We thank H. Tsutsui and D. Kurihara for providing us with transgenic plants, including the T. fournieriRPS5Ap::H2B-tdTomato line and the A. thaliana UBQ10pro::H2B-mClover line, respectively. This work was supported by the Institute of Transformative Bio-Molecules of Nagoya University and the Japan Advanced Plant Science Network. Financial support for this work was provided by grants from the Japan Science and Technology Agency (ERATO project grant no. JPMJER1004 for T.H.), a Grant-in-Aid for Scientific Research on Innovative Areas (Nos. JP16H06465 and JP16H06464 for T.H.), and Japan Society for the Promotion of Science (JSPS) Grants-in-Aid for challenging Exploratory Research (grant no. 26600061 for N.Y. and grant nos. 25650075 and 15K14542 for Y.S.).

1. Fabrication of the PDMS Microdevice to Examine Growing Pollen Tubes and Moss Protonemata
NOTE: We used a maskless photolithography instrument to prepare PDMS molds on silicon wafers. The details regarding the operation of the system are omitted in this manuscript. A standard photolithography technique9 using a photomask may also be used to create the PDMS molds described in this manuscript.
<ol>
	<li>Pour 11&#160;g of pre-polymer PDMS mixture (elastomer base:curing agent at a ratio of 10:1) into each 4-inch mold.</li>
	<li>Degas the mold prepared in step 1.1 for 20 min in a vacuum chamber.</li>
	<li>After curing at 65 &#176;C for 90 min in a non-convection oven, peel the PDMS layer off the mold, and punch access holes into the fluidic channels using biopsy punches. 
	NOTE: For the pollen tube device, the size of the hole must be adjusted to reflect the diameter of the pistil. This value will therefore vary by species. In this experiment, we punched a 1 mm hole for the pistil inlets and 1.5 mm holes for the liquid medium reservoirs. For the moss protonemata device, a 4 mm hole was used for the sample reservoir.</li>
	<li>Expose the PDMS layer and a glass bottom dish (3.5 -&#160;5 cm in diameter) to air plasma for 50 s.</li>
	<li>Press the PDMS layer into the glass bottom dish and heat at 65 &#176;C for 30 min in a non-convection oven to completely seal off the microfluidic network.</li>
</ol>
2. Fabrication of the PDMS Microdevice for Root Hairs
<ol>
	<li>Repeat steps 1.1 - 1.3 using the molds to prepare two PDMS layers for the root and root hair microdevices. 
	NOTE: We used a 2 mm hole for the liquid medium reservoirs in the root microdevice.</li>
	<li>Expose both PDMS layers to air plasma for 50 s.</li>
	<li>Assemble the two PDMS layers under a stereomicroscope using a custom-made desktop aligner. 
	NOTE: The microchannels on these PDMS layers must face each other. Before assembling, make sure that the alignment marks on both layers match up.</li>
	<li>Heat at 65 &#176;C for 30 min in a non-convection oven to completely seal off the microfluidic network.</li>
	<li>Remove the cover slip from the constructed microdevice and place the device on a glass bottom dish that is 5 cm in diameter.</li>
</ol>
3. Preparation of In Vitro Cell Culture Medium for Pollen Tubes (Torenia fournieri)
<ol>
	<li>Prepare modified Nitsch&#39;s medium as described previously15 and autoclave the medium at 121 &#176;C for 20 min. 
	NOTE: The composition of the modifed Nitsch&#39;s medium is NH4NO3 (80 mg/L), KNO3 (125 mg/L), Ca(NO3)2&#8231;4H2O (500 mg/L), MgSO4&#8231;7H2O (125 mg/L), KH2PO4 (125 mg/L), MnSO4&#8231;4H2O (3 mg/L), ZnSO4&#8231;7H2O (0.5 mg/L), H3BO3 (10 mg/L), CuSO4&#8231;5H2O (0.025 mg/L), Na2MoO4&#8231;2H2O (0.025 mg/L), sucrose (50,000 mg/L), and casein (500 mg/L). The prepared medium can be stored at 4 &#176;C for 4 months at least.</li>
	<li>Prepare 26% (w/v) polyethylene glycol using autoclaved deionized water and filter the solution with a 0.3 &#181;m pore filter. 
	NOTE: The prepared medium can be stored at 4 &#176;C for 1 month.</li>
	<li>Mix the reagents prepared in step 3.1 and 3.2 in a 1:1 (v/v) ratio. 
	NOTE: This medium should be prepared fresh for each use.</li>
</ol>
4. Preparation of In Vitro Cell Culture Medium for Root Hairs (Arabidopsis thaliana)
<ol>
	<li>Prepare a solution of 0.215% (w/v) Murashige &#38; Skoog Medium, 0.05% (w/v) MES, 1% (w/v) sucrose, and 1% (w/v) agar in deionized water. Autoclave the solution at 121 &#176;C for 20 min.</li>
</ol>
5. Preparation of In Vitro Cell Culture Medium for Moss Protonemata (Physcomitrella patens)
<ol>
	<li>Pre-incubate the moss protonemata in BCDAT medium16 in a Petri dish. 
	NOTE: The composition of BCDAT medium is 1 mM MgSO4, 10 mM KNO3, 45 &#181;M FeSO4, 1.8 mM KH2PO4 (pH adjusted to 6.5 with KOH), trace element solution (0.22 &#181;M CuSO4, 0.19 &#181;M ZnSO4, 10 &#181;M H3BO3, 0.1 &#181;M Na2MoO4, 2 &#181;M MnCl2, 0.23 &#181;M CoCl2, and 0.17 &#181;M KI), 1 mM CaCl2, and 5 mM diammonium(+)-tartrate.</li>
	<li>Culture the moss in the BCDATG medium in the microdevice. 
	NOTE: The BCDATG medium is BCDAT medium with 0.5 % (w/v) glucose. The prepared medium can be stored at 4 &#176;C for 1 month.</li>
</ol>
