Name: studying the supramolecular organization of photosynthetic membranes within freeze-fractured leaf tissues by cryo-scanning electron microscopy
Uploaded: 2016-06-23T14:00:00
Description: Watch this Scientific Journal Video about Studying the Supramolecular Organization of Photosynthetic Membranes within Freeze-fractured Leaf Tissues by Cryo-scanning Electron Microscopy at JoVE.com

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.
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	<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 p...

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	<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. 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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. 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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.

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			<td height="21" style="height:21px;width:312px;">ethanol abs</td>
			<td style="width:359px;">Bio-Lab</td>
			<td style="width:119px;">052505</td>
			<td style="width:489px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:312px;">isopropanol</td>
			<td style="width:359px;">Bio-Lab</td>
			<td style="width:119px;">162605</td>
			<td style="width:489px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:312px;">1-hexadecene</td>
			<td style="width:359px;">Sigma-Aldrich</td>
			<td style="width:119px;">H7009</td>
			<td></td>
		</tr>
		<tr height="63">
			<td height="63" style="height:63px;width:312px;">0.1/0.2 platelets</td>
			<td style="width:359px;">Engineering Office M. Wohlwend GmbH, Switzerland</td>
			<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>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:312px;">high-precision-grade tweezers</td>
			<td style="width:359px;">Electron Microscopy Sciences</td>
			<td style="width:119px;">72706-01</td>
			<td style="width:489px;">Dumont (Switzerland) Durostar style #5 tweezers; Can be substituted with other high-precision tweezers.</td>
		</tr>
		<tr height="63">
			<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>
		</tr>
		<tr height="21">
			<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>
			<td style="width:489px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:312px;">cryo preparation loading stage</td>
			<td style="width:359px;">Leica Microsystems</td>
			<td style="width:119px;">16770228</td>
			<td style="width:489px;"></td>
		</tr>
		<tr height="21">
			<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>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:312px;">vacuum cryo-transfer shuttle</td>
			<td style="width:359px;">Leica Microsystems</td>
			<td style="width:119px;">EM VCT 100</td>
			<td></td>
		</tr>
		<tr height="21">
			<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>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:312px;">cryo SEM stage</td>
			<td style="width:359px;">Leica Microsystems</td>
			<td style="width:119px;">16770299905</td>
			<td></td>
		</tr>
		<tr height="21">
			<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>
		</tr>
		<tr height="21">
			<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>
		</tr>
	</tbody>
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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 (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 ...

