This work was supported by USDA in-house funding.

1. Preparation
<ol>
	<li>Gather equipment, materials, and software to record and process the video of plant freezing.
	<ol>
		<li>Start a programmable freezer by setting the power switch to On, and set the temperature to 0 &#176;C. Program the freezer to reach -8 &#176;C at 1 &#176;C/h.</li>
	</ol>
	</li>
	<li>Place one 6-weeks-old strawberry plant with 2 - 5 flowers that was grown in a 1 L container into the freezer.</li>
	<li>Set up 2 IR cameras (e.g., FLIR T620 cameras) using fastening straps and a small block of wood to produce the correct convergence angle of the lenses. 
	Note: The optimum distance to space the center of the lenses of the 2 cameras is generally considered to be the same as the distance between the eyes1 or approximately 7 cm.</li>
	<li>Mount both cameras to a 10 x 10 cm laboratory jack and position the jack in the freezer close enough to the plant to allow the image to be focused. Adjust the cameras vertically and horizontally so that the same part of the plant is visible from both cameras. Use the jack to position both cameras vertically in such a way that the 2 images contain the whole plant and a portion of the soil.</li>
	<li>Connect the 2 cameras using a USB connector to the USB outlets on the computer.</li>
	<li>Plug the 2 A/C outlets into both cameras to allow a continuous monitoring of the plants.</li>
</ol>
2. Computer and Software Setup for Recording
<ol>
	<li>Open 2 windows (1 window for each camera) of the software by double-clicking the icon for the IR camera software 2x. Follow the instructions in the help menu to connect the left camera in the left window and the right camera in the right window. 
	Note: The details for using the software can be accessed through the help menu. A monochromatic palette is best suited for this example due to the necessity of using red-blue tinting for the 3D rendering.</li>
	<li>Open the screen capture software by double-clicking the icon for the program. Adjust the capture frame by clicking and dragging the frame so it includes both cameras to allow a screen capture of both cameras simultaneously. 
	Note: Screen capturing the video stream from both cameras simultaneously is crucial because it allows a flawless synchronization of both left and right views.</li>
	<li>Record the video in 3-h increments for an easier processing in the video processing software. 
	Note: It is impossible to know exactly when the plant will freeze, so it is important to record for some time prior to the freeze event. The option to record in segments is a feature of this software, so it is recommended that this is set to record for 3 h. The software will automatically save the 3-h recording and then begin a new recording. The file for each 3-h recording will automatically be given a numerical sequence after the name. Each video file will be from 10 to 20 GB, so ensure that sufficient space is available on a hard drive for multiple files of this size.</li>
	<li>Start the freezer program by selecting Run in the controller menu and begin the screen capture. Then press the Rec button on the window. Ensure that the outline showing the region of the screen being captured turns red.</li>
	<li>Record the strawberry plant freezing down to -8 &#176;C and hold the temperature of the freezer for 1 h.</li>
	<li>Raise the temperature of the freezer at 2 &#176;C/h until the freezer is at +2 &#176;C. Stop the recording. 
	Note: The total freezing time is 14 h.</li>
	<li>Convert the file(s) of interest from .mp4 format to .mov using a file conversion software. 
	Note: In this case, a single 3 h file containing 1 or more freeze events will be used.</li>
</ol>
3. Processing Video Using a Video Imaging Software
Note: Video imaging software will be used in this example. Tutorials on how to use the software are available online. This example will assume a basic knowledge of the software. The understanding of terms such as &#8220;composition&#8221;, &#8220;layer&#8221;, and &#8220;render queue&#8221;, as well as the various panels and how to manipulate them, is assumed.
<ol>
	<li>Double-click anywhere inside the project panel to import the .mov file of interest into the imaging software and drag the file to the Composition icon at the bottom of the project panel. Then save the project to the same folder containing the original videos. 
	Note: The video recorded will be visible in the preview pane.</li>
	<li>Click on the Region of Interest icon along the bottom of the preview window and, using the cursor, outline only the recording from the left camera.</li>
	<li>Drag the same .mov video to the Composition icon to create a second composition of the same video. Repeat step 3.3, but this time, use the cursor to select only the right camera.</li>
	<li>Select Composition &#62; Crop Comp to Region of Interest for the left view. Repeat this for the right view. Rename each composition to indicate which is left and right.</li>
	<li>Highlight the left composition by clicking on it and, in the main menu at the top, selecting Composition &#62; Add to Render Queue.</li>
	<li>In the render queue, click on Output Module and make sure the video will be rendered as a video (e.g., a QuickTime video). Click on the Render Settings to reduce the resolution to allow a faster render. Click on Output to, name the video Strawberry Left, and save it to the same folder as the original recording and the project. Click Save, and then click the Render button at the top right side of the render panel.</li>
	<li>Repeat step 3.6 for the Strawberry Right composition.</li>
	<li>Double-click the project panel and import the Strawberry Left and Strawberry Right videos that were just rendered.</li>
	<li>Highlight both videos and drag them to the composition icon at the bottom of the project panel. In the pop-up screen asking for the Still Duration, enter 3 with 5 zeros for a 3-h duration. 
	Note: Both videos, precisely synchronized, will be in the project panel, but only the topmost video in the composition panel will be visible.</li>
	<li>To view the other image, click on the little eyeball to turn off the layer. Press Control/W to allow a rotational control of the images in the preview panel using the cursor. Using the cursor and clicking the top layer on and off, adjust the rotational aspect of either the top or the bottom view to make sure both images are in the same rotational plane. Then adjust the X- and Y-plane directly within the 3D glasses subroutine.</li>
	<li>Highlight the top layer of the composition panel and select Effect &#62; Perspective &#62; 3D Glasses from the menu at the top. 
	Note: The parameters for the 3D Glasses Effect will pop up inside the control panel.</li>
	<li>In the control panel, click the box to the right of &#8220;left view&#8221; &#8212; if the control panel is not separated from the project panel, click the control panel tab at the top of the project panel. List the 2 videos in the composition panel in a drop-down menu, highlight the video in the list for the &#8220;left view&#8221;. Repeat this step for &#8220;right view&#8221;.</li>
	<li>In the box to the right of 3D View, select Red Blue LR.</li>
	<li>Using red-blue glasses, inspect the view in the project panel. If the 3D view seems to be incorrect, try clicking Swap Left-Right. Adjust Scene convergence and Vertical alignment to eliminate any ghosting and eye strain.</li>
	<li>When the 3D aspect of the video is acceptable, highlight the composition by clicking on it and select Composition &#62; Add to render queue as was done in step 3.7. Render the video into the same folder as the other files in the project. 
	Note: This file will be rather large. Once the file has been rendered, it can be re-rendered to a smaller file size using the video processing software.</li>
</ol>

