We thank Sara Simonini and Stefano Bencivenga for donating the pFG:roGFP2-ORP1-NLS construct and Christof Eichenberger, Johann Almendinger, Vincent Sutter, and Celia Baroux for their advice on microscopy. We kindly acknowledge advice from Ravi Palanivelu, Philipp Denninger, Sharon Kessler, Mark Johnson, Tomokazu Kawashima, and everyone else at the International Conference on Sexual Plant Reproduction 2022 who showed interest in a protocol on SIV cum septum. This work was supported by the University of Zurich and grants from the Swiss National Science Foundation to U.G.

NOTE: See the Table of Materials for a list of the materials and equipment used in this protocol.
1. Considerations for designing the imaging experiment
<ol>
	<li>Predetermine the imaging duration and sampling frequency required to capture the desired phenomenon to be observed. For example, following Nyquist&#39;s sampling theorem, the sampling frequency must be greater than twice the highest frequency of the input sample. Therefore, to accurately capture synergid [Ca2+]cyt oscillations, perform imaging approximately every 10 s, but to measure the velocity of the sperm cells upon pollen tube discharge, ensure a temporal resolution of less than 1 s10.</li>
	<li>Select fluorescent markers for the cells of interest that are sufficiently bright beyond the background autofluorescence. For the use of biosensors, select the brightest lines that do not perturb the cell function, as they are generally more sensitive and require less exposure time.</li>
	<li>Consider which type of fluorescence microscopy and magnification are most suitable for capturing the desired phenomenon. Use widefield microscopy and low-magnification objectives to capture light from a greater depth of field, and maintain the focus over time, image larger objects such as whole ovules, or use ratiometric sensors that are sensitive to chromatic shift artifacts. Use CLSM or 2PEM with high-magnification objectives for better resolution of cellular and subcellular objects, but note the difficulty in maintaining focus over time.</li>
	<li>Estimate the number of samples to be imaged per experiment. For example, use a 10x objective with a minimum sampling time of ~30 s for imaging three septa and imaging up to 40 fertilization events per experiment because of the time required for the autofocus and multi-stage acquisition functions.</li>
</ol>
2. Pollen germination medium preparation
<ol>
	<li>Prepare liquid pollen germination medium (5mM CaCl2, 1mM MgSO4, 5mM KCl, 0.01% [w/v] H3BO3, and 10% [w/v] sucrose) using 1 M aliquot stocks of each micronutrient, which are stored at &#8722;20 &#176;C. Adjust the pH to 7.5 with 0.1 M KOH. Store the medium at 4 &#176;C for up to 2 weeks. 
	NOTE: The pH may drop over time.</li>
	<li>Add 2.5 mL of liquid medium to a 10 mL beaker, and slowly add 32 mg of ultra-low melting point agarose (~1.3%) with a fast-rotating stir bar to avoid clumping.</li>
	<li>Melt agarose on a hotplate at 65 &#176;C, and nutate the beaker to collect any condensation from the sides.</li>
	<li>Pipette 130 &#181;L of medium into the 14 mm well of a 35 mm glass bottom dish, and rotate the dish until the medium covers the well.</li>
	<li>Aspirate a total of 40 &#181;L with a pipette from the center of the dish, leaving behind a total volume of 90 &#181;L. 
	NOTE: More medium can be aspirated out of the well to achieve a lower total volume; this may be necessary for high-magnification objectives with low working distances. However, the thinner the agar pad is, the quicker it will dry out.</li>
	<li>Cool the dish on an aluminum block in a humidity chamber to ensure even cooling. Store the plates at 4 &#176;C overnight for use the next morning.</li>
</ol>
3. Floral staging of septum donors, stigma donors, and pollen donors
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/65156/65156fig01.jpg" /> 
Figure 1:&#160;Floral staging of septum donors, stigma donors, and pollen donors. (A)&#160;Stages of Arabidopsis flowers (Ler-0)&#160;within an inflorescence. Buds at stage 12B, which have petals that are about to open and yellow indehiscent anthers, should be emasculated by removing all the sepals, petals, and stamens. Pistils (harboring the female gametophyte maker) are then usable as septum donors 24 h later. (B)&#160;Stages of Arabidopsis flowers (Col-0) within an inflorescence. Buds at stage 12B, which have yellow indehiscent anthers and stigmas that are just barely emerging from the petals, should be emasculated by removing all the sepals, petals, and stamens. The pistils are then usable as stigma donors 24 h later, when they should be pollinated by flowers (harboring the male gametophyte marker) that are open and shedding pollen. Pollinated stigmas should be dissected within 1 h of pollination. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65156/65156fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<ol>
	<li>The morning before imaging, choose Arabidopsis plants to be used as the septum donors and stigma donors; ensure that they are healthy and have recently begun to produce siliques. 
	NOTE: Landsberg erecta (Ler-0) is a good accession for use as a septum donor, since its septa are sturdy, and there are small internodes between the ovules. Columbia (Col-0) is a good accession to use as a stigma donor because they seem to be conducive to pollen tube growth.</li>
	<li>Emasculate at least three stage 12B septum donor flowers and 12 stage 12B stigma donor flowers by removing their stamens, sepals, and petals (Figure 1A, B). Remove also older flowers on the same inflorescence, which could potentially pollinate the emasculated flower.</li>
	<li>In the morning, 24 h later, pollinate the stigma donors with the pollen donor flowers (e.g., harboring pollen tube or sperm cell markers) that are readily shedding pollen. Pollination is complete when the stigmas are almost completely covered with pollen (Figure 1B).</li>
</ol>
4. Septum dissection
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/65156/65156fig02.jpg" /> 
Figure 2: A quick guide to septum and stigma dissection. (A)&#160;Steps of septum dissection. Pin the pistil on double-sided tape with an insulin syringe needle, and make cuts at the style-ovary junction and the ovary-pedicle junction, followed by a shallow cut along the septum of either carpel (Step 1). Peel the carpel walls back onto the tape (Step 2). Cut the replum under the top septum (Step 3). Cut the septum at the style, and remove using forceps at the pedicel (Step 4). Place the septum on agar media, and embed gently with forceps. (B)&#160;Steps of stigma dissection. Pin the pistil (pollinated &#60;1 h before) on double-sided tape, and cut at the style-ovary junction with a razor blade (Step 6). Place the stigma on agar medium with an insulin needle next to the septum, and adjust the distance to about 250 &#181;m (Steps 7-8). Ensure that a semi-liquid pool forms around the micropyles and base of the styles (Step 9). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65156/65156fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<ol>
