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/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/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/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 n...

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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>1 mm glass slide&#160;</td>
			<td>Epredia&#160;</td>
			<td>16211551</td>
			<td></td>
		</tr>
		<tr>
			<td>35 mm glass bottom dish (14 mm well)</td>
			<td>Mattek&#160;</td>
			<td>P35G-1.5-14-C</td>
			<td></td>
		</tr>
		<tr>
			<td>Calcium Chloride&#160;</td>
			<td>Roth&#160;</td>
			<td>CN93.1</td>
			<td></td>
		</tr>
		<tr>
			<td>Columbia (Col-0)&#160;</td>
			<td>Nottingham Arabidopsis Stock Centre (NASC)</td>
			<td></td>
			<td>stigma donor</td>
		</tr>
		<tr>
			<td>Dissecting Scope&#160;</td>
			<td>Olympus&#160;</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 (Ler-0)&#160;</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&#160;</td>
			<td>Scotch</td>
			<td>768720</td>
			<td>Less thick and good for stigma dissection</td>
		</tr>
		<tr>
			<td>Sodium Chloride&#160;</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&#160;</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 poll...

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

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

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 ...

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 ...

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

The problem of acquiring high-resolution images deep into biological samples is widely acknowledged1. In air-filled tissue such as the spongy mesophyll of plant leaves or vertebrate lungs further difficulties arise from multiple transitions in refractive index between cellular components, between cells and airspaces and between the biological tissue and the rest of the optical system. Moreover, refractive index mismatches lead to attenuation of fluorophore excitation and signal emission in fluorescence microscopy. We describe here the application of the perfluorocarbon, perfluorodecalin (PFD), as an infiltrative imaging medium which optically improves laser scanning confocal microscopy (LSCM) sample imaging at depth, without resorting to damaging increases in laser power and has minimal physiological impact2. We describe the protocol for use of PFD with Arabidopsis thaliana leaf tissue, which is optically complex as a result of its structure (Figure 1). PFD has a number of attributes that make it suitable for this use3. The refractive index of PFD (1.313) is comparable with that of water (1.333) and is closer to that of cytosol (approx. 1.4) than air (1.000). In addition, PFD is readily available, non-fluorescent and is non-toxic. The low surface tension of PFD (19 dynes cm-1) is lower than that of water (72 dynes cm-1) and also below the limit (25 - 30 dyne cm-1) for stomatal penetration4, which allows it to flood the spongy mesophyll airspaces without the application of a potentially destructive vacuum or surfactant. Finally and crucially, PFD has a great capacity for dissolving CO2 and O2, which allows gas exchange to be maintained in the flooded tissue, thus minimizing the physiological impact on the sample. These properties have been used in various applications which include partial liquid breathing and lung inflation5,6, surgery7, artificial blood8, oxygenation of growth media9, and studies of ice crystal formation in plants10. Currently, it is common to mount tissue in water or aqueous buffer for live confocal imaging. We consider that the use of PFD as a mounting medium represents an improvement on existing practice and allows the simple preparation of live whole leaf samples for imaging.

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

We describe the use of perfluorodecalin as an infiltrative mounting medium. This is a simple method for improving depth of imaging in Arabidopsis thaliana leaf tissue with minimal physiological impact.

The problem of acquiring high-resolution images deep into biological samples is widely acknowledged1. In air-filled tissue such as the spongy mesophyll of ...

Live Imaging of Apoptotic Cell Clearance during Drosophila Embryogenesis

The proper elimination of unwanted or aberrant cells through apoptosis and subsequent phagocytosis (apoptotic cell clearance) is crucial for normal development in all metazoan organisms. Apoptotic cell clearance is a highly dynamic process intimately associated with cell death; unengulfed apoptotic cells are barely seen in vivo under normal conditions. In order to understand the different steps of apoptotic cell clearance and to compare 'professional' phagocytes - macrophages and dendritic cells to 'non-professional' - tissue-resident neighboring cells, in vivo live imaging of the process is extremely valuable. Here we describe a protocol for studying apoptotic cell clearance in live Drosophila embryos. To follow the dynamics of different steps in phagocytosis we use specific markers for apoptotic cells and phagocytes. In addition, we can monitor two phagocyte systems in parallel: 'professional' macrophages and 'semi-professional' glia in the developing central nervous system (CNS). The method described here employs the Drosophila embryo as an excellent model for real time studies of apoptotic cell clearance.

Live Imaging of Apoptotic Cell Clearance during Drosophila Embryogenesis

Here we describe an effective method for studying dynamics of apoptotic cell clearance in vivo. This method employs live Drosophila embryos as a powerful model for monitoring phagocytosis of apoptotic cells using specific labeling of apoptotic cells and phagocytes.

The  proper elimination of unwanted or aberrant cells through apoptosis and  subsequent phagocytosis (apoptotic cell clearance) is crucial for normal  ...

