This work was supported by a PhD fellowship from the University of Antwerp to V.A.; a PhD fellowship from the Flemish Science Foundation (FWO, 11ZI916N) to K.S.; project grants from the FWO (G0D0514N); a concerted research activity (GOA) research grant, &#34;A Systems Biology Approach of Leaf Morphogenesis&#34; from the research council of the University of Antwerp; and the Interuniversity Attraction Poles (IUAP VII/29, MARS), &#34;Maize and Arabidopsis Root and Shoot Growth&#34; from the Belgian Federal Science Policy Office (BELSPO) to G.T.S.B. Han Asard, Bulelani L. Sizani and Hamada AbdElgawad all contributed to the video.

NOTE: The following protocol for kinematic analysis is only valid for leaves during steady-state growth. This implies a stable leaf elongation rate and spatial patterns of cell length and expansion in a leaf during a period of several days6.
1. Plant Growth and Measurements of Leaf Elongation Rate (LER)
<ol>
	<li>Choose a leaf in steady-state growth and a developmental stage of interest. 
	NOTE: There is a difference between steady-state growth and repetitive growth, which implies similar spatial patterns on successive leaves on the same axis. During the early stages of seedling growth, successive leaves typically grow increasingly faster due to the increasing size of the growth zone7. Although a few higher leaf positions can have a similar growth pattern8, this is a transient phase that may be affected by treatments under investigation. It is therefore important to compare lines and treatments strictly on the same leaf position, even though it may be developing at a different time. Even at a constant elongation rate, the growth rate profile is not necessarily the same at different developmental stages. Therefore, it is important to analyze leaves at the same developmental stage8, typically defined by the number of days after emergence.</li>
	<li>To perform a full kinematic analysis of leaf growth in monocots, grow at least 15 plants for each treatment and genotype under controlled conditions in a growth room.</li>
	<li>At the time the leaf of interest appears (emergence from the whorl of surrounding leaves), start measuring the length of the leaf daily with a ruler until the leaf is fully expanded (Figure 1i). Leaf length implies the length from soil level to the tip of the leaf. Be careful not to break or damage the leaf, since this might alter its growth.</li>
</ol>
<img alt="Figure 1" src="/files/ftp_upload/54887/54887fig1.jpg" /> 
Figure 1: Schematic overview of a kinematic analysis of maize leaves. The leaf of interest is measured with a ruler for three consecutive days to calculate the Leaf Elongation Rate (LER). Thereafter, the leaf is harvested and a three-centimeter segment is used for the determination of the meristem size. This is done by measuring the length from the base up to the most distal mitotic figure after DAPI staining. (A) Examples of proliferative mitotic figures and (B) formative mitotic figures. The first eleven centimeters from the leaf base on the other side of the mid vein are used to cut ten one-centimeter segments for cell length measurements. These measurements provide the basis for creating the cell length profile, which serves to determine the mature cell length (lmat) and the length of cells leaving the meristem (ldiv). The LER and lmat are used to calculate the cell production rate (P), while ldiv and Lmer are used to calculate the number of cells in the meristem (Nmer). In turn, P and Nmer are used to calculate the average cell division rate (D), which is the inverse of cell cycle duration (Tc). Arrows of the same color indicate parameters that are used to calculate the parameter following on these arrows. Scale bars = 40 &#181;m. Roman numbers are used to refer to specific experimental procedures described in the protocol. <a href="http://cloudfront.jove.com/files/ftp_upload/54887/54887fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
2. Harvesting
<ol>
	<li>At the developmental stage of interest (e.g., the third day after emergence), choose at least five representative plants from the batch on which to conduct the kinematic analysis. Continue measuring the rest of the plants as explained in step 1.3 to determine the final leaf length.</li>
	<li>Cut the above ground part of the plant. To keep the meristematic part intact, cut as close as possible to the roots (Figure 1ii).</li>
	<li>Starting from the outer leaves, remove all leaves up to the leaf of interest by gently unrolling them one by one. If necessary, remove a few extra millimeters from the base to detach the leaves. Also remove the apex and small leaves enclosed by the leaf of interest (Figure 1iii).</li>
	<li>Cut a 3 cm segment, starting from the base on one side of the mid vein, and store it in a 1.5 ml test tube filled with 3:1 (v:v) absolute ethanol:acetic acid solution (CAUTION: wear gloves) at 4 &#176;C for 24 hr up to several months (Figure 1iv). This segment will later be used to determine the length of the meristem.</li>
	<li>From the other side of the vein, cut a 11 cm segment from the base (Figure 1 v) and place it in a 15 ml tube filled with absolute ethanol at 4 &#176;C for at least 6 hr to remove pigments (Figure 1 vi). 
	NOTE: Later on, use only the first 10 cm to determine the cell length profile (see Discussion).</li>
	<li>Renew the absolute ethanol for another round of cleaning at 4 &#176;C for at least 24 hr (Figure 1vi).</li>
	<li>Finally, substitute the absolute ethanol with pure lactic acid (CAUTION: wear gloves) for cleaning and storage at 4 &#176;C for 24 h or until further use (Figure 1vi).</li>
</ol>
3. Meristem Length Measurements
<ol>
	<li>Prepare a rinsing buffer containing 50 mM sodium chloride (NaCl), 5 mM ethylenediaminetetraacetic acid (EDTA; CAUTION: wear gloves) and 10 mM Tris(hydroxymethyl)aminomethane-hydrochloric acid (TRIS-HCl; pH 7).</li>
	<li>Take the 3 cm segment from section 2.4 and soak it in the buffer for 20 min (Figure 1vii).</li>
	<li>While waiting, use the rinsing buffer to prepare a 4&#39;,6-diamidino-2-phenylindole (DAPI) staining solution of 1 &#181;g/ml, keeping it on ice and in the dark.</li>
