Funding was provided by a Fiske Award and a Wellesley College Faculty Award.

1. Preparing leaves for a run
<ol>
	<li>Place 8 A. thaliana plants in the dark overnight (&#62; 6 h works for most species) to ensure that the chloroplasts move into their dark position. All replicas start with comparable transmission values.</li>
	<li>Alternatively, place 8 complete leaves into a Petri dish with a moist filter paper at the bottom, close the Petri dish, and wrap it with aluminum foil.</li>
</ol>
2. Testing if the transmission device works
<ol>
	<li>Connect the transmission device to a stable power source and press the power switch (ON/OFF button) of the device to reset the instrument (Figure 1A,B).</li>
	<li>Connect the iPad to a stable power source, press the Home Screen button to activate the screen, and enter the passcode to log in.</li>
	<li>Press the Settings icon, press the Display &#38; Brightness icon, press Auto-Lock, and select Never by pressing this option to ensure that the screen stays on permanently. Otherwise the program will stop running when the screen goes to sleep. Press the Home Screen button to return to the main screen.</li>
	<li>Double press the Home Screen button to see which applications are open, and close all of them by swiping them towards the top of the screen. Press the Home Screen button to return to the main screen.
	<ol>
		<li>Find the LeafSensor app on the main screen or by swiping left or right. Press the icon of the app to open it (see Supplementary File). A green screen with text and white fields will appear to enter the information.</li>
		<li>Ensure the word Connected is seen in the lower part of the screen as it indicates that the app is communicating with the transmission device. If the message &#39;Adafruit NOT Found&#39; appears, check that the device is plugged in and press the start button on the device again.</li>
	</ol>
	</li>
	<li>Fill in the first 4 fields on the app page to name the experiment and to set up the conditions for a brief test run without leaves and with the leaf clips open:
	<ol>
		<li>Name the experiment (use 8 or fewer uppercase letters or numbers) e.g., by typing TEST into the field named Expt Name.</li>
		<li>Choose how many different blue light intensities will be used in the experiment, e.g., by typing 3 into the field named # Light Intensities.</li>
		<li>Choose the blue light intensities for this run (choose an integer between 0 and 3000 and separate each number from the next with a comma; see Supplementary File on how to convert these numbers into actual blue light intensities of LEDs) e.g., by typing 0,100,1000 into the field named Blue Intensities.</li>
		<li>Choose the length of time each blue light intensity will shine onto the leaf (separate each number from the next with a comma) e.g., by typing 2,2,2 into the field named Blue Duration (minutes).</li>
	</ol>
	</li>
	<li>Press Start Experiment in the middle section of the screen. In the lower part of the screen 8 hyphens and the message Starting Experiment will appear.
	<ol>
		<li>Ensure that no light is being emitted from the LEDs for the first two minutes, then weak blue light is being emitted, and after 2 min the blue light intensity is increasing.</li>
		<li>Ensure that intense red light is being emitted from the LEDs once a minute for the measurements. 
		&#8203;NOTE: As the experiment is running, numbers will appear on the app page for each of the eight sensors, and data will be updated once a minute. Ensure the output numbers from the photodiodes are around 1000-1023 (if the room is dark). An update on the lower left shows how many measurements were done so far e.g., Done 1 of 6 measurements (one measurement each minute).</li>
		<li>When the experiment is done, check for the appearance of the Experiment finished on the lower left of the app page. Now the instrument is ready for a run with leaves.</li>
	</ol>
	</li>
	<li>Press twice the Home Screen button, swipe out of the app and open it again. Press the ON/OFF button on the instrument to reset it.</li>
</ol>
3. Setting up leaves in the leaf clips
NOTE: This step has to be done in the dark with a green light source (e.g., place a green filter in front of a light bulb) to avoid inducing chloroplast movement. Alternatively, use very low white light and an extended dark period in the leaf clips. Remember, one part of the leaf clip holds the LED (larger opening), while the other holds the phototransistor (Figure 1C).
<ol>
	<li>If the plants are dark-adapted entire, pick 8 leaves wide enough to cover the LEDs. Otherwise, remove the leaves from the Petri dish. Prepare 8 strips of filter paper about the length of a leaf clip and with a hole punched out at the top to not cover the LED.
	<ol>
		<li>Moisten the filter paper and place it onto the leaf clip part that holds the LED. Repeat this for each of the eight leaf clips.</li>
