Name: a flame-free method for sterilizing c. elegans picks, spatulas, and scalpels
Uploaded: 2022-03-04T15:00:00.000+00:00
Duration: 4 min 30 s
Description: Watch this Scientific Journal Video about A Flame-Free Method for Sterilizing C. elegans Picks, Spatulas, and Scalpels at JoVE.com

The authors would like to acknowledge Suzanne Howard and Justin Finne. This work was funded by the Wellesley College Neuroscience Department. N2 worms were provided by the CGC which is funded by the NIH Office of Research Infrastructure Programs (P40 OD010440).&#160;The publication fees for this article were supported by the Wellesley College Library and Technology Services Open Access Fund.

1. Prepare the micro-incinerator
<ol>
	<li>Attach a loop holder accessory guide to the micro-incinerator by clipping it onto the outer barrel.</li>
	<li>Plug in micro-incinerator to a standard 120 V or 230 V electrical outlet as appropriate. 
	NOTE: This can be done on a benchtop or in a laminar flow hood.</li>
	<li>Turn the micro-incinerator to the high setting and allow it to warm up for 10-20 min depending on the manufacturer&#39;s instructions to reach an optimum temperature of 800-825 oC. 
	&#8203;NOTE: If keeping the incinerator on for longer than 3 h, the lower temperature setting (500 oC) can be used as a standby setting. According to manufacturers&#39; user manuals, this extends the usable life of the equipment.</li>
</ol>
2. Sterilize the pick, spatula, or scalpel
<ol>
	<li>Insert the instrument into the cylindrical sterilization area without touching the sides by sliding along the guide21. 
	NOTE: If sterilizing a scalpel, it is critical to avoid touching the ceramic wall with the blade. Scraping the ceramic wall may compromise the integrity of the heating unit.</li>
	<li>Hold the instrument in the sterilization area for 5-7 s.</li>
	<li>Remove the instrument without touching the sides by sliding backward along the guide.</li>
	<li>For picks, allow the instrument to cool for 3-5 s before touching a worm to avoid burning it. 
	NOTE: Scalpels or spatulas that are not allowed to cool will singe the agar.</li>
	<li>After picking worms, re-insert the pick in the chamber for 5-7 s to incinerate any worms on the pick.</li>
</ol>
3. Comparative method - sterilization of instruments using a Bunsen burner
<ol>
	<li>Connect Bunsen burner to the gas line using rubber tubing. Make sure to secure the tubing tightly and position the burner away from overhead objects.</li>
	<li>Turn on gas by turning the knob on the gas line.</li>
	<li>Ignite the burner using a striker or lighter.</li>
	<li>Adjust flame using the gas knob and the air intake until a blue cone is visible.</li>
	<li>Hold the pick, spatula, or scalpel in the flame until it glows red.</li>
	<li>For picks, allow the instrument to cool for 3-5 seconds before touching a worm to avoid burning it. 
	NOTE: Scalpels or spatulas that are not allowed to cool will singe the agar.</li>
	<li>After picking worms, re-insert the pick in the flame to incinerate any worms on the pick.</li>
</ol>
4. Experiment
<ol>
	<li>Culture OP50 Escherichia coli (E. coli) bacteria in Luria Broth22 overnight on a 37 oC shaker.</li>
	<li>After overnight culture, dilute the culture in sterile water at a ratio of 1:100. 
	NOTE: This dilution factor was chosen to ensure the separation of colonies after plating.</li>
	<li>Dip a worm pick sterilized using a Bunsen burner in the bacterial solution, re-sterilize with the Bunsen burner, and swirl it in 100 mL of sterile water.</li>
	<li>Plate the 100 &#181;L of water on a 10 cm LB agar22,23 Petri dish, spread the water using a sterile cell spreader, and incubate at room temperature for 24 h with the lid on.</li>
	<li>Perform steps 4.3-4.4 using a worm pick sterilized using a micro-incinerator.</li>
	<li>As a positive control, perform steps 4.3-4.4 without re-sterilization.</li>
	<li>As a negative control, perform steps 4.3-4.4 using sterile water in place of the bacterial solution.</li>
	<li>Count the colonies manually after the incubation period.</li>
</ol>

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No conflicts of interest were disclosed.