	<li>Stain the nuclei by placing the meristem segment for 2-5 min in the DAPI staining solution. Work on ice and in the dark (Figure 1vii).</li>
	<li>Check for fluorescence signal by quickly mounting the segment on a microscopy glass and covering it with a cover glass. The epidermal cells should show fluorescence, while the underlying cell layers should not.</li>
	<li>If the staining is not sufficient, put the segment back in the DAPI staining solution for some extra minutes.</li>
	<li>To stop the staining, mount the segments in a drop of rinsing buffer on a microscopy slide and cover with a cover glass.</li>
	<li>Use a microscope equipped with UV-fluorescence at a 20X magnification, allowing for visualization of around 1,000 epidermal cells at once. Scroll throughout the segment and look for proliferative mitotic figures (metaphase, anaphase, telophase, and cytokinesis), but avoid formative cell division of the developing stomata (Figure 1viii)9. Define where the most distal mitotic figure is located.</li>
	<li>Determine the length of the meristem by measuring the distance between the base of the leaf and the most distal epidermal mitotic figure. Use an image-analysis software (e.g., ImageJ) to measure the full length of the image frame.
	<ol>
		<li>Count the number of frames that cover the full meristem length (from the leaf base to the most distal mitotic figure) and multiply this number by the length of one frame to obtain the full meristem length (Figure 1ix).</li>
	</ol>
	</li>
</ol>
4. Cell Length Profile
<ol>
	<li>Take the segment that is stored in lactic acid (step 2.5) and place it carefully on the bench. Cut the segments with a scalpel in 10 segments of 1 cm each (Figure 1x).</li>
	<li>Mount the successive leaf segments on a microscopy slide in a small drop of lactic acid. Make sure to consistently face either the adaxial or abaxial side up. In principle, there is no preference for a particular side.</li>
	<li>Use a microscope equipped with differential interference contrast (DIC) optics to analyze the segments, starting from the leaf base. Measure with an image analysis software the length of at least 20 replicate epidermal cells in files directly adjacent to the stomatal files in order to consistently select the same cell type.
	<ol>
		<li>Do this at equally spaced positions along each of the segments (4 positions per segment suffice), and make sure to write down the corresponding position for each measurement throughout the leaf (Figure 1xi).</li>
	</ol>
	</li>
	<li>Determine the average cell length at each mm along the leaf axis by using a local polynomial smoothing procedure, implemented in an R-script (Figure 1xii; Supplementary File 1). 
	NOTE: The R-script provides a series of data with increasing smoothing. The amount of smoothing required is somewhat arbitrary and ideally should just remove the local noise, but not affect the overall curve. Make sure to use the same amount of smoothing for all samples within one experiment.</li>
	<li>Average the cell length at each position between plants and calculate the standard error to create a cell length profile along the leaf axis.</li>
</ol>
5. Calculations of Kinematic Parameters (See Supplementary File 2)
<ol>
	<li>Calculate the LER by taking the change in leaf length between two successive time points (e.g., 24 hr, as in step 1.3) and dividing it by the time interval.</li>
	<li>Calculate the length of the growth zone (Lgz) corresponding to the position distal from the base where cells reach 95% of their mature cell length on the smoothened cell length profile.
	<ol>
		<li>Take for each position on the smoothened cell length profile 95% of the average of all cell lengths following that position (Figure 2).</li>
		<li>Compare the smoothened cell lengths (step 4.4) with the calculated 95% cell length at each position. Starting from the base of the leaf, the growth zone ends at the position where the actual cell length equals 95% of the following cell lengths (Figure 2; see Supplementary Data 2).</li>
	</ol>
	</li>
</ol>
<img alt="Figure 2" src="/files/ftp_upload/54887/54887fig2.jpg" /> 
Figure 2: Determining the end of the growth zone. Meristem: At the position indicated with a red star, the actual cell size is smaller than 95% (red dotted line) of the average cell size of all cells following this position (red solid line). The end of the growth zone (Lgz; indicated with a blue star) is located where 95% (dotted blue line) of the average cell size of all cells following this position (solid blue line) equals the actual cell size. Division zone (D), elongation zone (E), and mature zone (M). Dashed arrows indicate the convergence between the local size and 95% of the average size over the distal portion of the leaf when moving from the basal positions to the tip of the leaf. <a href="http://cloudfront.jove.com/files/ftp_upload/54887/54887fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<ol start="3">
	<li>Calculate the length of elongation zone (Lel) as the difference between the length of the growth zone (Lgz) and the meristem size (Lmer; determined in step 3).</li>
	<li>Calculate the mature cell length (lmat) as the average cell length in the mature zone.</li>
	<li>Divide the LER by lmat to obtain the cell production rate (P).</li>
	<li>Calculate the number of cells in the elongation zone (Nel) as the difference between Ngz and Nmer. The number of cells in the meristem (Nmer) equals the cumulative number of cells located in the intervals corresponding to the meristem. The number of cells in the growth zone (Ngz) equals the cumulative number of cells located in the intervals corresponding to the growth zone.</li>
	<li>Calculate the average cell division rate (D) as P/Nmer. The cell cycle duration (Tc) equals ln(2)/D.</li>
	<li>Calculate the time in the elongation zone (Tel) by dividing Nel by P. The time in the division zone equals log2(Nmer)*Tc. The length of cells leaving the meristem (ldiv) equals the cell length from the smoothened cell length profile at the end of the meristem.</li>
	<li>Calculate the average cell expansion rate (Rel) using following formula: ln(lmat)-ln(ldiv)]/Tel.</li>
</ol>