	</ol>
	</li>
	<li>Place each leaf on the top of the wet filter paper of its leaf clip. Ensure the correct leaf side is facing towards the LED (usually experiments are done with the adaxial leaf surface facing the LED).
	<ol>
		<li>Avoid placing the midribs of the leaf on top of the LEDs, and for more consistent results, place similar parts of each leaf (e.g., the widest section of the leaf) over the LED.</li>
		<li>Place the other leaf clip part with the phototransistor on top. Use a rubber band to hold the two leaf clip parts together if needed (Figure 1C,D).</li>
	</ol>
	</li>
	<li>Place each leaf clip into its &#39;boat&#39; and fill the reservoirs with water using a pipette. Ensure the leaf or at least the filter paper touches the water to avoid dehydration of the leaves during the run (Figure 1D).</li>
</ol>
4. Conducting a run
NOTE: For a standard exploratory run, start out with 4 h of darkness (0 &#181;mol photon m-2 s-1), followed by 7 h of low blue light (2 &#181;mol photon m-2 s-1), followed by 60 min each of 5, 10, 30, 40, 50, 60, 90, 100 &#181;mol photon m-2 s-1 of blue light. This will induce the leaves to exhibit their dark transmission, induce chloroplast movement into the maximum accumulation, and show different degrees of avoidance response.
<ol>
	<li>Set up the LeafSensor app on the iPad using the steps described below.
	<ol>
		<li>Type EXPLORA1 into the field named Expt Name.</li>
		<li>Type 10 into the field named # Light Intensities.</li>
		<li>Type 0,1,60,160,550,750,950,1150,1350,1950 into the field named Blue intensities.</li>
		<li>Type 240,420,60,60,60,60,60,60,60,60 into the field named Blue Duration.</li>
	</ol>
	</li>
	<li>Press Start Experiment. After the first min, the output values (usually between 990 and 820) will appear on the screen. If the values are far off, check if the leaves were placed correctly into the leaf clips.</li>
	<li>When the run is done, ensure that the message Experiment Finished appears at the bottom left of the screen. Data will be saved automatically.
	<ol>
		<li>Put the screen in the upright position (not horizontal). Two new options will appear on the screen, namely Save and Utilities.</li>
		<li>Press Utilities and a list of saved files will appear. Select the file of interest, in this case, EXPLORA1.</li>
		<li>Below the list of files , look for Selected Expt: EXPLORA1. Press Email, enter an email address, and the data file will be automatically attached to the message. Press Send. 
		NOTE: If there is a long delay until the files arrive, restart the app and send the file again.</li>
	</ol>
	</li>
	<li>If a run was aborted, but data up to that point are of interest, press Save before selecting Utilities. After several runs, clean up the memory space: Press Utilities, select one file at a time, swipe left next to file, and press Delete to remove file.</li>
	<li>Press Back to run another experiment or if done, press the Home Screen button, swipe to the main screen, press Settings, press Display &#38; Brightness, press Auto-Lock, and press 2 Minutes.</li>
</ol>
5. Data analysis
<ol>
	<li>Download the file EXPLORA1 from the email, add extension .csv to the file, double-click on the file. Data will be sorted into a spreadsheet with the data for the eight different sensors sorted into separate columns. The last column shows the time (in seconds) at which the data were collected. Delete the first row under the headings (Sensor1-8) if it contains nonsensical data.</li>
	<li>Set up a master data sheet that will contain the results for each sensor in a separate sheet and that converts the output values into % transmission values using the equations obtained from the calibration of each leaf clip and sensor (see Supplementary File).
	<ol>
		<li>Copy each data set into a separate data sheet (e.g., column A contains the time; column C contains the data of Sensor1).</li>
		<li>Set up column B so that it converts time from seconds to minutes. Set up column D so that it contains the formula to convert voltage to % transmission using the equation from the calibration.</li>
		<li>Set up a similar data sheet for each sensor and remember the formula used to convert the voltage output into % transmission values may be different for each sensor.</li>
	</ol>
	</li>
	<li>Plot % transmission (T) against time, min (Figure 2).</li>
	<li>Save the data sheet under a new name so that the master data sheet can be reused.</li>
	<li>To further analyze data (Figure 2), calculate &#916;T (%) accumulation (e.g., change of T at maximum accumulation compared to T at dark), &#916;T (%) avoidance (e.g., change of T at maximum avoidance compared to T at dark), or dT/dt (%/h) (e.g., change in T during the fastest part of the accumulation or avoidance reaction). For further details see 8.</li>
</ol>

