This manuscript has been authored by UT-Battelle, LLC under Contract No. DE-AC05-00OR22725 with the U.S. Department of Energy. The United States Government retains and the publisher, by accepting the article for publication, acknowledges that the United States Government retains a non-exclusive, paid-up, irrevocable, world-wide license to publish or reproduce the published form of this manuscript, or allow others to do so, for United States Government purposes. The Department of Energy will provide public access to these results of federally sponsored research in accordance with the DOE Public Access Plan (http://energy.gov/downloads/doe-public-access-plan).
This work was supported in part by the Genomic Science Program, U.S. Department of Energy, Office of Science, Biological and Environmental Research, as part of the Plant Microbe Interfaces Scientific Focus Area (http://pmi.ornl.gov). The fabrication of the microfluidic platforms was carried out in the Nanofabrication Research Laboratory at the Center for Nanophase Materials Sciences, which is a DOE Office of Science User Facility. JAA is supported by an NSF graduate research fellowship DGE -1452154

1. Two-layer Platform Fabrication
<ol>
	<li>Fabrication of multilayer masters
	<ol>
		<li>Spin coat epoxy-based negative photoresist (~63.45% solids, 1,250 cSt) according to the manufacturer&#39;s specifications (2,000 rpm for 45 s) onto a 4 inch diameter silicon wafer to obtain the desired height of 20 &#181;m for the first design layer.</li>
		<li>Soft-bake the resist coated wafers for 4 min at 95 &#176;C. Allow wafer to cool for 5 min. Expose the wafer to UV light for 15 s (~150 mJ/cm2 at 365 nm) through a photomask in a UV contact aligner to define the geometry of the bottom layer.</li>
		<li>Post-exposure bake the wafer for 5 min at 95 &#176;C. Without developing, return the wafer to the spin coater and spin on a second layer of epoxy-based negative photoresist (~76.75% solids, 80,000 cSt) at 3,000 rpm to achieve a second layer height of 150 - 200 &#181;m.</li>
		<li>Allow the wafer to rest for 5 min before transferring to a 95 &#176;C hotplate for 45 min. After baking on the hotplate, remove the wafer and allow the resist to cool and harden at room temperature for another 5 min on a flat level surface. Align and expose the wafer to the second layer photomask for 30 s in a UV contact aligner (dose of approximately 300 mJ/cm2).</li>
		<li>Perform a post-exposure bake by placing the wafer on a hotplate for 15 min at 95 &#176;C, and then allow the wafer to cool on a flat level surface for 5 min before developing.</li>
		<li>Develop both resist layers simultaneously by placing the wafer into a plastic dish and submerging the wafer in the appropriate developer (see Materials Table). Gently rock the dish occasionally to wash fresh developer over the wafer.</li>
		<li>After 17 min, rinse the wafer with isopropanol (IPA). If a white film appears, continue to iterate between rinsing the wafer with developer and IPA until the film disappears. Dry the patterned silicon wafer with nitrogen.</li>
	</ol>
	</li>
	<li>Polydimethylsiloxane Soft-lithography
	<ol>
		<li>Expose the silicon wafer to an air plasma for 30 s in a plasma cleaner set on high (see Materials Table) to clean. Silanize the wafer with trichloro (1H,1H,2H,2H-perfluoro-n-octyl) silane in a chemical hood on a hotplate below the flashpoint of the silane (85 &#176;C) for 2 h.</li>
