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 be cultured in the microdevice for a couple of weeks. Without sterilizing the device, microorganisms may flourish, which could affect the growth of these tip-growing cells. Sterilization of the pollen tube device is not as critical, since the experiment is concluded within half a day. We have regularly used the pollen tube device without sterilization and did not observe any undesirable effects during the experiment.
The microchannel design should be modified according to the cells of interest. In our work, the channel design used for the A. thaliana root hair experiment was different from the one used for the T. fournieri pollen tube and the P. patens protonemata experiments. Different devices are required because the samples have different dimensions; for instance, root hairs are much shorter than pollen tubes and moss protonemata. Therefore, our root hair device was designed such that the microgap regions were adjacent to the main root channel. In addition to the channel design, the procedure for preparing the root hair device also varies from the procedures used for pollen tubes or moss protonemata. For the pollen tube and moss protonemata studies, we used a microdevice containing a single PDMS layer with the microchannels sealed against a glass bottom dish. However, the device used to study root hairs comprises two PDMS layers: the top layer with a microchannel (depth: 200 &#181;m) for the main root is sealed against the bottom layer, containing microchannels (depth: 4 &#181;m) for root hairs. The standard photolithography technique that was employed to fabricate the silicon mold is limited to an aspect ratio of around 5:1. Therefore, to create a 1 &#181;m wide gap, the channel height will be limited to approximately 5 &#181;m. It is also technically challenging to precisely align a deep channel (e.g., 200 &#181;m) near the 1 &#181;m wide gap on a silicon mold. Therefore, we prepared separate PDMS layers for the main root and the root hairs, and constructed the device by sealing them together.
While this protocol was successfully used to visualize the tip growing process of pollen tubes through the microgaps (Supplementary Movie 1 and 2), time-lapse imaging of root hair and moss protonemata elongation at the microgap region would be more challenging. In the microdevice used for culturing A. thaliana root hairs, the height of the side channels where the microgaps are located (4 &#181;m) is much shallower than that of the root growth chamber (200 &#181;m), so most root hairs are unable to proceed into the side channels. Even if the root hairs do find the side channel entrance, it is hard to predict when they will enter it. To capture time-lapse images of root hair growth at the microgap, we recommend using a microscope equipped with an automated stage that can be programmed to be situated at multiple positions for image acquisition. When such instrumentation is unavailable, we suggest searching for the root hairs that have already entered the side channels but have not yet crossed the microgaps. Since the growth rate of A. thaliana root hairs is much slower (typically 1 - 2.5 &#181;m/min20) than that of T. fournieri pollen tubes (typically 22 - 23 &#181;m/min19), it may be possible to capture time-lapse images of those root hairs that are about to cross the microgaps. On the other hand, moss protonemata observations require a time-lapse system that can be exclusively used for this study, because a prolonged observation period is required due to the slow growth rate.
The microfluidic approach presented here is the first method that allows us to study the capability of tip-growing plant cells to elongate through extremely narrow spaces. These microdevices may be useful for screening assays that investigate the functions of genes associated with cell wall biosynthesis and the integrity of tip-growing cells through their penetration capability. In addition, Denais et al.21 recently demonstrated a nuclear envelope rupture and repair mechanism by squeezing a cancer cell through a confined space prepared in a microfluidic device, and it may be possible to study whether a similar mechanism exists in pollen tubes using our device. This newly developed experimental platform can be used to investigate how individual cells respond to physically constrained spaces and may thus enhance our understanding of tip-growth mechanisms.

