Name: Preparing Lamellae from Vitreous Biological Samples Using a Dual-Beam Scanning Electron Microscope for Cryo-Electron Tomography (Video) | JoVE Journal
Uploaded: 2021-08-05T13:30:00.000+00:00
Duration: 7 min
Description: Learn cryo-lamella preparation for cryo-electron tomography using Plasmodium falciparum-infected erythrocytes. This protocol enables high-resolution structural analysis of malaria parasites within host cells.

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5 minutes

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/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/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

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

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

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

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

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

Research

JoVE Journal

Biochemistry

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.

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

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.

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

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.

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

This report describes a sample preparation protocol and specific imaging conditions for performing scanning transmission electron tomography of thick biological specimens.

Preparation and Observation of Thick Biological Samples by Scanning Transmission 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 Saccharomyces cerevisiae.

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

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.

Strategies for Optimization of Cryogenic Electron Tomography Data Acquisition

Here, we present a pipeline for 3D-correlative focused ion beam milling on guiding the preparation of cellular samples for cryo-electron tomography. The 3D position of fluorescently tagged proteins of interest is first determined by cryo-fluorescence microscopy, and then targeted for milling. The protocol is suitable for mammalian, yeast, and bacterial cells.

Sample Preparation by 3D-Correlative Focused Ion Beam Milling for High-Resolution Cryo-Electron Tomography

This workflow enables lamella production targeting fluorescently labeled biological structures that are small (&lt;1 μm in axial extent) and rare (1 copy per cell) using a cryogenic tri-coincident imaging platform. This platform integrates fluorescence microscopy, focused ion beam milling, and scanning electron microscopy at a single focal position and enables simultaneous fluorescence microscopy while milling.

Workflow Using a Cryogenic Coincident Fluorescence, Electron, and Ion Beam Microscope for Targeted Milling of Cells

This video protocol illustrates how to perform scanning transmission electron microscopy tomography of virological specimens. For optimal outcomes, samples are prepared by high-pressure freezing and subsequent freeze substitution.

Scanning Transmission Electron Microscopy Tomography in Virology: 3D Imaging of High-pressure Frozen, Freeze-substituted Samples

This protocol standardizes cryo-electron tomography (cryo‑ET) for thick tissues by integrating high-pressure freezing (HPF) with the waffle method, cryogenic focus ion beam (cryo‑FIB) lift‑out, and dose‑symmetric, parallel acquisition, yielding uniformly thin lamellae and vitrified specimens for reproducible in situ analysis in the mouse hippocampus.

Cryo-Electron Tomography Workflow for Thick Tissues Demonstrated in Mouse Hippocampus

Learn cryo-lamella preparation for cryo-electron tomography using Plasmodium falciparum-infected erythrocytes. This protocol enables high-resolution structural analysis of malaria parasites within host cells.

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

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