		<li>Pour a 10:1 ratio of polydimethylsiloxane (PDMS) to curing agent onto the silicon master wafer. Degas the mixed PDMS in a vacuum chamber and then cure the polymer in a 70 &#176;C oven for 1 h or until the PDMS is fully cured.</li>
		<li>Using a scalpel, cut the PDMS devices and peel them from the silicon master wafer. Use a 1.5 mm biopsy punch to create seed and treatment inlets, punching the seed inlet at a 45&#176; angle to encourage root growth into the main channel.</li>
		<li>Use clear adhesive tape to remove debris from the PDMS device and place the device design side down on a glass coverslip. Autoclave the assembled devices.</li>
	</ol>
	</li>
</ol>
2. Planting Devices
<ol>
	<li>Arabidopsis thaliana seed preparation
	<ol>
		<li>Surface sterilize A. thaliana seeds in a microfuge tube with a solution of 30% commercially available bleach diluted in de-ionized water and 0.1% Triton X detergent for 7 min. Wash seeds 4 times with sterile water.</li>
		<li>Stratify seeds by refrigerating microfuge tube overnight or up to a week at 4 &#176;C.</li>
	</ol>
	</li>
	<li>Device Preparation
	<ol>
		<li>Place sterilized devices in a vacuum degassing chamber to remove air from the gas permeable PDMS and fluidic channels to improve ease of filling.</li>
		<li>Remove the devices from the vacuum chamber and immediately submerge the devices in a Petri dish filled with liquid &#188; strength plant based media at pH 5.7.</li>
		<li>Using a pipette, pull liquid through the inlets to fill the device. Make sure no air bubbles remain within channels by visual inspection.</li>
		<li>Transfer individual devices to new dry Petri dishes. Pour or pipette hot agar around the device until the agar level is almost flush with the top of the PDMS device. Allow agar to solidify. 
		NOTE: Devices are now ready for planting seeds or can be stored at 4 &#176;C until needed.</li>
	</ol>
	</li>
	<li>Planting seeds within devices
	<ol>
		<li>In a sterile environment, transfer one sterilized seed to the inlet of each device using a small pipette.</li>
		<li>Cover the Petri dish with wax film (see Materials Table) and place in a light/dark cycling growth chamber or windowsill at room temperature. Orient the Petri dish so that the devices are vertical and gravity will encourage the roots to grow through the channel.</li>
	</ol>
	</li>
</ol>
3. Treatment
<ol>
	<li>Experimental Treatments
	<ol>
		<li>At the desired time in the seedling&#39;s development, add the experimental treatment to the seedling by adding a prescribed amount of the treatment to each of the 8 side ports via pipette.</li>
		<li>Reseal the Petri dish and return to growth chamber or windowsill.</li>
		<li>Proceed with imaging samples at the desired time to capture individual time points or begin time lapse imaging.</li>
	</ol>
	</li>
</ol>
4. Optical Imaging
<ol>
	<li>Lower Resolution (4 - 20X) Imaging
	<ol>
		<li>Place the entire Petri dish containing the device and seedling under an inverted bright field microscope for lower resolution (4 - 20X) bright field or differential interference contrast (DIC) imaging. Optimize lighting conditions to elucidate morphological features of interest by adjusting exposure times, lamp brightness, aperture, and the polarity of light. Drive the stage to the desired area of the root and focus on the root or root hair(s) of interest.</li>
		<li>Acquire a single time point or a time series of images. To visualize the growth of root hairs, image growing root hairs once per min. To visualize the growth of the main root, acquire one image every 30 min. Return the Petri dish and the seedlings to growth chamber or windowsill in a vertical position after imaging is complete.</li>