C. elegans is a model organism well suited for exercises in undergraduate teaching laboratories. The use of micro-incinerators in place of open flames provides benefits in both research laboratories and classroom laboratories. In fact, undergraduate laboratory courses may pose a higher risk of accidental fires given the number of newly trained scientists in the room. In addition, the risk of fire increases when ethanol is used to sterilize instruments near the source of flame as ethanol vapors are ignitable. The portability also confers an advantage for classrooms where gas lines are not installed at each bench. This method has been employed in teaching and research laboratories at our institution, resulting in no increases in contamination and zero laboratory safety accidents since its incorporation in 2016. 
To ensure compatibility with micro-incinerators, a range of picks with different mountings, wire compositions, and wire gauges were tested. Wire composition included 100% platinum as well as 90% platinum/10% iridium with a wire thickness of 30-32 G, and regardless of thickness and composition, the heating method did not compromise wire integrity. Mountings included two different types of commercially available pick handles and in-house-made glass mountings from Pasteur pipettes. Note that picks do not glow red-hot as they do in the flame. However, sufficient sterilization is still achieved as long as the sterilizer has reached the proper temperature. Thus, it is critical to allow the micro-incinerator to warm up for 10 or 20 min, as indicated in the manufacturer's instructions. Keeping the instrument in the chamber for at least 5 s to achieve sterilization is also critical. Leaving the instrument in the chamber longer than 7 s will not cause harm to the instrument but is unnecessary. While this is a relatively straightforward procedure with minimal steps and is unlikely to need troubleshooting, it may take some practice to learn to steady the instrument in the barrel without touching the sides.
In order to replace an open flame, a sterilization method must cover all of the applications used in a laboratory. In addition to sterilizing instruments, C. elegans laboratories may also use Bunsen burners to create a sterile field to do other tasks such as pouring plates or inoculating cultures13. However, whether this creates a sterile field or draws in still-viable contaminants remains controversial14. Although not an option at all institutions, a biosafety cabinet or laminar flow hood can be used for these purposes, allowing a laboratory to function without the use of open flames. Instruction manuals for most micro-incinerators recommend against use for sterilization of scalpel blades because scraping of the inner walls damages the sterilizer. However, if carefully using a guide, one can sterilize a scalpel blade or spatula without touching the inner walls. This extends the use of the technique to allow for chunking without using a flame and allows a C. elegans laboratory to function without switching techniques between sterilizing different objects.
As outlined in Table 1, micro-incinerators offer increased safety, increased compatibility with laminar flow hoods, and increased portability over flame-based methods but have limitations. They are more costly than the other methods and require a warm-up time before sterilization temperatures are achieved. In conclusion, this method which is routinely used in many microbiology laboratories, may be applicable in some C. elegans research and teaching laboratories seeking to improve laboratory safety without compromising sterility.

The model organism C. elegans is well-suited for both research and education at primarily undergraduate institutions (PUIs) due to the low cost, ease of maintenance, and range of applications1,2,3,4. In order to handle worms - for example, to move a worm from one plate to another, experimenters can use a worm pick. A variety of picks can be made or purchased for use with C. elegans. Picks are most commonly made using a platinum or platinum/iridium tip, mounted in a glass, metal, or wooden handle. Glass handles can be made in-house by melting a Pasteur pipette around a platinum wire until the wire is secure. Additional information on C. elegans husbandry, including how to grow and maintain worms and their food sources, can be found in WormBook5 and other sources6,7,8.
When working with C. elegans, aseptic techniques are typically used to prevent contamination with microbes and fungi. Examples of aseptic techniques include sterilization of instruments, autoclaving of reagents, and conducting work in sterile fields. Worm picks are typically sterilized using an open flame9. Additionally, sterilization of the worm pick incinerates worms, thus preventing accidental mix-ups of strains when working with multiple worm strains. Typical methods for sterilizing worm picks involve an open flame from either a Bunsen burner, ethanol lantern, or standard lighter (Table 1). We were motivated to seek safer alternatives to existing methods in the laboratory when an undergraduate student unknowingly spilled ethanol while filling an ethanol lantern and accidentally started a small fire when igniting the lantern. Unfortunately, many accidents have been reported using ethanol lanterns10,11,12. Fortunately, alternative sterilization methods have been validated for use in microbiology, and the purpose of this article is to demonstrate how to use this equipment to sterilize instruments for use with C. elegans.
In microbiology laboratories, the aseptic technique is also critical. Serological loops and wires made of platinum are sterilized either using an open flame13 or a micro-incinerator14,15,16. Other names for micro-incinerators include micro-sterilizers or bacto-incinerators. Advantages of the micro-incinerator over traditional flame methods include reduced fire hazard, elimination of spattering of incinerated materials, and ability to work in a laminar flow hood/biosafety cabinet16,17,18. In fact, both the American Society of Microbiology and the World Health Organization recommends using micro-incinerators over the use of an open flame17,19,20. In comparison with Bunsen burners, micro-incinerators also do not require a gas line, which some laboratories might not have, or might not have located at each bench for students to use. Inspired by these advantages, a protocol was developed to replace the use of flame with micro-incinerators for sterilization of commonly used instruments such as picks, spatulas, and scalpels in the C. elegans laboratory. This method may be appropriate for instructors and researchers seeking to enhance safety and/or flexibility when working with C. elegans.

