The authors would like to acknowledge Dr. Kevin Bittman for his guidance in establishing the Seahorse XFp in the lab. National Institutes of Health grant GM088213 supported this work.

NOTE: Figure 2 provides a schematic overview of the full protocol.
1. Growth and synchronization of nematode population9,10
<ol>
	<li>Transfer L4 larvae of desired genetic backgrounds (e.g., N2 [wild type] and sel-12 animals) onto nematode growth media (NGM) plates (see Table 1 for recipe) freshly seeded with a lawn of Escherichia coli (OP50)11. Use at least two 100 mm or three 60 mm plates for each strain. Incubate the worms at the appropriate growth temperature (between 15&#8211;25 &#176;C) for 3&#8211;4 days or until plates are concentrated with large number of eggs and gravid worms.</li>
	<li>Wash the eggs and worms off the plates using approximately 6 mL of M9 buffer (see Table 1 for recipe) per 100 mm plate of nematodes and transfer them with glass Pasteur pipettes into individual 15 mL centrifuge tubes for each strain. Spin these tubes down for 3 min at 6,180 x g and aspirate out the M9 buffer, retaining just the animals and eggs pellet.</li>
	<li>Add 3&#8211;4 mL of bleach solution (see Table 1 for recipe) to each tube and intermittently vortex for 6 min. Add M9 buffer to fill each tube and spin at 6,180 x g for 1 min and aspirate supernatant. Repeat this wash with M9 three times and move the egg pellet to a fresh 15 mL tube containing approximately 9 mL of M9 buffer.</li>
	<li>Synchronize the freshly hatched worms by nutating at 20 &#176;C for 16&#8211;48 h. Spin these tubes down at 6,180 x g for 1.5 min and put the synchronized L1 animals down on individual NGM plates freshly seeded with a lawn of OP50 (approximately 6,000&#8211;10,000 animals per 100 mm plate) and keep at 20 &#176;C.</li>
	<li>These animals will reach the L4 larval stage after ~42 h. At this time, move the L4 larvae using a platinum pick to OP50 seeded NGM plates containing 0.5 mg/mL 5-fluoro-2&#8217;-deoxyuridine (FUdR) to prevent them from producing progeny. 
	NOTE: Make sure that within an experiment all strains are synchronized for a similar amount of time. FUdR treatment sterilizes the animals and prevents egg laying and progeny production, which could influence OCR. These sterilized animals will be analyzed the next day (day 1 adult animals at ~66 h) for the assay. FUdR has been reported to impact physiology and lifespan of certain mutant worms. Therefore, this should be taken into consideration when the drug is used for sterilization12,13,14. Worms can also be sterilized by the means of feminizing mutations such as fem-1(hc17) or fem-3(e2006). However, these mutations might impact mitochondrial function.</li>
</ol>
2. Preparation&#160;of compounds to be injected and hydration of probes
NOTE: During the assay run, both basal and maximal respiration rates of the nematodes are measured. Maximal respiration is triggered in the animals upon the addition of carbonyl cyanide-4 (trifluormethoxy)phenylhydrazone (FCCP), an uncoupling ionophore that disturbs the mitochondrial membrane potential and thus ATP synthesis by transporting protons through the mitochondrial membrane, while allowing proton pumping, electron transport, and oxygen consumption to proceed4,15 uncoupled from ATP synthesis (Figure 1). The final step in the assay involves the addition of sodium azide (NaN3), a drug that inhibits complexes IV and V in the ETC, allowing one to determine non-mitochondrial respiration16 (Figure 1). The following steps can be performed the day before the actual assay run.
<ol>
	<li>Prepare 1 mL stock solutions of the FCCP (10 mM in dimethyl sulfoxide [DMSO]: 1000x the final assay concentration) and NaN3 (400 mM in dH2O: 10x final assay concentration) and store at -20 &#176;C. 
	NOTE: Run a concentration curve to optimize the concentration of FCCP required to elicit maximal OCR response for each instrument and laboratory setting.</li>
	<li>Hydrate the sensor cartridge by adding 200 &#181;L of dH2O to each well (and the surrounding reservoir) in the plate, ensuring that the sensor probes are submerged in the dH2O and store overnight at room temperature. 
	NOTE: The cartridge probes can be left submerged for up to 72 h but in the case of a prolonged hydration, the plates should be wrapped in paraffin film and stored at 4 &#176;C. If overnight hydration is not possible, the sensor cartridge should be hydrated for at least 4 h before assay.</li>
