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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B., Krieg, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Using+a+microfluidics+device+for+mechanical+stimulation+and+high+resolution+imaging+of+C.+elegans.">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...

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 border="0" cellpadding="0" cellspacing="0" style="width:1151px;" width="1152">
	<tbody>
		<tr height="40">
			<td height="40" style="height:40px;width:223px;">C. elegans wild-type strain</td>
			<td style="width:125px;">Caenorhabditis Genetics Center (CGC) , Minnesota, USA</td>
			<td style="width:353px;">N2</td>
			<td style="width:449px;">Wild-type C. elegans strain generally used in this study</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:223px;">C. elegans unc-119(e2498) III mutant strain</td>
			<td style="width:125px;">Caenorhabditis Genetics Center (CGC) , Minnesota, USA</td>
			<td style="width:353px;">CB4845</td>
			<td style="width:449px;">C. elegans strain only employed as an example of mutants with abnormal body shape&#160;</td>
		</tr>
		<tr height="64">
			<td height="64" style="height:64px;width:223px;">C. elegans transgenic strain HBR4</td>
			<td style="width:125px;">Caenorhabditis Genetics Center (CGC) , Minnesota, USA</td>
			<td style="width:353px;">HBR4</td>
			<td style="width:449px;">The genotype of this transgenic C. elegans strain is HBR4:goeIs3[pmyo-3::GCamP3.35:: unc-54&#8211;3&#8217;utr, unc-119(+)]V. This strain was only employed for imaging observation.</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:223px;">E. coli strain</td>
			<td style="width:125px;">Caenorhabditis Genetics Center (CGC) , Minnesota, USA</td>
			<td style="width:353px;">OP50</td>
			<td style="width:449px;">E. coli strain used as food for C. elegans</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Worm Sheet IR (50/60)</td>
			<td style="width:125px;">Biocosm, Inc., Hyogo, Japan</td>
			<td>BCM17-0001</td>
			<td style="width:449px;">Microfluidic chip with 25 straight 50/60-&#181;m width channels used in all experiments and observation in this paper&#160;</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Worm Sheet 60</td>
			<td style="width:125px;">Biocosm, Inc., Hyogo, Japan</td>
			<td>BCM18-0001</td>
			<td style="width:449px;">Microfluidic chip with 20 straight 60 &#181;m-width channels. This is sitable for adults 3-5 days after hatching at 20&#176;C.&#160;</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Worm Sheet 50</td>
			<td style="width:125px;">Biocosm, Inc., Hyogo, Japan</td>
			<td>BCM18-0002</td>
			<td style="width:449px;">Microfluidic chip with 20 straight 50 &#181;m-width channels. This is sitable for youg adults ~3 days after hatching at 20&#176;C.&#160;</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">MICRO COVER GLASS</td>
			<td style="width:125px;">MATSUNAMI GLASS IND. LTD.</td>
			<td>C030401</td>
			<td style="width:449px;">Cover glass (thickness: 130-170 &#181;m) used in locomotion assays in Protocol 3</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Polystyrene Film</td>
			<td style="width:125px;">Biocosm, Inc., Hyogo, Japan</td>
			<td>BCM18-0001/ BCM18-0002</td>
			<td style="width:449px;">Bundled items of Worm Sheets. PS filim (thickness: ~130 &#181;m) used in locomotion assays in Protocol 3.</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Polyester Film Lumirror</td>
			<td style="width:125px;">TORAY INDUSTRIES, INC., Tokyo, Japan</td>
			<td>Lumirror T60 (t 125 &#181;m)</td>
			<td style="width:449px;">PET filim (thickness: 125 &#181;m) used in locomotion assays in Protocol 3</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">IWAKI 60 mm/non-treated dish</td>
			<td style="width:125px;">AGC Techno Glass Co., Ltd., Shizuoka, Japan).</td>
			<td>1010-060</td>
			<td style="width:449px;">Non-treated dish used in incuvation of C. elegans in Protocol 1</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">IWAKI 35 mm/non-treated dish</td>
			<td style="width:125px;">AGC Techno Glass Co., Ltd., Shizuoka, Japan).</td>
			<td>1010-035</td>
			<td style="width:449px;">Non-treated dish used in locomotion assays in Protocol 3</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Milli-Q</td>
			<td style="width:125px;">Merck, France</td>
			<td></td>
			<td style="width:449px;">Ultrapure water</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Kimwipe S-200</td>
			<td style="width:125px;">Nippon Paper Crecia Co., Ltd., Tokyo, Japan</td>
			<td>62020</td>
			<td style="width:449px;">120 mm x 215 mm; 200 sheets/ box</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">WormStuff Worm Pick</td>
			<td style="width:125px;">Genesee Scientific Corporation, CA, USA)</td>
			<td>59-AWP</td>
			<td style="width:449px;">Platina picker specilized for picking up C. elegans</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:223px;">Research Stereo Microscope System</td>
			<td style="width:125px;">OLYMPUS CORPORATION, Tokyo, Japan</td>
			<td>SZX16</td>
			<td style="width:449px;">Micriscope used in all experiments and observation in this paper</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:223px;">Motorized Focus Stand for SZX16</td>
			<td style="width:125px;">OLYMPUS CORPORATION, Tokyo, Japan</td>
			<td style="width:353px;">SZX2-ILLB</td>
			<td style="width:449px;">This was used for bright field observation in Protocol 3-8.</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Objective Lens (&#215;1)</td>
			<td style="width:125px;">OLYMPUS CORPORATION, Tokyo, Japan</td>
			<td>SDFPLAPO1&#215;PF</td>
			<td style="width:449px;">NA: 0.15; W.D.: 60 mm. This lends was used for bright field observation in Protocol 3-8.</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Objective Lens (&#215;2)</td>
			<td style="width:125px;">OLYMPUS CORPORATION, Tokyo, Japan</td>
			<td>SDFPLAPO2XPFC</td>
			<td style="width:449px;">NA: 0.3; W.D.: 20 mm. This&#160; lends was used for imaging observations.</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: ~...