<ol>
	<li>Geng, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Three-dimensional+display+technologies.">Three-dimensional display technologies.</a> Advances in Optics and Photonics. 5, 456-535 (2013).</li><li>Ceccardi, T. L., Heath, R. L., Ting, I. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Low-temperature+exotherm+measurement+using+infrared+thermography.">Low-temperature exotherm measurement using infrared thermography.</a> HortScience. 30, 140-142 (1995).</li><li>Wimmer, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> History of thermal imaging, Security Sales and Integration. , (2011).</li><li>Wisniewski, M., Lindow, S. E., Ashworth, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Observations+of+ice+nucleation+and+propagation+in+plants+using+infrared+video+thermography.">Observations of ice nucleation and propagation in plants using infrared video thermography.</a> Plant Physiology. 113, 327-334 (1997).</li><li>Kuprian, E., Tuong, T., Pfaller, K., Livingston, D. P., Neuner, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Persistent+supercooling+of+reproductive+shoots+is+enabled+by+structural+ice+barriers+being+active+despite+an+intact+xylem+connection.">Persistent supercooling of reproductive shoots is enabled by structural ice barriers being active despite an intact xylem connection.</a> Public Library of Science ONE. 11, e0163160 (2016).</li><li>Livingston, D. P., Tuong, T. D., Murphy, J. P., Gusta, L., Wisniewski, M. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=High-definition+infrared+thermography+of+ice+nucleation+and+propagation+in+wheat+under+natural+frost+conditions+and+controlled+freezing.">High-definition infrared thermography of ice nucleation and propagation in wheat under natural frost conditions and controlled freezing.</a> Planta. 247, 791-806 (2017).</li><li>Boyles, R. P., Raman, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Analysis+of+climate+patterns+and+trends+in+North+Carolina+(1949-1998).">Analysis of climate patterns and trends in North Carolina (1949-1998).</a> Environment International. 29 (2-3), 263-275 (2003).</li><li>Poling, E. B., Poling, E. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Managing+Cold+Events.">Managing Cold Events.</a> A Growers' Guide to Production, Economics and Marketing. , 75-97 (2015).</li><li>Hacker, J., Neuner, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Ice+porpagaion+in+plants+visualized+at+the+tissue+level+by+infrared+differential+thermal+analysis+(IDTA).">Ice porpagaion in plants visualized at the tissue level by infrared differential thermal analysis (IDTA).</a> Tree Physiology. 27, 1661-1670 (2007).</li><li>He, J. Q., Harrison, R. J., Li, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+novel+3D+imaging+system+for+strawberry+phenotyping.">A novel 3D imaging system for strawberry phenotyping.</a> Plant Methods. 13, 93-101 (2017).</li></ol>

