The authors would like to thank Heather Currey for her initial contribution to the study design, and Dr. Swarup Mitra for his critical review of the manuscript. We would also like to thank Dr. Michael B. Harris for comments, refinements and assistance in producing the demonstration of this methodology. The strains were provided by the Caenorhabditis Genetics Center, which is funded by the NIH Office of Research Infrastructure Programs (P40 OD010440). The research reported in this publication was supported by the National Institute Of General Medical Sciences of the National Institutes of Health under Award Numbers UL1GM118991, TL4GM118992, or RL5GM118990 and by an Institutional Development Award (IDeA) from the National Institute of General Medical Sciences of the National Institutes of Health under grant number 5P20GM103395-15. The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institutes of Health. UA is an AA/EO employer and educational institution and prohibits illegal discrimination against any individual: www.alaska.edu/titleIXcompliance/nondiscrimination.

1. Caenorhabditis Sieve Construction and Use
<ol>
	<li>Construction protocol
	<ol>
		<li>Acquire 2 lids from 50 mL conical tubes (Figure 1A).</li>
		<li>Remove the center area inside the inner lip of the lids (when viewed from the bottom, Figure 1B) using a Bunsen burner and a hot metal probe or a soldering iron or stepped drill bit. 
		NOTE: Using heat to cut the plastic lid is preferable to a blade because there is less risk of injury.</li>
		<li>Clean and sand the cut edges and top the surface with a curved file or a rotary grinding tool (e.g., Dremel). See Figure 1C.</li>
		<li>Cut the circle of the monofilament mesh to the appropriate diameter (Figure 1D). For this, trace a lid onto a monofilament nylon mesh sheet and cut just inside the line drawn.</li>
		<li>Cut grooves/slashes into the top surface of the lids to enhance the adhesion of the two lids when glue is applied to the plastic (Figure 1E).</li>
		<li>Clean the lids with ethanol and let them dry.</li>
		<li>Apply cyanoacrylate glue to the top surface of both lids, keeping to the outer edge. 
		NOTE: A little glue goes a long way.</li>
		<li>Lay the monofilament mesh, according to Table 1, on one glued lid (Figure 1F). Place the second lid inverted on top of the mesh; both lids must have their tops together. Press firmly together (Figure 1G). Ensure that the mesh is taut. 
		NOTE: As a safety measure, use tweezers to place the mesh on the lids.</li>
		<li>Once the initial layer of glue has dried, apply a ring of cyanoacrylate glue around the outer gap between the lids. Be generous as this adds integrity and prevents any peeling apart or leaking.</li>
		<li>Label the filter with the mesh pore size of the monofilament mesh. 
		NOTE: Here, we use two mesh pore sizes&#8212;20 &#181;m and 50 &#181;m.</li>
	</ol>
	</li>
	<li>Use protocol
	<ol>
		<li>Pre-wet the sieve.
		<ol>
			<li>Pipette a saline solution, such as M9 [5 g of NaCl, 6 g of Na2HPO4, 3 g of KH2PO4, 1 L of ultrapure H2O, and 1 mL of MgSO4 (1 M)]17, through the center of the sieve until a droplet forms, condenses, and drips off the center of the filter bottom (Figure 2A). Optionally, apply a lint-free wipe (e.g., Kimwipe) to the bottom to shape or spread a moisture droplet on the mesh.</li>
			<li>Place the sieve over a 50 mL conical tube. Label the tube as &#39;Waste tube&#39; (Figure 2B).</li>
		</ol>
		</li>
		<li>Wash a population of C. elegans off an agar plate.
		<ol>
			<li>Wash the plate with the M9 buffer and place the worm-containing medium on the topside of the monofilament mesh 1 mL at a time (Figure 2C). Make sure to operate in the center of the mesh. Wash all worms from the plate. 
			NOTE: Typically, 3 mL of M9 for a 60 mm plate is sufficient.</li>
			<li>To minimize the number of worms lost in pipetting, use a glass Pasteur pipette to move the worms off the plate and onto the mesh center (repeat as needed).</li>
			<li>Rinse the filter with the M9 buffer from the top. Again, make sure to operate in the center of the mesh and rinse all worms in that area. Wash as many times as necessary to ensure that all the bacteria and smaller worms have passed through the mesh.</li>
		</ol>
		</li>
		<li>Harvest the size-matched animals from the sieve.