	</ol>
	</li>
	<li>Higher Resolution (20 - 63X) Imaging
	<ol>
		<li>Once the seedling has grown for a desired time, use forceps to remove coverslip and device from the agar. Clean off the bottom of the coverslip using an ethanol wetted laboratory tissue.</li>
		<li>Apply the recommended immersion media to the objective as suggested by the microscope lens manufacturer. Place the coverslip down on the microscope stage and raise the stage to contact the immersion media on the objective. Optimize the lighting conditions by adjusting exposure times, lamp brightness, aperture, and the polarity of light. Drive the stage to the desired area and focus on root hair(s) of interest. 
		NOTE: DIC is needed to visualize cytoplasmic streaming, and fluorescence markers are needed to visualize organelle movement.</li>
		<li>To quantify root hair growth over time, acquire one image per min. To visualize organelle movement, minimize the fluorescence exposure time while still retaining an identifiable fluorescence signal from the organelles. Acquire images of the organelles as quickly as the exposure time allows. For longer time-lapse imaging, use a live-cell imaging stage incubator to maintain ambient temperature and moisture. 
		NOTE: A 63X oil objective is recommended for imaging root hair organelles.</li>
	</ol>
	</li>
</ol>
5. Non-Optical Imaging
<ol>
	<li>Device preparation
	<ol>
		<li>Once seedling has grown for desired time, remove the device and coverslip from the Petri dish.</li>
		<li>Turn device upside down and gently peel away the coverslip so that the root stays within the PDMS channel.</li>
		<li>Proceed to use the PDMS device as a substrate for the root in higher resolution atomic force or scanning electron microscopy.</li>
	</ol>
	</li>
	<li>Scanning Electron Microscopy
	<ol>
		<li>Deposit a thin (~20 nm) layer of chromium onto the root and surrounding PDMS using a Dual Gun Electron Beam Evaporation Chamber.</li>
		<li>Transfer the root and PDMS device to a scanning electron microscope chamber. 
		NOTE: Imaging conditions, i.e. voltage, current and working distance will need to be optimized to achieve the desired resolution for a given application.</li>
	</ol>
	</li>
	<li>Atomic Force Microscopy (AFM)
	<ol>
		<li>Mount the PDMS device root side up on an AFM specimen holder. To increase contrast in the camera and more easily distinguish the root from the PDMS, place the PDMS device on a flat reflective substrate (such as mica coated in gold) before mounting the device on the AFM specimen holder.</li>
		<li>Secure a liquid well attachment on top of the PDMS device and fill it with water to keep the roots hydrated during imaging.</li>
		<li>Load the specimen holder into the AFM. Adjust the z-control for the thickness of the PDMS device and drive the cantilever to the region of interest on the root using a camera for guidance.</li>
		<li>Align the laser with the tip of the cantilever. For best results with contact mode imaging, use cantilevers with spring constants of 0.01 or 0.03 N/m to exert minimal force on the root during scanning.</li>
		<li>Slowly lower the scanning mechanism until the cantilever just makes contact with the sample. Adjust the scan size for the desired region and choose a scan speed of 1 line/s with 256 voltage points per line. Acquire the scan.</li>
	</ol>
	</li>
</ol>

