 90 mm) that has been previously seeded with E. coli OP50 bacteria.</li>
		<li>Place the new plate with worms in a controlled environment at 20 &#176;C and maintain humidity &#62;95% for 3 days (72 h allow most of the worms to reach the gravid adult stage). 
		NOTE: The growth temperature may need to be adjusted depending on the C. elegans strain(s) being used.</li>
		<li>Using a Pasteur pipette, collect the worms from the prepared plate with distilled H2O and transfer them to a 50 mL centrifuge tube.</li>
		<li>Wait for the sedimentation of the worms by gravity (approximately 15 min) and then remove the supernatant. Repeat this process 3x to eliminate the bacteria. After the last wash, reduce the volume to 5 mL.</li>
		<li>Add 10 mL of bleaching solution (Supplementary Table 1)&#160;and vigorously shake using the hands for approximately 6 min.</li>
		<li>Immediately after the shaking is complete, fill the tubes to 50 mL capacity with M9 buffer.</li>
		<li>Centrifuge at 1400 x g for 3 min, remove the supernatant up to 5 mL and then add 45 mL of M9 buffer again. Repeat this process 4x.</li>
		<li>After the last wash, reduce the volume to 15 mL and maintain it at a controlled temperature (20 &#176;C) and humidity (&#62;95%) for approximately 14 h. 
		NOTE: The synchronization process can be replaced by obtaining eggs through oviposition: Transfer approximately 20 adult pregnant worms to an NGM plate (6 mm x 15 mm) previously seeded with 200 &#181;L of E. coli OP50. Allow the worms to lay eggs for 1 day. After that, remove the pregnant worms and wait for the eggs to hatch.</li>
	</ol>
	</li>
	<li>Preparation of worms
	<ol>
		<li>Using an automatic pipette add 10 &#181;L of the synchronized worms (step 1.2.8) to a microscope slide and count the number of worms. Calculate how much volume must be pipetted to obtain 500-1000 worms/&#181;L.</li>
		<li>Transfer 500-1000 L1 synchronized worms (step 1.2.8) to NGM agar plates (150 mm x 90 mm, Supplementary Table 1) seeded with E. coli OP50&#160;bacteria as a food source, until they reach the larvae stage 4 (L4) at 20 &#176;C. 
		NOTE: Worms can be transferred using an automatic pipette. If an automatic pipette is not available, use a glass Pasteur pipette, and volumes can be controlled using microtube labels or glass graduated pipettes.</li>
		<li>After 48 h of synchronization20: collect the L4 larvae from the plates with M9 buffer (Supplementary Table 1), with the aid of a plastic Pasteur pipette to a clean conical tube 50 mL and wash them 3x with fresh M9 buffer or repeat the washes until all remaining E. coli OP50 bacteria are completely removed (at room temperature). 
		NOTE: The washing step is important because some worms will experience starvation during the procedure.</li>
	</ol>
	</li>
	<li>Running the assay
	<ol>
		<li>Transfer 500-1000 L4 worms (step 1.3.1) from each of the new NGM agar plates (60mm x 15mm) previously prepared (steps 1.1.2 and 1.1.3). Keep them at 20 &#176;C for 48 h until the test day. This schedule will result in three experimental groups as following, in duplicate (Figure 1): 
		A- (starvation): worms will grow on regular NGM agar plates without E. coli OP50 bacteria; 
		B- (regular): worms will grow on regular NGM agar plates seeded with E. coli OP50 bacteria; 
		C- (high glucose): worms will grow on NGM agar plates seeded with E. coli OP50 bacteria and containing 0.025 M D-glucose.</li>
		<li>On the test day (after 48 h): collect the worms and give them three brief washes in the M9 buffer (steps 1.2.2 and 1.2.3) 
		NOTE: Here, prepare NGM agar plates with the addition of D-glucose (step 1.3) and set them aside until it is time to add the worms.
		<ol>
			<li>Iodine staining: prepare a 5% (v/v) Lugol&#39;s Iodine solution (iodine/potassium iodide solution) using fresh M9 buffer. 
			NOTE: Lugol&#39;s Iodine solution can be obtained from local reagent suppliers or pharmacies, therefore the initial concentration may vary. If necessary, calculate to obtain a 5% (v/v) solution.</li>
		</ol>
		</li>
		<li>Transfer approximately 100 &#181;L of washed worms (step 4.2) from each group to the 1.5 mL microtube containing diluted Lugol solution (5% v/v). Using an automatic pipette in a 1:5 proportion transfer 100 &#181;L of worms to 400 &#181;L of Lugol solution, followed by gentle agitation in a mixer for 5 min. 
		NOTE: If a mixer is unavailable, it is possible to shake the microtubes carefully by hands.</li>
		<li>Right after these 5 min, centrifuge the 1.5 mL microtube (worms + Lugol solution) at 1400 x g for 3 min. 
		NOTE: If this assay is performed on worms from the L4 stage or beyond, it is possible to achieve gravity sedimentation instead of using a centrifuge. Leave the microtubes open in racks on the benchtop for 10 min until all the worms have settled at the bottom of the microtube.</li>
		<li>Remove the supernatant and wash the worms with 1.0 mL of fresh M9 buffer. Repeat the wash until all remaining iodine is removed from the solution (minimum 3x).</li>
		<li>After the last washing step, remove the supernatant except for a residual of approximately 100 &#181;L. Use this to gently resuspend the worms and for microscopic analysis.</li>
		<li>Transfer approximately 50 &#181;L of the resuspended worm solution to microscopy slides and cover them with cover slips. Observe the bright field images using a stereomicroscope (at 1.5x). 
		NOTE: If a stereomicroscope is unavailable, a regular microscope can be used, and images can be captured with a cell phone camera (at 3.4x) using an adapter. To prevent errors from light/brightness/exposure, it is crucial that all cell phone camera settings remain consistent for all pictures.</li>
	</ol>
	</li>
	<li>Data inspection
	<ol>
		<li>Quantitative data: Calculate the glycogen content based on the iodine staining of worms.
		<ol>
			<li>Save images from stereomicroscope containing a minimum of 10 worms per group as a .jpeg file for further processing. Download the free ImageJ software to do this. Details are provided in Supplementary File 1, Supplementary Figure 1, Supplementary Figure 2.</li>