		<ol>
			<li>Attach a new 50 mL conical tube to the top of the sieve, with the adult worms from step 1.2.2.3 facing the inside of the new collection tube (Figure 2D).</li>
			<li>Remove the first tube (waste tube) and quickly flip over the sieve and new tube to keep the droplet from migrating (Figure 2E). 
			NOTE: If sterility is not required, use a lint-free wipe (e.g., Kimwipe) to wick fluid from bottom of the mesh prior to flipping.</li>
			<li>Rinse the mesh with M9 into the new 50 mL tube from the new topside (Figure 2F). Again, operate in the mesh center and maintain a droplet on the underside of the mesh.</li>
			<li>Allow the worms to settle or gently spin them down (e.g., &#60;16 x g) for approximately 1 min (Figure 2G).</li>
			<li>Aspirate the buffer solution from the pellet of worms, ideally down to &#62;0.5 mL, or remove as much fluid as possible without disturbing the pellet.</li>
			<li>Pipette the worms using a glass Pasteur pipette onto an NGM plate and let them dry. Space multiple droplets out on a new plate so they dry faster (Figure 2H).</li>
		</ol>
		</li>
		<li>Clean the sieve.
		<ol>
			<li>Rinse the sieve gently and thoroughly with ethanol and reverse-osmosis water. Let it dry.</li>
			<li>Store it in a clean container for indefinite future use.</li>
			<li>Discard it when the mesh develops a &#34;sag&#34; appearance.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>
2. Validation of Caenorhabditis Sieve Sorting Method
<ol>
	<li>General maintenance
	<ol>
		<li>For all experiments, culture worms on a standard Nematode Growth Media17 (1 L of NGM consists of 2.5 g of peptone, 17 g of agar, 3 g of NaCl, 975 mL of double-distilled water, 1 mL of 5 mg/mL cholesterol, 1 mL of 1 M CaCl2, 1 mL of 1 M MgSO4, 25 mL of 1 M KHPO4, and 0.5 mL of 100 mg/mL streptomycin) at 25 &#176;C.</li>
	</ol>
	</li>
	<li>Experimental treatment administration
	<ol>
		<li>Compare the three treatment groups: pick, fluorodeoxyuridine (FUDR), and Caenorhabditis Sieve.
		<ol>
			<li>For the FUDR treatment group, add 100 mg/mL FUDR to the NGM media to a final concentration of 100 &#956;M to prevent any progeny production, and transfer the worms every other day to a fresh NGM plate to avoid food depletion.</li>
			<li>For the pick group, select and transfer the worms manually using a platinum loop.</li>
			<li>For the Caenorhabditis Sieve treatment group, follow step 1.2 and pipette the worms onto an NGM plate.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Caenorhabditis Sieve percentage yield
	<ol>
		<li>To quantify how effective the sieve is at sorting C. elegans, grow age-synchronous N2 animals to day 1 of adulthood at 25 &#176;C (i.e., 48 h after egg laying) and then transfer them by picking them to fresh NGM plates (totaling N = 50 or N = 100 animals per treatment group).</li>
		<li>After 24 h of recovery, transfer the population to new NGM plates with the Caenorhabditis Sieve following the above protocol (see step 1.2) and count the number of successfully transferred animals.</li>
		<li>Calculate the percentage yield as the ratio of the number of animals transferred in comparison to the starting number at the beginning of the transfer multiplied by 100 (%).</li>
	</ol>
	</li>
	<li>Healthspan assays 
	NOTE: Score healthspan parameters of the motility class, the pharyngeal pump rate, and both the anterior and the posterior gentle touch response on days 2, 4, 6, and 8 of adulthood for age-synchronized N2 animals maintained at 25 &#176;C.
	<ol>
		<li>Motility tracking
		<ol>
			<li>Assign motility scores based on a class-based system (classes A, B, and C) following the methods of Herndon et al.18. Compare the effects of the three experimental groups using an ordinal logistic statistics model in statistical analysis software. 
			NOTE: Class A individuals move spontaneously in a normal, sinusoidal pattern. Class B individuals move in markedly non-sinusoidal movements and may require prodding to encourage movement. Class C individuals move their head and/or tail in response to prodding but are unable to move across the agar.</li>
		</ol>
		</li>
		<li>Pharyngeal pump rate
		<ol>
			<li>Count the grinder movement of the animal&#8217;s terminal pharyngeal bulb visually under a stereomicroscope at a 600X final magnification for 1 min.</li>