The authors declare that they have no competing financial interests.

A full kinematic analysis on maize leaves enables the determination of the cellular basis of leaf growth and allows for the design of efficient sampling strategies. Although the protocol is relatively straightforward, some caution is recommended in the following critical steps: (1) It is important to detach the younger, enclosed leaves (step 2.3) without damaging the meristem, since meristem length determination (step 3) requires the complete meristem to be present. Some practice beforehand might be needed. (2) Meristema...

Growth analysis depends on a set of tools that are commonly used by plant scientists to describe genotype determined growth differences and/or phenotypic responses to environmental factors. They include size and weight measurements of the whole plant or an organ and calculations of growth rates to explore the underlying mechanisms of growth. Organ growth is determined by cell division and expansion at the cellular level. Therefore, including the quantification of these two processes in growth analyses is key to understanding differences in whole-organ growth1. Consequently, it is crucial to have an appropriate methodology to determine cellular growth parameters that is relatively easy to use by non-specialized laboratories.
Kinematic analysis has already been established as an approach providing a powerful framework for the development of organ growth models2. The technique has been optimized for linear systems, such as Arabidopsis thaliana roots and monocotyledonous leaves, but also for non-linear systems, such as dicotyledonous leaves3. Nowadays, this methodology is increasingly being used to study how genetic, hormonal, developmental, and environmental factors influence cell division and expansion in various organs (Table 1). Moreover, it also provides a framework to link cellular processes to their underlying biochemical, molecular, and physiological regulations (Table 2), although limitations can be imposed by organ size and spatial organization for techniques that require higher amounts of plant material (e.g., metabolite measurements, proteomics, etc.).
Monocotyledonous leaves, such as the maize (Zea mays) leaf, represent linear systems in which cells move from the base of the leaf towards the tip, sequentially passing through the meristem and elongation zone to reach the mature zone. This makes it an ideal model system for quantitative studies of the spatial patterns of growth4. Moreover, maize leaves have large growth zones (meristem and elongation zone spanning several centimeters5) and provide possibilities for studies at other organizational levels. This allows for the investigation of the (putative) regulatory mechanisms controlling cell division and expansion, quantified by kinematic analysis through a range of molecular techniques, physiological measurements, and cell biology approaches (Table 2).
Here, we provide a protocol for performing a kinematic analysis in monocot leaves. First, we explain how to conduct a proper analysis of both cell division and cell elongation as a function of position along the leaf axis and how to calculate kinematic parameters. Secondly, we also show how this can be used as a basis for sampling design. Here, we discuss two cases: high-resolution sampling and focused sampling, enabling improved data interpretation and the saving of time/money, respectively.
Table 1. Overview of kinematic analyses methods for quantification of cell division and expansion in various organs.
<table border="1" fo:keep-together.within-page="1">
	<tbody>
		<tr>
			<td>organ</td>
			<td>reference</td>
		</tr>
		<tr>
			<td>monocotyledonous leaves</td>
			<td>16, 20, 21, 22</td>
		</tr>
		<tr>
			<td>root tips</td>
			<td>2, 23, 24, 25, 26, 27, 28, 29</td>
		</tr>
		<tr>
			<td>dicotyledonous leaves</td>
			<td>21, 30, 31</td>
		</tr>
		<tr>
			<td>shoot apical meristem</td>
			<td>32</td>
		</tr>
	</tbody>
</table>
Table 1. Overview of kinematic analyses methods for quantification of cell division and expansion in various organs.
<img alt="Figure 3" src="/files/ftp_upload/54887/54887table2.jpg" />
Table 2. Link between cellular processes quantified by the kinematic analysis to their regulation at the molecular level. References to various studies linking the quantification of cellular processes to results from biochemical and molecular assays in various species and organs. Xyloglucan endotransglucosylase (XET), malondialdehyde (MDA), cyclin-dependent kinases (CDK). <a href="http://cloudflare.jove.com/files/ftp_upload/54887/54887table2large.jpg" target="_blank">Please click here to view a larger version of this table.</a>