	<li>Turn off the heating element within the respirometer interface and store the instrument inside a 15 &#176;C incubator overnight to lower core temperature within the instrument to prevent the animals from overheating during the assay run. 
	NOTE: The respirometer is equipped with a heating element but cannot be cooled. Within a 15 &#176;C incubator, the respirometer will have a stable temperature between 18&#8211;22 &#176;C, which is a healthy temperature for C. elegans maintenance.</li>
</ol>
3. Drug loading and cartridge equilibration
<ol>
	<li>Pipette out and discard the dH2O used to hydrate the sensor probes and replace it with 200 &#181;L of the calibrant solution (pH 7.4) in each well. Dilute the FCCP stock solution to 100 &#181;M FCCP in dH2O and add 20 &#181;L of the diluted solution to the injection port A in the sensor cartridge. Add 22 &#181;L of 400 mM sodium azide to port B of the senor cartridge.</li>
	<li>Turn the respirometer on and on the home screen select Start; the templates page will appear. On the templates page, select Blank or a previously designed template; the Groups page will appear. On the Groups page, select the wells A and H as the background wells and assign the remaining 6 wells into appropriate groups according to the experimental plan.</li>
	<li>Push the arrow on the lower right corner to go to the Protocol page. On the Protocol page, ensure that the Equilibrate, Basal and Injections 1 and 2 buttons are selected. On this page, adjust the number of readings of basal OCR, as well as maximal OCR (after injection 1 with FCCP) and non-mitochondrial OCR (after injection 2 with sodium azide). 
	NOTE: Each measurement is preceded by a mix and wait step and the time frames for these parameters are shown in Table 2.</li>
	<li>Select the arrow on the lower right corner and a prompt to load the sensor cartridge plate will appear. Ensure that the sensor cartridge plate is loaded in the right orientation, following the instructions on the reminder prompt screen. The respirometer will now equilibrate the cartridge. 
	NOTE: This equilibration provides ample time to place the animals to be assayed into appropriate wells of the cell plate.</li>
</ol>
4. Preparation of worm plate and assay run
<ol>
	<li>Add 200 &#181;L of M9 buffer into each of the 8 wells of the cell plate and into the reservoirs surrounding the wells.</li>
	<li>Pick ~100 worms from each strain onto unseeded NGM plates and allow to rest for 2-3 min. Wet the end of the platinum pick with M9 buffer and pick 20 age-synchronized animals into wells B-G, leaving the background wells A and H empty. 
	NOTE: After loading each well, wait approximately 2 min to allow the animals to settle into the bottom of the wells. It is also advisable that a worm number curve be run to optimize the worm number per well for each instrument and laboratory setting.</li>
	<li>By now, the respirometer should be calibrated and by clicking OK on the screen, the plate containing calibration buffer is ejected and can be removed from the respirometer, while the sensor cartridge stays inside the instrument. Replace the calibrant plate with the plate containing animals. Load plate and close the door by hitting Continue and allow the assay to run.</li>
</ol>
5. Post-experiment data analysis
<ol>
	<li>Once the assay run is complete, follow the prompts on the screen and remove the cell plate and sensor cartridge and insert a flash drive into the USB port to save the run data in the .war format. Remove the sensor cartridge and allow the animals to settle to the bottom of the wells for approximately 2 min. Place the cell plates under a stereo dissecting microscope and count the number of animals per well.</li>
	<li>Open the analysis software on the computer and open the normalization tab to normalize OCR to animal number. Apply appropriate labels for the various groups under the modify tab and export the file as a Prism file for further analysis. 
	NOTE: The average of the first five measurements, before the addition of FCCP is the basal OCR, while the average of the five measurements after FCCP addition is the maximal OCR and the average of the last five measurements (minimum of two measurements should be done) after sodium azide addition is the non-mitochondrial respiration rate. The assay should be repeated three times to ensure reproducibility.</li>
</ol>