			<li>Conduct a statistical analysis with a one-way ANOVA with &#945; = 0.05 and Bonferroni post-tests with &#945; = 0.05.</li>
		</ol>
		</li>
		<li>Touch response
		<ol>
			<li>Record a gentle touch response and compare between the three treatment groups. Perform the assays based on the methods described by Calixto et al.19.</li>
			<li>Record the anterior and posterior touch response by gently stroking an eyelash pick perpendicularly across the tail or the head (5x each, alternating head and tail) of the animals.</li>
			<li>Score any movement in the opposite direction of the stroke as 1 point on a scale of 0 to 5 for both the anterior and the posterior response.</li>
			<li>Conduct statistical analysis with a one-way ANOVA with &#945; = 0.05 and Bonferroni post-tests with &#945; = 0.05.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Fecundity assay
	<ol>
		<li>To determine the use of the Caenorhabditis Sieve&#39;s impact on reproduction, grow age-synchronous N2 animals to day 2 of adulthood at 25 &#176;C.</li>
		<li>60 h after egg lay, transfer animals to new NGM plates with either a platinum pick or the Caenorhabditis Sieve and give them 4 h to recover (dry the sieved plates for 20&#8211;30 min).</li>
		<li>After recovery, individually plate the animals via an eyelash pick to NGM plates, give them 24 h to lay eggs, and remove the animals. Allow the progeny on each plate to develop under normal conditions at 25 &#176;C for another 24 h. 
		NOTE: An eyelash pick is a human eyelash secured to the end of a Pasteur pipette with nail polish and sterilized with ethanol.</li>
		<li>Count the number of viable F1 generation individuals. Perform a statistical analysis using a T-test with &#945; = 0.05. 
		Note: Viable offspring are considered to be eggs that successfully hatch and begin their development through the regular larval cycles.</li>
	</ol>
	</li>
	<li>Fluorescent reporter stress response assays
	<ol>
		<li>Perform three commonly used fluorescent reporter assays to detect potential markers of stress: a DAF-16::GFP translocation into the nuclei of cells in a strain [TJ356- zIs356 (pDAF-16::DAF-16-GFP;rol-6)]20; an hsp-16.2 expression [TJ375- gpIs1 (hsp-16.2p::GFP)]21; and a sod-3 expression [CF1553-muIs84([pAD76]sod-3p::GFP+rol-6[su1006])]22.</li>
		<li>For each assay, culture age-synchronous animals from the three treatment groups at 20 &#176;C and examine them at day 3 of adulthood: use a negative control (on a daily basis, transfer a group of animals manually with a platinum pick), a positive control (on a daily basis, transfer a group of animals manually with a platinum pick plus an established stressor), and the Caenorhabditis Sieve (pass the animals through the sieve and allow them to recover for 30 min on NGM just before imaging).</li>
		<li>In the DAF-16::GFP assay, heat shock the positive control animals at 37 &#176;C for 30 min before imaging20. For the hsp-16.2 assay, heat shock the positive control animals for 90 min, 20 h before imaging21. For the sod-3 assay, treat the positive control animals with 100 mM paraquat for 4 h before imaging22.</li>
		<li>Harvest the worms immediately with an eyelash pick and mount them to a coverslip with 1 &#956;L of a 36% poloxamer 407 surfactant solution to immobilize the worms.</li>
		<li>Sandwich the mounted worms with another coverslip. Mount the two coverslips to a standard glass microscopy slide and image the worms using an 8X magnification on an inverted fluorescent microscope (for an overall magnification of 80X) and a constant exposure with a FITC filter.</li>
		<li>To detect any differences between the three groups in the DAF-16::GFP assay, categorize the animals based on the location of the DAF-16::GFP reporter (nuclear if the reporter translocated to nuclei, cytosolic if the reporter located in cytosol, and intermediate if the reporter located in both nuclei and cytosol).</li>
		<li>Compare the results using an ordinal logistics statistical model in statistical analysis software. For the hsp-16.2 and the sod-3 expression assays, use a one-way ANOVA with &#945; = 0.05 and Tukey&#39;s post hoc tests with &#945; = 0.05 to compare the total fluorescence of the head region.</li>
	</ol>
	</li>
</ol>