6. In Vitro Culturing of T. fournieri Pollen Tubes in the Microdevice
<ol>
	<li>Place the pollen tube microdevice in a vacuum chamber and degas for 20 min.</li>
	<li>Remove the microdevice from the vacuum chamber and introduce the growth medium to the pistil inlet using a micropipette with a very fine tip. Fill the remaining wells to their tops with the same medium. Wait for a few minutes until all the microchannels are filled with the medium by the vacuum created inside of the microchannels.</li>
	<li>Place some wet paper towel in the glass bottom dish to maintain the humidity in the dish.</li>
	<li>Transfer the pollen grains from a wild-type T. fournieri &#39;blue and white&#39; flower to its stigma using a dissection needle. 
	NOTE: We also conducted this experiment with a transgenic T. fournieri &#39;Crown violet&#39; flower (the RPS5Ap::H2B-tdTomato line17), with pollen tubes containing fluorescently labeled sperm cells and vegetative nuclei.</li>
	<li>Cut the pollinated style (1 cm long) using a blade.</li>
	<li>Insert the cut style into the inlet of the device as shown in Figure 2.</li>
	<li>Put a lid on the dish and seal it with tape.</li>
	<li>Place the device in an incubator at 28 &#176;C for 5 - 6 h in darkness.</li>
</ol>
7. In Vitro Culturing of A. thaliana Root Hairs in the Microdevice
NOTE: Steps 7.1 - 7.9 (except for 7.3 and 7.5) should be performed in a laminar flow hood.
<ol>
	<li>Sterilize seeds from A. thaliana Columbia (Col-0) and the transgenic line UBQ10pro::H2B-mClover (containing the fluorescent nuclear marker) by soaking them in sterile liquid (5% (v/v) household bleach and 0.02 % (v/v) Triton X-100) for 5 min.</li>
	<li>Thoroughly rinse the sterilized seeds with autoclaved water.</li>
	<li>Store the rinsed seeds in water for 2 days at 4 &#176;C in darkness.</li>
	<li>Sterilize the microdevice under UV light overnight.</li>
	<li>Place the microdevice in a vacuum chamber and degas for 20 min.</li>
	<li>Introduce the growth medium to the wells in the device using a micropipette. Wait a few minutes until the vacuum fills all the microchannels with the medium.</li>
	<li>Place autoclaved wet paper towel in the glass bottom dish to maintain the humidity in the dish.</li>
	<li>Transfer a sterilized seed into the inlet of the device.</li>
	<li>Put a lid on the dish and seal it with tape.</li>
	<li>Place the device vertically in an incubator at 22 &#176;C under continuous white light.</li>
	<li>Check root hair growth after 4 - 10 days of incubation. Obtain both bright field and fluorescent images using an inverted microscope.</li>
</ol>
8. In Vitro Culturing of P. patens (moss) Protonemata in the Microfluidic Device
NOTE: Steps 8.2 - 8.6 (except for 8.3) should be performed in a laminar flow hood.
<ol>
	<li>Preculture P. patens strain Gransden 200418 on BCDAT medium covered with cellophane in a Petri dish for one week under continuous white light at 25 &#176;C.</li>
	<li>Sterilize the microdevice under UV light overnight in a laminar flow hood.</li>
	<li>Place the microdevice in a vacuum chamber and degas for 20 min.</li>
	<li>Introduce the growth medium to the wells using a micropipette. Wait a few minutes until the vacuum fills all the microchannels with the medium.</li>
	<li>Add autoclaved water into the dish surrounding the microdevice to maintain the humidity in the dish.</li>
	<li>Transfer a small piece of moss protonemata tissue to the inlet of the microdevice and culture the tissue in an incubator at 25 &#176;C under continuous white light.</li>
	<li>Check the moss protonemata growth after 2 - 3 weeks of incubation. Obtain bright field images using a microscope.</li>
</ol>
9. Time-lapse Imaging of T. fournieri Pollen Tube Growth
<ol>
	<li>Place the microdevice on an inverted fluorescence microscope equipped with image acquisition hardware (e.g., a CCD camera) and software. Find the microgap locations. 
	NOTE: For the image acquisition software, we used a commercially available product (Table of Materials). Open source microscopy software such as &#181;Manager is also available online (https://micro-manager.org/wiki/Micro-Manager). We recommend installing a microscope condenser on the microscope to obtain higher resolution images.</li>
	<li>Capture bright field images every 10 s using microscope image acquisition software.</li>
	<li>To observe the fluorescently labeled sperm cells and vegetative nucleus in the pollen tube of the RPS5Ap::H2B-tdTomato line, irradiate the specimen with 561 nm laser and use a bandpass optical filter (578/105 nm). 
	NOTE: Although the time-lapse images of pollen tube growth were captured frequently, imaging did not appear to affect growth. However, it is always recommended to minimize the laser intensity, exposure time, and time-lapse interval, to minimize the phototoxicity and photobleaching of the fluorophore.</li>
	<li>Adjust the brightness and contrast of the images and prepare a video file using ImageJ software (https://imagej.nih.gov/ij/).</li>
</ol>

