We thank Maty&#225;&#353; Fendrych for critical reading/viewing and Stephan Stadlbauer for the audio equipment. Thanks to the Miba Machine Shop at IST Austria for their contribution to the OpenSPIM. The research leading to these results has received funding from the People Programme (Marie Curie Actions) of the European Union&#39;s Seventh Framework Programme (FP7/2007-2013) under REA grant agreement n&#176; [291734] and European Research Council (project ERC-2011-StG-20101109-PSDP).

1. Arabidopsis Culturing Prior to Imaging
<ol>
	<li>Prepare &#189; MS medium (half-strength Murashige and Skoog medium) by adding 2.15 g MS-medium, 10 g sucrose, 0.97 g MES (2-(N-morpholino)ethanesulfonic acid) and 1 L ddH2O (double-distilled water) into a 1 L bottle. Adjust the pH to 5.8 using KOH.</li>
	<li>Add 15 g/L gellan gum to the &#189; MS medium and autoclave it for 20 min at 121 &#176;C.</li>
	<li>Pour 30 mL of the hot medium into square Petri dishes (245 x 245 x 25 mm) to create a layer of gel with a thickness of about 2 mm. Let the dishes cool down to room temperature to allow the medium to solidify.</li>
	<li>Put sterilized Arabidopsis seeds into a 1.5 mL reaction tube containing 1 mL sterile H2O. Pick up the seeds using a glass pipet or a 1,000 &#181;L pipet tip and sow them onto the surface of the gel. Place the seeds separately about 10 mm apart. Seal the plate with tape.</li>
	<li>Incubate the plate for 24 h at 4 &#176;C (stratification).</li>
	<li>Cultivate the plate in a growth incubator, e.g. at 22 &#176;C in a 16/8 h day/night cycle with 120-140 &#181;mol/m&#178;/s amount of light for 6 days. Up to 10 days old plants can be used.</li>
</ol>
2. Plant Sample Preparation Method
Note: The sample holder can be either 3D printed or hand made in a machine workshop using the dimensions depicted in Figure 2C. The 3D model file is provided in the supplemental material (Supplemental_File_-_3D_Sample_Holder.stl). See the link for ordering the 3D print in the material list.
<ol>
	<li>Add 5 g low-melt agarose into a 50 mL bottle containing 50 mL &#189; MS medium and autoclave it for 20 min at 121 &#176;C.</li>
	<li>Aliquot the 1% low-melt agarose solution in 1.5 mL reaction tubes. Store at 4 &#176;C (can be used for at least two months).</li>
	<li>Melt an aliquot of 1%% low-melt agarose at 80 &#176;C and let it cool down to 33 &#176;C.</li>
	<li>Clean the sample holder in an ultrasound unit. Sterilize the sample holder with 70% ethanol and wash with sterile water.</li>
	<li>Cut the gel around the plant using a scalpel.</li>
	<li>Lift the block with a flat spatula and slide it carefully on the sample holder using a second spatula.</li>
	<li>Glue the gel on the sample holder with 1% agarose (at 33 &#176;C) using a 100 &#181;L pipette.</li>
	<li>Glue the plant on the gel with 1% agarose using a 10 &#181;L pipette. Use a stereo microscope to verify that the leaves are not covered with gel. Do not position the gel directly on the region of interest.</li>
	<li>To prevent the plant from drying out, work uninterruptedly. Insert the sample holder into a 1,000 &#181;L pipet tip whenever possible. Use the pipet tip as a cap and slide it carefully over one end of the sample holder where the plant is located.</li>
	<li>Put the sample holder in a pipette tip box and prepare more plants if necessary. Plants can be directly imaged or put back in the growth incubator.</li>
</ol>
3. Set up the Microscope
NOTE: The LED illumination system is a custom-built lamp. The technical details required to build the LED-ring can be found in Figure 3 and the material list. See the supplemental material file (Supplemental_File_-_LED_Ring_Board.brd) for the board design.
<ol>
	<li>Screw or glue (e.g. double sided tape) the LED ring on the lower side of the OpenSPIM x/y/z/&#952;-stage arm.</li>
	<li>Connect the LED ring with an adjustable power supply (0-30 V, max 2 A).</li>
	<li>Adjust the voltage to the desired light intensity (for Arabidopsis, 120-140 &#181;mol/m2/s, Figure 3D).</li>
	<li>Sterilize the sample chamber with 70% ethanol and wash with sterile water.</li>
	<li>Clean the objective lens with benzine and lens cleaning tissue. Sterilize the objective lens with 70% ethanol.</li>
	<li>Connect the perfusion tubes to the sample chamber in a one-way arrangement. Put a 1 L bottle containing fresh &#189; MS medium and another empty bottle next to the peristaltic perfusion pump. Connect the bottle with medium with the lower inlet of the sample chamber using one perfusion tube. Connect the upper outlet of the sample chamber with the empty bottle to trash the used medium using another perfusion tube.</li>
	<li>Set the speed of flow to 1 mL/min. 
	NOTE: To not overspill the sample chamber, it is important to have a higher outflow than the inflow. Either increase the pumping rate or use a tube with a larger inner diameter for the outflow.</li>
	<li>Cut a black plastic foil into 3 mm small squares, wash with 70% ethanol and let them dry before placing them in the sample chamber on the water surface.</li>
	<li>Remove the pipet tip from the sample holder and insert the sample holder in the sample chamber. If the newly manufactured sample holder does not fit into the stage arm, use a fine sandpaper to make it thinner or use an O-ring (&#8709; 6 mm) in case it is too thin.</li>
	<li>To create the lids, cut the black aluminum foil into two 50 x 25 mm pieces. Make a 5 mm cut in the middle of one side of each piece. Fold to create a triangle indentation (Figure 1D). Close the sample chamber with the two lids by putting the lids on top of the sample chamber with the triangle indentations facing the sample holder. Make sure that the plant leaves are not in shade of the lids and receive light from the illumination system.</li>
	<li>Find the region of interest using the x/y/z and rotation stage to position an emerging lateral root in the field of view.</li>
	<li>Before recording, allow the system to equilibrate for at least 15 min.</li>
	<li>Setup the image acquisition.
	<ol>
		<li>Set a stack of 217 images with 3 &#181;m z-spacing (650 &#181;m) and set the time lapse with 15 min imaging interval for a total period of 17 h. 
		NOTE: A detailed documentation on how to operate the OpenSPIM software can be found on (http://openspim.org/Acquisition#Acquiring_a_Stack).</li>
	</ol>
	</li>
	<li>Start recording.</li>
</ol>