<table><tbody><tr><td>Pots</td><td>Any</td><td>Any</td><td>We use pots with the following measures, but can be different depending on the treatment/study: bottom diameter: 11 cm, opening diameter: 15 cm, height: 12 cm. We grow one maize plant per pot.</td></tr><tr><td>Planting substrate</td><td>Any</td><td>Any</td><td>We use potting medium (Jiffy, The Netherlands), but other substrates can be used, depending on treatment/study.</td></tr><tr><td>Ruler</td><td>Any</td><td>Any</td><td>An extension ruler that covers at least 1.5 meters is needed to measure the final leaf length of the plants.</td></tr><tr><td>Seeds</td><td>Any</td><td>NA</td><td>Seeds can be ordered from a breeder.</td></tr><tr><td>Scalpel</td><td>Any</td><td>Any</td><td>The scalpel is used during leaf harvesting to detach the leaf of interest from its surrounding leaves and right after harvesting to cut a proper sample for cell length and meristem length measurements.&amp;nbsp;</td></tr><tr><td>15 mL falcon tubes</td><td>Any</td><td>Any</td><td>The 15 mL falcon tubes are used for storing samples used for cell length measurements during sample clearing with absolute ethanol and lactic acid.</td></tr><tr><td>Eppendorf tubes</td><td>Any</td><td>Any</td><td>The eppendorf tubes are used for storing samples used for meristem length measurements in ethanol:acetic acid 3:1 (v:v) solution.</td></tr><tr><td>Gloves</td><td>Any</td><td>Any</td><td>Latex gloves, which protect against corrosive reagents.</td></tr><tr><td>Acetic acid</td><td>Any</td><td>Any</td><td>CAUTION: Corrosive to metals, category 1 Skin corrosion, categories 1A,1B,1C Serious eye damage, category 1; Flammable liquids, categories 1,2,3</td></tr><tr><td>Absolute ethanol</td><td>Any</td><td>Any</td><td>CAUTION: Hazardous in case of skin contact (irritant), of eye contact (irritant), of inhalation. Slightly hazardous in case of skin contact (permeator), of ingestion</td></tr><tr><td>Lactic acid&amp;nbsp;&gt;98%</td><td>Any</td><td>Any</td><td>CAUTION: Corrosive to metals, category 1 Skin corrosion, categories 1A,1B,1C Serious eye damage, category 1</td></tr><tr><td>Sodium chloride (NaCl)</td><td>Any</td><td>Any</td><td></td></tr><tr><td>Ethylenediaminetetraacetic acid (EDTA)</td><td>Any</td><td>Any</td><td>CAUTION: Acute toxicity (oral, dermal, inhalation), category 4 Skin irritation, category 2 Eye irritation, category 2 Skin sensitisation, category 1 Specific Target Organ Toxicity &amp;ndash; Single exposure, category 3</td></tr><tr><td>Tris(hydroxymethyl)aminomethane hydrochloride (Tris-HCl)</td><td>Any</td><td>Any</td><td>This material can be an irritant, contact with eyes and skin should be avoided. Inhalation of dust may be irritating to the respiratory tract.</td></tr><tr><td>4&amp;prime;,6-Diamidine-2&amp;prime;-phenylindole dihydrochloride (DAPI)</td><td>Any</td><td>Any</td><td>Cell permeable fluorescent minor groove-binding probe for DNA. Causes skin irritation. May cause an allergic skin reaction. May cause respiratory irritation.</td></tr><tr><td>Ice</td><td>Any</td><td>NA</td><td>The DAPI solution has to be kept on ice.</td></tr><tr><td>Fluorescent microscope</td><td>AxioScope A1, Axiocam ICm1 from Zeiss or other</td><td></td><td>Any fluorescent microscope can be used for determining meristem length.</td></tr><tr><td>Microscopic slide</td><td>Any</td><td>Any</td><td></td></tr><tr><td>Cover glass</td><td>Any</td><td>Any</td><td></td></tr><tr><td>Tweezers</td><td>Any</td><td>Any</td><td>Tweezers are needed for unfolding the rolled maize leaf right after harvesting in order to cut a proper sample for cell length and meristem length measurements.&amp;nbsp;</td></tr><tr><td>Image-analysis software</td><td>Axiovision (Release 4.8) from Zeiss</td><td>NA</td><td>The software can be downloaded at: http://www.zeiss.com/microscopy/en_de/downloads/axiovision.html. Other softwares such as ImageJ (https://imagej.nih.gov/ij/) could be used as well.</td></tr><tr><td>Microscope equipped with DIC</td><td>AxioScope A1, Axiocam ICm1 from Zeiss or other</td><td></td><td>Any microscope, equipped with differential interference contrast (DIC) can be used to measure cell lengths.</td></tr><tr><td>R statistical analysis software</td><td>R Foundation for Statistical Computing</td><td>NA</td><td>Open source; Could be downloaded at https://www.r-project.org/</td></tr><tr><td>R script</td><td>NA</td><td>NA</td><td>We use the kernel smoothing function locpoly of the Kern Smooth package (Wand MP, Jones MC.&amp;nbsp; Kernel Smoothing: Chapman &amp;amp; Hall/CRC (1995)). The script is available for Mac and Windows upon inquiry with the corresponding author.</td></tr></tbody></table>