<ol>
	<li>Pritchard, S. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Soil+organisms+and+global+climate+change.">Soil organisms and global climate change.</a> Plant Pathol. 60 (1), 82-99 (2011).</li><li>Norby, R. J., Ledford, J., Reilly, C. D., Miller, N. E., O'Neill, E. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fine-root+production+dominates+response+of+a+deciduous+forest+to+atmospheric+CO2+enrichment.">Fine-root production dominates response of a deciduous forest to atmospheric CO2 enrichment.</a> Proc. Natll. Acad. Sci. USA. 101 (26), 9689-9693 (2004).</li><li>McCormack, M. L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Redefining+fine+roots+improves+understanding+of+below-ground+contributions+to+terrestrial+biosphere+processes.">Redefining fine roots improves understanding of below-ground contributions to terrestrial biosphere processes.</a> New Phytol. 207 (3), 505-518 (2015).</li><li>Mangano, S., Juarez, S. P. D., Estevez, J. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=ROS+regulation+of+polar-growth+in+plant+cells.">ROS regulation of polar-growth in plant cells.</a> Plant Physiol. 171 (3), 1593-1605 (2016).</li><li>Ketelaar, T., Emons, A. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+Actin+Cytoskeleton+in+Root+Hairs:+A+Cell+Elongation+Device.">The Actin Cytoskeleton in Root Hairs: A Cell Elongation Device.</a> Root Hairs. , 211-232 (2009).</li><li>Grossmann, G., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+RootChip:+an+integrated+microfluidic+chip+for+plant+science.">The RootChip: an integrated microfluidic chip for plant science.</a> Plant Cell. 23 (12), 4234-4240 (2011).</li><li>Grossmann, G., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Time-lapse+fluorescence+imaging+of+Arabidopsis+root+growth+with+rapid+manipulation+of+the+root+environment+using+the+RootChip.">Time-lapse fluorescence imaging of Arabidopsis root growth with rapid manipulation of the root environment using the RootChip.</a> J. Vis. Exp. (65), (2012).</li><li>Jiang, H., Xu, Z., Aluru, M. R., Dong, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Plant+chip+for+high-throughput+phenotyping+of+Arabidopsis.">Plant chip for high-throughput phenotyping of Arabidopsis.</a> Lab Chip. 14 (7), 1281 (2014).</li><li>Busch, W., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+microfluidic+device+and+computational+platform+for+high-throughput+live+imaging+of+gene+expression.">A microfluidic device and computational platform for high-throughput live imaging of gene expression.</a> Nat. Methods. 9 (11), (2012).</li><li>Meier, M., Lucchettta, E., Ismagilov, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Chemical+Stimulation+of+the+Arabidopsis+thaliana+Root+using+Multi-Laminar+Flow+on+a+Microfluidic+Chip.">Chemical Stimulation of the Arabidopsis thaliana Root using Multi-Laminar Flow on a Microfluidic Chip.</a> Lab Chip. 10 (16), 2147-2153 (2010).</li><li>Ozoe, K., Hida, H., Kanno, I., Higashiyama, T., Notaguchi, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Early+characterization+method+of+plant+root+adaptability+to+soil+environments.">Early characterization method of plant root adaptability to soil environments.</a> Proc. of 28th IEEE Interntl. Conf. Micro. Electro Mech. Syst. , (2015).</li><li>Sanati Nezhad, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Microfluidic+platforms+for+plant+cells+studies.">Microfluidic platforms for plant cells studies.</a> Lab on a chip. , 3262-3274 (2014).</li><li>Parashar, A., Pandey, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Plant-in-chip:+Microfluidic+system+for+studying+root+growth+and+pathogenic+interactions+in+Arabidopsis.">Plant-in-chip: Microfluidic system for studying root growth and pathogenic interactions in Arabidopsis.</a> App. Phys. Lett. 98 (26), 2009-2012 (2011).</li><li>Rigas, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Root+gravitropism+and+root+hair+development+constitute+coupled+developmental+responses+regulated+by+auxin+homeostasis+in+the+Arabidopsis+root+apex.">Root gravitropism and root hair development constitute coupled developmental responses regulated by auxin homeostasis in the Arabidopsis root apex.</a> New Phytolol. 197 (4), 1130-1141 (2013).</li><li>Bengough, A. G., McKenzie, B. M., Hallett, P. D., Valentine, T. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Root+elongation,+water+stress,+and+mechanical+impedance:+A+review+of+limiting+stresses+and+beneficial+root+tip+traits.">Root elongation, water stress, and mechanical impedance: A review of limiting stresses and beneficial root tip traits.</a> J. Exp. Bot. 62 (1), 59-68 (2011).</li><li>Sia, S. K., Whitesides, G. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Microfluidic+devices+fabricated+in+poly(dimethylsiloxane)+for+biological+studies.">Microfluidic devices fabricated in poly(dimethylsiloxane) for biological studies.</a> Electrophor. 24 (21), 3563-3576 (2003).</li><li>Millet, L. J., Stewart, M. E., Sweedler, J. V., Nuzzo, R. G., Gillette, M. U. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Microfluidic+devices+for+culturing+primary+mammalian+neurons+at+low+densities.">Microfluidic devices for culturing primary mammalian neurons at low densities.</a> Lab chip. 7 (8), 987-994 (2007).</li><li>Nelson, B. K., Cai, X., Nebenf&#252;hr, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+multicolored+set+of+in+vivo+organelle+markers+for+co-localization+studies+in+Arabidopsis+and+other+plants.">A multicolored set of in vivo organelle markers for co-localization studies in Arabidopsis and other plants.</a> Plant J. 51 (6), 1126-1136 (2007).</li><li>Talbot, M. J., White, R. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cell+surface+and+cell+outline+imaging+in+plant+tissues+using+the+backscattered+electron+detector+in+a+variable+pressure+scanning+electron+microscope.">Cell surface and cell outline imaging in plant tissues using the backscattered electron detector in a variable pressure scanning electron microscope.</a> Plant Methods. 9 (1), 40 (2013).</li></ol>

