This research was supported by the National Science Foundation through grants PHY-1205494 and MCB-1413134 (EL) and by the National Research Foundation of Korea grant 2017R1D1A1B03035671 (KSL).

The protocol below uses WormSpa5, a previously-described microfluidic device for longitudinal imaging of worms. Fabrication of WormSpa (starting with CAD files that can be obtained from the authors upon request) is straightforward but requires some expertise. In most cases, fabrication can readily be done by a core facility or by a commercial company that provides such services. When fabricating the device, make sure to specify that the height of the features is 50 &#181;m.
1. Experimental Setup
<ol>
	<li>Mount the microfluidic device on an inverted fluorescence microscope with a motorized stage.</li>
	<li>Place an orbital shaker close to the microscope, but make sure the vibration from the shaker does not directly affect the microscope. For example, put the shaker on a shelf fixed on a wall that makes no contact with the optical table.</li>
	<li>Place a syringe pump on the orbital shaker. Tape the pump to the shaker so that it does not jiggle while shaking.</li>
</ol>
2. Preparation of the Microfluidic Environment
NOTE: During the experiment, four syringes are connected to the microfluidic device via syringe tubes, as outlined in Figure 1. This step describes the preparation of these syringes. Unless mentioned otherwise, keep the syringe tubes as short as possible without making them taut.
<ol>
	<li>Prepare an outlet syringe and a syringe tube. This syringe (S1 in Figure 1) will later be connected to the outlet of the device through the syringe tube, and used for temporary storage of fluid coming out of the device.
	<ol>
		<li>Make an adapter needle by removing plastic parts of a 20 G x 1/2 inch syringe tip, keeping only the needle (the metal part). Do so by holding the plastic and metal parts with pliers and pulling them apart. Remaining plastic residue on the needle can be burnt off with a Bunsen burner.</li>
		<li>Bend the needle according to the geometry of the experimental setup while holding its two ends with pliers. In the design described here, bending the needle in its middle to an angle of ~110&#176; is most convenient.</li>
		<li>Plug the adapter needle half way into the tube. The other half will later be plugged into the device. The syringe tube should be compatible with a 20 G syringe tip without a leak (for example, tubing with inner diameter of 0.86 mm and outer diameter of 1.32 mm). The recommended tube length is ~50 cm.</li>
		<li>Connect a 10 mL luer-lock syringe to the other (open) end of the syringe tube via a 20 G x 1/2 inch syringe tip.</li>
	</ol>
	</li>
	<li>Prepare buffer-inlet syringes and syringe tubes. One of these syringes (S2 in Figure 1) is used as a reservoir for the fluid that goes into the device and to control the injection flow. The other syringe (S3 in Figure 1) is required for buffer exchange, as explained in Step 3.2.
	<ol>
		<li>Connect a 10 mL luer-lock syringe (S2) directly to a 3-way valve and connect another syringe (S3) to the same valve via a syringe tube (Figure 1): connect S3 to the tube via a syringe tip, and connect the tube to the valve via another syringe tip. 
		NOTE: The order in which these materials are connected does not matter.</li>
		<li>Connect a syringe tube to the remaining port of the 3-way valve. Prepare another adapter needle as in Step 2.1.2, and plug the needle half-way into the other (open) end of the syringe tube.</li>
		<li>Set the valve such that the syringe tube in Step 2.2.2 and S2 are connected while S3 is closed (Figure 1).</li>
	</ol>
	</li>