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The authors have nothing to disclose.

Light Sheet Fluorescence Microscopy has the great advantage to combine low phototoxicity and ultrafast acquisition speed, which can be used to capture a large volume with a high spatio-temporal resolution while keeping the sample in a physiological state. The resolution of a light sheet microscope can be compared to that of a confocal microscope9. However, light scattering and absorption occurs along the excitation and emission path individually and the overall image quality can be significantly lower inside opaque tissues compared to the surface. To circumvent this complication one can use the possibility to rotate the sample along the vertical axis and observe the same volume from different directions. But this is not always advantageous, e.g. lateral roots emerge on one side of the root and imaging from behind results in a low image quality without gaining more information. However, the rotation can be principally used to position the sample in the best way. The classic horizontal arrangement of the objective lenses allows for new ways of sample mounting. Plants benefit from a vertical position. Presented here, the &#34;on the surface of the gel&#34; mounting method has several advantages compared to other mounting methods such as embedding the root inside of a gel24,25. 1) The root system is in direct contact with the liquid medium. The sample chamber is connected to a perfusion system which provides continuously fresh medium. It can also be used to rapidly exchange the entire volume of the sample chamber to apply different media or drugs. 2) Prior to sample preparation plants grow as they are used to grow in laboratories. Plants can be selected under a fluorescence microscope and only the desired plants need to be prepared. 3) The plant is transferred from the Petri dish to the sample holder without being touched. Thereby the plant can further develop on the same gel it was growing on in the growth incubator and mechanical stress is reduced to a minimum. 4) The view on the specimen is unobstructed and optical aberrations are minimized because the space between the sample and the detection objective is solely filled with medium and no other materials with differing refractive indices.
In order to perform long-term imaging, the plant illumination system is necessary to ensure photosynthetic activity of the plant. In most laboratories plants grow on a transparent gel, i.e. the roots are exposed to light. This may cause different responses to their environment and induces changes in their biochemistry and development26,27. In order to reduce the amount of light on the root system, black plastic foil was used to cover the water surface as well as a lid made of black aluminum foil covered the sample chamber. Light can reach the plant leaves through the central hole in the lid. In this setup, no increase in background light was observed, suggesting that the amount of stray light from the red and blue LEDs was significantly reduced by the GFP filter and the shading approaches. This allowed keeping the light turned on during image acquisition without increasing the camera background noise.
The sample holder is designed for 3D printing. However, the choice of material is crucial as several plastics that were tested were not 100% stable, resulting in a drift of the sample. Therefore, it is recommended to use resins instead or build the sample holder by milling a Polyethylene (PEP) rod. When using a light sheet microscope setup with double-sided illumination system the sample holder might interfere with one of the light sheets depending on the rotation angle. To reduce mechanical stress during scooping the plant from the plate, use a flat angle of the spatula. The plant can quickly dry out and experience air flow for the very first time. Try to avoid any air draft (rapid movements, air-condition flow), work uninterruptedly and slide the sample holder into a 1,000 &#181;L pipet tip whenever possible. Inside the microscope, it is crucial to not dip the whole plant in liquid and keep the leaves dry.
The technique is ideal for imaging early stages of lateral root formation. When performing long term imaging of mature root tips one must keep in mind that Arabidopsis roots grow with 100-300 &#181;m/h rapidly out of the field of view. A very useful future implementation could be an automated tracking algorithm, which would allow following root tip growth over prolonged periods of time. The ability to control environmental conditions such as light and nutrient composition of the medium during the acquisition process allows investigating how plants adapt to changes. The root is in direct contact with the liquid medium, which can be used to apply drugs to chemically activate gene expression, for example using the dexamethasone inducible28 or the &#946;-estradiol inducible system29. However, it takes time to exchange the entire volume of the sample chamber to wash out a drug. The setup could be improved by minimizing the volume of the sample chamber to accelerate medium exchange. Nevertheless, this technique has a great potential. The combination of mounting procedure, standardized growing conditions and the gentle image acquisition using light-sheet microscopy allows long-term studies of plant development with high resolution at a physiological level. This will help researchers to explore fundamental mechanisms of plant development.

One of the key questions in understanding plant development is how single cells behave during organ differentiation and growth. Ideally, cellular events, like gene expression patterns and intracellular protein localization, can be seen in the light of a larger context of the tissue. This objective poses technical challenges and requires whole organ observation with a high spatial as well as temporal resolution over prolonged periods of time, which may cause photo-toxicity effects. Since plants quickly adapt to environmental changes, the growing conditions must be tightly controlled. In order to do long-term imaging without interfering with the physiological state of the plant, three things have to be ensured, 1) growing conditions in the sample chamber, 2) stable sample mounting over long periods of time, and 3) the imaging with low light intensities to avoid photo-damage and non-physiological conditions.
Physiological growing conditions in the microscope specimen chamber are crucial for long-term experiments. There are a number of protocols available that describe imaging growth chambers for confocal microscopes1-3. However, confocal microscopy introduces high light-intensity to the plant, which can cause stress responses and usually inhibits the growth4. In addition, most conventional microscopes allow only horizontal positioning of the sample, which is not optimal for plants since they try to reorient themselves and grow towards the vector of gravity. Over the past ten years, light-sheet microscopy has emerged as a powerful tool to capture the development of large specimens at cellular resolution for time periods of up to several days5-9. Light-sheet microscopy allows positioning the specimen vertically and is increasingly used in plant research studying root development10-21, recently reviewed by Berthet and Maizel22. Many of the mentioned studies10,13-18,21 were optimized and conducted in the laboratory of Ernst H.K. Stelzer employing a special way of sample mounting characterized by growing the root on the surface of a gel17. In these studies, a custom-made microscope was used, in which the plant is held from the bottom. In contrast, the majority of broadly available light-sheet microscopes hold the sample from the top. Thus, this particular preparation method cannot be readily applied. The method presented here provides a protocol for the well-established on-surface mounting method applicable for the OpenSPIM23, an open access platform for applying and enhancing Selective Plane Illumination Microscopy (SPIM).
The overall goal of this protocol is to enable long-term imaging of Arabidopsis roots in the OpenSPIM light-sheet microscope. This is accomplished by growing a plant upright on the surface of a gel with the roots 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 but not the roots (Figure 1).

