Name: preparing lamellae from vitreous biological samples using a dual-beam scanning electron microscope for cryo-electron tomography
Uploaded: 2021-08-05T13:30:00.000+00:00
Duration: 7 min
Description: Watch this Scientific Journal Video about Preparing Lamellae from Vitreous Biological Samples Using a Dual-Beam Scanning Electron Microscope for Cryo-Electron Tomography at JoVE.com

This research was funded in whole, or in part, by the Wellcome Trust (212916/Z/18/Z). For the purpose of open access, the author has applied a CC BY public copyright licence to any Author Accepted Manuscript version arising from this submission.
This project for which this method was developed was funded by a Medical Research Council grant MR/P010288/1 awarded to Helen R. Saibil, Roland A. Fleck,&#160;and Michael J. Blackman. Cultures of P. falciparum were grown at The Francis Crick Institute, with the support of members of Michael J. Blackman&#39;s group. The authors would like to thank Dr. Ser Ying (Michele) Tan for providing the images of compound 2 and E64-treated schizonts in thin blood smears. Most of the lamellae were produced with support from staff at eBIC and we are grateful for access to the Scios dual-beam cryoFIB-SEM on research proposal NT21004. The authors also acknowledge the support of the Royal Society Industry Fellowship scheme (INF\R2\202061) in continuing the development of the FIB-milling technique within the CUI. The authors would also like to thank Helen R. Saibil for useful discussions in relation to this methods paper and supervising the project.