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R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+phototropin+family+of+photoreceptors.">The phototropin family of photoreceptors.</a> Plant Cell. 13, 993-997 (2001).</li><li>Jarillo, J. A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Phototropin-related+NPL1+controls+chloroplast+relocation+induced+by+blue+light.">Phototropin-related NPL1 controls chloroplast relocation induced by blue light.</a> Nature. 410, 952-954 (2001).</li><li>Sakai, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Arabidopsis+nph1+and+npl1:+blue+light+receptors+that+mediate+both+phototropism+and+chloroplast+relocation.">Arabidopsis nph1 and npl1: blue light receptors that mediate both phototropism and chloroplast relocation.</a> Proceedings of the National Academy of Sciences, USA. 98 (12), 6969-6974 (2001).</li><li>Wada, M., Kong, S. -. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Actin-mediated+movement+of+chloroplasts.">Actin-mediated movement of chloroplasts.</a> Journal of Cell Science. 131, 1-8 (2018).</li><li>Wen, F., Xing, D., Zhang, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Hydrogen+peroxide+is+involved+in+high+blue+light-induced+chloroplast+avoidance+movements+in+Arabidopsis.">Hydrogen peroxide is involved in high blue light-induced chloroplast avoidance movements in Arabidopsis.</a> Journal of Experimental Botany. 59 (10), 2891-2901 (2008).</li><li>Tlalka, M., Fricker, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+role+of+calcium+in+blue-light+dependent+chloroplast+movement+in+Lemna+trisulca+L.">The role of calcium in blue-light dependent chloroplast movement in Lemna trisulca L.</a> The Plant Journal. 20, 461-473 (1999).</li><li>K&#246;niger, M., Bollinger, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Chloroplast+movement+behavior+varies+widely+among+species+and+does+not+correlate+with+high+light+stress+tolerance.">Chloroplast movement behavior varies widely among species and does not correlate with high light stress tolerance.</a> Planta. 236, 411-426 (2012).</li><li>Higa, T., Wada, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Chloroplast+avoidance+movement+is+not+functional+in+plants+grown+under+strong+sunlight.">Chloroplast avoidance movement is not functional in plants grown under strong sunlight.</a> Plant, Cell and Environment. 39, 871-882 (2016).</li><li>. 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The authors have no conflicts of interest.