The authors have nothing to disclose.

Two IR cameras are necessary for this protocol, and they must be aimed at the subject from slightly different angles1. This will require the lenses to be from 5 - 8 cm apart, but both must be aimed at the same place on the subject to be filmed. Think of the 2 camera lenses as a kind of surrogate for the viewer&#39;s eyes. The left camera is analogous to the left eye and the right camera to the right eye. The post-processing software will tint the left image to a red color and the right image to a blue color, so by wearing red-blue glasses, the left eye can only see the left image and the right eye only the right image. This means that it is important to use the gray-scale palette of the IR camera software when recording the freeze event. The brain will combine the 2 images which the viewer will observe in 3D1.
Another critical step to this protocol is the use of screen capture software to capture the output of both cameras simultaneously. By capturing the output of both cameras simultaneously, a perfect synchronization of the output from both cameras is guaranteed. Synchronizing the right and left images is a crucial aspect of producing 3D films and are discussed elsewhere in detail.1
To prevent any eye strain, it is important that the vertical and the horizontal convergence of the left and right images are correct. While the cameras should be positioned to ensure a correct convergence prior to the recording, they do not have to be perfect. The post-production software described here will allow adjustments in right-left, up-down, and rotational convergence. The software will also allow a red-green anaglyph video to be produced if red-blue glasses are not available.
One limitation of the technique is the requirement of red-blue glasses to view the 3D video. It is likely that many individuals will not have red-blue glasses readily available. Also, while producing a red-blue anaglyph video is the easiest and least expensive way to produce a 3D video, red-blue anaglyph videos can only convey a limited chromatic view of their subject. However, this is arguably an insignificant limitation since IR radiation, in reality, can only be observed in grayscale. Colors are only perceived by humans in the visible portion of the electromagnetic spectrum.
The limited resolution in early IR technology made it difficult to determine the precise locations of ice nucleation as well as which tissues the ice propagated into. Differential thermal analysis9 has improved the ability to detect sites of ice nucleation; however, it remains a 2-dimensional perspective that lacks information about depth. The lack of information provides a limited perspective and does not fully represent freezing as it occurs in the real world.
Commercial films use various techniques for visualizing images in 3D, the most common being polarization-interlacing1. The most popular techniques require headgear that is specific to the interlacing process, but auto-stereoscopic techniques that do not require headgear are in developmental stages1. None of the 3D rendering techniques, however, are available for viewing the IR video in 3 dimensions. In addition, while these techniques provide the clearest 3D video available, they require synchronization and special projection devices as well as reflective surfaces on which to project the images1.
Communicating scientific findings in the clearest possible manner is essential for creating a community that will promote an efficient and timely progress in scientific discoveries. Observations of the world we live in are always in 3 dimensions, but it is difficult to accurately represent those observations using only 2D images. For example, it would be difficult, if not impossible, to determine precisely which root(s) had frozen in the IR imaging of the freezing in wheat roots (Figure 2B). However, using a 3D anaglyph process makes it relatively simple to determine exactly which root froze at what time (Figure 2D). Admittedly, it remains to be determined what new information (not obtainable from 2D videography) might be gleaned from a 3D perspective of freezing in plants. However, it is not unusual for unique information to be obtained when analyzing plant material in 3D10. By using screen capture software to precisely synchronize the right-left images and commercially available software to create an anaglyph video, any laboratory that uses visual data to understand biological processes can generate images and video in 3D.