Human blood was obtained from anonymized donors through the UK National Blood and Transplant service and is used within 2 weeks of receipt. No ethical approval is required for its use.
1. Preparation and plunge freezing of&#160; Plasmodium falciparum&#160; infected red blood cells
<ol>
	<li>Isolate mature schizonts by centrifugation (1,580 x g) over a 70% (v/v) isotonic density gradient medium (for standard procedures for how to culture asexual blood stages of 3D7 Plasmodium falciparum in human erythrocytes see Blackman M.J., 199537).</li>
	<li>Fix the air-dried thin blood films on a glass slide with 100% methanol to check the morphological homogeneity of the schizonts before staining with 10% Giemsa stain in 6.7 mM phosphate buffer, pH 7.1. 
	NOTE: To enrich the preparation for schizonts stalled at specific points of egress, schizonts can be further synchronized with compound 2 and E64 inhibitors (see Representative Results and Figure 1 for an explanation of the effect of these inhibitors). 
	CAUTION: Plasmodium falciparum is a human pathogen and must only be handled in a suitable containment facility following local health and safety guidelines.</li>
	<li>Gently centrifuge schizonts (240 x g) to pellet them and resuspend in 2x volume of the cell pellet of RPMI media, resulting in a 50% haematocrit suspension.</li>
	<li>On a manual plunge-freezing rig, apply 2.5 &#956;L of schizonts to the carbon side of a glow-discharged (Use glow-discharge unit settings of 60 s, 30 mA in air, treating the carbon side of the grid only) 200 mesh copper grid with a 2/4 holey carbon film and blot for ~20 s from the back of the grid using grade 1 filter paper with a torn edge. Plunge into liquid nitrogen-cooled liquid ethane and transfer the grids to storage (see&#160;Table of Materials&#160;for equipment used in this study). 
	NOTE: The experiment can be paused here, and grids can be stored under liquid nitrogen indefinitely. 
	CAUTION: Liquid nitrogen is an asphyxiant and causes frostbite; handle with care in a suitable environment with oxygen monitoring. 
	CAUTION: Liquid ethane causes severe burns and is flammable; use in a fume hood away from sources of ignition. 
	NOTE: Plunge frozen schizonts are no longer viable. This was determined by incubating human blood with several gold grids of air-thawed plunge-frozen schizonts and observing no parasite growth after several days, in comparison to un-frozen controls. Frozen grids of schizonts are therefore safe to handle outside of containment facilities using normal safety and decontamination procedures (gloves, surface/tool sterilization with &#62;70% ethanol and disposal of grids in &#62;70% ethanol).</li>
	<li>Screen grids using a cryo-stage for a light microscope, paying particular attention to the gradient of ice across the grid. In the thinner areas of ice, check individual grid squares for cell coverage. 
	&#8203;NOTE: The best squares should be one cell thick at the center of the grid square (Figure 2). This ensures that a lamella can be milled across the center of the square without hitting the thicker ice at the edges against the grid bars. The experiment can be paused here, and grids can be stored under liquid nitrogen indefinitely. If cells carry a fluorescent marker, grids can also be screened by cryoCLEM to locate X/Y positions of interest, which can be correlated with grid locations in the cryoFIB-SEM to direct milling.</li>
</ol>
2. On-grid FIB-milling of plunge frozen cells
<ol>
	<li>Mark the front of cryoFIB-specific autogrid rims with a black indelible marker pen to indicate the center of the cutaway section and the opposite side of the rim (Figure 3). Setup the clipping station and clip grids into the marked rings carbon side down.</li>
	<li>Load grids into the cryoFIB-SEM shuttle (typically 2 grids, depending on the shuttle) carbon side up and apply a platinum sputter coat in an argon atmosphere (5 x 10-2 mbar) (5 mA for 60 s - thickness is variable) or a carbon/platinum e-beam rotary coat (~4 nm thickness) to the surface of the cells. 
	NOTE: Both types of coatings aid charge dispersal during SEM imaging. The benefit of the e-beam rotary coater is that the exact thickness of the coating can be specified.</li>
	<li>Load the shuttle into the cryoFIB-SEM and assess the cell distribution on each grid by SEM at 5 kV (13 pA or 25 pA). Take a low-magnification overview (~100x) to look at ice gradients across the grid. Then, take higher-magnification (~5,000x) images to look at individual grid squares and identify the areas of the grid with visible cellular features and low surface contamination to mill.</li>
	<li>Apply a &#62;2 &#181;m organoplatinum coat to the surface of each grid using the gas injection system (GIS). To do this, insert the GIS needle into the chamber above the grid and warm the organoplatinum source to a set temperature for a set time (for example, ~27 &#176;C for 3-10 s) to produce a flow of vapor. 
	NOTE: The angle of application, temperature, and timing of the organoplatinum coat via the GIS needle should be optimized to maximize an even coating. This will depend on the specific cryoFIB-SEM used (see Representative Results for further explanation).</li>
	<li>Tilt the sample so that the plane of the grid is ~10&#176; from the incidence angle of the ion beam and move the center of a suitable grid square to be seen in both the SEM and FIB images.</li>
	<li>Survey the grid using the ion beam at low current (nominally 1.5 pA, 30 kV) and go to high enough magnification (~7,000x) to visualize the cells at the center of a grid square. Bring the sample into focus at the first milling current (300 pA) and correct for astigmatism. Adjust the brightness and the contrast, then mark out two rectangular patterns to mill, one above and one below a 3 &#181;m thick protected region, the center of which is the desired location of the final lamella. 
	NOTE: The chosen width of the patterns will depend on the topography of the surrounding cell layer and the cell size. 7-20 &#181;m is a suitable width for schizonts, but wider lamellae take longer to mill. The chosen height of the patterns for the first milling step depends on the sample thickness, starting around 6 &#181;m; this may need to be adjusted during milling. Milling is carried out directionally from the outside top and bottom edges of the patterns toward the face of the lamella. This can be done in parallel, where both patterns are milled concurrently or sequentially, first removing material from above the lamella and then from underneath.</li>
	<li>Start milling at the first current; monitor the live progress in the ion beam view and intermittently by SEM (5 kV, 13 or 25 pA). Check that the ion beam has broken through the sample above and below the protected region. If not, increase the height of the rectangular patterns to remove more material. Stop when the surface above and below the lamella is completely smooth in the ion beam view. 
	NOTE: The ion beam has broken through the sample above and below the protected region when the inside of the rectangular patterns contains no features in the focal plane. Some features, such as grid bars, may be visible out of focus in the background.</li>
	<li>Change to the next milling current (100 pA); focus and adjust the brightness/contrast. Decrease the space between the two rectangular patterns to 1.5 &#181;m and decrease the pattern height to only cover the un-milled material. Begin milling at the new current until the surface above and below the lamella is completely smooth.</li>
	<li>Repeat this process stepwise until a thickness of 0.3 &#181;m is reached, reducing the ion beam current each time according to the milling scheme in Table 1.</li>
	<li>Mill several lamellae (the amount is dependent on the thickness of the sample and the time available) on one or both grids to a thickness of 0.3 &#181;m, record the X/Y/Z position of each lamella and reserve ~1-2 h at the end of the session to polish the lamellae to their final thickness (60-200 nm).</li>
	<li>Take a low-magnification SEM overview of the entire grid and plan a polishing route starting from lamellae at the front of the grid (closer to the ion beam source) to lamellae at the back of the grid (furthest away from the ion beam source) (Figure 5). 
	NOTE: This reduces redeposition of ablated material back onto the surface of polished lamellae.</li>
	<li>Reduce the space between the two rectangular patterns to 100-200 nm and begin the final polishing step at an ion beam current of 30 pA. Monitor the progress by SEM at 2-3 kV (13 pA, dwell = 300 n, 3072 x 2048, ~2 s for a full frame) and stop polishing when the contrast is lost in the lamella by SEM or when the organoplatinum coat of the lamella itself begins to lose integrity.</li>
	<li>Before removing the grids, acquire a low-magnification SEM image of the entire grid and save images of each lamella. Use them to cross-check the grid later in the TEM. Pause the experiment here and store the grids with lamellae under liquid nitrogen - handle with care. 
	&#8203;NOTE: Grids can be lightly sputter coated upon removal from the cryoFIB-SEM, which can help limit drift and charging in the TEM at high-magnification, but this should be done cautiously as too much sputter coating can obscure the biological contents inside the lamellae. It is possible to screen lamellae for fluorescence at this stage; however, obtaining enough signal will depend on the abundance of the labeled protein within the thickness of the lamella. Great care must be taken handling the grids to limit damage to the lamellae and prevent surface contamination.</li>
</ol>
3. Tilt-series acquisition and general overview of data processing
<ol>
	<li>Load the grids into the TEM, aligning the milling direction perpendicular to the tilt axis of the stage. 
	NOTE: Alignment of the grids is done by eye using the marks on the autogrid rim. A ~10&#176; margin of error is acceptable, otherwise the walls of the trench either side of the lamella may obscure the lamella as the grid is tilted.</li>
	<li>Acquire a low-magnification map (~150x) of the entire grid and locate the lamellae; then, acquire a medium-magnification map (~1,500x, depending on lamella size) of each lamella and locate the areas of interest.</li>
	<li>Pre-tilt the grid by &#177;10&#176;&#160;to make the plane of the lamella (not the grid) perpendicular to the optical axis 
	NOTE: The direction of the pre-tilt can be determined by the position of the front edge of the lamellae (looking for the left over organoplatinum coat) in the low and medium magnification maps. For the TEM used here, grids require&#160;a +10&#176; pre-tilt if the front edges of the lamellae point&#160;up in the maps and require&#160;a -10&#176; pre-tilt if they point&#160;down. Each grid may be different due to how they were picked up and inserted into the autoloader.</li>
	<li>Acquire dose-symmetric tilt-series38 (for example, -54&#176; to +54&#176;&#160;with an increment of 3-5&#176;) at a pixel size that permits both the field of view and resolution required for the region of interest. Use a range of defocus values between -2 and -5 &#181;m. Collect movies with 3-10 frames at each increment and for this, adjust these parameters depending on the pixel size to accumulate a total dose of ~150 e-/&#197;2 (for a 300 kV TEM). 
	NOTE: Cracks in the lamella should be avoided as these regions may drift. Surface contamination should also be avoided as it may obscure the region of interest or the focus area at high-tilt.</li>
	<li>Motion correct the movies using a program such as MotionCor239. Apply a dose weighting filter to the corrected images (accumulated e-/image)40 and estimate the defocus of each image using a program such as CTFFIND441.</li>
	<li>Use fiducial-less alignment (patch tracking) in a program such at etomo (IMOD)42 to calculate the alignment and angular relationship of the images in tilt-series.</li>
	<li>Input the alignment and rotation information, along with the defocus values into a program that can apply three dimensional CTF correction, for example, NovaCTF43. Calculate a corrected tomogram to get output tomograms with binning factors relevant to downstream analysis.</li>
	<li>Analyze the reconstructions and prepare for any downstream processing, for example, filtering, segmentation, or sub-tomogram averaging.</li>
</ol>

