We thank Andres Kaech (University of Zurich) for his helpful advice on scanning electron microscopy imaging. This work was supported by the United States-Israel Binational Agricultural Research and Development Fund (grant no. US-4334-10, Z.R.), the Israel Science Foundation (grant no. 1034/12, Z.R.), and the Human Frontier Science Program (RGP0005/2013, Z.R.). The electron microscopy studies were conducted at the Irving and Cherna Moskowitz Center for Nano and Bio-Nano Imaging at the Weizmann Institute of Science.

1. Cryo-fixation of Leaf Tissues by High-pressure Freezing
Note: This section describes how to carry out high-pressure freezing of leaf tissues for a freeze-fracture experiment. For considerations related to plant samples see29. This can be adapted for other types of tissues or samples with some modification.
<ol>
	<li>Using the corner of a razor blade, scratch the bottom of the 0.1 mm cavity of a 0.1/0.2 mm aluminum platelet (Figure 1). Prepare at least a dozen platelets (6 samples). Take caution not to create knife marks on the circumference of the disc.
	<ol>
		<li>After scratching, wash discs in absolute ethanol, let them dry and keep in a clean dish.</li>
	</ol>
	</li>
	<li>Prepare the high pressure freezing machine according to the manufacturer&#39;s instructions. Fill the high pressure freezing machine&#39;s alcohol chamber with isopropanol.</li>
	<li>Prepare a home-made vacuum-assisted infiltration device to replace gas found in the leaf tissue with liquid.
	<ol>
		<li>Tightly connect a 200 &#181;l pipette tip to the end of a 10 ml syringe. Cut off ~1 cm of the tip. Make a hole in the cap of a 1.5 ml microcentrifuge tube to tightly fit the cut tip. Vacuum can be created inside the tube by pulling the plunger of the syringe.</li>
	</ol>
	</li>
	<li>Cut a small piece of leaf from the plant using a razor blade and with tweezers place the leaf inside a microcentrifuge tube half-filled with 1-hexadecene. Infiltrate the intercellular spaces of the leaf with the liquid by gently pulling the plunger to create vacuum, hold for about 30 sec, and then slowly release the plunger.</li>
	<li>Remove the leaf piece from the tube using tweezers. Trim the leaf with a razor blade in case it is thicker than 0.2 mm and cut a small piece that will fit into a platelet.</li>
	<li>Place a clean platelet with the scratched side up into the freezing machine&#39;s holder and then place the cut piece of leaf into the platelet. Fill the remaining space surrounding the leaf with 1-hexadecene. Close the sandwich using another platelet, with the scratches facing the leaf.</li>
	<li>Close and tighten the holder and insert it into the machine&#39;s chamber. Then press the &#39;Jet&#39; button to freeze (~2,100 bars for 300 - 500 msec), and quickly transfer the holder into liquid nitrogen. Carefully remove the frozen sandwich from the holder using pre-cooled tweezers, taking care not to fracture the sandwich. 
	Note: Closed frozen sandwiches can be freeze fractured immediately or kept in liquid nitrogen for later use. Use protective clothing and eyewear when handling liquid nitrogen.</li>
</ol>
2. Freeze-fracture and Double-layer Coating27,28
<ol>
	<li>Prepare the freeze fracture machine for use, including both guns for evaporation of platinum/carbon (Pt/C) and carbon, according to the manufacturer&#39;s instructions. Place the Pt/C gun at 45 degrees and the carbon gun at 90 degrees.
	<ol>
		<li>Pre-program a coating scheme of a 2-nm layer of Pt/C (coating rate 0.02 - 0.04 nm/sec) and a 6-nm layer of carbon (at ~0.1 nm/sec).
		<ol>
			<li>Enter the setup on the freeze-fracture machine&#39;s screen. Select a user number where the coating scheme will be saved.</li>
			<li>Choose Pt/C for &#39;Layer 1&#39;. Select a time frame,&#160;e.g., 5 min (to exceed the necessary time needed to complete the coating). Use the up-down arrows to define a thickness of 2 nm.</li>
			<li>Press the &#39;Layer&#39; button to move to Layer 2. Repeat step 2.1.1.2, except select carbon and define a thickness of 6 nm.</li>
		</ol>
		</li>
		<li>Test and adjust gun operation.
		<ol>
			<li>Select &#39;Layer 1&#39;. Select the Pt/C gun by pressing the &#39;GUN1&#39; button and press the &#39;ON&#39; button on the evaporation control unit of the freeze-fracture machine. Monitor the evaporation rate and adjust it using the voltage and emission knobs until reaching a rate of 0.02 - 0.04 nm/sec. Press &#39;OFF&#39; to turn off the Pt/C gun.</li>
			<li>Select &#39;Layer 2&#39;. Repeat 2.1.2.1, except select &#39;GUN2&#39; and adjust for an evaporation rate of ~0.1 nm/sec.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Cool the freeze fracture system stage to -140 &#730;C or -160 &#730;C (for hydrated or dehydrated leaves, respectively). Attach the vacuum cryo transfer shuttle to the machine and fill its chamber with liquid nitrogen. Pressure in the cooled freeze-fracture chamber is usually between 1 - 8 x 10-7 mbar.</li>
	<li>Insert a freeze fracturing specimen holder (suitable for 3-mm platelets) into the loading station. Flood the loading station chamber with liquid nitrogen.</li>
	<li>Load two frozen sandwiches onto the holder one by one using high-precision grade tweezers. Tighten the first sandwich into the holder and then flip the holder over to check that it fits securely in place. Repeat for the second sandwich.</li>
	<li>Detach the cooled shuttle from the freeze fracture machine and connect it to the loading station. Open the shuttle valve, introduce the retracting arm into the holder and lock it into place. Re-introduce the arm with the holder into the shuttle and close the shuttle valve using the loading station knob. Attach the shuttle to the freeze-fracture machine and introduce the holder into the chamber with the retracting arm. Detach the shuttle from the machine.</li>
	<li>Align the freeze fracture microtome knife to a height such that it will knock off the top platelets of the sandwiches.</li>
	<li>With the microtome knife, quickly break the sandwiches, and then immediately raise it up to prevent contamination of the samples by debris clinging to the knife, and turn the cooled shutter above the newly exposed plane to shield the samples from possible contamination by water molecules in the chamber.</li>
	<li>Coat the samples with platinum and carbon.
	<ol>