<table><tbody><tr><td>Agarose, low melting</td><td>VWR</td><td>AFFY3282125GM</td><td></td></tr><tr><td>Black aluminum foil</td><td>Thorlabs</td><td>BKF12</td><td></td></tr><tr><td>Black plastic foil</td><td>Carl Roth</td><td>HT83.2</td><td></td></tr><tr><td>LED blue (453 nm)</td><td>OSRAM</td><td>LD&amp;nbsp;CN5M-1R1S-35-1</td><td></td></tr><tr><td>LED red (625 nm)</td><td>OSRAM</td><td>LR T66F-ABBB-1-1</td><td></td></tr><tr><td>LED board - PCB design software</td><td>Cadsoft Eagle</td><td></td><td></td></tr><tr><td>MES monohydrate</td><td>Duchefa</td><td>M1503.0100</td><td></td></tr><tr><td>Micropore Surgical Tape</td><td>3M</td><td>1530-1</td><td></td></tr><tr><td>Murashige &amp;amp; Skoog Medium (MS-Medium)</td><td>Duchefa</td><td>M0221</td><td></td></tr><tr><td>Phytagel</td><td>Sigma-Aldrich</td><td>P8169</td><td></td></tr><tr><td>Sample holder 3D print</td><td>i.materialise</td><td>&lt;a href=&quot;https://i.materialise.de/shop/item/sampleholder-openspim-zeisslightsheetz1&quot;&gt;https://i.materialise.de/shop/item/sampleholder-openspim-zeisslightsheetz1&lt;/a&gt;</td><td></td></tr><tr><td>Square Petri dishes (245 x 245 x 25 mm)</td><td>VWR</td><td>734-2179</td><td></td></tr><tr><td>Sucrose</td><td>Sigma-Aldrich</td><td>84097-1KG</td><td></td></tr></tbody></table>