The authors have nothing to disclose.

While cryoFIB-milling is becoming more routine, preparation of optimal samples for milling has not; therefore, most of the critical steps in this protocol occur before the sample reaches the cryoFIB-SEM. Sample optimization by multiple rounds of cell preparation, plunge freezing, and screening by light and electron microscopy are required to produce the best possible sample before milling is attempted. Having the best sample possible not only increases the chances of success but also optimizes the use of the equipment. For this reason, most national facilities require evidence that sufficient optimization has been carried out before granting time on their machines. Once inside the cryoFIB-SEM, maximizing the effectiveness of the organoplatinum coat is critical in order to produce uniformly thin lamellae. Finally, patience and good sample handling is a prerequisite skill from the user, who will be required to sit for multiple days producing and then transferring grids containing delicate lamellae. This may change with the introduction of automated milling strategies47,48, but as yet, the entire milling process is still largely manual at most facilities.
While the work focused solely on P. falciparum schizonts, the method presented here could be easily modified to optimize grid preparation and milling for other cell types. Important factors to consider are the density of the cells being applied to the grid, the grid type (copper is toxic to some cells), the size and spacing of holes in the carbon film, the blotting time, and the method of blotting (single/double sided, manual, or automated). Additionally, if the cells are being grown on the grids (usually gold grids with a holey carbon film), confluency can be checked by light microscopy before blotting and freezing. The milling strategy depends on how the cells are deposited on the grid. For malaria infected red blood cells, the grid is essentially coated by an unbroken layer of cells, thicker in some areas and thinner in other areas. Milling is carried out at multiple points at one region along this gradient where the ice thickness generates lamellae that are a suitable size for tomography. This approach works well for smaller cells as you can produce lamellae containing a slice through multiple cells. For larger cells or clumps of cells, it might be the case that one cell or cell clump may produce one lamella.
Grid handling and sample transfer remain one of the main challenges of this workflow. Lamellae can be lost due to breakages or cracks, curtaining artefacts obscuring the biology, excessive surface contamination, as well as grid orientation problems within the TEM, requiring an extra handling step to reposition the grid. The degree of losses varies from day-to-day and grid-to-grid, but it is improved through practice and handling experience over time. In this study, it was found that the grids can be carefully reorientated in the autoloader multiple times without destroying lamellae and this sometimes has the benefit of washing off surface ice. Another main limitation of this workflow is the time it takes to produce lamellae. As production is slow, it is critical to have a properly optimized sample, making the milling as efficient as possible.
A number of adaptations to FIB-milling of vitreous biological samples have recently been introduced. A game-changer is the implementation of cryogenically cooled lift-out tools within the cryoFIB-SEM chamber that enable larger blocks of material to be milled from high-pressure frozen samples. The blocks can be attached to a metal rod or picked up in a gripper and moved to a second sample position, containing a specially modified EM grid. The blocks can then be coated with organoplatinum and milled to generate lamellae. The ability to mill lamellae from high-pressure frozen material means that much larger cells and tissues can be processed, specifically targeting areas by correlative fluorescence microscopy12. Other recent adaptations to the FIB-milling method include reducing curtaining artifacts by wedge pre-milling the sample16, microfluidic cryo-fixation49 and photo-micropatterning of electron microscopy grids to improve cell distribution50. Also, it has been demonstrated that milling micro-expansion gaps either side of a lamella can alleviate compression by the surrounding sample as it reaches its final thinness51. This may be particularly useful when milling continuous cell layers, such as the sample in this study, where bending of lamellae is sometimes seen during the final polishing step.
The future of electron microscopy will likely be the determination of in situ molecular structures by sub-tomogram averaging and FIB-milling is an important tool that will facilitate the production of vitreous biological samples for these types of workflows. While FIB-milling is still in its infancy for biological applications, methods development is happening at a rapid pace thanks to the hard work of researchers, both academic and at national facilities, plus commercial investment in developing cryoFIB-SEM technology to support research.