		<li>Enter &#39;Layer 1&#39; on the screen. Turn on the Pt/C gun by pressing the &#39;GUN1&#39; button followed by the &#39;ON&#39; button. Examine the evaporation rate and once it reaches 0.02 - 0.04 nm/sec, simultaneously expose the samples and press the &#39;Measure&#39; button to monitor the thickness of the deposited layer. 
		Note: Gun will turn off automatically when the thickness has reached the pre-selected value.</li>
		<li>Cover the samples with the shutter when Pt/C evaporation is completed.</li>
		<li>Press the &#39;Layer&#39; button to move to &#39;Layer 2&#39;. Repeat steps 2.8.1 and 2.8.2, with the exception of selecting &#39;GUN2&#39; (for carbon) and an evaporation rate of ~0.1 nm/sec.</li>
	</ol>
	</li>
	<li>Warm the freeze fracture system stage to -120 &#730;C.</li>
</ol>
3. Cryo-scanning Electron Microscopy
<ol>
	<li>Prepare the scanning electron microscope for cryo work.
	<ol>
		<li>Select Stage/Stage Initialise from the main menu and confirm.</li>
		<li>Select Tools/Goto Panel, and open the &#39;Airlock&#39; panel by selecting it from the list. Click the &#39;Close Column Chamber Valve&#39; button. Vent the microscope chamber by clicking the Vent button under the &#39;Vacuum&#39; tab.</li>
		<li>Select Tools/Goto Panel, and open the &#39;Stage Points List&#39; panel. Select the cryo-exchange position to move the stage to the proper coordinates for accepting the sample holder. Note: The &#39;cryo-exchange position&#39; is preset to be in alignment with the position of the vacuum transfer unit.</li>
		<li>Open the specimen chamber with a clean pair of gloves and insert the cryo-stage onto the main stage of the microscope. Close the chamber and click the Pump button under the &#39;Vacuum tab&#39;.</li>
		<li>Cool the microscope to -120 &#730;C. Fill the microscope Dewar with liquid nitrogen. Activate the heat function on the cryo-transfer control unit to -120 &#730;C once the temperature of the stage drops below -120 &#730;C.</li>
	</ol>
	</li>
	<li>Attach the cryo-transfer shuttle to the microscope and transfer the sample holder using the retracting arm into the microscope cryo stage (as in step 2.5). Detach the shuttle from the microscope.</li>
	<li>Click on the &#39;Open Column Chamber Valve&#39; button in the Airlock Panel.</li>
	<li>Carefully move the specimen holder close to the objective lens (working distance 1 - 2 mm) using the joystick. Select a 10-&#181;m aperture in the &#39;Apertures&#39; Tab. Double-click the EHT field in the &#39;Gun&#39; tab. Enter 10 (kV) for the EHT target and click OK. Click on the bottom EHT tab and select &#39;EHT On&#39;.</li>
	<li>Select &#39;InLens&#39; from the Detectors drop-down list in the &#39;Detectors&#39; tab. Optimize imaging conditions (focus, brightness and contrast, beam alignment, astigmatism) as appropriate.</li>
	<li>Click on the &#39;Scanning&#39; tab. Choose a fast scan (&#39;Scan Speed = 1&#39;, corresponding to a dwell time of 100 nsec per pixel) to minimize beam damage, and a &#39;Store resolution&#39; of 1,024 x 768 pixels. Select &#39;Line Avg&#39; in the &#39;Noise Reduction&#39; drop-down list. Adjust the value of N to 20 - 30 lines for searching. Move the sample using the &#39;Centre Point&#39; function (by pressing control-tab on the keyboard) to search for regions with fractured cells and chloroplasts (Figures 2A and 2B).</li>
	<li>Acquire images of fractured chloroplasts at low magnifications (50 - 70 kX) (Figure 2C and 2D). Use the &#39;Centre Feature&#39; function (by pressing control-shift-tab on the keyboard) to zoom into interesting chloroplast fracture faces and acquire images at high magnifications (100 - 200 kX, Figures 3 and 4). Use the same parameters as in step 3.6 for image acquisition, but change the Line Avg value to N &#62;100 for noise reduction30.
	<ol>
		<li>Press Freeze on the main microscope control unit or in the &#39;Scanning&#39; tab to stop the scanning and save the image by pressing the &#39;Save TIFF&#39; button.</li>
	</ol>
	</li>
</ol>
4. Image Analysis
Note: This section describes a short procedure for segmentation of membrane particles from freeze-fracture SEM images using the Fiji31 open-source package. Similar results can be obtained with other image analysis software.
<ol>
	<li>Open image with Fiji by dragging image file into the main program window. Convert the image to 8-bit by selecting Image/Type/8-bit. Select Image/Adjust/Brightness/Contrast and adjust as necessary (Figure 5A).</li>
	<li>Using the Straight Line tool, draw a line the size of the embedded image scale bar. Select Analyze/Set Scale. Enter the proper scale information (Known distance and Unit of length) and click OK. Save this adjusted scaled image.</li>
	<li>Select Process/Filters/Gaussian blur (1.5 nm radius) and click OK (Figure 5B).</li>
	<li>Select Image/Adjust/Auto Local Threshold (using the Niblack method) and click OK (Figure 5C).</li>
	<li>Select Edit/Invert and then Process/Binary/Watershed (Figure 5D).</li>
	<li>Draw a border around the region of interest (ROI) with the Freehand Selection tool. Add the selection to the ROI manager by selecting Edit/Selection/Add to Manager or by clicking &#39;T&#39;.</li>
	<li>Select Edit/Clear Outside (Figure 5E).</li>
	<li>Select Analyze/Set Measurements, choose the appropriate parameters (e.g., area, fit ellipse).</li>
	<li>Select Analyze/Analyze Particles. Enter an appropriate range for the size of particles and mark &#39;Display results&#39;, &#39;Add to Manager&#39; and, if necessary, the &#39;Exclude on edges&#39; option and click OK. The result is shown in Figure 5F.</li>
	<li>Open the saved scaled imaged file (step 4.2). Select &#39;Show All&#39; and deselect &#39;Labels&#39; in the ROI Manager. Review the selected particles laid onto the original image and manually add or remove selections by using the &#39;Add&#39; or &#39;Delete&#39; buttons of the ROI Manager. Correct any selections necessary by using the Freehand Selection tool and the &#39;Update&#39; button of the ROI Manager (Figures 5G and 5H).</li>
	<li>Analyze the data using the obtained measurements. In the Results window, click Results/Summarize to obtain, for example, the mean Area, and mean Major and Minor axes of the particles listed in the ROI Manager. Click Results/Distribution, select a Parameter (such as Area) and click OK to view a histogram for this parameter.</li>
</ol>