			<li>Open the .jpeg image using ImageJ software (which can be downloaded for free at https://imagej.nih.gov/ij/download.html). Click on&#160;&#8203;Segmented line and outline the worm (one worm at a time). Click on Analyze, and then select Measure to obtain the stain quantification. The data will be displayed as the Mean in the table. Mean indicates the data calculated from staining density per worm area.</li>
		</ol>
		</li>
		<li>Qualitative data: Obtain images from the stereomicroscope of each group and visually compare the staining of the worms. Create a scoring system, if needed, to analyze the staining: 0 colorless; 1 lightly stained; 2 stained and 3 heavily stained. Using the stereomicroscope, manually count approximately 10 worms per group (minimum) with a hand counter.</li>
	</ol>
	</li>
</ol>
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/66084/66084fig01.jpg" /> 
Figure 1: Overall glycogen content assay schematic in C. elegans. A schematic of the experimental performed here to carry out the glycogen content assay. <a href="https://www.jove.com/files/ftp_upload/66084/66084fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
2. Evaluation of intestinal permeability
<ol>
	<li>Preparation of worms
	<ol>
		<li>Approximately 14 h after synchronization (step 1.2.8), transfer 500-1000 worms (step 1.3.1 and NOTE) at the first larval stage (L1) to each new NGM agar plate (60 mm x 15 mm) previously prepared and keep them at 20 &#176;C until the day of the test. This schedule will lead to two experimental groups as follows (Figure 2): 
		A- (young): worms will grow on NGM agar plates seeded with the bacteria E. coli OP50 until larval stage 4 (L4); 
		B- (old): the worms will grow on NGM agar plates seeded with the bacteria E. coli OP50 until the 7th day of adulthood. 
		NOTE: The worms maintained until the 7th day of adulthood must be washed with M9 buffer every day and transferred to new NGM agar plates (60 mm x 15mm) with 200 &#181;L of E. coli OP50 bacteria previously seeded for food replacement and removal of the progeny. Younger worms float during decantation and can be separated by removing supernatant. The proportion of smurf worms increases with age. However, the evaluation of intestinal permeability can be performed at any stage and age, depending on the purpose of the study.</li>
	</ol>
	</li>
	<li>Running the assay
	<ol>
		<li>On the test day, collect L4 worms (2 days after step 2.1.1) and adulthood worms on 7th day (now in their 9th day) with M9 buffer (Supplementary Table 1) and transfer to labeled 1.5 mL microtubes.</li>
		<li>Centrifuge at 1400 x g for 3 min, remove the supernatant, add 1.0 mL of M9, and mix gently. Repeat this process 3x to remove the bacteria. After the last wash, reduce the volume to 500 &#181;L.</li>
		<li>Using an automatic pipette, add 10 &#181;L of the worm sediment to a microscope slide and count the number of worms. Calculate the volume to be pipetted to obtain 100 worms/&#181;L.</li>
		<li>Smurf staining: prepare a 25% Erioglaucine disodium salt (Supplementary Table 1) solution using distilled water. If necessary, use the vortex for better solubilization.</li>
		<li>In new previously identified microtubes, using an automatic pipette, add the volume containing 100 worms, 100 &#181;L of 25% Erioglaucine disodium salt solution, 200 &#181;L of E. coli OP50 and complete with M9 buffer for a final volume of 500 &#181;L. 
		NOTE: The volumes of Erioglaucine disodium, E. coli and number of worms are fixed. The volume of the M9 buffer is variable and can be used to bring the final volume to 500 &#181;L.</li>
		<li>Incubate for 3 h with agitation in a mixer, protected from light, at room temperature. 
		NOTE: If a mixer is unavailable, shake the microtubes carefully with your hands every 15 min.</li>
		<li>After 3 h, centrifuge the 1.5 mL microtube at 1400 x g for 3 min. Remove 1.0 mL of supernatant and wash the worms with 1.0 mL of M9 buffer.</li>
		<li>Repeat washes until the remaining Erioglaucine disodium salt solution is removed from the solution. After the last washing step, remove the supernatant except for a residual of approximately 250 &#181;L. Use this to carefully resuspend worms and process them for microscopic analysis.</li>
		<li>Transfer approximately 50 worms of resuspended worm solution to microscopy slides and cover them with cover slips. Incubate the microscopy slides in the refrigerator at -20 &#176;C for 10 min to paralyze the worms. 
		NOTE: Another alternative to paralyze the worms is to add 10 &#181;L of Levamisole hydrochloride solution (10 mM ; Supplementary Table 1) and cover them with cover slips.</li>
		<li>Observe and count the total number of worms and the totally dyed worms using the bright field in a stereomicroscope (at 1.5x). Images can be obtained from a cell phone camera (at 3.4x) coupled with an adapter. To prevent errors from light/brightness/exposure, it is crucial that all cell phone camera settings remain consistent for all pictures.</li>
	</ol>
	</li>
	<li>Data inspection
	<ol>
		<li>Qualitative data: Obtain images from stereomicroscope of each group (L4 and 7th day of adulthood) and visually compare the staining of the worms.</li>
		<li>Count the total number of worms and the number of smurf worms (blue worms) using a stereomicroscope.</li>
		<li>Express results as the percentage of smurf worms by comparing the young worms (L4, as a control group) and old worms (7th day of adulthood): 
		A x X = 100 (%) x B 
		X = 100 x B / A 
		Where, A = total number of worms 
		B= number of smurf worms</li>
	</ol>
	</li>
</ol>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/66084/66084fig02.jpg" /> 
Figure 2: Overall intestinal permeability assay schematic in C. elegans. (A) C. elegans preparation. (B) Staining with Erioglaucine disodium salt. <a href="https://www.jove.com/files/ftp_upload/66084/66084fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Evaluating Metabolism and Aging Pattern in Caenorhabditis elegans