Despite living in a world of three physical dimensions, researchers are often limited to reporting visual observations in 2D. Although 2D images are generally sufficient to convey important information, this lack of information about depth restricts our ability to perceive and understand the complexity of real-world objects.1
This deficiency in information about the depth provided an incentive to produce 3D videos mainly in the commercial film industry since the early 1900s1. However, generating clear 3D information in still images and video is hindered by the complexities involved in producing those images. The simplest approach to generating 3D film is based on principles used in stereoscopic photography. Stereoscopic photography utilizes two images of the same object from slightly different angles that conveys a 3D image in the brain.To make this possible, each eye must look only at its respective image (i.e., the left eye at the left image and the right eye at the right image). Since the eyes will not naturally do this, stereoscopic headgear was designed to make this possible1. Several stereoscopic viewing techniques, as well as polarization-interlaced, time-multiplexed, and head-mount display techniques, have been used during the development of 3D films, but the color-interlacing or anaglyph method using red and green (or cyan) glasses is one of the simplest and least expensive techniques. For a comprehensive review of 3D imaging and the various techniques involved, see the review by Geng1.
Monitoring freezing in plants using IR thermography is based on the principle that when water freezes, it must give up internal energy2. This energy is in the form of heat, which is detectable in the IR region of the electromagnetic spectrum. Cameras able to record the IR energy have been in use since 19293. The first published report using IR technology to film freezing in plants is from Cecardi et al.2, but the resolution of the camera used makes it difficult to accurately determine the tissue where the freezing is initiated. Wisniewski et al.4 determined more precise sites of ice nucleation in several plant species using a higher resolution camera. As the technology used in IR thermography improved, higher resolution images led to discoveries such as barriers to freezing5 and the precise cellular localization of ice formation6.
One difficulty in filming subjects in IR is caused by small differences in temperatures. This will cause most objects in the field of view to be a similar color, making it difficult to determine precisely which object(s) is/are freezing. This can be important when determining the order of freezing in specific tissues, such as leaves or roots in wheat6. If the IR video of plants freezing could be imaged in 3D, the accuracy of determining which part of the plant is freezing at a certain point in time could be improved.
Strawberry is a crop in certain areas of the United States in which freezing temperatures are of considerable concern for the growers. Under some growing conditions, it is common for strawberry flowers to appear 2 - 3 weeks before the average last spring freeze. A freeze event can occur as late as June in some areas of the Appalachian Mountains7 and usually results in the death of the flower. Frost protection is, therefore, critical for strawberry growers in areas subject to these freeze events. Strawberry growers in North Carolina, for example, must frost-protect, on average, between 4 - 6 frost events before bloom and 1 - 2 hard freezes during the early bloom period8. To help develop strawberry genotypes that are more freezing tolerant, it is important to understand various aspects of the freezing, such as the sites of ice nucleation and propagation into other parts of the plant. IR thermography provides an effective means to address these issues.
Here, we use strawberry to illustrate a technique for recording freezing events in 3D using the anaglyph method. Strawberry is well suited for this example because the leaves and flowers are widely distributed in the 3D space and can be difficult to differentiate when viewed in 2D infrared videos.

<table><tbody><tr><td>T620 Infrared Camera and software</td><td>FLIR</td><td>55903-5122</td><td>2 cameras are needed. Software works only on a Windows-based computer</td></tr><tr><td>After Effects</td><td>Adobe</td><td>15.0.1.73</td><td>Post-Production Video Editing Software</td></tr><tr><td>Bandicam</td><td>Bandisoft</td><td>4.1.2.1385</td><td>Screen Capture Software</td></tr><tr><td>Laboratory Scissor Jack&amp;nbsp;&amp;nbsp;</td><td>Eisco</td><td>CH0642A</td><td>Steel Platform 13X15 cm</td></tr><tr><td>Fastening Strap</td><td>Velcro</td><td>90441</td><td>To hold camera on jack.&amp;nbsp; Should be at least 60cm long by 2cm wide</td></tr><tr><td>Media Converter</td><td>iSkysoft</td><td>10.0.6</td><td>Software to convert mp4 files to .mov&amp;nbsp;</td></tr></tbody></table>