	<li>Prepare a worm-inlet syringe and a syringe tube. This syringe (S4 in Figure 1) is used for loading worms into the device.
	<ol>
		<li>Connect a long syringe tube to a 10 mL luer-lock syringe (S4). Determine the length of this tube from the volume of the fluid that will be used for loading; for example, 100 cm of tubing supports over 2 mL of liquid (also, see step 4.2).</li>
		<li>Prepare another adapter needle as in Step 2.1.2, and plug one half of the needle to the other (open) end of the syringe tube.</li>
	</ol>
	</li>
	<li>Rinse all the syringes and tubing with S-medium12 twice. Fill the syringes and the tubes with S-medium, taking care to remove all air bubbles.</li>
	<li>Connect the outlet syringe tube to the device outlet.
	<ol>
		<li>Plug in the adapter needle at the end of the syringe tube to the outlet port.</li>
		<li>Inject a small volume of S-medium through the outlet to fill up the device, until a little S-medium comes out of both the buffer inlet and the worm inlet.</li>
	</ol>
	</li>
	<li>Connect the buffer-inlet syringe tube to the buffer-inlet port of the device, while plugging the worm-inlet with a pin (stainless steel dowel pin with a 1/32-inch diameter).</li>
	<li>Inject a constant flow of buffer into the device. Load the buffer-inlet syringe S2 onto a syringe pump13, and set the pump such that the medium in S2 is continuously injected into the device at a flow rate of 3 &#181;L/min.</li>
</ol>
3. Loading Bacteria into the Microfluidic Device
<ol>
	<li>Prepare Escherichia coli OP50 suspended in S-medium12.
	<ol>
		<li>Following a standard protocol, inoculate a 250 mL flask with 50 mL of LB and a single colony, and incubate overnight at 37 &#176;C.</li>
		<li>Centrifuge the overnight culture at 4000 rpm for 10 minutes and resuspend the pellet in 30 mL of S-medium.</li>
		<li>Filter the suspension through a 5 &#181;m syringe filter. Measure the bacterial concentration by its optical density at 600 nm (OD600), and adjust the concentration to an OD600 of 3. This high density is required to keep the worms well-fed.</li>
	</ol>
	</li>
	<li>Exchange the buffer in the device with the OP50 suspension.
	<ol>
		<li>Prepare a clean syringe (S5) filled with the OP50 suspension. Hold the syringe vertically and tap it several times to collect all the air bubbles at the top.</li>
		<li>Turn the 3-way valve of the buffer-inlet syringes to close the connection to the device, and connect the two syringes, S2 and S3. Disengage S2 from the syringe pump.</li>
		<li>Disconnect S2 from the valve and connect S5 to the valve. Push out all the air collected at the top of S5 to S3 through the valve until a little bit of the OP50 buffer goes into S3. In doing so, make sure that no air bubble remains in S5 or the valve.</li>
		<li>Turn the 3-way valve back to the original position to close the connection between S3 and S5 and connect S5 to the device. Load S5 onto the syringe pump. Turn on the orbital shaker that holds the pump. Set the shaking speed to approximately 200 rpm.</li>
		<li>Set the flow rate to be 100 &#181;L/min. The time it takes for the new buffer to arrive to the worms in the device depends on the length of the buffer-inlet syringe tube (around 5 minutes with the tubing described above). A higher flow rate can be employed if a quicker buffer exchange is required.</li>
		<li>Once the OP50 buffer spreads throughout the device, set the flow rate back to 3 &#181;L/min.</li>
	</ol>
	</li>
</ol>
4. Loading Worms into the Microfluidic Device
<ol>