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B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=High-resolution+cryoscanning+electron+microscopy+of+biological+samples.">High-resolution cryoscanning electron microscopy of biological samples.</a> Biological Low-Voltage Scanning Electron Microscopy. , 245-261 (2008).</li><li>Walther, P., M&#252;ller, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Double-layer+coating+for+field-emission+cryo-scanning+electron+microscopy--present+state+and+applications.">Double-layer coating for field-emission cryo-scanning electron microscopy--present state and applications.</a> Scanning. 19 (5), 343-348 (1997).</li><li>Walther, P., Wehrli, E., Hermann, R., M&#252;ller, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Double-layer+coating+for+high-resolution+low-temperature+scanning+electron+microscopy.">Double-layer coating for high-resolution low-temperature scanning electron microscopy.</a> J. Microsc. 179, 229-237 (1995).</li><li>Hess, M. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cryopreparation+methodology+for+plant+cell+biology.">Cryopreparation methodology for plant cell biology.</a> Cell. Electron Microsc. 79, 57-100 (2007).</li><li>Schertel, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cryo+FIB-SEM:+Volume+imaging+of+cellular+ultrastructure+in+native+frozen+specimens.">Cryo FIB-SEM: Volume imaging of cellular ultrastructure in native frozen specimens.</a> J. Struct. Biol. 184 (2), 355-360 (2013).</li><li>Schindelin, J., 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=Fiji:+an+open-source+platform+for+biological-image+analysis.">Fiji: an open-source platform for biological-image analysis.</a> Nat. Methods. 9 (7), 676-682 (2012).</li><li>Walther, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Recent+progress+in+freeze-fracturing+of+high-pressure+frozen+samples.">Recent progress in freeze-fracturing of high-pressure frozen samples.</a> J. Microsc. 212 (1), 34-43 (2003).</li><li>Nevo, R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Architecture+of+Thylakoid+Membrane+Networks.">Architecture of Thylakoid Membrane Networks.</a> Lipids Photosynth. 30, 295-328 (2009).</li></ol>

The authors declare they have no competing financial interests.

The technique described in this paper allows investigation of freeze-fractured membranes within the context of well-preserved high-pressure frozen plant tissues by cryo-scanning electron microscopy. The major advantage of using these procedures is that sample preparation is purely physical; no steps involving chemicals or dehydration are necessary. Thus, it allows studying biological structures at a near-native state26,32. The benefit of using leaf tissues is that one can obtain information on the overall cellular organization, as well as study specific membranes, such as the thylakoid membranes, within their native physiological context, i.e., within the chloroplast. Another advantage, imperative to studying C. pumilum plants at different relative water contents (RWC), is that their hydration state is not altered during preparation. This is as opposed to utilizing isolated chloroplasts or thylakoid membranes as the starting material for a freeze-fracture experiment. In the latter, one also has to take into account that some structural and/or supramolecular alterations likely take place during organelle/membrane isolation.
Working with plant tissue samples, which are relatively thick [&#62;50 - 100 &#181;m, and can be significantly thicker depending on the type of tissue and species], dictates the use of high-pressure freezing (HPF) for sample vitrification. Application of high pressure [2,100 bar], at which the melting temperature of water is about &#8722;22 &#176;C, affects the hydrogen bond network of the water such that it slows down the rate of ice crystal formation. Therefore, the chance of obtaining a vitrified sample is high even at relatively slow cooling rates. HPF is currently the only method available for vitrification of thick, yet not exceeding 200 &#181;m, samples. A critical step for HPF of plant tissues is infiltration of their intercellular air spaces with an inert &#34;space filler&#34; prior to freezing, to prevent tissue collapse under the high pressure (see reviews29,33). In the case of C. pumilum leaves, whose thickness can be up to 1 mm, the leaves also have to be trimmed or thinned prior to HPF.
Essentially, the technique of replica preparation18 can also be applied to freeze-fractured leaf tissues. However, in our experience, sufficient yield of intact fractured areas was not obtained, since replicas often fragment to small pieces. Moreover, searching for a specific region of interest such as chloroplasts can be impractical within fragmented replicas of plant tissues. This is mainly because mesophyll (photosynthetic) cells are comprised of a large central vacuole surrounded by a narrow cytoplasmic strip (Figure 2A), which includes the nucleus, chloroplasts and all other organelles. Thus, chloroplasts represent only a very small fraction of the total fractured area of mesophyll tissues. Cryo-SEM imaging of freeze-fractured samples offers a solution to this problem since samples are directly visualized following fracture (and coating/shadowing). However, this method suffers from the limitation that frozen-hydrated samples are beam-sensitive and thus scanning at high magnifications readily damages the sample on the first scan. Double-layer coating (DLC)27,28 offers a solution to this serious drawback.
In DLC, fractured samples are coated in a way similar to how they are for replica preparation. First, a thin layer (1 - 3 nm) of Pt/C is evaporated onto the sample, providing contrast and conductivity. This is followed by a thicker (5 - 10 nm), protective layer of carbon. DLC combined with a high accelerating voltage (10 kV) was used to image samples using the backscattered electron (BSE) signal28. The BSE signal, originating from the platinum layer allows obtaining surface information through the carbon and, possibly, water vapor contaminant layer, which are practically transparent to the BSE at high acceleration voltages (water contamination accumulates in experiments in which a vacuum-cryo transfer unit is not used and the fractured surface is exposed to the atmosphere, even if this is for a fraction of a second). In addition, imaging using the BSE signal minimized the problem of charging effects which are apparent in secondary electron (SE) detection27. With the microscope setup described here, imaging of the SE signal with the In-lens SE detector at 10 kV yielded information equivalent to that obtained when samples were coated solely with 2 - 3 nm of Pt/C and imaged at low accelerating voltages. The difference is that samples prepared by DLC and imaged as described were considerably less beam-sensitive. This enabled us to scan them at high magnifications and in many cases even repeatedly, and allowed obtaining valuable information on photosynthetic protein supramolecular organization and other cellular constituents (Figures 2 - 4)24.
Finally, the procedures described here can be modified to study the structure and/or membrane protein organization in other types of samples. We have successfully employed the technique to study the supramolecular organization of photosynthetic membranes in cyanobacterial and algal cells (Shperberg-Avni et al. unpublished data). The main consideration is finding the balance between having as thin a sample as possible, in order to increase the chance for vitrification, while keeping sample intactness. For different types of samples, which can be suspensions or animal/plant tissues, different packaging should be used (e.g., in terms of the type of platelets utilized for high-pressure freezing). Once successful cryo-immobilization is achieved, the combination of freeze-fracture, DLC and cryo-SEM imaging provides an excellent means for obtaining information on membrane protein organization at high magnification with minimal beam damage.