protocol for producing three-dimensional infrared video of freezing in plants

Freezing in plants can be monitored using infrared (IR) thermography, because when water freezes, it gives off heat. However, problems with color contrast make 2-dimensions (2D) infrared images somewhat difficult to interpret. Viewing an IR image or the video of plants freezing in 3 dimensions (3D) would allow a more accurate identification of sites for ice nucleation as well as the progression of freezing. In this paper, we demonstrate a relatively simple means to produce a 3D infrared video of a strawberry plant freezing. Strawberry is an economically important crop that is subjected to unexpected spring freeze events in many areas of the world. An accurate understanding of the freezing in strawberry will provide both breeders and growers with more economical ways to prevent any damage to plants during freezing conditions.
The technique involves a positioning of two IR cameras at slightly different angles to film the strawberry freezing. The two video streams will be precisely synchronized using a screen capture software that records both cameras simultaneously. The recordings will then be imported into the imaging software and processed using an anaglyph technique. Using red-blue glasses, the 3D video will make it easier to determine the precise site of ice nucleation on leaf surfaces.

Surprisingly, the IR video of the strawberry plant freezing (Supplemental Video 1) indicated that not all leaves/flowers froze at the same time. The leaves and flowers both froze individually at different temperatures, but the leaves froze earlier than the flowers and at a higher temperature. In addition, the freezing began in the leaves but not necessarily at the same position on each leaf. While these results have not been described previously in strawberry, similar results have been found in other plant species6. Once the leaves were frozen, the ice progressed down the petiole to the crown of the plant. When the freezer temperature became 1 or 2 degrees colder, the flowers froze beginning at the calyx and quickly spreading into the petals and receptacle (Figure 1). The receptacle remained a lighter color (warmer) longer than most other plant parts, suggesting a greater amount of water freezing.
When comparing the 2D infrared image with the 3D (wearing glasses), the 3D image makes it easier to precisely determine the order in which the leaves and the flowers froze (Figure 1). When viewing the video in 3D, it is also easier to determine the exact position on the leaves where the freezing began (Supplemental Video 1).
The survival results (not shown) indicated that despite the freezing, the leaves were not killed (not shown) by the freeze. The flowers that froze, on the other hand, died within 3 or 4 days.
A second video, this time of wheat roots (Supplemental Video 2), showed an interesting sequence of freezing. The base of these roots was submerged in growing medium consisting primarily of peat. Ice shavings were added prior to the freezing to ensure the roots would freeze. Freeze nucleation occurred at about -0.5 &#176;C midway along a root on the right side. The freezing then progressed upwards to the crown of the plant causing the base of the outer leaves to freeze. The freezing then progressed downwards in the roots at the back of the plant. Note that, without the 3D perspective, it is nearly impossible to determine the order in which the specific roots froze (Figure 2).
When considering freezing in roots (Figure 2 and Supplemental Video 2), if only a 2D perspective was viewed, it would be nearly impossible to determine which root was freezing due to the lack of information about depth. The 3D perspective of this freeze event represents the event as it occurred in the real world and greatly improves the ability of the viewer to distinguish the sequence of freezing in individual roots.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/58025/58025fig1.jpg" /> 
Figure 1: A comparison of an image of strawberries in 2D to the same image in 3D. These images are freeze-frames from Supplemental Video 1 showing 2 leaves and a single flower of a strawberry plant freezing. (A) This panel shows the left view only, in 2D. (B) This panel shows the 3D anaglyph view. Red-blue glasses must be worn to see this image in true 3D. A comparison between the two panels illustrates the improvement in visual perception when the subject has been captured in 3D. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58025/58025fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/58025/58025fig2.jpg" /> 
Figure 2. A comparison of images of a root mass of wheat in 2D to the same image in 3D. These images are freeze-frames from Supplemental Video 2. Panels A and B show the root mass in 2D. (A) This is an image of the roots prior to freezing, while (B) this is about midway through the freezing event. Panels C and D show the same images as Panels A and B but in anaglyph format. (C) This panel shows the root mass prior to the freezing (corresponding to panel A). (B) This is an image of the roots at the same point in the freeze event as in panel D. Panels C and D must be viewed with red-blue glasses to see the images in 3D. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58025/58025fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Supplemental Video 1: A red-blue anaglyph video showing freezing in a strawberry plant in 3D. This video was generated using the protocol demonstrated here. Note that red-blue glasses are necessary to observe the video in 3D. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58025/Supplimental_Video_1.mov">Please click here to download this file.</a>
Supplemental Video 2: A red-blue anaglyph video showing freezing in wheat roots in 3D. This video was generated using the protocol demonstrated here. Note that red-blue glasses are necessary to observe the video in 3D. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58025/Supplimental_Video_2.mov">Please click here to download this file.</a>