	<li>Thirty minutes after pollination, collect healthy and mature septum donor pistils at the pedicle, and place them onto fresh double-sided sticky tape mounted on a glass slide, with the stigma sticking just off the edge of the tape. 
	NOTE: Limit the dissection time for which the pistils are on the sticky tape to ensure it is as short as possible. The dissection should be conducted at high humidity, which can be achieved by placing a humidifier next to the slide.</li>
	<li>Pin the pistil securely onto the tape by pressing gently on the pedicle and along the ovary using the back side of a new insulin syringe needle before removing the sepal, petal, and stamen debris left over from the emasculation.</li>
	<li>Under a stereoscope, use an insulin syringe needle to make two cuts per carpel at the style-ovary junction and the ovary-pedicle junction, followed by shallow cuts along the septum of each carpel (Figure 2, step 1). 
	NOTE: Cutting too deeply along the septum will sever ovules at the funiculus.</li>
	<li>Peel back the ovary walls carefully onto the tape (Figure 2, step 2), and slide the needle gently between the two septa to cut the replum along the entire length of the pistil without disturbing the ovules (Figure 2, step 3). 
	NOTE: Cut the replum in the direction from the style to the pedicle to ensure that the ovules on the top septum are disturbed as little as possible.</li>
	<li>Cut the top septum gently at both the junction with the style and with the pedicle, and remove the septum with forceps from the pedicle side (Figure 2, step 4). 
	NOTE: Keep this cut straight and gentle so that the septum stays flat and does not curve.</li>
	<li>Place the septum as flat as possible on the medium such that the micropyles of the ovules are face up and in the same focal plane. Press the septum gently into the medium until the ovules are just slightly embedded in the medium (Figure 2, step 5). 
	NOTE: It is critical that the septum be placed such that the micropyles of the ovules are accessible to the pollen tubes (any inaccessible ovules can be gently rearranged with a needle). A small pool of liquid should form around the septum after embedding.</li>
	<li>Repeat steps 4.1-4.6 for up to two more pistils, placing the septa end to end.</li>
</ol>
5. Stigma dissection
<ol>
	<li>Place the pollinated stigma donor pistils on fresh double-sided sticky tape mounted on a glass slide, such that the ovary-style junction is off the edge of the tape (Figure 2, step 6).</li>
	<li>Use a fresh razor blade to cut the style straight down and lift away to keep the stigma attached to the blade (Figure 2, step 6). 
	NOTE: Cut at the edge of the tape for the cleanest cut.</li>
	<li>Remove the stigma from the blade with an insulin syringe needle, and place it flat approximately 250-300 &#181;m from the ovules (Figure 2, steps 7-8). 
	NOTE: The style should be oriented horizontally on its side; otherwise, most of the pollen tubes will grow down under the septum. A small pool of liquid at the entry of the style should appear after it has been lying on the medium for 1 min; this is a good sign, as the pollen tubes will have a smooth exit from the style.</li>
	<li>Repeat steps 5.1-5.3 for up to 12 stigmas, placing two on either side of each septum. Look for a small pool of liquid forming around the micropyle of the ovules; this aids the accessibility and targeting of the pollen tubes to the embryo sacs (Figure 2, step 9). 
	&#8203;NOTE: Limit the amount of time the dish is open to prevent the medium and ovules from drying out.</li>
</ol>
6. Imaging
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/65156/65156fig04.jpg" /> 
Figure 3: SIV cum septum imaging scheme. (A)&#160;Full SIV cum septum setup with 12 pollinated stigmas and 3 septa, as seen through a stereoscope. (B)&#160;Merged images of the full SIV cum septum setup seen in A 5 h after incubation using a 10x objective, with the five overview areas (~1 mm each) marked for multistage acquisition. The green boxes show the ovules that received pollen tubes undergoing an explosive burst in the synergids. (C)&#160;Closer view of overview area 2 at different time points. Approximately 3 h after incubating in the humidity chamber on the microscope, the pollen tubes should arrive near the micropyles, and by 6 h of imaging, most ovules should have properly received pollen tubes (green squares); these ovules then undergo explosive pollen tube burst (asterisk). Scale bars = (A) 5 mm, (B,C) 500 &#181;m. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65156/65156fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<ol>
	<li>Once the dissection is done and the dish is ready to be imaged, let it incubate in a humidity chamber on the microscope maintained at 92% relative humidity and 21 &#176;C. 
	NOTE: If no humidity chamber is available for the microscope, lining the outer edges of the lid and dish with a small volume of water will help to maintain a high level of humidity in the dish.</li>
	<li>Bring the samples into focus under brightfield at a low light intensity, and then switch to fluorescent light, marking the areas of the sample on the stage overview to be imaged. 
	NOTE: At 10x magnification, about five areas can be marked for a total of three septa (Figure 3). Limit the amount of time used for marking the overview areas to minimize phototoxicity or the warming of the sample by the fluorescent light.</li>
	<li>Begin imaging using a multi-stage acquisition scheme with an autofocus function at the beginning of each overview area. As pollen tubes will emerge from the style ~1 h after incubating the sample, delay the acquisition by 2 h or until the pollen tubes have reached the micropyle. Ensure the autofocus is set with a 100 &#181;m range (7 &#181;m large steps and 1 &#181;m fine steps) to keep the ovules in focus as the septa slightly move and sink over time.</li>
	<li>Stop the image acquisition ~9 h after starting the sample incubation, as by this time, most of the ovules should have attracted and received a pollen tube. 
	NOTE: If a low number of ovules attract pollen tubes, careful observation of the dish under a stereoscope is useful to determine how the ovule or style orientation could be improved next time.</li>
</ol>

