The authors thank Dr. Atsushi Higashitani for kind advice regarding treatment of C. elegans and Drs. Yuya Hattori, Yuichiro Yokota, and Yasuhiko Kobayashi for valuable discussions. The authors thank the Caenorhabditis Genetic Center for providing strains of C. elegans and E. coli. We thank the crew of the cyclotron of TIARA at QST-Takasaki for their kind assistance with the irradiation experiments. We thank Dr. Susan Furness for editing a draft of this manuscript. This study was supported in part by KAKENHI (Grant Numbers JP15K11921 and JP18K18839) from JSPS to M.S.

1. Strains and maintenance
<ol>
	<li>Select a suitable strain of C. elegans and Escherichia coli (food) depending on the purpose of the experiment. 
	NOTE: In the present paper, wild-type N210C. elegans (Figure 2A) is generally used, and HBR4:goeIs3[pmyo-3::GCamP3.35::unc-54 - 3&#39;utr, unc-119(+)]V11 is only employed for imaging assay. E. coli OP50 was used as food for C. elegans. Some mutants with abnormal body shape, such as the unc-119(e2498) III mutant with a coiled shape (Figure 2B), also can be enclosed in the straight microfluidic channels (Figure 2C).</li>
	<li>Maintain the C. elegans at 20 &#176;C on 6 cm Petri dishes containing 10 mL of nematode growth medium (NGM) spread with overnight-incubated (37 &#176;C) E. coli as described previously10. If possible, synchronize the developmental stages of C. elegans from the embryo stage.</li>
	<li>Use well-fed adult animals, approximately 3-4 days after hatching with a width of about 50-60 &#181;m, which are optimally suited to the size of the microfluidic channels in the worm sheets.</li>
</ol>
2. Selection of buffer solution for on-chip immobilization
<ol>
	<li>For wettable worm sheets, use any buffer solution depending on the purpose of the experiments. 
	NOTE: Suitable buffer solutions for on-chip immobilization on PDMS microfluidic chips have been defined previously9 and the following buffer solutions were shown to have no effect on the motility of the animals after immobilization: S basal buffer solution (5.85 g of NaCl, 1 mL of cholesterol (5 mg/mL in ethanol), 50 mL of 1 M pH 6.0 potassium phosphate, H2O to 1 L; sterilized by autoclaving)10 containing a large amount of NaCl, M9 phosphate buffer solution (5 g of NaCl, 3 g of KH2PO4, 6 g of Na2HPO4, 1 mL of 1 M MgSO4, H2O to 1 L; sterilized by autoclaving)10, wash buffer solution (5 mL of 1 M pH 6.0 potassium phosphate, 1 mL of 1 M CaCl2, 1 mL of 1 M MgSO4, 0.5 g pf gelatin, H2O to 1 L; sterilized by autoclaving)12 containing gelatin, and ultrapure water9. In the present paper, wash buffer solution is typically used, and S basal buffer solution is only employed for evaluating on-chip immobilization using different cover films in the section 3.</li>
	<li>For conventional microfluidic chips without wettability, use a wash buffer solution, which provides the most effective means of maintaining moisture in the microfluidic channels of the chip and thus prevents drying of C. elegans individuals (regardless of the wettability of the microfluidic chips).</li>
</ol>
3. Selection of suitable cover film for on-chip immobilization
<ol>
	<li>Prepare different cover films as follows. Based on the ease of handling, cut the following transparent, biocompatible cover films to a suitable size (i.e., width of 10-15 mm and length of 30-50 mm): 130-170 &#181;m thick cover glass, 125 &#181;m thick polyester (PET) film, and ~130 &#181;m thick polystyrene (PS) film.</li>
	<li>Perform 3 h on-chip immobilization using different cover films as follows. Based on steps 4.1-4.6, enclose &#8805;10 washed adult animals&#160;(3.5 days after hatching) in the worm sheet using each cover film, respectively, and leave for 3 h.</li>
	<li>For comparison purposes, allow 3 h of free movement as follows. Place &#8805;10 washed adult animals on a 3.5 cm plate containing 3 mL fresh NGM, cover with a lid, and leave for 3 h.</li>
	<li>Collect animals immediately after 3 h of on-chip immobilization according to steps 5.1-5.3. Remove the cover film from the worm sheet and add a drop of buffer solution to each animal.&#160;Pick up animals swimming in the droplet using a platina picker and transfer them to a 3.5 cm plate containing 3 mL fresh NGM without food (assay plate).</li>
	<li>Collect animals after 3 h of free movement. Add a drop of buffer solution to each animal. Pick up animals swimming in the droplet on the NGM plate and transfer them to an assay plate using a platina picker.</li>
	<li>Evaluate the effects of cover film on motility (locomotion assay).
	<ol>
		<li>At least 5 min after transfer to the assay plate, count &#39;body bends&#39; (de&#64257;ned as the number of bends in the anterior body region at 20-s intervals) manually under a microscope13.</li>
		<li>Carry out locomotion assays five times independently for each cover film. 
		NOTE: In the experiments shown in the representative results, 10 animals were evaluated for each experiment in which multiple animals were enclosed simultaneously, compared with only five animals counted in the previous study13.</li>
		<li>Calculate the average number of body bends for the 10 animals in each group for each experiment. Then average the values from five independent experiments to evaluate the e&#64256;ects of on-chip immobilization. Analyze the data statistically using a one-way analysis of variance (ANOVA) test in a spreadsheet software at significance levels of 0.01 and 0.05. 
		NOTE: Based on these experiments, a PS cover film with high oxygen permeability was deemed suitable (see the representative results). These PS films are included with the worm sheets. PET film, which has low oxygen permeability, was not deemed suitable because the animals tended to suffocate if covered with it during long-term immobilization on the worm sheets. It is also important that the cover film does not break during a series of procedures; glass covers&#160;were therefore also unsuitable (see the representative results).</li>
	</ol>
	</li>
</ol>
4. On-chip immobilization
NOTE: Disposable, sterile gloves should be worn to avoid contaminating the worm sheets.
<ol>
	<li>Place a thin transparent sheet such as a PS or glass cover film which has no autofluorescence, for use as a bottom cover film, onto the experimental desk or the microscopic stage. Place a worm sheet gently onto the bottom cover film using flat tweezers. 
	NOTE: In addition to the worm sheets with 60 &#181;m-wide microfluidic channels (suitable for adults 3-4 days after hatching), sheets with 50 &#181;m-wide channels (suitable for young adults 3 days after hatching and mutants with small body in adult stage) can also be employed. Select a suitable sheet based on the size of the animals to be used.</li>
	<li>To collect animals, pick up an individual adult C. elegans from the culture plate under a stereomicroscope using a platina picker. Repeat the process if multiple animals are needed.</li>
	<li>Wash animals to remove food (bacteria).
	<ol>
		<li>Place at least three droplets (5 &#181;L drops) of buffer solution on the surface of a 6 cm non-treated (water-repellent) Petri dish.</li>
		<li>Transfer the animals to a droplet using a platina picker and allow them to remove any food by swimming. Wash them twice in two separate droplets, and then rinse the animals in another droplet. 
		NOTE: It is necessary to practice this procedure to be able to perform it quickly before carrying out any experiments.</li>
	</ol>
	</li>
	<li>Drop 2-3 &#181;L of buffer solution onto the surface of a worm sheet. Pick up the washed animals from the droplet on the Petri dish and transfer them to the droplet on the worm sheet.</li>
	<li>Enclose animals in microfluidic channels as follow. Place a PS cover film over the worm sheet using flat tweezers and press gently over the channels from one end of the sheet to the other to maintain humidity (as described previously4,9). 
	NOTE: Animals are randomly enclosed in the channels. Droplets containing animals are spread across the worm sheet by covering the sheet, resulting in droplets being widely extended over several channels. Each animal can be enclosed in any one of these channels. The important point regarding the use of the worm sheet for on-chip immobilization is that multiple channels are available over a wide area.</li>
	<li>Immediately after enclosing the animals in the microfluidic channels, confirm that they are alive by checking for movement of the head under a microscope (e.g., 1x or 2x magnification).</li>
	<li>Record animal&#39;s position, at the same time as carrying out step 4.6 and note the following on a dedicated sheet of paper (Supplementary File 1): number of the channel in which the animal is enclosed; position of each animal in the channel (left/center/right); and direction of the head.</li>
	<li>Draw an arrow to mark the position of each animal in the channel, with the direction of the arrow corresponding to the animal&#39;s head. 