Oxygenic photosynthesis, originating in ancient cyanobacteria, was inherited by algae and land plants by endosymbiotic events that led to development of the chloroplast organelle. In all modern-day oxygenic phototrophs, photosynthetic electron transport and the generation of proton-motive force and reducing power are carried out within flattened sac-like vesicles termed &#39;thylakoid&#39; membranes. These membranes house the protein complexes that carry out the light-driven reactions of photosynthesis and provide a medium for energy transduction. The thylakoid membranes of plants and (some) algae are differentiated into two distinct morphological domains: tightly appressed membrane regions called &#39;grana&#39; and unstacked membranes that interconnect the grana, called &#39;stroma lamellae&#39;1. Various freeze-fracture studies of plant and algal thylakoid membranes have been conducted, starting in the early 1970s. When freeze-fractured, membranes split along their hydrophobic core2, generating an exoplasmic face (EF) and a protoplasmic face (PF), depending on the cellular compartment which the half-membrane borders, as originally coined by Branton et al. in 19753. Plant and algal thylakoids have four different fracture faces: EFs, EFu, PFs and PFu, with &#39;s&#39; and &#39;u&#39; denoting &#39;stacked&#39; and &#39;unstacked&#39; membrane regions, respectively. The membrane protein complexes, which are not split or broken, have the tendency to remain with either the E or P side of the membrane. The initial observations that the different fracture faces of the thylakoids contain particles of different sizes and densities4, and the numerous investigations that followed, led to identification and correlation between the observed particles and the membrane protein complexes that carry out the light reactions5-13 (see also reviews14,15).
Freeze-fracture experiments of thylakoid membranes are typically carried out on preparations of chloroplasts or isolated thylakoid membranes (but see16,17), at the risk of any alteration in structural and/or supramolecular organization that may occur during the isolation procedure. Following fracture, replicas are prepared by evaporation of platinum/carbon (Pt/C), then by a thick layer of carbon (C), and finally digestion of the biological material18. Replicas are visualized by transmission electron microscopy (TEM). The traditional freeze-fracture-replica technique continues to serve as an important tool for studying the supramolecular organization of photosynthetic membranes and their adaption to different, e.g., light, conditions19-23.
In our recent study of the homoiochlorophyllous resurrection plant Craterostigma pumilum24, we aimed to investigate the changes in the supramolecular organization of thylakoid membranes, as well as in overall cellular organization, during dehydration and rehydration. The uniqueness of homoiochlorophyllous resurrection species is that they are able to survive conditions of desiccation in their vegetative tissues (leaves), while retaining their photosynthetic apparatus. Once water is available, these plants recover and resume photosynthetic activity within hours to a few days25. For this study, cryo-scanning EM (SEM) imaging of freeze-fractured leaf samples was combined with high-pressure freezing for sample cryo-immobilization. These procedures provide a means to visualize frozen-hydrated biological samples at a state close to their native state26. One main benefit is that samples are examined directly after freeze-fracture and coating with no successive steps. This is particularly relevant to the investigation of plants at different relative water contents (RWC), as their hydration state is maintained during preparation. However, one critical disadvantage is that frozen-hydrated samples may suffer from beam damage during imaging, especially when scanned at high magnifications, required for accurate measurement of the size of photosynthetic complexes. To overcome this, a method called &#39;double-layer coating&#39; (DLC)27,28 combined with specific cryo-SEM imaging conditions were utilized. These resulted in samples that are significantly less beam-sensitive and allowed for the elucidation of valuable information on photosynthetic protein supramolecular organization and other cellular constituents of the resurrection plant C. pumilum at high magnifications in situ.

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			<td height="63" style="height:63px;width:312px;">0.1/0.2 platelets</td>
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			<td style="width:119px;">241</td>
			<td style="width:489px;">Platelets are of 3-mm diameter and 0.5-mm-thick (Type A) with 0.1/0.2-mm-deep cavities (of diamater 2 mm). Similar platelets can be obtained from Leica Microsystems.</td>
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			<td style="width:489px;">Dumont (Switzerland) Durostar style #5 tweezers; Can be substituted with other high-precision tweezers.</td>
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			<td height="63" style="height:63px;width:312px;">high-pressure freezing machine</td>
			<td style="width:359px;">Bal-Tec</td>
			<td style="width:119px;">HPM 010</td>
			<td style="width:489px;">High-pressure freezing alternatives: 1. HPF Compact 02, Wohlwend GmbH; 2. HPM 010, RMC Boeckeler; 3. EM PACT2, Leica Microsystems; 4. EM HPM 100, Leica Microsystems; 5. EM ICE, Leica Microsystems.</td>
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			<td height="21" style="height:21px;width:312px;">freeze-fracture system</td>
			<td style="width:359px;">Leica Microsystems</td>
			<td style="width:119px;">EM BAF 060</td>
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			<td height="21" style="height:21px;width:312px;">cryo preparation loading stage</td>
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			<td height="21" style="height:21px;width:312px;">specimen holder for univeral freeze fracturing</td>
			<td style="width:359px;">Leica Microsystems</td>
			<td style="width:119px;">16LZ04746VN</td>
			<td style="width:489px;">Clamp holder for specimen carriers of diameter 3 mm</td>
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			<td height="21" style="height:21px;width:312px;">vacuum cryo-transfer shuttle</td>
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			<td height="21" style="height:21px;width:312px;">scanning electron microscope</td>
			<td style="width:359px;">Zeiss</td>
			<td style="width:119px;">Ultra 055</td>
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			<td height="21" style="height:21px;width:312px;">cryo SEM stage</td>
			<td style="width:359px;">Leica Microsystems</td>
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			<td height="21" style="height:21px;width:312px;">image acquisiton software</td>
			<td style="width:359px;">SmartSEM, Carl Zeiss Microscopy GmbH</td>
			<td style="width:119px;"></td>
			<td></td>
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			<td height="21" style="height:21px;width:312px;">image analysis software</td>
			<td style="width:359px;">Fiji/Image J, National Institute of Health</td>
			<td style="width:119px;">http://fiji.sc/Fiji</td>
			<td></td>
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studying the supramolecular organization of photosynthetic membranes within freeze-fractured leaf tissues by cryo-scanning electron microscopy