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 sample preparation method allows the cultivation of the plant inside the microscope sample chamber while observing the root system with a light-sheet microscope (Figure 1). The plant grows on the surface of a layer of gel (&#189; MS medium containing 1.5% gellan gum) mounted on a custom designed sample holder (Figure 2). The sample holder is 3D printed using a transparent resin as material. A manufactured version highlighting the dimensions is depicted in Figure 2C. The roots are immersed in liquid (&#189; MS medium), which is continuously refreshed by a perfusion system. The leaves remain in the air and are continuously illuminated with a light intensity of 130 &#181;mol/m&#178;/s coming from blue and red LEDs that are arranged in a ring above the plant (Figure 1A, B and Figure 3A-C). The LED ring is manufactured in our machine workshop and we provide technical details on how to build the LED-ring in the Figure 3 and the material list. The light intensity can be continuously adjusted ranging from 30-250 &#181;mol/m2/s (Figure 3D). The root system is shaded by small sheets of a black plastic foil covering the water surface (Figure 1). Any stray light from the illumination that is collected by the detection lens is filtered by the GFP filter (Figure 3E).
With this setup, a time lapse of a growing Arabidopsis lateral root was recorded for 17 h using a 20X/0.5 lens (Figure 4). The lateral root has its origin in the pericycle cell layer, which is located deep inside the primary root. In order to demonstrate the imaging capabilities even deeper inside of a tissue for prolonged time periods, a higher magnification (40X/0.75) was used to capture the formation of a lateral root from the first stage primordium until the emergence out of the primary root within a time period of 38 h (Figure 5). An exemplary 2D slice of the data set is shown in Fig. 5B. This recording allows us to follow the dynamics of lateral root formation in 3D (Figure 5A) with a cellular resolution.
<img alt="Figure 1" src="/files/ftp_upload/55044/55044fig1.jpg" /> 
Figure 1: Growing conditions inside the OpenSPIM sample chamber. A) Sketch of the imaging chamber. The plant is growing upright on the surface of a gel, mounted on a custom built sample holder (see also Figure 2). The roots are growing in a liquid medium, which is continuously exchanged by a perfusion system. The plant leaves grow in air and are illuminated by red and blue LEDs (see also Figure 3). The root system is shaded with small sheets of a black plastic foil covering the water surface. A lid made of two pieces of black aluminum foil further reduces the amount of light below the water surface and maintains humidity in the sample chamber. The magnified panel on the right highlights the plant growing on the surface of a block of gel immersed in the liquid medium. A drop of agarose mounts the root onto the gel. The dashed box indicates the region of interest observed by the microscope. B) Photograph of the imaging chamber (without lid). Numbers (1)-(10) in A and B represent: (1): x/y/z/&#952;-stage with LED ring, (2): sample holder, (3): lid, (4): Arabidopsis thaliana, (5): sheets of black plastic foil, (6): perfusion system, (7): detection objective lens, (8): liquid medium, (9): sample chamber, (10): illumination objective lens. C) The lid is made of two pieces of black aluminum foil. <a href="http://cloudflare.jove.com/files/ftp_upload/55044/55044fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" src="/files/ftp_upload/55044/55044fig2.jpg" /> 