	NOTE: The channel numbers (e.g., 1, 5, 10, 15, 20, 25) are engraved on the surface of the worm sheet near the left and right edges of the microfluidic channels (Figure 1B). This information on the dedicated sheet allows animals to be located quickly, making the process more efficient, and also helps to prevent radiation leakage during subsequent steps.</li>
</ol>
5. Collection of animals from a worm sheet
<ol>
	<li>Immediately after immobilization, remove the cover film from the worm sheet using flat tweezers.</li>
	<li>Drop 10-15 &#181;L buffer solution onto the microfluidic channels enclosing the animals and observe the animals starting to swim in the droplets under a stereomicroscope. 
	NOTE: The animals start swimming in the new droplet on their own without any additional pressure, help, or push. Just dropping some buffer on top of them is enough to make them swim out of the channel. An animal may be unable to swim in a droplet if it has been vitally damaged by irradiation in step 7.9 or by the immobilization process in steps 4.2-4.5, though this is rare. In addition, if the animal remains attached to the cover film when it is removed, add the droplets onto the cover film instead of into the channels.</li>
	<li>Pick up the animals swimming in the droplet using a platina picker and place them on an assay plate.</li>
</ol>
6. Application of worm sheets for imaging observations
NOTE: Worm sheets can be widely used in microscopic observations. The chip can retain water and does not affect the motility of C. elegans individuals after 3 h of on-chip immobilization9. In addition, the chip itself has no autofluorescence, making it suitable for use in fluorescence imaging assays. A sample application for fluorescence imaging assay is given below.
<ol>
	<li>Select a fluorescence microscope and mount a digital camera or digital video camera on the microscope to capture images or videos, respectively (Figure 3A). 
	NOTE: The working distance (WD) of microscope is more than 0.2 mm depending on objective lens specification (see Figure 3B,C). The specification of the fluorescence microscope system used in this paper is shown in the Table of Materials. However, the specification is not limited to our example because it depends on the purpose of observation or/and users.</li>
	<li>Select any fluorescent strain of C. elegans and maintain as described in section 1. 
	NOTE: If it is necessary to observe the body-wall muscular contraction (see representative results), use young adult HBR411C. elegans, in which a reporter gene is used to express the calcium indicator GCaMP3.35 in all body-wall muscle cells. It is important to use young adults (&#8804;3 days after hatching) that are thinner than the width of the microfluidic channels (50 or 60 &#181;m) of the worm sheet. The small degree of clearance means that the animals can bend slightly, making it possible to observe the muscular contraction and extension during crawling, using a calcium-ion indicator.</li>
	<li>Carry out enclosure of animals according to the immobilization procedure in section 4.</li>
	<li>Observe fluorescent spots of animals (e.g., green fluorescent protein-labeled strains) using a fluorescence microscope, and capture images using a digital camera mounted on the microscope. 
	NOTE: Follow the previously established methods14,15,16, since the microscope observation methods (including fluorescence observation) and the specification of the microscope system depends on the purpose of the observation.</li>
	<li>Image calcium-ion wave propagation using video acquisition to observe dynamic activities, such as the body-wall muscular contraction and extension in the HBR4 worms (Video 1).</li>
</ol>
7. Application of worm sheets for microbeam irradiation
NOTE: The collimating microbeam irradiation system5 can use several heavy-ion particles accelerated from the azimuthally varying field cyclotron installed at the Takasaki Ion Accelerators for Advanced Radiation Application (TIARA) facility of QST-Takasaki (Figure 4A). There is an automatic stage for irradiation under the beam exit (Figure 4B). The procedure for heavy-ion microbeam irradiation of C. elegans using this system is as follows.
<ol>
	<li>Locate the worm sheet enclosing multiple animals on an aluminum frame custom-made for the microbeam-irradiation facility and set it on the automatic stage for irradiation (Figure 4C).</li>
	<li>Raise the irradiation sample (i.e., animals enclosed in a worm sheet on a frame), to immediately under (~2 mm) the beam exit based on the monitor image from a microscope located under the automatic stage (Figure 4D).</li>
	<li>After vertical positioning, locate each animal to target with the microbeam irradiation using the custom-made software for targeted irradiation of animals and cell cultures. To locate each animal enclosed in the microfluidic channel, refer to the dedicated sheet described in step 4.7-4.8 (see Supplementary File 1) and confirm the channel number near the left and right edges of the channels.</li>
	<li>Control the automatic stage in the X and Y directions to position the animals just under the beam exit using a remote-control system and a laser mouse operated with the irradiation software.</li>
	<li>Roughly tune the irradiation area by watching the projection from the microscope located under the automatic stage of the sample and moving the cursor to the irradiation position of the animal to be targeted.</li>
	<li>After rough positioning, exit and close the irradiation room and move to the adjacent control room to avoid irradiating the operators.</li>
	<li>Set the desired number of ion particles for one irradiation procedure, corresponding to irradiation of a specific region of the animal, using the console linked to an ion-counter system. 
	NOTE: This consists of a plastic scintillator and a photoelectron multiplier in the collimating microbeam irradiation system.</li>
	<li>From the irradiation-control room, fine-tune the position of the animals based on the monitor image from the microscope located under the automatic stage, which is the same image projected on the monitor in the irradiation room (Figure 4E).</li>
	<li>Target the microbeam irradiation by click the irradiation button of the software for irradiation. 
	NOTE: In the example shown in Figure 4F, we target the pharynx in the head region and irradiate with the appropriate number of microbeam carbon ions. Because the number of ion particles passing through the sample is counted using an ion-counter system linked to the console and the irradiation software, the irradiation is stopped after delivery of the desired number of ion particles.</li>
	<li>Locate each animal recorded on the dedicated sheet and carry out targeted irradiation to each animal. Repeat steps 7.8 and 7.9 until all animals have been irradiated.</li>
	<li>Immediately after irradiation, enter the irradiation room and lower the automatic irradiation stage. Remove the sample on the custom-made frame located on the automatic stage.</li>
	<li>Collect the animals as described in steps 5.1-5.3.</li>
	<li>Carry out behavioral and/or molecular analyses as required to evaluate the effects of the targeted irradiation, depending on the purpose of the study. 
	NOTE: The locomotion assay described in step 3.6 is an effective method for evaluating the effects of irradiation on motility4,9.</li>
</ol>
8. Treatment of worm sheets for repeated use
NOTE: Worm sheets can be used repeatedly at least 10 times9 with no adverse effects on the animals if cleaned and sterilized properly after use as follows.
<ol>
	<li>Cleaning
	<ol>
		<li>Place the used worm sheet on a 6 cm Petri dish and drop about 100 &#181;L of sterilized ultrapure water onto the whole sheet.</li>
		<li>Paddle the water on the surface of the sheet using gloved fingers to wash off dirt such as dust, bacterial food, and any eggs laid in the channels.</li>
		<li>Wipe the moisture off the worm sheet thoroughly using disposable wipes.</li>
	</ol>
	</li>
	<li>Sterilization
	<ol>
		<li>Inject about 5 mL of 70% ethanol into the Petri dish containing the worm sheet and paddle the surface of the sheet using gloved fingers to wash off the dirt.</li>
	</ol>
	</li>
	<li>Drying
	<ol>
		<li>Remove the chips from the Petri dish filled with 70% ethanol and allow to dry naturally.</li>
	</ol>
	</li>
	<li>Storage
	<ol>
		<li>After drying, place the worm sheet on a sterile Petri dish and cover it. Sheets can also be stored on a sterile Petri dish filled with 70% ethanol. 
		NOTE: It is better to dispose of the cover films after use, but if they are to be re-used, clean them as done for the worm sheet. However, if folds develop on the cover film after use it will no longer adhere to the chip, resulting in dehydration of the animals, and it should therefore be replaced.</li>
	</ol>
	</li>
</ol>