kinematic analysis of cell division and expansion: quantifying the cellular basis of growth and sampling developmental zones in zea mays leaves

Growth analyses are often used in plant science to investigate contrasting genotypes and the effect of environmental conditions. The cellular aspect of these analyses is of crucial importance, because growth is driven by cell division and cell elongation. Kinematic analysis represents a methodology to quantify these two processes. Moreover, this technique is easy to use in non-specialized laboratories. Here, we present a protocol for performing a kinematic analysis in monocotyledonous maize (Zea mays) leaves. Two aspects are presented: (1) the quantification of cell division and expansion parameters, and (2) the determination of the location of the developmental zones. This could serve as a basis for sampling design and/or could be useful for data interpretation of biochemical and molecular measurements with high spatial resolution in the leaf growth zone. The growth zone of maize leaves is harvested during steady-state growth. Individual leaves are used for meristem length determination using a DAPI stain and cell-length profiles using DIC microscopy. The protocol is suited for emerged monocotyledonous leaves harvested during steady-state growth, with growth zones spanning at least several centimeters. To improve the understanding of plant growth regulation, data on growth and molecular studies must be combined. Therefore, an important advantage of kinematic analysis is the possibility to correlate changes at the molecular level to well-defined stages of cellular development. Furthermore, it allows for a more focused sampling of specified developmental stages, which is useful in case of limited budget or time.

Here, we show a comparison between well-watered plants (control, 54% soil water content, (SWC)) and plants subjected to drought stress conditions (drought, 34% SWC) in terms of their leaf growth. All plants were grown in a growth chamber under controlled conditions (16 hr day/8 hr night, 25 &#176;C/18 &#176;C day/night, 300-400 &#181;Em-2sec-1 photosynthetically active radiation (PAR). The drought conditions were established by withholding water until the correct SWC...

Watch this Scientific Journal Video about Kinematic Analysis of Cell Division and Expansion: Quantifying the Cellular Basis of Growth and Sampling Developmental Zones in Zea mays Leaves at JoVE.com

Kinematic Analysis of Cell Division and Expansion: Quantifying the Cellular Basis of Growth and Sampling Developmental Zones in Zea mays Leaves

Quantifying cell division and expansion is of crucial importance to the understanding of whole-plant growth. Here, we present a protocol to calculate cellular parameters determining maize leaf growth rates and highlight the use of these data for investigating molecular growth regulatory mechanisms by directing developmental stage-specific sampling strategies.