<table><tbody><tr><td>90% platinum 10% iridium wire</td><td>Tritech</td><td>PT-9010</td><td>Other sources and wire compositions may be used.</td></tr><tr><td>Agarose</td><td>Sigma Aldrich</td><td>A6013</td><td>LB agar ingredient</td></tr><tr><td>Bunsen burner</td><td>Fisher Scientific</td><td>50-110-1225</td><td>Other Bunsen burners may be used</td></tr><tr><td>Ethanol Lamp</td><td>Carolina</td><td>706604</td><td>Included here as a reference to Table 1</td></tr><tr><td>Lighter</td><td>Carolina</td><td>706636</td><td>Included here as a reference to Table 1</td></tr><tr><td>Loop holder accessory</td><td>Fisher</td><td>22-630-002</td><td>Referred to in the manuscript as &quot;guide&quot;</td></tr><tr><td>Micro-incinerator</td><td>Thomas Scientific</td><td>1154J15</td><td>There are many companies that sell similar equipment. Similar models also sold by Benchmark Scientific (B1001),&amp;nbsp; Fisher Scientific (22-630-001), Carolina (703400), and BT Lab Systems (BT1702).</td></tr><tr><td>N2 worms</td><td>CGC</td><td>N2</td><td></td></tr><tr><td>NaCl</td><td>Sigma Aldrich</td><td>S5886</td><td>LB ingredient</td></tr><tr><td>OP50 E. coli</td><td>CGC</td><td>OP50</td><td></td></tr><tr><td>Petri dish</td><td>Fisher</td><td>08-772B</td><td></td></tr><tr><td>Pick handle</td><td>Tritech</td><td>TWPH1</td><td></td></tr><tr><td>Scalpel blade</td><td>Fisher</td><td>12-000-161</td><td></td></tr><tr><td>Scalpel handle</td><td>Fisher</td><td>12-000-164</td><td></td></tr><tr><td>Spatula</td><td>Fisher</td><td>14-374</td><td>Other spatulas will work</td></tr><tr><td>Sterile cell spreaders</td><td>VWR</td><td>76206-438</td><td>Other cell spreaders may be used as long as they are sterile</td></tr><tr><td>Tryptone</td><td>Sigma Aldrich</td><td>T7293</td><td>LB ingredient</td></tr><tr><td>Yeast Extract</td><td>Sigma Aldrich</td><td>Y1625</td><td>LB ingredient</td></tr></tbody></table>

a flame-free method for sterilizing c. elegans picks, spatulas, and scalpels

Caenorhabtidis elegans (C. elegans) is an optimal model organism for research and education at primarily undergraduate institutions. Undergraduates can quickly learn the sterile technique required to maintain C. elegans cultures. Sterilization of platinum picks used to transfer worms from one plate to another is traditionally done by holding the pick in a flame from a Bunsen burner or ethanol lantern. However, Bunsen burners require a gas source, and both pieces of equipment pose the risk of accidental fire associated with an open flame. Demonstrated here is a technique for sterilizing worm picks, spatulas, and scalpels using an infrared bacteriological loop micro-incinerator. This equipment requires only an electrical outlet and minimizes potential fire hazards. By lowering risk and gas requirements, this technique is well suited for research and teaching in an undergraduate setting.