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The authors declare that they have no competing financial interests.

Several critical steps in the protocol need to be followed precisely to obtain the results presented above. First, the PDMS layer and glass bottom dish surfaces must both be treated with plasma for a sufficient amount of time before bonding. Otherwise, the PDMS layer may locally detach from the glass surface while tip-growing cells are crossing the microgaps. Another crucial step in the root hair and moss protonemata protocol is the sterilization of the microdevice. Normally, root hairs and moss protonemata cells need to...

After pollen grains germinate on a stigma, each grain produces a single pollen tube that carries sperm cells to the egg cell and the central cell in the ovule for double fertilization. Pollen tubes elongate through the style and eventually reach the ovule by sensing multiple guidance cues along their way1. During the elongation, pollen tubes encounter a series of physical barriers; the transmitting track is filled with cells, and pollen tubes must enter the minute micropylar opening of the ovule to reach their target (Figure 1A)2. Therefore, pollen tubes must have the ability to penetrate physical obstacles, while tolerating the compressive stress from their surroundings. Root hairs are another type of tip-growing plant cell that must withstand physical obstacles in the environment, in the form of packed soil particles (Figure 1B).
Various mechanical properties of the pollen tube have been studied, including turgor pressure and stiffness of the cell&#39;s apical region, which can be measured using the incipient plasmolysis method3,4 and cellular force microscopy (CFM)5,6, respectively. However, these methods alone do not reveal whether pollen tubes are capable of elongating through physical barriers along their growth paths. An alternative technique that allows pollen tube elongation to be monitored in vivo is two photon microscopy7. However, with this method, it is difficult to observe the morphological changes in individual pollen tubes deep inside the ovule tissue. Additionally, root hair growth in soil can be visualized using X-ray computed tomography (CT) and magnetic resonance imaging (MRI)8, albeit with low resolution. Here, we present a method that can be used to acquire high-resolution images of a cell&#39;s deformation process on a conventional microscope.
The overall goal of the method described here is to visualize the elongation capability of tip-growing plant cells, including pollen tubes, root hairs, and moss protonemata, in extremely small spaces. As the poly-dimethylsiloxane (PDMS) microdevices presented in this manuscript are optically transparent and air permeable, we can culture living cells inside the device and observe their growth behaviors under a microscope. It is also possible to create micro ~ nanometer scale spaces by the soft lithography technique9 with the use of molds. These features allow us to study the elongation capability of tip-growing plant cells in a physically confined environment.
In this work, we constructed 1 &#181;m wide gaps (4 &#181;m in height) in microfluidic devices and examined the ability of pollen tubes to penetrate these artificial obstacles that are much smaller than the diameter of the cylindrical pollen tube (approximately 8 &#181;m). This experimental platform enables us to visualize the pollen tube&#39;s response to microgaps and capture time-lapse images of the response, which track the cell&#39;s deformation process. We also developed the microdevices that can be used to investigate the penetration capability of root hairs and moss protonemata. Several microdevices have been reported to date that enable the visualization of plant root10,11,12,13 and moss protonemata14 growth at high resolution. In our device, a series of root hair growth channels are perpendicularly connected to a root growth chamber, and individual root hairs (approximately 7 &#181;m in diameter) are guided to fluidic channels with a 1 &#181;m wide gap. We also cultured moss protonemata (approximately 20 &#181;m in diameter) in a microdevice containing microgaps to examine their responses to these physical barriers. The proposed microfluidic-based approach allows us to explore the capability of various tip-growing plant cells to elongate through extremely small spaces, which cannot be examined by any other currently available method.