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The authors have nothing to disclose. The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Herein, we introduced the design and use of the accessible, effective Caenorhabditis Sieve as a tool for sorting and maintaining C. elegans. This tool has several advantages to manually picking individual animals, washing populations, chemical treatments (e.g., FUDR), and more expensive methods of segregating animals. First, the Caenorhabditis Sieve efficiently and quickly (less than 20 min) sorts progeny from large mixed populations of animals (Table 2). Also, the use...

The nematode worm, Caenorhabditis elegans, is a premier model organism. In addition to the straightforward and controlled nature of their cultivation in the laboratory, their entire genome is sequenced1 and the developmental fate of each cell is known2. Due to these features, C. elegans is a widely used model organism for genetic studies. However, along with these beneficial characteristics come some challenges for researchers. Due to their rapid generation time, C. elegans populations can quickly run out of food and/or become mixed populations with multiple generations and developmental stages present at once. Thus, experiments performed on solid nematode growth media (NGM) require researchers to physically move animals to fresh plates before the bacterial food source depletes and new larvae develop. This can be tedious as a frequent transferring of the animals is required to prevent the experimental populations from becoming mixed with offspring generations. Still, some experiments require both large numbers of animals and extended time points (e.g., DNA or RNA extraction in adulthood). This compounds the challenges of accurately maintaining a synchronized population and transferring large numbers of animals.
Current methods of transferring C. elegans cultured on NGM are picking or washing the animals from plate to plate; chemically treating the animals (e.g., with the DNA replication inhibitor fluorodeoxyuridine or FUDR); or using flow cytometry to sort the animals in multi-well plates. Picking involves the use of a hand tool, made with either a thin platinum wire or an eyelash, to manually transfer individual or multiple animals3,4. This method is accurate but requires both skill and time and is a limitation for studies involving large numbers of animals. Picking may also be physically damaging and stressful to the animals by potentially subjecting individuals to unnatural and inconsistent amounts of disturbance and force. Washing involves rinsing a culture dish with a buffer solution and transferring the solution with the animals via glass Pasteur pipette to a new culture plate. This method is rapid and efficient but is not accurate as multiple generations and developmental stages of animals are transferred in bulk. Chemical treatments, such as FUDR, can be dissolved in the culturing media to prevent the production of offspring through blocking any DNA replication, and thus, the gamete production and egg development. While effective, this method must be applied after developmental maturation as to not disrupt the normal developmental processes, and this means that there is still a requirement to transfer the animals prior to its administration3. This method also influences multiple cellular signaling pathways, resulting in noticeable effects on the animals as they age (e.g., a lifespan extension or an altered proteostasis) depending on the strain of C. elegans used5,6,7,8,9,10. Flow cytometry methods automatically sort and transfer individual C. elegans from one multi-well plate to another11. While this method is very effective and efficient, flow cytometry equipment is prohibitively expensive and inaccessible to many researchers. An alternative to transferring animals is to use mutant models that are temperature sensitive, such as fer-15 and fem-1, which become sterile with temperature adjustment12. While using mutant animals is useful in some situations, these specific strains grow slower than wild-type animals and they rely on an altered genome, serving as poor representatives for aging or healthy worms. In addition, the reliance on a temperature shift to induce sterility also results in the absence of a static environment, and temperature changes have been readily shown to influence gene expressions13,14,15. Research groups have previously published techniques describing the use of a mesh to filter C. elegans by size16. However, we were unable to find previous work testing for any changes in the overall health outcomes that may be associated with the use of such filters.
There is, thus, a need within the C. elegans research community for an affordable, efficient, rapid, and accurate method for transferring large numbers of animals between culture plates. We have developed an improved, accessible piece of equipment (named the Caenorhabditis Sieve) and an associated protocol for its manufacture and operation that meets the needs of the C. elegans research community. Herein, we share the design of the Caenorhabditis Sieve and methods for its use, and we demonstrate that its use does not impact the common health or any stress markers when compared to standard manual picking and a treatment with the commonly-used, fertility-restricting chemical FUDR.

<table><tbody><tr><td>Safety glasses</td><td>Uline</td><td>S-21076</td><td></td></tr><tr><td>Protective heat resistant glove</td><td>Grainger</td><td>Item # 3AT17 Mfr. Model # 3AT17 Catalog Page # 1703</td><td></td></tr><tr><td>50 mL conical tube</td><td>Falcon</td><td>14-432-22</td><td></td></tr><tr><td>Synthetic Nylon mesh</td><td>Dynamic&lt;br/&gt; Aqua-Supply Ltd</td><td>NTX20 and NTX50</td><td></td></tr><tr><td>Cyanoacrylate glue</td><td>Scotch Super Glue Liquid</td><td>SAD114</td><td></td></tr><tr><td>Pliers</td><td>Vampliers</td><td>VMPVT-001-8</td><td></td></tr><tr><td>Dremmel tool with circular file</td><td>Lowe&#039;s</td><td>Item # 525945 Model # 100-LG</td><td></td></tr><tr><td>FUDR</td><td>Sigma</td><td>F0503</td><td></td></tr><tr><td>M9 chemicals ( NaCl, Na2HPO4, KH2PO4, MgSO4)&amp;nbsp;</td><td>Sigma&amp;nbsp;</td><td>S7653, RES20908-A7, 1551139, M7506</td><td></td></tr><tr><td>NGM plate chemicals (Bactopeptone, Agar, KH2PO4, K2HPO4, CaCl2,Cholesterol, Streptomycin)</td><td>BD Biosciences (bactopeptone) , Lab express (agar), Sigma ( rest)</td><td>BD bioscience 211677, Lab Express 1001,&amp;nbsp; Sigma 1551139, 1551128, C1016, C8667, S6501</td><td></td></tr><tr><td>Pluronic F-127</td><td>Sigma</td><td>P2443</td><td></td></tr><tr><td>Paraquat dichloride hydrate</td><td>Sigma</td><td>36541</td><td></td></tr><tr><td>Inverted fluorescence microscope</td><td>Olympus</td><td>FSX100</td><td></td></tr></tbody></table>