An erratum was issued for:&#160;<a href="https://www-jove-com.remotexs.ntu.edu.sg/t/62350">Preparing Lamellae from Vitreous Biological Samples using a Dual-Beam Scanning Electron Microscope for Cryo-Electron Tomography</a>. The Authors section was updated from:
Claudine Bisson1,2 
Corey W. Hecksel3,4 
James B. Gilchrist3 
Roland A. Fleck1 
1Centre for Ultrastructural Imaging, New Hunt&#8217;s House, Guy&#8217;s Campus, King&#8217;s College London 
2Department of Biological Science, Birkbeck College, University of London 
3Electron Bio-Imaging Centre, Diamond Light Source, Harwell Science and Innovation Campus 
4SLAC National Accelerator Laboratory, Stanford University
to:
Claudine Bisson1,2 
Corey W. Hecksel3,4 
James B. Gilchrist3 
M. Alejandra Carbajal1 
Roland A. Fleck1 
1Centre for Ultrastructural Imaging, New Hunt&#8217;s House, Guy&#8217;s Campus, King&#8217;s College London 
2Department of Biological Science, Birkbeck College, University of London 
3Electron Bio-Imaging Centre, Diamond Light Source, Harwell Science and Innovation Campus 
4SLAC National Accelerator Laboratory, Stanford University

An erratum was issued for:&#160;<a href="https://www-jove-com.remotexs.ntu.edu.sg/t/62350">Preparing Lamellae from Vitreous Biological Samples using a Dual-Beam Scanning Electron Microscope for Cryo-Electron Tomography</a>. The Authors section was updated.

