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>

<ol>
	<li>Anderson, J. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Insights+into+the+consequences+of+grana+stacking+of+thylakoid+membranes+in+vascular+plants:+a+personal+perspective.">Insights into the consequences of grana stacking of thylakoid membranes in vascular plants: a personal perspective.</a> Aust. J. Plant Physiol. 26 (7), 625-639 (1999).</li><li>Branton, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fracture+Faces+of+Frozen+Membranes.">Fracture Faces of Frozen Membranes.</a> Proc. Natl. Acad. Sci. U. S. A. 55 (5), 1048-1056 (1966).</li><li>Branton, D., 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=Freeze-Etching+Nomenclature.">Freeze-Etching Nomenclature.</a> Science. 190 (4209), 54-56 (1975).</li><li>Goodenough, U. W., Staehelin, L. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Structural+Differentiation+of+Stacked+and+Unstacked+Chloroplast+Membranes+-+Freeze-Etch+Electron+Microscopy+of+Wild-Type+and+Mutant+Strains+of+Chlamydomonas.">Structural Differentiation of Stacked and Unstacked Chloroplast Membranes - Freeze-Etch Electron Microscopy of Wild-Type and Mutant Strains of Chlamydomonas.</a> J. Cell Biol. 48 (3), 594-619 (1971).</li><li>Simpson, D. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Freeze-Fracture+Studies+on+Barley+Plastid+Membranes+.6.+Location+of+the+P700-Chlorophyll+a-Protein-1.">Freeze-Fracture Studies on Barley Plastid Membranes .6. Location of the P700-Chlorophyll a-Protein-1.</a> Eur. J. Cell Biol. 31 (2), 305-314 (1983).</li><li>Staehelin, L. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Reversible+Particle+Movements+Associated+with+Unstacking+and+Restacking+of+Chloroplast+Membranes.">Reversible Particle Movements Associated with Unstacking and Restacking of Chloroplast Membranes.</a> Invitro. J. Cell Biol. 71 (1), 136-158 (1976).</li><li>Miller, K. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+Chloroplast+Membrane+Lacking+Photosystem-I+-+Changes+in+Unstacked+Membrane+Regions.">A Chloroplast Membrane Lacking Photosystem-I - Changes in Unstacked Membrane Regions.</a> Biochim. Biophys. Acta. 592 (1), 143-152 (1980).</li><li>Miller, K. R., Cushman, R. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Chloroplast+Membrane+Lacking+Photosystem-II+-+Thylakoid+Stacking+in+the+Absence+of+the+Photosystem-II+Particle.">Chloroplast Membrane Lacking Photosystem-II - Thylakoid Stacking in the Absence of the Photosystem-II Particle.</a> Biochim. Biophys. Acta. 546 (3), 481-497 (1979).</li><li>Miller, K. R., Staehelin, L. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Analysis+of+Thylakoid+Outer+Surface+-+Coupling+Factor+Is+Limited+to+Unstacked+Membrane+Regions.">Analysis of Thylakoid Outer Surface - Coupling Factor Is Limited to Unstacked Membrane Regions.</a> J. Cell Biol. 68 (1), 30-47 (1976).</li><li>Simpson, D. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Freeze-Fracture+Studies+on+Barley+Plastid+Membranes+.3.+Location+of+the+Light-Harvesting+Chlorophyll-Protein.">Freeze-Fracture Studies on Barley Plastid Membranes .3. Location of the Light-Harvesting Chlorophyll-Protein.</a> Carlsberg Res. Commun. 44 (5), 305-336 (1979).</li><li>Olive, J., Recouvreur, M., Girardbascou, J., Wollman, F. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Further+Identification+of+the+Exoplasmic+Face+Particles+on+the+Freeze-Fractured+Thylakoid+Membranes+-+a+Study+Using+Double+and+Triple+Mutants+from+Chlamydomonas-Reinhardtii+Lacking+Various+Photosystem-Ii+Subunits+and+the+Cytochrome+B6/F+Complex.">Further Identification of the Exoplasmic Face Particles on the Freeze-Fractured Thylakoid Membranes - a Study Using Double and Triple Mutants from Chlamydomonas-Reinhardtii Lacking Various Photosystem-Ii Subunits and the Cytochrome B6/F Complex.</a> Eur. J. Cell Biol. 59 (1), 176-186 (1992).</li><li>Olive, J., Vallon, O., Wollman, F. A., Recouvreur, M., Bennoun, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Studies+on+the+Cytochrome+B6/F+Complex+.2.+Localization+of+the+Complex+in+the+Thylakoid+Membranes+from+Spinach+and+Chlamydomonas-Reinhardtii+by+Immunocytochemistry+and+Freeze-Fracture+Analysis+of+B6/F+Mutants.">Studies on the Cytochrome B6/F Complex .2. Localization of the Complex in the Thylakoid Membranes from Spinach and Chlamydomonas-Reinhardtii by Immunocytochemistry and Freeze-Fracture Analysis of B6/F Mutants.</a> Biochim. Biophys. Acta. 851 (2), 239-248 (1986).</li><li>Armond, P. A., Staehelin, L. A., Arntzen, C. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Spatial+Relationship+between+Light+Harvesting+Complex+and+Photosystem-1+and+Photosystem-2+in+Stacked+and+Unstacked+Chloroplast+Membranes.">Spatial Relationship between Light Harvesting Complex and Photosystem-1 and Photosystem-2 in Stacked and Unstacked Chloroplast Membranes.</a> J. Cell Biol. 70 (2), 400-418 (1976).</li><li>Staehelin, L. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Chloroplast+structure:+from+chlorophyll+granules+to+supra-molecular+architecture+of+thylakoid+membranes.">Chloroplast structure: from chlorophyll granules to supra-molecular architecture of thylakoid membranes.</a> Photosynth. Res. 76 (1-3), 185-196 (2003).</li><li>Nevo, R., Charuvi, D., Tsabari, O., Reich, Z. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Composition,+architecture+and+dynamics+of+the+photosynthetic+apparatus+in+higher+plants.">Composition, architecture and dynamics of the photosynthetic apparatus in higher plants.</a> Plant J. 70 (1), 157-176 (2012).</li><li>Platt, K. A., Oliver, M. J., Thomson, W. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Membranes+and+Organelles+of+Dehydrated+Selaginella+and+Tortula+Retain+Their+Normal+Configuration+and+Structural+Integrity+-+Freeze-Fracture+Evidence.">Membranes and Organelles of Dehydrated Selaginella and Tortula Retain Their Normal Configuration and Structural Integrity - Freeze-Fracture Evidence.</a> Protoplasma. 178 (1-2), 57-65 (1994).</li><li>Platt-Aloia, K. A., Thomson, W. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Advantages+of+the+use+of+intact+plant+tissues+in+freeze-fracture+electron+microscopy.">Advantages of the use of intact plant tissues in freeze-fracture electron microscopy.</a> J. Electron Microsc. Tech. 13 (4), 288-299 (1989).</li><li>Carson, J. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fundamental+technical+elements+of+freeze-fracture/freeze-etch+in+biological+electron+microscopy.">Fundamental technical elements of freeze-fracture/freeze-etch in biological electron microscopy.</a> J. Vis. Exp. (91), e51694 (2014).</li><li>Kirchhoff, H., 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=Low-light-induced+formation+of+semicrystalline+photosystem+II+arrays+in+higher+plant+chloroplasts.">Low-light-induced formation of semicrystalline photosystem II arrays in higher plant chloroplasts.</a> Biochemistry. 46 (39), 11169-11176 (2007).</li><li>Johnson, M. 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=Photoprotective+Energy+Dissipation+Involves+the+Reorganization+of+Photosystem+II+Light-Harvesting+Complexes+in+the+Grana+Membranes+of+Spinach+Chloroplasts.">Photoprotective Energy Dissipation Involves the Reorganization of Photosystem II Light-Harvesting Complexes in the Grana Membranes of Spinach Chloroplasts.</a> Plant Cell. 23 (4), 1468-1479 (2011).</li><li>Kirchhoff, H., Tremmel, I., Haase, W., Kubitscheck, U. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Supramolecular+photosystem+II+organization+in+grana+thylakoid+membranes:+evidence+for+a+structured+arrangement.">Supramolecular photosystem II organization in grana thylakoid membranes: evidence for a structured arrangement.</a> Biochemistry. 43 (28), 9204-9213 (2004).</li><li>Belgio, E., Ungerer, P., Ruban, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=V+Light-harvesting+superstructures+of+green+plant+chloroplasts+lacking+photosystems.">V Light-harvesting superstructures of green plant chloroplasts lacking photosystems.</a> Plant Cell Environ. , (2015).</li><li>Goral, T. K., 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=Light-harvesting+antenna+composition+controls+the+macrostructure+and+dynamics+of+thylakoid+membranes+in+Arabidopsis.">Light-harvesting antenna composition controls the macrostructure and dynamics of thylakoid membranes in Arabidopsis.</a> Plant J. 69 (2), 289-301 (2012).</li><li>Charuvi, D., 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=Photoprotection+Conferred+by+Changes+in+Photosynthetic+Protein+Levels+and+Organization+during+Dehydration+of+a+Homoiochlorophyllous+Resurrection+Plant.">Photoprotection Conferred by Changes in Photosynthetic Protein Levels and Organization during Dehydration of a Homoiochlorophyllous Resurrection Plant.</a> Plant Physiol. 167 (4), 1554-1565 (2015).</li><li>Farrant, J. M., Brandt, W., Lindsey, G. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=An+Overview+of+Mechanisms+of+Desiccation+Tolerance+in+Selected+Angiosperm+Resurrection+Plants.">An Overview of Mechanisms of Desiccation Tolerance in Selected Angiosperm Resurrection Plants.</a> Plant Stress. 1 (1), 72-84 (2007).</li><li>Walther, P., Schatten, H., Pawley, J. 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 cell...