Figure 2: The sample holder. A) 3D model. The 3D model file is provided in the supplemental material. B) Photograph of 3D prints using different materials (1)-(3): transparent acrylic plastics, (4) and (5): resin, (6): transparent resin. C) Technical drawing of the sample holder, numbers represent millimeter. D) Photograph of the manufactured sample holder with a plant mounted. The dashed area can be observed by the microscope. <a href="http://cloudflare.jove.com/files/ftp_upload/55044/55044fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" src="/files/ftp_upload/55044/55044fig3.jpg" /> 
Figure 3: Plant illumination setup. A) Schematic circuit diagram of the lamp. Pairs of LEDs can be switched on/off individually for directional lightning. LED: light-emitting diode, R: resistance, T: transistor, JP: pinhead. B) The final design of the illumination lamp was drawn using a PCB-software (PCB: printed circuit board). We provide the board design file in the supplemental material. The board was then manufactured and assembled in our institute&#39;s MIBA machine shop. C) Photograph of the LED ring switched on. Four pairs of a red and a blue LED are arranged in a ring. D) The range of voltage can be adjusted between 3.5 V and 14.0 V. Resistances were used to reach the amount of light ranging from 30-250 &#181;mol/m2/s (R1-8: 220 Ohm, R9-12: 1,220 Ohm). E) The emission spectrum of the lamp, GFP and YFP. <a href="http://cloudflare.jove.com/files/ftp_upload/55044/55044fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" src="/files/ftp_upload/55044/55044fig4.jpg" /> 
Figure 4: Time lapse recording of Arabidopsis thaliana lateral root. The 5 days old seedling expresses a membrane marker (pUBQ10::YFP-PIP1;4) and a nuclear reporter (pGATA23::nls-GUS-GFP) specifically marking pericycle cells that develop into a lateral root. A stack of 217 images (3 &#181;m z-spacing) was captured every 15 min for 17 h recording using a 20X/0.5 lens. A) Four time points out of 69 are shown in a maximum intensity projection. B) Six out of 217 single slices of a z-stack of one time point are shown. Scale bars in A and B represent 100 &#181;m. <a href="http://cloudflare.jove.com/files/ftp_upload/55044/55044fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" src="/files/ftp_upload/55044/55044fig5.jpg" /> 
Figure 5: Time lapse recording of Arabidopsis thaliana lateral root. The 6 days old seedling expresses a membrane marker (pUBQ10::YFP-PIP1;4) and a nuclear reporter (pGATA23::nls-GUS-GFP) specifically marking pericycle cells that develop into a lateral root. A stack of 200 images (1.5 &#181;m z-spacing) was captured every 15 min for 38 h recording using a 40X/0.75 lens. A) 3D rendering of four time points, the numbers in the grid represent &#181;m, B) single slice through the central plane of the main root. Scale bar represents 50 &#181;m. <a href="http://cloudflare.jove.com/files/ftp_upload/55044/55044fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Supplemental_File_-_3D_Sample_Holder.stl. The 3D model file is provided. <a href="http://cloudflare.jove.com/files/ftp_upload/55044/Supplemental_File-3D_Sample_Holder.stl" target="_blank">Please click here to download this file.</a>
Supplemental File LED Ring Board.brd. The board design file is provided. <a href="http://cloudflare.jove.com/files/ftp_upload/55044/Supplemental_File-LED_Ring_Board.brd" target="_blank">Please click here to download this file.</a>

Watch this Scientific Journal Video about Light Sheet Fluorescence Microscopy of Plant Roots Growing on the Surface of a Gel at JoVE.com

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

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 ...

light-sheet-fluorescence-microscopy-plant-roots-growing-on-surface

Research

JoVE Journal

Biology

14.6K Views.  Institute of Science and Technology Austria. 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.