The device is extremely easy to use but it is of crucial importance to calibrate each leaf clip set-up of the transmission device independently since the positioning of the LEDs and phototransistors may slightly vary from leaf clip to leaf clip. Ensure the LEDs and phototransistors are inserted stably and re-check the calibration if the data seem off. Avoid getting water onto the device. The leaves in the leaf clips are placed into &#39;boats&#39; filled with water to avoid water stress. Place these boats e.g., into a lo...

Light is essential for photosynthesis, plant growth, and development. It is one of the most dynamic abiotic factors as light intensities not only change over the course of a season or day, but also rapidly and in unpredictable ways depending on the cloud cover. At the leaf level, light intensities are also influenced by the density and nature of the surrounding vegetation and the plant&#39;s own canopy. One important mechanism that allows plants to optimize light interception under variable light conditions is the ability of chloroplasts to move in response to blue light stimuli1,2. Under low light conditions, chloroplasts spread out perpendicular to the light (along the periclinal cell walls) in a so-called accumulation response, maximizing light interception and hence photosynthesis. Under high light conditions, chloroplasts move towards the anticlinal cell wall in a so-called avoidance response, minimizing light interception and the danger of photoinhibition. In many species, chloroplasts also assume a specific dark position, which is distinct from the accumulation and avoidance positions and often intermediary between those two3,4. Various studies have demonstrated that chloroplast movement is not only important for the short-term stress tolerance of leaves5,6,7, but also for the growth and reproductive success of plants, especially under variable light conditions8,9.
Chloroplast movement is readily observed in real time in certain live specimens (e.g., algae or thin-leafed plants like Elodea) using light microscopy1. Studying chloroplast movement in most leaves, however, requires a pretreatment to induce chloroplast movement, chemical fixation, and preparation of cross sections before viewing the samples under a light microscope10. With the introduction of confocal laser microscopy, it also became possible to image the 3D arrangement of chloroplasts in intact or fixed leaves4,11,12. These imaging techniques greatly aid the understanding of chloroplast movement by providing important qualitative information. Quantifying chloroplast positioning (e.g., as a percentage of chloroplasts in the periclinal or anticlinal positions in these images or the percentage of area covered by chloroplasts per total cell surface) is also possible but quite time-consuming, especially if conducted at the intervals necessary to capture rapid changes in positioning10,8. The simplest way to show whether dark-adapted leaves of a certain species or mutants are capable of chloroplast movement into the avoidance response is by covering most of the area of a leaf to keep the chloroplasts in the dark while exposing a strip of the leaf to high light. After a minimum of 20 min of high light exposure, the chloroplasts in the exposed area will have moved into the avoidance position, and the exposed strip will be visibly lighter in color than the rest of the leaf. This is true for wild type A. thaliana but not for some of the chloroplast movement mutants described in more detail later on13. This method and modifications of it (e.g., reversing what parts of the leaf are exposed, changing light intensities) are useful for screening large numbers of mutants and to identify null mutants that lack either the ability to exhibit an avoidance or accumulation response or both. However, it does not provide information about the dynamic changes in chloroplast movement.
In contrast, the method described here allows for the quantification of chloroplast movement in intact leaves using changes in light transmission through a leaf as a proxy for overall chloroplast movement: under conditions when chloroplasts are spread out in the mesophyll cells in the accumulation response, less light is transmitted through the leaf than when many chloroplasts are in an avoidance response, positioning themselves along the anticlinal cell walls. Hence, changes in transmission can be used as a proxy for the overall chloroplast movement in leaves14. The details of the instrument are described elsewhere (see Supplementary File), but basically, the instrument uses blue light to trigger chloroplast movement and measures how much red light is transmitted through that leaf at set intervals.&#160;More recently, a modification of this system has been described, which uses a modified 96-well microplate reader, a blue LED, a computer, and a temperature-controlled incubator15.
The option to use a combination of methods, including the optical assessment of leaves for screening, followed by measuring dynamic changes in transmission and the use of microscopy, has greatly aided our understanding of both the underlying mechanisms and the physiological/ecological significance of chloroplast movement. For example, it led to the discovery and characterization of various mutants, which are impaired in specific aspects of their movements. For example, A. thaliana phot 1 mutants lack the ability to accumulate their chloroplasts in low light, while phot 2 mutants lack the ability to perform an avoidance reaction. These phenotypes are due to an impairment in two respective blue light receptors16,17,18. In contrast, chup1 mutants lack the ability to form proper actin filaments around the chloroplasts which are essential to move the chloroplasts into the desired position within a cell11,19. In addition to mutant studies, researchers have assessed the effects of various inhibitors on chloroplast movement to elucidate the mechanistic aspects of the process. For example, chemicals such as H2O2 and various antioxidants were used to investigate the effects of this signaling molecule on chloroplast movement20. Various inhibitors were used to elucidate the role of calcium in chloroplast movement21. In addition to helping to uncover the mechanisms of chloroplast movement, these methods can be used to compare chloroplast movement in various species or mutants grown in different conditions in an attempt to understand the ecological and evolutionary context of this behavior. For example, it has been shown that the extent of the effects of various mutations in the chloroplast movement pathway are dependent on the growth conditions7,9, and that sun-adapted plants do not seem to move their chloroplasts much. In contrast, movement is very important for shade plants10,22,23.
This methods paper, focused on the model plant A. thaliana, describes how to use a transmission device which is an updated version of a previously developed instrument9. While this instrument is not commercially available, people with a basic understanding of electronics or the help of engineering or physics colleagues and students will be able to build the instrument using affordable parts and following the detailed instructions (see Supplementary File). The open-source platform used to build the instrument has extensive web support and a community forum which offers help should problems arise24.
The protocol focuses on how to use the instrument to determine changes in leaf transmission in a standard exploratory run that exposes a leaf to a wide range of light conditions and captures the dark, accumulation, and avoidance reactions of A. thaliana. These runs can be modified depending on the goal of the experiment and can be used with most plant species. The paper provides examples of transmission data of A. thaliana wildtype and several mutants and shows how to further analyze the data.

<table><tbody><tr><td>Aluminum foil</td><td></td><td></td><td></td></tr><tr><td>Dark adapted leaves</td><td></td><td></td><td></td></tr><tr><td>Filter paper</td><td></td><td></td><td></td></tr><tr><td>iPad with &lt;em&gt;LeafSensor &lt;/em&gt;app installed (see Supplemental Info)</td><td></td><td></td><td></td></tr><tr><td>Pipette</td><td>Any</td><td></td><td></td></tr><tr><td>Petri dish</td><td>Any</td><td></td><td></td></tr><tr><td>Transmission device (see Supplemental info)</td><td></td><td></td><td></td></tr><tr><td>Water</td><td></td><td></td><td></td></tr></tbody></table>

using changes in leaf transmission to investigate chloroplast movement in arabidopsis thaliana

Chloroplast movement in leaves has been shown to help minimize photoinhibition and increase growth under certain conditions. Much can be learned about chloroplast movement by studying the chloroplast positioning in leaves using e.g., confocal fluorescence microscopy, but access to this type of microscopy is limited. This protocol describes a method that uses the changes in leaf transmission as a proxy for chloroplast movement. If chloroplasts are spread out in order to maximize light interception, the transmission will be low. If chloroplasts move towards the anticlinal cell walls to avoid light, the transmission will be higher. This protocol describes how to use a straightforward, home-built instrument to expose leaves to different blue light intensities and quantify the dynamic changes in leaf transmission. This approach allows researchers to quantitatively describe chloroplast movement in different species and mutants, study the effects of chemicals and environmental factors on it, or screen for novel mutants e.g., to identify missing components in the process that leads from light perception to the movement of chloroplasts.