<table border="0" cellpadding="0" cellspacing="0" width="927">
	<tbody>
		<tr height="18">
			<td height="18" style="height:19px;width:216px;">PDMS</td>
			<td style="width:165px;">Dow Corning Co.</td>
			<td style="width:119px;">Sylgard184</td>
			<td style="width:427px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Murashige &#38; Skoog Medium</td>
			<td style="width:165px;">Wako Pure Chemical</td>
			<td style="width:119px;">392-00591</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">MES</td>
			<td style="width:165px;">Dojindo</td>
			<td style="width:119px;">345-01625</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Sucrose</td>
			<td style="width:165px;">Wako Pure Chemical</td>
			<td style="width:119px;">196-00015</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">50 mm glass-bottom dish</td>
			<td style="width:165px;">Matsunami Glass</td>
			<td style="width:119px;">D210402</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">35 mm glass-bottom dish</td>
			<td style="width:165px;">Iwaki&#160;</td>
			<td style="width:119px;">3971-035</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Surgical blade</td>
			<td style="width:165px;">Feather</td>
			<td style="width:119px;">No.11</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">biopsy punches</td>
			<td style="width:165px;">Harris</td>
			<td style="width:119px;"></td>
			<td>Uni-Core</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Gel loading tips</td>
			<td style="width:165px;">Bio-Bik</td>
			<td style="width:119px;">124-R-204</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Inverted Microscope</td>
			<td style="width:165px;">Olympus</td>
			<td style="width:119px;">IX83</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:22px;width:216px;">CSU-W1</td>
			<td style="width:165px;">Yokogawa Electric</td>
			<td style="width:119px;"></td>
			<td>No Catalog number is avairable for this customized microscope</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">MetaMorph imaging software</td>
			<td style="width:165px;">Molecular Devices</td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
	</tbody>
</table>

development of microfluidic devices to study the elongation capability of tip-growing plant cells in extremely small spaces

In vivo, tip-growing plant cells need to overcome a series of physical barriers; however, researchers lack the methodology to visualize cellular behavior in such restrictive conditions. To address this issue, we have developed growth chambers for tip-growing plant cells that contain a series of narrow, micro-fabricated gaps (~1 &#181;m) in a poly-dimethylsiloxane (PDMS) substrate. This transparent material allows the user to monitor tip elongation processes in individual cells during microgap penetration by time-lapse imaging. Using this experimental platform, we observed morphological changes in pollen tubes as they penetrated the microgap. We captured the dynamic changes in the shape of a fluorescently labeled vegetative nucleus and sperm cells in a pollen tube during this process. Furthermore, we demonstrated the capability of root hairs and moss protonemata to penetrate the 1 &#181;m gap. This in vitro platform can be used to study how individual cells respond to physically constrained spaces and may provide insights into tip-growth mechanisms.

As illustrated in Figure 1, tip-growing plant cells encounter a series of physical barriers along their growth paths in vivo. The microfluidic in vitro cell culture platforms presented in this study enabled the examination the of tip-growing process in three types of plant cells (pollen tubes, root hairs, and moss protonemata) through 1 &#181;m artificial gaps (Figure 3, Figure 4, <...

Watch this Scientific Journal Video about Development of Microfluidic Devices to Study the Elongation Capability of Tip-growing Plant Cells in Extremely Small Spaces at JoVE.com

Development of Microfluidic Devices to Study the Elongation Capability of Tip-growing Plant Cells in Extremely Small Spaces

We describe a method to investigate the capability of tip-growing plant cells, including pollen tubes, root hairs, and moss protonemata, to elongate through extremely narrow gaps (~1 &#181;m) in a microfluidic device.