Cryo-scanning electron microscopy (SEM) of freeze-fractured samples allows investigation of biological structures at near native conditions. Here, we describe a technique for studying the supramolecular organization of photosynthetic (thylakoid) membranes within leaf samples. This is achieved by high-pressure freezing of leaf tissues, freeze-fracturing, double-layer coating and finally cryo-SEM imaging. Use of the double-layer coating method allows acquiring high magnification (&#62;100,000X) images with minimal beam damage to the frozen-hydrated samples as well as minimal charging effects. Using the described procedures we investigated the alterations in supramolecular distribution of photosystem and light-harvesting antenna protein complexes that take place during dehydration of the resurrection plant Craterostigma pumilum, in situ.

Figure 1 shows cryo-SEM images of platelets containing high-pressure frozen, freeze-fractured Craterostigma pumilum leaf pieces. In some samples, large regions of fractured cells are obtained (Figure 1A). In others, the leaf piece stays tightly bound to the upper disc and is knocked off along with it (Figure 1B). However, even in the second case, some leaf tissue may remain attached to the knife grooves on the platelet (Figure 1B, arrowheads) and a fair amount of data can still be collected by imaging the leaf &#34;remnants&#34;, provided they were fractured. After successful fracture is found, low magnification images of cells are acquired to identify regions of interest, in this case chloroplasts (Figure 2).
Higher-magnification images of thylakoid membranes in C. pumilum leaves are shown in Figures 3 and 4. The four different fracture faces, EFs, EFu, PFs and PFu, can be distinguished (Figure 3). Photosystem II (PSII), which is the largest protein complex found in the membranes, is located in both the EFs (grana) and EFu (stroma lamellae). At hydrated conditions, PSII density is ~3 times higher in EFs than in EFu (~1,550 complexes &#181;m-2 vs. ~550 complexes &#181;m-2, Figure 3). This is one basis for differentiating between these two continuous faces. Another is the difference in their background. While the EFs background appears smooth, the EFu face is rough and contains holes that are the footprints of detached PSI complexes, which fracture to the complementary face, the PFu10 (Figure 3). An example for the segmentation of PSII complexes from the EFs face of the thylakoid membrane is shown in Figure 5.
During dehydration of C. pumilum, the density of PSII in the grana membranes (EFs) gradually decreases, reaching roughly half (~700 complexes &#181;m-2 at ~15% RWC, Figure 4C) of their density at hydrated conditions (100% RWC, Figure 4A). Notably, upon further dehydration, to 5 - 10% RWC, PSII complexes organize into rows and arrays (arrows, Figures 4D and 4E). For complete dataset, see24. Formation of such PSII arrays necessitates that some of their bound antenna complexes, LHCII, detach from PSII. An example for LHCII complexes organized into rows in the PFs that would be complementary to organized PSII complexes (in the EFs) is shown in Figure 4E (arrowhead). PSII complexes in the arrays, as well as the detached LHCII, are likely to be in a photochemically quenched state, serving a photo-protective role. These structural rearrangements are part of the mechanisms utilized by the resurrection plant C. pumilum to protect itself in the dehydrated state24.
<img alt="Figure 1" src="/files/ftp_upload/54066/54066fig1.jpg" /> 
Figure 1. Low Magnification Cryo-SEM Images of Platelets with Freeze-fractured C. pumilum Leaf Pieces. (A) A large region of fractured cells (dashed outline) of a C. pumilum leaf. (B) The leaf piece was knocked off with the top platelet, but some leaf tissue remained attached to the knife marks at the bottom platelet (arrowheads mark some of these). The regions surrounding the leaf pieces are frozen 1-hexadecene (asterisks). Scale bars: 200 &#181;m. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54066/54066fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" src="/files/ftp_upload/54066/54066fig2.jpg" /> 
Figure 2. Zooming into Chloroplasts. (A) A group of fractured photosynthetic (mesophyll) cells (asterisks) of hydrated leaf tissue. Most of the cell volume is comprised of a large central vacuole, with all cytoplasmic constituents surrounding the vacuole in a narrow strip. (B) Two neighboring fractured cells - the cell wall (cw) marks the border between the cells. Five chloroplasts (c) are apparent in the image, only one of which has been fractured (fractured plane marked by the arrowhead). (C and D) Examples for single fractured chloroplasts from hydrated (C) and dehydrated (D, ~15% relative water content [RWC]) plants. The small white dots (e.g., arrowheads) are the membrane protein complexes found within the thylakoid membranes of the chloroplast. Note the massive vesiculation surrounding the chloroplast found in the dehydrated leaf (D). Scale bars: 10 &#181;m (A); 1 &#181;m (B); 200 nm (C and D). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54066/54066fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" src="/files/ftp_upload/54066/54066fig3.jpg" /> 
Figure 3. The Supramolecular Organization of Thylakoid Membranes within Plant Tissues. A fractured chloroplast in which the different thylakoid fracture faces can be seen: the exoplasmic fracture faces (EF) of both stacked (EFs) and unstacked (EFu) membrane regions contain photosystem II (PSII) complexes; The protoplasmic fracture face of stacked regions (PFs) contains the peripheral light-harvesting antenna complexes of PSII, LHCII; Photosystem I (PSI) and ATP synthase fracture to the protoplasmic fracture face of unstacked regions (PFu); Cytochrome b6f fractures to both PFs and PFu. Image was scanned at a magnification of 203 kX (horizontal field of view = 1.45 &#181;m). Scale bar: 100 nm. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54066/54066fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" src="/files/ftp_upload/54066/54066fig4.jpg" /> 
Figure 4. Changes in Photosystem II Organization during Dehydration of C. pumilum. EFs faces of thylakoid membranes of plants at different RWC: (A) 100%, (B) ~40%, (C) ~15%, and (D and E) 5 - 10%. During dehydration, the density of PSII in the EFs face gradually decreases from ~1,500 complexes &#181;m-2 (A) to ~700 complexes &#181;m-2 (~15% RWC, C). Notably, at drier conditions (5 - 10% RWC), some PSII complexes organize into rows and arrays (D and E, arrows). The arrowhead (E) marks the PFs face, with LHCII antenna appearing to also be organized in rows, in between which PSII rows would be found in the complementary EFs. For additional information on this data see24. Scale bars: 100 nm. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54066/54066fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" src="/files/ftp_upload/54066/54066fig5.jpg" /> 
Figure 5. An Example for Segmentation of Photosystem II Particles. Using the Fiji open-source package, protein complexes can be segmented from the images with a few steps (complete detailed procedure found in part 4 of the Protocol section). Images (A, original image) are blurred with the Gaussian filter (B); the image is thresholded using an auto local threshold tool (C); the image is inverted and the watershed algorithm is applied in order to separate objects found in close contact (D); a region of interest is selected and the outside is cleared (E); particles (within a user-defined size range) are selected using the particle analyzer function. The resulting mask is shown in (F). Particles are added to the ROI Manager and overlaid onto the original image (G) to check for errors or for faulty or missed particles. The list of ROIs can be manually edited, replacing, deleting or adding new ROIs, as appropriate. The final user edited selection is shown in (H). Scale bar: 200 nm. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54066/54066fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