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The authors have nothing to disclose.

In summary, this protocol provides a qualitative evaluation of glycogen content in individual C. elegans worms using Lugol staining: a straightforward, robust, and swift assay. Lugol staining is a label-free and non-invasive approach that facilitates the acquisition of molecular data at subcellular resolutions, allowing for the monitoring of glycogen content fluctuations within single worms10. Furthermore, the assay offers the advantage of cost-effectiveness due to its minimal equipment r...

Biological sciences teaching in high school is a continuous challenge. Notably, the access and use of technology have brought important advances in the teaching-learning process, however, tools such as artificial intelligence chatbots make rationalizing and seeking evidence more difficult due to easy (and sometimes incorrect) responses obtained1. Because of that, the use of a scientific method with practical experimentation in an inquiry-based approach in the classroom is an important strategy to develop or stimulate critical thinking, creativity, and technical skills in the students2.
In this context, the free living nematode Caenorhabditis elegans has been successfully used in experimentation for teaching purposes3 because of its particular advantages: It is not a parasite and the Escherichia coli used for feeding is biosafety level-1, therefore reducing near to zero the biological hazard; it has an elegant and quantifiable locomotion movement, which is interesting for students to observe; and it is transparent, which allows organ observation, but also staining with pigments that can indicate the presence of biomolecules or the occurrence of physiological alterations4. Therefore, it is possible to hypothesize and test in the classroom simple postulations related to biochemistry and physiological changes such as aging.
Glycogen is a storage carbohydrate, formed by a long and branched chain of glucose molecules formed by glucosyl residues with (1&#8594;4)-&#945; glycosidic linear linkages and (1&#8594;6)-&#945; glycosidic bonds at branch points and is particularly important for muscle contraction, cell differentiation and glycemia maintenance5. Glycogen is synthesized after feeding due to insulin activation of the enzyme glycogen-synthase. During exercise or fasting, epinephrine or glucagon, respectively, activate glycogen phosphorylase and, therefore, break down the polysaccharide to provide glucose-6-phosphate to the muscle cells or release free glucose to circumvent hypoglycemia6,7. Alterations in glycogen levels impacts cell differentiation, signaling, redox regulation, and stemness under various physiological and pathophysiological conditions, including cancer8. In C. elegans, glycogen is mainly found in esophageal muscle, hypodermis, intestine, neurons and mainly in body wall muscles9. The glycogen content can be measured by using Lugol&#39;s Iodine solution, since iodine binds into the helical coils forming an iodine-glycogen complex, giving a visible sharp blue-black or brown-black color, which has been successfully used to demonstrate glycogen content in C. elegans10. It has been demonstrated that glycogen accumulation caused by high glucose feeding can reduce worm&#39;s lifespan, therefore accelerating the aging process11,12. In addition, metabolic disturbances, other hormones, and exposure to xenobiotics can alter glycogen metabolism as well13,14. Therefore, experimentation on glycogen content in C. elegans is quite interesting, since diverse factors may disturb its metabolism and can stimulate an in-class discussion on basic biochemistry associated with transversal themes such as exercise, diets, diseases, and aging.
Aging is a time-dependent functional decline caused by cellular damage. This damage can be associated with oxidative stress, telomere attrition, loss of proteostasis, inflammation and even by accumulation of insoluble polyglucosan bodies15, just to name a few. One of the hallmarks of aging is the reduction of intestinal integrity, associated with several chronic conditions that occur during the life of an organism16. Maintenance of intestinal homeostasis depends on the integrity of the intestinal epithelium, which is supported by junctional proteins forming a physical barrier and connecting adjacent epithelial cells. When there is damage to this epithelium, leakage of luminal content into the interstitium occurs17. Based on this mechanism, the smurf test has been used to verify intestinal integrity in several animal models, since this blue dye Erioglaucine disodium salt does not cross the intestinal membrane, remaining in the lumen18. When worms are infected with a pathogen, contaminated with some toxicants or age, altering the interstitial integrity, the dye crosses the barrier and spreads all over the worm, which becomes all blue. This assay allows discussion on the physiology of aging and experimenting on factors that can accelerate or delay this process by exposing worms to different conditions. The protocols here will describe in detail these two, dye-based methods that can be easily done in class to instigate and stimulate students to formulate and test hypotheses related to biochemistry and physiology.
The first part of protocol shows its applicability to analyze qualitatively and quantitatively the glycogen content in C. elegans model10. The purpose of second part of the protocol is to assess the integrity of the C. elegans intestine. This technique allows for the monitoring of C. elegans aging by evaluating the integrity of the intestinal membranes. Furthermore, it allows evaluating whether a substance accelerates or delays aging and whether any substances have toxic potential on the intestinal barrier19.
The C. elegans strain used for the present study was Bristol N2 wild type. However, the procedure can be replicated using strains that exhibit comparable growth rates, or the method must be adjusted based on need of equipment replacement, considering they have the same or similar function, or depending on the strain used, as certain strains have specific maintenance and/or sensitivity requirements; this information can be obtained from the Caenorhabditis Genetics Center (CGC) or WormBase website. These changes should not impact the reproducibility of the method.