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

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, ...

Bioengineering

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

Light Sheet-based Fluorescence Microscopy of Living or Fixed and Stained Tribolium castaneum Embryos

The red flour beetle Tribolium castaneum has become an important insect model organism in developmental genetics and evolutionary developmental biology. The observation of Tribolium embryos with light sheet-based fluorescence microscopy has multiple advantages over conventional widefield and confocal fluorescence microscopy. Due to the unique properties of a light sheet-based microscope, three dimensional images of living specimens can be recorded with high signal-to-noise ratios and significantly reduced photo-bleaching as well as photo-toxicity along multiple directions over periods that last several days. With more than four years of methodological development and a continuous increase of data, the time seems appropriate to establish standard operating procedures for the usage of light sheet technology in the Tribolium community as well as in the insect community at large. This protocol describes three mounting techniques suitable for different purposes, presents two novel custom-made transgenic Tribolium lines appropriate for long-term live imaging, suggests five fluorescent dyes to label intracellular structures of fixed embryos and provides information on data post-processing for the timely evaluation of the recorded data. Representative results concentrate on long-term live imaging, optical sectioning and the observation of the same embryo along multiple directions. The respective datasets are provided as a downloadable resource. Finally, the protocol discusses quality controls for live imaging assays, current limitations and the applicability of the outlined procedures to other insect species.
This protocol is primarily intended for developmental biologists who seek imaging solutions that outperform standard laboratory equipment. It promotes the continuous attempt to close the gap between the technically orientated laboratories/communities, which develop and refine microscopy methodologically, and the life science laboratories/communities, which require 'plug-and-play' solutions to technical challenges. Furthermore, it supports an axiomatic approach that moves the biological questions into the center of attention.

Light Sheet-based Fluorescence Microscopy of Living or Fixed and Stained Tribolium castaneum Embryos

Imaging the morphogenesis of insect embryos with light sheet-based fluorescence microscopy has become state of the art. This protocol outlines and compares three mounting techniques appropriate for Tribolium castaneum embryos, introduces two novel custom-made transgenic lines well suited for live imaging, discusses essential quality controls and indicates current experimental limitations.

The red flour beetle Tribolium castaneum has become an important insect model organism in developmental genetics and evolutionary developmental biology. ...

Measuring Interactions between Fluorescent Probes and Lignin in Plant Sections by sFLIM Based on Native Autofluorescence

In lignocellulosic biomass (LB), the activity of enzymes is limited by the appearance of non-specific interactions with lignin during the hydrolysis process, which maintains enzymes far from their substrate. Characterization of these complex interactions is thus a challenge in complex substrates such as LB. The method here measures molecular interactions between fluorophore-tagged molecules and native autofluorescent lignin, to be revealed by F&#246;rster resonance energy transfer (FRET). Contrary to FRET measurements in living cells using two exogenous fluorophores, FRET measurements in plants using lignin is not trivial due to its complex autofluorescence. We have developed an original acquisition and analysis pipeline with correlated observation of two complementary properties of fluorescence: fluorescence emission and lifetime. sFLIM (spectral and fluorescent lifetime imaging microscopy) provides the quantification of these interactions with high sensitivity, revealing different interaction levels between biomolecules and lignin.

This protocol describes an original setup combining spectral and fluorescence lifetime measurements to evaluate F&#246;rster resonance energy transfer (FRET) between rhodamine-based fluorescent probes and lignin polymer directly in thick plant sections.

In lignocellulosic biomass (LB), the activity of enzymes is limited by the appearance of non-specific interactions with lignin during the hydrolysis ...