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

Only the cellular contents of biological samples &#60;500 nm thick can be imaged effectively by transmission electron microscopy (TEM) at cryogenic temperatures, limiting the range of specimens to viruses, prokaryotes, simple single-cellular organisms, and thinner regions of larger eukaryotic cells1. On-grid focused ion beam (FIB)-milling enables thicker plunge-frozen biological samples to be thinned to electron transparent lamellae at cryogenic temperatures (&#60; -150 &#176;C). The resulting lamellae are then transferred to a TEM for visualization and tomographic data collection, enabling high-resolution 3D reconstructions of the cellular and molecular features inside of cells (for reviews see Rigort et al., 20122, Oikonomou et al., 20163, and Wagner et al., 20204).
FIB-milling emerged from the field of materials sciences, where samples are routinely thinned to prepare them for downstream analysis5. It is carried out in a scanning electron microscope (SEM), which has two optical columns: conventional scanning electron microscope optics and a second column containing optics able to generate and finely control a focused ion beam (FIB) - called a FIB-SEM. This allows a specific region of the sample to be ablated by ions generated by a gallium source, removing excess material and leaving behind a lamella6. The milling process is guided by SEM imaging of the topography of the sample, which is used to locate regions of interest and monitor milling progress. For biological applications, the basic setup is largely the same, but milling is carried out at cryogenic temperatures. This has required standard FIB-SEMs to be adapted to have cryo-cooled stages that maintain constant temperatures and low surface contamination rates, as well as airlocks to facilitate sample transfer without devitrification or contamination. Sample shuttles have also been modified to permit a range of different carriers to be mounted inside the cryoFIB-SEM, including TEM grids, planchettes, and capillaries. Several key groups of researchers have been central to the development of these methods and the continuing technological advances in this area7,8,9,10,11,12. Commercial solutions are now more widely available for biological FIB-milling at cryogenic temperature and on-grid milling of lamellae is becoming more routine, given an optimized sample.
A range of room temperature and cryoEM techniques can be used to visualize cellular information across all scales of life, from whole multicellular organisms at modest resolution to understanding the context of complex processes at the cellular level and in even greater detail, to the determination of in situ molecular structures13,14,15,16,17,18,19. Classical room temperature techniques include sectioning fixed and stained, resin imbedded cells and tissues by ultramicrotomy for analysis of cellular morphology by TEM (for review see Studer et al., 200820). Alternative techniques have been developed exploiting secondary electron scattering by SEM to image the surface of blocks of preserved cells, before progressively removing material either with a knife (serial block face imaging) or a focused ion beam21,22,23. This technique has also been successfully implemented at cryogenic temperatures (referred to as cryo-volume imaging) with a cryoFIB-SEM on vitreous, unstained blocks of cells or tissues24. Alternatively, thicker lamellae (~15 &#181;m thick) can be milled and studied by STEM imaging25. Using these techniques, whole blocks containing many cells can be imaged to gather population information or a whole organ/organism can be imaged and reconstructed in 3D. However, to access high-resolution molecular information from cells, samples need to be preserved in a near-native, frozen-hydrated state and, therefore, need to be prepared under cryogenic conditions. Cryo-electron microscopy of vitreous sections (CEMOVIS) is a technique whereby high-pressure frozen blocks of biological material are sectioned under cryogenic conditions with an ultramicrotome. This produces ribbons of cryo-sections (40-100 nm thick)26, which are attached to an EM grid and imaged in the TEM. However, the physical interaction of the knife with the vitreous sample causes crevassing and compression artifacts that can severely distort cellular structure27,28,29,30. Thicker sections are more prone to these artefacts, making it impractical to use sections thicker than ~70 nm26. This limitation greatly restricts the 3D view of the biological structure in the tomogram. FIB-milling at cryogenic temperatures does not experience these problems, but has its own artifacts caused by differential milling rates across parts of the specimen, leading to variable thicknesses within a lamella - called curtaining. This problem is mitigated by the application of an organoplatinum coat applied via a gas injection system (GIS), which protects the front edge of the lamella during milling31. The upper limit of sample thickness for on-grid FIB-milling is defined by the ability to plunge freeze the specimen while keeping it vitreous32, although, with the introduction of cryo lift-out techniques and adapted sample carriers for biological specimens, FIB-milling can also be used to process high-pressure frozen samples31,33,34,35. Additionally, plunge frozen samples cannot be too thin as there must be enough biological material to generate a reasonably sized lamella that will provide enough surface area to collect tilt-series at the required magnification. This problem can be alleviated by milling clumps of smaller cells, such as bacteria or yeast. Final lamella thickness (~100-300 nm) is usually dictated by the sample integrity and milling strategy. Thinner lamellae are better for high-resolution structural work, such as sub-tomogram averaging, but thicker lamellae contain much larger cellular volumes than can be achieved by CEMOVIS, providing more cellular context in a near to natively preserved sample. FIB-milling can also be used to thin plunge-frozen protein crystals for electron diffraction studies36.
FIB-milling of vitreous cells is worth the time and effort to carry out if the scientific question requires high-resolution molecular detail of near-native specimens in situ. With access to more facilities for the routine production of lamellae, the rate limiting step is often the optimization of the sample prior to milling, where time must be taken to ensure the sample is vitreous and an appropriate thickness to produce robust and uniformly thin lamellae. Described here is the sample optimization for FIB-milling plunge-frozen human red blood cells infected with Plasmodium falciparum parasites, the causative agent of malaria, but this approach could be adapted for any given sample.