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

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, Figure 5). For the pollen tube study, live-cell imaging was used to monitor morphological changes in the apical region of pollen tubes as well as the vegetative nucleus and sperm cells in response to encountering an extremely small space (Supplemental Movie 1 and 2).
Although most of the microgaps were successfully fabricated by employing the method described here, we noticed that a few of them were completely closed (Figure 6). Because the 1 &#181;m wide channels created on the silicon mold are fragile, repeated use of this mold might damage the gaps, leading to gap blocking on the PDMS layer. Therefore, before performing the experiment, it is important to check that the PDMS microgaps are intact.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/57262/57262fig1.jpg" /> 
Figure 1: Penetration of tip-growing plant cells into physically constrained spaces. (A) Pollen tube elongating through several physical barriers on its way to an ovule. Barriers include transmitting tract tissue in the style and the integuments surrounding the micropyle. (B) Root hairs penetrating through dense soil. Insets in (A) and (B) show enlargements of the boxed regions. <a href="/files/ftp_upload/57262/57262fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/57262/57262fig2.jpg" /> 
Figure 2: Characteristics of the microdevices used to study T. fournieri pollen tubes, A. thaliana root hairs, and P. patens protonemata. In the schematic drawings of microchannels, a sample is placed at the location indicated by an asterisk mark (*). Microchannel designs, photographs of each device, channel structures, and cell culturing periods are summarized. This figure has been modified with permission from Yanagisawa et al. 201719. Scale bar, 20 &#181;m. <a href="/files/ftp_upload/57262/57262fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/57262/57262fig3.jpg" /> 
Figure 3: T. fournieri pollen tube elongation through a 1 &#181;m gap. (A) Pollen tube passing through a 1 &#181;m wide and 4 &#181;m high gap. The time-lapse bright field images were captured using an inverted microscope equipped with a spinning-disk confocal system. Scale bar, 20 &#181;m. (B) Time lapse images of a fluorescently labeled vegetative nucleus and sperm cells in a pollen tube of the RPS5Ap::H2B-tdTomato line crossing the microgap. Scale bar, 20 &#181;m. Figure (A) and (B) have been modified with permission from Yanagisawa et al. 201719. <a href="/files/ftp_upload/57262/57262fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/57262/57262fig4.jpg" /> 
Figure 4. A. thaliana root hair elongation through a 1 &#181;m gap. (A) Root and root hair growth in the microdevice. The microgaps (1 &#181;m in width and 4 &#181;m in height) are located on the left side of the root growth channel. The microchannels on the right side do not contain microgaps and can be used as a control. Scale bar, 100 &#181;m. (B) Bright field, (C) fluorescence, and (D) merged images of root hairs passing through the microgaps. The nuclei in root hairs are fluorescently labeled in the UBQ10pro::H2B-mClover line. The tip of each root hair is indicated by an arrow. Scale bar, 30 &#181;m. These figures have been modified with permission from Yanagisawa et al. 201719. <a href="/files/ftp_upload/57262/57262fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/57262/57262fig5.jpg" /> 
Figure 5. P. patens protonemata elongation through a 1 &#181;m gap. Moss protonemata cells crossing through a 1 &#181;m gap (4 &#181;m in height), which was widened due to turgor pressure of the protonemata. Scale bar, 20 &#181;m. The figure has been modified with permission from Yanagisawa et al. 201719. <a href="/files/ftp_upload/57262/57262fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/57262/57262fig6.jpg" /> 
Figure 6. Closed microgaps. SEM image of the 1 &#181;m gap region. The microgap on the right is completely closed. <a href="/files/ftp_upload/57262/57262fig6large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Supplementary Movie 1. Pollen tube growth through a 1 &#181;m gap.Time-lapse bright field images were captured every 10 s using an inverted microscope equipped with a spinning disk confocal microscope. Scale bar, 20 &#181;m. <a href="/files/ftp_upload/57262/movie1.avi">Please click here to download this file.</a>
Supplementary Movie 2. Penetration of the vegetative nucleus and sperm cells through a 1 &#181;m gap. Time-lapse bright field and fluorescence images were captured every 20 s using an inverted fluorescence microscope. Scale bar, 20 &#181;m. <a href="/files/ftp_upload/57262/movie2_3fps.avi">Please click here to download this file.</a>

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 San Sebastian

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

A Microfluidic Device with Groove Patterns for Studying Cellular Behavior

We describe a microfluidic device with microgrooved patterns for studying cellular behavior. This microfluidic platform consists of a top fluidic channel and a bottom microgrooved substrate. To fabricate the microgrooved channels, a top poly(dimethylsiloxane) (PDMS) mold containing the impression of the microfluidic channels was aligned and bonded to a microgrooved substrate. Using this device, mouse fibroblast cells were immobilized and patterned within microgrooved substrates (25, 50, 75, and 100 &mu;m wide). To study apoptosis in a microfluidic device, media containing hydrogen peroxide, Annexin V, and propidium iodide was perfused into the fluidic channel for 2 hours. We found that cells exposed to the oxidative stress became apoptotic. These apoptotic cells were confirmed by Annexin V that bound to phosphatidylserine at the outer leaflet of the plasma membrane during the apoptosis process. Using this microfluidic device with microgrooved patterns, the apoptosis process was observed in real-time and analyzed by using an inverted microscope containing an incubation chamber (37&deg;C, 5% CO2). Therefore, this microfluidic device incorporated with microgrooved substrates could be useful for studying the cellular behavior and performing high-throughput drug screening.