In vivo, tip-growing plant cells need to overcome a series of physical barriers; however, researchers lack the methodology to visualize cellular behavior ...

development-microfluidic-devices-to-study-elongation-capability-tip

Universidad de Cartagena UDEC

Research

JoVE Journal

Bioengineering

7.3K Views.  Nagoya University. We describe a method to investigate the capability of tip-growing plant cells, including pollen tubes, root hairs, and moss protonemata, to elongate through extremely narrow gaps (~1 µm) in a microfluidic device.

Video: Development of Microfluidic Devices to Study the Elongation Capability of Tip-growing Plant Cells in Extremely Small Spaces

Separating Beads and Cells in Multi-channel Microfluidic Devices Using Dielectrophoresis and Laminar Flow

Microfluidic devices have advanced cell studies by providing a dynamic fluidic environment on the scale of the cell for studying, manipulating, sorting and counting cells. However, manipulating the cell within the fluidic domain remains a challenge and requires complicated fabrication protocols for forming valves and electrodes, or demands specialty equipment like optical tweezers. Here, we demonstrate that conventional printed circuit boards (PCB) can be used for the non-contact manipulation of cells by employing dielectrophoresis (DEP) for bead and cell manipulation in laminar flow fields for bioactuation, and for cell and bead separation in multichannel microfluidic devices. First, we present the protocol for assembling the DEP electrodes and microfluidic devices, and preparing the cells for DEP. Then, we characterize the DEP operation with polystyrene beads. Lastly, we show representative results of bead and cell separation in a multichannel microfluidic device. In summary, DEP is an effective method for manipulating particles (beads or cells) within microfluidic devices.

Dielectrophoresis (DEP) is an effective method to manipulate cells. Printed circuit boards (PCB) can provide inexpensive, reusable and effective electrodes for contact-free cell manipulation within microfluidic devices. By combining PDMS-based microfluidic channels with coverslips on PCBs, we demonstrate bead and cell manipulation and separation within multichannel microfluidic devices.

Microfluidic devices have advanced cell studies by providing a dynamic fluidic environment on the scale of the cell for studying, manipulating, sorting ...

Design of a Cyclic Pressure Bioreactor for the Ex Vivo Study of Aortic Heart Valves

The aortic valve, located between the left ventricle and the aorta, allows for unidirectional blood flow, preventing backflow into the ventricle. Aortic valve leaflets are composed of interstitial cells suspended within an extracellular matrix (ECM) and are lined with an endothelial cell monolayer. The valve withstands a harsh, dynamic environment and is constantly exposed to shear, flexion, tension, and compression. Research has shown calcific lesions in diseased valves occur in areas of high mechanical stress as a result of endothelial disruption or interstitial matrix damage1-3. Hence, it is not surprising that epidemiological studies have shown high blood pressure to be a leading risk factor in the onset of aortic valve disease4. 
The only treatment option currently available for valve disease is surgical replacement of the diseased valve with a bioprosthetic or mechanical valve5. Improved understanding of valve biology in response to physical stresses would help elucidate the mechanisms of valve pathogenesis. In turn, this could help in the development of non-invasive therapies such as pharmaceutical intervention or prevention. Several bioreactors have been previously developed to study the mechanobiology of native or engineered heart valves6-9. Pulsatile bioreactors have also been developed to study a range of tissues including cartilage10, bone11 and bladder12. The aim of this work was to develop a cyclic pressure system that could be used to elucidate the biological response of aortic valve leaflets to increased pressure loads. 
The system consisted of an acrylic chamber in which to place samples and produce cyclic pressure, viton diaphragm solenoid valves to control the timing of the pressure cycle, and a computer to control electrical devices. The pressure was monitored using a pressure transducer, and the signal was conditioned using a load cell conditioner. A LabVIEW program regulated the pressure using an analog device to pump compressed air into the system at the appropriate rate. The system mimicked the dynamic transvalvular pressure levels associated with the aortic valve; a saw tooth wave produced a gradual increase in pressure, typical of the transvalvular pressure gradient that is present across the valve during diastole, followed by a sharp pressure drop depicting valve opening in systole. The LabVIEW program allowed users to control the magnitude and frequency of cyclic pressure. The system was able to subject tissue samples to physiological and pathological pressure conditions. This device can be used to increase our understanding of how heart valves respond to changes in the local mechanical environment.