	<li>Prepare a small low binding microcentrifuge tube (650 &#181;L) and fill it with 100 &#181;L of OP50 suspension. Transfer around 20 - 30 age-synchronized young adult worms (46 hours post L1 larval arrest at 25 &#176;C) from an NGM (nematode growth medium) agar plate to the tube by picking them one by one with a worm pick. Age-synchronization can be done following a standard protocol12.</li>
	<li>Fill the worm-inlet syringe and syringe tube (S4 in Figure 1) with the OP50 suspension. Inject ~500 &#181;L of the fluid into the microcentrifuge tube with worms. Draw the suspension with worms into the worm-inlet syringe tube. Make sure that the syringe tube is long enough (&#62;1 m) and that drawn worms stay in the syringe tube and do not make all the way to the syringe S4.</li>
	<li>Connect S4 to the worm-inlet. Inject all the worms in the worm-inlet syringe tube into the microfluidic device. Disconnect S4 and the syringe tube. Plug the worm-inlet with a pin.</li>
	<li>Disengage the buffer-inlet syringe S2 from the mechanical pump ( Figure 1) and manually control S1 and S2 to push all the worms into separate channels. Pressing S2 will move worms into the channels, while pressing S1 will move them in the reverse direction. Once a channel is loaded, the worm in each channel will block the entry of other worms.</li>
	<li>Let worms adjust to the new environment for 2 - 3 hours.</li>
</ol>
5. Setting Up an Imaging Protocol
<ol>
	<li>Find all worms that are securely located in the device, and set an imaging position for each of them in the image acquisition software that controls your microscope. Note that in some cases (e.g. when higher magnification is desired, or when using cameras with a smaller CCD sensor) multiple imaging positions are required for each channel. 
	NOTE: While this can be done manually, some acquisition software packages can load a text file that lists the positions where images should be taken. Since these positions are ordered regularly in WormSpa, a user can write a small script (e.g., in Python or commercial software) that automatically creates such a list. The precise format of this script would depend on the file format expected by the acquisition software (see an example in Supplementary Material 1).</li>
	<li>Set the required frequency of time-lapse imaging. For example, imaging of immune gene response to infection is done at a frequency of 10 minutes per frame. Make sure that all necessary channels (e.g. bright-field and the GFP fluorescence) are acquired at every time point.</li>
</ol>
6. Host Response to Pseudomonas Infection
Note: This step is specific for studying host-pathogen interactions. Alternatively, one can prepare a buffer that contains other environmental cues of interest (biotic and abiotic stressors, drugs, signaling molecules, etc.).
<ol>
	<li>Prepare Pseudomonas aeruginosa PA14 suspended in SK-medium14.
	<ol>
		<li>Inoculate a 250 mL flask with 50 mL of LB and a single colony, and incubate overnight at 37 &#176;C.</li>
		<li>Inoculate another 250 mL flask with 50 mL of LB and an aliquot of the culture, and incubate overnight at 37 &#176;C.</li>
		<li>Centrifuge the overnight culture at 4000 rpm for 10 minutes and resuspend the pellet in 10 mL of SK-medium. Measure the bacterial concentration and adjust the concentration to OD600 = 4.</li>
		<li>Transfer the suspension to a clean flask, and incubate at 37 &#176;C for 24 hours and at 25 &#176;C for another 24 hours while shaking. This step mimics the plate-incubation step in the standard killing assay14.</li>
		<li>Filter the suspension through a 5 &#181;m syringe filter. This is to prevent bacterial aggregates from clogging the device.</li>
	</ol>
	</li>
	<li>Replace the OP50 suspension in the device with the PA14 suspension, following the same procedure as in Step 3.2. Keep imaging for 10 hours.</li>
</ol>