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 ...

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7.7K Views.  National Institutes for Quantum and Radiological Science and Technology (QST-Takasaki). 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.

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 ...

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&#039; 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 ...

A Microfluidic Chip for the Versatile Chemical Analysis of Single Cells

We present a microfluidic device that enables the quantitative determination of intracellular biomolecules in multiple single cells in parallel. For this purpose, the cells are passively trapped in the middle of a microchamber. Upon activation of the control layer, the cell is isolated from the surrounding volume in a small chamber. The surrounding volume can then be exchanged without affecting the isolated cell. However, upon short opening and closing of the chamber, the solution in the chamber can be replaced within a few hundred milliseconds. Due to the reversibility of the chambers, the cells can be exposed to different solutions sequentially in a highly controllable fashion, e.g. for incubation, washing, and finally, cell lysis. The tightly sealed microchambers enable the retention of the lysate, minimize and control the dilution after cell lysis. Since lysis and analysis occur at the same location, high sensitivity is retained because no further dilution or loss of the analytes occurs during transport. The microchamber design therefore enables the reliable and reproducible analysis of very small copy numbers of intracellular molecules (attomoles, zeptomoles) released from individual cells. Furthermore, many microchambers can be arranged in an array format, allowing the analysis of many cells at once, given that suitable optical instruments are used for monitoring. We have already used the platform for proof-of-concept studies to analyze intracellular proteins, enzymes, cofactors and second messengers in either relative or absolute quantifiable manner.

In this article we present a microfluidic chip for single cell analysis. It allows the quantification of intracellular proteins, enzymes, cofactors, and second messengers by means of fluorescent assays or immunoassays.&#160;

We present a microfluidic device that enables the quantitative determination of intracellular biomolecules in multiple single cells in parallel. For this ...

A Microfluidic Chip for ICPMS Sample Introduction

This protocol discusses the fabrication and usage of a disposable low cost microfluidic chip as sample introduction system for inductively coupled plasma mass spectrometry (ICPMS). The chip produces monodisperse aqueous sample droplets in perfluorohexane (PFH). Size and frequency of the aqueous droplets can be varied in the range of 40 to 60 &#181;m and from 90 to 7,000&#160;Hz, respectively. The droplets are ejected from the chip with a second flow of PFH and remain intact during the ejection. A custom-built desolvation system removes the PFH and transports the droplets into the ICPMS. Here, very stable signals with a narrow intensity distribution can be measured, showing the monodispersity of the droplets. We show that the introduction system can be used to quantitatively determine iron in single bovine red blood cells. In the future, the capabilities of the introduction device can easily be extended by the integration of additional microfluidic modules.