A simple experiment (section 4) was devised to demonstrate the relative contamination rates using a micro-incinerator versus a flame (Figure 1, Table 2). While these results represent contamination rates across methods, it would not be necessary to repeat this method to use the micro-incinerator technique. The experiment was run in triplicate. Negative control was sterile water with no bacteria. The positive control used neither sterilization technique: the pick was dipped in the diluted OP50 culture then transferred directly to the sterile water. All replicates and conditions were run in parallel on the same day. In all three negative control plates, zero colonies were counted. A mean count of 4.7 (&#177; 0.3 SEM) colonies was obtained when the Bunsen burner was used to sterilize picks. A mean count of 3.3 (&#177;1.2 SEM) colonies was observed in the micro-incinerator condition. However, a mean count of 298.3 (&#177;17.9 SEM) colonies was obtained in the positive control condition. A 2-tailed t-test assuming equal variance comparing the Bunsen burner to micro-incinerator yielded no statistically significant difference, p = 0.35. Thus, the micro-incinerator achieved sterility results equally effective to the Bunsen burner.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/63578/63578fig01.jpg" /> 
Figure 1: Bacterial counts across sterilization methods.&#160;Picks were dipped in 1:100 OP50 culture in sterile water and then sterilized using a Bunsen burner or micro-incinerator. Picks were then dipped in sterile water, plated on LB agar22,23, and incubated for 24 h at room temperature, and colonies were counted manually. Positive control had no sterilization, and negative control used sterile water with no bacteria. n = 3 replicates per condition. Note: y-axis is broken to allow separation of data points in relevant portions of the graph. <a href="https://www.jove.com/files/ftp_upload/63578/63578fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Sterilization Method</td>
			<td>Cost</td>
			<td>Lab Requirements</td>
			<td colspan="2">Advantages</td>
			<td colspan="3">Disadvantages</td>
		</tr>
		<tr>
			<td>Micro-incinerator</td>
			<td>$365&#8211;530</td>
			<td>120 V or 230 V outlet</td>
			<td colspan="2">&#8226; Portable 
			&#8226; No open flame or exposed heating element 
			&#8226; Useable in laminar flow hoods and biological safety cabinets</td>
			<td colspan="3">&#8226; Warm up time 
			&#8226; Higher cost</td>
		</tr>
		<tr>
			<td>Bunsen Burner</td>
			<td>$24&#8211;169</td>
			<td>Gas Line</td>
			<td colspan="2">&#8226; Fast set up 
			&#8226; Low cost</td>
			<td colspan="3">&#8226; Open Flame 
			&#8226; Not recommended for use in laminar flow hood or biological safety cabinet</td>
		</tr>
		<tr>
			<td>Ethanol Lamp</td>
			<td>$11</td>
			<td>None</td>
			<td colspan="2">&#8226; Fast set up 
			&#8226; Low cost 
			&#8226; Refillable</td>
			<td colspan="3">&#8226; Open Flame 
			&#8226; Safety hazard when refilling 
			&#8226; Not recommended for use in laminar flow hood or biological safety cabinet 
			&#8226; Not allowed at certain institutions</td>
		</tr>
		<tr>
			<td>Lighter</td>
			<td>$5&#8211;8</td>
			<td>None</td>
			<td colspan="2">&#8226; Low cost 
			&#8226; Accessible 
			&#8226; Disposable 
			&#8226; Refillable</td>
			<td colspan="3">&#8226; Open Flame 
			&#8226; Hand operated 
			&#8226; Not recommended for use in laminar flow hood or biological safety cabinet</td>
		</tr>
	</tbody>
</table>
Table 1: Comparison of methods for sterilizing instruments. Four methods for sterilizing instruments were compared based on advantages, disadvantages, costs, and laboratory requirements.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td rowspan="2">Replicate</td>
			<td colspan="4">Condition</td>
		</tr>
		<tr>
			<td>Micro-incinerator</td>
			<td>Bunsen Burner</td>
			<td>Negative Control</td>
			<td>Positve Control</td>
		</tr>
		<tr>
			<td>1</td>
			<td>1</td>
			<td>5</td>
			<td>0</td>
			<td>278</td>
		</tr>
		<tr>
			<td>2</td>
			<td>4</td>
			<td>4</td>
			<td>0</td>
			<td>334</td>
		</tr>
		<tr>
			<td>3</td>
			<td>5</td>
			<td>5</td>
			<td>0</td>
			<td>283</td>
		</tr>
		<tr>
			<td>Mean</td>
			<td>3.3</td>
			<td>4.7</td>
			<td>0.0</td>
			<td>298.3</td>
		</tr>
		<tr>
			<td>SEM</td>
			<td>1.2</td>
			<td>0.3</td>
			<td>0.0</td>
			<td>17.9</td>
		</tr>
	</tbody>
</table>
Table 2: Raw data of bacterial counts across sterilization methods.&#160;Picks were dipped in 1:100 OP50 culture in sterile water and then sterilized using a Bunsen burner or micro-incinerator. Picks were then dipped in sterile water, plated on LB agar22,23, and incubated for 24 h at room temperature, and colonies were counted manually. Positive control had no sterilization, and negative control used sterile water with no bacteria. n = 3 replicates per condition. Mean and SEM reported below individual counts.