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The authors have nothing to disclose.

Microfluidic tools provide multiple benefits in studying worms. Imaging in a PDMS device offers higher imaging quality as compared with a standard NGM agar plate. Multiple images can be taken from a single worm, in contrast with traditional methods in which animals are picked from the plate and mounted on a microscope slide for imaging. In addition, the microenvironment in which worms reside can be kept constant or modulated as desired, permitting precise mapping between the composition of the environment and the respons...

Changes in environmental conditions may lead to activation of genetic programs accompanied by induction and repression of the expression of specific genes1,2. These kinetic changes may be variable among tissues in the same animals and between different animals. Studies of such genetic programs therefore call for methods that allow longitudinal imaging of individual animals and provide precise dynamical control of environmental conditions.
In recent years, microfabricated fluidic devices have been used to study many aspects of response and behavior in small animals, including worms, flies, water bears and more3,4,5,6,7. Applications include, for example, deep phenotyping, optogenetic recording of neuronal activity in response to chemical stimuli, and tracking of motor behaviors such as locomotion and pumping8,9,10,11.
Microfluidic-based approaches hold many properties that could benefit long-term longitudinal imaging of response to environmental cues, including precise dynamical control of the local microenvironment, flexible design that allows maintenance of individual animals in separate quarters, and favorable attributes for imaging. However, maintaining animals in a microfluidic chamber for a long time with minimal adverse impact on their well-beings is a challenge, which requires particular care in the design of the microfluidic device as well as in the execution of the experiment.
Here we demonstrate the use of WormSpa, a microfluidic device for longitudinal imaging of Caenorhabditis elegans.5 Individual worms are confined in chambers. A constant low flow of liquid and bacterial suspension guarantees that worms are well-fed and sufficiently active to maintain good health and alleviate stress, and the structure of the chambers allows worms to lay eggs. The simplicity of the design and operation of WormSpa should allow researchers with no previous experience in microfluidics to incorporate this device into their own research plans.

<table border="0" cellpadding="0" cellspacing="0" style="width:1012px;" width="1011">
	<colgroup>
		<col />
		<col />
		<col />
		<col />
	</colgroup>
	<tbody>
		<tr height="40">
			<td height="40" style="height:40px;width:216px;">WormSpa</td>
			<td style="width:123px;">N/A</td>
			<td style="width:344px;">N/A</td>
			<td style="width:329px;">The CAD file for WormSpa is available from the Levine lab.</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Compound Microscope</td>
			<td>Zeiss</td>
			<td>AxioObserver Z1</td>
			<td>An inverted fluorescence microscope with a motorized stage</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Syringe Pump</td>
			<td>New Era Pump Systems</td>
			<td>NE-501</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Tubing</td>
			<td>SCI Scientific Commodities Inc.</td>
			<td>BB31695-PE/5</td>
			<td>0.034&#8221; (0.86 mm) I.D. x 0.052&#8221; (1.32 mm) O.D</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Syringe Tip</td>
			<td>CMLsupply</td>
			<td>901-20-050</td>
			<td>20 Gauge x 1/2&#8221; blunt tip stainless steel canula</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Syringe Filter</td>
			<td>PALL</td>
			<td>4650</td>
			<td>Acrodisc 32 mm Syringe Filter with 5 um Supor Membrane</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Syringe</td>
			<td>Qosina</td>
			<td>C3307</td>
			<td>10 mL Male Luer Lock Syringe</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">3 Way Valve</td>
			<td>ColeParmer</td>
			<td>FF-30600-23</td>
			<td>Large-bore 3-way, male-lock, stopcocks, 10/pack, Non-sterile</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Dowel Pin</td>
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			<td height="40" style="height:40px;">Low Binding Microcentrifuge Tube</td>
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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.

Age-synchronized young adult worms (46 hours post L1 larval arrest at 25 &#176;C)12 were loaded in the device, as described in the protocol. The worms were individually located in separate channels, enabling longitudinal measurement of animals&#39; response to the pathogen. When the experiment is successful, most worms remain in their channels for the duration of the experiment. In this case, images of individual worms are taken simply by placing their channel in t...

Watch this Scientific Journal Video about A Microfluidic Platform for Longitudinal Imaging in Caenorhabditis elegans at JoVE.com

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.

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 ...

a-microfluidic-platform-for-longitudinal-imaging-caenorhabditis

Research

JoVE Journal

Bioengineering

6.4K Views.  Kongju National University. 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.

Video: A Microfluidic Platform for Longitudinal Imaging in Caenorhabditis elegans

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 ...