The authors have nothing to disclose.

The method described in this article for creating a plant-on-a-chip platform is unique in that the two-layer design confines the root hairs to a single imaging plane and the platform may be deconstructed and used as a substrate for high resolution non-optical imaging. Using high-resolution non-optical imaging can provide valuable information about the plant tissue that could not be obtained from optical imaging alone. For example, AFM imaging can provide force measurements to calculate the elasticity of root tissues duri...

Fine root features increase water and nutrient acquisition for the plant, exploring new soil spaces and increasing the total root surface area. The turnover of these fine root features plays a major role in stimulating the underground food chain1 and the number of fine roots in certain plant species is expected to double under elevated atmospheric carbon dioxide2. Fine roots are generally defined as those smaller than 2 mm in diameter, although new definitions advocate for characterizing fine roots by their function3. Like many fine roots, root hairs provide the function of uptake and absorption but occupy a much smaller space with diameters on the order of microns. Because of their small size, root hairs are difficult to image in situ and are often overlooked as a part of the overall root architecture in field scale experiments and models.
Ex terra root hair studies, such as from seedlings grown on agar plates, have provided the scientific community with valuable information on cellular growth and transport4,5. While agar plates allow root systems to be imaged non-destructively and in real time, they do not offer high environmental control for the addition of experimental treatments such as nutrients, plant hormones, or bacteria. An emerging solution to facilitating high resolution imaging while also affording dynamic environmental control has been the advent of microfluidic platforms for plant studies. These platforms have enabled the non-destructive growth and visualization of several plant species for high throughput phenotyping6,7,8,9, isolated chemical treatments10, force measurements11,12, and the addition of microorganisms13. Microfluidic platform designs have focused on the use of single open space fluidic layers in which the roots may propagate, permitting the root hairs to drift in and out of optical focus during growth or treatment.
Here we present a procedure for developing a two-layer microfluidic platform using photo and soft-lithography methods that builds upon previous plant-on-a-chip designs by confining the seedling root hairs to the same imaging plane as the main root. This allows us to track root hair development in real time, at high resolution, and throughout the experimental treatment process. Our culturing methods allow Arabidopsis thaliana seedlings to be germinated from seed within the platform and cultured for up to a week in a hydrated and sterile environment that does not require the use of syringe pump equipment. Once the time-lapse imaging experiment has concluded, the platform presented here can be opened without disturbing the position of the finer root features. This allows the use of other high resolution imaging methods. Here we provide representative results for the quantification and visualization of root hair morphology in this platform by optical, scanning electron microscopy (SEM), and atomic force microscopy techniques (AFM).

<table><tbody><tr><td>Silicon Wafer</td><td>WRS Materials</td><td>100mm diameter, 500-550um thickness, Prime, 10-20 resistivity, N/Phos&amp;lt;100&gt;</td><td></td></tr><tr><td>Quintel Contact Aligner</td><td>Neutronix Quintel Corp</td><td>NXQ 7500 Mask Aligner</td><td></td></tr><tr><td>Fluorescent Microscope</td><td>Nikon</td><td>Eclipse Ti-U</td><td></td></tr><tr><td>laboratory tissue</td><td>Kimberly Clark</td><td>Kimwipe KIMTECH SCIENCE Brand, 34155</td><td></td></tr><tr><td>Negative Photoresist Epoxy</td><td>Microchem</td><td>SU-8 2000s series</td><td></td></tr><tr><td>Photoresist developer</td><td>Microchem</td><td>Su-8 developer</td><td></td></tr><tr><td>trichloro(1H,1H,2H,2H-perfluoro-octyl)silane</td><td>Sigma Aldrich</td><td></td><td>use in chemical hood</td></tr><tr><td>Air Plasma Cleaner</td><td>Harrick Plasma</td><td></td><td></td></tr><tr><td>PDMS</td><td>Dow Corning</td><td>Sylgard 184 Silicone elastomer base</td><td></td></tr><tr><td>PDMS curing agent</td><td>Dow Corning</td><td>Sylgard 184 Silicone elastomer curing agent</td><td></td></tr><tr><td>Dessicator</td><td>Bel-Art</td><td>F42010-000</td><td></td></tr><tr><td>Scalpel</td><td>X-acto knife</td><td></td><td></td></tr><tr><td>Biopsy Punch</td><td>Ted Pella</td><td>15110-15</td><td></td></tr><tr><td>Adhesive tape</td><td>Staples</td><td>Invisible Tape</td><td></td></tr><tr><td>Microfuge tube</td><td>Eppendorf</td><td></td><td></td></tr><tr><td>Triton X</td><td>J.T.Baker XI98-07</td><td></td><td></td></tr><tr><td>Bleach</td><td>Chlorox</td><td>concentrated</td><td></td></tr><tr><td>Plant-Based Media</td><td>Phyto Technology Laboratories</td><td>M524</td><td></td></tr><tr><td>Agar</td><td>Teknova</td><td>A7777</td><td></td></tr><tr><td>Wax film</td><td>Parafilm</td><td></td><td></td></tr><tr><td>microscope</td><td>Olympus</td><td>IX51</td><td></td></tr><tr><td>Atomic Force Microscope</td><td>Keysight Technologies</td><td>5500 PicoPlus AFM</td><td></td></tr><tr><td>Petri dish</td><td>VWR</td><td></td><td></td></tr><tr><td>Scanning Electron Microscope</td><td>JEOL</td><td>7400</td><td></td></tr><tr><td>Dual Gun Electron Beam Evaporator</td><td>Thermionics</td><td>Custom Dual Electron Gun Evaporation System</td><td></td></tr></tbody></table>