The different parts of the transmission device are shown in Figure 1. The microcontroller is the control unit of the device and controls the light conditions that the leaves, secured in black leaf clips, are experiencing, and stores the light transmission data it receives (Figure 1A,B). A close-up of the control unit of the instrument shows the ON/OFF button, the SD card for data storage capability, the Bluetooth shield (which sends the data to ...

Watch this Scientific Journal Video about Using Changes in Leaf Transmission to Investigate Chloroplast Movement in Arabidopsis thaliana at JoVE.com

Using Changes in Leaf Transmission to Investigate Chloroplast Movement in Arabidopsis thaliana

Many plant species change the positioning of chloroplasts to optimize light absorption. This protocol describes how to use a straightforward, home-built instrument to investigate chloroplast movement in Arabidopsis thaliana leaves using changes in the transmission of light through a leaf as a proxy.

Using Changes in Leaf Transmission to Investigate Chloroplast Movement in Arabidopsis thaliana

Chloroplast movement in leaves has been shown to help minimize photoinhibition and increase growth under certain conditions. Much can be learned about ...

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2.2K Views.  Wellesley College. Many plant species change the positioning of chloroplasts to optimize light absorption. This protocol describes how to use a straightforward, home-built instrument to investigate chloroplast movement in Arabidopsis thaliana leaves using changes in the transmission of light through a leaf as a proxy.

Video: Using Changes in Leaf Transmission to Investigate Chloroplast Movement in Arabidopsis thaliana

Investigating Tissue- and Organ-specific Phytochrome Responses using FACS-assisted Cell-type Specific Expression Profiling in Arabidopsis thaliana

Light mediates an array of developmental and adaptive processes throughout the life cycle of a plant. Plants utilize light-absorbing molecules called photoreceptors to sense and adapt to light. The red/far-red light-absorbing phytochrome photoreceptors have been studied extensively. Phytochromes exist as a family of proteins with distinct and overlapping functions in all higher plant systems in which they have been studied1. Phytochrome-mediated light responses, which range from seed germination through flowering and senescence, are often localized to specific plant tissues or organs2. Despite the discovery and elucidation of individual and redundant phytochrome functions through mutational analyses, conclusive reports on distinct sites of photoperception and the molecular mechanisms of localized pools of phytochromes that mediate spatial-specific phytochrome responses are limited. We designed experiments based on the hypotheses that specific sites of phytochrome photoperception regulate tissue- and organ-specific aspects of photomorphogenesis, and that localized phytochrome pools engage distinct subsets of downstream target genes in cell-to-cell signaling. We developed a biochemical approach to selectively reduce functional phytochromes in an organ- or tissue-specific manner within transgenic plants. Our studies are based on a bipartite enhancer-trap approach that results in transactivation of the expression of a gene under control of the Upstream Activation Sequence (UAS) element by the transcriptional activator GAL43. The biliverdin reductase (BVR) gene under the control of the UAS is silently maintained in the absence of GAL4 transactivation in the UAS-BVR parent4. Genetic crosses between a UAS-BVR transgenic line and a GAL4-GFP enhancer trap line result in specific expression of the BVR gene in cells marked by GFP expression4. BVR accumulation in Arabidopsis plants results in phytochrome chromophore deficiency in planta5-7. Thus, transgenic plants that we have produced exhibit GAL4-dependent activation of the BVR gene, resulting in the biochemical inactivation of phytochrome, as well as GAL4-dependent GFP expression. Photobiological and molecular genetic analyses of BVR transgenic lines are yielding insight into tissue- and organ-specific phytochrome-mediated responses that have been associated with corresponding sites of photoperception4, 7, 8. Fluorescence Activated Cell Sorting (FACS) of GFP-positive, enhancer-trap-induced BVR-expressing plant protoplasts coupled with cell-type-specific gene expression profiling through microarray analysis is being used to identify putative downstream target genes involved in mediating spatial-specific phytochrome responses. This research is expanding our understanding of sites of light perception, the mechanisms through which various tissues or organs cooperate in light-regulated plant growth and development, and advancing the molecular dissection of complex phytochrome-mediated cell-to-cell signaling cascades.