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<li>Fehlauer, H., Nekimken, A. L., Kim, A. A., Pruitt, B. L., Goodman, M. B., Krieg, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Using+a+microfluidics+device+for+mechanical+stimulation+and+high+resolution+imaging+of+C.+elegans%5BTitle%5D)+AND+%22Journal+of+Visualized+Experiments%22%5BJournal%5D)">Using a microfluidics device for mechanical stimulation and high resolution imaging of C. elegans.</a> Journal of Visualized Experiments. (132), e56530(2018).</li>
</ol>

The authors have nothing to disclose.

On-chip immobilization of C. elegans under live conditions using a wettable PDMS microfluidic chip enables the efficient targeted microbeam irradiation of multiple animals. The ease of handling and features to prevent drying make this system suitable for applications not only in microbeam irradiation, but also in several behavioral assays. These worm sheets have already been commercialized and can be easily obtained. Conventional microfluidic chips, such as olfactory chips, are associated with problems including clogging of animals and eggs in the closed microfluidic channels making it difficult to collect the animals, and thus such chips have tended to be disposable, thereby increasing the cost. In contrast, the microfluidic channels in the current worm sheet are open, making it easier to collect the animals. These worm sheets can therefore be used repeatedly, making them more economical.
Recent technological innovations in PDMS microfluidic chips have shown an increasing trend in structural complexity and multifunctionality16,18,19,20,21,22,23. However, we believe that it is important to make the system simple and easy to use. Indeed, in contrast to the use of conventional large microfluidic chips19,20,21,22,23 that require the attachment of a vacuum pump, the small size and simple design of the worm sheets allow procedures to be conducted easily within a limited space.
The water retention performance of the worm sheets enables long-term observations to be carried out. In addition, the thickness of the sheet allows ion particles to pass through the samples, thus enabling targeted irradiation to be applied to live C. elegans individuals with a precise number of ion particles. These advantages of the worm sheets have greatly expanded the possibilities for biological experiments.
We believe that it is important for biologists to develop new equipment and methods in order to improve the efficiency of their experiments and analyses. The worm sheets and microbeam irradiation software have been developed with this aim in mind, and have the potential to contribute to the success of future innovative experiments beyond their original objectives.
To the best of our knowledge, our group is the first to develop this technology worldwide. However, its standardized use in the future will facilitate the application of targeted microbeam irradiation to animals under live conditions, thus helping to identify the roles of specific cells/tissues in internal processes.

Radiation, including X-rays, gamma rays, and heavy-ion beam, is widely used for biological applications such as in cancer diagnosis and treatment, and for ion-beam breeding. Numerous studies and technical developments are currently focusing on the effects of radiation1,2,3. Microbeam irradiation is a powerful means of identifying radiosensitive sites in living organisms4. The Takasaki Advanced Radiation Research Institute of National Institutes for Quantum and Radiological Science and Technology (QST-Takasaki) has been developing a technology to irradiate individual cells under microscopic observation using heavy-ion microbeams5, and has established methods to enable targeted microbeam irradiation of several model animals, such as the nematode Caenorhabditis elegans4,6, silkworms7, and Oryzias latipes (Japanese medaka)8. Targeted microbeam irradiation of the nematode C. elegans allows the effective knockdown of specific regions, such as the nerve ring in the head region, thus helping to identify the roles of these systems in processes such as locomotion.
A method for on-chip immobilization of C. elegans individuals without the need for anesthesia has been developed to allow for microbeam irradiation4. In addition, to improve microfluidic chips used in the previous study4, we have recently developed wettable, ion-penetrable, polydimethylsiloxane (PDMS) microfluidic chips, referred to as worm sheets (see Table of Materials), for immobilizing C. elegans individuals9. These comprise of ultra-thin soft sheets (thickness = 300 &#181;m; width = 15 mm; length = 15 mm) with multiple (20 or 25) straight microfluidic channels (depth = 70 &#181;m; width = 60 &#181;m or 50 &#181;m; length = 8 mm) at the surface (Figure 1A-D). The microfluidic channels are open and allow multiple animals to be enclosed in them simultaneously (Figure 1E). The sheets are resilient to allow the microfluidic channels to be expanded (by ~10%, Figure 1F), and the elasticity allows animals to be enveloped gently. Also, owing to the self-adsorption capacity of the PDMS, animals can be sealed in the channels by covering the surface of the worm sheet with a thin cover film, in which animals are not pushed into the channels for enclosure. By turning the cover film over, we can easily collect the animals.
The channels do not hurt the worms when they are being enclosed or when they are collected. Furthermore, the sheets are made from PDMS, which is essentially hydrophobic, but water retention can be achieved by imparting hydrophilicity to the material. The water retention and thickness are favorable characteristics of the worm sheets. The water-retention capacity prevents dehydration of the animals after prolonged immobilization and enables long-term observations to be carried out.
In addition, as described previously9, the sheets are only 300 &#181;m thick, allowing heavy ions such as carbon ions (with a range of about 1 mm in water) to pass through the sheet enclosing the animals. This allows the ion particles to be detected and the applied radiation dose to be measured accurately. Moreover, the worm sheets can be reused and are thus economical. With the conventional injection method, the animals enclosed are sometimes dead and they cannot be taken out of the channel; their eggs can also clog the channels. This makes the chip unusable. Chips are, therefore, basically disposable and the cost-benefit ratio is poor.
In the present paper, we describe in detail a series of methods for on-chip immobilization of live C. elegans individuals using worm sheets. Through locomotion assays of animals 3 h after on-chip immobilization, we evaluated the suitable cover film. In addition, we showed the examples of on-chip immobilization for both imaging observations and microbeam irradiation.