We describe a protocol for the fabrication of microfluidic devices that can enable cell capture and culture. In this approach patterned microstructures such as grooves within microfluidic channels are used to create low shear stress regions within which cell can dock.

We describe a microfluidic device with microgrooved patterns for studying cellular behavior.  This microfluidic platform consists of a top fluidic channel ...

Biology

A Microfluidic Device for Studying Multiple Distinct Strains

The study of cell responses to environmental changes poses many experimental challenges: cells need to be imaged under changing conditions, often in a comparative manner. Multiwell plates are routinely used to compare many different strains or cell lines, but allow limited control over the environment dynamics. Microfluidic devices, on the other hand, allow exquisite dynamic control over the surrounding conditions, but it is challenging to image and distinguish more than a few strains in them. Here we describe a method to easily and rapidly manufacture a microfluidic device capable of applying dynamically changing conditions to multiple distinct yeast strains in one channel. The device is designed and manufactured by simple means without the need for soft lithography. It is composed of a Y-shaped flow channel attached to a second layer harboring microwells. The strains are placed in separate microwells, and imaged under the exact same dynamic conditions. We demonstrate the use of the device for measuring protein localization responses to pulses of nutrient changes in different yeast strains.

We present a simple method to produce microfluidic devices capable of applying similar dynamic conditions to multiple distinct strains, without the need for a clean room or soft lithography.

The study of cell responses to environmental changes poses many experimental challenges: cells need to be imaged under changing conditions, often in a ...

Time-lapse Fluorescence Imaging of Arabidopsis Root Growth with Rapid Manipulation of The Root Environment Using The RootChip

The root functions as the physical anchor of the plant and is the organ responsible for uptake of water and mineral nutrients such as nitrogen, phosphorus, sulfate and trace elements that plants acquire from the soil. If we want to develop sustainable approaches to producing high crop yield, we need to better understand how the root develops, takes up a wide spectrum of nutrients, and interacts with symbiotic and pathogenic organisms. To accomplish these goals, we need to be able to explore roots in microscopic detail over time periods ranging from minutes to days.
We developed the RootChip, a polydimethylsiloxane (PDMS)- based microfluidic device, which allows us to grow and image roots from Arabidopsis seedlings while avoiding any physical stress to roots during preparation for imaging1 (Figure 1). The device contains a bifurcated channel structure featuring micromechanical valves to guide the fluid flow from solution inlets to each of the eight observation chambers2. This perfusion system allows the root microenvironment to be controlled and modified with precision and speed. The volume of the chambers is approximately 400 nl, thus requiring only minimal amounts of test solution.
Here we provide a detailed protocol for studying root biology on the RootChip using imaging-based approaches with real time resolution. Roots can be analyzed over several days using time lapse microscopy. Roots can be perfused with nutrient solutions or inhibitors, and up to eight seedlings can be analyzed in parallel. This system has the potential for a wide range of applications, including analysis of root growth in the presence or absence of chemicals, fluorescence-based analysis of gene expression, and the analysis of biosensors, e.g. FRET nanosensors3.

This article provides a protocol for cultivation of Arabidopsis seedlings in the RootChip, a microfluidic imaging platform that combines automated control of growth conditions with microscopic root monitoring and FRET-based measurement of intracellular metabolite levels.

The root functions as the physical anchor of the plant and is the organ responsible for uptake of water and mineral nutrients such as nitrogen, ...

Microfluidic Picoliter Bioreactor for Microbial Single-cell Analysis: Fabrication, System Setup, and Operation