High-throughput Protein Expression Generator Using a Microfluidic Platform

Rapidly increasing fields, such as systems biology, require the development and implementation of new technologies, enabling high-throughput and high-fidelity measurements of large systems. Microfluidics promises to fulfill many of these requirements, such as performing high-throughput screening experiments on-chip, encompassing biochemical, biophysical, and cell-based assays1. Since the early days of microfluidics devices, this field has drastically evolved, leading to the development of microfluidic large-scale integration2,3. This technology allows for the integration of thousands of micromechanical valves on a single device with a postage-sized footprint (Figure 1). We have developed a high-throughput microfluidic platform for generating in vitro expression of protein arrays (Figure 2) named PING (Protein Interaction Network Generator). These arrays can serve as a template for many experiments such as protein-protein 4, protein-RNA5 or protein-DNA6 interactions.
The device consist of thousands of reaction chambers, which are individually programmed using a microarrayer. Aligning of these printed microarrays to microfluidics devices programs each chamber with a single spot eliminating potential contamination or cross-reactivity Moreover, generating microarrays using standard microarray spotting techniques is also very modular, allowing for the arraying of proteins7, DNA8, small molecules, and even colloidal suspensions. The potential impact of microfluidics on biological sciences is significant. A number of microfluidics based assays have already provided novel insights into the structure and function of biological systems, and the field of microfluidics will continue to impact biology.

We present a microfluidic approach for the expression of protein arrays. The device consists of thousands of reaction chambers controlled by micro-mechanical valves. The microfluidic device is mated to a microarray-printed gene library. These genes are then transcribed and translated on-chip, resulting in a protein array ready for experimental use.

Rapidly increasing fields, such as systems biology, require the development and implementation of new technologies, enabling high-throughput and ...

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 ...

Cell Squeezing as a Robust, Microfluidic Intracellular Delivery Platform

Rapid mechanical deformation of cells has emerged as a promising, vector-free method for intracellular delivery of macromolecules and nanomaterials. This technology has shown potential in addressing previously challenging applications; including, delivery to primary immune cells, cell reprogramming, carbon nanotube, and quantum dot delivery. This vector-free microfluidic platform relies on mechanical disruption of the cell membrane to facilitate cytosolic delivery of the target material. Herein, we describe the detailed method of use for these microfluidic devices including, device assembly, cell preparation, and system operation. This delivery approach requires a brief optimization of device type and operating conditions for previously unreported applications. The provided instructions are generalizable to most cell types and delivery materials as this system does not require specialized buffers or chemical modification/conjugation steps. This work also provides recommendations on how to improve device performance and trouble-shoot potential issues related to clogging, low delivery efficiencies, and cell viability.

Rapid mechanical deformation of cells has emerged as a promising, vector-free method for intracellular delivery of macromolecules and nanomaterials. This protocol provides detailed steps on how to use the system for a broad range of applications.

Rapid mechanical deformation of cells has emerged as a promising, vector-free method for intracellular delivery of macromolecules and nanomaterials. This ...

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 ...

A Microfluidic Platform for High-throughput Single-cell Isolation and Culture

Studying the heterogeneity of single cells is crucial for many biological questions, but is technically difficult. Thus, there is a need for a simple, yet high-throughput, method to perform single-cell culture experiments. Here, we report a microfluidic chip-based strategy for high-efficiency single-cell isolation (~77%) and demonstrate its capability of performing long-term single-cell culture (up to 7 d) and cellular heterogeneity analysis using clonogenic assay. These applications were demonstrated with KT98 mouse neural stem cells, and A549 and MDA-MB-435 human cancer cells. High single-cell isolation efficiency and long-term culture capability are achieved by using different sizes of microwells on the top and bottom of the microfluidic channel. The small microwell array is designed for precisely isolating single-cells, and the large microwell array is used for single-cell clonal culture in the microfluidic chip. This microfluidic platform constitutes an attractive approach for single-cell culture applications, due to its flexibility of adjustable cell culture spaces for different culture strategies, without decreasing isolation efficiency.

Here, we present a protocol for isolating and culturing single cells with a microfluidic platform, which utilizes a new microwell design concept to allow for high-efficiency single cell isolation and long-term clonal culture.

Studying the heterogeneity of single cells is crucial for many biological questions, but is technically difficult. Thus, there is a need for a simple, yet ...

