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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<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.

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

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

Research

JoVE Journal

Biochemistry

3.1K Views.  New Hunt’s House, Guy’s Campus, King’s College London. Using focused ion beam milling to produce vitreous on-grid lamellae from plunge frozen biological samples for cryo-electron tomography.

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

Use of Electron Paramagnetic Resonance in Biological Samples at Ambient Temperature and 77 K

The accurate and specific detection of reactive oxygen species (ROS) in different cellular and tissue compartments is essential to the study of redox-regulated signaling in biological settings. Electron paramagnetic resonance spectroscopy (EPR) is the only direct method to assess free radicals unambiguously. Its advantage is that it detects physiologic levels of specific species with a high specificity, but it does require specialized technology, careful sample preparation, and appropriate controls to ensure accurate interpretation of the data. Cyclic hydroxylamine spin probes react selectively with superoxide or other radicals to generate a nitroxide signal that can be quantified by EPR spectroscopy. Cell-permeable spin probes and spin probes designed to accumulate rapidly in the mitochondria allow for the determination of superoxide concentration in different cellular compartments.
In cultured cells, the use of cell permeable 1-hydroxy-3-methoxycarbonyl-2,2,5,5-tetramethylpyrrolidine (CMH) along with and without cell-impermeable superoxide dismutase (SOD) pretreatment, or use of cell-permeable PEG-SOD,&#160;allows for the differentiation of extracellular from cytosolic superoxide. The mitochondrial 1-hydroxy-4-[2-triphenylphosphonio)-acetamido]-2,2,6,6-tetramethyl-piperidine,1-hydroxy-2,2,6,6-tetramethyl-4-[2-(triphenylphosphonio)acetamido] piperidinium dichloride (mito-TEMPO-H) allows for measurement of mitochondrial ROS (predominantly superoxide).
Spin probes and EPR spectroscopy can also be applied to in vivo models. Superoxide can be detected in extracellular fluids such as blood and alveolar fluid, as well as tissues such as lung tissue. Several methods are presented to process and store tissue for EPR measurements and deliver intravenous 1-hydroxy-3-carboxy-2,2,5,5-tetramethylpyrrolidine (CPH) spin probe in vivo. While measurements can be performed at room temperature, samples obtained from in vitro and in vivo models can also be stored at -80 &#176;C and analyzed by EPR at 77 K. The samples can be stored in specialized tubing stable at -80 &#176;C and run at 77 K to enable a practical, efficient, and reproducible method that facilitates storing and transferring samples.

Electron paramagnetic resonance (EPR) spectroscopy is an unambiguous method to measure free radicals. The use of selective spin probes allows for detection of free radicals in different cellular compartments. We present a practical, efficient method to collect biological samples that facilitate treating, storing, and transferring samples for EPR measurements.

The accurate and specific detection of reactive oxygen species (ROS) in different cellular and tissue compartments is essential to the study of ...

Fast Grid Preparation for Time-Resolved Cryo-Electron Microscopy

The field of cryo-electron microscopy (cryo-EM) is rapidly developing with new hardware and processing algorithms, producing higher resolution structures and information on more challenging systems. Sample preparation for cryo-EM is undergoing a similar revolution with new approaches being developed to supersede the traditional blotting systems. These include the use of piezo-electric dispensers, pin printing and direct spraying. As a result of these developments, the speed of grid preparation is going from seconds to milliseconds, providing new opportunities, especially in the field of time-resolved cryo-EM where proteins and substrates can be rapidly mixed before plunge freezing, trapping short lived intermediate states. Here we describe, in detail, a standard protocol for making grids on our in-house time-resolved EM device both for standard fast grid preparation and also for time-resolved experiments. The protocol requires a minimum of about 50 &#181;L sample at concentrations of &#8805; 2 mg/mL for the preparation of 4 grids. The delay between sample application and freezing can be as low as 10 ms. One limitation is increased ice thickness at faster speeds and compared to the blotting method. We hope this protocol will aid others in designing their own grid making devices and those interested in designing time-resolved experiments.