Watch this Scientific Journal Video about A Flame-Free Method for Sterilizing C. elegans Picks, Spatulas, and Scalpels at JoVE.com

A Flame-Free Method for Sterilizing C. elegans Picks, Spatulas, and Scalpels

This article describes a method for sterilizing worm picks, spatulas, and scalpels using a micro-incinerator in place of an open flame.

A Flame-Free Method for Sterilizing C. elegans Picks, Spatulas, and Scalpels

Caenorhabtidis elegans (C. elegans) is an optimal model organism for research and education at primarily undergraduate institutions. Undergraduates can ...

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Biology

4.6K Views.  Wellesley College. This article describes a method for sterilizing worm picks, spatulas, and scalpels using a micro-incinerator in place of an open flame.

Video: A Flame-Free Method for Sterilizing C. elegans Picks, Spatulas, and Scalpels

The Nematode Caenorhabditis Elegans - A Versatile In Vivo Model to Study Host-microbe Interactions

We demonstrate a method using Caenorhabditis elegans as a model host to study microbial interaction. Microbes are introduced via the diet making the intestine the primary location for disease. The nematode intestine structurally and functionally mimics mammalian intestines and is transparent making it amenable to microscopic study of colonization. Here we show that pathogens can cause disease and death. We are able to identify microbial mutants that show altered virulence. Its conserved innate response to biotic stresses makes C. elegans an excellent system to probe facets of host innate immune interactions. We show that hosts with mutations in the dual oxidase gene cannot produce reactive oxygen species and are unable to resist microbial insult. We further demonstrate the versatility of the presented survival assay by showing that it can be used to study the effects of inhibitors of microbial growth. This assay may also be used to discover fungal virulence factors as targets for the development of novel antifungal agents, as well as provide an opportunity to further uncover host-microbe interactions. The design of this assay lends itself well to high throughput whole-genome screens, while the ability to cryo-preserve worms for future use makes it a cost-effective and attractive whole animal model to study.

The Nematode Caenorhabditis Elegans - A Versatile In Vivo Model to Study Host-microbe Interactions

Here, we present the nematode Caenorhabditis elegans as a versatile host model to study microbial interaction.

We demonstrate a method using Caenorhabditis elegans as a model host to study microbial interaction. Microbes are introduced via the diet making the ...

Genetics

In Vitro Reconstitution of Spatial Cell Contact Patterns with Isolated Caenorhabditis elegans Embryo Blastomeres and Adhesive Polystyrene Beads

In multicellular systems, individual cells are surrounded by the various physical and chemical cues coming from neighboring cells and the environment. This tissue complexity confounds the identification of causal link between extrinsic cues and cellular dynamics. A synthetically reconstituted multicellular system overcomes this problem by enabling researchers to test for a specific cue while eliminating others. Here, we present a method to reconstitute cell contact patterns with isolated Caenorhabditis elegans blastomere and adhesive polystyrene beads. The procedures involve eggshell removal, blastomere isolation by disrupting cell-cell adhesion, preparation of adhesive polystyrene beads, and reconstitution of cell-cell or cell-bead contact. Finally, we present the application of this method to investigate the orientation of cellular division axes that contributes to the regulation of spatial cellular patterning and cell fate specification in developing embryos. This robust, reproducible, and versatile in vitro method enables the study of direct relationships between spatial cell contact patterns and cellular responses.