<table><tbody><tr><td>&lt;em&gt;C. elegans &lt;/em&gt;wild-type strain</td><td>Caenorhabditis Genetics Center (CGC) , Minnesota, USA</td><td>N2</td><td>Wild-type &lt;em&gt;C. elegans&lt;/em&gt; strain generally used in this study</td></tr><tr><td>&lt;em&gt;C. elegans unc-119(e2498) III &lt;/em&gt;mutant strain</td><td>Caenorhabditis Genetics Center (CGC) , Minnesota, USA</td><td>CB4845</td><td>&lt;em&gt;C. elegans&lt;/em&gt; strain only employed as an example of mutants with abnormal body shape&amp;nbsp;</td></tr><tr><td>&lt;em&gt;C. elegans transgenic strain&lt;/em&gt; HBR4</td><td>Caenorhabditis Genetics Center (CGC) , Minnesota, USA</td><td>HBR4</td><td>The genotype of this transgenic &lt;em&gt;C. elegans&lt;/em&gt; strain is HBR4:&lt;em&gt;goeIs3[pmyo-3::GCamP3.35:: unc-54&amp;ndash;3&amp;rsquo;utr, unc-119(+)]V&lt;/em&gt;. This strain was only employed for imaging observation.</td></tr><tr><td>&lt;em&gt;E. coli strain&lt;/em&gt;</td><td>Caenorhabditis Genetics Center (CGC) , Minnesota, USA</td><td>OP50</td><td>&lt;em&gt;E. coli&lt;/em&gt; strain used as food for &lt;em&gt;C. elegans&lt;/em&gt;</td></tr><tr><td>Worm Sheet IR (50/60)</td><td>Biocosm, Inc., Hyogo, Japan</td><td>BCM17-0001</td><td>Microfluidic chip with 25 straight 50/60-&amp;micro;m width channels used in all experiments and observation in this paper&amp;nbsp;</td></tr><tr><td>Worm Sheet 60</td><td>Biocosm, Inc., Hyogo, Japan</td><td>BCM18-0001</td><td>Microfluidic chip with 20 straight 60 &amp;micro;m-width channels. This is sitable for adults 3-5 days after hatching at 20&amp;deg;C.&amp;nbsp;</td></tr><tr><td>Worm Sheet 50</td><td>Biocosm, Inc., Hyogo, Japan</td><td>BCM18-0002</td><td>Microfluidic chip with 20 straight 50 &amp;micro;m-width channels. This is sitable for youg adults ~3 days after hatching at 20&amp;deg;C.&amp;nbsp;</td></tr><tr><td>MICRO COVER GLASS</td><td>MATSUNAMI GLASS IND. LTD.</td><td>C030401</td><td>Cover glass (thickness: 130-170 &amp;micro;m) used in locomotion assays in Protocol 3</td></tr><tr><td>Polystyrene Film</td><td>Biocosm, Inc., Hyogo, Japan</td><td>BCM18-0001/ BCM18-0002</td><td>Bundled items of Worm Sheets. PS filim (thickness: ~130 &amp;micro;m) used in locomotion assays in Protocol 3.</td></tr><tr><td>Polyester Film Lumirror</td><td>TORAY INDUSTRIES, INC., Tokyo, Japan</td><td>Lumirror T60 (t 125 &amp;micro;m)</td><td>PET filim (thickness: 125 &amp;micro;m) used in locomotion assays in Protocol 3</td></tr><tr><td>IWAKI 60 mm/non-treated dish</td><td>AGC Techno Glass Co., Ltd., Shizuoka, Japan).</td><td>1010-060</td><td>Non-treated dish used in incuvation of &lt;em&gt;C. elegans&lt;/em&gt; in Protocol 1</td></tr><tr><td>IWAKI 35 mm/non-treated dish</td><td>AGC Techno Glass Co., Ltd., Shizuoka, Japan).</td><td>1010-035</td><td>Non-treated dish used in locomotion assays in Protocol 3</td></tr><tr><td>Milli-Q</td><td>Merck, France</td><td></td><td>Ultrapure water</td></tr><tr><td>Kimwipe S-200</td><td>Nippon Paper Crecia Co., Ltd., Tokyo, Japan</td><td>62020</td><td>120 mm x 215 mm; 200 sheets/ box</td></tr><tr><td>WormStuff Worm Pick</td><td>Genesee Scientific Corporation, CA, USA)</td><td>59-AWP</td><td>Platina picker specilized for picking up &lt;em&gt;C. elegans&lt;/em&gt;</td></tr><tr><td>Research Stereo Microscope System</td><td>OLYMPUS CORPORATION, Tokyo, Japan</td><td>SZX16</td><td>Micriscope used in all experiments and observation in this paper</td></tr><tr><td>Motorized Focus Stand for SZX16</td><td>OLYMPUS CORPORATION, Tokyo, Japan</td><td>SZX2-ILLB</td><td>This was used for bright field observation in Protocol 3-8.</td></tr><tr><td>Objective Lens (&amp;times;1)</td><td>OLYMPUS CORPORATION, Tokyo, Japan</td><td>SDFPLAPO1&amp;times;PF</td><td>NA: 0.15; W.D.: 60 mm. This lends was used for bright field observation in Protocol 3-8.</td></tr><tr><td>Objective Lens (&amp;times;2)</td><td>OLYMPUS CORPORATION, Tokyo, Japan</td><td>SDFPLAPO2XPFC</td><td>NA: 0.3; W.D.: 20 mm. This&amp;nbsp; lends was used for imaging observations.</td></tr><tr><td>Mercury Light Source</td><td>OLYMPUS CORPORATION, Tokyo, Japan</td><td>U-LH100HG</td><td>The broad emission spectrum enables a range of fluorescence imaging experiments to be conducted using all common fluorophores.</td></tr><tr><td>SZX16 Fluorescent filter unit (High performance for YFP)</td><td>OLYMPUS CORPORATION, Tokyo, Japan</td><td>SZX2-FYFPHQ</td><td>Ex: 490-500 nm; Em: 510-560. This was used for imaging observation of HBR4 strain.</td></tr><tr><td>Digital Camera High Speed EXILIM</td><td>CASIO COMPUTER Co., Ltd, Tokyo, Japan</td><td></td><td>EX-F1 Figure 1B, 1E, 1F, Figure 2A-C, and Video 1 were obtained by using this digital camera.</td></tr><tr><td>Office 2013</td><td>Microsoft Co. Ltd, Redmond, WA, USA</td><td></td><td>EXCEL Software used for statistical analyses</td></tr></tbody></table>

immobilization of live caenorhabditis elegans individuals using an ultra-thin polydimethylsiloxane microfluidic chip with water retention