Kinematic Analysis of Cell Division and Expansion: Quantifying the Cellular Basis of Growth and Sampling Developmental Zones in Zea mays Leaves

Growth analyses are often used in plant science to investigate contrasting genotypes and the effect of environmental conditions. The cellular aspect of ...

kinematic-analysis-cell-division-expansion-quantifying-cellular-basis

Research

JoVE Journal

Biology

10.8K Views.  University of Antwerp. Quantifying cell division and expansion is of crucial importance to the understanding of whole-plant growth. Here, we present a protocol to calculate cellular parameters determining maize leaf growth rates and highlight the use of these data for investigating molecular growth regulatory mechanisms by directing developmental stage-specific sampling strategies. 

Video: Kinematic Analysis of Cell Division and Expansion: Quantifying the Cellular Basis of Growth and Sampling Developmental Zones in Zea mays Leaves

Experimental Design for Laser Microdissection RNA-Seq: Lessons from an Analysis of Maize Leaf Development

Genes with important roles in development frequently have spatially and/or temporally restricted expression patterns. Often these gene transcripts are not detected or are not identified as differentially expressed (DE) in transcriptomic analyses of whole plant organs. Laser Microdissection RNA-Seq (LM RNA-Seq) is a powerful tool to identify genes that are DE in specific developmental domains. However, the choice of cellular domains to microdissect and compare, and the accuracy of the microdissections are crucial to the success of the experiments. Here, two examples illustrate design considerations for transcriptomics experiments; a LM RNA-seq analysis to identify genes that are DE along the maize leaf proximal-distal axis, and a second experiment to identify genes that are DE in liguleless1-R (lg1-R) mutants compared to wild-type. Key elements that contributed to the success of these experiments were detailed histological and in situ hybridization analyses of the region to be analyzed, selection of leaf primordia at equivalent developmental stages, the use of morphological landmarks to select regions for microdissection, and microdissection of precisely measured domains. This paper provides a detailed protocol for the analysis of developmental domains by LM RNA-Seq. The data presented here illustrate how the region selected for microdissection will affect the results obtained.

Many developmentally important genes have cell- or tissue-specific expression patterns. This paper describes LM RNA-seq experiments to identify genes that are differentially expressed at the maize leaf blade-sheath boundary and in lg1-R mutants compared to wild-type. The experimental considerations discussed here apply to transcriptomic analyses of other developmental phenomena.

Genes with important roles in development frequently have spatially and/or temporally restricted expression patterns. Often these gene transcripts are not ...

Biochemistry

Observing Mitotic Division and Dynamics in a Live Zebrafish Embryo

Mitosis is critical for organismal growth and differentiation. The process is highly dynamic and requires ordered events to accomplish proper chromatin condensation, microtubule-kinetochore attachment, chromosome segregation, and cytokinesis in a small time frame. Errors in the delicate process can result in human disease, including birth defects and cancer. Traditional approaches investigating human mitotic disease states often rely on cell culture systems, which lack the natural physiology and developmental/tissue-specific context advantageous when studying human disease. This protocol overcomes many obstacles by providing a way to visualize, with high resolution, chromosome dynamics in a vertebrate system, the zebrafish. This protocol will detail an approach that can be used to obtain dynamic images of dividing cells, which include: in vitro transcription, zebrafish breeding/collecting, embryo embedding, and time-lapse imaging. Optimization and modifications of this protocol are also explored. Using H2A.F/Z-EGFP (labels chromatin) and mCherry-CAAX (labels cell membrane) mRNA-injected embryos, mitosis in AB wild-type, auroraBhi1045, and esco2hi2865 mutant zebrafish is visualized. High resolution live imaging in zebrafish allows one to observe multiple mitoses to statistically quantify mitotic defects and timing of mitotic progression. In addition, observation of qualitative aspects that define improper mitotic processes (i.e., congression defects, missegregation of chromosomes, etc.) and improper chromosomal outcomes (i.e., aneuploidy, polyploidy, micronuclei, etc.) are observed. This assay can be applied to the observation of tissue differentiation/development and is amenable to the use of mutant zebrafish and pharmacological agents. Visualization of how defects in mitosis lead to cancer and developmental disorders will greatly enhance understanding of the pathogenesis of disease.

Mitosis is critical to every living organism and defects often lead to cancer and developmental disorders. Using this imaging protocol and zebrafish as a model system, researchers can visualize mitosis in a live vertebrate organism and the multitude of defects that arise when mitotic processes are defective.

Mitosis is critical for organismal growth and differentiation. The process is highly dynamic and requires ordered events to accomplish proper chromatin ...