In Vitro Reconstitution of Spatial Cell Contact Patterns with Isolated Caenorhabditis elegans Embryo Blastomeres and Adhesive Polystyrene Beads

Tissue complexities of multicellular systems confound the identification of causal relationship between extracellular cues and individual cellular behaviors. Here, we present a method to study the direct link between contact-dependent cues and division axes using C. elegans embryo blastomeres and adhesive polystyrene beads.

In multicellular systems, individual cells are surrounded by the various physical and chemical cues coming from neighboring cells and the environment. ...

Developmental Biology

Tracking Calcium in the Early Developmental Phases of C. elegans

In Vivo Visualization of Calcium Transients during Fertilization and Early Development in C. elegans

Calcium is an important signaling molecule during the oocyte-to-embryo transition (OET) and early embryogenesis. The hermaphroditic nematode Caenorhabditis elegans provides several unique advantages for the study of the OET as it is transparent and has an ordered gonad that produces one mature oocyte every ~23 min at 20 &#176;C. We have modified the genetically encoded calcium indicator jGCaMP7s to fluorescently indicate the moment of fertilization within a living organism. We have termed this reporter &#34;CaFE&#34; for Calcium during Fertilization in C. elegans. The CaFE reporter was&#160;engineered into a safe harbor locus in single copy, has no significant impact on physiology or fecundity, and produces a robust signal upon fertilization. Here, a series of protocols is presented for utilizing the CaFE reporter as an in vivo tool for dissecting the OET and embryogenesis. We include methods to synchronize worms, examine the effects of RNAi knockdown, mount worms for imaging, and to visualize calcium in oocytes and embryos. Additionally, we present the generation of additional worm strains to aid in this type of analysis. Demonstrating the utility of the CaFE reporter to visualize the timing of fertilization, we report that double ovulation occurs when ipp-5 is targeted by RNAi and that only the first oocyte undergoes immediate fertilization. Furthermore, the discovery of single-cell calcium transients during early embryogenesis is reported here, demonstrating that the CaFE reporter persists into early development. Importantly, the CaFE reporter in worms is simple enough to use for incorporation into course-based undergraduate research (CURE) laboratory classes. The CaFE reporter, coupled with the ordered gonad and ease of RNAi in worms, facilitates inquiry into the cell-cell dynamics required to regulate internal fertilization and early embryogenesis.

Here, we present protocols and tools for visualizing calcium during fertilization and early embryogenesis using a genetically encoded calcium reporter expressed in the germline of the model nematode C. elegans.

Calcium is an important signaling molecule during the oocyte-to-embryo transition (OET) and early embryogenesis. The hermaphroditic nematode ...

A Method for Culturing Embryonic C. elegans Cells

C. elegans is a powerful model system, in which genetic and molecular techniques are easily applicable. Until recently though, techniques that require direct access to cells and isolation of specific cell types, could not be applied in C. elegans. This limitation was due to the fact that tissues are confined within a pressurized cuticle which is not easily digested by treatment with enzymes and/or detergents. Based on early pioneer work by Laird Bloom, Christensen and colleagues 1 developed a robust method for culturing C. elegans embryonic cells in large scale. Eggs are isolated from gravid adults by treatment with bleach/NaOH and subsequently treated with chitinase to remove the eggshells. Embryonic cells are then dissociated by manual pipetting and plated onto substrate-covered glass in serum-enriched media. Within 24 hr of isolation cells begin to differentiate by changing morphology and by expressing cell specific markers. C. elegans cells cultured using this method survive for up 2 weeks in vitro and have been used for electrophysiological, immunochemical, and imaging analyses as well as they have been sorted and used for microarray profiling.

A Method for Culturing Embryonic C. elegans Cells

We describe here a revised protocol for large-scale culture of embryonic C. elegans cells. Embryonic C. elegans cells cultured in vitro using this method, appear to differentiate and recapitulate the expression of genes in a cell specific manner. Techniques that require direct access to the cells or isolation of specific cell types from the other tissues can be applied on C. elegans cultured cells.

