Name: Development of Microfluidic Devices to Study the Elongation Capability of Tip-growing Plant Cells in Extremely Small Spaces (Video) | JoVE Journal
Uploaded: 2018-05-22T12:30:00.000+00:00
Duration: 7 min 1 s
Description: Learn microfluidic techniques to study elongation of tip-growing plant cells through narrow gaps. This method enables high-resolution imaging of pollen tubes and moss protonemata deformation at micrometer scales.

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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><tbody><tr><td>PDMS</td><td>Dow Corning Co.</td><td>Sylgard184</td><td></td></tr><tr><td>Murashige &amp;amp; Skoog Medium</td><td>Wako Pure Chemical</td><td>392-00591</td><td></td></tr><tr><td>MES</td><td>Dojindo</td><td>345-01625</td><td></td></tr><tr><td>Sucrose</td><td>Wako Pure Chemical</td><td>196-00015</td><td></td></tr><tr><td>50 mm glass-bottom dish</td><td>Matsunami Glass</td><td>D210402</td><td></td></tr><tr><td>35 mm glass-bottom dish</td><td>Iwaki&amp;nbsp;</td><td>3971-035</td><td></td></tr><tr><td>Surgical blade</td><td>Feather</td><td>No.11</td><td></td></tr><tr><td>biopsy punches</td><td>Harris</td><td></td><td>Uni-Core</td></tr><tr><td>Gel loading tips</td><td>Bio-Bik</td><td>124-R-204</td><td></td></tr><tr><td>Inverted Microscope</td><td>Olympus</td><td>IX83</td><td></td></tr><tr><td>CSU-W1</td><td>Yokogawa Electric</td><td></td><td>No Catalog number is avairable for this customized microscope</td></tr><tr><td>MetaMorph imaging software</td><td>Molecular Devices</td><td></td><td></td></tr></tbody></table>

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

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 µm artificial gaps (Figure 3, Figure 4, <stron...

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 ...

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We demonstrate a microfluidics-based assay to measure the timescale for cells to transit through a sequence of micron-scale constrictions.

A Microfluidic Technique to Probe Cell Deformability

Micro-fabricated devices integrated with fluidic components provide an in vitro platform for cell studies mimicking the in vivo micro-environment. We developed polymethylmethacrylate-based microfluidic chips for studying cellular responses under single or coexisting chemical/electrical/shear stress stimuli.

Designing Microfluidic Devices for Studying Cellular Responses Under Single or Coexisting Chemical/Electrical/Shear Stress Stimuli

A microfluidic channel with deformable sidewalls offers flow control, particle handling, channel dimension customization and other reconfigurations while in use. We describe a method for fabricating a microfluidic channel with sidewalls made of an array of pins that allows their shape to change.

Reconfigurable Microfluidic Channel with Pin-discretized Sidewalls

We describe a method to construct devices for 3D culture and experimentation with cells and multicellular organoids. This device allows analysis of cellular responses to soluble signals in 3D microenvironments with defined chemoattractant gradients. Organoids are better than single cells at detection of weak noisy inputs.

3D Analysis of Multi-cellular Responses to Chemoattractant Gradients

Here, we provide a microfluidic chip and an automatically controlled, highly efficient circulation microfluidic system that recapitulates the initial microenvironment of neovascularization, allowing endothelial cells (ECs) to be stimulated by high luminal shear stress, physiological level of transendothelial flow, and various vascular endothelial growth factor (VEGF) distribution simultaneously.

Microfluidic Model to Mimic Initial Event of Neovascularization

Owing to the opacity of soil, interactions between its constituent microbes cannot easily be visualised with cellular resolution. Here, two microfluidic tools are presented, which offer new opportunities for investigating fungal-microbial interactions. The devices are versatile and simple to use, enabling high spatiotemporal control and high-resolution imaging at the cellular level.

Microfluidic Tools for Probing Fungal-Microbial Interactions at the Cellular Level

This protocol details the design and fabrication of a microfluidic device suitable for investigating microtubule polymer mechanics. The synthesis of microfabrication, automated flow control, and computational modeling techniques enables a flexible system ideally suited for probing the cellular cytoskeleton in vitro.

Development of a Microfluidics-Based Approach for Investigating Microtubule Polymer Mechanics

Learn microfluidic techniques to study elongation of tip-growing plant cells through narrow gaps. This method enables high-resolution imaging of pollen tubes and moss protonemata deformation at micrometer scales.

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

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