Design of a Cyclic Pressure Bioreactor for the Ex Vivo Study of Aortic Heart Valves

A cyclic pressure bioreactor capable of subjecting heart valve tissue to physiological and pathological pressure conditions has been designed. A LabVIEW program allows users to control pressure magnitude, amplitude and frequency. This device can be used to study the mechanobiology of heart valve tissue or isolated cells.

The aortic valve, located between the left ventricle and the aorta, allows for unidirectional blood flow, preventing backflow into the ventricle. Aortic ...

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 ...

Layer-by-layer Collagen Deposition in Microfluidic Devices for Microtissue Stabilization

Although microfluidics provides exquisite control of the cellular microenvironment, culturing cells within microfluidic devices can be challenging. 3D culture of cells in collagen type I gels helps to stabilize cell morphology and function, which is necessary for creating microfluidic tissue models in microdevices. Translating traditional 3D culture techniques for tissue culture plates to microfluidic devices is often difficult because of the limited channel dimensions. In this method, we describe a technique for modifying native type I collagen to generate polycationic and polyanionic collagen solutions that can be used with layer-by-layer deposition to create ultrathin collagen assemblies on top of cells cultured in microfluidic devices. These thin collagen layers stabilize cell morphology and function, as shown using primary hepatocytes as an example cell, allowing for the long term culture of microtissues in microfluidic devices.

The creation of functional microtissues within microfluidic devices requires the stabilization of cell phenotypes by adapting traditional cell culture techniques to the limited spatial dimensions in microdevices. Modification of collagen allows the layer-by-layer deposition of ultrathin collagen assemblies that can stabilize primary cells, such as hepatocytes, as microfluidic tissue models.

Although microfluidics provides exquisite control of the cellular microenvironment, culturing cells within microfluidic devices can be challenging. 3D ...

Construction of Defined Human Engineered Cardiac Tissues to Study Mechanisms of Cardiac Cell Therapy

Human cardiac tissue engineering can fundamentally impact therapeutic discovery through the development of new species-specific screening systems that replicate the biofidelity of three-dimensional native human myocardium, while also enabling a controlled level of biological complexity, and allowing non-destructive longitudinal monitoring of tissue contractile function. Initially, human engineered cardiac tissues (hECT) were created using the entire cell population obtained from directed differentiation of human pluripotent stem cells, which typically yielded less than 50% cardiomyocytes. However, to create reliable predictive models of human myocardium, and to elucidate mechanisms of heterocellular interaction, it is essential to accurately control the biological composition in engineered tissues. 
To address this limitation, we utilize live cell sorting for the cardiac surface marker SIRP&#945; and the fibroblast marker CD90 to create tissues containing a 3:1 ratio of these cell types, respectively, that are then mixed together and added to a collagen-based matrix solution. Resulting hECTs are, thus, completely defined in both their cellular and extracellular matrix composition.
Here we describe the construction of defined hECTs as a model system to understand mechanisms of cell-cell interactions in cell therapies, using an example of human bone marrow-derived mesenchymal stem cells (hMSC) that are currently being used in human clinical trials. The defined tissue composition is imperative to understand how the hMSCs may be interacting with the endogenous cardiac cell types to enhance tissue function. A bioreactor system is also described that simultaneously cultures six hECTs in parallel, permitting more efficient use of the cells after sorting.

This manuscript describes the creation of defined engineered cardiac tissues using surface marker expression and cell sorting. The defined tissues can then be used in a multi-tissue bioreactor to investigate mechanisms of cardiac cell therapy in order to provide a functional, yet controlled, model system of the human heart.

Human cardiac tissue engineering can fundamentally impact therapeutic discovery through the development of new species-specific screening systems that ...

Using Adhesive Patterning to Construct 3D Paper Microfluidic Devices

We demonstrate the use of patterned aerosol adhesives to construct both planar and nonplanar 3D paper microfluidic devices. By spraying an aerosol adhesive through a metal stencil, the overall amount of adhesive used in assembling paper microfluidic devices can be significantly reduced. We show on a simple 4-layer planar paper microfluidic device that the optimal adhesive application technique and device construction style depends heavily on desired performance characteristics. By moderately increasing the overall area of a device, it is possible to dramatically decrease the wicking time and increase device success rates while also reducing the amount of adhesive required to keep the device together. Such adhesive application also causes the adhesive to form semi-permanent bonds instead of permanent bonds between paper layers, enabling single-use devices to be non-destructively disassembled after use. Nonplanar 3D origami devices also benefit from the semi-permanent bonds during folding, as it reduces the likelihood that unrelated faces may accidently stick together. Like planar devices, nonplanar structures see reduced wicking times with patterned adhesive application vs uniformly applied adhesive.