C. elegans is a powerful model system, in which genetic and molecular techniques are easily applicable. Until recently though, techniques that require ...

Ablation of a Single Cell From Eight-cell Embryos of the Amphipod Crustacean Parhyale hawaiensis

The amphipod Parhyale hawaiensis is a small crustacean found in intertidal marine habitats worldwide. Over the past decade, Parhyale has emerged as a promising model organism for laboratory studies of development, providing a useful outgroup comparison to the well studied arthropod model organism Drosophila melanogaster. In contrast to the syncytial cleavages of Drosophila, the early cleavages of Parhyale are holoblastic. Fate mapping using tracer dyes injected into early blastomeres have shown that all three germ layers and the germ line are established by the eight-cell stage. At this stage, three blastomeres are fated to give rise to the ectoderm, three are fated to give rise to the mesoderm, and the remaining two blastomeres are the precursors of the endoderm and germ line respectively. However, blastomere ablation experiments have shown that Parhyale embryos also possess significant regulatory capabilities, such that the fates of blastomeres ablated at the eight-cell stage can be taken over by the descendants of some of the remaining blastomeres. Blastomere ablation has previously been described by one of two methods: injection and subsequent activation of phototoxic dyes or manual ablation. However, photoablation kills blastomeres but does not remove the dead cell body from the embryo. Complete physical removal of specific blastomeres may therefore be a preferred method of ablation for some applications. Here we present a protocol for manual removal of single blastomeres from the eight-cell stage of Parhyale embryos, illustrating the instruments and manual procedures necessary for complete removal of the cell body while keeping the remaining blastomeres alive and intact. This protocol can be applied to any Parhyale cell at the eight-cell stage, or to blastomeres of other early cleavage stages. In addition, in principle this protocol could be applicable to early cleavage stage embryos of other holoblastically cleaving marine invertebrates.

Ablation of a Single Cell From Eight-cell Embryos of the Amphipod Crustacean Parhyale hawaiensis

The amphipod Parhyale hawaiensis is a promising model organism for studies of crustacean embryology and comparative arthropod development and evolution. This protocol describes a method for manual removal of single blastomeres from early cleavage stage embryos of Parhyale.

The amphipod Parhyale hawaiensis is a small crustacean found in intertidal marine habitats worldwide. Over the past decade, Parhyale has emerged as a ...

Using Single-Worm Data to Quantify Heterogeneity in Caenorhabditis elegans-Bacterial Interactions

The nematode Caenorhabditis elegans is a model system for host-microbe and host-microbiome interactions. Many studies to date use batch digests rather than individual worm samples to quantify bacterial load in this organism. Here it is argued that the large inter-individual variability seen in bacterial colonization of the C. elegans intestine is informative, and that batch digest methods discard information that is important for accurate comparison across conditions. As describing the variation inherent to these samples requires large numbers of individuals, a convenient 96-well plate protocol for disruption and colony plating of individual worms is established.

Using Single-Worm Data to Quantify Heterogeneity in Caenorhabditis elegans-Bacterial Interactions

This protocol describes a 96-well disruption of individual bacterially colonized Caenorhabditis elegans following cold paralysis and surface bleaching&#160;to remove external bacteria. The resulting suspension is plated on agar plates to allow accurate, medium-throughput quantification of bacterial load in large numbers of individual worms.

The nematode Caenorhabditis elegans is a model system for host-microbe and host-microbiome interactions. Many studies to date use batch digests rather ...

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, ...

Long-Term Culture and Monitoring of Isolated Caenorhabditis elegans on Solid Media in Multi-Well Devices