We present a discrete droplet sample introduction system for inductively coupled plasma mass spectrometry (ICPMS). It is based on a cheap and disposable microfluidic chip that generates highly monodisperse droplets in a size range of 40&#8722;60 &#181;m at frequencies from 90 to 7,000 Hz.

This protocol discusses the fabrication and usage of a disposable low cost microfluidic chip as sample introduction system for inductively coupled plasma ...

Electrotaxis Studies of Lung Cancer Cells using a Multichannel Dual-electric-field Microfluidic Chip

The behavior of directional cell migration under a direct current electric-field (dcEF) is referred to as electrotaxis. The significant role of physiological dcEF in guiding cell movement during embryo development, cell differentiation, and wound healing has been demonstrated in many studies. By applying microfluidic chips to an electrotaxis assay, the investigation process is shortened and experimental errors are minimized. In recent years, microfluidic devices made of polymeric substances (e.g., polymethylmethacrylate, PMMA, or acrylic) or polydimethylsiloxane (PDMS) have been widely used in studying the responses of cells to electrical stimulation. However, unlike the numerous steps required to fabricate a PDMS device, the simple and rapid construction of the acrylic micro&#64258;uidic chip makes it suitable for both device prototyping and production. Yet none of the reported devices facilitate the efficient study of the simultaneous chemical and dcEF effects on cells. In this report, we describe our design and fabrication of an acrylic-based multichannel dual-electric-field (MDF) chip to investigate the concurrent effect of chemical and electrical stimulation on lung cancer cells. The MDF chip provides eight combinations of electrical/chemical stimulations in a single test. The chip not only greatly shortens the required experimental time but also increases accuracy in electrotaxis studies.

Many microfluidic devices have been developed for use in the study of electrotaxis. Yet, none of these chips allows the efficient study of the simultaneous chemical and electric-field (EF) effects on cells. We developed a polymethylmethacrylate-based device that offers better-controlled coexisting EF and chemical stimulation for use in electrotaxis research.

The behavior of directional cell migration under a direct current electric-field (dcEF) is referred to as electrotaxis. The significant role of ...

Three-Dimensionally Printed Microfluidic Cross-flow System for Ultrafiltration/Nanofiltration Membrane Performance Testing

Minimization and management of membrane fouling is a formidable challenge in diverse industrial processes and other practices that utilize membrane technology. Understanding the fouling process could lead to optimization and higher efficiency of membrane based filtration. Here we show the design and fabrication of an automated three-dimensionally (3-D) printed microfluidic cross-flow filtration system that can test up to 4 membranes in parallel. The microfluidic cells were printed using multi-material photopolymer 3-D printing technology, which used a transparent hard polymer for the microfluidic cell body and incorporated a thin rubber-like polymer layer, which prevents leakages during operation. The performance of ultrafiltration (UF), and nanofiltration (NF) membranes were tested and membrane fouling could be observed with a model foulant bovine serum albumin (BSA). Feed solutions containing BSA showed flux decline of the membrane. This protocol may be extended to measure fouling or biofouling with many other organic, inorganic or microbial containing solutions. The microfluidic design is especially advantageous for testing materials that are costly or only available in small quantities, for example polysaccharides, proteins, or lipids due to the small surface area of the membrane being tested. This modular system may also be easily expanded for high throughput testing of membranes.&#160;

Design and fabrication of a three-dimensionally (3-D) printed microfluidic cross-flow filtration system is demonstrated. The system is used to test performance and observe fouling of ultrafiltration and nanofiltration (thin film composite) membranes.

Minimization and management of membrane fouling is a formidable challenge in diverse industrial processes and other practices that utilize membrane ...