caenorhabditis sieve: a low-tech instrument and methodology for sorting small multicellular organisms

Caenorhabditis elegans (C. elegans) is a well-established model organism used across a range of basic and biomedical research. Within the nematode research community, there is a need for an affordable and effective way to maintain large, age-matched populations of C. elegans. Here, we present a methodology for mechanically sorting and cleaning C. elegans. Our aim is to provide a cost-effective, efficient, fast, and simple process to obtain animals of uniform sizes and life stages for their use in experiments. This tool, the Caenorhabditis Sieve, uses a custom-built lid system that threads onto common conical lab tubes and sorts C. elegans based on body size. We also demonstrate that the Caenorhabditis Sieve effectively transfers animals from one culture plate to another allowing for a rapid sorting, synchronizing, and cleaning without impacting markers of health, including motility and stress-inducible gene reporters. This accessible and innovative tool is a fast, efficient, and non-stressful option for maintaining C. elegans populations.

The Caenorhabditis Sieve consists of 2 screw caps, securing an area of woven nylon monofilament mesh smaller than the body diameter of the desired developmental age, used to extract live populations of organisms using a simple washing technique. It attaches to standard conical tubes and uses the mesh screen to mechanically sort animals by body diameter, leaving the desired animals in the tube ready for further maintenance and experimentation (e.g., transfer or genetic ha...

Watch this Scientific Journal Video about Caenorhabditis Sieve: A Low-tech Instrument and Methodology for Sorting Small Multicellular Organisms at JoVE.com

Caenorhabditis Sieve: A Low-tech Instrument and Methodology for Sorting Small Multicellular Organisms

The current protocol includes a methodology for the sorting and cleaning of age-matched populations of Caenorhabditis elegans. It uses a simple, inexpensive, and efficient custom-made tool to obtain a large experimental population of nematodes for research.

Caenorhabditis Sieve: A Low-tech Instrument and Methodology for Sorting Small Multicellular Organisms

Caenorhabditis elegans (C. elegans) is a well-established model organism used across a range of basic and biomedical research. Within the nematode ...

caenorhabditis-sieve-low-tech-instrument-methodology-for-sorting

Research

JoVE Journal

Developmental Biology

6.2K Views.  University of Alaska Fairbanks. The current protocol includes a methodology for the sorting and cleaning of age-matched populations of Caenorhabditis elegans. It uses a simple, inexpensive, and efficient custom-made tool to obtain a large experimental population of nematodes for research.

Video: Caenorhabditis Sieve: A Low-tech Instrument and Methodology for Sorting Small Multicellular Organisms

Culture and Assay of Large-Scale Mixed-Stage Caenorhabditis elegans Populations

Caenorhabditis elegans (C. elegans) has been and remains a valuable model organism to study developmental biology, aging, neurobiology, and genetics. The large body of work on C. elegans makes it an ideal candidate to integrate into large-population, whole-animal studies to dissect the complex biological components and their relationships with another organism. In order to use C. elegans in collaborative -omics research, a method is needed to generate large populations of animals where a single sample can be split and assayed across diverse platforms for comparative analyses.
Here, a method to culture and collect an abundant mixed-stage C. elegans population on a large-scale culture plate (LSCP) and subsequent phenotypic data is presented. This pipeline yields sufficient numbers of animals to collect phenotypic and population data, along with any data needed for -omics experiments (i.e., genomics, transcriptomics, proteomics, and metabolomics). In addition, the LSCP method requires minimal manipulation to the animals themselves, less user preparation time, provides tight environmental control, and ensures that handling of each sample is consistent throughout the study for overall reproducibility. Lastly, methods to document population size and population distribution of C. elegans life stages in a given LSCP are presented.

Culture and Assay of Large-Scale Mixed-Stage Caenorhabditis elegans Populations

To use Caenorhabditis elegans (C. elegans) in omics research, a method is needed to generate large populations of worms where a single sample can be measured across platforms for comparative analyses. Here, a method to culture C. elegans populations on large-scale culture plates (LSCPs) and to document population growth is presented.