The nematode Caenorhabditis elegans is among the most common model systems used in aging research owing to its simple and inexpensive culture techniques, rapid reproduction cycle (~3 days), short lifespan (~3 weeks), and numerous available tools for genetic manipulation and molecular analysis. The most common approach for conducting aging studies in C. elegans, including survival analysis, involves culturing populations of tens to hundreds of animals together on solid nematode growth media (NGM) in Petri plates. While this approach gathers data on a population of animals, most protocols do not track individual animals over time. Presented here is an optimized protocol for the long-term culturing of individual animals on microfabricated polydimethylsiloxane (PDMS) devices called WorMotels. Each device allows up to 240 animals to be cultured in small wells containing NGM, with each well isolated by a copper sulfate-containing moat that prevents the animals from fleeing. Building on the original WorMotel description, this paper provides a detailed protocol for molding, preparing, and populating each device, with descriptions of common technical complications and advice for troubleshooting. Within this protocol are techniques for the consistent loading of small-volume NGM, the consistent drying of both the NGM and bacterial food, options for delivering pharmacological interventions, instructions for and practical limitations to reusing PDMS devices, and tips for minimizing desiccation, even in low-humidity environments. This technique allows the longitudinal monitoring of various physiological parameters, including stimulated activity, unstimulated activity, body size, movement geometry, healthspan, and survival, in an environment similar to the standard technique for group culture on solid media in Petri plates. This method is compatible with high-throughput data collection when used in conjunction with automated microscopy and analysis software. Finally, the limitations of this technique are discussed, as well as a comparison of this approach to a recently developed method that uses microtrays to culture isolated nematodes on solid media.

Long-Term Culture and Monitoring of Isolated Caenorhabditis elegans on Solid Media in Multi-Well Devices

Presented here is an optimized protocol for culturing isolated individual nematodes on solid media in microfabricated multi-well devices. This approach allows individual animals to be monitored throughout their lives for a variety of phenotypes related to aging and health, including activity, body size and shape, movement geometry, and survival.

The nematode Caenorhabditis elegans is among the most common model systems used in aging research owing to its simple and inexpensive culture techniques, ...

Aseptic Technique in Environmental Science

Source: Laboratories of Dr. Ian Pepper and Dr. Charles Gerba - The University of Arizona 
Demonstrating Author: Luisa Ikner
Aseptic technique is a fundamental skill widely practiced in the field of environmental microbiology that requires a balance of mindfulness and practice in the laboratory. Proper use of this technique reduces the likelihood of bacterial or fungal contamination of reagents, culture media, and environmental samples. Aseptic technique is also vital to ensure data integrity and maintain the purity of culture libraries that may be comprised of very rare and difficult to culture isolates. Sources of contamination in the laboratory environment include airborne microorganisms (including those adhering to dust and lint particles), microbes present on the laboratory bench workspace or on unsterilized glassware or equipment, and microbes transferred from the body and hair of the researcher. The use of aseptic technique is also a safety measure that lowers the potential for the transmission of microorganisms to researchers, which is particularly important when working with pathogens.

Source: Laboratories of Dr. Ian Pepper and Dr. Charles Gerba - The University of Arizona
Demonstrating Author: Luisa Ikner
Aseptic technique is a ...

Education

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Environmental Sciences

Environmental Microbiology

C. elegans Maintenance

Ceanorhabditis elegans has been, and is still, used to great success as a model organism for studying a variety of developmental, genetic, molecular and even physical phenomena. In order to use C. elegans to its full potential, proper care and attention to the basic maintenance of this powerful organism is essential.
In this video you will learn the basic housing and feeding requirements of C. elegans, how to correctly handle and manipulate worms using a worm pick and how to freeze and recover important worm stocks. Towards the end of the video we will visit a few applications of modifying the housing, feeding and manipulation of these important animals.

Maintenance of Caenorhabditis elegans

Ceanorhabditis elegans has been, and is still, used to great success as a model organism for studying a variety of developmental, genetic, molecular and ...

A Flame-Free Method for Sterilizing C. elegans Picks, Spatulas, and Scalpels

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The Nematode Caenorhabditis Elegans - A Versatile In Vivo Model to Study Host-microbe Interactions

In Vitro Reconstitution of Spatial Cell Contact Patterns with Isolated Caenorhabditis elegans Embryo Blastomeres and Adhesive Polystyrene Beads

In Vivo Visualization of Calcium Transients during Fertilization and Early Development in C. elegans

A Method for Culturing Embryonic C. elegans Cells

Ablation of a Single Cell From Eight-cell Embryos of the Amphipod Crustacean Parhyale hawaiensis

Using Single-Worm Data to Quantify Heterogeneity in Caenorhabditis elegans-Bacterial Interactions

Hand Dissection of Caenorhabditis elegans Intestines

Long-Term Culture and Monitoring of Isolated Caenorhabditis elegans on Solid Media in Multi-Well Devices

Aseptic Technique in Environmental Science

C. elegans Maintenance