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.

<table><tbody><tr><td>Ethanol abs</td><td>Bio-Lab</td><td>052505</td><td></td></tr><tr><td>Isopropanol</td><td>Bio-Lab</td><td>162605</td><td></td></tr><tr><td>1-hexadecene</td><td>Sigma-Aldrich</td><td>H7009</td><td></td></tr><tr><td>0.1/0.2 Platelets</td><td>Engineering Office M. Wohlwend GmbH, Switzerland</td><td>241</td><td>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></tr><tr><td>High-precision-grade tweezers</td><td>Electron Microscopy Sciences</td><td>72706-01</td><td>Dumont (Switzerland) Durostar style #5 tweezers; Can be substituted with other high-precision tweezers.</td></tr><tr><td>High-pressure freezing machine</td><td>Bal-Tec</td><td>HPM 010</td><td>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></tr><tr><td>Freeze-fracture system</td><td>Leica Microsystems</td><td>EM BAF 060</td><td></td></tr><tr><td>Cryo preparation loading stage</td><td>Leica Microsystems</td><td>16770228</td><td></td></tr><tr><td>Specimen holder for univeral freeze fracturing</td><td>Leica Microsystems</td><td>16LZ04746VN</td><td>Clamp holder for specimen carriers of diameter 3 mm</td></tr><tr><td>Vacuum cryo-transfer shuttle</td><td>Leica Microsystems</td><td>EM VCT 100</td><td></td></tr><tr><td>Scanning electron microscope</td><td>Zeiss</td><td>Ultra 055</td><td></td></tr><tr><td>Cryo SEM stage</td><td>Leica Microsystems</td><td>16770299905</td><td></td></tr><tr><td>Image acquisiton software</td><td>SmartSEM, Carl Zeiss Microscopy GmbH</td><td></td><td></td></tr><tr><td>Image analysis software</td><td>Fiji/Image J, National Institute of Health</td><td>http://fiji.sc/Fiji</td><td></td></tr></tbody></table>

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 (F...

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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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12.1K 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

Cell Culture on Silicon Nitride Membranes and Cryopreparation for Synchrotron X-ray Fluorescence Nano-analysis

Very little is known about the distribution of metal ions at the subcellular level. However, those chemical elements have essential regulatory functions and their disturbed homeostasis is involved in various diseases. State-of-the-art synchrotron X-ray fluorescence nanoprobes provide the required sensitivity and spatial resolution to elucidate the two-dimensional (2D) and three-dimensional (3D) distribution and concentration of metals inside entire cells at the organelle level. This opens new exciting scientific fields of investigation on the role of metals in the physiopathology of the cell. The cellular preparation is a key and often complex procedure, particularly for basic analysis. Although X-ray fluorescence techniques are now widespread and various preparation methods have been used, very few studies have investigated the preservation of the elemental content of cells at best, and no stepwise detailed protocol for the cryopreparation of adherent cells for X-ray fluorescence nanoprobes has been released so far. This is a description of a protocol that provides the stepwise cellular preparation for fast cryofixation to enable synchrotron X-ray fluorescence nano-analysis of cells in a frozen hydrated state when a cryogenic environment and transfer is available. In case nano-analysis has to be performed at room temperature, an additional procedure for freeze-drying the cryofixed adherent cellular preparation is provided. The proposed protocols have been successfully used in previous works, most recently in studying the 2D and 3D intracellular distribution of an organometallic compound in breast cancer cells.