<table><tbody><tr><td>1.5 mL microtubes&amp;nbsp;</td><td>Local suppliers</td><td>-</td><td></td></tr><tr><td>37-degree incubator</td><td>KS 4000i&amp;nbsp;</td><td>97014-816</td><td></td></tr><tr><td>50 mL conical tube</td><td>Local suppliers</td><td>-</td><td></td></tr><tr><td>6 cm Petri plates&amp;nbsp;</td><td>Local suppliers</td><td>-</td><td></td></tr><tr><td>Agar bacteriological</td><td>Din&amp;acirc;mica Qu&amp;iacute;mica Contempor&amp;acirc;nea Ltda.</td><td>9002-18-0</td><td></td></tr><tr><td>C. elegans Bristol N2 (wild type)</td><td>Caenorhabditis elegans Genetic Center (CGC, Minnesota, USA)</td><td>-</td><td></td></tr><tr><td>CaCL2</td><td>Din&amp;acirc;mica Qu&amp;iacute;mica Contempor&amp;acirc;nea Ltda.</td><td>10035-04-8</td><td></td></tr><tr><td>Cholesterol</td><td>Sigma-Aldrich Brasil Ltda</td><td>57-88-5</td><td></td></tr><tr><td>D-(+)-Glucose anhydrous&amp;nbsp;</td><td>Neon</td><td>50-99-7</td><td></td></tr><tr><td>Distilled H2O</td><td>Local suppliers</td><td>-</td><td></td></tr><tr><td>Erioglaucine disodium salt</td><td>Sigma-Aldrich Brasil Ltda</td><td>3844-45-9</td><td></td></tr><tr><td>&lt;em&gt;Escherichia coli&lt;/em&gt; OP50</td><td>Caenorhabditis elegans Genetic Center (CGC, Minnesota, USA)</td><td>-</td><td></td></tr><tr><td>Flow hood</td><td>Mylabor</td><td></td><td></td></tr><tr><td>Incubator</td><td>Panasonic Healthcare company of North America, MIR-254-PA.</td><td>-</td><td></td></tr><tr><td>KH2PO4</td><td>Din&amp;acirc;mica Qu&amp;iacute;mica Contempor&amp;acirc;nea Ltda.</td><td>7778-77-0</td><td></td></tr><tr><td>Levamisole hydrochloride</td><td>RIPERCOL L 150F</td><td>-</td><td></td></tr><tr><td>Lugol solution</td><td>Sigma-Aldrich Brasil Ltda</td><td>L6146</td><td></td></tr><tr><td>MgSO4</td><td>Synth</td><td>S1063-01-AH</td><td></td></tr><tr><td>Microcentrifuge</td><td>Centrifuge 5425R Eppendorf SE, Germany</td><td></td><td></td></tr><tr><td>Na2HPO4</td><td>Din&amp;acirc;mica Qu&amp;iacute;mica Contempor&amp;acirc;nea Ltda.</td><td>7558-79-4</td><td></td></tr><tr><td>NaCl</td><td>Din&amp;acirc;mica Qu&amp;iacute;mica Contempor&amp;acirc;nea Ltda.</td><td>7647-14-5</td><td></td></tr><tr><td>Nystatin</td><td>Sigma-Aldrich Brasil Ltda</td><td>N6261</td><td></td></tr><tr><td>Peptone bacteriological</td><td>&amp;ecirc;xodo cient&amp;iacute;fica</td><td>91079-38-8</td><td></td></tr><tr><td>Stereomicroscope</td><td>Leica S8 Apo Stereomicroscope (S&amp;atilde;o Paulo, Brazil)</td><td></td><td></td></tr><tr><td>Streptomycin Sulfate</td><td>Estreptomax</td><td>-</td><td></td></tr></tbody></table>