<ol>
	<li>Huck, N., Moore, J. M., Federer, M., Grossniklaus, U. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+Arabidopsis+mutant+feronia+disrupts+the+female+gametophytic+control+of+pollen+tube+reception.">The Arabidopsis mutant feronia disrupts the female gametophytic control of pollen tube reception.</a> Development. 130 (10), 2149-2159 (2003).</li><li>Rotman, N., 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=Female+control+of+male+gamete+delivery+during+fertilization+in+Arabidopsis+thaliana.">Female control of male gamete delivery during fertilization in Arabidopsis thaliana.</a> Current Biology. 13 (5), 432-436 (2003).</li><li>Palanivelu, R., Preuss, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Distinct+short-range+ovule+signals+attract+or+repel+Arabidopsis+thaliana+pollen+tubes+in+vitro.">Distinct short-range ovule signals attract or repel Arabidopsis thaliana pollen tubes in vitro.</a> BMC Plant Biology. 6, 7 (2006).</li><li>Susaki, D., Maruyama, D., Yelagandula, R., Berger, F., Kawashima, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Live-cell+imaging+of+F-actin+dynamics+during+fertilization+in+Arabidopsis+thaliana.">Live-cell imaging of F-actin dynamics during fertilization in Arabidopsis thaliana.</a> Methods in Molecular Biology. 1669, 47-54 (2017).</li><li>Iwano, M., 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=Cytoplasmic+Ca2++changes+dynamically+during+the+interaction+of+the+pollen+tube+with+synergid+cells.">Cytoplasmic Ca2+ changes dynamically during the interaction of the pollen tube with synergid cells.</a> Development. 139 (22), 4202-4209 (2012).</li><li>Ngo, Q., Vogler, H., Lituiev, D., Nestorova, A., Grossniklaus, U. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+calcium+dialog+mediated+by+the+FERONIA+signal+transduction+pathway+controls+plant+sperm+delivery.">A calcium dialog mediated by the FERONIA signal transduction pathway controls plant sperm delivery.</a> Developmental Cell. 29 (4), 491-500 (2014).</li><li>Denninger, P., 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=Male-female+communication+triggers+calcium+signatures+during+fertilization+in+Arabidopsis.">Male-female communication triggers calcium signatures during fertilization in Arabidopsis.</a> Nature Communications. 5, 4645 (2014).</li><li>Hamamura, Y., 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=Live+imaging+of+calcium+spikes+during+double+fertilization+in+Arabidopsis.">Live imaging of calcium spikes during double fertilization in Arabidopsis.</a> Nature Communications. 5, 4722 (2014).</li><li>Ponvert, N., Johnson, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Synergid+calcium+ion+oscillations+define+a+new+feature+of+pollen+tube+reception+critical+for+blocking+interspecific+hybridization.">Synergid calcium ion oscillations define a new feature of pollen tube reception critical for blocking interspecific hybridization.</a> bioRxiv. , (2020).</li><li>Hamamura, Y., 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=Live-cell+imaging+reveals+the+dynamics+of+two+sperm+cells+during+double+fertilization+in+Arabidopsis+thaliana.">Live-cell imaging reveals the dynamics of two sperm cells during double fertilization in Arabidopsis thaliana.</a> Current Biology. 21 (6), 497-502 (2011).</li></ol>

The authors report no conflicts of interest.

This manuscript introduces an efficient protocol for the imaging of pollen tube reception and double fertilization in Arabidopsis. The improved method, SIV cum septum,&#160;greatly increases the percentage and total number of successful pollen tube reception events that are observable per imaging session. The representative results shown here demonstrate an imaging session with 41 successful pollen tube reception events and 10 ovules showing reception defects (~80% efficiency). This is over double the number of successful pollen tube reception events captured in a total of eight imaging sessions using the original SIV excised ovules method (Video 1). Using the SIV cum septum method in five technical replicates for a total of eight septa, we achieved an average pollen tube reception efficiency of 79% in comparison to an average of 28% using the SIV excised septum method with eight technical replicates (Figure 4).
While the efficiency of pollen tube reception is much greater for the SIV cum septum method, instances of pollen tube overgrowth still occur at varying rates, depending on several factors. Most importantly, the synergid cells must be kept alive and receptive for the several hours over which they are being imaged. Maintaining a humid environment during dissection and imaging, using freshly made medium, embedding the ovules and septum into the semi-liquid agar, and avoiding heat generated by excessive light exposure during image acquisition are all crucial factors to ensure successful pollen tube reception. The accessibility and attraction of pollen tubes to the micropyles of the ovules are also key factors that limit the number of successful pollen tube reception events. Minimizing the disturbance of the ovules on the septum during dissection, orienting the septum such that the micropyles are facing upward, and the formation of a semi-liquid pool between the ovules and the opening of the style are key for pollen tube attraction. The formation of this pool could vary between brands of low-melting point agar, and it is, therefore, recommended to begin the experiments with varying percentages of agar from 1% to 1.5%. Finally, the difficulty of the dissection means it requires some practice, and it is recommended to stay patient, practice relaxing and deep breathing before the dissection, and, importantly, to keep note of the techniques that improve stability and consistency (e.g., hand positioning, slide orientation).
This detailed protocol of the SIV cum septum method should greatly improve the efficiency and sample number for experiments aimed at the live imaging of pollen tube reception and double fertilization. Applications include the use of genetically encoded biosensors, mutant analysis, and descriptive observations of cytological events before, during, and just after double fertilization. We hope that this method may be expanded upon and extended to other flowering plant species where maintaining ovules on the ovary&#39;s placenta could be of benefit.