Here, we provide a detailed protocol for the use of a rapid grid making device for both fast grid-making and for rapid mixing and freezing to conduct time-resolved experiments.

The field of cryo-electron microscopy (cryo-EM) is rapidly developing with new hardware and processing algorithms, producing higher resolution structures ...

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 ...

Micropatterning Transmission Electron Microscopy Grids to Direct Cell Positioning within Whole-Cell Cryo-Electron Tomography Workflows

Whole-cell cryo-electron tomography (cryo-ET) is a powerful technology that is used to produce nanometer-level resolution structures of macromolecules present in the cellular context and preserved in a near-native frozen-hydrated state. However, there are challenges associated with culturing and/or adhering cells onto TEM grids in a manner that is suitable for tomography while retaining the cells in their physiological state. Here, a detailed step-by-step protocol is presented on the use of micropatterning to direct and promote eukaryotic cell growth on TEM grids. During micropatterning, cell growth is directed by depositing extra-cellular matrix (ECM) proteins within specified patterns and positions on the foil of the TEM grid while the other areas remain coated with an anti-fouling layer. Flexibility in the choice of surface coating and pattern design makes micropatterning broadly applicable for a wide range of cell types. Micropatterning is useful for studies of structures within individual cells as well as more complex experimental systems such as host-pathogen interactions or differentiated multi-cellular communities. Micropatterning may also be integrated into many downstream whole-cell cryo-ET workflows, including correlative light and electron microscopy (cryo-CLEM) and focused-ion beam milling (cryo-FIB).

The goal of this protocol is to direct cell adhesion and growth to targeted areas of grids for cryo-electron microscopy. This is achieved by applying an anti-fouling layer that is ablated in user-specified patterns followed by deposition of extra-cellular matrix proteins in the patterned areas prior to cell seeding.

Whole-cell cryo-electron tomography (cryo-ET) is a powerful technology that is used to produce nanometer-level resolution structures of macromolecules ...

A Robust Single-Particle Cryo-Electron Microscopy (cryo-EM) Processing Workflow with cryoSPARC, RELION, and Scipion

Recent advances in both instrumentation and image processing software have made single-particle cryo-electron microscopy (cryo-EM) the preferred method for structural biologists to determine high-resolution structures of a wide variety of macromolecules. Multiple software suites are available to new and expert users for image processing and structure calculation, which streamline the same basic workflow: movies acquired by the microscope detectors undergo correction for beam-induced motion and contrast transfer function (CTF) estimation. Next, particle images are selected and extracted from averaged movie frames for iterative 2D and 3D classification, followed by 3D reconstruction, refinement, and validation. Because various software packages employ different algorithms and require varying levels of expertise to operate, the 3D maps they generate often differ in quality and resolution. Thus, users regularly transfer data between a variety of programs for optimal results. This paper provides a guide for users to navigate a workflow across the popular software packages: cryoSPARC v3, RELION-3, and Scipion 3 to obtain a near-atomic resolution structure of the adeno-associated virus (AAV). We first detail an image processing pipeline with cryoSPARC v3, as its efficient algorithms and easy-to-use GUI allow users to quickly arrive at a 3D map. In the next step, we use PyEM and in-house scripts to convert and transfer particle coordinates from the best quality 3D reconstruction obtained in cryoSPARC v3 to RELION-3 and Scipion 3 and recalculate 3D maps. Finally, we outline steps for further refinement and validation of the resultant structures by integrating algorithms from RELION-3 and Scipion 3. In this article, we describe how to effectively utilize three processing platforms to create a single and robust workflow applicable to a variety of data sets for high-resolution structure determination.

A Robust Single-Particle Cryo-Electron Microscopy cryo-EM Processing Workflow with cryoSPARC, RELION, and Scipion

This article describes how to effectively utilize three cryo-EM processing platforms, i.e., cryoSPARC v3, RELION-3, and Scipion 3, to create a single and robust workflow applicable to a variety of single-particle data sets for high-resolution structure determination.