using low-cost dyes to visualize glycogen accumulation and gut integrity in caenorhabditis elegans

Caenorhabditis elegans (C. elegans) is a transparent, non-parasitic nematode with a simple biology, which makes it a great tool for biological sciences teaching through the staining of the cells or their molecular content. Lugol dye (iodine-potassium iodide solution) has been widely used in biochemistry to stain glycogen stores. In this context, it is possible to observe differences between fed and starved animals, besides the effects of different conditions, such as different diets and oxygen levels. Erioglaucine is a blue dye that indicates the loss of the intestinal barrier. When the intestinal barrier is intact, the blue dye stains inside the lumen; however, when this integrity is disrupted, the dye leaks into the body cavity. Using a stereomicroscope or a microscope, teachers can demonstrate physiological and biochemical alterations, or they can instigate students to ask a scientific question and hypothesize and test their hypothesis using these assays. The present protocol describes two staining techniques in C. elegans that can be easily carried out by students.

The glycogen content assay provides a robust and rapid method for screening various testing conditions, such as comparative studies of different strains that may influence glycogen synthesis or degradation. In this study, L4 worms were subjected to three distinct test conditions: fasting, feeding, and glucose-enriched groups. The assay was performed three times, with each condition replicated twice in each assay; a representative image is displayed in Figure 3. Following each assay, worms st...

Read this Scientific Article Protocol about Using Low-cost Dyes to Visualize Glycogen Accumulation and Gut Integrity in Caenorhabditis elegans at JoVE.com

Author Spotlight: Exploring Metabolic and Aging Processes in C. elegans Using Low-Cost, High-Impact Assays

Teaching biological sciences can be made more stimulating for students through the use of experimentation. This manuscript presents two different yet complementary protocols that can be utilized in the classroom to encourage students to formulate and test hypotheses related to high-calorie diets, starvation, and aging.

Using Low-Cost Dyes to Visualize Glycogen Accumulation and Gut Integrity in Caenorhabditis elegans

Caenorhabditis elegans (C. elegans) is a transparent, non-parasitic nematode with a simple biology, which makes it a great tool for biological sciences ...

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Research

JoVE Journal

Biology

958 Views.  Federal University of Pampa. Teaching biological sciences can be made more stimulating for students through the use of experimentation. This manuscript presents two different yet complementary protocols that can be utilized in the classroom to encourage students to formulate and test hypotheses related to high-calorie diets, starvation, and aging.

Video: Author Spotlight: Exploring Metabolic and Aging Processes in C. elegans  Using Low-Cost, High-Impact Assays

Small-Scale Colorimetric Assays of Intracellular Lactate and Pyruvate in the Nematode Caenorhabditis elegans

Lactate and pyruvate are key intermediates of intracellular energy metabolic pathways. Monitoring the lactate/pyruvate ratio in cells helps to determine whether there is an imbalance in age-related energy metabolism between mitochondrial oxidative phosphorylation and aerobic glycolysis. Here, we show the utilization of commercial colorimetric assay kits for lactate and pyruvate in the model organism C. elegans. Recently, the sensitivity and accuracy of the colorimetric/fluorimetric assay kits have been improved greatly by the research and development conducted by reagent manufacturers. The improved reagents have enabled the use of small-scale assays with a 96-well plate in C. elegans. In general, a fluorimetric assay is superior in sensitivity to a colorimetric assay; however, the colorimetric approach is more suitable for the use in common laboratories. Another important issue in these assays for quantitative determination is protein precipitation of homogenized C. elegans samples. In our protein precipitation method, common precipitants (e.g., trichloroacetic acid, perchloric acid and metaphosphoric acid) are used for sample preparation. A protein-free assay sample is prepared by directly adding cold precipitant (final concentration of 5%) during homogenization.

Small-Scale Colorimetric Assays of Intracellular Lactate and Pyruvate in the Nematode Caenorhabditis elegans

We describe a modified small-scale extraction and colorimetric assays of lactate and pyruvate in the nematode C. elegans. When utilizing commercial assay kits, the technical development of their sensitivity and accuracy is important. Protein precipitation in extraction is the most critical step for the quantitative determination of intracellular metabolites.