Caenorhabditis elegans (C. elegans) has been and remains a valuable model organism to study developmental biology, aging, neurobiology, and genetics. The ...

Genetics

Automated Separation of C. elegans Variably Colonized by a Bacterial Pathogen

The wormsorter is an instrument analogous to a FACS machine that is used in studies of Caenorhabditis elegans, typically to sort worms based on expression of a fluorescent reporter. Here, we highlight an alternative usage of this instrument, for sorting worms according to their degree of colonization by a GFP-expressing pathogen. This new usage allowed us to address the relationship between colonization of the worm intestine and induction of immune responses. While C. elegans immune responses to different pathogens have been documented, it is still unknown what initiates them. The two main possibilities (which are not mutually exclusive) are recognition of pathogen-associated molecular patterns, and detection of damage caused by infection. To differentiate between the two possibilities, exposure to the pathogen must be dissociated from the damage it causes. The wormsorter enabled separation of worms that were extensively-colonized by the Gram-negative pathogen Pseudomonas aeruginosa, with the damage likely caused by pathogen load, from worms that were similarly exposed, but not, or marginally, colonized. These distinct populations were used to assess the relationship between pathogen load and the induction of transcriptional immune responses. The results suggest that the two are dissociated, supporting the possibility of pathogen recognition.

Automated Separation of C. elegans Variably Colonized by a Bacterial Pathogen

The wormsorter facilitates genetic screens in Caenorhabditis elegans by sorting worms according to expression of fluorescent reporters. Here, we describe a new usage:&#160;sorting according to colonization by a GFP-expressing pathogen, and we employ it to examine the poorly understood role of pathogen recognition in initiating immune responses.

The wormsorter is an instrument analogous to a FACS machine that is used in studies of Caenorhabditis elegans, typically to sort worms based on expression ...

Immunology and Infection

Novel Metrics to Characterize Embryonic Elongation of the Nematode Caenorhabditis elegans

Dissecting the signaling pathways that control the alteration of morphogenic processes during embryonic development requires robust and sensitive metrics. Embryonic elongation of the nematode Caenorhabditis elegans is a late developmental stage consisting of the elongation of the embryo along its longitudinal axis. This developmental stage is controlled by intercellular communication between hypodermal cells and underlying body-wall muscles. These signaling mechanisms control the morphology of hypodermal cells by remodeling the cytoskeleton and the cell-cell junctions. Measurement of embryonic lethality and developmental arrest at larval stages as well as alteration of cytoskeleton and cell-cell adhesion structures in hypodermal and muscle cells are classical phenotypes that have been used for more than 25 years to dissect these signaling pathways. Recent studies required the development of novel metrics specifically targeting either early or late elongation and characterizing morphogenic defects along the antero-posterior axis of the embryo. Here, we provide detailed protocols enabling the accurate measurement of the length and the width of the elongating embryos as well as the length of synchronized larvae. These methods constitute useful tools to identify genes controlling elongation, to assess whether these genes control both early and late phases of this stage and are required evenly along the antero-posterior axis of the embryo.

Novel Metrics to Characterize Embryonic Elongation of the Nematode Caenorhabditis elegans

Here we detail protocols specifically designed to monitor morphogenic defects that occur during early and late phases of embryonic elongation of the nematode Caenorhabditis elegans. Ultimately, these protocols are designed to identify genes that regulate these phases and to characterize their differential requirements along the antero-posterior axis of the embryo.

Dissecting the signaling pathways that control the alteration of morphogenic processes during embryonic development requires robust and sensitive metrics. ...

Neuroscience

Basic Caenorhabditis elegans Methods: Synchronization and Observation

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

Basic Caenorhabditis elegans Methods: Synchronization and Observation

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

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

Biology

Sorting of Streptomyces Cell Pellets Using a Complex Object Parametric Analyzer and Sorter

Streptomycetes are filamentous soil bacteria that are used in industry for the production of enzymes and antibiotics. When grown in bioreactors, these organisms form networks of interconnected hyphae, known as pellets, which are heterogeneous in size. Here we describe a method to analyze and sort mycelial pellets using a Complex Object Parametric Analyzer and Sorter (COPAS). Detailed instructions are given for the use of the instrument and the basic statistical analysis of the data. We furthermore describe how pellets can be sorted according to user-defined settings, which enables downstream processing such as the analysis of the RNA or protein content. Using this methodology the mechanism underlying heterogeneous growth can be tackled. This will be instrumental for improving streptomycetes as a cell factory, considering the fact that productivity correlates with pellet size.