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The authors have nothing to disclose.

Mitochondrial respiration is an insightful indicator of mitochondrial function; therefore, being able to measure the oxygen consumption rates in a biological system, whether in vitro or in vivo is highly valuable. Respirometers sense oxygen levels using porphyrin-based phosphors that get quenched by oxygen or via amperometric oxygen sensors that rely on the generation of an electric current proportional to oxygen pressure. Clark electrode falls into the latter category and has been used extensively in literature, especia...

Adenosine triphosphate (ATP), the main source of cellular energy, is produced in the mitochondria by enzymes in the electron transport chain (ETC) located in the inner mitochondrial membrane. Pyruvate, a key metabolite utilized for mitochondrial ATP production, is imported into the mitochondrial matrix where it is decarboxylated to produce acetyl coenzyme A (CoA). Subsequently, acetyl CoA enters the citric acid cycle resulting in the generation of nicotinamide adenine dinucleotide (NADH), a key electron carrier molecule. As electrons from NADH are passed to oxygen via the ETC, protons build up in the mitochondrial intermembrane space, which results in the generation of an electrochemical gradient across the membrane. These protons will then flow from the intermembrane space across this electrochemical gradient back into the mitochondrial matrix through the proton pore of the ATP synthase, driving its rotation and the synthesis of ATP1 (Figure 1).
Mitochondrial function is not limited to energy production but is also crucial for calcium homeostasis, reactive oxygen species (ROS) scavenging, and apoptosis, critically positioning their function in organismal health2. Mitochondrial function can be assessed using a variety of assays, including but not limited to analyses that measure mitochondrial membrane potential, ATP and ROS levels, and mitochondrial calcium concentrations. However, these assays provide a single snapshot of mitochondrial function and therefore might not provide a comprehensive view of mitochondrial health. Since oxygen consumption during ATP generation is reliant on a myriad of sequential reactions, it serves as a superior indicator of mitochondrial function. Interestingly, variations in oxygen consumption rates have been observed as a result of mitochondrial dysfunction3,4,5.
Oxygen consumption rates (OCR) of living samples can be measured using techniques that can be broadly divided into two groups: amperometric oxygen sensors and porphyrin-based phosphors that can be quenched by oxygen6. Amperometric oxygen sensors have been used extensively to measure OCR in cultured cells, tissues, and in model systems, such as C. elegans. However, porphyrin-based phosphors containing respirometers possess the following advantages: (1) they allow for a side by side comparison of two samples in triplicate, (2) they require smaller sample size (e.g., 20 worms per well versus ~2,000&#8722;5,000 worms in the chamber)7, and (3) the respirometer can be programmed to do four different compound injections at desired times throughout the experimental run, eliminating the need for manual application.
In this protocol, steps involved in using a porphyrin-based oxygen-sensing respirometer to measure OCR in live, intact C. elegans are described. While there is a written protocol for the use of the large format, high throughput respirometer8, this protocol has been adapted for use with a more budget friendly, accessible, and smaller scale instrument. This protocol is particularly useful for assessing the difference in OCR between two strains, where high-throughput screening is not required and its use would be excessive.

<table><tbody><tr><td>100 mm, 60 mm Petri dishes</td><td>Kord-Valmark Labware Products</td><td>2900, 2901</td><td></td></tr><tr><td>1.5 mL centrifuge tubes</td><td>Globe Scientific</td><td>6285</td><td></td></tr><tr><td>15 mL conical tubes</td><td>Corning</td><td>430791</td><td></td></tr><tr><td>22 &amp;times; 22 mm coverslip</td><td>Globe Scientific</td><td>1404-10</td><td></td></tr><tr><td>50 mL conical tubes</td><td>Corning</td><td>430829</td><td></td></tr><tr><td>Agar</td><td>Fisher Scientific</td><td>BP1423-2</td><td></td></tr><tr><td>Bacto peptone</td><td>BD, Bacto</td><td>211677</td><td></td></tr><tr><td>Bacto tryptone</td><td>BD, Bacto</td><td>211705</td><td></td></tr><tr><td>Bacto yeast extract</td><td>BD, Bacto</td><td>212705</td><td></td></tr><tr><td>Bleach</td><td>Generic</td><td></td><td></td></tr><tr><td>Calcium chloride dihydrate (CaCl2&amp;middot;2H&lt;sub&gt;2&lt;/sub&gt;O)</td><td>Fisher Scientific</td><td>C79-500</td><td></td></tr><tr><td>Carbonyl cyanide 4-(trifluoromethoxy)phenylhydrazone (FCCP)</td><td>Abcam</td><td>ab120081</td><td></td></tr><tr><td>Cholesterol</td><td>Fisher Scientific</td><td>C314-500</td><td></td></tr><tr><td>Deionized water (dH&lt;sub&gt;2&lt;/sub&gt;O)</td><td></td><td></td><td></td></tr><tr><td>Dimethyl sulfoxide (DMSO)</td><td>Thomas Scientific</td><td>C987Y85</td><td></td></tr><tr><td>Glass Pasteur pipettes</td><td>Krackeler Scientific</td><td>6-72050-900</td><td></td></tr><tr><td>Magnesium sulfate heptahydrate (MgSO4&amp;middot;7H2O)</td><td>Fisher Scientific</td><td>BP213-1</td><td></td></tr><tr><td>Potassium phosphate dibasic (K2HPO4)</td><td>Fisher Scientific</td><td>BP363-1</td><td></td></tr><tr><td>Potassium phosphate monobasic (KH2PO4)</td><td>Fisher Scientific</td><td>P285-500</td><td></td></tr><tr><td>Sodium chloride</td><td>Fisher Scientific</td><td>BP358-10</td><td></td></tr><tr><td>Sodium hydroxide (NaOH)</td><td>Fisher Scientific</td><td>BP359-500</td><td></td></tr><tr><td>Sodium phosphate dibasic anhydrous (Na2HPO4)</td><td>Fisher Scientific</td><td>BP332-1</td><td></td></tr><tr><td>Seahorse XFp Analyzer</td><td>Agilent</td><td></td><td></td></tr><tr><td>Seahorse XFp FluxPak</td><td>Agilent</td><td>103022-100</td><td></td></tr><tr><td>Sodium Azide</td><td>Sigma-Aldrich</td><td>S2002</td><td></td></tr></tbody></table>