In this protocol the fabrication, experimental setup and basic operation of the recently introduced microfluidic picoliter bioreactor (PLBR) is described in detail. The PLBR can be utilized for the analysis of single bacteria and microcolonies to investigate biotechnological and microbiological related questions concerning, e.g.&#160;cell growth, morphology, stress response, and metabolite or protein production on single-cell level. The device features continuous media flow enabling constant environmental conditions for perturbation studies, but in addition allows fast medium changes as well as oscillating conditions to mimic any desired environmental situation. To fabricate the single use devices, a silicon wafer containing sub micrometer sized SU-8 structures served as the replication mold for rapid polydimethylsiloxane casting. Chips were cut, assembled, connected, and set up onto a high resolution and fully automated microscope suited for time-lapse imaging, a powerful tool for spatio-temporal cell analysis. Here, the biotechnological platform organism Corynebacterium glutamicum&#160;was seeded into the PLBR and cell growth and intracellular fluorescence were followed over several hours unraveling time dependent population heterogeneity on single-cell level, not possible with conventional analysis methods such as flow cytometry. Besides insights into device fabrication, furthermore, the preparation of the preculture, loading, trapping of bacteria, and the PLBR cultivation of single cells and colonies is demonstrated. These devices will add a new dimension in microbiological research to analyze time dependent phenomena of single bacteria under tight environmental control. Due to the simple and relatively short fabrication process the technology can be easily adapted at any microfluidics lab and simply tailored towards specific needs.

In this protocol the fabrication, setup and basic operation of a microfluidic picoliter bioreactor (PLBR) for single-cell analysis of prokaryotic microorganisms is introduced. Industrially relevant microorganisms were analyzed as proof of principle allowing insights into growth rate, morphology, and phenotypic heterogeneity over certain time periods, hardly possible with conventional methods.

In this protocol the fabrication, experimental setup and basic operation of the recently introduced microfluidic picoliter bioreactor (PLBR) is described ...

A Simple Chamber for Long-term Confocal Imaging of Root and Hypocotyl Development

Several aspects of plant development, such as lateral root morphogenesis, occur on time spans of several days. To study underlying cellular and subcellular processes, high resolution time-lapse microscopy strategies that preserve physiological conditions are required. Plant tissues must have adequate nutrient and water supply with sustained gaseous exchange but, when submerged and immobilized under a coverslip, they are particularly susceptible to anoxia. One strategy that has been successfully employed is the use of a perfusion system to maintain a constant supply of oxygen and nutrients. However, such arrangements can be complicated, cumbersome, and require specialized equipment. Presented here is an alternative strategy for a simple imaging system using perfluorodecalin as an immersion medium. This system is easy to set up, requires minimal equipment, and is easily mounted on a microscope stage, allowing several imaging chambers to be set up and imaged in parallel. In this system, lateral root growth rates are indistinguishable from growth rates under standard conditions on agar plates for the first two days, and lateral root growth continues at reduced rates for at least another day. Plant tissues are supplied with nutrients via an agar slab that can be used also to administer a range of pharmacological compounds. The system was established to monitor lateral root development but is readily adaptable to image other plant organs such as hypocotyls and primary roots.

Presented here is a simple technique for high-resolution confocal time-lapse imaging of root and hypocotyl development for up to 3 days using high numerical-aperture objectives and perfluorodecalin as an immersion medium.

Several aspects of plant development, such as lateral root morphogenesis, occur on time spans of several days. To study underlying cellular and ...

Developmental Biology

A Microfluidic Technique to Probe Cell Deformability

Here we detail the design, fabrication, and use of a microfluidic device to evaluate the deformability of a large number of individual cells in an efficient manner. Typically, data for ~102 cells can be acquired within a 1 hr experiment. An automated image analysis program enables efficient post-experiment analysis of image data, enabling processing to be complete within a few hours. Our device geometry is unique in that cells must deform through a series of micron-scale constrictions, thereby enabling the initial deformation and time-dependent relaxation of individual cells to be assayed. The applicability of this method to human promyelocytic leukemia (HL-60) cells is demonstrated. Driving cells to deform through micron-scale constrictions using pressure-driven flow, we observe that human promyelocytic (HL-60) cells momentarily occlude the first constriction for a median time of 9.3 msec before passaging more quickly through the subsequent constrictions with a median transit time of 4.0 msec per constriction. By contrast, all-trans retinoic acid-treated (neutrophil-type) HL-60 cells occlude the first constriction for only 4.3 msec before passaging through the subsequent constrictions with a median transit time of 3.3 msec. This method can provide insight into the viscoelastic nature of cells, and ultimately reveal the molecular origins of this behavior.

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

Here we detail the design, fabrication, and use of a microfluidic device to evaluate the deformability of a large number of individual cells in an ...