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Studying the Supramolecular Organization of Photosynthetic Membranes within Freeze-fractured Leaf Tissues by Cryo-scanning Electron Microscopy | Biology | Video

Studying the Supramolecular Organization of Photosynthetic Membranes within Freeze-fractured Leaf Tissues by Cryo-scanning Electron Microscopy

Here we describe a procedure for studying freeze-fractured plant tissues. High-pressure frozen leaf samples are freeze-fractured and double-layer coated, yielding well preserved frozen-hydrated samples that are imaged using the cryo-scanning electron microscope at high magnifications with minimal beam damage.

Cryo-scanning electron microscopy (SEM) of freeze-fractured samples allows investigation of biological structures at near native conditions. Here, we ...

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11.9K Views.  Weizmann Institute of Science. Here we describe a procedure for studying freeze-fractured plant tissues. High-pressure frozen leaf samples are freeze-fractured and double-layer coated, yielding well preserved frozen-hydrated samples that are imaged using the cryo-scanning electron microscope at high magnifications with minimal beam damage.

Video: Studying the Supramolecular Organization of Photosynthetic Membranes within Freeze-fractured Leaf Tissues by Cryo-scanning Electron Microscopy

Cryo-electron Microscopy Specimen Preparation By Means Of a Focused Ion Beam

Here we present a protocol used to prepare cryo-TEM samples of Aspergillus niger spores, but which can easily be adapted for any number of microorganisms or solutions. We make use of a custom built cryo-transfer station and a modified cryo-SEM preparation chamber2. The spores are taken from a culture, plunge-frozen in a liquid nitrogen slush and observed in the cryo-SEM to select a region of interest. A thin lamella is then extracted using the FIB, attached to a TEM grid and subsequently thinned to electron transparency. The grid is transferred to a cryo-TEM holder and into a TEM for high resolution studies. Thanks to the introduction of a cooled nanomanipulator tip and a cryo-transfer station, this protocol is a straightforward adaptation to cryogenic temperature of the routinely used FIB preparation of TEM samples. As such it has the advantages of requiring a small amount of modifications to existing instruments, setups and procedures; it is easy to implement; it has a broad range of applications, in principle the same as for cryo-TEM sample preparation. One limitation is that it requires skillful handling of the specimens at critical steps to avoid or minimize contaminations.

Cryo Electron Microscopes, either Scanning (SEM) or Transmission (TEM), are widely used for characterization of biological samples or other materials with a high water content1. A SEM/Focused Ion Beam (FIB) is used to identify features of interest in samples and extract a thin, electron-transparent lamella for transfer to a cryo-TEM.

Here we present a protocol used to prepare cryo-TEM samples of Aspergillus niger spores, but which can easily be adapted for any number of microorganisms ...

In Vivo 4-Dimensional Tracking of Hematopoietic Stem and Progenitor Cells in Adult Mouse Calvarial Bone Marrow

Through a delicate balance between quiescence and proliferation, self renewal and production of differentiated progeny, hematopoietic stem cells (HSCs) maintain the turnover of all mature blood cell lineages. The coordination of the complex signals leading to specific HSC fates relies upon the interaction between HSCs and the intricate bone marrow microenvironment, which is still poorly understood[1-2].
We describe how by combining a newly developed specimen holder for stable animal positioning with multi-step confocal and two-photon in vivo imaging techniques, it is possible to obtain high-resolution 3D stacks containing HSPCs and their surrounding niches and to monitor them over time through multi-point time-lapse imaging. High definition imaging allows detecting ex vivo labeled hematopoietic stem and progenitor cells (HSPCs) residing within the bone marrow. Moreover, multi-point time-lapse 3D imaging, obtained with faster acquisition settings, provides accurate information about HSPC movement and the reciprocal interactions between HSPCs and stroma cells.
Tracking of HSPCs in relation to GFP positive osteoblastic cells is shown as an exemplary application of this method. This technique can be utilized to track any appropriately labeled hematopoietic or stromal cell of interest within the mouse calvarium bone marrow space.