measurement of oxygen consumption rates in intact caenorhabditis elegans

Optimal mitochondrial function is critical for healthy cellular activity, particularly in cells that have high energy demands like those in the nervous system and muscle. Consistent with this, mitochondrial dysfunction has been associated with a myriad of neurodegenerative diseases and aging in general. Caenorhabditis elegans have been a powerful model system for elucidating the many intricacies of mitochondrial function. Mitochondrial respiration is a strong indicator of mitochondrial function and recently developed respirometers offer a state-of-the-art platform to measure respiration in cells. In this protocol, we provide a technique to analyze live, intact C. elegans. This protocol spans a period of ~7 days and includes steps for (1) growing and synchronization of C. elegans, (2) preparation&#160;of compounds to be injected and hydration of probes, (3) drug loading and cartridge equilibration, (4) preparation of worm assay plate and assay run, and (5) post-experiment data analysis.

Using the protocol described herein, OCR of wild type animals and three different sel-12 mutant strains were determined. sel-12 encodes the C. elegans ortholog of presenilin17. Mutations in human presenilin are the most common genetic aberration associated with the development of familial Alzheimer&#39;s disease18. Our studies have shown elevated mitochondrial calcium levels in sel-12 mutant animals compar...

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Measurement of Oxygen Consumption Rates in Intact Caenorhabditis elegans

Mitochondrial respiration is critical for organismal survival; therefore, oxygen consumption rate is an excellent indicator of mitochondrial health. In this protocol, we describe the use of a commercially available respirometer to measure basal and maximal oxygen consumption rates in live, intact, and freely-motile Caenorhabditis elegans.

Measurement of Oxygen Consumption Rates in Intact Caenorhabditis elegans

Optimal mitochondrial function is critical for healthy cellular activity, particularly in cells that have high energy demands like those in the nervous ...

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8.9K Views.  Albany Medical College. Mitochondrial respiration is critical for organismal survival; therefore, oxygen consumption rate is an excellent indicator of mitochondrial health. In this protocol, we describe the use of a commercially available respirometer to measure basal and maximal oxygen consumption rates in live, intact, and freely-motile Caenorhabditis elegans.

Video: Measurement of Oxygen Consumption Rates in Intact Caenorhabditis elegans

Measurement of Mitochondrial Oxygen Consumption in Permeabilized Fibers of Drosophila Using Minimal Amounts of Tissue

The fruit fly, Drosophila melanogaster, represents an emerging model for the study of metabolism. Indeed, drosophila have structures homologous to human organs, possess highly conserved metabolic pathways and have a relatively short lifespan that allows the study of different fundamental mechanisms in a short period of time. It is, however, surprising that one of the mechanisms essential for cellular metabolism, the mitochondrial respiration, has not been thoroughly investigated in this model. It is likely because the measure of the mitochondrial respiration in Drosophila usually requires a very large number of individuals and the results obtained are not highly reproducible. Here, a method allowing the precise measurement of mitochondrial oxygen consumption using minimal amounts of tissue from Drosophila is described. In this method, the thoraxes are dissected and permeabilized both mechanically with sharp forceps and chemically with saponin, allowing different compounds to cross the cell membrane and modulate the mitochondrial respiration. After permeabilization, a protocol is performed to evaluate the capacity of the different complexes of the electron transport system (ETS) to oxidize different substrates, as well as their response to an uncoupler and to several inhibitors. This method presents many advantages compared to methods using mitochondrial isolations, as it is more physiologically relevant because the mitochondria are still interacting with the other cellular components and the mitochondrial morphology is conserved. Moreover, sample preparations are faster, and the results obtained are highly reproducible. By combining the advantages of Drosophila as a model for the study of metabolism with the evaluation of mitochondrial respiration, important new insights can be unveiled, especially when the flies are experiencing different environmental or pathophysiological conditions.