Developmental Biology

Confocal Live Imaging of Shoot Apical Meristems from Different Plant Species

The shoot apical meristem (SAM) functions as a conserved stem cell reservoir and it generates almost all aboveground tissues during the postembryonic development. The activity and morphology of SAMs determine important agronomic traits, such as shoot architecture, size and number of reproductive organs, and most importantly, grain yield. Here, we provide a detailed protocol for analyzing both the surface morphology and the internal cellular structure of the living SAMs from different species through laser scanning confocal microscope. The whole procedure from the sample preparation to the acquisition of high resolution three-dimensional (3D) images can be accomplished within as short as 20 minutes. We demonstrate that this protocol is highly efficient for studying not only the inflorescence SAMs of the model species but also the vegetative meristems from different crops, providing a simple but powerful tool to study the organization and development of meristems across different plant species.

This protocol presents how to live image and analyze the shoot apical meristems from different plant species using laser scanning confocal microscopy.

The shoot apical meristem (SAM) functions as a conserved stem cell reservoir and it generates almost all aboveground tissues during the postembryonic ...

Use of Atomic Force Microscopy to Measure Mechanical Properties and Turgor Pressure of Plant Cells and Plant Tissues

We present here the use of atomic force microscopy to indent plant tissues and recover its mechanical properties. Using two different microscopes in indentation mode, we show how to measure an elastic modulus and use it to evaluate cell wall mechanical properties. In addition, we also explain how to evaluate turgor pressure. The main advantages of atomic force microscopy are that it is non-invasive, relatively rapid (5~20 min), and that virtually any type of living plant tissue that is superficially flat can be analyzed without the need for treatment. The resolution can be very good, depending on the tip size and on the number of measurements per unit area. One limitation of this method is that it only gives direct access to the superficial cell layer.

Here, we present atomic force microscopy (AFM), operated as a nano- and micro-indentation tool on cells and tissues. The instrument allows the simultaneous acquisition of 3D surface topography of the sample and its mechanical properties, including cell wall Young's modulus as well as turgor pressure.

We present here the use of atomic force microscopy to indent plant tissues and recover its mechanical properties. Using two different microscopes in ...

Live Cell Imaging of Microtubule Cytoskeleton and Micromechanical Manipulation of the Arabidopsis Shoot Apical Meristem

Understanding cell and tissue level regulation of growth and morphogenesis has been at the forefront of biological research for many decades. Advances in molecular and imaging technologies allowed us to gain insights into how biochemical signals influence morphogenetic events. However, it is increasingly evident that apart from biochemical signals, mechanical cues also impact several aspects of cell and tissue growth. The Arabidopsis shoot apical meristem (SAM) is a dome-shaped structure responsible for the generation of all aboveground organs. The organization of the cortical microtubule cytoskeleton that mediates apoplastic cellulose deposition in plant cells is spatially distinct. Visualization and quantitative assessment of patterns of cortical microtubules are necessary for understanding the biophysical nature of cells at the SAM, as cellulose is the stiffest component of the plant cell wall. The stereotypical form of cortical microtubule organization is also a consequence of tissue-wide physical forces existing at the SAM. Perturbation of these physical forces and subsequent monitoring of cortical microtubule organization allows for the identification of candidate proteins involved in mediating mechano-perception and transduction. Here we describe a protocol that helps investigate such processes.

Live Cell Imaging of Microtubule Cytoskeleton and Micromechanical Manipulation of the Arabidopsis Shoot Apical Meristem

Here we describe a protocol for live cell imaging of the cortical microtubule cytoskeleton at the shoot apical meristem and monitoring its response to changes in physical forces.

Understanding cell and tissue level regulation of growth and morphogenesis has been at the forefront of biological research for many decades. Advances in ...

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

In vivo, tip-growing plant cells need to overcome a series of physical barriers; however, researchers lack the methodology to visualize cellular behavior in such restrictive conditions. To address this issue, we have developed growth chambers for tip-growing plant cells that contain a series of narrow, micro-fabricated gaps (~1 &#181;m) in a poly-dimethylsiloxane (PDMS) substrate. This transparent material allows the user to monitor tip elongation processes in individual cells during microgap penetration by time-lapse imaging. Using this experimental platform, we observed morphological changes in pollen tubes as they penetrated the microgap. We captured the dynamic changes in the shape of a fluorescently labeled vegetative nucleus and sperm cells in a pollen tube during this process. Furthermore, we demonstrated the capability of root hairs and moss protonemata to penetrate the 1 &#181;m gap. This in vitro platform can be used to study how individual cells respond to physically constrained spaces and may provide insights into tip-growth mechanisms.

We describe a method to investigate the capability of tip-growing plant cells, including pollen tubes, root hairs, and moss protonemata, to elongate through extremely narrow gaps (~1 &#181;m) in a microfluidic device.