Presented here is a protocol for cell culture on silicon nitride membranes and plunge-freezing prior to X-ray fluorescence imaging with a synchrotron cryogenic X-ray nanoprobe. When only room temperature nano-analysis is provided, the frozen samples can be further freeze-dried. These are critical steps to obtain information on the intracellular elemental composition.

Very little is known about the distribution of metal ions at the subcellular level. However, those chemical elements have essential regulatory functions ...

Biochemistry

Preparing Lamellae from Vitreous Biological Samples Using a Dual-Beam Scanning Electron Microscope for Cryo-Electron Tomography

Presented here is a protocol for preparing cryo-lamellae from plunge-frozen grids of Plasmodium falciparum-infected&#160;human erythrocytes, which could easily be adapted for other biological samples. The basic principles for preparing samples, milling, and viewing lamellae are common to all instruments and the protocol can be followed as a general guide to on-grid cryo-lamella preparation for cryo-electron microscopy (cryoEM) and cryo-electron tomography (cryoET). Electron microscopy grids supporting the cells are plunge-frozen into liquid nitrogen-cooled liquid ethane using a manual or automated plunge freezer, then screened on a light microscope equipped with a cryo-stage. Frozen grids are transferred into a cryo-scanning electron microscope equipped with a focused ion beam (cryoFIB-SEM). Grids are routinely sputter coated prior to milling, which aids dispersal of charge build-up during milling. Alternatively, an e-beam rotary coater can be used to apply a layer of carbon-platinum to the grids, the exact thickness of which can be more precisely controlled. Once inside the cryoFIB-SEM an additional coating of an organoplatinum compound is applied to the surface of the grid via a gas injection system (GIS). This layer protects the front edge of the lamella as it is milled, the integrity of which is critical for achieving uniformly thin lamellae. Regions of interest are identified via SEM and milling is carried out in a step-wise fashion, reducing the current of the ion beam as the lamella reaches electron transparency, in order to avoid excessive heat generation. A grid with multiple lamellae is then transferred to a transmission electron microscope (TEM) under cryogenic conditions for tilt-series acquisition. A robust and contamination-free workflow for lamella preparation is an essential step for downstream techniques, including cellular cryoEM, cryoET, and sub-tomogram averaging. Development of these techniques, especially for lift-out and milling of high-pressure frozen samples, is of high-priority in the field.

Using focused ion beam milling to produce vitreous on-grid lamellae from plunge frozen biological samples for cryo-electron tomography.

Presented here is a protocol for preparing cryo-lamellae from plunge-frozen grids of Plasmodium falciparum-infected&#160;human erythrocytes, which could ...

Preparation and Cryo-FIB micromachining of Saccharomyces cerevisiae for Cryo-Electron Tomography

Today, cryo-electron tomography (cryo-ET) is the only technique that can provide near-atomic resolution structural data on macromolecular complexes in situ. Owing to the strong interaction of an electron with the matter, high-resolution cryo-ET studies are limited to specimens with a thickness of less than 200 nm, which restricts the applicability of cryo-ET only to the peripheral regions of a cell. A complex workflow that comprises the preparation of thin cellular cross-sections by cryo-focused ion beam micromachining (cryo-FIBM) was introduced during the last decade to enable the acquisition of cryo-ET data from the interior of larger cells. We present a protocol for the preparation of cellular lamellae from a sample vitrified by plunge freezing utilizing Saccharomyces cerevisiae as a prototypical example of a eukaryotic cell with wide utilization in cellular and molecular biology research. We describe protocols for vitrification of S. cerevisiae into isolated patches of a few cells or a continuous monolayer of the cells on a TEM grid and provide a protocol for lamella preparation by cryo-FIB for these two samples.

Preparation and Cryo-FIB micromachining of Saccharomyces cerevisiae for Cryo-Electron Tomography

We present a protocol for lamella preparation of plunge frozen biological specimens by cryo-focused ion beam micromachining for high-resolution structural studies of macromolecules in situ with cryo-electron tomography. The presented protocol provides guidelines for the preparation of high-quality lamellae with high reproducibility for structural characterization of macromolecules inside the&#160;Saccharomyces cerevisiae.