Biochemistry

Two-Dimensional Visualization and Quantification of Labile, Inorganic Plant Nutrients and Contaminants in Soil

We describe a method for two-dimensional (2D) visualization and quantification of the distribution of labile (i.e., reversibly adsorbed) inorganic nutrient (e.g., P, Fe, Mn) and contaminant (e.g., As, Cd, Pb) solute species in the soil adjacent to plant roots (the &#8216;rhizosphere&#8217;) at sub-millimeter (~100 &#181;m) spatial resolution. The method combines sink-based solute sampling by the diffusive gradients in thin films (DGT) technique with spatially resolved chemical analysis by laser ablation inductively coupled plasma mass spectrometry (LA-ICP-MS). The DGT technique is based on thin hydrogels with homogeneously distributed analyte-selective binding phases. The variety of available binding phases allows for the preparation of different DGT gel types following simple gel fabrication procedures. For DGT gel deployment in the rhizosphere, plants are grown in flat, transparent growth containers (rhizotrons), which enable minimal invasive access to a soil-grown root system. After a pre-growth period, DGT gels are applied to selected regions of interest for in situ solute sampling in the rhizosphere. Afterwards, DGT gels are retrieved and prepared for subsequent chemical analysis of the bound solutes using LA-ICP-MS line-scan imaging. Application of internal normalization using 13C and external calibration using matrix-matched gel standards further allows for the quantification of the 2D solute fluxes. This method is unique in its capability to generate quantitative, sub-mm scale 2D images of multi-element solute fluxes in soil-plant environments, exceeding the achievable spatial resolution of other methods for measuring solute gradients in the rhizosphere substantially. We present the application and evaluation of the method for imaging multiple cationic and anionic solute species in the rhizosphere of terrestrial plants and highlight the possibility of combining this method with complementary solute imaging techniques.

This protocol presents a workflow for sub-mm 2D visualization of multiple labile inorganic nutrient and contaminant solute species using diffusive gradients in thin films (DGT) combined with mass spectrometry imaging. Solute sampling and high-resolution chemical analysis are described in detail for quantitative mapping of solutes in the rhizosphere of terrestrial plants.

We describe a method for two-dimensional (2D) visualization and quantification of the distribution of labile (i.e., reversibly adsorbed) inorganic ...

Environment

Fluorescence Activated Cell Sorting of Plant Protoplasts

High-resolution, cell type-specific analysis of gene expression greatly enhances understanding of developmental regulation and responses to environmental stimuli in any multicellular organism. In situ hybridization and reporter gene visualization can to a limited extent be used to this end but for high resolution quantitative RT-PCR or high-throughput transcriptome-wide analysis the isolation of RNA from particular cell types is requisite. Cellular dissociation of tissue expressing a fluorescent protein marker in a specific cell type and subsequent Fluorescence Activated Cell Sorting (FACS) makes it possible to collect sufficient amounts of material for RNA extraction, cDNA synthesis/amplification and microarray analysis.
An extensive set of cell type-specific fluorescent reporter lines is available to the plant research community. In this case, two marker lines of the Arabidopsis thaliana root are used: PSCR::GFP (endodermis and quiescent center) and PWOX5::GFP (quiescent center). Large numbers (thousands) of seedlings are grown hydroponically or on agar plates and harvested to obtain enough root material for further analysis. Cellular dissociation of plant material is achieved by enzymatic digestion of the cell wall. This procedure makes use of high osmolarity-induced plasmolysis and commercially available cellulases, pectinases and hemicellulases to release protoplasts into solution.
FACS of GFP-positive cells makes use of the visualization of the green versus the red emission spectra of protoplasts excited by a 488 nm laser. GFP-positive protoplasts can be distinguished by their increased ratio of green to red emission. Protoplasts are typically sorted directly into RNA extraction buffer and stored for further processing at a later time.
This technique is revealed to be straightforward and practicable. Furthermore, it is shown that it can be used without difficulty to isolate sufficient numbers of cells for transcriptome analysis, even for very scarce cell types (e.g. quiescent center cells). Lastly, a growth setup for Arabidopsis seedlings is demonstrated that enables uncomplicated treatment of the plants prior to cell sorting (e.g. for the cell type-specific analysis of biotic or abiotic stress responses). Potential supplementary uses for FACS of plant protoplasts are discussed.

A method for isolating specific cell types from plant material is demonstrated. This technique employs transgenic marker lines expressing fluorescent proteins in particular cell types, cellular dissociation and Fluorescence Activated Cell Sorting. Additionally, a growth setup is established here that facilitates treatment of Arabidopsis thaliana seedlings prior to cell sorting.