The generation of genetically unique offspring in sexually reproducing organisms is dependent on the successful fusion of male and female gametes. In flowering plants, the interaction of two male gametes (sperm cells) with two female gametes (egg cell and central cell) during double fertilization depends on sperm release from the pollen tube (the male gametophyte). This process, called pollen tube reception, is largely controlled by the synergid cells that reside within the embryo sac (the female gametophyte)1,2. As pollen tube reception takes place deep inside the flower, a method allowing for live-cell imaging of the process, called semi-in vitro (SIV) pollen tube reception, has been established3. With this method, excised Arabidopsis ovules are placed on semi-liquid pollen germination medium and targeted by pollen tubes that grow through the stigma and style of a pistil severed at the style-transmitting tract junction3,4. Since the development of this technique, detailed observations have led to several discoveries surrounding pollen tube guidance, reception, and fertilization. Among others, these discoveries include the acquisition of pollen tube targeting competence by growth through the stigma3, the onset of intracellular calcium oscillations in the synergids upon pollen tube arrival5,6,7,8,9, and the dynamics of sperm cell release and fertilization upon pollen tube burst10. Nevertheless, because this technique relies on the excision of ovules, the observations of fertilization are limited in number, and pollen tube reception is often aberrant, resulting in the failure of pollen tube rupture (Video 1 and Supplementary File 1). Therefore, there is a need for a more efficient approach allowing for high-throughput analyses of pollen tube reception and fertilization.
In developing this protocol, several new approaches to analyze pollen tube reception, spanning from the most &#34;in vitro&#34; to the most &#34;in vivo&#34; methods, were tested, and an efficient technique based on the excision of the entire septum was settled upon, which allows for up to 40 observations of fertilization per day. Here, the nuances and critical points of the technique are outlined, including flower staging, dissection, medium preparation, and imaging settings. By following this protocol, research focusing on pollen tube guidance, pollen tube reception, and double fertilization should be facilitated. The higher sample sizes the method allows for are expected to bolster the scientific soundness of the conclusions drawn from live imaging experiments. The potential applications of this technique include, but are not limited to, performing observations of the molecular and physiological changes in cytosolic calcium concentrations ([Ca2+]cyt), pH, or H2O2 during gametophyte interactions through the use of genetically encoded biosensors. Furthermore, cytological changes, such as degeneration of the receptive synergid, sperm cell migration, or karyogamy, can be more easily observed using this improved method. Finally, the timing of the different stages of fertilization can be monitored under widefield microscopy, and then more detailed analyses using confocal laser scanning microscopy (CLSM) or two-photon excitation microscopy (2PEM) can be conducted for higher resolution and 3D reconstruction.

<table><tbody><tr><td>1 mm glass slide&amp;nbsp;</td><td>Epredia&amp;nbsp;</td><td>16211551</td><td></td></tr><tr><td>35 mm glass bottom dish (14 mm well)</td><td>Mattek&amp;nbsp;</td><td>P35G-1.5-14-C</td><td></td></tr><tr><td>Calcium Chloride&amp;nbsp;</td><td>Roth&amp;nbsp;</td><td>CN93.1</td><td></td></tr><tr><td>Columbia (Col-0)&amp;nbsp;</td><td>Nottingham Arabidopsis Stock Centre (NASC)</td><td></td><td>stigma donor</td></tr><tr><td>Dissecting Scope&amp;nbsp;</td><td>Olympus&amp;nbsp;</td><td>SZX2-ILLT</td><td></td></tr><tr><td>Insulin needle (0.3 G)</td><td>BD</td><td>304000</td><td></td></tr><tr><td>Landsberg erecta (L&lt;em&gt;er&lt;/em&gt;-0)&amp;nbsp;</td><td>Nottingham Arabidopsis Stock Centre (NASC)</td><td></td><td>septum donor</td></tr><tr><td>Magnesium Sulfate</td><td>Merck</td><td>5886</td><td></td></tr><tr><td>Potassium Chloride</td><td>Roth</td><td>6781.1</td><td></td></tr><tr><td>Razor blade</td><td>Beldura</td><td>7026797</td><td></td></tr><tr><td>Scotch double sided tape&amp;nbsp;</td><td>Scotch</td><td>768720</td><td>Less thick and good for stigma dissection</td></tr><tr><td>Sodium Chloride&amp;nbsp;</td><td>Roth</td><td>3957.1</td><td></td></tr><tr><td>Sucrose</td><td>ITW reagents</td><td>A2211,1000</td><td></td></tr><tr><td>Tesa double sided tape&amp;nbsp;</td><td>Tesa</td><td>05681-00018</td><td>Very sticky and good for septum dissection</td></tr><tr><td>Ultra low gelling temperature agarose</td><td>FMC SeaPrep</td><td>50302</td><td></td></tr></tbody></table>

live imaging of arabidopsis pollen tube reception and double fertilization using the semi-in vitro cum septum method

In flowering plants, the growth and guidance of the pollen tube (male gametophyte) within the pistil and the reception of the pollen tube by the female gametophyte are essential for double fertilization and subsequent seed development. The interactions between male and female gametophytes during pollen tube reception culminate in pollen tube rupture and the release of two sperm cells to effect double fertilization. As pollen tube growth and double fertilization are deeply hidden within the tissues of the flower, this process is difficult to observe in vivo.
A semi-in vitro (SIV) method for the live-cell imaging of fertilization in the model plant Arabidopsis thaliana has been developed and implemented in several investigations. These studies have helped to elucidate the fundamental features of how the fertilization process occurs in flowering plants and which cellular and molecular changes occur during the interaction of the male and female gametophytes. However, because these live cell imaging experiments involve the excision of individual ovules, they are limited to a low number of observations per imaging session, making this approach tedious and very time-consuming. Among other technical difficulties, a failure of the pollen tubes to fertilize the ovules in vitro is&#160;often reported, which severely confounds such analyses.
Here, a detailed video protocol for the imaging of pollen tube reception and fertilization in an automated and high-throughput manner is provided, allowing for up to 40 observations of pollen tube reception and rupture per imaging session. Coupled with the use of genetically encoded biosensors and marker lines, this method enables the generation of large sample sizes with a reduced time investment. Nuances and critical points of the technique, including flower staging, dissection, medium preparation, and imaging, are clearly detailed in video format to facilitate future research on the dynamics of pollen tube guidance, reception, and double fertilization.