Recent advances in both instrumentation and image processing software have made single-particle cryo-electron microscopy (cryo-EM) the preferred method ...

Preparation of Nucleosome Core Particles Complexed with DNA Repair Factors for Cryo-Electron Microscopy Structural Determination

DNA repair in the context of chromatin is poorly understood. Biochemical studies using nucleosome core particles, the fundamental repeating unit of chromatin, show most DNA repair enzymes remove DNA damage at reduced rates as compared to free DNA. The molecular details on how base excision repair (BER) enzymes recognize and remove DNA damage in nucleosomes have not been elucidated. However, biochemical BER data of nucleosomal substrates suggest the nucleosome presents different structural barriers dependent on the location of the DNA lesion and the enzyme. This indicates the mechanisms employed by these enzymes to remove DNA damage in free DNA may be different than those employed in nucleosomes. Given that the majority of genomic DNA is assembled into nucleosomes, structural information of these complexes is needed. To date, the scientific community lacks detailed protocols to perform technically feasible structural studies of these complexes. Here, we provide two methods to prepare a complex of two genetically fused BER enzymes (Polymerase &#946; and AP Endonuclease1) bound to a single-nucleotide gap near the entry-exit of the nucleosome for cryo-electron microscopy (cryo-EM) structural determination. Both methods of sample preparation are compatible for vitrifying quality grids via plunge freezing. This protocol can be used as a starting point to prepare other nucleosomal complexes with different BER factors, pioneer transcription factors, and chromatin-modifying enzymes.

This protocol outlines in detail the preparation of nucleosomal complexes using two methods of sample preparation for freezing TEM grids.

DNA repair in the context of chromatin is poorly understood. Biochemical studies using nucleosome core particles, the fundamental repeating unit of ...

Extracting Modified Microtubules from Mammalian Cells to Study Microtubule-Protein Complexes by Cryo-Electron Microscopy

Microtubules are an important part of the cytoskeleton and are involved in intracellular organization, cell division, and migration. Depending on the posttranslational modifications, microtubules can form complexes with various interacting proteins. These microtubule-protein complexes are often implicated in human diseases. Understanding the structure of such complexes is useful for elucidating their mechanisms of action and can be studied by cryo-electron microscopy (cryo-EM). To obtain such complexes for structural studies, it is important to extract microtubules containing or lacking specific posttranslational modifications. Here, we describe a simplified protocol to extract endogenous tubulin from genetically modified mammalian cells, involving microtubule polymerization, followed by sedimentation using ultracentrifugation. The extracted tubulin can then be used to prepare cryo-electron microscope grids with microtubules that are bound to a purified microtubule-binding protein of interest. As an example, we demonstrate the extraction of fully tyrosinated microtubules from cell lines engineered to lack the three known tubulin-detyrosinating enzymes. These microtubules are then used to make a protein complex with enzymatically inactive microtubule-associated tubulin detyrosinase on cryo-EM grids.

Here, we describe a protocol to extract endogenous tubulin from mammalian cells, which can lack or contain specific microtubule-modifying enzymes, to obtain microtubules enriched for a specific modification. We then describe how the extracted microtubules can be decorated with purified microtubule-binding proteins to prepare grids for cryo-electron microscopy.

Microtubules are an important part of the cytoskeleton and are involved in intracellular organization, cell division, and migration. Depending on the ...