High-resolution, cell type-specific analysis of gene expression greatly enhances understanding of developmental regulation and responses to environmental ...

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 ...

3-D Time-Lapse Imaging of Cell Wall Dynamics Using Calcofluor in the Moss Physcomitrium patens

Time-lapse imaging with fluorescence microscopy allows observation of the dynamic changes of growth and development at cellular and subcellular levels. In general, for observations over a long period, the technique requires transformation of a fluorescent protein; however, for most systems, genetic transformation is either time-consuming or technically unavailable. This manuscript presents a protocol for 3-D time-lapse imaging of cell wall dynamics over a 3 day period using calcofluor dye (which stains cellulose in the plant cell wall), developed in the moss Physcomitrium patens. The calcofluor dye signal from the cell wall is stable and can last for 1 week without obvious decay. Using this method, it has been shown that the detachment of cells in ggb mutants (in which the protein geranylgeranyltransferase-I beta subunit is knocked out) is caused by unregulated cell expansion and cell wall integrity defects. Moreover, the patterns of calcofluor staining change over time; less intensely stained regions correlate with the future cell expansion/branching sites in the wild type. This method can be applied to many other systems that contain cell walls and that can be stained by calcofluor.

3-D Time-Lapse Imaging of Cell Wall Dynamics Using Calcofluor in the Moss Physcomitrium patens

This manuscript presents a detailed protocol to image the 3-D cell wall dynamics of living moss tissue, allowing the visualization of the detachment of cell walls in ggb mutants and thickening cell wall patterns in the wild type during development over a long period.

Time-lapse imaging with fluorescence microscopy allows observation of the dynamic changes of growth and development at cellular and subcellular levels. In ...

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 ...

Real-time Imaging of Plant Cell Surface Dynamics with Variable-angle Epifluorescence Microscopy

A plant&#8217;s cell surface is its interface for perceiving environmental cues; it responds with cell biological changes such as membrane trafficking and cytoskeletal rearrangement. Real-time and high-resolution image analysis of such intracellular events will increase the understanding of plant cell biology at the molecular level. Variable angle epifluorescence microscopy (VAEM) is an emerging technique that provides high-quality, time-lapse images of fluorescently-labeled proteins on the plant cell surface. In this article, practical procedures are described for VAEM specimen preparation, adjustment of the VAEM optical system, movie capturing and image analysis. As an example of VAEM observation, representative results are presented on the dynamics of PATROL1. This is a protein essential for stomatal movement, thought to be involved in proton pump delivery to plasma membranes in the stomatal complex of Arabidopsis thaliana. VAEM real-time observation of guard cells and subsidiary cells in A. thaliana cotyledons showed that fluorescently-tagged PATROL1 appeared as dot-like structures on plasma membranes for several seconds and then disappeared. Kymograph analysis of VAEM movie data determined the time distribution of the presence (termed &#8216;residence time&#8217;) of the dot-like structures. The use of VAEM is discussed in the context of this example.

The goal of this protocol is to demonstrate how to monitor fluorescently-tagged protein dynamics on plant cell surfaces with variable-angle epifluorescence microscopy, showing blinking dots of GFP-tagged PATROL1, a membrane trafficking protein, in the cell cortex of the stomatal complex in Arabidopsis thaliana.

A plant&#8217;s cell surface is its interface for perceiving environmental cues; it responds with cell biological changes such as membrane trafficking and ...

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

Summary

Explore More Videos

Chapters in this video

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

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

Light Sheet-based Fluorescence Microscopy of Living or Fixed and Stained Tribolium castaneum Embryos

Measuring Interactions between Fluorescent Probes and Lignin in Plant Sections by sFLIM Based on Native Autofluorescence

Two-Dimensional Visualization and Quantification of Labile, Inorganic Plant Nutrients and Contaminants in Soil

Fluorescence Activated Cell Sorting of Plant Protoplasts

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

3-D Time-Lapse Imaging of Cell Wall Dynamics Using Calcofluor in the Moss Physcomitrium patens

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

Real-time Imaging of Plant Cell Surface Dynamics with Variable-angle Epifluorescence Microscopy