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.

The two-layer PDMS microfluidic devices described here have a 200 &#181;m high channel for the main Arabidopsis root and a 20 &#181;m high chamber to confine laterally growing root hairs (Figure 1A). This design may be used for plant species with similar root diameters as Arabidopsis thaliana and can be readily modified to accommodate species of different sizes. The design incorporates an inlet for the plant as well as 8 side inlets...

Watch this Scientific Journal Video about Imaging the Root Hair Morphology of Arabidopsis Seedlings in a Two-layer Microfluidic Platform at JoVE.com

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.

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 ...

imaging-root-hair-morphology-arabidopsis-seedlings-two-layer

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Bioengineering

8.5K Views.  University of Tennessee. Dit artikel laat zien hoe cultuur Arabidopsis thaliana zaailingen in een twee-laag microfluidic platform dat de belangrijkste wortel en root haren op een interne optische vlak beperkt. Dit platform kan worden gebruikt voor real-time optische beeldvorming van fijne wortel morfologie alsook wat betreft hoge resolutie beeldvorming door andere middelen.

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

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, ...

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

Atomic Force Microscopy to Study the Physical Properties of Epidermal Cells of Live Arabidopsis Roots

A method is described here to characterize the physical properties of the cell wall of epidermal cells of living Arabidopsis roots through nanoindentations with an atomic force microscope (AFM) coupled with an optical inverted fluorescence microscope. The method consists of applying controlled forces to the sample while measuring its deformation, allowing quantifying parameters such as the apparent Young's modulus of cell walls at subcellular resolutions. It requires a careful mechanical immobilization of the sample and correct selection of indenters and indentation depths. Although it can be used only in external tissues, this method allows characterizing mechanical changes in plant cell walls during development and enables the correlation of these microscopic changes with the growth of an entire organ.

Atomic Force Microscopy to Study the Physical Properties of Epidermal Cells of Live Arabidopsis Roots

The atomic force microscopy indentation protocol offers the possibility to dissect the role of the physical properties of the cell wall of a particular cell of a tissue or organ during normal or constrained growth (i.e., under water deficit).

A method is described here to characterize the physical properties of the cell wall of epidermal cells of living Arabidopsis roots through ...

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.

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 ...

A Simple Method for Imaging Arabidopsis Leaves Using Perfluorodecalin as an Infiltrative Imaging Medium