In this paper, a method to measure oxygen consumption using high-resolution respirometry in permeabilized thoraxes of Drosophila is described. This technique requires a minimal amount of tissue compared to the classic mitochondrial isolation technique and the results obtained are more physiologically relevant.

The fruit fly, Drosophila melanogaster, represents an emerging model for the study of metabolism. Indeed, drosophila have structures homologous to human ...

Biochemistry

Comparative Analysis of Experimental Methods to Quantify Animal Activity in Caenorhabditis elegans Models of Mitochondrial Disease

Caenorhabditis elegans is widely recognized for its central utility as a translational animal model to efficiently interrogate mechanisms and therapies of diverse human diseases. Worms are particularly well-suited for high-throughput genetic and drug screens to gain deeper insight into therapeutic targets and therapies by exploiting their fast development cycle, large brood size, short lifespan, microscopic transparency, low maintenance costs, robust suite of genomic tools, mutant repositories, and experimental methodologies to interrogate both in vivo and ex vivo physiology. Worm locomotor activity represents a particularly relevant phenotype that is frequently impaired in mitochondrial disease, which is highly heterogeneous in causes and manifestations but collectively shares an impaired capacity to produce cellular energy. While a suite of different methodologies may be used to interrogate worm behavior, these vary greatly in experimental costs, complexity, and utility for genomic or drug high-throughput screens. Here, the relative throughput, advantages, and limitations of 16 different activity analysis methodologies were compared that quantify nematode locomotion, thrashing, pharyngeal pumping, and/or chemotaxis in single worms or worm populations of C. elegans at different stages, ages, and experimental durations. Detailed protocols were demonstrated for two semi-automated methods to quantify nematode locomotor activity that represent novel applications of available software tools, namely, ZebraLab (a medium-throughput approach) and WormScan (a high-throughput approach). Data from applying these methods demonstrated similar degrees of reduced animal activity occurred at the L4 larval stage, and progressed in day 1 adults, in mitochondrial complex I disease (gas-1(fc21)) mutant worms relative to wild-type (N2 Bristol) C. elegans. This data validates the utility for these novel applications of using the ZebraLab or WormScan software tools to quantify worm locomotor activity efficiently and objectively, with variable capacity to support high-throughput drug screening on worm behavior in preclinical animal models of mitochondrial disease.

Comparative Analysis of Experimental Methods to Quantify Animal Activity in Caenorhabditis elegans Models of Mitochondrial Disease

This study presents protocols for two semi-automated locomotor activity analysis approaches in C. elegans complex I disease gas-1(fc21) worms, namely, ZebraLab (a medium-throughput assay) and WormScan (a high-throughput assay) and provide comparative analysis among a wide array of research methods to quantify nematode behavior and integrated neuromuscular function.

Caenorhabditis elegans is widely recognized for its central utility as a translational animal model to efficiently interrogate mechanisms and therapies of ...

Behavior

High-Throughput Assay to Measure Caenorhabditis elegans Food Intake and Lifespan

Quantifying Food Intake in Caenorhabditis elegans by Measuring Bacterial Clearance

Feeding is an essential biological process for an organism&#39;s growth, reproduction, and survival. This assay aims to measure the food intake of Caenorhabditis elegans&#160;(C. elegans), an important parameter when studying the genetics of aging or metabolism. In most species, feeding is determined by measuring the difference between the amount of food provided and the amount left after a given time interval. The method presented here uses the same strategy to determine the feeding of C. elegans. It measures the amount of bacteria, the food source of C. elegans, cleared within 72 h. This method uses 96-well microtiter plates and has allowed the screening of hundreds of drugs for their ability to modulate food intake at a speed and depth not possible in other animal models. The strength of this assay is that it allows to measure feeding and lifespan simultaneously and directly measures the disappearance of food and, thus, is based on the same principles used for other organisms, facilitating species-to-species comparison.

Author Spotlight: Exploring the Relationship Between Lipotoxicity and HFpEF 

This protocol describes an assay to quantify Caenorhabditis elegans feeding rate based on measuring the clearance of bacteria in liquid culture.

Feeding is an essential biological process for an organism&#39;s growth, reproduction, and survival. This assay aims to measure the food intake of ...