Sorting of Streptomyces Cell Pellets Using a Complex Object Parametric Analyzer and Sorter

Liquid-grown Streptomyces cultures are characterized by mycelial pellets that are heterogeneous in size. We here describe a method to analyze and sort such pellets in a high-throughput manner. These pellets can be used for further analyses, which will provide leads to understand and control growth heterogeneity.

Streptomycetes are filamentous soil bacteria that are used in industry for the production of enzymes and antibiotics. When grown in bioreactors, these ...

Automating Aggregate Quantification in Caenorhabditis elegans

A rise in the prevalence of neurodegenerative protein conformational diseases (PCDs) has fostered a great interest in this subject over the years. This increased attention has called for the diversification and improvement of animal models capable of reproducing disease phenotypes observed in humans with PCDs. Though murine models have proven invaluable, they are expensive and are associated with laborious, low-throughput methods. Use of the Caenorhabditis elegans nematode model to study PCDs has been justified by its relative ease of maintenance, low cost, and rapid generation time, which allow for high-throughput applications. Additionally, high conservation between the C. elegans and human genomes makes this model an invaluable discovery tool. Nematodes that express fluorescently tagged tissue-specific polyglutamine (polyQ) tracts exhibit age- and polyQ length-dependent aggregation characterized by fluorescent foci. Such reporters are often employed as proxies to monitor changes in proteostasis across tissues. Manual aggregate quantification is time-consuming, limiting experimental throughput. Furthermore, manual foci quantification can introduce bias, as aggregate identification can be highly subjective. Herein, a protocol consisting of worm culturing, image acquisition, and data processing was standardized to support high-throughput aggregate quantification using C. elegans that express intestine-specific polyQ. By implementing a C. elegans-based image processing pipeline using CellProfiler, an image analysis software, this method has been optimized to separate and identify individual worms and enumerate their respective aggregates. Though the concept of automation is not entirely unique, the need to standardize such procedures for reproducibility, elimination of bias from manual counting, and increase throughput is high. It is anticipated that these methods can drastically simplify the screening process of large bacterial, genomic, or drug libraries using the C. elegans model.

Automating Aggregate Quantification in Caenorhabditis elegans

The following protocol describes the development and optimization of a high-throughput workflow for worm culturing, fluorescence imaging, and automated image processing to quantify polyglutamine aggregates as an assessment of changes in proteostasis.

A rise in the prevalence of neurodegenerative protein conformational diseases (PCDs) has fostered a great interest in this subject over the years. This ...

A Highly Scalable Approach to Perform Ecological Surveys of Selfing Caenorhabditis Nematodes

Caenorhabditis elegans is one of the major model organisms in biology, but only recently have researchers focused on its natural ecology. The relative sparsity of information about C. elegans in its natural context comes from the challenges involved in the identification of the small nematode in nature. Despite these challenges, an increasing focus on the ecology of C. elegans has caused a wealth of new information regarding its life outside of the laboratory. The intensified search for C. elegans in nature has contributed to the discovery of many new Caenorhabditis species and revealed that congeneric nematodes frequently cohabitate in the wild, where they feed on microbial blooms associated with rotting plant material. The identification of new species has also revealed that the androdioecious mating system of males and self-fertilizing hermaphrodites has evolved three times independently within Caenorhabditis. The other two selfing species, C. briggsae and C. tropicalis, share the experimental advantages of C. elegans and have enabled comparative studies into the mechanistic basis of important traits, including self-fertilization. Despite these advances, much remains to be learned about the ecology and natural diversity of these important species. For example, we still lack functional information for many of their genes, which might only be attained through an understanding of their natural ecology. To facilitate ecological research of selfing Caenorhabditis nematodes, we developed a highly scalable method to collect nematodes from the wild. Our method makes use of mobile data collection platforms, cloud-based databases, and the R software environment to enhance researchers&#39; ability to collect nematodes from the wild, record associated ecological data, and identify wild nematodes using molecular barcodes.

A Highly Scalable Approach to Perform Ecological Surveys of Selfing Caenorhabditis Nematodes

This protocol can be used to perform large-scale ecological surveys of selfing Caenorhabditis nematodes. The primary advantage of this method is the efficient organization and analysis of ecological and molecular data associated with the nematodes collected from nature.

Caenorhabditis elegans is one of the major model organisms in biology, but only recently have researchers focused on its natural ecology. The relative ...