In Vivo 4-Dimensional Tracking of Hematopoietic Stem and Progenitor Cells in Adult Mouse Calvarial Bone Marrow

The nature of the interactions between hematopoietic stem and progenitor cells (HSPCs) and bone marrow niches is poorly understood. Custom hardware modifications and a multi-step acquisition protocol allow the use of two-photon and confocal microscopy to image ex vivo labeled HSPCs homed within bone marrow areas, tracking interactions and movement.

Through a delicate balance between quiescence and proliferation, self renewal and production of differentiated progeny, hematopoietic stem cells (HSCs) ...

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

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

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

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

Scanning Electron Microscopy of Macerated Tissue to Visualize the Extracellular Matrix

Fibrosis is a component of all forms of heart disease regardless of etiology, and while much progress has been made in the field of cardiac matrix biology, there are still major gaps related to how the matrix is formed, how physiological and pathological remodeling differ, and most importantly how matrix dynamics might be manipulated to promote healing and inhibit fibrosis. There is currently no treatment option for controlling, preventing, or reversing cardiac fibrosis. Part of the reason is likely the sheer complexity of cardiac scar formation, such as occurs after myocardial infarction to immediately replace dead or dying cardiomyocytes. The extracellular matrix itself participates in remodeling by activating resident cells and also by helping to guide infiltrating cells to the defunct lesion. The matrix is also a storage locker of sorts for matricellular proteins that are crucial to normal matrix turnover, as well as fibrotic signaling. The matrix has additionally been demonstrated to play an electromechanical role in cardiac tissue. Most techniques for assessing fibrosis are not qualitative in nature, but rather provide quantitative results that are useful for comparing two groups but that do not provide information related to the underlying matrix structure. Highlighted here is a technique for visualizing cardiac matrix ultrastructure. Scanning electron microscopy of decellularized heart tissue reveals striking differences in structure that might otherwise be missed using traditional quantitative research methods.

Shown here is a method for visualizing extracellular matrix ultrastructure in decellularized cardiac tissues.

Fibrosis is a component of all forms of heart disease regardless of etiology, and while much progress has been made in the field of cardiac matrix ...

Sample Preparation and Imaging of Exosomes by Transmission Electron Microscopy

Exosomes are nano-sized extracellular vesicles secreted by body fluids and are known to represent the characteristics of cells that secrete them. The contents and morphology of the secreted vesicles reflect cell behavior or physiological status, for example cell growth, migration, cleavage, and death. The exosomes&#39; role may depend highly on size, and the size of exosomes varies from 30 to 300 nm. The most widely used method for exosome imaging is negative staining, while other results are based on Cryo-Transmission Electron Microscopy, Scanning Electron Microscopy, and Atomic Force Microscopy. The typical exosome&#39;s morphology assessed through negative staining is a cup-shape, but further details are not yet clear. An exosome well-characterized through structural study is necessary particular in medical and pharmaceutical fields. Therefore, function-dependent morphology should be verified by electron microscopy techniques such as labeling a specific protein in the detailed structure of exosome. To observe detailed structure, ultrathin sectioned images and negative stained images of exosomes were compared. In this protocol, we suggest transmission electron microscopy for the imaging of exosomes including negative staining, whole mount immuno-staining, block preparation, thin section, and immuno-gold labelling.

This protocol describes the various techniques necessary for transmission electron microscopy including negative staining, ultrathin sectioning for detailed structure, and immuno-gold labelling to determine the positions of specific proteins in exosomes.

Exosomes are nano-sized extracellular vesicles secreted by body fluids and are known to represent the characteristics of cells that secrete them. The ...

Biological Sample Preparation by High-pressure Freezing, Microwave-assisted Contrast Enhancement, and Minimal Resin Embedding for Volume Imaging

The described sample preparation technique is designed to combine the best quality of ultrastructural preservation with the most suitable contrast for the imaging modality in a focused ion beam scanning electron microscope (FIB-SEM), which is used to obtain stacks of sequential images for 3D reconstruction and modelling. High-pressure freezing (HPF) allows close to native structural preservation, but the subsequent freeze substitution often does not provide sufficient contrast, especially for a bigger specimen, which is needed for high-quality imaging in the SEM required for 3D reconstruction. Therefore, in this protocol, after the freeze substitution, additional contrasting steps are carried out at room temperature. Although these steps are performed in a microwave, it is also possible to follow traditional bench processing, which requires longer incubation times.The subsequent embedding in minimal amounts of resin allows for faster and more precise targeting and preparation inside the FIB-SEM. This protocol is especially useful for samples that require preparation by high-pressure freezing for a reliable ultrastructural preservation but do not gain enough contrast during the freeze substitution for volume imaging using FIB-SEM. In combination with the minimal resin embedding, this protocol provides an efficient workflow for the acquisition of high-quality volume data.

Here we present a protocol for combining two sample processing techniques, high-pressure freezing and microwave-assisted sample processing, followed by minimal resin embedding for acquiring data with a focused ion beam scanning electron microscope (FIB-SEM). This is demonstrated using a mouse tibial nerve sample and Caenorhabditis elegans.

The described sample preparation technique is designed to combine the best quality of ultrastructural preservation with the most suitable contrast for the ...

The CryoAPEX Method for Electron Microscopy Analysis of Membrane Protein Localization Within Ultrastructurally-Preserved Cells

Key cellular events like signal transduction and membrane trafficking rely on proper protein location within cellular compartments. Understanding precise subcellular localization of proteins is thus important for answering many biological questions. The quest for a robust label to identify protein localization combined with adequate cellular preservation and staining has been historically challenging. Recent advances in electron microscopy (EM) imaging have led to the development of many methods and strategies to increase cellular preservation and label target proteins. A relatively new peroxidase-based genetic tag, APEX2, is a promising leader in cloneable EM-active tags. Sample preparation for transmission electron microscopy (TEM) has also advanced in recent years with the advent of cryofixation by high pressure freezing (HPF) and low-temperature dehydration and staining via freeze substitution (FS). HPF and FS provide excellent preservation of cellular ultrastructure for TEM imaging, second only to direct cryo-imaging of vitreous samples. Here we present a protocol for the cryoAPEX method, which combines the use of the APEX2 tag with HPF and FS. In this protocol, a protein of interest is tagged with APEX2, followed by chemical fixation and the peroxidase reaction. In place of traditional staining and alcohol dehydration at room temperature, the sample is cryofixed and undergoes dehydration and staining at low temperature via FS. Using cryoAPEX, not only can a protein of interest be identified within subcellular compartments, but also additional information can be resolved with respect to its topology within a structurally preserved membrane. We show that this method can provide high enough resolution to decipher protein distribution patterns within an organelle lumen, and to distinguish the compartmentalization of a protein within one organelle in close proximity to other unlabeled organelles. Further, cryoAPEX is procedurally straightforward and amenable to cells grown in tissue culture. It is no more technically challenging than typical cryofixation and freeze substitution methods. CryoAPEX is widely applicable for TEM analysis of any membrane protein that can be genetically tagged.