In vivo, tip-growing plant cells need to overcome a series of physical barriers; however, researchers lack the methodology to visualize cellular behavior ...

Bioengineering

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 ...

AFM-based Mapping of the Elastic Properties of Cell Walls: at Tissue, Cellular, and Subcellular Resolutions

We describe a recently developed method to measure mechanical properties of the surfaces of plant tissues using atomic force microscopy (AFM) micro/nano-indentations, for a JPK&#160;AFM. Specifically, in this protocol we measure the apparent Young&#8217;s modulus of cell walls at subcellular resolutions across regions of up to 100 &#181;m&#160;x 100 &#181;m in floral meristems, hypocotyls, and roots. This requires careful preparation of the sample, the correct selection of micro-indenters and indentation depths. To account for cell wall properties only, measurements are performed in highly concentrated solutions of mannitol in order to plasmolyze the cells and thus remove the contribution of cell turgor pressure.
In contrast to other extant techniques, by using different indenters and indentation depths, this method allows simultaneous multiscale measurements, i.e. at subcellular resolutions and across hundreds of cells comprising a tissue. This means that it is now possible&#160;to spatially-temporally characterize the changes that take place in the mechanical properties of cell walls during development, enabling these changes to be correlated with growth and differentiation. This represents a key step to understand how coordinated microscopic cellular changes bring about macroscopic morphogenetic events.
However, several limitations remain: the method can only be used on fairly small samples (around 100 &#181;m in diameter) and only on external tissues; the method is sensitive to tissue topography; it measures only certain aspects of the tissue&#8217;s complex mechanical properties. The technique is being developed rapidly and it is likely that most of these limitations will be resolved in the near future.

We describe a method to map mechanical properties of plant tissues using an atomic force microscope (AFM). We focus on how to record mechanical changes that take place in cell walls during plant development at wide-field mesoscale, enabling these changes to be correlated with growth and morphogenesis.

We describe a recently developed method to measure mechanical properties of the surfaces of plant tissues using atomic force microscopy (AFM) ...

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 ...

Quantitative Analysis of Cell Edge Dynamics during Cell Spreading

Cell spreading is a dynamic process in which a cell suspended in media attaches to a substrate and flattens itself from a rounded to a thin and spread-out shape. Following the cell-substrate attachment, the cell forms a thin sheet of lamellipodia emanating from the cell body. In the lamellipodia, globular actin (G-actin) monomers polymerize into a dense filamentous actin (F-actin) meshwork that pushes against the plasma membrane, thereby providing the mechanical forces required for the cell to spread. Notably, the molecular players that control the actin polymerization in lamellipodia are essential for many other cellular processes, such as cell migration and endocytosis. 
Since spreading cells form continuous lamellipodia that span the entire cell periphery and persistently expand outward, cell spreading assays have become an efficient tool to assess the kinetics of lamellipodial protrusions. Although several technical implementations of the cell spreading assay have been developed, a detailed description of the workflow, which would include both a step-by-step protocol and computational tools for data analysis, is currently lacking. Here, we describe the experimental procedures of the cell spreading assay and present an open-source tool for quantitative and unbiased analysis of cell edge dynamics during spreading. When combined with pharmacological manipulations and/or gene-silencing techniques, this protocol is amenable to a large-scale screen of molecular players regulating lamellipodial protrusions.

In this protocol, we present the experimental procedures of a cell spreading assay that is based on live-cell microscopy. We provide an open-source computational tool for the unbiased segmentation of fluorescently labeled cells and quantitative analysis of lamellipodia dynamics during cell spreading.

Cell spreading is a dynamic process in which a cell suspended in media attaches to a substrate and flattens itself from a rounded to a thin and spread-out ...

Kinematic Analysis of Cell Division and Expansion: Quantifying the Cellular Basis of Growth and Sampling Developmental Zones in Zea mays Leaves

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Experimental Design for Laser Microdissection RNA-Seq: Lessons from an Analysis of Maize Leaf Development

Observing Mitotic Division and Dynamics in a Live Zebrafish Embryo

Confocal Live Imaging of Shoot Apical Meristems from Different Plant Species

Use of Atomic Force Microscopy to Measure Mechanical Properties and Turgor Pressure of Plant Cells and Plant Tissues

Live Cell Imaging of Microtubule Cytoskeleton and Micromechanical Manipulation of the Arabidopsis Shoot Apical Meristem

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

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

AFM-based Mapping of the Elastic Properties of Cell Walls: at Tissue, Cellular, and Subcellular Resolutions

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

Quantitative Analysis of Cell Edge Dynamics during Cell Spreading