Microfluidic Platform with Multiplexed Electronic Detection for Spatial Tracking of Particles

Microfluidic processing of biological samples typically involves differential manipulations of suspended particles under various force fields in order to spatially fractionate the sample based on a biological property of interest. For the resultant spatial distribution to be used as the assay readout, microfluidic devices are often subjected to microscopic analysis requiring complex instrumentation with higher cost and reduced portability. To address this limitation, we have developed an integrated electronic sensing technology for multiplexed detection of particles at different locations on a microfluidic chip. Our technology, called Microfluidic CODES, combines Resistive Pulse Sensing with Code Division Multiple Access to compress 2D spatial information into a 1D electrical signal. In this paper, we present a practical demonstration of the Microfluidic CODES technology to detect and size cultured cancer cells distributed over multiple microfluidic channels. As validated by the high-speed microscopy, our technology can accurately analyze dense cell populations all electronically without the need for an external instrument. As such, the Microfluidic CODES can potentially enable low-cost integrated lab-on-a-chip devices that are well suited for the point-of-care testing of biological samples.

We demonstrate a microfluidic platform with an integrated surface electrode network that combines resistive pulse sensing (RPS) with code division multiple access (CDMA), to multiplex detection and sizing of particles in multiple microfluidic channels.

Microfluidic processing of biological samples typically involves differential manipulations of suspended particles under various force fields in order to ...

Imaging the Root Hair Morphology of Arabidopsis Seedlings in a Two-layer Microfluidic Platform

Root hairs increase root surface area for better water uptake and nutrient absorption by the plant. Because they are small in size and often obscured by their natural environment, root hair morphology and function are difficult to study and often excluded from plant research. In recent years, microfluidic platforms have offered a way to visualize root systems at high resolution without disturbing the roots during transfer to an imaging system. The microfluidic platform presented here builds on previous plant-on-a-chip research by incorporating a two-layer device to confine the Arabidopsis thaliana main root to the same optical plane as the root hairs. This design enables the quantification of root hairs on a cellular and organelle level and also prevents z-axis drifting during the addition of experimental treatments. We describe how to store the devices in a contained and hydrated environment, without the need for fluidic pumps, while maintaining a gnotobiotic environment for the seedling. After the optical imaging experiment, the device may be disassembled and used as a substrate for atomic force or scanning electron microscopy while keeping fine root structures intact.

Imaging the Root Hair Morphology of Arabidopsis Seedlings in a Two-layer Microfluidic Platform

This article demonstrates how to culture Arabidopsis thaliana seedlings in a two-layer microfluidic platform that confines the main root and root hairs to a single optical plane. This platform can be used for real-time optical imaging of fine root morphology as well as for high-resolution imaging by other means.

Root hairs increase root surface area for better water uptake and nutrient absorption by the plant. Because they are small in size and often obscured by ...

An Ultrahigh-throughput Microfluidic Platform for Single-cell Genome Sequencing

Sequencing technologies have undergone a paradigm shift from bulk to single-cell resolution in response to an evolving understanding of the role of cellular heterogeneity in biological systems. However, single-cell sequencing of large populations has been hampered by limitations in processing genomes for sequencing. In this paper, we describe a method for single-cell genome sequencing (SiC-seq) which uses droplet microfluidics to isolate, amplify, and barcode the genomes of single cells. Cell encapsulation in microgels allows the compartmentalized purification and tagmentation of DNA, while a microfluidic merger efficiently pairs each genome with a unique single-cell oligonucleotide barcode, allowing &gt;50,000 single cells to be sequenced per run. The sequencing data is demultiplexed by barcode, generating groups of reads originating from single cells. As a high-throughput and low-bias method of single-cell sequencing, SiC-seq will enable a broader range of genomic studies targeted at diverse cell populations.

Single-cell sequencing reveals genotypic heterogeneity in biological systems, but current technologies lack the throughput necessary for the deep profiling of community composition and function. Here, we describe a microfluidic workflow for sequencing &gt;50,000 single-cell genomes from diverse cell populations.

Sequencing technologies have undergone a paradigm shift from bulk to single-cell resolution in response to an evolving understanding of the role of ...

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.

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.

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