This protocol describes the cryoAPEX method, in which an APEX2-tagged membrane protein can be localized by transmission electron microscopy within optimally-preserved cell ultrastructure.

Key cellular events like signal transduction and membrane trafficking rely on proper protein location within cellular compartments. Understanding precise ...

Cryo-Electron Microscopic Grid Preparation for Time-Resolved Studies using a Novel Robotic System, Spotiton

The capture of short-lived molecular states triggered by the early encounter of two or more interacting particles continues to be an experimental challenge of great interest to the field of cryo-electron microscopy (cryo-EM). A few methodological strategies have been developed that support these &#34;time-resolved&#34; studies, one of which, Spotiton-a novel robotic system-combines the dispensing of picoliter-sized sample droplets with precise temporal and spatial control. The time-resolved Spotiton workflow offers a uniquely efficient approach to interrogate early structural rearrangements from minimal sample volume. Fired from independently controlled piezoelectric dispensers, two samples land and rapidly mix on a nanowire EM grid as it plunges toward the cryogen. Potentially hundreds of grids can be prepared in rapid succession from only a few microliters of a sample. Here, a detailed step-by-step protocol of the operation of the Spotiton system is presented with a focus on troubleshooting specific problems that arise during grid preparation.

The protocol presented here describes the use of Spotiton, a novel robotic system, to deliver two samples of interest onto a self-wicking, nanowire grid that mix for a minimum of 90 ms prior to vitrification in liquid cryogen.

The capture of short-lived molecular states triggered by the early encounter of two or more interacting particles continues to be an experimental ...

Strategies for Optimization of Cryogenic Electron Tomography Data Acquisition

Cryogenic electron tomography (cryoET) is a powerful method to study the 3D structure of biological samples in a close-to-native state. Current state-of-the-art cryoET combined with subtomogram averaging analysis enables the high-resolution structural determination of macromolecular complexes that are present in multiple copies within tomographic reconstructions. Tomographic experiments usually require a vast amount of tilt series to be acquired by means of high-end transmission electron microscopes with important operational running-costs. Although the throughput and reliability of automated data acquisition routines have constantly improved over the recent years, the process of selecting regions of interest at which a tilt series will be acquired cannot be easily automated and it still relies on the user's manual input. Therefore, the set-up of a large-scale data collection session is a time-consuming procedure that can considerably reduce the remaining microscope time available for tilt series acquisition. Here, the protocol describes the recently developed implementations based on the SerialEM package and the PyEM software that significantly improve the time-efficiency of grid screening and large-scale tilt series data collection. The presented protocol illustrates how to use SerialEM scripting functionalities to fully automate grid mapping, grid square mapping, and tilt series acquisition. Furthermore, the protocol describes how to use PyEM to select additional acquisition targets in off-line mode after automated data collection is initiated. To illustrate this protocol, its application in the context of high-end data collection of Sars-Cov-2 tilt series is described. The presented pipeline is particularly suited to maximizing the time-efficiency of tomography experiments that require a careful selection of acquisition targets and at the same time a large amount of tilt series to be collected.

The increasing demand for large-scale data collection in cryogenic electron tomography requires high-throughput image acquisition routines. Described here is a protocol that implements the recent developments of advanced acquisition strategies aimed at maximizing the time-efficiency and throughput of tomographic data collection.

Cryogenic electron tomography (cryoET) is a powerful method to study the 3D structure of biological samples in a close-to-native state. Current ...

Super-Resolution Microscopy of the Synaptonemal Complex Within the Caenorhabditis elegans Germline

During meiosis, homologous chromosomes must recognize and adhere to one another to allow for their correct segregation. One of the key events that secures the interaction of homologous chromosomes is the assembly of the synaptonemal complex (SC) in meiotic prophase I. Even though there is little sequence homology between protein components within the SC among different species, the general structure of the SC has been highly conserved during evolution. In electron micrographs, the SC appears as a tripartite, ladder-like structure composed of lateral elements or axes, transverse filaments, and a central element. 
However, precisely identifying the localization of individual components within the complex by electron microscopy to determine the molecular structure of the SC remains challenging. By contrast, fluorescence microscopy allows for the identification of individual protein components within the complex. However, since the SC is only ~100 nm wide, its substructure cannot be resolved by diffraction-limited conventional fluorescence microscopy. Thus, determining the molecular architecture of the SC requires super-resolution light microscopy techniques such as structured illumination microscopy (SIM), stimulated-emission depletion (STED) microscopy, or single-molecule localization microscopy (SMLM). 
To maintain the structure and interactions of individual components within the SC, it is important to observe the complex in an environment that is close to its native environment in the germ cells. Therefore, we demonstrate an immunohistochemistry and imaging protocol that enables the study of the substructure of the SC in intact, extruded Caenorhabditis elegans germline tissue with SMLM and STED microscopy. Directly fixing the tissue to the coverslip reduces the movement of the samples during imaging and minimizes aberrations in the sample to achieve the high resolution necessary to visualize the substructure of the SC in its biological context.

Super-Resolution Microscopy of the Synaptonemal Complex Within the Caenorhabditis elegans Germline

Super-resolution microscopy can provide a detailed insight into the organization of components within the synaptonemal complex in meiosis. Here, we demonstrate a protocol to resolve individual proteins of the Caenorhabditis elegans synaptonemal complex.

During meiosis, homologous chromosomes must recognize and adhere to one another to allow for their correct segregation. One of the key events that secures ...