Lactate and pyruvate are key intermediates of intracellular energy metabolic pathways. Monitoring the lactate/pyruvate ratio in cells helps to determine ...

Biochemistry

Plate-based Large-scale Cultivation of Caenorhabditis elegans: Sample Preparation for the Study of Metabolic Alterations in Diabetes

Culturing Caenorhabditis elegans (C. elegans) in a large-scale manner on agar plates can be time-consuming and difficult. This protocol describes a simple and inexpensive method to obtain a large number of animals for the isolation of proteins to proceed with a western blot, mass spectrometry, or further proteomics analyses. Furthermore, an increase of nematode numbers for immunostainings and the integration of multiple analyses under the same culturing conditions can easily be achieved. Additionally, a transfer between plates with different experimental conditions is facilitated. Common techniques in plate culture involve the transfer of a single C. elegans using a platinum wire and the transfer of populated agar chunks using a scalpel. However, with increasing nematode numbers, these techniques become overly time-consuming. This protocol describes the large-scale culture of C. elegans including numerous steps to minimize the impact of the sample preparation on the physiology of the worm. Fluid and shear stress can alter the lifespan of and metabolic processes in C. elegans, thus requiring a detailed description of the critical steps in order to retrieve reliable and reproducible results. C. elegans is a model organism, consisting of neuronal cells for up to one-third, but lacking blood vessels, thus providing the possibility to investigate solely neuronal alterations independent of vascular control. Recently, early neurodegeneration in diabetic retinopathy was found prior to vascular alterations. Thus, C. elegans is of special interest for studying general mechanisms of diabetic complications. For example, an increased formation of advanced glycation end products (AGEs) and reactive oxygen species (ROS) is observed, which are reproducibly found in C. elegans. Protocols to handle samples of adequate size for a broader spectrum of investigations are presented here, exemplified by the study of diabetes-induced biochemical alterations. In general, this protocol can be useful for studies requiring large C. elegans numbers and in which liquid culture is not suitable.

Plate-based Large-scale Cultivation of Caenorhabditis elegans: Sample Preparation for the Study of Metabolic Alterations in Diabetes

This protocol describes a method for the large-scale cultivation of Caenorhabditis elegans on solid media. As an alternative to liquid culture, this protocol allows obtaining parameters of different scales under plate-based cultivation. This increases the comparability of results by omitting the morphological and metabolic differences between liquid and solid media culture.

Culturing Caenorhabditis elegans (C. elegans) in a large-scale manner on agar plates can be time-consuming and difficult. This protocol describes a simple ...

Assessing Lysosomal Alkalinization in the Intestine of Live Caenorhabditis elegans

The nematode Caenorhabditis elegans (C. elegans) is a model system that is widely used to study longevity and developmental pathways. Such studies are facilitated by the transparency of the animal, the ability to do forward and reverse genetic assays, the relative ease of generating fluorescently labeled proteins, and the use of fluorescent dyes that can either be microinjected into the early embryo or incorporated into its food (E. coli strain OP50) to label cellular organelles (e.g. 9-diethylamino-5H-benzo(a)phenoxazine-5-one and (3-{2-[(1H,1&#39;H-2,2&#39;-bipyrrol-5-yl-kappaN(1))methylidene]-2H-pyrrol-5-yl-kappaN}-N-[2-(dimethylamino)ethyl]propanamidato)(difluoro)boron). Here, we present the use of a fluorescent pH-sensitive dye that stains intestinal lysosomes, providing a visual readout of dynamic, physiological changes in lysosomal acidity in live worms. This protocol does not measure lysosomal pH, but rather aims to establish a reliable method of assessing physiological relevant variations in lysosomal acidity. cDCFDA is a cell-permeant compound that is converted to the fluorescent fluorophore 5-(and-6)-carboxy-2&#39;,7&#39;-dichlorofluorescein (cDCF) upon hydrolysis by intracellular esterases. Protonation inside lysosomes traps cDCF in these organelles, where it accumulates. Due to its low pKa of 4.8, this dye has been used as a pH sensor in yeast. Here we describe the use of cDCFDA as a food supplement to assess the acidity of intestinal lysosomes in C. elegans. This technique allows for the detection of alkalinizing lysosomes in live animals, and has a broad range of experimental applications including studies on aging, autophagy, and lysosomal biogenesis.

Assessing Lysosomal Alkalinization in the Intestine of Live Caenorhabditis elegans

A step-by-step guide to probe loss of lysosomal acidity in the intestine of C. elegans using the pH-sensitive vital dye 5(6)-carboxy-2&#39;,7&#39;-dichlorofluorescein diacetate (cDCFDA)

The nematode Caenorhabditis elegans (C. elegans) is a model system that is widely used to study longevity and developmental pathways. Such studies are ...