The problem of acquiring high-resolution images deep into biological samples is widely acknowledged1. In air-filled tissue such as the spongy mesophyll of plant leaves or vertebrate lungs further difficulties arise from multiple transitions in refractive index between cellular components, between cells and airspaces and between the biological tissue and the rest of the optical system. Moreover, refractive index mismatches lead to attenuation of fluorophore excitation and signal emission in fluorescence microscopy. We describe here the application of the perfluorocarbon, perfluorodecalin (PFD), as an infiltrative imaging medium which optically improves laser scanning confocal microscopy (LSCM) sample imaging at depth, without resorting to damaging increases in laser power and has minimal physiological impact2. We describe the protocol for use of PFD with Arabidopsis thaliana leaf tissue, which is optically complex as a result of its structure (Figure 1). PFD has a number of attributes that make it suitable for this use3. The refractive index of PFD (1.313) is comparable with that of water (1.333) and is closer to that of cytosol (approx. 1.4) than air (1.000). In addition, PFD is readily available, non-fluorescent and is non-toxic. The low surface tension of PFD (19 dynes cm-1) is lower than that of water (72 dynes cm-1) and also below the limit (25 - 30 dyne cm-1) for stomatal penetration4, which allows it to flood the spongy mesophyll airspaces without the application of a potentially destructive vacuum or surfactant. Finally and crucially, PFD has a great capacity for dissolving CO2 and O2, which allows gas exchange to be maintained in the flooded tissue, thus minimizing the physiological impact on the sample. These properties have been used in various applications which include partial liquid breathing and lung inflation5,6, surgery7, artificial blood8, oxygenation of growth media9, and studies of ice crystal formation in plants10. Currently, it is common to mount tissue in water or aqueous buffer for live confocal imaging. We consider that the use of PFD as a mounting medium represents an improvement on existing practice and allows the simple preparation of live whole leaf samples for imaging.

A Simple Method for Imaging Arabidopsis Leaves Using Perfluorodecalin as an Infiltrative Imaging Medium

We describe the use of perfluorodecalin as an infiltrative mounting medium. This is a simple method for improving depth of imaging in Arabidopsis thaliana leaf tissue with minimal physiological impact.

The problem of acquiring high-resolution images deep into biological samples is widely acknowledged1. In air-filled tissue such as the spongy mesophyll of ...

Biology

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 ...

Long-term, High-resolution Confocal Time Lapse Imaging of Arabidopsis Cotyledon Epidermis during Germination

Imaging in vivo dynamics of cellular behavior throughout a developmental sequence can be a powerful technique for understanding the mechanics of tissue patterning. During animal development, key cell proliferation and patterning events occur very quickly. For instance, in Caenorhabditis elegans all cell divisions required for the larval body plan are completed within six hours after fertilization, with seven mitotic cycles1; the sixteen or more mitoses of Drosophila embryogenesis occur in less than 24 hr2. In contrast, cell divisions during plant development are slow, typically on the order of a day 3,4,5 . This imposes a unique challenge and a need for long-term live imaging for documenting dynamic behaviors of cell division and differentiation events during plant organogenesis. Arabidopsis epidermis is an excellent model system for investigating signaling, cell fate, and development in plants. In the cotyledon, this tissue consists of air- and water-resistant pavement cells interspersed with evenly distributed stomata, valves that open and close to control gas exchange and water loss. Proper spacing of these stomata is critical to their function, and their development follows a sequence of asymmetric division and cell differentiation steps to produce the organized epidermis (Fig. 1).
This protocol allows observation of cells and proteins in the epidermis over several days of development. This time frame enables precise documentation of stem-cell divisions and differentiation of epidermal cells, including stomata and epidermal pavement cells. Fluorescent proteins can be fused to proteins of interest to assess their dynamics during cell division and differentiation processes. This technique allows us to understand the localization of a novel protein, POLAR6, during the proliferation stage of stomatal-lineage cells in the Arabidopsis cotyledon epidermis, where it is expressed in cells preceding asymmetric division events and moves to a characteristic area of the cell cortex shortly before division occurs. Images can be registered and streamlined video easily produced using public domain software to visualize dynamic protein localization and cell types as they change over time.

Long-term, High-resolution Confocal Time Lapse Imaging of Arabidopsis Cotyledon Epidermis during Germination

We describe a protocol using chamber slides and media to immobilize plant cotyledons for confocal imaging of the epidermis over several days of development, documenting stomatal differentiation. Fluorophore-tagged proteins can be tracked dynamically by expression and subcellular localization, increasing understanding of their possible roles during cell division and cell-type differentiation.

Imaging in vivo dynamics of cellular behavior throughout a developmental sequence can be a powerful technique for understanding the mechanics of tissue ...