Investigating Tissue- and Organ-specific Phytochrome Responses using FACS-assisted Cell-type Specific Expression Profiling in Arabidopsis thaliana

The molecular basis of spatial-specific phytochrome responses is being investigated using transgenic plants that exhibit tissue- and organ-specific phytochrome deficiencies. The isolation of specific cells exhibiting induced phytochrome chromophore depletion by Fluorescence-Activated Cell Sorting followed by microarray analyses is being utilized to identify genes involved in spatial-specific phytochrome responses.

Light mediates an array of developmental and adaptive processes throughout the life cycle of a plant. Plants utilize light-absorbing molecules called ...

Biology

Evaluation of Photosynthetic Behaviors by Simultaneous Measurements of Leaf Reflectance and Chlorophyll Fluorescence Analyses

Chlorophyll a fluorescence analysis is widely used to measure photosynthetic behaviors in intact plants, and has resulted in the development of many parameters that efficiently measure photosynthesis. Leaf reflectance analysis provides several vegetation indices in ecology and agriculture, including the photochemical reflectance index (PRI), which can be used as an indicator of thermal energy dissipation during photosynthesis because it correlates with non-photochemical quenching (NPQ). However, since NPQ is a composite parameter, its validation is required to understand the nature of the PRI parameter. To obtain physiological evidence for evaluation of the PRI parameter, we simultaneously measured chlorophyll fluorescence and leaf reflectance in xanthophyll cycle defective mutant (npq1) and wild-type Arabidopsis plants. Additionally, the qZ parameter, which likely reflects the xanthophyll cycle, was extracted from the results of chlorophyll fluorescence analysis by monitoring relaxation kinetics of NPQ after switching the light off. These simultaneous measurements were carried out using a pulse-amplitude modulation (PAM) chlorophyll fluorometer and a spectral radiometer. The fiber probes from both instruments were positioned close to each other to detect signals from the same leaf position. An external light source was used to activate photosynthesis, and the measuring lights and saturated light were provided from the PAM instrument. This experimental system enabled us to monitor light-dependent PRI in the intact plant and revealed that light-dependent changes in PRI differ significantly between the wild type and npq1 mutant. Furthermore, PRI was strongly correlated with qZ, meaning that qZ reflects the xanthophyll cycle. Together, these measurements demonstrated that simultaneous measurement of leaf reflectance and chlorophyll fluorescence is a valid approach for parameter evaluation.

We describe a new technical approach to study photosynthetic responses in higher plants involving simultaneous measurements of chlorophyll a fluorescence and leaf reflectance using a PAM and a spectral radiometer for the detection of signals from the same leaf area in Arabidopsis.

Chlorophyll a fluorescence analysis is widely used to measure photosynthetic behaviors in intact plants, and has resulted in the development of many ...

Bioengineering

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 ...

Cell Specific Analysis of Arabidopsis Leaves Using Fluorescence Activated Cell Sorting

After initiation of the leaf primordium, biomass accumulation is controlled mainly by cell proliferation and expansion in the leaves1. However, the Arabidopsis leaf is a complex organ made up of many different cell types and several structures. At the same time, the growing leaf contains cells at different stages of development, with the cells furthest from the petiole being the first to stop expanding and undergo senescence1. Different cells within the leaf are therefore dividing, elongating or differentiating; active, stressed or dead; and/or responding to stimuli in sub-sets of their cellular type at any one time. This makes genomic study of the leaf challenging: for example when analyzing expression data from whole leaves, signals from genetic networks operating in distinct cellular response zones or cell types will be confounded, resulting in an inaccurate profile being generated.
To address this, several methods have been described which enable studies of cell specific gene expression. These include laser-capture microdissection (LCM)2 or GFP expressing plants used for protoplast generation and subsequent fluorescence activated cell sorting (FACS)3,4, the recently described INTACT system for nuclear precipitation5 and immunoprecipitation of polysomes6.
FACS has been successfully used for a number of studies, including showing that the cell identity and distance from the root tip had a significant effect on the expression profiles of a large number of genes3,7. FACS of GFP lines have also been used to demonstrate cell-specific transcriptional regulation during root nitrogen responses and lateral root development8, salt stress9 auxin distribution in the root10 and to create a gene expression map of the Arabidopsis shoot apical meristem11. Although FACS has previously been used to sort Arabidopsis leaf derived protoplasts based on autofluorescence12,13, so far the use of FACS on Arabidopsis lines expressing GFP in the leaves has been very limited4. In the following protocol we describe a method for obtaining Arabidopsis leaf protoplasts that are compatible with FACS while minimizing the impact of the protoplast generation regime. We demonstrate the method using the KC464 Arabidopsis line, which express GFP in the adaxial epidermis14, the KC274 line, which express GFP in the vascular tissue14 and the TP382 Arabidopsis line, which express a double GFP construct linked to a nuclear localization signal in the guard cells (data not shown; Figure 2). We are currently using this method to study both cell-type specific expression during development and stress, as well as heterogeneous cell populations at various stages of senescence.