To assess the timing of nuclear degeneration in the receptive synergid with respect to pollen tube rupture in Arabidopsis, as well as to observe whether the left or right synergid is predestined to become the receptive synergid,&#160;the SIV cum septum method described here was employed using a female gametophyte nuclear marker stacked with a synergid cytosolic marker (pFG:roGFP2-ORP1-NLS, pMYB98:roGFP2-ORP1) as the septum donor and a male gametophyte marker (pLAT52:R-GECO) as the pollen donor. The septa and pollinated stigmas were dissected 24 h after emasculation, according to sections 4-5 of the protocol, and placed onto glass bottom dishes with a medium prepared the day before. Each dish was placed in a humidity chamber on the microscope, and a multistage experiment was performed using a 10x objective lens (numerical aperture: 0.4) with five overview areas (Figure 3). Images of each area were taken every 60 s for 9 h with four fluorescent channels: Channel (1) 560/640 for R-GECO (25 ms), Channel (2) 488/520 for roGFP2-ORP1 (60 ms), Channel (3) 387/520 for roGFP2-ORP1 (60 ms), and Channel (4) 387/640 for background subtraction (60 ms) (Table 1). An autofocus function using a 100 &#181;m range with 7 &#181;m coarse stepping and 1 &#181;m fine stepping was employed for each minute before image acquisition. The five image files were opened in FIJI, the channels were separated, and then the images were manually tiled together (Video 2 and Supplementary File 1).
In Video 2 (also see Supplementary File 1), 41 examples of explosive pollen tube rupture in the synergids can be seen within the ovules, with green squares marking the regions of interest. Using the calcium biosensor line pLAT52:RGECO as the pollen donor, instances of explosive pollen tube rupture were seen at the tips of the pollen tubes where they lyse. Notably, there was a rapid increase in the brightness of the biosensor as a result of its sudden exposure to high levels of calcium. Explosive pollen tube rupture can also be analyzed with GFP-based or RFP-based sperm cell or cytosolic markers, which also show rapid release of the sperm or cytoplasmic content. Upon pollen tube rupture, the nucleus of the receptive synergid was observed to degrade simultaneously (within the 1 min time resolution in the experiment), and migration of the egg cell nucleus toward the micropyle was also seen upon pollen tube rupture as a result of karyogamy (Video 3 and Supplementary File 1). Both the left and right synergids were able to serve as the receptive synergid, and instances of two pollen tube bursts were observed, firstly in the receptive synergid and secondly in the persistent synergid. Video 2 (also see Supplementary File 1) shows 10 examples of defective pollen tube reception, which can be seen in the ovules, with the regions of interest marked by red squares. Thus, the overall efficiency of pollen tube reception was ~80% with respect to the ovules that attracted pollen tubes. In these unsuccessful cases, the pollen tubes either arrest their growth in the micropyle, arrest their growth after the fast growth into the synergids, or fail to arrest their growth and continue growing and coiling within the embryo sac. The ovules without any indicated regions of interest failed to attract pollen tubes or were not in an accessible orientation. Enlarged examples of negative and positive outcomes can be seen in Video 3 (also see Supplementary File 1), showing two instances of defective pollen tube reception (Video 3A,C and Supplementary File 1), and two instances of successful pollen tube reception (Video 3B,D and Supplementary File 1).
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/65156/65156fig03.jpg" /> 
Figure 4: Comparison of SIV method efficiencies. Eight technical replicates of the SIV excised ovules method (71 total ovules) showed only ~28% ovules with explosive pollen tube bursts, resulting in an efficiency ranging from ~10%-50%. Five technical replicates of the SIV cum septum method (8 septa and 102 ovules) showed ~79% ovules with explosive pollen tube burst, resulting in an efficiency ranging from ~70%-100%. Only ovules that attracted pollen tubes to the micropyle and ovules that had enough recorded time to receive the attracted pollen tube were counted. The &#34;X&#34; denotes the mean and the center line the median value (50th percentile), respectively. The lines outside the upper and lower quartiles show the minimum and maximum percentages of successful pollen tube reception. The percentage of ovules with pollen tube burst was significantly different (p &#60; 0.001) between the two methods based on an unpaired t-test. Abbreviation: SIV = semi-in vitro. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65156/65156fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Equipment/Setting</td>
			<td>Supplemental Video 2 &#38; 3</td>
		</tr>
		<tr>
			<td>Microscope</td>
			<td>Olympus IX-81F-ZDC2 inverted microscope</td>
		</tr>
		<tr>
			<td>Temperature control</td>
			<td>Life Imaging Services GmbH Cube &#38; Box (22C)</td>
		</tr>
		<tr>
			<td>Humidity control&#160;</td>
			<td>Life Imaging Services GmbH Brick (92% relative humidity)</td>
		</tr>
		<tr>
			<td>Software</td>
			<td>Olympus cellSens Dimension 2.3</td>
		</tr>
		<tr>
			<td>Real time controller</td>
			<td>Olympus U-RTC</td>
		</tr>
		<tr>
			<td>Laser</td>
			<td>MT20- &#160;150 W xenon arc burner&#160;</td>
		</tr>
		<tr>
			<td>Objective lens&#160;</td>
			<td>10x air immersion U Plan SApo objective lens (0.4 NA), working distance 3.1 mm</td>
		</tr>
		<tr>
			<td>Mirror Cube</td>
			<td>GFP/RFP&#160;</td>
		</tr>
		<tr>
			<td rowspan="5">Filter Wheel</td>
			<td>Olympus U-FFWO</td>
		</tr>
		<tr>
			<td>Channel 1 (25 ms) : reflected (560/25) observation (624/40)</td>
		</tr>
		<tr>
			<td>Channel 2 (60 ms): reflected (485/20) observation (525/30)</td>
		</tr>
		<tr>
			<td>Channel 3 (60 ms): reflected (387/11) observation (525/30)</td>
		</tr>
		<tr>
			<td>Channel 4 (60 ms):&#160; reflected (387/11) observation (624/30)</td>
		</tr>
		<tr>
			<td>Detector&#160;</td>
			<td>Hamamatsu ORCA-Fusion Digital CMOS camera</td>
		</tr>
		<tr>
			<td>Autofocus&#160;</td>
			<td>100 &#181;m range, 7 &#181;m coarse step, 1 &#181;m fine step</td>
		</tr>
		<tr>
			<td>Time interval</td>
			<td>1 min</td>
		</tr>
	</tbody>
</table>
Table 1: The microscope systems and settings used in this manuscript. 
Video 1: Timelapses of the SIV excised ovules method. Merged timelapse from eight technical replicates of SIV using excised ovules showing the growth of pollen tubes (harboring pLAT52:R-GECO) and pollen tube reception within the ovules. The 20 green boxes show instances of explosive pollen tube burst in the synergids, and the 51 red boxes show instances of failed pollen tube burst and overgrowth. The time lapses are between 260 min and 400 min, and the scale bar is 100 &#181;m. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65156/Movie 1.mp4" target="_blank">Please click here to download this Video.</a>
Video 2: SIV cum septum timelapse. (A)&#160;Merged timelapse from all five overview areas (see Figure 3) of pollen tube growth and reception within the ovules with just the RFP channel (Channel 1). The pollen is harboring pLAT52:R-GECO and the ovules pFG:roGFP2-ORP1-NLS; pMYB98:roGFP2-ORP1. The 41 green boxes show instances of explosive pollen tube burst, and the 10 red boxes show instances of failed pollen tube burst and pollen tube overgrowth. (B)&#160;The same timelapse with both the RFP and GFP channels (Channel 1 and Channel 2), showing the nuclear dynamics in the gametes and synergid cells during pollen tube reception. The numbers on the top right indicate the time (h:min). Scale bar = 500 &#181;m. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65156/Movie 2.mp4" target="_blank">Please click here to download this Video.</a>
Video 3: Examples of positive and negative outcomes from SIV cum septum. (A,C) Example ovules taken from Video 2 showing two instances of failed fertilization. In A, the pollen tubes fail to arrest growth and continue coiling in the embryo sac. In C, a second category of pollen tube overgrowth is seen, where the first pollen tube undergoes fast growth as it grows into the synergid but arrests growth without rupturing. (B,D)&#160;Example ovules taken from Video 2 showing two instances of successful pollen tube reception. The wild-type-like pollen tube burst shows the explosive rupture of the pollen tube in the synergids, and the migration of the egg cell nucleus toward the micropyle indicates karyogamy. The numbers on the top right indicate the time (h:min). The arrowheads indicate the receptive synergid nuclei (rSN), and the arrows indicate the migration of the egg cell nucleus (EN) after pollen tube burst (lysis). Scale bar = 30 &#181;m. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65156/Movie 3.mp4" target="_blank">Please click here to download this Video.</a>
Supplementary File 1: Still images of Video 1, Video 2, and Video 3. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65156/65156_Supplementary File 1.pdf" target="_blank">Please click here to download this File.</a>