Automated Lipid Bilayer Membrane Formation Using a Polydimethylsiloxane Thin Film

An artificial lipid bilayer, or black lipid membrane (BLM), is a powerful tool for studying ion channels and protein interactions, as well as for biosensor applications. However, conventional BLM formation techniques have several drawbacks and they often require specific expertise and laborious processes. In particular, conventional BLMs suffer from low formation success rates and inconsistent membrane formation time. Here, we demonstrate a storable and transportable BLM formation system with controlled thinning-out time and enhanced BLM formation rate by replacing conventionally used films (polytetrafluoroethylene, polyoxymethylene, polystyrene) to polydimethylsiloxane (PDMS). In this experiment, a porous-structured polymer such as PDMS thin film is used. In addition, as opposed to conventionally used solvents with low viscosity, the use of squalene permitted a controlled thinning-out time via slow solvent absorption by PDMS, prolonging membrane lifetime. In addition, by using a mixture of squalene and hexadecane, the freezing point of the lipid solution was increased (~16 &#176;C), in addition, membrane precursors were produced that can be indefinitely stored and readily transported. These membrane precursors have reduced BLM formation time of &#60; 1 hr and achieved a BLM formation rate of ~80%. Moreover, ion channel experiments with gramicidin A demonstrated the feasibility of the membrane system.

We demonstrate a storable, transportable lipid bilayer formation system. A lipid bilayer membrane can be formed within 1 hr with over 80% success rate when a frozen membrane precursor is brought to ambient temperature. This system will reduce laborious processes and expertise associated with ion channels.

An artificial lipid bilayer, or black lipid membrane (BLM), is a powerful tool for studying ion channels and protein interactions, as well as for ...

Rapid Subtractive Patterning of Live Cell Layers with a Microfluidic Probe

The microfluidic probe (MFP) facilitates performing local chemistry on biological substrates by confining nanoliter volumes of liquids. Using one particular implementation of the MFP, the hierarchical hydrodynamic flow confinement (hHFC), multiple liquids are simultaneously brought in contact with a substrate. Local chemical action and liquid shaping using the hHFC, is exploited to create cell patterns by locally lysing and removing cells. By utilizing the scanning ability of the MFP, user-defined patterns of cell monolayers are created. This protocol enables rapid, real-time and spatially controlled cell patterning, which can allow selective cell-cell and cell-matrix interaction studies.

We present a protocol to perform subtractive patterning of live cell monolayers on a surface. This is achieved by local and selective lysis of adherent cells using a microfluidic probe (MFP). The cell lysate retrieved from local regions can be used for downstream analysis, enabling molecular profiling studies.

The microfluidic probe (MFP) facilitates performing local chemistry on biological substrates by confining nanoliter volumes of liquids. Using one ...

Polydimethylsiloxane-polycarbonate Microfluidic Devices for Cell Migration Studies Under Perpendicular Chemical and Oxygen Gradients

This paper reports a microfluidic device made of polydimethylsiloxane (PDMS) with an embedded polycarbonate (PC) thin film to study cell migration under combinations of chemical and oxygen gradients. Both chemical and oxygen gradients can greatly affect cell migration in vivo; however, due to technical limitations, very little research has been performed to investigate their effects in vitro. The device developed in this research takes advantage of a series of serpentine-shaped channels to generate the desired chemical gradients and exploits a spatially confined chemical reaction method for oxygen gradient generation. The directions of the chemical and oxygen gradients are perpendicular to each other to enable straightforward migration result interpretation. In order to efficiently generate the oxygen gradients with minimal chemical consumption, the embedded PC thin film is utilized as a gas diffusion barrier. The developed microfluidic device can be actuated by syringe pumps and placed into a conventional cell incubator during cell migration experiments to allow for setup simplification and optimized cell culture conditions. In cell experiments, we used the device to study migrations of adenocarcinomic human alveolar basal epithelial cells, A549, under combinations of chemokine (stromal cell-derived factor, SDF-1&#945;) and oxygen gradients. The experimental results show that the device can stably generate perpendicular chemokine and oxygen gradients and is compatible with cells. The migration study results indicate that oxygen gradients may play an essential role in guiding cell migration, and cellular behavior under combinations of gradients cannot be predicted from those under single gradients. The device provides a powerful and practical tool for researchers to study interactions between chemical and oxygen gradients in cell culture, which can promote better cell migration studies in more in vivo-like microenvironments.

The control of chemical and oxygen gradients is essential for cell cultures. This paper reports a polydimethylsiloxane-polycarbonate (PDMS-PC) microfluidic device capable of reliably generating combinations of chemical and oxygen gradients for cell migration studies, which can be practically utilized in biological labs without sophisticated instrumentation.

This paper reports a microfluidic device made of polydimethylsiloxane (PDMS) with an embedded polycarbonate (PC) thin film to study cell migration under ...