Today, cryo-electron tomography (cryo-ET) is the only technique that can provide near-atomic resolution structural data on macromolecular complexes in ...

Visualization of ATP Synthase Dimers in Mitochondria by Electron Cryo-tomography

Electron cryo-tomography is a powerful tool in structural biology, capable of visualizing the three-dimensional structure of biological samples, such as cells, organelles, membrane vesicles, or viruses at molecular detail. To achieve this, the aqueous sample is rapidly vitrified in liquid ethane, which preserves it in a close-to-native, frozen-hydrated state. In the electron microscope, tilt series are recorded at liquid nitrogen temperature, from which 3D tomograms are reconstructed. The signal-to-noise ratio of the tomographic volume is inherently low. Recognizable, recurring features are enhanced by subtomogram averaging, by which individual subvolumes are cut out, aligned and averaged to reduce noise. In this way, 3D maps with a resolution of 2 nm or better can be obtained. A fit of available high-resolution structures to the 3D volume then produces atomic models of protein complexes in their native environment. Here we show how we use electron cryo-tomography to study the in situ organization of large membrane protein complexes in mitochondria. We find that ATP synthases are organized in rows of dimers along highly curved apices of the inner membrane cristae, whereas complex I is randomly distributed in the membrane regions on either side of the rows. By subtomogram averaging we obtained a structure of the mitochondrial ATP synthase dimer within the cristae membrane.

We present a protocol of how to collect and process electron cryo-tomograms of whole mitochondria. The technique provides detailed insights into the structure, function, and organization of large membrane protein complexes in native biological membranes.

Electron cryo-tomography is a powerful tool in structural biology, capable of visualizing the three-dimensional structure of biological samples, such as ...

Fundamental Technical Elements of Freeze-fracture/Freeze-etch in Biological Electron Microscopy

Freeze-fracture/freeze-etch describes a process whereby specimens, typically biological or nanomaterial in nature, are frozen, fractured, and replicated to generate a carbon/platinum &#8220;cast&#8221; intended for examination by transmission electron microscopy. Specimens are subjected to ultrarapid freezing rates, often in the presence of cryoprotective agents to limit ice crystal formation, with subsequent fracturing of the specimen at liquid nitrogen cooled temperatures under high vacuum. The resultant fractured surface is replicated and stabilized by evaporation of carbon and platinum from an angle that confers surface three-dimensional detail to the cast. This technique has proved particularly enlightening for the investigation of cell membranes and their specializations and has contributed considerably to the understanding of cellular form to related cell function. In this report, we survey the instrument requirements and technical protocol for performing freeze-fracture, the associated nomenclature and characteristics of fracture planes, variations on the conventional procedure, and criteria for interpretation of freeze-fracture images. This technique has been widely used for ultrastructural investigation in many areas of cell biology and holds promise as an emerging imaging technique for molecular, nanotechnology, and materials science studies.

Basic techniques and refinements of freeze-fracture processing of biological specimens and nanomaterials for examination by transmission electron microscopy are described. This technique is a preferred method for revealing ultrastructural features and specializations of biological membranes and for obtaining ultrastructural level dimensional and spatial data in materials sciences and nanotechnology products.

Freeze-fracture/freeze-etch describes a process whereby specimens, typically biological or nanomaterial in nature, are frozen, fractured, and replicated ...

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, ...

Plunge Freezing: A Tool for the Ultrastructural and Immunolocalization Studies of Suspension Cells in Transmission Electron Microscopy

Transmission Electron Microscopy (TEM) is an extraordinary tool for studying cell ultrastructure, in order to localize proteins and visualize macromolecular complexes at very high resolution. However, to get as close as possible to the native state, perfect sample preservation is required. Conventional electron microscopy (EM) fixation with aldehydes, for instance, does not provide good ultrastructural preservation. The slow penetration of fixatives induces cell reorganization and loss of various cell components. Therefore, conventional EM fixation does not allow for an instantaneous stabilization and preservation of structures and antigenicity. The best choice for examining intracellular events is to use cryofixation followed by the freeze-substitution fixation method that keeps cells in their native state. High-pressure freezing/freeze-substitution, which preserves the integrity of cellular ultrastructure, is the most commonly used method, but requires expensive equipment. Here, an easy-to-use and low-cost freeze fixation method followed by freeze-substitution for suspension cell cultures is presented.