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

Microfluidic devices are capable of creating a precise and controllable cellular micro-environment of pH, temperature, salt concentration, and other physical or chemical stimuli. They have been commonly used for in vitro cell studies by providing in vivo like surroundings. Especially, how cells response to chemical gradients, electrical fields, and shear stresses has drawn many interests since these phenomena are important in understanding cellular properties and functions. These microfluidic chips can be made of glass substrates, silicon wafers, polydimethylsiloxane (PDMS) polymers, polymethylmethacrylate (PMMA) substrates, or polyethyleneterephthalate (PET) substrates. Out of these materials, PMMA substrates are cheap and can be easily processed using laser ablation and writing. Although a few microfluidic devices have been designed and fabricated for generating multiple, coexisting chemical and electrical stimuli, none of them was considered efficient enough in reducing experimental repeats, particular for screening purposes. In this report, we describe our design and fabrication of two PMMA-based microfluidic chips for investigating cellular responses, in the production of reactive oxygen species and the migration, under single or coexisting chemical/electrical/shear stress stimuli. The first chip generates five relative concentrations of 0, 1/8, 1/2, 7/8, and 1 in the culture regions, together with a shear stress gradient produced inside each of these areas. The second chip generates the same relative concentrations, but with five different electric field strengths created within each culture area. These devices not only provide cells with a precise, controllable micro-environment but also greatly increase the experimental throughput.

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.

Microfluidic devices are capable of creating a precise and controllable cellular micro-environment of pH, temperature, salt concentration, and other ...

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 ...

Microfluidic Model to Mimic Initial Event of Neovascularization

Neovascularization is usually initialized from an existing normal vasculature and the biomechanical microenvironment of endothelial cells (ECs) in the initial stage varies dramatically from the following process of neovascularization. Although there are plenty of models to simulate different stages of neovascularization, an in vitro 3D model that capitulates the initial process of neovascularization under the corresponding stimulations of normal vasculature microenvironments is still lacking. Here, we reconstructed an in vitro 3D model that mimics the initial event of neovascularization (MIEN). The MIEN model contains a microfluidic sprouting chip and an automatic control, highly efficient circulation system. A functional, perfusable microchannel coated with endothelium was formed and the process of sprouting was simulated in the microfluidic sprouting chip. The initially physiological microenvironment of neovascularization was recapitulated with the microfluidic control system, by which ECs would be exposed to high luminal shear stress, physiological transendothelial flow, and various vascular endothelial growth factor (VEGF) distributions simultaneously. The MIEN model can be readily applied to the study of neovascularization mechanism and holds a potential promise as a low-cost platform for drug screening and toxicology applications.

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.

Neovascularization is usually initialized from an existing normal vasculature and the biomechanical microenvironment of endothelial cells (ECs) in the ...

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

Filamentous fungi are successful inhabitants of soil and play a major role in soil ecosystems, such as in the decomposition of organic and inorganic matter, as well as regulation of nutrient levels. There they also find numerous opportunities to interact with a variety of other microbes such as bacteria or other fungi. Studying fungal interactions at the cellular level, however, can be challenging owing to the black box-like nature of soil. New microfluidic tools are being developed for the study of fungal interactions; two platforms designed to study bacterial-fungal and fungal-fungal interactions are highlighted. Within these microchannels, fungal-microbial interactions can be monitored in controlled physico-chemical environments at higher temporal and spatial resolution than previously possible. Application of these tools have yielded numerous novel biological insights, such as the observation of bacterial polar attachment to hyphae or revealing uncharacterised fungal-fungal antagonisms. A key feature of these methodologies regards the ease of use of this tool by non-experts, yielding highly translatable technologies for use in microbiology labs.

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.

Filamentous fungi are successful inhabitants of soil and play a major role in soil ecosystems, such as in the decomposition of organic and inorganic ...

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

Summary

Explore More Videos

Chapters in this video

A Microfluidic Device with Groove Patterns for Studying Cellular Behavior

A Microfluidic Device for Studying Multiple Distinct Strains

Time-lapse Fluorescence Imaging of Arabidopsis Root Growth with Rapid Manipulation of The Root Environment Using The RootChip

Microfluidic Picoliter Bioreactor for Microbial Single-cell Analysis: Fabrication, System Setup, and Operation

A Simple Chamber for Long-term Confocal Imaging of Root and Hypocotyl Development

A Microfluidic Technique to Probe Cell Deformability

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

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

Microfluidic Model to Mimic Initial Event of Neovascularization

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