Watch this Scientific Journal Video about Protocol for Producing Three-Dimensional Infrared Video of Freezing in Plants at JoVE.com

Protocol for Producing Three-Dimensional Infrared Video of Freezing in Plants

Here, we present a protocol to image a strawberry plant freezing in 3 dimensions. Two infrared cameras positioned at slightly different angles are used to produce a red-blue anaglyph video to observe the freezing of the plant in 3 dimensions.

Freezing in plants can be monitored using infrared (IR) thermography, because when water freezes, it gives off heat. However, problems with color contrast ...

protocol-for-producing-three-dimensional-infrared-video-freezing

Nanyang Technological University

Research

JoVE Journal

Environment

6.1K Views.  United States Department of Agriculture and North Carolina State University. Hier presenteren we een protocol om de afbeelding van een aardbei plant bevriezing in 3 dimensies. Twee infrarood camera's geplaatst in iets verschillende hoeken worden gebruikt om een rood-blauw anaglyph video om te zien hoe de bevriezing van de fabriek in 3 dimensies te produceren.

Video: Protocol for Producing Three-Dimensional Infrared Video of Freezing in Plants

The Use of High-resolution Infrared Thermography (HRIT) for the Study of Ice Nucleation and Ice Propagation in Plants

Freezing events that occur when plants are actively growing can be a lethal event, particularly if the plant has no freezing tolerance. Such frost events often have devastating effects on agricultural production and can also play an important role in shaping community structure in natural populations of plants, especially in alpine, sub-arctic, and arctic ecosystems. Therefore, a better understanding of the freezing process in plants can play an important role in the development of methods of frost protection and understanding mechanisms of freeze avoidance. Here, we describe a protocol to visualize the freezing process in plants using high-resolution infrared thermography (HRIT). The use of this technology allows one to determine the primary sites of ice formation in plants, how ice propagates, and the presence of ice barriers. Furthermore, it allows one to examine the role of extrinsic and intrinsic nucleators in determining the temperature at which plants freeze and evaluate the ability of various compounds to either affect the freezing process or increase freezing tolerance. The use of HRIT allows one to visualize the many adaptations that have evolved in plants, which directly or indirectly impact the freezing process and ultimately enables plants to survive frost events.

The Use of High-resolution Infrared Thermography HRIT for the Study of Ice Nucleation and Ice Propagation in Plants

Here we present a protocol that allows one to visualize sites of ice formation and avenues of ice propagation in plants utilizing high resolution infrared thermography (HRIT).

Freezing events that occur when plants are actively growing can be a lethal event, particularly if the plant has no freezing tolerance. Such frost events ...

RGB and Spectral Root Imaging for Plant Phenotyping and Physiological Research: Experimental Setup and Imaging Protocols

Better understanding of plant root dynamics is essential to improve resource use efficiency of agricultural systems and increase the resistance of crop cultivars against environmental stresses. An experimental protocol is presented for RGB and hyperspectral imaging of root systems. The approach uses rhizoboxes where plants grow in natural soil over a longer time to observe fully developed root systems. Experimental settings are exemplified for assessing rhizobox plants under water stress and studying the role of roots. An RGB imaging setup is described for cheap and quick quantification of root development over time. Hyperspectral imaging improves root segmentation from the soil background compared to RGB color based thresholding. The particular strength of hyperspectral imaging is the acquisition of chemometric information on the root-soil system for functional understanding. This is demonstrated with high resolution water content mapping. Spectral imaging however is more complex in image acquisition, processing and analysis compared to the RGB approach. A combination of both methods can optimize a comprehensive assessment of the root system. Application examples integrating root and aboveground traits are given for the context of plant phenotyping and plant physiological research. Further improvement of root imaging can be obtained by optimizing RGB image quality with better illumination using different light sources and by extension of image analysis methods to infer on root zone properties from spectral data.