The overall goal of this technique is to study the supramolecular organization of photosynthetic membranes using cryo-scanning electron microscopy of freeze-fractured leaf tissue samples. This method can help answer key questions in cell biology, such as the morphological characterization of cells and organelles, the distribution of membrane-integral protein complexes, and localization of biominerals. The main advantage of this technique is that freeze-fracture is performed on leaf tissues, and so thylakoid membrane supramolecular organazation is investigated within the native organelle and cellular context.While this method is demonstrated here for a biological sample, it can also be applied to such as gels, polymers, and drug delivery systems. Generally, individuals new to these procedures should be aware that not all samples will be well-frozen or properly fractured. To carry out cryofixation of leaf tissues by high-pressure freezing, begin by using the corner of a razor blade to scratch the bottom of the 0.1-millimeter cavity of a 0.1-to-0.2-millimeter aluminum platelet.Prepare at least a dozen platelets for six samples. Then use absolute ethanol to wash the platelets, and keep them in a clean dish. After preparing the high-pressure freezing machine, fill the samples box with liquid nitrogen.With a razor blade, cut a small piece of leaf from the plant and use tweezers to place the tissue inside a microcentrifuge tube half filled with one hexadecane. To infiltrate the intercellular spaces of the leaf tissue, insert the tip of the infiltration device, prepared as described in the text protocol into the hole in the lid of the microcentrifuge tube, and pull the plunger to create vacuum. After about 30 seconds, release the plunger.Next, using tweezers, remove the leaf piece from the tube. Use the razor blade to cut a piece small enough to fit into a platelet. And if necessary, trim the leaf to a thickness of less than 0.2 millimeters.To prevent tissue collapse, leaves are infiltrated with liquid prior to high-pressure freezing. Sample thickness is kept below 200 microns in order to maximize the chance for obtaining a vitrified sample. Place a clean platelet with the scratch side up into the freezing machine's holder.And then place the cut piece of leaf into the platelet. Use one hexadecane to fill the remaining space of the platelet before using another platelet with the scratches facing the leaf to close the sandwich. Close and tighten the holder.Freeze the sample by pressing the jet button. Then, quickly transfer the holder into liquid nitrogen. Use precool tweezers to remove the sandwich from the holder, taking care not to fracture it.There are two critical steps in the procedure:freeze-fracture, which is a random process and its success can only be assessed in the cryo-SEM'and double-layer coating, which is required for obtaining high-magnification images with minimal beam damage. While preparing and programming the freeze-fracture machine according to the manufacturer's instructions, place the platinum carbon, or Pt/C gun, at 45 degrees, and the carbon gun at 90 degrees. Cool the freeze-fracture system stage to minus 140 or minus 160 degrees Celsius.Then attach the vacuum cryo transfer shuttle to the machine and use liquid nitrogen to fill the shuttle chamber. Insert a freeze-fracturing specimen holder into the loading station, and with liquid nitrogen flood the loading station chamber. Next, using high-precision-grade tweezers, load a frozen sandwich onto the holder and tighten it in place.Then flip the holder over to check that the sandwich is secured in place, and repeat for a second sandwich. Detach the cooled shuttle from the freeze-fracture machine and connect it to the loading station. Then open the shuttle valve, introduce the retracting arm into the holder, lock it in place, and reintroduce the arm with the holder into the shuttle.After reattaching the shuttle to the freeze-fracture machine introduce the holder into the chamber, using the retracting arm. Now align the freeze-fracture microtome knife to height such that it will knock off the top platelets of the sandwiches. With the microtome knife, quickly break the sandwiches.And then immediately raise up the knife to prevent contamination of the samples by debris clinging to the knife. Position the cooled shutter above the newly-exposed plane to shield the samples from possible contamination by water molecules in the chamber. Next, to coat the samples with platinum, press Setup to show Layer 1 on the screen.Then turn on the Pt/C gun by pressing the GUN 1 button followed by the ON button. Examine the evaporation rate, and once it reaches 0.02 to 0.04 nanometers per second, press the Measure button to monitor the thickness of the deposited layer while simultaneously exposing the samples. To coat with carbon, select GUN 2 followed by the ON button.Once the evaporation rate is approximately 0.1 nanometers per second, press the Measure button and at the same time move the shutter to expose the samples. When the coating is complete, warm the stage to minus 120 degrees Celsius. To prepare the scanning electron microscope for cryo work, from the main menu select Stage, Stage Initialize, and confirm.Under the Vacuum tab, click the Vent button to vent the microscope chamber. Then select Tools, Goto Panel, and open the Stage Points List panel. Select the CRYO EXCHANGE position to move the stage to the proper coordinates for accepting the sample holder.While wearing a clean pair of gloves, open the specimen chamber and insert the cryostage onto the main stage of the microscope. Close the chamber, and under the Vacuum tab, click the Pump button. Next, cool the microscope by filling the microscope duor with liquid nitrogen.Once the temperature of the stage drops below minus 120 degrees Celsius, activate the heat function on the cryo transfer unit to warm the stage to minus 120 degrees Celsius. Now attach the cryo transfer shuttle to the microscope and transfer the sample holder using the retracting arm into the microscope cryo stage. Using the joystick, carefully move the specimen holder close to the objective lens.In the Apertures tab, select the 10-micron aperture. Then, in the Gun tab, double-click the EHT Target field, enter 10 kilovolts for the EHT target, and click OK.Then, to turn on accelerating voltage, click the bottom EHT tab and select EHT On.In the Detectors tab, select InLens from the drop-down list. Then click on the Scanning tab and choose a scan speed equal to one, which is a fast scan to minimize beam damage, and a store resolution of 1024 by 768 pixels.In the Noise Reduction dropdown list, select Line Average, and adjust the value of n to 20 to 30 lines for searching. Then, by clicking the Control and Tab keys on the keyboard, use the centerpoint function to move the sample to areas of interest while simultaneously magnifying to identify regions with fractured cells. After identifying regions with fractured cells, scan the sample while magnifying to find a fractured chloroplast.Then change the line average n value to greater than 100, for noise reduction. In the Scanning tab, press Freeze to stop the scanning. By pressing Control Shift Tab on the keyboard, the center-feature function is used to zoom into interesting chloroplast-fracture faces;and high-magnification images are acquired using the same parameters as before.Finally, press the Save TIFF button to save the image. Carry out image analysis according to the text protocol. This figure shows a cryo-SEM image of a platelet containing a high-pressure frozen freeze-fractured Craterostigma Pumilum leaf piece with a large region of fractured cells.In this example, the leaf piece is missing;but some leaf tissue remained, attached to the knife grooves on the platelet. After a successful fracture is found, low-magnification images of cells are acquired to identify regions of interest, such as fractured chloroplasts. Images of single-fractured chloroplasts are acquired to identify thylakoid membrane fracture faces.Shown in this example is a high-magnification image of thylakoid membranes in C.Pumilum leaves. The four different fracture faces can be distinguished. The exoplasmic fracture faces of stacked and unstacked membranes, EFs and EFu respectively, and the protoplasmic faces of stacked and unstacked membranes, PFs and PFu respectively.Photosystem II, or PS2, is located in both the EFs stacked grana membrane regions and EFu stromalamellar membrane regions. When hydrated, PS2 density is approximately three times higher in the EFs than in the EFu, which differentiates the two faces. During dehydration of C.Pumilum, the density of PS2 in the EFs gradually decreases, reaching roughly half of its density at hydrated conditions.Notably, upon further dehydration to five to 10%relative water content, PS2 complexes organize into rows and arrays. Development of the freeze-fracture replica technique paved the way for studying the size of membrane protein complexes and their arrangement within biological membranes. The combination of freeze-fracture with high-pressure freezing in cryo-SEM allows investigating various kinds of samples at several size scales, from the tissue and cellular levels down to the macromolecular level at a near-native state.In addition to the technique shown here, methods such as Energy-Dispersive x-ray Spectroscopy, EDS, can also be applied in the cryo-SEM;for instance, for alkalizing biominerals.

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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 ...