Radiation is widely used for biological applications and for ion-beam breeding, and among these methods, microbeam irradiation represents a powerful means of identifying radiosensitive sites in living organisms. This paper describes a series of on-chip immobilization methods developed for the targeted microbeam irradiation of live individuals of Caenorhabditis elegans. Notably, the treatment of the polydimethylsiloxane (PDMS) microfluidic chips that we previously developed to immobilize C. elegans individuals without the need for anesthesia is explained in detail. This chip, referred to as a worm sheet, is resilient to allow the microfluidic channels to be expanded, and the elasticity allows animals to be enveloped gently. Also, owing to the self-adsorption capacity of the PDMS, animals can be sealed in the channels by covering the surface of the worm sheet with a thin cover film, in which animals are not pushed into the channels for enclosure. By turning the cover film over, we can easily collect the animals. Furthermore, the worm sheet shows water retention and allows C. elegans individuals to be subjected to microscopic observation for long periods under live conditions. In addition, the sheet is only 300 &#181;m thick, allowing heavy ions such as carbon ions to pass through the sheet enclosing the animals, thus allowing the ion particles to be detected and the applied radiation dose to be measured accurately. Because selection of the cover films used for enclosing the animals is very important for successful long-term immobilization, we conducted the selection of the suitable cover films and showed a recommended one among some films. As an application example of the chip, we introduced imaging observation of muscular activities of animals enclosing the microfluidic channel of the worm sheet, as well as the microbeam irradiation. These examples indicate that the worm sheets have greatly expanded the possibilities for biological experiments.

Active C. elegans individuals could be immobilized successfully using an ultra-thin, wettable PDMS, microfluidic chip (worm sheet). We investigated the suitability of different cover films for sealing the worm sheet, as described in protocol section 3. To evaluate the sealing effects of the cover films, we determined the motility of animals 3 h after on-chip immobilization using cover glass (thickness: 130-170 &#181;m), PET film (thickness: 125 &#181;m), and PS film (thickness: ~130 &#181;m), respectively. As shown in Figure 5, there was no significant difference in motility (body bends) between control animals allowed to move freely for 3 h and animals enclosed in the worm sheet with PS film. In contrast, motility was significantly reduced in animals enclosed under a cover glass. Some animals appeared to have dried out, suggesting that the cover glass repelled the droplet, preventing a close seal and allowing the animals to partially dry out, resulting in reduced motility. The motility of animals enclosed using a PET film was also significantly decreased; although no drying was observed, the animals&#39; motility tended to decrease uniformly, suggesting that the low oxygen transmission rate (~30 mL/[24 h&#183;m&#178;&#183;MPa]), which is about 100 times lower than that of PS, caused the animals to suffocate. These results suggest that PS cover films should be used to enclose animals&#160;in the worm sheet.
We also applied the worm sheet technique for imaging observations and to perform region-specific microbeam irradiation. On-chip immobilization using a worm sheet with water retention and no autofluorescence was suitable for microscopic observation under live conditions. For example, we applied the technique to the HBR4 strain11 of C. elegans, in which a reporter gene expressed the calcium indicator GCaMP3.35 in all body-wall muscle cells. We observed the activities of all body-wall muscle cells in young adult animals at &#8804;3 days post-hatching in worm sheets with 50 &#181;m-wide microfluidic channels, which allowed the animals space to bend slightly. The GCaMP3.35 signal intensity in the HBR4 strain corresponds to the contraction of the body-wall muscle cells. The Ca2+ wave propagation corresponding to the muscular activity was clearly observed (Video 1). Additionally, we confirmed that the worm sheet had no autofluorescence as shown in the last stage (last ~10 s) of Video 1. In this way, the lack of need for anesthesia allowed the physiological activities of the muscle cells to be observed under live conditions.
Furthermore, we applied the worm sheet for region-specific microbeam irradiation of C. elegans individuals. The multiple straight microfluidic channels on the worm sheet allowed multiple animals to be immobilized simultaneously, without the need for anesthesia, thus allowing sequential irradiation of &#8805;20 animals (enough for a group assay) in a short time (30 min for 20 individuals).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/59008/59008fig1.jpg" /> 
Figure 1: Schematic of a worm sheet. (A) Overview of a worm sheet with an American 1 cent coin for scale. The worm sheet was 300 &#181;m thick, 15 mm wide, and 15 mm long. (B) The surface of the worm sheet contained 25 straight microfluidic channels (depth = 70 &#181;m; width = 60 &#181;m; length = 8 mm). (C) Schematic of the samples consisting of the bottom cover film, the worm sheet, and the cover film. (D) The worm sheet is a soft, ultra-thin sheet made from PDMS, and can be bent by pinching with flat tweezers. (E) Example of multiple animals enclosed in multiple channels.(F) Expansion of a microfluidic channel by pushing with a platina picker. The elasticity of the channel allows animals to be enveloped gently. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/59008/59008fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/59008/59008fig2.jpg" /> 
Figure 2: Body form of C. elegans. (A) Wild-type (N2) C. elegans on an NGM plate. (B) An unc-119 mutant with abnormal shape on an NGM plate. (C) The unc-119 mutants enclosed in the microfluidic channels of the worm sheet. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/59008/59008fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/59008/59008fig3.jpg" /> 
Figure 3: Schematics of microscope observation of live C. elegans individuals enclosed in the worm sheet. (A) Schematic of the stereomicroscope system for imaging observations. (B) Schematic of microscope observation of the worm sheet enclosing live C. elegans individuals. (C) Sectional view of the worm sheet enclosing live C. elegans individuals placed on the microscope stage. W.D. indicates the working distance of microscope. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/59008/59008fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/59008/59008fig4.jpg" /> 
Figure 4: Schematic of collimating microbeam system at QST-Takasaki and targeted microbeam irradiation procedure for live C. elegans individuals. (A) Overview of the collimating microbeam irradiation system5, which can use several heavy-ion particles accelerated from the azimuthally varying field cyclotron installed at the TIARA of QST-Takasaki. (B) Overview of the beam exit and the automatic stage for irradiation. (C) Sample setting on the automatic stage of the collimating microbeam system. (D) Vertical positioning of the irradiation sample conducted in the irradiation room. (E) Fine-tuning of irradiation area conducted in the irradiation-control room. (F) Targeted microbeam irradiation of live C. elegans. The pharynx was clicked-on as the targeted position and irradiated by pushing the irradiation button. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/59008/59008fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/59008/59008fig5.jpg" /> 
Figure 5: Motility of C. elegans after on-chip immobilization using cover glass, polyester (PET) film, and polystyrene (PS) film. Bars indicate mean body bends of animals 3 h after on-chip immobilization or after free movement for 3 h on an NGM plate (control). Ten animals were examined and body bends were averaged among each group. Finally, data from five independent experiments were averaged for each group. Error bars represent standard error of the mean of five independent experiments. All data were analyzed using one-way ANOVA at the 0.05 (*) or 0.01 (**) significance level. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/59008/59008fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Video 1" src="/files/ftp_upload/59008/59008video1.jpg" /> 
Video 1: Examples of imaging observations of muscular activities in C. elegans enclosed in a worm sheet. Calcium-ion wave propagation corresponding to contraction of the body-wall muscle cells during crawling in HBR4 C. elegans individuals enclosed in a worm sheet. The last ~10 s were observed under bright-field illumination. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/59008/Video_1_R1.mp4" target="_blank">Please click here to view this video. (Right-click to download.)</a>
<img alt="Supplementary Figure 1" src="/files/ftp_upload/59008/59008supfig1.jpg" /> 
Supplementary File 1: Example of dedicated sheet of paper (microbeam irradiation version). Draw an arrow to indicate the position of each animal in the channel. The direction of the arrow corresponds to the head. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/59008/Supplement_1_R2.tif" target="_blank">Please click here to download this file.</a>