Watch this Scientific Journal Video about Live Imaging of Arabidopsis Pollen Tube Reception and Double Fertilization Using the Semi-In Vitro Cum Septum Method at JoVE.com

Live Imaging of Arabidopsis Pollen Tube Reception and Double Fertilization Using the Semi-In Vitro Cum Septum Method

Here, we describe an improvement of the semi-in vitro (SIV) method for observing pollen tube guidance and reception in Arabidopsis thaliana,&#160;which increases the receptivity of ovules. The high-throughput SIV cum septum method may be coupled with gametophyte marker lines and genetically encoded biosensors to monitor the dynamic process of fertilization.

Live Imaging of Arabidopsis Pollen Tube Reception and Double Fertilization Using the Semi-In Vitro Cum Septum Method

In flowering plants, the growth and guidance of the pollen tube (male gametophyte) within the pistil and the reception of the pollen tube by the female ...

live-imaging-arabidopsis-pollen-tube-reception-double-fertilization

Nanyang Technological University

Research

JoVE Journal

Biology

3.8K Views.  University of Zurich. Here, we describe an improvement of the semi-in vitro (SIV) method for observing pollen tube guidance and reception in Arabidopsis thaliana, which increases the receptivity of ovules. The high-throughput SIV cum septum method may be coupled with gametophyte marker lines and genetically encoded biosensors to monitor the dynamic process of fertilization.

Video: Live Imaging of Arabidopsis Pollen Tube Reception and Double Fertilization Using the Semi-In Vitro Cum Septum Method

Live Confocal Imaging of Developing Arabidopsis Flowers

The study of plant growth and development has long relied on experimental techniques using dead, fixed tissues and lacking proper cellular resolution. Recent advances in confocal microscopy, combined with the development of numerous fluorophores, have overcome these issues and opened the possibility to study the expression of several genes simultaneously, with a good cellular resolution, in live samples. Live confocal imaging provides plant biologists with a powerful tool to study development, and has been extensively used to study root growth and the formation of lateral organs on the flanks of the shoot apical meristem. However, it has not been widely applied to the study of flower development, in part due to challenges that are specific to imaging flowers, such as the sepals that grow over the flower meristem, and filter out the fluorescence from underlying tissues. Here, we present a detailed protocol to perform live confocal imaging on live, developing Arabidopsis flower buds, using either an upright or an inverted microscope.

Live confocal imaging provides biologists with a powerful tool to study development. Here, we present a detailed protocol for the live confocal imaging of developing Arabidopsis flowers.

The study of plant growth and development has long relied on experimental techniques using dead, fixed tissues and lacking proper cellular resolution. ...