Cell Specific Analysis of Arabidopsis Leaves Using Fluorescence Activated Cell Sorting

A method for producing Arabidopsis leaf protoplasts that are compatible with fluorescence activated cell sorting (FACS), allowing for studies of specific cell populations. This method is compatible with any Arabidopsis line that expresses GFP in a subset of cells.

After initiation of the leaf primordium, biomass accumulation is controlled mainly by cell proliferation and expansion in the leaves1. However, the ...

Automated Stomatal Response Measurement to Bacterial Invasion in Arabidopsis

Direct Observation and Automated Measurement of Stomatal Responses to Pseudomonas syringae pv. tomato DC3000 in Arabidopsis thaliana

Stomata are microscopic pores found in the plant leaf epidermis. Regulation of stomatal aperture is pivotal not only for balancing carbon dioxide uptake for photosynthesis and transpirational water loss but also for restricting bacterial invasion. While plants close stomata upon recognition of microbes, pathogenic bacteria, such as Pseudomonas syringae pv. tomato DC3000 (Pto), reopen the closed stomata to gain access into the leaf interior. In conventional assays for assessing stomatal responses to bacterial invasion, leaf epidermal peels, leaf discs, or detached leaves are floated on bacterial suspension, and then stomata are observed under a microscope followed by manual measurement of stomatal aperture. However, these assays are cumbersome and may not reflect stomatal responses to natural bacterial invasion in a leaf attached to the plant. Recently, a portable imaging device was developed that can observe stomata by pinching a leaf without detaching it from the plant, together with a deep learning-based image analysis pipeline designed to automatically measure stomatal aperture from leaf images captured by the device. Here, building on these technical advances, a new method to assess stomatal responses to bacterial invasion in Arabidopsis thaliana is introduced. This method consists of three simple steps: spray inoculation of Pto mimicking natural infection processes, direct observation of stomata on a leaf of the Pto-inoculated plant using the portable imaging device, and automated measurement of stomatal aperture by the image analysis pipeline. This method was successfully used to demonstrate stomatal closure and reopening during Pto invasion under conditions that closely mimic the natural plant-bacteria interaction.

Author Spotlight: Advancing Stomatal Research with Automated Aperture Measurement

Here, we present a simple method for direct observation and automated measurement of stomatal responses to bacterial invasion in Arabidopsis thaliana. This method leverages a portable stomatal imaging device, together with an image analysis pipeline designed for leaf images captured by the device.

Stomata are microscopic pores found in the plant leaf epidermis. Regulation of stomatal aperture is pivotal not only for balancing carbon dioxide uptake ...

Immunology and Infection

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 ...

Real-time In Vivo Recording of Arabidopsis Calcium Signals During Insect Feeding Using a Fluorescent Biosensor

Calcium ions are predicted to be key signaling entities during biotic interactions, with calcium signaling forming an established part of the plant defense response to microbial elicitors and to wounding caused by chewing insects, eliciting systemic calcium signals in plants. However, the role of calcium in vivo during biotic stress is still unclear. This protocol describes the use of a genetically-encoded calcium sensor to detect calcium signals in plants during feeding by a hemipteran pest. Hemipterans such as aphids pierce a small number of cells with specialized, elongated sucking mouthparts, making them the ideal tool to study calcium dynamics when a plant is faced with a biotic stress, which is distinct from a wounding response. In addition, fluorescent biosensors are revolutionizing the measurement of signaling molecules in vivo in both animals and plants. Expressing a GFP-based calcium biosensor, GCaMP3, in the model plant Arabidopsis thaliana allows for the real-time imaging of plant calcium dynamics during insect feeding, with a high spatial and temporal resolution. A repeatable and robust assay has been developed using the fluorescence microscopy of detached GCaMP3 leaves, allowing for the continuous measurement of cytosolic calcium dynamics before, during, and after insect feeding. This reveals a highly-localized rapid calcium elevation around the aphid feeding site that occurs within a few minutes. The protocol can be adapted to other biotic stresses, such as additional insect species, while the use of Arabidopsis thaliana allows for the rapid generation of mutants to facilitate the molecular analysis of the phenomenon.