Watch this Scientific Journal Video about Immobilization of Live Caenorhabditis elegans Individuals Using an Ultra-thin Polydimethylsiloxane Microfluidic Chip with Water Retention at JoVE.com

Immobilization of Live Caenorhabditis elegans Individuals Using an Ultra-thin Polydimethylsiloxane Microfluidic Chip with Water Retention

A series of immobilization methods has been established to allow the targeted irradiation of live Caenorhabditis elegans individuals using a recently developed ultra-thin polydimethylsiloxane microfluidic chip with water retention. This novel on-chip immobilization is also adequate for imaging observations. The detailed treatment and application examples of the chip are explained.

Immobilization of Live Caenorhabditis elegans Individuals Using an Ultra-thin Polydimethylsiloxane Microfluidic Chip with Water Retention

Radiation is widely used for biological applications and for ion-beam breeding, and among these methods, microbeam irradiation represents a powerful means ...

immobilization-live-caenorhabditis-elegans-individuals-using-an-ultra

Nanyang Technological University

Research

JoVE Journal

Bioengineering

7.7K Views.  National Institutes for Quantum and Radiological Science and Technology (QST-Takasaki). Een voordeel van dit protocol is dat het wormblad gemakkelijk te hanteren is. Het blad heeft functies die drogen voorkomen, waardoor we microbeam bestraling kunnen uitvoeren naar levende dieren, verschillende gedragstesten kunnen uitvoeren en observatie op lange termijn kunnen uitvoeren. De microfluïde kanalen in dit wormenblad zijn open, waardoor het makkelijker wordt om dieren te verzamelen.Deze worm platen kunnen ook herhaaldelijk worden gebruikt, waardoor ze zuiniger. Toen we deze...

Video: Immobilization of Live Caenorhabditis elegans Individuals Using an Ultra-thin Polydimethylsiloxane Microfluidic Chip with Water Retention

Fluorescence detection methods for microfluidic droplet platforms

The development of microfluidic platforms for performing chemistry and biology has in large part been driven by a range of potential benefits that accompany system miniaturisation. Advantages include the ability to efficiently process nano- to femoto- liter volumes of sample, facile integration of functional components, an intrinsic predisposition towards large-scale multiplexing, enhanced analytical throughput, improved control and reduced instrumental footprints.1
In recent years much interest has focussed on the development of droplet-based (or segmented flow) microfluidic systems and their potential as platforms in high-throughput experimentation.2-4 Here water-in-oil emulsions are made to spontaneously form in microfluidic channels as a result of capillary instabilities between the two immiscible phases. Importantly, microdroplets of precisely defined volumes and compositions can be generated at frequencies of several kHz. Furthermore, by encapsulating reagents of interest within isolated compartments separated by a continuous immiscible phase, both sample cross-talk and dispersion (diffusion- and Taylor-based) can be eliminated, which leads to minimal cross-contamination and the ability to time analytical processes with great accuracy. Additionally, since there is no contact between the contents of the droplets and the channel walls (which are wetted by the continuous phase) absorption and loss of reagents on the channel walls is prevented.
Once droplets of this kind have been generated and processed, it is necessary to extract the required analytical information. In this respect the detection method of choice should be rapid, provide high-sensitivity and low limits of detection, be applicable to a range of molecular species, be non-destructive and be able to be integrated with microfluidic devices in a facile manner. To address this need we have developed a suite of experimental tools and protocols that enable the extraction of large amounts of photophysical information from small-volume environments, and are applicable to the analysis of a wide range of physical, chemical and biological parameters. Herein two examples of these methods are presented and applied to the detection of single cells and the mapping of mixing processes inside picoliter-volume droplets. We report the entire experimental process including microfluidic chip fabrication, the optical setup and the process of droplet generation and detection.

Droplet-based microfluidic platforms are promising candidates for high throughput experimentation since they are able to generate picoliter, self-compartmentalized vessels inexpensively at kHz rates. Through integration with fast, sensitive and high resolution fluorescence spectroscopic methods, the large amounts of information generated within these systems can be efficiently extracted, harnessed and utilized.

The development of microfluidic platforms for performing chemistry and biology has in large part been driven by a range of potential benefits that ...

Simple Microfluidic Devices for in vivo Imaging of C. elegans, Drosophila and Zebrafish

Micro fabricated fluidic devices provide an accessible micro-environment for in vivo studies on small organisms. Simple fabrication processes are available for microfluidic devices using soft lithography techniques 1-3. Microfluidic devices have been used for sub-cellular imaging 4,5, in vivo laser microsurgery 2,6 and cellular imaging 4,7. In vivo imaging requires immobilization of organisms. This has been achieved using suction 5,8, tapered channels 6,7,9, deformable membranes 2-4,10, suction with additional cooling 5, anesthetic gas 11, temperature sensitive gels 12, cyanoacrylate glue 13 and anesthetics such as levamisole 14,15. Commonly used anesthetics influence synaptic transmission 16,17 and are known to have detrimental effects on sub-cellular neuronal transport 4. In this study we demonstrate a membrane based poly-dimethyl-siloxane (PDMS) device that allows anesthetic free immobilization of intact genetic model organisms such as Caenorhabditis elegans (C. elegans), Drosophila larvae and zebrafish larvae. These model organisms are suitable for in vivo studies in microfluidic devices because of their small diameters and optically transparent or translucent bodies. Body diameters range from ~10 &mu;m to ~800 &mu;m for early larval stages of C. elegans and zebrafish larvae and require microfluidic devices of different sizes to achieve complete immobilization for high resolution time-lapse imaging. These organisms are immobilized using pressure applied by compressed nitrogen gas through a liquid column and imaged using an inverted microscope. Animals released from the trap return to normal locomotion within 10 min.
We demonstrate four applications of time-lapse imaging in C. elegans namely, imaging mitochondrial transport in neurons, pre-synaptic vesicle transport in a transport-defective mutant, glutamate receptor transport and Q neuroblast cell division. Data obtained from such movies show that microfluidic immobilization is a useful and accurate means of acquiring in vivo data of cellular and sub-cellular events when compared to anesthetized animals (Figure 1J and 3C-F 4).
Device dimensions were altered to allow time-lapse imaging of different stages of C. elegans, first instar Drosophila larvae and zebrafish larvae. Transport of vesicles marked with synaptotagmin tagged with GFP (syt.eGFP) in sensory neurons shows directed motion of synaptic vesicle markers expressed in cholinergic sensory neurons in intact first instar Drosophila larvae. A similar device has been used to carry out time-lapse imaging of heartbeat in ~30 hr post fertilization (hpf) zebrafish larvae. These data show that the simple devices we have developed can be applied to a variety of model systems to study several cell biological and developmental phenomena in vivo.