Developmental Biology

A Method for Characterizing Embryogenesis in Arabidopsis

Given the highly predictable nature of their development, Arabidopsis embryos have been used as a model for studies of morphogenesis in plants. However, early stage plant embryos are small and contain few cells, making them difficult to observe and analyze. A method is described here for characterizing pattern formation in plant embryos under a microscope using the model organism Arabidopsis. Following the clearance of fresh ovules using Hoyer's solution, the cell number in and morphology of embryos could be observed, and their developmental stage could be determined by differential interference contrast microscopy using a 100X oil immersion lens. In addition, the expression of specific marker proteins tagged with Green Fluorescent Protein (GFP) was monitored to annotate cell identity specification during embryo patterning by confocal laser scanning microscopy. Thus, this method can be used to observe pattern formation in wild-type plant embryos at the cellular and molecular levels, and to characterize the role of specific genes in embryo patterning by comparing pattern formation in embryos from wild-type plants and embryo-lethal mutants. Therefore, the method can be used to characterize embryogenesis in Arabidopsis.

A Method for Characterizing Embryogenesis in Arabidopsis

This protocol outlines a method for observing embryogenesis in Arabidopsis via ovule clearance followed by the inspection of embryo pattern formation under a microscope.

Given the highly predictable nature of their development, Arabidopsis embryos have been used as a model for studies of morphogenesis in plants. However, ...

In Vitro Ovule Cultivation for Live-cell Imaging of Zygote Polarization and Embryo Patterning in Arabidopsis thaliana

In most flowering plants, the zygote and embryo are hidden deep in the mother tissue, and thus it has long been a mystery of how they develop dynamically; for example, how the zygote polarizes to establish the body axis and how the embryo specifies various cell fates during organ formation. This manuscript describes an in vitro ovule culture method to perform live-cell imaging of developing zygotes and embryos of Arabidopsis thaliana. The optimized cultivation medium allows zygotes or early embryos to grow into fertile plants. By combining it with a poly(dimethylsiloxane) (PDMS) micropillar array device, the ovule is held in the liquid medium in the same position. This fixation is crucial to observe the same ovule under a microscope for several days from the zygotic division to the late embryo stage. The resulting live-cell imaging can be used to monitor the real-time dynamics of zygote polarization, such as nuclear migration and cytoskeleton rearrangement, and also the cell division timing and cell fate specification during embryo patterning. Furthermore, this ovule cultivation system can be combined with inhibitor treatments to analyze the effects of various factors on embryo development, and with optical manipulations such as laser disruption to examine the role of cell-cell communication.

In Vitro Ovule Cultivation for Live-cell Imaging of Zygote Polarization and Embryo Patterning in Arabidopsis thaliana

This manuscript describes an in vitro ovule cultivation method that enables live-cell imaging of Arabidopsis zygotes and embryos. This method is utilized to visualize the intracellular dynamics during zygote polarization and the cell fate specification in developing embryos.

In most flowering plants, the zygote and embryo are hidden deep in the mother tissue, and thus it has long been a mystery of how they develop dynamically; ...

The Preparation of Drosophila Embryos for Live-Imaging Using the Hanging Drop Protocol

Green fluorescent protein (GFP)-based timelapse live-imaging is a powerful technique for studying the genetic regulation of dynamic processes such as tissue morphogenesis, cell-cell adhesion, or cell death. Drosophila embryos expressing GFP are readily imaged using either stereoscopic or confocal microscopy. A goal of any live-imaging protocol is to minimize detrimental effects such as dehydration and hypoxia. Previous protocols for preparing Drosophila embryos for live-imaging analysis have involved placing dechorionated embryos in halocarbon oil and sandwiching them between a halocarbon gas-permeable membrane and a coverslip1-3. The introduction of compression through mounting embryos in this manner represents an undesirable complication for any biomechanical-based analysis of morphogenesis. Our method, which we call the hanging drop protocol, results in excellent viability of embryos during live imaging and does not require that embryos be compressed. Briefly, the hanging drop protocol involves the placement of embryos in a drop of halocarbon oil that is suspended from a coverslip, which is, in turn, fixed in position over a humid chamber. In addition to providing gas exchange and preventing dehydration, this arrangement takes advantage of the buoyancy of embryos in halocarbon oil to prevent them from drifting out of position during timelapse acquisition. This video describes in detail how to collect and prepare Drosophila embryos for live imaging using the hanging drop protocol. This protocol is suitable for imaging dechorionated embryos using stereomicroscopy or any upright compound fluorescence microscope.

The Preparation of Drosophila Embryos for Live-Imaging Using the Hanging Drop Protocol

A simple, inexpensive, and effective method of preparing Drosophila embryos for live-imaging analysis is presented. Our protocol provides humidity and gas exchange and does not compress the Drosophila embryo. This method is suitable for GFP-based live imaging of Drosophila embryos using a stereomicroscope or upright compound microscope.

Green fluorescent protein (GFP)-based timelapse live-imaging is a powerful technique for studying the genetic regulation of dynamic processes such as ...

A High-Throughput Method For Zebrafish Sperm Cryopreservation and In Vitro Fertilization

This is a method for zebrafish sperm cryopreservation that is an adaptation of the Harvey method (Harvey et al., 1982).  We have introduced two changes to the original protocol that both streamline the procedure and increase sample uniformity.  First, we normalize all sperm volumes using freezing media that does not contain the cryoprotectant. Second, cryopreserved sperm are stored in cryovials instead of capillary tubes.  The rates of sperm freezing and thawing (&delta;&deg;C/time) are probably the two most critical variables to control in this procedure.  For this reason, do not substitute different tubes for those specified.  Working in teams of 2 it is possible to freeze the sperm of 100 males per team in ~2 hrs.  Sperm cryopreserved using this protocol has an average of 25% fertility (measured as the number of viable embryos generated in an in vitro fertilization divided by the total number of eggs fertilized) and this percent fertility is stable over many years.

A High-Throughput Method For Zebrafish Sperm Cryopreservation and In Vitro Fertilization

This is a high-throughput sperm cryopreservation protocol for zebrafish. Sperm cryopreserved using this protocol has an average of 25% fertility in subsequent vitro fertilization and is stable over many years.

This is a method for zebrafish sperm cryopreservation that is an adaptation of the Harvey method (Harvey et al., 1982).  We have introduced two changes to ...