Real-time In Vivo Recording of Arabidopsis Calcium Signals During Insect Feeding Using a Fluorescent Biosensor

This protocol outlines a simple method for analyzing calcium signals in plants generated by feeding hemipteran insects, such as aphids. Arabidopsis thaliana transformed with the GFP calcium biosensor GCaMP3 allow for the real-time in vivo imaging of calcium dynamics with a high temporal and spatial resolution.

Calcium ions are predicted to be key signaling entities during biotic interactions, with calcium signaling forming an established part of the plant ...

Analysis of Arabidopsis thaliana Growth Behavior in Different Light Qualities

Plant biologists often need to observe the growth behavior of their chosen species. To this end, the plants need constant environmental and stable light conditions, which are preferably variable in quantity and quality so that studies under different setups can be conducted. These requirements are met by climatic chambers featuring light emitting diodes (LED) lights, which can &#8211; in contrast to fluorescent lights &#8211; be set to different wavelengths. LEDs are energy conserving and emit virtually no heat even at light intensities, which often constitutes a problem with other light sources. The presented protocol provides a step-by-step guidance of how to program a climatic chamber equipped with variable LED lights as well as describing several approaches for in depth analysis of growth phenotypes. Depending on the experimental set-up various characteristics of the growing plants can be observed and analyzed. Here we describe how to determine fresh weight, leaf area, photosynthetic activity, and stomatal density. We demonstrate that in order to obtain reliable data and draw valid conclusions it is mandatory to use a sufficient number of individuals for statistical evaluation. Taking too few plants for this kind of analysis results in high statistical errors and consequently in less clear interpretations of the data.

Analysis of Arabidopsis thaliana Growth Behavior in Different Light Qualities

Here, we present a protocol for studying plant growth behavior and especially phenotypes in a reproducible manner. We show how to provide variable and at the same time stable light conditions. Proper analyses depend on sufficient sample numbers and valid statistical evaluations.

Plant biologists often need to observe the growth behavior of their chosen species. To this end, the plants need constant environmental and stable light ...

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 ...

High-Resolution Imaging of Chlorophyll Biosynthesis in Arabidopsis Seedling

Non-invasive Assay for Chlorophyll Biosynthesis Kinetics Determination during Early Stages of Arabidopsis De-etiolation

Chlorophyll biosynthesis is a hallmark of de-etiolation, one of the most dramatic stages in the plant life cycle. The tightly controlled and highly dynamic process of chlorophyll biosynthesis is triggered during the shift from the dark to the light in flowering plants. At the moment when etiolated seedlings are exposed to the first traces of sunlight, rapid (in order of seconds) conversion of protochlorophyllide into chlorophyllide is mediated by unique light-accepting protein complexes, leading via subsequent metabolic steps to the production of fully functional chlorophyll. Standard techniques for chlorophyll content analysis include pigment extraction from detached plant tissues, which does not apply to studying such fast processes. To investigate chlorophyll kinetics in vivo with high accuracy and spatiotemporal resolution in the first hours after light-induced de-etiolation, an instrument and protocol were developed. Here, we present a detailed procedure designed for statistically robust quantification of chlorophyll in the early stages of Arabidopsis de-etiolation.

Author Spotlight: Non-Invasive High-Resolution Measurement of Chlorophyll Synthesis During De-Etiolation

Here, we describe an advanced tool designed for chlorophyll biosynthesis monitoring during the early stages of Arabidopsis seedling de-etiolation. The novel methodology provides non-invasive real-time chlorophyll fluorescence imaging at high spatial and temporal resolution.

Chlorophyll biosynthesis is a hallmark of de-etiolation, one of the most dramatic stages in the plant life cycle. The tightly controlled and highly ...

Using Changes in Leaf Transmission to Investigate Chloroplast Movement in Arabidopsis thaliana

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Investigating Tissue- and Organ-specific Phytochrome Responses using FACS-assisted Cell-type Specific Expression Profiling in Arabidopsis thaliana

Evaluation of Photosynthetic Behaviors by Simultaneous Measurements of Leaf Reflectance and Chlorophyll Fluorescence Analyses

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

Cell Specific Analysis of Arabidopsis Leaves Using Fluorescence Activated Cell Sorting

Direct Observation and Automated Measurement of Stomatal Responses to Pseudomonas syringae pv. tomato DC3000 in Arabidopsis thaliana

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

Real-time In Vivo Recording of Arabidopsis Calcium Signals During Insect Feeding Using a Fluorescent Biosensor

Analysis of Arabidopsis thaliana Growth Behavior in Different Light Qualities

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

Non-invasive Assay for Chlorophyll Biosynthesis Kinetics Determination during Early Stages of Arabidopsis De-etiolation