Laser Microdissection for Species-Agnostic Single-Tissue Applications

Single-cell methodologies have revolutionized the analysis of the transcriptomes of specific cell types. However, they often require species-specific genetic &#34;toolkits,&#34; such as promoters driving tissue-specific expression of fluorescent proteins. Further, protocols that disrupt tissues to isolate individual cells remove cells from their native environment (e.g., signaling from neighbors) and may result in stress responses or other differences from native gene expression states. In the present protocol, laser microdissection (LMD) is optimized to isolate individual nematode tail tips for the study of gene expression during male tail tip morphogenesis.
LMD allows the isolation of a portion of the animal without the need for cellular disruption or species-specific toolkits and is thus applicable to any species. Subsequently, single-cell RNA-seq library preparation protocols such as CEL-Seq2 can be applied to LMD-isolated single tissues and analyzed using standard pipelines, given that a well-annotated genome or transcriptome is available for the species. Such data can be used to establish how conserved or different the transcriptomes are that underlie the development of that tissue in different species.
Limitations include the ability to cut out the tissue of interest and the sample size. A power analysis shows that as few as 70 tail tips per condition are required for 80% power. Tight synchronization of development is needed to obtain this number of animals at the same developmental stage. Thus, a method to synchronize animals at 1 h intervals is also described.

A protocol is described that uses laser microdissection to isolate individual nematode tissues for RNA-sequencing. The protocol does not require species-specific genetic toolkits, allowing gene expression profiles to be compared between different species at the level of single-tissue samples.

Single-cell methodologies have revolutionized the analysis of the transcriptomes of specific cell types. However, they often require species-specific ...

Hand Dissection of Caenorhabditis elegans Intestines

Comprised of only 20 cells, the Caenorhabditis elegans intestine&#160;is the nexus of many life-supporting functions, including digestion, metabolism, aging, immunity, and environmental response. Critical interactions between the C. elegans host and its environment converge within the intestine, where gut microbiota concentrate. Therefore, the ability to isolate intestine tissue away from the rest of the worm is necessary to assess intestine-specific processes. This protocol describes a method for hand dissecting adult C. elegans intestines. The procedure can be performed in fluorescently labeled strains for ease or training purposes. Once the technique is perfected, intestines can be collected from unlabeled worms of any genotype. This microdissection approach allows for the simultaneous capture of host intestinal tissue and gut microbiota, a benefit to many microbiome studies. As such, downstream applications for the intestinal preparations generated by this protocol can include but are not limited to RNA isolation from intestinal cells and DNA isolation from captured microbiota. Overall, hand dissection of C. elegans intestines affords a simple and robust method to investigate critical aspects of intestine biology.

Hand Dissection of Caenorhabditis elegans Intestines

The present protocol describes a procedure for isolating intestines from adult Caenorhabditis elegans nematode worms by hand for input in genomics, proteomics, microbiome, or other assays.

Comprised of only 20 cells, the Caenorhabditis elegans intestine&#160;is the nexus of many life-supporting functions, including digestion, metabolism, ...

Single Cell Dissociation of Caenorhabditis elegans: A Method to Isolate Live Cells

Source: Germany, E. M., et al. <a href="https://www-jove-com.bdigitaluss.remotexs.co/video/60145/isolation-specific-neuron-populations-from-roundworm-caenorhabditis">Isolation of Specific Neuron Populations from Roundworm Caenorhabditis elegans</a>. J. Vis. Exp. (2019).
This video describes a method to dissociate whole worms into a single-cell suspension suitable for FACS or immunoprecipitation of intact cells.

Source: Germany, E. M., et al. Isolation of Specific Neuron Populations from Roundworm Caenorhabditis elegans. J. Vis. Exp. (2019).
This video describes a ...

Caenorhabditis Sieve: A Low-tech Instrument and Methodology for Sorting Small Multicellular Organisms

Summary

Explore More Videos

Chapters in this video

Culture and Assay of Large-Scale Mixed-Stage Caenorhabditis elegans Populations

Automated Separation of C. elegans Variably Colonized by a Bacterial Pathogen

Novel Metrics to Characterize Embryonic Elongation of the Nematode Caenorhabditis elegans

Basic Caenorhabditis elegans Methods: Synchronization and Observation

Sorting of Streptomyces Cell Pellets Using a Complex Object Parametric Analyzer and Sorter

Automating Aggregate Quantification in Caenorhabditis elegans

A Highly Scalable Approach to Perform Ecological Surveys of Selfing Caenorhabditis Nematodes

Laser Microdissection for Species-Agnostic Single-Tissue Applications

Hand Dissection of Caenorhabditis elegans Intestines

Single Cell Dissociation of Caenorhabditis elegans: A Method to Isolate Live Cells