Simple Microfluidic Devices for in vivo Imaging of C. elegans, Drosophila and Zebrafish

A simple microfluidic device has been developed to perform anesthetic free in vivo imaging of C. elegans, intact Drosophila larvae and zebrafish larvae. The device utilizes a deformable PDMS membrane to immobilize these model organisms in order to perform time lapse imaging of numerous processes such as heart beat, cell division and sub-cellular neuronal transport. We demonstrate the use of this device and show examples of different types of data collected from different model systems.

Micro fabricated fluidic devices provide an accessible micro-environment for in vivo studies on small organisms. Simple fabrication processes are ...

Microfluidic-based Electrotaxis for On-demand Quantitative Analysis of Caenorhabditis elegans' Locomotion

The nematode Caenorhabditis elegans is a versatile model organism for biomedical research because of its conservation of disease-related genes and pathways as well as its ease of cultivation. Several C. elegans disease models have been reported, including neurodegenerative disorders such as Parkinson's disease (PD), which involves the degeneration of dopaminergic (DA) neurons 1. Both transgenes and neurotoxic chemicals have been used to induce DA neurodegeneration and consequent movement defects in worms, allowing for investigations into the basis of neurodegeneration and screens for neuroprotective genes and compounds 2,3.
Screens in lower eukaryotes like C. elegans provide an efficient and economical means to identify compounds and genes affecting neuronal signaling. Conventional screens are typically performed manually and scored by visual inspection; consequently, they are time-consuming and prone to human errors. Additionally, most focus on cellular level analysis while ignoring locomotion, which is an especially important parameter for movement disorders.
We have developed a novel microfluidic screening system (Figure 1) that controls and quantifies C. elegans' locomotion using electric field stimuli inside microchannels. We have shown that a Direct Current (DC) field can robustly induce on-demand locomotion towards the cathode ("electrotaxis") 4. Reversing the field's polarity causes the worm to quickly reverse its direction as well. We have also shown that defects in dopaminergic and other sensory neurons alter the swimming response 5. Therefore, abnormalities in neuronal signaling can be determined using locomotion as a read-out. The movement response can be accurately quantified using a range of parameters such as swimming speed, body bending frequency and reversal time.
Our work has revealed that the electrotactic response varies with age. Specifically, young adults respond to a lower range of electric fields and move faster compared to larvae 4. These findings led us to design a new microfluidic device to passively sort worms by age and phenotype 6.
We have also tested the response of worms to pulsed DC and Alternating Current (AC) electric fields. Pulsed DC fields of various duty cycles effectively generated electrotaxis in both C. elegans and its cousin C. briggsae 7. In another experiment, symmetrical AC fields with frequencies ranging from 1 Hz to 3 KHz immobilized worms inside the channel 8.
Implementation of the electric field in a microfluidic environment enables rapid and automated execution of the electrotaxis assay. This approach promises to facilitate high-throughput genetic and chemical screens for factors affecting neuronal function and viability.

Microfluidic-based Electrotaxis for On-demand Quantitative Analysis of Caenorhabditis elegans' Locomotion

A semi-automated micro-electro-fluidic method to induce on-demand locomotion in Caenorhabditis elegans is described. This method is based on the neurophysiologic phenomenon of worms responding to mild electric fields (&#x201C;electrotaxis&#x201D;) inside microfluidic channels. Microfluidic electrotaxis serves as a rapid, sensitive, low-cost, and scalable technique to screen for factors affecting neuronal health.

The nematode Caenorhabditis elegans is a versatile model organism for biomedical research because of its conservation of disease-related genes and ...

Microfluidic Chip Fabrication and Method to Detect Influenza

Fast and effective diagnostics play an important role in controlling infectious disease by enabling effective patient management and treatment. Here, we present an integrated microfluidic thermoplastic chip with the ability to amplify influenza A virus in patient nasopharyngeal (NP) swabs and aspirates. Upon loading the patient sample, the microfluidic device sequentially carries out on-chip cell lysis, RNA purification and concentration steps within the solid phase extraction (SPE), reverse transcription (RT) and polymerase chain reaction (PCR) in RT-PCR chambers, respectively. End-point detection is performed using an off-chip Bioanalyzer (Agilent Technologies, Santa Clara, CA). For peripherals, we used a single syringe pump to drive reagent and samples, while two thin film heaters were used as the heat sources for the RT and PCR chambers. The chip is designed to be single layer and suitable for high throughput manufacturing to reduce the fabrication time and cost. The microfluidic chip provides a platform to analyze a wide variety of virus and bacteria, limited only by changes in reagent design needed to detect new pathogens of interest.

An integrated microfluidic thermoplastic chip has been developed for use as a molecular diagnostic. The chip performs nucleic acid extraction, reverse transcriptase, and PCR. Methods for fabricating and running the chip are described.

Fast  and effective diagnostics play an important role in controlling infectious  disease by enabling effective patient management and treatment. Here, we ...

Using Microfluidics Chips for Live Imaging and Study of Injury Responses in Drosophila Larvae

Live imaging is an important technique for studying cell biological processes, however this can be challenging in live animals. The translucent cuticle of the Drosophila larva makes it an attractive model organism for live imaging studies. However, an important challenge for live imaging techniques is to noninvasively immobilize and position an animal on the microscope. This protocol presents a simple and easy to use method for immobilizing and imaging Drosophila larvae on a polydimethylsiloxane (PDMS) microfluidic device, which we call the 'larva chip'. The larva chip is comprised of a snug-fitting PDMS microchamber that is attached to a thin glass coverslip, which, upon application of a vacuum via a syringe, immobilizes the animal and brings ventral structures such as the nerve cord, segmental nerves, and body wall muscles, within close proximity to the coverslip. This allows for high-resolution imaging, and importantly, avoids the use of anesthetics and chemicals, which facilitates the study of a broad range of physiological processes. Since larvae recover easily from the immobilization, they can be readily subjected to multiple imaging sessions. This allows for longitudinal studies over time courses ranging from hours to days. This protocol describes step-by-step how to prepare the chip and how to utilize the chip for live imaging of neuronal events in 3rd instar larvae. These events include the rapid transport of organelles in axons, calcium responses to injury, and time-lapse studies of the trafficking of photo-convertible proteins over long distances and time scales. Another application of the chip is to study regenerative and degenerative responses to axonal injury, so the second part of this protocol describes a new and simple procedure for injuring axons within peripheral nerves by a segmental nerve crush.