We demonstrate the use of patterned aerosol adhesives to construct 3D paper microfluidic devices. This method of adhesive application forms semi-permanent bonds between layers, enabling single-use devices to be non-destructively disassembled after use and to ease folding complex nonplanar structures.

We demonstrate the use of patterned aerosol adhesives to construct both planar and nonplanar 3D paper microfluidic devices. By spraying an aerosol ...

The Arteriovenous (AV) Loop in a Small Animal Model to Study Angiogenesis and Vascularized Tissue Engineering

A functional blood vessel network is a prerequisite for the survival and growth of almost all tissues and organs in the human body. Moreover, in pathological situations such as cancer, vascularization plays a leading role in disease progression. Consequently, there is a strong need for a standardized and well-characterized in vivo model in order to elucidate the mechanisms of neovascularization and develop different vascularization approaches for tissue engineering and regenerative medicine.
We describe a microsurgical approach for a small animal model for induction of a vascular axis consisting of a vein and artery that are anastomosed to an arteriovenous (AV) loop. The AV loop is transferred to an enclosed implantation chamber to create an isolated microenvironment in vivo, which is connected to the living organism only by means of the vascular axis. Using 3D imaging (MRI, micro-CT) and immunohistology, the growing vasculature can be visualized over time. By implanting different cells, growth factors and matrices, their function in blood vessel network formation can be analyzed without any disturbing influences from the surroundings in a well controllable environment. 
In addition to angiogenesis and antiangiogenesis studies, the AV loop model is also perfectly suited for engineering vascularized tissues. After a certain prevascularization time, the generated tissues can be transplanted into the defect site and microsurgically connected to the local vessels, thereby ensuring immediate blood supply and integration of the engineered tissue. By varying the matrices, cells, growth factors and chamber architecture, it is possible to generate various tissues, which can then be tailored to the individual patient's needs.

The Arteriovenous AV Loop in a Small Animal Model to Study Angiogenesis and Vascularized Tissue Engineering

We describe a microsurgical approach for the generation of an arteriovenous (AV) loop as a model for analyzing vascularization in vivo in an isolated and well-characterized environment. This model is not only useful for the investigation of angiogenesis, but is also optimally suited for engineering axially vascularized and transplantable tissues.

A functional blood vessel network is a prerequisite for the survival and growth of almost all tissues and organs in the human body. Moreover, in ...

Automated Robotic Liquid Handling Assembly of Modular DNA Devices

Recent advances in modular DNA assembly techniques have enabled synthetic biologists to test significantly more of the available &#34;design space&#34; represented by &#34;devices&#34; created as combinations of individual genetic components. However, manual assembly of such large numbers of devices is time-intensive, error-prone, and costly. The increasing sophistication and scale of synthetic biology research necessitates an efficient, reproducible way to accommodate large-scale, complex, and high throughput device construction.
Here, a DNA assembly protocol using the Type-IIS restriction endonuclease based Modular Cloning (MoClo) technique is automated on two liquid-handling robotic platforms. Automated liquid-handling robots require careful, often times tedious optimization of pipetting parameters for liquids of different viscosities (e.g. enzymes, DNA, water, buffers), as well as explicit programming to ensure correct aspiration and dispensing of DNA parts and reagents. This makes manual script writing for complex assemblies just as problematic as manual DNA assembly, and necessitates a software tool that can automate script generation. To this end, we have developed a web-based software tool, http://mocloassembly.com, for generating combinatorial DNA device libraries from basic DNA parts uploaded as Genbank files. We provide access to the tool, and an export file from our liquid handler software which includes optimized liquid classes, labware parameters, and deck layout. All DNA parts used are available through Addgene, and their digital maps can be accessed via the Boston University BDC ICE Registry. Together, these elements provide a foundation for other organizations to automate modular cloning experiments and similar protocols.
The automated DNA assembly workflow presented here enables the repeatable, automated, high-throughput production of DNA devices, and reduces the risk of human error arising from repetitive manual pipetting. Sequencing data show the automated DNA assembly reactions generated from this workflow are ~95% correct and require as little as 4% as much hands-on time, compared to manual reaction preparation.

Here, an automated workflow to perform modular DNA &#34;device&#34; assembly using a modular cloning DNA assembly method on liquid-handling robots is presented. The protocol uses an intuitive software tool for generating liquid handler picklists for combinatorial DNA device library generation, which we demonstrate using two liquid handling platforms.