An Assay for Measuring the Effects of Ethanol on the Locomotion Speed of Caenorhabditis elegans

Alcohol use disorders are a significant public health concern, for which there are few effective treatment strategies. One difficulty that has delayed the development of more effective treatments is the relative lack of understanding of the molecular underpinnings of the effects of ethanol on behavior. The nematode, Caenorhabditis elegans (C.&#160;elegans), provides a useful model in which to generate and test hypotheses about the molecular effects of ethanol. Here, we describe an assay that has been developed and used to examine the roles of particular genes and environmental factors in behavioral responses to ethanol, in which locomotion is the behavioral output. Ethanol dose-dependently causes an acute depression of crawling on an agar surface. The effects are dynamic; animals exposed to a high concentration demonstrate an initial strong depression of crawling, referred to here as initial sensitivity, and then partially recover locomotion speed despite the continued presence of the drug. This ethanol-induced behavioral plasticity is referred to here as the development of acute functional tolerance. This assay has been used to demonstrate that these two phenotypes are distinct and genetically separable. The straightforward locomotion assay described here is suitable for examining the effects of both genetic and environmental manipulations on these acute behavioral responses to ethanol in C.&#160;elegans.

An Assay for Measuring the Effects of Ethanol on the Locomotion Speed of Caenorhabditis elegans

C. elegans is a useful model for studying the effects of ethanol on behavior. We present a behavioral assay that quantifies the effects of ethanol on the locomotion speed of crawling worms; both initial sensitivity and the development of acute functional tolerance to ethanol can be measured with this assay.

Alcohol use disorders are a significant public health concern, for which there are few effective treatment strategies. One difficulty that has delayed the ...

Neuroscience

Isolation and In vitro Activation of Caenorhabditis elegans Sperm

Males and hermaphrodites are the two naturally found sexual forms in the nematode C. elegans. The amoeboid sperm are produced by both males and hermaphrodites. In the earlier phase of gametogenesis, the germ cells of hermaphrodites differentiate into limited number of sperm - around 300 - and are stored in a small 'bag' called the spermatheca. Later on, hermaphrodites continually produce oocytes1. In contrast, males produce exclusively sperm throughout their adulthood. The males produce so much sperm that it accounts for &gt;50% of the total cells in a typical adult worm2. Therefore, isolating sperm from males is easier than from that of hermaphrodites.
Only a small proportion of males are naturally generated due to spontaneous non-disjunction of X chromosome3. Crossing hermaphrodites with males or more conveniently, the introduction of mutations to give rise to Him (High Incidence of Males) phenotype are some of strategies through which one can enrich the male population3.
Males can be easily distinguished from hermaphrodites by observing the tail morphology4. Hermaphrodite's tail is pointed, whereas male tail is rounded with mating structures.
Cutting the tail releases vast number of spermatids stored inside the male reproductive tract. Dissection is performed under a stereo microscope using 27 gauge needles. Since spermatids are not physically connected with any other cells, hydraulic pressure expels internal contents of male body, including spermatids2.
Males are directly dissected on a small drop of 'Sperm Medium'. Spermatids are sensitive to alteration in the pH. Hence, HEPES, a compound with good buffering capacity is used in sperm media. Glucose and other salts present in sperm media help maintain osmotic pressure to maintain the integrity of sperm.
Post-meiotic differentiation of spermatids into spermatozoa is termed spermiogenesis or sperm activation. Shakes5, and Nelson6 previously showed that round spermatids can be induced to differentiate into spermatozoa by adding various activating compounds including Pronase E. Here we demonstrate in vitro spermiogenesis of C. elegans spermatids using Pronase E.
Successful spermiogenesis is pre-requisite for fertility and hence the mutants defective in spermiogenesis are sterile. Hitherto several mutants have been shown to be defective specifically in spermiogenesis process7. Abnormality found during in vitro activation of novel Spe (Spermatogenesis defective) mutants would help us discover additional players participating in this event.

Isolation and In vitro Activation of Caenorhabditis elegans Sperm

A protocol for isolating and activating spermatids from male C. elegans is described here. Cutting the posterior end of male releases spermatids. The spermatids can be activated by addition of protease.

Males and hermaphrodites are the two naturally found sexual forms in the nematode C. elegans. The amoeboid sperm are produced by both males and ...

Solid Plate-based Dietary Restriction in Caenorhabditis elegans

Reduction of food intake without malnutrition or starvation is known to increase lifespan and delay the onset of various age-related diseases in a wide range of species, including mammals. It also causes a decrease in body weight and fertility, as well as lower levels of plasma glucose, insulin, and IGF-1 in these animals. This treatment is often referred to as dietary restriction (DR) or caloric restriction (CR). The nematode Caenorhabditis elegans has emerged as an important model organism for studying the biology of aging. Both environmental and genetic manipulations have been used to model DR and have shown to extend lifespan in C. elegans. However, many of the reported DR studies in C. elegans were done by propagating animals in liquid media, while most of the genetic studies in the aging field were done on the standard solid agar in petri plates. Here we present a DR protocol using standard solid NGM agar-based plate with killed bacteria.