An experimental protocol is presented for assessment of soil grown plant root systems with RGB and hyperspectral imaging. Combination of RGB image time series with chemometric information from hyperspectral scans optimizes insights into plant root dynamics.

Better understanding of plant root dynamics is essential to improve resource use efficiency of agricultural systems and increase the resistance of crop ...

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

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 ...

Tandem High-pressure Freezing and Quick Freeze Substitution of Plant Tissues for Transmission Electron Microscopy

Since the 1940s transmission electron microscopy (TEM) has been providing biologists with ultra-high resolution images of biological materials. Yet, because of laborious and time-consuming protocols that also demand experience in preparation of artifact-free samples, TEM is not considered a user-friendly technique. Traditional sample preparation for TEM used chemical fixatives to preserve cellular structures. High-pressure freezing is the cryofixation of biological samples under high pressures to produce very fast cooling rates, thereby restricting ice formation, which is detrimental to the integrity of cellular ultrastructure. High-pressure freezing and freeze substitution are currently the methods of choice for producing the highest quality morphology in resin sections for TEM. These methods minimize the artifacts normally associated with conventional processing for TEM of thin sections. After cryofixation the frozen water in the sample is replaced with liquid organic solvent at low temperatures, a process called freeze substitution. Freeze substitution is typically carried out over several days in dedicated, costly equipment. A recent innovation allows the process to be completed in three hours, instead of the usual two days. This is typically followed by several more days of sample preparation that includes infiltration and embedding in epoxy resins before sectioning. Here we present a protocol combining high-pressure freezing and quick freeze substitution that enables plant sample fixation to be accomplished within hours. The protocol can readily be adapted for working with other tissues or organisms. Plant tissues are of special concern because of the presence of aerated spaces and water-filled vacuoles that impede ice-free freezing of water. In addition, the process of chemical fixation is especially long in plants due to cell walls impeding the penetration of the chemicals to deep within the tissues. Plant tissues are therefore particularly challenging, but this protocol is reliable and produces samples of the highest quality.

Obtaining high-quality transmission electron microscopy images is challenging, especially in the case of plant cells, which have abundant large water-filled vacuoles and aerated spaces. Tandem high-pressure freezing and quick freeze substitution greatly reduce preparation time of plant samples for TEM while producing samples with excellent ultrastructural preservation.

Since the 1940s transmission electron microscopy (TEM) has been providing biologists with ultra-high resolution images of biological materials. Yet, ...

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 ...

A Versatile Method for Mounting Arabidopsis Leaves for Intravital Time-lapse Imaging

The plant immune response associated with a genome-wide transcriptional reprogramming is initiated at the site of infection. Thus, the immune response is regulated spatially and temporally. The use of a fluorescent gene under the control of an immunity-related promoter in combination with an automated fluorescence microscopy is a simple way to understand spatiotemporal regulation of plant immunity. In contrast to the root tissues that have been used for a number of various intravital fluorescent imaging experiments, there exist few fluorescent live-imaging examples for the leaf tissues that encounter an array of airborne microbial infections. Therefore, we developed a simple method to mount leaves of Arabidopsis thaliana plants for live-cell imaging over an extended period of time. We used transgenic Arabidopsis plants expressing the yellow fluorescent protein (YFP) genefused to the nuclear localization signal (NLS) under the control of the promoter of a defense-related marker gene, Pathogenesis-Related 1 (PR1). We infiltrated a transgenic leaf with Pseudomonas syringae pv. tomato DC3000 (avrRpt2) strain (Pst_a2) and performed in vivo time-lapse imaging of the YFP signal for a total of 40 h using an automated fluorescence stereomicroscope. This method can be utilized not only for studies on plant immune responses but also for analyses of various developmental events and environmental responses occurring in leaf tissues.

A Versatile Method for Mounting Arabidopsis Leaves for Intravital Time-lapse Imaging

We report a simple and versatile method for performing fluorescent live-imaging of Arabidopsis thaliana leaves over an extended period of time. We use a transgenic Arabidopsis plant expressing a fluorescent reporter gene under the control of an immunity-related promoter as an example for gaining spatiotemporal understanding of plant immune responses.