Coating CryoEM Grids with Monolayer Graphene to Improve Cryo-Sample Preparation

Application of Monolayer Graphene to Cryo-Electron Microscopy Grids for High-resolution Structure Determination

In cryogenic electron microscopy (cryoEM), purified macromolecules are applied to a grid bearing a holey carbon foil; the molecules are then blotted to remove excess liquid and rapidly frozen in a roughly 20-100 nm thick layer of vitreous ice, suspended across roughly 1 &#181;m wide foil holes. The resulting sample is imaged using cryogenic transmission electron microscopy, and after image processing using suitable software, near-atomic resolution structures can be determined. Despite cryoEM&#39;s widespread adoption, sample preparation remains a severe bottleneck in cryoEM workflows, with users often encountering challenges related to samples behaving poorly in the suspended vitreous ice. Recently, methods have been developed to modify cryoEM grids with a single continuous layer of graphene, which acts as a support surface that often increases particle density in the imaged area and can reduce interactions between particles and the air-water interface. Here, we provide detailed protocols for the application of graphene to cryoEM grids and for rapidly assessing the relative hydrophilicity of the resulting grids. Additionally, we describe an EM-based method to confirm the presence of graphene by visualizing its characteristic diffraction pattern. Finally, we demonstrate the utility of these graphene supports by rapidly reconstructing a 2.7 &#197; resolution density map of a Cas9 complex using a pure sample at a relatively low concentration.

Author Spotlight: Enhancing CryoEM Sample Preparation Using Graphene Monolayer on Microscopy Grids 

The application of support layers to cryogenic electron microscopy (cryoEM) grids can increase particle density, limit interactions with the air-water interface, reduce beam-induced motion, and improve the distribution of particle orientations. This paper describes a robust protocol for coating cryoEM grids with a monolayer of graphene for improved cryo-sample preparation.

In cryogenic electron microscopy (cryoEM), purified macromolecules are applied to a grid bearing a holey carbon foil; the molecules are then blotted to ...

Accelerating Macromolecular Structural Analysis Using Smart Leginon Autoscreen

Cryo-Electron Microscopy Screening Automation Across Multiple Grids Using Smart Leginon

Advancements in cryo-electron microscopy (cryoEM) techniques over the past decade have allowed structural biologists to routinely resolve macromolecular protein complexes to near-atomic resolution. The general workflow of the entire cryoEM pipeline involves iterating between sample preparation, cryoEM grid preparation, and sample/grid screening before moving on to high-resolution data collection. Iterating between sample/grid preparation and screening is typically a major bottleneck for researchers, as every iterative experiment must optimize for sample concentration, buffer conditions, grid material, grid hole size, ice thickness, and protein particle behavior in the ice, amongst other variables. Furthermore, once these variables are satisfactorily determined, grids prepared under identical conditions vary widely in whether they are ready for data collection, so additional screening sessions prior to selecting optimal grids for high-resolution data collection are recommended. This sample/grid preparation and screening process often consumes several dozen grids and days of operator time at the microscope. Furthermore, the screening process is limited to operator/microscope availability and microscope accessibility. Here, we demonstrate how to use Leginon and Smart Leginon Autoscreen to automate the majority of cryoEM grid screening. Autoscreen combines machine learning, computer vision algorithms, and microscope-handling algorithms to remove the need for constant manual operator input. Autoscreen can autonomously load and image grids with multi-scale imaging using an automated specimen-exchange cassette system, resulting in unattended grid screening for an entire cassette. As a result, operator time for screening 12 grids may be reduced to ~10 min with Autoscreen compared to ~6 h using previous methods which are hampered by their inability to account for high variability between grids. This protocol first introduces basic Leginon setup and functionality, then demonstrates Autoscreen functionality step-by-step from the creation of a template session to the end of a 12-grid automated screening session.

Author Spotlight: Enhancing Cryo-Electron Microscopy by Automated Data Collection and Analysis Techniques

Cryo-electron microscopy (cryoEM) multi-grid screening is often a tedious process that demands hours of attention. This protocol shows how to set up a standard Leginon collection and Smart Leginon Autoscreen to automate this process. This protocol can be applied to the majority of cryoEM holey foil grids.

Advancements in cryo-electron microscopy (cryoEM) techniques over the past decade have allowed structural biologists to routinely resolve macromolecular ...