<table><tbody><tr><td>c-clips</td><td>Thermo Fisher Scientific</td><td>1036171</td><td></td></tr><tr><td>Clipping station</td><td>Thermo Fisher Scientific</td><td>n/a</td><td>Direct quote from Thermo</td></tr><tr><td>Clipping station</td><td>Sub-angstrom</td><td>CSA-01</td><td></td></tr><tr><td>Compound 2</td><td>n/a</td><td>n/a</td><td>Synthesised by Dr Simon A. Osborne, LifeArc</td></tr><tr><td>Cryo-FIB specific autogrid rims</td><td>Thermo Fisher Scientific</td><td>1205101</td><td></td></tr><tr><td>E64</td><td>Sigma</td><td>E3132</td><td></td></tr><tr><td>Emitech K100X glow discharge unit</td><td>Quorum</td><td>n/a</td><td></td></tr><tr><td>Gibco RPMI 1640 media</td><td>Thermo Fisher Scientific</td><td>12633012</td><td>Formulation used for culturing is custom made (REF 041-91762 A) and includes Albumax, glutamine, HEPES and hypoxanthine supplements</td></tr><tr><td>Giemsa Stain</td><td>VWR International</td><td>350864X</td><td></td></tr><tr><td>Glass slides</td><td>Thermo Fisher Scientific</td><td>11562203</td><td></td></tr><tr><td>Grid boxes</td><td>Sub-angstrom</td><td>PB</td><td>For clipped grids</td></tr><tr><td>Grid boxes</td><td>Thermo Fisher Scientific</td><td>n/a</td><td>For clipped grids - direct quote from Thermo Fisher Scientific</td></tr><tr><td>Grid boxes</td><td>Agar Scientific</td><td>AGG3727</td><td></td></tr><tr><td>Home-made manual plunge freezing rig</td><td>n/a</td><td>n/a</td><td>With an insulated ethane pot (high-density foam) and liquid nitrogen bath (polystyrene) on a bench top in a containment facility.</td></tr><tr><td>Human blood</td><td>n/a</td><td>n/a</td><td>UK National Blood and Transplant service</td></tr><tr><td>JEOL 4700F Z JSM with a Leica VCT500 stage cooling system</td><td>JEOL</td><td>n/a</td><td>FIB-SEM</td></tr><tr><td>Leica EM ACE600 with a VCT500 cryostage</td><td>Leica</td><td>n/a</td><td>sputter coater</td></tr><tr><td>Leica EM ACE900</td><td>Leica</td><td>n/a</td><td>e-beam rotary coater. In a humidity controlled room.</td></tr><tr><td>Linkam cryo-stage for light microscope</td><td>Linkam</td><td>Model No. CMS196</td><td>Cassettes for clipped and un-clipped grids. In a humidity controlled room.</td></tr><tr><td>Methanol</td><td>Sigma</td><td>179957</td><td></td></tr><tr><td>Nikon Eclipse E200 light microscope</td><td>Nikon</td><td>n/a</td><td>with Linkam cryo-stage in a humidity controlled room.</td></tr><tr><td>Percoll</td><td>VWR International</td><td>17-0891-01</td><td>Solution for percoll cushion is 35 ml 10x PBS, 150 ml RPMI 1640 media and 315 ml Percoll</td></tr><tr><td>Quantifoil copper 200 mesh 2/4 holey carbon EM Finder grids</td><td>Quantifoil</td><td>N1-C17nCuH2-01</td><td>100 pack</td></tr><tr><td>Quantifoil copper 200 mesh 2/4 holey carbon EM grids</td><td>Quantifoil</td><td>N1-C17nCu20-01</td><td>100 pack</td></tr><tr><td>Quorum PP3010 prep-chamber</td><td>Quorum</td><td>n/a</td><td>sputter coater</td></tr><tr><td>Scios dual beam equipped with a Quorum PP3010 transfer stage.</td><td>FEI</td><td>n/a</td><td>FIB-SEM</td></tr><tr><td>Staedtler Lumocolor fine black permanent marker pen</td><td>Viking Direct</td><td>ND538522</td><td></td></tr><tr><td>TFS Titan Krios 300 kV TEM</td><td>Thermo Fisher Scientific</td><td>n/a</td><td>TEM equipped with a K2 or K3 camera.</td></tr><tr><td>Whatmann grade 1 filter paper</td><td>Sigma</td><td>WHA1001150</td><td></td></tr><tr><td>Zeiss Axio Scope A1 light microscope</td><td>Zeiss</td><td>n/a</td><td>with Linkam cryo-stage in a humidity controlled room.</td></tr></tbody></table>

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.

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

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 ...

preparing-lamellae-from-vitreous-biological-samples-using-dual-beam

Nanyang Technological University

Research

JoVE Journal

Biochemistry

3.2K Views.  New Hunt’s House, Guy’s Campus, King’s College London. Dit protocol schetst de bereiding van glasvochtlamellen van met Plasmodium-falciparum geïnfecteerde rode bloedcellen voor cryo-elektronentomografie. Dit maakt directe observatie van gedetailleerde structurele informatie mogelijk over hoe de parasiet malaria veroorzaakt. Cryo-elektronentomografie registreert informatie met hoge resolutie over de binnenkant van cellen.Het is krachtig genoeg om afzonderlijke eiwitcomplexen in situ te observeren, die computationeel kunnen worden geëxtrah...

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

Preparation of Primary Neurons for Visualizing Neurites in a Frozen-hydrated State Using Cryo-Electron Tomography

Neurites, both dendrites and axons, are neuronal cellular processes that enable the conduction of electrical impulses between neurons. Defining the structure of neurites is critical to understanding how these processes move materials and signals that support synaptic communication. Electron microscopy (EM) has been traditionally used to assess the ultrastructural features within neurites; however, the exposure to organic solvent during dehydration and resin embedding can distort structures. An important unmet goal is the formulation of procedures that allow for structural evaluations not impacted by such artifacts.
Here, we have established a detailed and reproducible protocol for growing and flash-freezing whole neurites of different primary neurons on electron microscopy grids followed by their examination with cryo-electron tomography (cryo-ET). This technique allows for 3-D visualization of frozen, hydrated neurites at nanometer resolution, facilitating assessment of their morphological differences. Our protocol yields an unprecedented view of dorsal root ganglion (DRG) neurites, and a visualization of hippocampal neurites in their near-native state. As such, these methods create a foundation for future studies on neurites of both normal neurons and those impacted by neurological disorders.