Live Cell Imaging of Early Autophagy Events: Omegasomes and Beyond

Autophagy is a cellular response triggered by the lack of nutrients, especially the absence of amino acids. Autophagy is defined by the formation of double membrane structures, called autophagosomes, that sequester cytoplasm, long-lived proteins and protein aggregates, defective organelles, and even viruses or bacteria. Autophagosomes eventually fuse with lysosomes leading to bulk degradation of their content, with the produced nutrients being recycled back to the cytoplasm. Therefore, autophagy is crucial for cell homeostasis, and dysregulation of autophagy can lead to disease, most notably neurodegeneration, ageing and cancer.	
	Autophagosome formation is a very elaborate process, for which cells have allocated a specific group of proteins, called the core autophagy machinery. The core autophagy machinery is functionally complemented by additional proteins involved in diverse cellular processes, e.g. in membrane trafficking, in mitochondrial and lysosomal biology. Coordination of these proteins for the formation and degradation of autophagosomes constitutes the highly dynamic and sophisticated response of autophagy. Live cell imaging allows one to follow the molecular contribution of each autophagy-related protein down to the level of a single autophagosome formation event and in real time, therefore this technique offers a high temporal and spatial resolution.	
	Here we use a cell line stably expressing GFP-DFCP1, to establish a spatial and temporal context for our analysis. DFCP1 marks omegasomes, which are precursor structures leading to autophagosomes formation. A protein of interest (POI) can be marked with either a red or cyan fluorescent tag. Different organelles, like the ER, mitochondria and lysosomes, are all involved in different steps of autophagosome formation, and can be marked using a specific tracker dye. Time-lapse microscopy of autophagy in this experimental set up, allows information to be extracted about the fourth dimension, i.e. time. Hence we can follow the contribution of the POI to autophagy in space and time.

Time-lapse microscopy of fluorescently labeled autophagy markers allows monitoring of the dynamic autophagy response with high temporal resolution. Using specific autophagy and organelle markers in a combination of 3 different colors, we can follow the contribution of a protein to autophagosome formation in a robust spatial and temporal context.

Autophagy is a  cellular response triggered by the lack of nutrients, especially the absence of  amino acids. Autophagy is defined by the formation of ...

The Tomato/GFP-FLP/FRT Method for Live Imaging of Mosaic Adult Drosophila Photoreceptor Cells

The Drosophila eye is widely used as a model for studies of development and neuronal degeneration. With the powerful mitotic recombination technique, elegant genetic screens based on clonal analysis have led to the identification of signaling pathways involved in eye development and photoreceptor (PR) differentiation at larval stages. We describe here the Tomato/GFP-FLP/FRT method, which can be used for rapid clonal analysis in the eye of living adult Drosophila. Fluorescent photoreceptor cells are imaged with the cornea neutralization technique, on retinas with mosaic clones generated by flipase-mediated recombination. This method has several major advantages over classical histological sectioning of the retina: it can be used for high-throughput screening and has proved an effective method for identifying the factors regulating PR survival and function. It can be used for kinetic analyses of PR degeneration in the same living animal over several weeks, to demonstrate the requirement for specific genes for PR survival or function in the adult fly. This method is also useful for addressing cell autonomy issues in developmental mutants, such as those in which the establishment of planar cell polarity is affected.

The Tomato/GFP-FLP/FRT Method for Live Imaging of Mosaic Adult Drosophila Photoreceptor Cells

The Tomato/GFP-FLP/FRT method involves visualizing mosaic photoreceptor cells in living Drosophila. It can be used to follow individual photoreceptor cell fates in the retina for days or weeks. This method is ideal for studies of retinal degeneration and neurodegenerative diseases or photoreceptor cell development.

The Drosophila eye is widely used as a model  for studies of development and neuronal degeneration. With the powerful mitotic  recombination technique, ...

Collection and Analysis of Arabidopsis Phloem Exudates Using the EDTA-facilitated Method

The plant phloem is essential for the long-distance transport of (photo-) assimilates as well as of signals conveying biotic or abiotic stress. It contains sugars, amino acids, proteins, RNA, lipids and other metabolites. While there is a large interest in understanding the composition and function of the phloem, the role of many of these molecules and thus, their importance in plant development and stress response has yet to be determined. One barrier to phloem analysis lies in the fact that the phloem seals itself upon wounding. As a result, the number of plants from which phloem sap can be obtained is limited. One method that allows collection of phloem exudates from several plant species without added equipment is the EDTA-facilitated phloem exudate collection described here. While it is easy to use, it does lead to the wounding of cells and care has to be taken to remove contents of damaged cells. In addition, several controls to prove purity of the exudate are necessary. Because it is an exudation rather than a direct collection of the phloem sap (not possible in many species) only relative quantification of its contents can occur. The advantage of this method over others is that it can be used in many herbaceous or woody plant species (Perilla, Arabidopsis, poplar, etc.) and requires minimal equipment and training. It leads to reasonably large amounts of exudates that can be used for subsequent analysis of proteins, sugars, lipids, RNA, viruses and metabolites. It is simple enough that it can be used in both a research as well as in a teaching laboratory.