Solid Plate-based Dietary Restriction in Caenorhabditis elegans

Here we present a protocol for performing solid plate-based dietary restriction in C. elegans with killed bacteria.

Reduction of food intake without malnutrition or starvation is known to increase lifespan and delay the onset of various age-related diseases in a wide ...

Creating Defined Gaseous Environments to Study the Effects of Hypoxia on C. elegans

Oxygen is essential for all metazoans to survive, with one known exception1. Decreased O2 availability (hypoxia) can arise during states of disease, normal development or changes in environmental conditions2-5. Understanding the cellular signaling pathways that are involved in the response to hypoxia could provide new insight into treatment strategies for diverse human pathologies, from stroke to cancer. This goal has been impeded, at least in part, by technical difficulties associated with controlled hypoxic exposure in genetically amenable model organisms.
The nematode Caenorhabditis elegans is ideally suited as a model organism for the study of hypoxic response, as it is easy to culture and genetically manipulate. Moreover, it is possible to study cellular responses to specific hypoxic O2 concentrations without confounding effects since C. elegans obtain O2 (and other gasses) by diffusion, as opposed to a facilitated respiratory system6. Factors known to be involved in the response to hypoxia are conserved in C. elegans. The actual response to hypoxia depends on the specific concentration of O2 that is available. In C. elegans, exposure to moderate hypoxia elicits a transcriptional response mediated largely by hif-1, the highly-conserved hypoxia-inducible transcription factor6-9. C .elegans embryos require hif-1 to survive in 5,000-20,000 ppm O27,10. Hypoxia is a general term for "less than normal O2". Normoxia (normal O2) can also be difficult to define. We generally consider room air, which is 210,000 ppm O2 to be normoxia. However, it has been shown that C. elegans has a behavioral preference for O2 concentrations from 5-12% (50,000-120,000 ppm O2)11. In larvae and adults, hif-1 acts to prevent hypoxia-induced diapause in 5,000 ppm O212. However, hif-1 does not play a role in the response to lower concentrations of O2 (anoxia, operational definition &lt;10 ppm O2)13. In anoxia, C. elegans enters into a reversible state of suspended animation in which all microscopically observable activity ceases10. The fact that different physiological responses occur in different conditions highlights the importance of having experimental control over the hypoxic concentration of O2.
Here, we present a method for the construction and implementation of environmental chambers that produce reliable and reproducible hypoxic conditions with defined concentrations of O2. The continual flow method ensures rapid equilibration of the chamber and increases the stability of the system. Additionally, the transparency and accessibility of the chambers allow for direct visualization of animals being exposed to hypoxia. We further demonstrate an effective method of harvesting C. elegans samples rapidly after exposure to hypoxia, which is necessary to observe many of the rapidly-reversed changes that occur in hypoxia10,14. This method provides a basic foundation that can be easily modified for individual laboratory needs, including different model systems and a variety of gasses.

Creating Defined Gaseous Environments to Study the Effects of Hypoxia on C. elegans

This paper details how to use continuous-flow hypoxia chambers to generate atmospheres with defined concentrations of O2 to understand biological responses to decreased O2. This system is easy to setup and maintain, and flexible enough to suit a wide range of O2 concentrations and model systems

Oxygen is essential for all metazoans to survive, with one known exception1. Decreased O2 availability (hypoxia) can arise during states of disease, ...

Measuring Oxidative Stress Resistance of Caenorhabditis elegans in 96-well Microtiter Plates

Oxidative stress, which is the result of an imbalance between production and detoxification of reactive oxygen species, is a major contributor to chronic human disorders, including cardiovascular and neurodegenerative diseases, diabetes, aging, and cancer. Therefore, it is important to study oxidative stress not only in cell systems but also using whole organisms. C. elegans is an attractive model organism to study the genetics of oxidative stress signal transduction pathways, which are highly evolutionarily conserved.
Here, we provide a protocol to measure oxidative stress resistance in C. elegans in liquid. Briefly, ROS-inducing reagents such as paraquat (PQ) and H2O2 are dissolved in M9 buffer, and solutions are aliquoted in the wells of a 96 well microtiter plate. Synchronized L4/young adult C. elegans animals are transferred to the wells (5-8 animals/well) and survival is measured every hour until most worms are dead. When performing an oxidative stress resistance assay using a low concentration of stressors in plates, aging might influence the behavior of animals upon oxidative stress, which could lead to an incorrect interpretation of the data. However, in the assay described herein, this problem is unlikely to occur since only L4/young adult animals are being used. Moreover, this protocol is inexpensive and results are obtained in one day, which renders this technique attractive for genetic screens. Overall, this will help to understand oxidative stress signal transduction pathways, which could be translated into better characterization of oxidative stress-associated human disorders.