To preserve neuronal processes for ultrastructural analysis, we describe a protocol for plating of primary neurons on electron microscopy grids followed by flash freezing, yielding samples suspended in a layer of vitreous ice. These samples can be examined with a cryo-electron microscope to visualize structures at the nanometer scale.

Neurites, both dendrites and axons, are neuronal cellular processes that enable the conduction of electrical impulses between neurons. Defining the ...

Neuroscience

Using Tomoauto: A Protocol for High-throughput Automated Cryo-electron Tomography

Cryo-electron tomography (Cryo-ET) is a powerful three-dimensional (3-D) imaging technique for visualizing macromolecular complexes in their native context at a molecular level. The technique involves initially preserving the sample in its native state by rapidly freezing the specimen in vitreous ice, then collecting a series of micrographs from different angles at high magnification, and finally computationally reconstructing a 3-D density map. The frozen-hydrated specimen is extremely sensitive to the electron beam and so micrographs are collected at very low electron doses to limit the radiation damage. As a result, the raw cryo-tomogram has a very low signal to noise ratio characterized by an intrinsically noisy image. To better visualize subjects of interest, conventional imaging analysis and sub-tomogram averaging in which sub-tomograms of the subject are extracted from the initial tomogram and aligned and averaged are utilized to improve both contrast and resolution. Large datasets of tilt-series are essential to understanding and resolving the complexes at different states, conditions, or mutations as well as obtaining a large enough collection of sub-tomograms for averaging and classification. Collecting and processing this data can be a major obstacle preventing further analysis. Here we describe a high-throughput cryo-ET protocol based on a computer-controlled 300kV cryo-electron microscope, a direct detection device (DDD) camera and a highly effective, semi-automated image-processing pipeline software wrapper library tomoauto developed in-house. This protocol has been effectively utilized to visualize the intact type III secretion system (T3SS) in Shigella flexneri minicells. It can be applicable to any project suitable for cryo-ET.

We present a protocol on how to utilize high-throughput cryo-electron tomography to determine high resolution in situ structures of molecular machines. The protocol permits large amounts of data to be processed, avoids common bottlenecks and reduces resource downtime, allowing the user to focus on important biological questions.

Cryo-electron tomography (Cryo-ET) is a powerful three-dimensional (3-D) imaging technique for visualizing macromolecular complexes in their native ...

Bioengineering

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 ...

Single Particle Cryo-Electron Microscopy: From Sample to Structure

Cryo-electron microscopy (cryoEM) is a powerful technique for structure determination of macromolecular complexes, via single particle analysis (SPA). The overall process involves i) vitrifying the specimen in a thin film supported on a cryoEM grid; ii) screening the specimen to assess particle distribution and ice quality; iii) if the grid is suitable, collecting a single particle dataset for analysis; and iv) image processing to yield an EM density map. In this protocol, an overview for each of these steps is provided, with a focus on the variables which a user can modify during the workflow and the troubleshooting of common issues. With remote microscope operation becoming standard in many facilities, variations on imaging protocols to assist users in efficient operation and imaging when physical access to the microscope is limited will be described.

Structure determination of macromolecular complexes using cryoEM has become routine for certain classes of proteins and complexes. Here, this pipeline is summarized (sample preparation, screening, data acquisition and processing) and readers are directed towards further detailed resources and variables that may be altered in the case of more challenging specimens.

Cryo-electron microscopy (cryoEM) is a powerful technique for structure determination of macromolecular complexes, via single particle analysis (SPA). The ...

Electron Cryotomography of Bacterial Cells

While much is already known about the basic metabolism of bacterial cells, many fundamental questions are still surprisingly unanswered, including for instance how they generate and maintain specific cell shapes, establish polarity, segregate their genomes, and divide. In order to understand these phenomena, imaging technologies are needed that bridge the resolution gap between fluorescence light microscopy and higher-resolution methods such as X-ray crystallography and NMR spectroscopy.
Electron cryotomography (ECT) is an emerging technology that does just this, allowing the ultrastructure of cells to be visualized in a near-native state, in three dimensions (3D), with "macromolecular" resolution (~4nm).1, 2 In ECT, cells are imaged in a vitreous, "frozen-hydrated" state in a cryo transmission electron microscope (cryoTEM) at low temperature (&lt; -180&deg;C). For slender cells (up to ~500 nm in thickness3), intact cells are plunge-frozen within media across EM grids in cryogens such as ethane or ethane/propane mixtures. Thicker cells and biofilms can also be imaged in a vitreous state by first "high-pressure freezing" and then, "cryo-sectioning" them. A series of two-dimensional projection images are then collected through the sample as it is incrementally tilted along one or two axes. A three-dimensional reconstruction, or "tomogram" can then be calculated from the images. While ECT requires expensive instrumentation, in recent years, it has been used in a few labs to reveal the structures of various external appendages, the structures of different cell envelopes, the positions and structures of cytoskeletal filaments, and the locations and architectures of large macromolecular assemblies such as flagellar motors, internal compartments and chemoreceptor arrays.1, 2
In this video article we illustrate how to image cells with ECT, including the processes of sample preparation, data collection, tomogram reconstruction, and interpretation of the results through segmentation and in some cases correlation with light microscopy.