Using Flatbed Scanners to Collect High-resolution Time-lapsed Images of the Arabidopsis Root Gravitropic Response

Research efforts in biology increasingly require use of methodologies that enable high-volume collection of high-resolution data. A challenge laboratories can face is the development and attainment of these methods. Observation of phenotypes in a process of interest is a typical objective of research labs studying gene function and this is often achieved through image capture. A particular process that is amenable to observation using imaging approaches is the corrective growth of a seedling root that has been displaced from alignment with the gravity vector. Imaging platforms used to measure the root gravitropic response can be expensive, relatively low in throughput, and/or labor intensive. These issues have been addressed by developing a high-throughput image capture method using inexpensive, yet high-resolution, flatbed scanners. Using this method, images can be captured every few minutes at 4,800 dpi. The current setup enables collection of 216 individual responses per day. The image data collected is of ample quality for image analysis applications.

This protocol describes a process for rapid collection of images of Arabidopsis seedlings responding to a gravity stimulus using commercially-available flatbed scanners. The method allows for inexpensive, high-volume capture of high-resolution images amenable for downstream analysis algorithms.

Research efforts in biology increasingly require use of methodologies that enable high-volume collection of high-resolution data. A challenge laboratories ...

Measuring Spatial and Temporal Ca2+ Signals in Arabidopsis Plants

Developmental and environmental cues induce Ca2+ fluctuations in plant cells. Stimulus-specific spatial-temporal Ca2+ patterns are sensed by cellular Ca2+ binding proteins that initiate Ca2+ signaling cascades. However, we still know little about how stimulus specific Ca2+ signals are generated. The specificity of a Ca2+ signal may be attributed to the sophisticated regulation of the activities of Ca2+ channels and/or transporters in response to a given stimulus. To identify these cellular components and understand their functions, it is crucial to use systems that allow a sensitive and robust recording of Ca2+ signals at both the tissue and cellular levels. Genetically encoded Ca2+ indicators that are targeted to different cellular compartments have provided a platform for live cell confocal imaging of cellular Ca2+ signals. Here we describe instructions for the use of two Ca2+ detection systems: aequorin based FAS (film adhesive seedlings) luminescence Ca2+ imaging and case12 based live cell confocal fluorescence Ca2+ imaging. Luminescence imaging using the FAS system provides a simple, robust and sensitive detection of spatial and temporal Ca2+ signals at the tissue level, while live cell confocal imaging using Case12 provides simultaneous detection of cytosolic and nuclear Ca2+ signals at a high resolution.

Measuring Spatial and Temporal Ca2+ Signals in Arabidopsis Plants

Ca2+ signaling regulates diverse biological processes in plants. Here we present approaches for monitoring abiotic stress induced spatial and temporal Ca2+ signals in Arabidopsis cells and tissues using the genetically encoded Ca2+ indicators Aequorin or Case12.

Developmental and environmental cues induce Ca2+ fluctuations in plant cells. Stimulus-specific spatial-temporal Ca2+ patterns are sensed by cellular Ca2+ ...

Light Sheet Fluorescence Microscopy of Plant Roots Growing on the Surface of a Gel

One of the key questions in understanding plant development is how single cells behave in a larger context of the tissue. Therefore, it requires the observation of the whole organ with a high spatial- as well as temporal resolution over prolonged periods of time, which may cause photo-toxic effects. This protocol shows a plant sample preparation method for light-sheet microscopy, which is characterized by mounting the plant vertically on the surface of a gel. The plant is mounted in such a way that the roots are submerged in a liquid medium while the leaves remain in the air. In order to ensure photosynthetic activity of the plant, a custom-made lighting system illuminates the leaves. To keep the roots in darkness the water surface is covered with sheets of black plastic foil. This method allows long-term imaging of plant organ development in standardized conditions.

This protocol shows a plant sample preparation method for light-sheet microscopy. The setup is characterized by mounting the plant vertically on the surface of a gel and letting it grow in controlled bright conditions. This allows long-term observation of plant organ development in standardized conditions.

One of the key questions in understanding plant development is how single cells behave in a larger context of the tissue. Therefore, it requires the ...

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

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