Developmental Biology

DiI Dye-Filling as a Simple and Inexpensive Tool to Visualize Ciliated Sensory Neurons in C. elegans

C. elegans have long been used as a simple and accessible model to study neuronal structure and the many functions of the nervous system. Of the 302 neurons within the adult hermaphrodite nervous system, 60 are classified as ciliated sensory neurons. These neurons are central to a number of C. elegans behaviors, including but not limited to chemo-, mechano-, and osmosensing, male mating, and dauer formation. For several decades now, members of the C. elegans community have used the red fluorescent lipophilic dye DiI to visualize a subset of the ciliated sensory neurons that are directly exposed to the external environment. This dye enters the ciliated ends of the neurons and distributes in a relatively uniform pattern throughout the dendrites, cell bodies, and axons. This simple and powerful method makes an excellent first-pass tool to identify genetic mutants that impart structural or functional defects in ciliated sensory neurons. Here, we present a streamlined version of this staining method to visualize the eight pairs of amphid and two pairs of phasmid neurons that are environmentally exposed in C. elegans. We discuss tips for using this inexpensive method for imaging cellular dye-filling patterns in anesthetized animals.

DiI dye-filling is a method commonly used in C. elegans to visualize a subset of the ciliated sensory neurons, allowing for the identification of genetic mutations that alter sensory neuron structure or function.

C. elegans have long been used as a simple and accessible model to study neuronal structure and the many functions of the nervous system. Of the 302 ...

Genetics

Visualization of Caenorhabditis elegans Cuticular Structures Using the Lipophilic Vital Dye DiI

The cuticle of C. elegans is a highly resistant structure that surrounds the exterior of the animal1-4. The cuticle not only protects the animal from the environment, but also determines body shape and plays a role in motility4-6. Several layers secreted by epidermal cells comprise the cuticle, including an outermost lipid layer7.
Circumferential ridges in the cuticle called annuli pattern the length of the animal and are present during all stages of development8. Alae are longitudinal ridges that are present during specific stages of development, including L1, dauer, and adult stages2,9. Mutations in genes that affect cuticular collagen organization can alter cuticular structure and animal body morphology5,6,10,11. While cuticular imaging using compound microscopy with DIC optics is possible, current methods that highlight cuticular structures include fluorescent transgene expression12, antibody staining13, and electron microscopy1. Labeled wheat germ agglutinin (WGA) has also been used to visualize cuticular glycoproteins, but is limited in resolving finer cuticular structures14. Staining of cuticular surface using fluorescent dye has been observed, but never characterized in detail15. We present a method to visualize cuticle in live C. elegans using the red fluorescent lipophilic dye DiI (1,1'-dioctadecyl-3,3,3',3'-tetramethylindocarbocyanine perchlorate), which is commonly used in C. elegans to visualize environmentally exposed neurons. This optimized protocol for DiI staining is a simple, robust method for high resolution fluorescent visualization of annuli, alae, vulva, male tail, and hermaphrodite tail spike in C. elegans.

Visualization of Caenorhabditis elegans Cuticular Structures Using the Lipophilic Vital Dye DiI

We present a method to visualize cuticle in live C. elegans using the red fluorescent lipophilic dye DiI (1,1'-dioctadecyl-3,3,3',3'-tetramethylindocarbocyanine perchlorate), which is commonly used in C. elegans to visualize environmentally exposed neurons. With this optimized protocol, alae and annular cuticular structures are stained by DiI and observed using compound microscopy.

The cuticle of C. elegans is a highly resistant structure that surrounds the exterior of the animal1-4. The cuticle not only protects the animal from the ...

Biochemical and High Throughput Microscopic Assessment of Fat Mass in Caenorhabditis Elegans

The nematode C. elegans has emerged as an important model for the study of conserved genetic pathways regulating fat metabolism as it relates to human obesity and its associated pathologies. Several previous methodologies developed for the visualization of C. elegans triglyceride-rich fat stores have proven to be erroneous, highlighting cellular compartments other than lipid droplets. Other methods require specialized equipment, are time-consuming, or yield inconsistent results. We introduce a rapid, reproducible, fixative-based Nile red staining method for the accurate and rapid detection of neutral lipid droplets in C. elegans. A short fixation step in 40% isopropanol makes animals completely permeable to Nile red, which is then used to stain animals. Spectral properties of this lipophilic dye allow it to strongly and selectively fluoresce in the yellow-green spectrum only when in a lipid-rich environment, but not in more polar environments. Thus, lipid droplets can be visualized on a fluorescent microscope equipped with simple GFP imaging capability after only a brief Nile red staining step in isopropanol. The speed, affordability, and reproducibility of this protocol make it ideally suited for high throughput screens. We also demonstrate a paired method for the biochemical determination of triglycerides and phospholipids using gas chromatography mass-spectrometry. This more rigorous protocol should be used as confirmation of results obtained from the Nile red microscopic lipid determination. We anticipate that these techniques will become new standards in the field of C. elegans metabolic research.

Biochemical and High Throughput Microscopic Assessment of Fat Mass in Caenorhabditis Elegans

We present robust biochemical and microscopic methods for studying Caenorhabditis elegans lipid stores. A rapid, simple, fixing-staining procedure for fluorescent lipid droplet imaging leverages the spectral properties of the lipophilic dye Nile red. We then present biochemical measurement of triglycerides and phospholipids using solid phase extraction and gas chromatography-mass spectrometry.