Recent advances in modular DNA assembly techniques have enabled synthetic biologists to test significantly more of the available &#34;design space&#34; ...

Multi-step Variable Height Photolithography for Valved Multilayer Microfluidic Devices

Microfluidic systems have enabled powerful new approaches to high-throughput biochemical and biological analysis. However, there remains a barrier to entry for non-specialists who would benefit greatly from the ability to develop their own microfluidic devices to address research questions. Particularly lacking has been the open dissemination of protocols related to photolithography, a key step in the development of a replica mold for the manufacture of polydimethylsiloxane (PDMS) devices. While the fabrication of single height silicon masters has been explored extensively in literature, fabrication steps for more complicated photolithography features necessary for many interesting device functionalities (such as feature rounding to make valve structures, multi-height single-mold patterning, or high aspect ratio definition) are often not explicitly outlined. 
Here, we provide a complete protocol for making multilayer microfluidic devices with valves and complex multi-height geometries, tunable for any application. These fabrication procedures are presented in the context of a microfluidic hydrogel bead synthesizer and demonstrate the production of droplets containing polyethylene glycol (PEG diacrylate) and a photoinitiator that can be polymerized into solid beads. This protocol and accompanying discussion provide a foundation of design principles and fabrication methods that enables development of a wide variety of microfluidic devices. The details included here should allow non-specialists to design and fabricate novel devices, thereby bringing a host of recently developed technologies to their most exciting applications in biological laboratories.

Multilayer microfluidic devices often involve the fabrication of master molds with complex geometries for functionality. This article presents a complete protocol for multi-step photolithography with valves and variable height features tunable to any application. As a demonstration, we fabricate a microfluidic droplet generator capable of producing hydrogel beads.

Microfluidic systems have enabled powerful new approaches to high-throughput biochemical and biological analysis. However, there remains a barrier to ...

Fabrication of Three-dimensional Paper-based Microfluidic Devices for Immunoassays

Paper wicks fluids autonomously due to capillary action. By patterning paper with hydrophobic barriers, the transport of fluids can be controlled and directed within a layer of paper. Moreover, stacking multiple layers of patterned paper creates sophisticated three-dimensional microfluidic networks that can support the development of analytical and bioanalytical assays. Paper-based microfluidic devices are inexpensive, portable, easy to use, and require no external equipment to operate. As a result, they hold great promise as a platform for point-of-care diagnostics. In order to properly evaluate the utility and analytical performance of paper-based devices, suitable methods must be developed to ensure their manufacture is reproducible and at a scale that is appropriate for laboratory settings. In this manuscript, a method to fabricate a general device architecture that can be used for paper-based immunoassays is described. We use a form of additive manufacturing (multi-layer lamination) to prepare devices that comprise multiple layers of patterned paper and patterned adhesive. In addition to demonstrating the proper use of these three-dimensional paper-based microfluidic devices with an immunoassay for human chorionic gonadotropin (hCG), errors in the manufacturing process that may result in device failures are discussed. We expect this approach to manufacturing paper-based devices will find broad utility in the development of analytical applications designed specifically for limited-resource settings.

We detail a method to fabricate three-dimensional paper-based microfluidic devices for use in the development of immunoassays. Our approach to device assembly is a type of multilayer, additive manufacturing. We demonstrate a sandwich immunoassay to provide representative results for these types of paper-based devices.

Paper wicks fluids autonomously due to capillary action. By patterning paper with hydrophobic barriers, the transport of fluids can be controlled and ...

Development of Microfluidic Devices to Study the Elongation Capability of Tip-growing Plant Cells in Extremely Small Spaces

Summary

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Chapters in this video

Separating Beads and Cells in Multi-channel Microfluidic Devices Using Dielectrophoresis and Laminar Flow

Design of a Cyclic Pressure Bioreactor for the Ex Vivo Study of Aortic Heart Valves

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

Layer-by-layer Collagen Deposition in Microfluidic Devices for Microtissue Stabilization

Construction of Defined Human Engineered Cardiac Tissues to Study Mechanisms of Cardiac Cell Therapy

Using Adhesive Patterning to Construct 3D Paper Microfluidic Devices

The Arteriovenous (AV) Loop in a Small Animal Model to Study Angiogenesis and Vascularized Tissue Engineering

Automated Robotic Liquid Handling Assembly of Modular DNA Devices

Multi-step Variable Height Photolithography for Valved Multilayer Microfluidic Devices

Fabrication of Three-dimensional Paper-based Microfluidic Devices for Immunoassays