Collection and Analysis of Arabidopsis Phloem Exudates Using the EDTA-facilitated Method

Knowledge of the composition of the phloem sap as well as the mechanism of its loading and long-distance transport is essential for the understanding of long-distance signaling in plant development and stress/pathogen response and of assimilate transport. This manuscript describes the collection of phloem exudates using the EDTA-facilitated method.

The plant phloem is essential for the long-distance transport of (photo-) assimilates as well as of signals conveying biotic or abiotic stress. It ...

Production of Haploid Zebrafish Embryos by In Vitro Fertilization

The zebrafish has become a mainstream vertebrate model that is relevant for many disciplines of scientific study. Zebrafish are especially well suited for forward genetic analysis of developmental processes due to their external fertilization, embryonic size, rapid ontogeny, and optical clarity &#8211; a constellation of traits that enable the direct observation of events ranging from gastrulation to organogenesis with a basic stereomicroscope. Further, zebrafish embryos can survive for several days in the haploid state. The production of haploid embryos in vitro is a powerful tool for mutational analysis, as it enables the identification of recessive mutant alleles present in first generation (F1) female carriers following mutagenesis in the parental (P) generation. This approach eliminates the necessity to raise multiple generations (F2, F3, etc.) which involves breeding of mutant families, thus saving the researcher time along with reducing the needs for zebrafish colony space, labor, and the husbandry costs. Although zebrafish have been used to conduct forward screens for the past several decades, there has been a steady expansion of transgenic and genome editing tools. These tools now offer a plethora of ways to create nuanced assays for next generation screens that can be used to further dissect the gene regulatory networks that drive vertebrate ontogeny. Here, we describe how to prepare haploid zebrafish embryos. This protocol can be implemented for novel future haploid screens, such as in enhancer and suppressor screens, to address the mechanisms of development for a broad number of processes and tissues that form during early embryonic stages.

Production of Haploid Zebrafish Embryos by In Vitro Fertilization

The zebrafish is a powerful model system for developmental biology and human disease research due to their genetic similarity with higher vertebrates. This protocol describes a methodology to create haploid zebrafish embryos that can be utilized for forward screen strategies to identify recessive mutations in genes essential for early embryogenesis.

The zebrafish has become a mainstream vertebrate model that is relevant for many disciplines of scientific study. Zebrafish are especially well suited for ...

Standardized Method for High-throughput Sterilization of Arabidopsis Seeds

Arabidopsis thaliana (Arabidopsis) seedlings often need to be grown on sterile media. This requires prior seed sterilization to prevent the growth of microbial contaminants present on the seed surface. Currently, Arabidopsis seeds are sterilized using two distinct sterilization techniques in conditions that differ slightly between labs and have not been standardized, often resulting in only partially effective sterilization or in excessive seed mortality. Most of these methods are also not easily scalable to a large number of seed lines of diverse genotypes. As technologies for high-throughput analysis of Arabidopsis continue to proliferate, standardized techniques for sterilizing large numbers of seeds of different genotypes are becoming essential for conducting these types of experiments. The response of a number of Arabidopsis lines to two different sterilization techniques was evaluated based on seed germination rate and the level of seed contamination with microbes and other pathogens. The treatments included different concentrations of sterilizing agents and times of exposure, combined to determine optimal conditions for Arabidopsis seed sterilization. Optimized protocols have been developed for two different sterilization methods: bleach (liquid-phase) and chlorine (Cl2) gas (vapor-phase), both resulting in high seed germination rates and minimal microbial contamination. The utility of these protocols was illustrated through the testing of both wild type and mutant seeds with a range of germination potentials. Our results show that seeds can be effectively sterilized using either method without excessive seed mortality, although detrimental effects of sterilization were observed for seeds with lower than optimal germination potential. In addition, an equation was developed to enable researchers to apply the standardized chlorine gas sterilization conditions to airtight containers of different sizes. The protocols described here allow easy, efficient, and inexpensive seed sterilization for a large number of Arabidopsis lines.

The objective of this study was to determine the effects of bleach and chlorine gas sterilization on seed germination of a range of Arabidopsis genotypes grown on sterile media. Optimized sterilization protocols have been developed to prevent the growth of microbial contaminants while providing satisfactory seed survival.

Arabidopsis thaliana (Arabidopsis) seedlings often need to be grown on sterile media. This requires prior seed sterilization to prevent the growth of ...

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

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

Fertilization of Xenopus oocytes using the Host Transfer Method

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

Live Imaging of Apoptotic Cell Clearance during Drosophila Embryogenesis

Live Cell Imaging of Early Autophagy Events: Omegasomes and Beyond

The Tomato/GFP-FLP/FRT Method for Live Imaging of Mosaic Adult Drosophila Photoreceptor Cells

Collection and Analysis of Arabidopsis Phloem Exudates Using the EDTA-facilitated Method

Production of Haploid Zebrafish Embryos by In Vitro Fertilization

Standardized Method for High-throughput Sterilization of Arabidopsis Seeds