The nematode C. elegans has emerged as an important  model for the study of conserved genetic pathways regulating fat metabolism as  it relates to human ...

Measuring the Effects of Bacteria and Chemicals on the Intestinal Permeability of Caenorhabditis elegans

In living organisms, intestinal hyperpermeability is a serious symptom that leads to many inflammatory bowel diseases (IBDs). Caenorhabditis elegans is a nonmammalian animal model that is widely used as an assay system due to its short lifespan, transparency, cost-effectiveness, and lack of animal ethics issues. In this study, a method was developed to investigate the effects of different bacteria and 3,3'-diindolylmethane (DIM) on the intestinal permeability of C. elegans with a high-throughput image analysis system. The worms were infected with different gut bacteria or cotreated with DIM for 48 h and fed with fluorescein isothiocyanate (FITC)-dextran overnight. Then, the intestinal permeability was examined by comparing the fluorescence images and the fluorescence intensity inside the worm bodies. This method may also have the potential to identify probiotic and pathogenic intestinal bacteria that affect intestinal permeability in the animal model and is effective for examining the effects of harmful or health-promoting chemicals on intestinal permeability and intestinal health. However, this protocol also has some considerable limitations at the genetic level, especially for determining which genes are altered to control illness, because this method is mostly used for phenotypic determination. In addition, this method is limited to determining exactly which pathogenic substrates cause inflammation or increase the permeability of the worms' intestines during infection. Therefore, further in-depth studies, including investigation of the molecular genetic mechanism using mutant bacteria and nematodes as well as chemical component analysis of bacteria, are required to fully evaluate the function of bacteria and chemicals in determining intestinal permeability.

Measuring the Effects of Bacteria and Chemicals on the Intestinal Permeability of Caenorhabditis elegans

This protocol describes how to measure intestinal permeability of Caenorhabditis elegans. This method is helpful for basic biological research on intestinal health related to the interaction between intestinal bacteria and their host and for screening to identify probiotic and chemical agents to cure leaky gut syndrome and inflammatory bowel diseases.

In living organisms, intestinal hyperpermeability is a serious symptom that leads to many inflammatory bowel diseases (IBDs). Caenorhabditis elegans is a ...

Medicine

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 ...

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.

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.

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

Enhancing Axonal Transport Analysis in Living C. elegans Using GFP-Tagged RAB-3

Optimizing Visualization of Axonal Transport of Endogenous Cargo by Fluorescence Microscopy in Living Caenorhabditis elegans

Axonal transport is a prerequisite to deliver axonal proteins from their site of synthesis in the neuronal cell body to their destination in the axon. Consequently, loss of axonal transport impairs neuronal growth and function. Studying axonal transport therefore improves our understanding of neuronal cell biology. With recent improvements in CRISPR Cas9 genome editing, endogenous labeling of axonal cargos has become accessible, enabling to move beyond ectopic expression-based visualization of transport. However, endogenous labeling often comes at the cost of low signal intensity and necessitates optimization strategies to obtain robust data. Here, we describe a protocol to optimize the visualization of axonal transport by discussing acquisition parameters and a bleaching approach to improve the signal of endogenous labeled cargo over diffuse cytoplasmic background. We apply our protocol to optimize the visualization of synaptic vesicle precursors (SVPs) labeled by green fluorescent protein (GFP)-tagged RAB-3 to highlight how fine-tuning acquisition parameters can improve the analysis of endogenously labeled axonal cargo in Caenorhabditis elegans (C. elegans).

Author Spotlight: Advanced Techniques for Visualizing Endogenous Axonal Transport Dynamics

The paper describes the optimization of fluorescence microscopy acquisition parameters to visualize the axonal transport of endogenous labeled cargos at single-neuron resolution in a living nematode.

Axonal transport is a prerequisite to deliver axonal proteins from their site of synthesis in the neuronal cell body to their destination in the axon. ...

Using Low-Cost Dyes to Visualize Glycogen Accumulation and Gut Integrity in Caenorhabditis elegans

Summary

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Small-Scale Colorimetric Assays of Intracellular Lactate and Pyruvate in the Nematode Caenorhabditis elegans

Plate-based Large-scale Cultivation of Caenorhabditis elegans: Sample Preparation for the Study of Metabolic Alterations in Diabetes

Assessing Lysosomal Alkalinization in the Intestine of Live Caenorhabditis elegans

DiI Dye-Filling as a Simple and Inexpensive Tool to Visualize Ciliated Sensory Neurons in C. elegans

Visualization of Caenorhabditis elegans Cuticular Structures Using the Lipophilic Vital Dye DiI

Biochemical and High Throughput Microscopic Assessment of Fat Mass in Caenorhabditis Elegans

Measuring the Effects of Bacteria and Chemicals on the Intestinal Permeability of Caenorhabditis elegans

Methods to Assess Subcellular Compartments of Muscle in C. elegans

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Optimizing Visualization of Axonal Transport of Endogenous Cargo by Fluorescence Microscopy in Living Caenorhabditis elegans