This manuscript describes an easy-to-use and low-cost cryofixation method for visualizing suspension cells by transmission electron microscopy.

Transmission Electron Microscopy (TEM) is an extraordinary tool for studying cell ultrastructure, in order to localize proteins and visualize ...

Flash-and-Freeze: A Novel Technique to Capture Membrane Dynamics with Electron Microscopy

Cells constantly change their membrane architecture and protein distribution, but it is extremely difficult to visualize these events at a temporal and spatial resolution on the order of ms and nm, respectively. We have developed a time-resolved electron microscopy technique, &#34;flash-and-freeze,&#34; that induces cellular events with optogenetics and visualizes the resulting membrane dynamics by freezing cells at defined time points after stimulation. To demonstrate this technique, we expressed channelrhodopsin, a light-sensitive cation channel, in mouse hippocampal neurons. A flash of light stimulates neuronal activity and induces neurotransmitter release from synaptic terminals through the fusion of synaptic vesicles. The optogenetic stimulation of neurons is coupled with high-pressure freezing to follow morphological changes during synaptic transmission. Using a commercial instrument, we captured the fusion of synaptic vesicles and the recovery of the synaptic vesicle membrane. To visualize the sequence of events, large datasets were generated and analyzed blindly, since morphological changes were followed in different cells over time. Nevertheless, flash-and-freeze allows the visualization of membrane dynamics in electron micrographs with ms temporal resolution.

We developed a novel technique in electron microscopy, &#34;flash-and-freeze,&#34; that enables the visualization of membrane dynamics with ms temporal resolution. This technique combines the optogenetic stimulation of neurons with high-pressure freezing. Here, we demonstrate the procedures and describe the protocols in detail.

Cells constantly change their membrane architecture and protein distribution, but it is extremely difficult to visualize these events at a temporal and ...

Visualization of DNA Compaction in Cyanobacteria by High-voltage Cryo-electron Tomography

This protocol describes how to visualize the transient DNA compaction in cyanobacteria. DNA compaction is a dramatic cytoplasmic event recently found to occur in some cyanobacteria before cell division. However, due to the large cell size and the transient character, it is difficult to investigate the structure in detail. To overcome the difficulties, first, DNA compaction is reproducibly produced in the cyanobacterium Synechococcus elongatus PCC 7942 by synchronous culture using 12 h each light/dark cycle. Second, DNA compaction is monitored by fluorescence microscopy and captured by rapid freezing. Third, the detailed structure of DNA compacted cells is visualized in three dimensions (3D) by high voltage cryo-electron tomography. This set of methods is widely applicable to investigate transient structures in bacteria, e.g. cell division, chromosome segregation, phage infection etc., which are monitored by fluorescence microscopy and directly visualized by cryo-electron tomography at appropriate time points.

This protocol describes how to visualize the transient DNA compaction in cyanobacteria. Synchronous cultivation, monitoring by fluorescence microscopy, rapid freezing, and high voltage cryo-electron tomography are used. A protocol for these methodologies is presented, and future applications and developments are discussed.

This protocol describes how to visualize the transient DNA compaction in cyanobacteria. DNA compaction is a dramatic cytoplasmic event recently found to ...

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 ...

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

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Chapters in this video

Cell Culture on Silicon Nitride Membranes and Cryopreparation for Synchrotron X-ray Fluorescence Nano-analysis

Preparing Lamellae from Vitreous Biological Samples Using a Dual-Beam Scanning Electron Microscope for Cryo-Electron Tomography

Preparation and Cryo-FIB micromachining of Saccharomyces cerevisiae for Cryo-Electron Tomography

Visualization of ATP Synthase Dimers in Mitochondria by Electron Cryo-tomography

Fundamental Technical Elements of Freeze-fracture/Freeze-etch in Biological Electron Microscopy

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

Plunge Freezing: A Tool for the Ultrastructural and Immunolocalization Studies of Suspension Cells in Transmission Electron Microscopy

Flash-and-Freeze: A Novel Technique to Capture Membrane Dynamics with Electron Microscopy

Visualization of DNA Compaction in Cyanobacteria by High-voltage Cryo-electron Tomography

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