Evaluation of Fertilization State by Tracing Sperm Nuclear Morphology in Arabidopsis Double Fertilization

Flowering plants have a unique sexual reproduction system called &#8216;double fertilization&#39;,&#160;in which each of the sperm cells precisely fuses with an egg cell or a central cell. Thus, two independent fertilization events take place almost simultaneously. The fertilized egg cell and central cell develop into zygote and endosperm, respectively. Therefore, precise control of double fertilization is essential for the ensuing seed development. Double fertilization occurs in the female gametophyte (embryo sac), which is deeply hidden and covered with thick ovule and ovary tissues. This pistil tissue construction makes observation and analysis of double fertilization quite difficult and has created the present situation in which many questions regarding the mechanism of double fertilization remain unanswered. For the functional evaluation of a potential candidate for fertilization regulator, phenotypic analysis of fertilization is important. To judge the completion of fertilization in Arabidopsis thaliana, the shapes of fluorescence signals labeling sperm nuclei are used as indicators. A sperm cell that fails to fertilize is indicated by a condensed fluorescence signal outside of the female gametes, whereas a sperm cell that successfully fertilizes is indicated by a decondensed signal due to karyogamy with the female gametes&#8217; nucleus. The method described here provides a tool to determine successful or failed fertilization under in vivo conditions.

Evaluation of Fertilization State by Tracing Sperm Nuclear Morphology in Arabidopsis Double Fertilization

We demonstrate a method to determine successful or failed fertilization on the basis of sperm nuclear morphology in Arabidopsis double fertilization using an epifluorescence microscope.

Flowering plants have a unique sexual reproduction system called &#8216;double fertilization&#39;,&#160;in which each of the sperm cells precisely fuses ...

Fertilization of Xenopus oocytes using the Host Transfer Method

Studying the contribution of maternally inherited molecules to vertebrate early development is often hampered by the time and expense necessary to generate maternal-effect mutant animals. Additionally, many of the techniques to overexpress or inhibit gene function in organisms such as Xenopus and zebrafish fail to sufficiently target critical maternal signaling pathways, such as Wnt signaling. In Xenopus, manipulating gene function in cultured oocytes and subsequently fertilizing them can ameliorate these problems to some extent. Oocytes are manually defolliculated from donor ovary tissue, injected or treated in culture as desired, and then stimulated with progesterone to induce maturation. Next, the oocytes are introduced into the body cavity of an ovulating host female frog, whereupon they will be translocated through the host's oviduct and acquire modifications and jelly coats necessary for fertilization. The resulting embryos can then be raised to the desired stage and analyzed for the effects of any experimental perturbations. This host-transfer method has been highly effective in uncovering basic mechanisms of early development and allows a wide range of experimental possibilities not available in any other vertebrate model organism.

Fertilization of Xenopus oocytes using the Host Transfer Method

Procedure for fertilizing Xenopus oocytes by the host transfer method.

Studying the contribution of maternally inherited molecules to vertebrate early development is often hampered by the time and expense necessary to ...

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 ...

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 ...

An Improved Method to Measure Pollen Hydration Profiles in Arabidopsis thaliana

A High-Resolution, Single-Grain, In Vivo Pollen Hydration Bioassay for Arabidopsis thaliana

Sexual reproduction in flowering plants requires initial interaction between the pollen grain and the stigmatic surface, where a molecular dialog is established between the interacting partners. Studies across a range of species have revealed that a series of molecular checkpoints regulate the pollen-stigma interaction to ensure that only compatible, generally intraspecific pollen is successful in effecting fertilization. In species that possess a 'dry stigma', such as the model plant Arabidopsis thaliana, the first post-pollination, prezygotic compatibility checkpoint is the establishment of pollen hydration. 
This phase of pollination is tightly regulated, whereby signals from the pollen grain elicit the release of water from the stigma, thus permitting pollen hydration. The ability to accurately measure and track pollen hydration over time is key to the design of experiments directed at understanding the regulation of this critical step in reproduction. Published protocols frequently utilize flowers that have been excised from the parent plant, maintained on liquid or solid media, and bulk pollinated. 
This paper describes a noninvasive, in vivo pollination bioassay that permits minute-by-minute hydration tracking of individual A. thaliana pollen grains at high resolution. The assay is highly reproducible, able to detect very subtle variations of pollen hydration profiles, and thus is suitable for the analysis of mutants that affect pathways regulating pollination. Although the protocol is lengthier than those described for bulk pollinations, the precision and reproducibility it provides, along with its in vivo nature, make it ideal for the detailed dissection of pollination phenotypes.

Author Spotlight: A High-Resolution, Single-Grain, In Vivo Pollen Hydration Bioassay for Arabidopsis thaliana  

An improved method to measure pollen hydration profiles in Arabidopsis thaliana is described here. The new method offers higher resolution, is noninvasive, and is highly reproducible. The protocol represents a new tool for a finer dissection of the processes that regulate the early stages of pollination.

Sexual reproduction in flowering plants requires initial interaction between the pollen grain and the stigmatic surface, where a molecular dialog is ...

Live Imaging of Arabidopsis Pollen Tube Reception and Double Fertilization Using the Semi-In Vitro Cum Septum Method

Podsumowanie

Przeglądaj więcej filmów

Rozdziały w tym wideo

Live Confocal Imaging of Developing Arabidopsis Flowers

A Method for Characterizing Embryogenesis in Arabidopsis

In Vitro Ovule Cultivation for Live-cell Imaging of Zygote Polarization and Embryo Patterning in Arabidopsis thaliana

The Preparation of Drosophila Embryos for Live-Imaging Using the Hanging Drop Protocol

A High-Throughput Method For Zebrafish Sperm Cryopreservation and In Vitro Fertilization

Evaluation of Fertilization State by Tracing Sperm Nuclear Morphology in Arabidopsis Double Fertilization

Fertilization of Xenopus oocytes using the Host Transfer Method

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

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

A High-Resolution, Single-Grain, In Vivo Pollen Hydration Bioassay for Arabidopsis thaliana