Measuring Oxidative Stress Resistance of Caenorhabditis elegans in 96-well Microtiter Plates

C. elegans is an attractive model organism to study signal transduction pathways involved in oxidative stress resistance. Here we provide a protocol to measure oxidative stress resistance of C. elegans animals in liquid phase, using several oxidizing agents in 96 well plates.

Oxidative stress, which is the result of an imbalance between production and detoxification of reactive oxygen species, is a major contributor to chronic ...

Adapting Microrespirometry for Drosophila - Insights into CASK Mutant Metabolism

Measuring O2 Consumption in Drosophila melanogaster Using Coulometric Microrespirometry

Coulometric microrespirometry is a straightforward, inexpensive method for measuring the O2 consumption of small organisms while maintaining a stable environment. A coulometric microrespirometer consists of an airtight chamber in which O2 is consumed and the CO2 produced by the organism is removed by an absorbent medium. The resulting pressure decrease triggers electrolytic O2 production, and the amount of O2 produced is measured by recording the amount of charge used to generate it. In the present study, the method has been adapted to Drosophila melanogaster tested in small groups, with the sensitivity of the apparatus and the environmental conditions optimized for high stability. The amount of O2 consumed by wildtype flies in this apparatus is consistent with that measured by previous studies. Mass-specific O2 consumption by CASK mutants, which are smaller and known to be less active, was not different from congenic controls. However, the small size of CASK mutants resulted in a significant reduction in O2 consumption on a per-fly basis. Therefore, the microrespirometer is capable of measuring O2 consumption in D. melanogaster, can distinguish modest differences between genotypes, and adds a versatile tool for measuring metabolic rates.

Author Spotlight: Influence of Temperature on Drosophila melanogaster and Desert-Adapted Beetles 

Coulometric respirometry is ideal for measuring the metabolic rate of small organisms. When adapted for Drosophila melanogaster in the present study, measured O2 consumption was within the range reported for wildtype D. melanogaster by previous studies. Per-fly O2 consumption by CASK mutants, which are smaller and less active, was significantly lower than the wildtype.

Coulometric microrespirometry is a straightforward, inexpensive method for measuring the O2 consumption of small organisms while maintaining a stable ...

Nematode Synchronization: A Method to Obtain Populations of Worms in Identical Stages of Development

Source: Sarasija, S. and Norman, K. R. <a href="https://www.jove.com/video/59277/measurement-oxygen-consumption-rates-intact-caenorhabditis">Measurement of Oxygen Consumption Rates in Intact&#160;Caenorhabditis elegans</a>.&#160;J. Vis. Exp. (2019).
In this video, we show a method of producing a developmentally synchronized population of sterile adult C. elegans. The example protocol synchronizes worms to study oxygen consumption rates in live animals.

Synchronizacja nicieni: metoda uzyskiwania populacji robaków w identycznych stadiach rozwoju

Source: Sarasija, S. and Norman, K. R. Measurement of Oxygen Consumption Rates in Intact&#160;Caenorhabditis elegans.&#160;J. Vis. Exp. (2019).
In this ...

Measurement of Oxygen Consumption Rates in Intact Caenorhabditis elegans

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Measurement of Mitochondrial Oxygen Consumption in Permeabilized Fibers of Drosophila Using Minimal Amounts of Tissue

Comparative Analysis of Experimental Methods to Quantify Animal Activity in Caenorhabditis elegans Models of Mitochondrial Disease

Quantifying Food Intake in Caenorhabditis elegans by Measuring Bacterial Clearance

An Assay for Measuring the Effects of Ethanol on the Locomotion Speed of Caenorhabditis elegans

Isolation and In vitro Activation of Caenorhabditis elegans Sperm

Solid Plate-based Dietary Restriction in Caenorhabditis elegans

Creating Defined Gaseous Environments to Study the Effects of Hypoxia on C. elegans

Measuring Oxidative Stress Resistance of Caenorhabditis elegans in 96-well Microtiter Plates

Measuring O₂ Consumption in Drosophila melanogaster Using Coulometric Microrespirometry

Synchronizacja nicieni: metoda uzyskiwania populacji robaków w identycznych stadiach rozwoju