The plant immune response associated with a genome-wide transcriptional reprogramming is initiated at the site of infection. Thus, the immune response is ...

Identification of Novel Regulators of Plant Transpiration by Large-Scale Thermal Imaging Screening in Helianthus Annuus

Plant adaptation to biotic and abiotic stresses is governed by a variety of factors, among which the regulation of stomatal aperture in response to water deficit or pathogens plays a crucial role. Identifying small molecules that regulate stomatal movement can therefore contribute to understanding the physiological basis by which plants adapt to their environment. Large-scale screening approaches that have been used to identify regulators of stomatal movement have potential limitations: some rely heavily on the abscisic acid (ABA) hormone signaling pathway, therefore excluding ABA-independent mechanisms, while others rely on the observation of indirect, long-term physiological effects such as plant growth and development. The screening method presented here allows the large-scale treatment of plants with a library of chemicals coupled with a direct quantification of their transpiration by thermal imaging. Since evaporation of water through transpiration results in leaf surface cooling, thermal imaging provides a non-invasive approach to investigate changes in stomatal conductance over time. In this protocol, Helianthus annuus seedlings are grown hydroponically and then treated by root feeding, in which the primary root is cut and dipped into the chemical being tested. Thermal imaging followed by statistical analysis of cotyledonary temperature changes over time allows for the identification of bioactive molecules modulating stomatal aperture. Our proof-of-concept experiments demonstrate that a chemical can be carried from the cut root to the cotyledon of the sunflower seedling within 10 minutes. In addition, when plants are treated with ABA as a positive control, an increase in leaf surface temperature can be detected within minutes. Our method thus allows the efficient and rapid identification of novel molecules regulating stomatal aperture.

Identification of Novel Regulators of Plant Transpiration by Large-Scale Thermal Imaging Screening in Helianthus Annuus

We provide a method for identifying modulators of foliar transpiration by large-scale screening of a compound library.

Plant adaptation to biotic and abiotic stresses is governed by a variety of factors, among which the regulation of stomatal aperture in response to water ...

Optical Clearing of Plant Tissues for Fluorescence Imaging

It is challenging to directly observe the internal structure of multi-layered and opaque plant specimens, without dissection, under a microscope. In addition, autofluorescence attributed to chlorophyll hampers the observation of fluorescent proteins in plants. For a long time, various clearing reagents have been used to make plants transparent. However, conventional clearing reagents diminish fluorescent signals; therefore, it has not been possible to observe the cellular and intracellular structures with fluorescent proteins. Reagents were developed that can clear plant tissues by removing chlorophyll while maintaining fluorescent protein stability. A detailed protocol is provided here for the optical clearing of plant tissues using clearing reagents, ClearSee (CS) or ClearSeeAlpha (CSA). The preparation of cleared plant tissues involves three steps: fixation, washing, and clearing. Fixation is a crucial step in maintaining the cellular structures and intracellular stability of fluorescent proteins. The incubation time for clearing depends on the tissue type and species. In Arabidopsis thaliana, the time required for clearing with CS was 4 days for leaves and roots, 7 days for seedlings, and 1 month for pistils. CS also required a relatively short time of 4 days to make the gametophytic leaves of Physcomitrium patens transparent. In contrast, pistils in tobacco and torenia produced brown pigment due to oxidation during CS treatment. CSA reduced the brown pigment by preventing oxidation and could make tobacco and torenia pistils transparent, although it took a relatively long time (1 or 2 months). CS and CSA were also compatible with staining using chemical dyes, such as DAPI (4',6-diamidino-2-phenylindole) and Hoechst 33342 for DNA and Calcofluor White, SR2200, and Direct Red 23 for the cell wall. This method can be useful for whole-plant imaging to reveal intact morphology, developmental processes, plant-microbe interactions, and nematode infections.

Here, a method is described for making plant tissues transparent while maintaining the stability of fluorescent proteins. This technique facilitates deep imaging of cleared plant tissues without physical sectioning.

It is challenging to directly observe the internal structure of multi-layered and opaque plant specimens, without dissection, under a microscope. In ...

Protocol for Producing Three-Dimensional Infrared Video of Freezing in Plants

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