Using Microfluidics Chips for Live Imaging and Study of Injury Responses in Drosophila Larvae

Drosophila larvae are an attractive model system for live imaging due to their translucent cuticle and powerful genetics. This protocol describes how to utilize a single-layer PDMS device, called the &#39;larva chip&#39; for live imaging of cellular processes within neurons of 3rd instar Drosophila larvae.

Live imaging is an important technique for studying cell biological processes, however this can be challenging in live animals. The translucent cuticle of ...

Immobilization of Caenorhabditis elegans to Analyze Intracellular Transport in Neurons

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

Immobilization of Caenorhabditis elegans to Analyze Intracellular Transport in Neurons

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

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

Developmental Biology

A Microfluidic Platform for Longitudinal Imaging in Caenorhabditis elegans

In the last decade, microfluidic techniques have been applied to study small animals, including the nematode Caenorhabditis elegans, and have proved useful as a convenient live imaging platform providing capabilities for precise control of experimental conditions in real time. In this article, we demonstrate live imaging of individual worms employing WormSpa, a previously-published custom microfluidic device. In the device, multiple worms are individually confined to separate chambers, allowing multiplexed longitudinal surveillance of various biological processes. To illustrate the capability, we performed proof-of-principle experiments in which worms were infected in the device with pathogenic bacteria, and the dynamics of expression of immune response genes and egg laying were monitored continuously in individual animals. The simple design and operation of this device make it suitable for users with no previous experience with microfluidic-based experiments. We propose that this approach will be useful for many researchers interested in longitudinal observations of biological processes under well-defined conditions.

A Microfluidic Platform for Longitudinal Imaging in Caenorhabditis elegans

In this article, we demonstrate live imaging of individual worms employing a custom microfluidic device. In the device, multiple worms are individually confined to separate chambers, allowing multiplexed longitudinal surveillance of various biological processes.

In the last decade, microfluidic techniques have been applied to study small animals, including the nematode Caenorhabditis elegans, and have proved ...

Using a Microfluidics Device for Mechanical Stimulation and High Resolution Imaging of C. elegans

One central goal of mechanobiology is to understand the reciprocal effect of mechanical stress on proteins and cells. Despite its importance, the influence of mechanical stress on cellular function is still poorly understood. In part, this knowledge gap exists because few tools enable simultaneous deformation of tissue and cells, imaging of cellular activity in live animals, and efficient restriction of motility in otherwise highly mobile model organisms, such as the nematode Caenorhabditis elegans. The small size of C. elegans makes them an excellent match to microfluidics-based research devices, and solutions for immobilization have been presented using microfluidic devices. Although these devices allow for high-resolution imaging, the animal is fully encased in polydimethylsiloxane (PDMS) and glass, limiting physical access for delivery of mechanical force or electrophysiological recordings. Recently, we created a device that integrates pneumatic actuators with a trapping design that is compatible with high-resolution fluorescence microscopy. The actuation channel is separated from the worm-trapping channel by a thin PDMS diaphragm. This diaphragm is deflected into the side of a worm by applying pressure from an external source. The device can target individual mechanosensitive neurons. The activation of these neurons is imaged at high-resolution with genetically-encoded calcium indicators. This article presents the general method using C. elegans strains expressing calcium-sensitive activity indicator (GCaMP6s) in their touch receptor neurons (TRNs). The method, however, is not limited to TRNs nor to calcium sensors as a probe, but can be expanded to other mechanically-sensitive cells or sensors.

Using a Microfluidics Device for Mechanical Stimulation and High Resolution Imaging of C. elegans

New tools for mechanobiology research are needed to understand how mechanical stress activates biochemical pathways and elicits biological responses. Here, we showcase a new method for selective mechanical stimulation of immobilized animals with a microfluidic trap allowing high-resolution imaging of cellular responses.

One central goal of mechanobiology is to understand the reciprocal effect of mechanical stress on proteins and cells. Despite its importance, the ...

Neuroscience

A Simple Microfluidic Chip for Long-Term Growth and Imaging of Caenorhabditis elegans

Caenorhabditis elegans (C. elegans) have proved to be a valuable model system for studying developmental and cell biological processes. Understanding these biological processes often requires long-term and repeated imaging of the same animal. Long recovery times associated with conventional immobilization methods done on agar pads have detrimental effects on animal health making it inappropriate to repeatedly image the same animal over long periods of time. This paper describes a microfluidic chip design, fabrication method, on-chip C. elegans culturing protocol, and three examples of long-term imaging to study developmental processes in individual animals. The chip, fabricated with polydimethylsiloxane and bonded on a cover glass, immobilizes animals on a glass substrate using an elastomeric membrane that is deflected using nitrogen gas. Complete immobilization of C. elegans enables robust time-lapse imaging of cellular and sub-cellular events in an anesthetic-free manner. A channel geometry with a large cross-section allows the animal to move freely within two partially sealed isolation membranes permitting growth in the channel with a continuous food supply. Using this simple chip, imaging of developmental phenomena such as neuronal process growth, vulval development, and dendritic arborization in the PVD sensory neurons, as the animal grows inside the channel, can be performed. The long-term growth and imaging chip operates with a single pressure line, no external valves, inexpensive fluidic consumables, and utilizes standard worm handling protocols that can easily be adapted by other laboratories using C. elegans.

A Simple Microfluidic Chip for Long-Term Growth and Imaging of Caenorhabditis elegans

The protocol describes a simple microfluidic chip design and microfabrication methodology used to grow C. elegans in presence of a continuous food supply for up to 36 h. The growth and imaging device also enables intermittent long-term high-resolution imaging of cellular and sub-cellular processes during development for several days.

Caenorhabditis elegans (C. elegans) have proved to be a valuable model system for studying developmental and cell biological processes. Understanding ...

Constructing and Operating a Cooling Stage to Immobilize C. elegans En Masse

Assembly and Operation of a Cooling Stage to Immobilize C. elegans on Their Culture Plates

High-resolution in vivo microscopy approaches can reveal subtle information and fine details inside the model animal Caenorhabditis elegans (C. elegans), but require strong animal immobilization to prevent motion blur in the images. Unfortunately, most current immobilization techniques require substantial manual effort, rendering high-resolution imaging low-throughput. Immobilization of C. elegans is greatly simplified by using a cooling approach that can easily immobilize entire populations directly on their cultivation plates. The cooling stage can establish and maintain a wide range of temperatures with a uniform distribution on the cultivation plate. In this article, the whole process of building the cooling stage is documented. The aim is that a typical researcher can build an operational cooling stage in their laboratory following this protocol without difficulty. Utilization of the cooling stage following three protocols is shown, and each protocol has advantages for different experiments. Also shown is an example cooling profile of the stage as it approaches its final temperature and some helpful tips in using cooling immobilization.

Author Spotlight: Assembly and Operation of a Cooling Stage to Immobilize C. elegans on Their Culture Plates  

This paper describes protocols for constructing and operating a cooling stage to immobilize C. elegans on their original cultivation plates en masse.

High-resolution in vivo microscopy approaches can reveal subtle information and fine details inside the model animal Caenorhabditis elegans (C. elegans), ...

Immobilization of Live Caenorhabditis elegans Individuals Using an Ultra-thin Polydimethylsiloxane Microfluidic Chip with Water Retention

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