We illustrate here how to use electron cryotomography (ECT) to study the ultrastructure of bacterial cells in near-native states, to "macromolecular" (~4 nm) resolution.

While much is already known about the basic metabolism of bacterial cells, many fundamental questions are still surprisingly unanswered, including for ...

Biology

Preparation of Electron Microscopy Grids for In Situ Cellular Cryotomography

Sample Preparation for In Situ Cryotomography of Mammalian Cells

In situ cellular cryotomography is a powerful technique for studying complex objects in their native frozen-hydrated cellular context, making it highly relevant to cellular biology and virology. The potential of combining cryotomography with other microscopy modalities makes it a perfect technique for integrative and correlative imaging. However, sample preparation for in situ cellular tomography is not straightforward, as cells do not readily attach and stretch over the electron microscopy grid. Additionally, the grids themselves are fragile and can break if handled too forcefully, resulting in the loss of imageable areas. The geometry of tissue culture dishes can also pose a challenge when manipulating the grids with tweezers. Here, we describe the tips and tricks to overcome these (and other) challenges and prepare good-quality samples for in situ cellular cryotomography and correlative imaging of adherent mammalian cells. With continued advances in cryomicroscopy technology, this technique holds enormous promise for advancing our understanding of complex biological systems.

Author Spotlight: Optimizing Grid Preparation for Enhanced Cryoelectron Tomography 

This method provides an accessible and flexible protocol for the preparation of electron microscopy (EM) grids for in situ cellular cryotomography and correlative light and electron microscopy (CLEM).

In situ cellular cryotomography is a powerful technique for studying complex objects in their native frozen-hydrated cellular context, making it highly ...

Correlative Light Electron Microscopy (CLEM) for Tracking and Imaging Viral Protein Associated Structures in Cryo-immobilized Cells

Due to its high resolution, electron microscopy (EM) is an indispensable tool for virologists. However, one of the main difficulties when analyzing virus-infected or transfected cells via EM are the low efficiencies of infection or transfection, hindering the examination of these cells. In order to overcome this difficulty, light microscopy (LM) can be performed first to allocate the subpopulation of infected or transfected cells. Thus, taking advantage of the use of fluorescent proteins (FPs) fused to viral proteins, LM is used here to record the positions of the "positive-transfected" cells, expressing a FP and growing on a support with an alphanumeric pattern. Subsequently, cells are further processed for EM via high pressure freezing (HPF), freeze substitution (FS) and resin embedding. The ultra-rapid freezing step ensures excellent membrane preservation of the selected cells that can then be analyzed at the ultrastructural level by transmission electron microscopy (TEM). Here, a step-by-step correlative light electron microscopy (CLEM) workflow is provided, describing sample preparation, imaging and correlation in detail. The experimental design can be also applied to address many cell biology questions.

Correlative Light Electron Microscopy CLEM for Tracking and Imaging Viral Protein Associated Structures in Cryo-immobilized Cells

A correlative light electron microscopy (CLEM) method is applied to image virus-induced intracellular structures via electron microscopy (EM) in cells that are previously selected by light microscopy (LM). LM and EM are combined as a hybrid imaging approach to achieve an integrated view of virus-host interactions.

Due to its high resolution, electron microscopy (EM) is an indispensable tool for virologists. However, one of the main difficulties when analyzing ...

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 ...

Array Tomography Workflow for the Targeted Acquisition of Volume Information using Scanning Electron Microscopy

Electron microscopy is applied in biology and medicine for imaging of cellular and structural details at nanometer resolution. Historically, Transmission Electron Microscopy (TEM) provided insight into cell ultrastructure, but in the recent decade, the development of modern Scanning Electron Microscopes (SEM) has changed the way of looking inside the cells. Even though the resolution of TEM is superior when protein-level structural details are needed, SEM-resolution is sufficient for the majority of the organelle-level cell biology-related questions. The advancement in technology enabled automatic volume acquisition solutions such as Serial block-face imaging (SBF-SEM) and Focused ion beam SEM (FIB-SEM). Nevertheless, to this day, these methods remain inefficient when the identification and navigation to areas of interest are crucial. Without the means for precise localization of target areas before imaging, operators need to acquire much more data than they need (in SBF-SEM), or, even worse, prepare many grids and image them all (in TEM). We propose the strategy of "lateral screening" using Array Tomography in SEM, which facilitates the localization of areas of interest, followed by automated imaging of the relevant fraction of the total sample volume. Array tomography samples are conserved during imaging, and they can be arranged into section libraries ready for repeated imaging. Several examples are shown in which lateral screening enables us to analyze structural details that are incredibly challenging to access with any other method.

We describe the preparation of ribbons of serial sections and their collection on large transfer support for use as Array Tomography samples, along with automated imaging procedures in a scanning electron microscope. The protocol allows screening, retrieval, and targeted imaging of local, rare events, and the acquisition of large data volumes.

Electron microscopy is applied in biology and medicine for imaging of cellular and structural details at nanometer resolution. Historically, Transmission ...

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.

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

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