We would like to thank the Mansky lab for access to plunge-freezing equipment. Parts of this work were carried out in the characterization facility of the University of Minnesota, which receives partial support from the National Science Foundation (NSF) through the Materials Research Science and Engineering Center (MRSEC; Award Number DMR-2011401) and the National Neuroscience Curriculum Initiative (NNCI; Award Number ECCS-2025124) programs. We would like to thank funding from the Behavior of HIV in Viral Environments center (B-HIVE; 1U54AI170855-01) and the Duke Center for HIV Structural Biology (DCHSB; U54AI170752) center.

1. Grid preparation
NOTE: In the experimental design, plan for a maximum of 8-12 grids total per plunge-freezing session and 4-5 grids per well. More than that will lead to a very long plunge-freezing session, which may cause increased cell stress, ice contamination, and user errors.
<ol>
	<li>Glow discharge
	<ol>
		<li>Load an appropriate number of grids with a perforated carbon support film to be glow discharged at 15 mA for 45 s at 0.39 mBar atmosphere in a glow discharge device.</li>
		<li>Make sure the carbon side of the grid is facing up. When finished, transfer the grids to a clean container lined with filter paper (Figure 1A1). 
		NOTE: Other grids may be used as long as the material is not toxic to cells (which excludes any copper grid). Finder grids are useful for correlative studies. Grids made of other materials are also possible. Many labs have had good results growing cells on top of SiO2 support grids8,9.</li>
	</ol>
	</li>
	<li>Adherent treatment
	<ol>
		<li>Transfer the grids, fibronectin, PBS, plates, and tweezers to a biosafety cabinet. Treat both sides of the grids with 20 &#181;g/mL bovine fibronectin by micropipetting two generously sized dots of adherent solution on a sterile surface, such as the lid of a 6 well plate. Ensure that the dots are large enough to envelop the entire grid (around 100 &#181;L is recommended). 
		NOTE: Prior to treatment with the selected adherent solution, there is the option of photo-micropatterning the grid here to optimize the attachment of cells on the center of grid squares. Reducing the ability of cells to attach to the bars will increase the number of imageable cells. This is ideal for experiments involving cryoFIB milling.</li>
		<li>Use 5/15 tweezers to manipulate the grids. Be sure to treat both sides of the grid in solution. 
		&#8203;NOTE: Other adhesive solutions may be used (fibrinogen, collagen, other extracellular matrix components, etc.). Fibronectin is used here, as it gives good results with a wide variety of cell lines. (U2OS, HeLa, Vero, Calu-3, Tzm-bl). The 5/15 tweezer is useful as its geometry allows improved maneuverability around the 6-well plates, 35 mm dishes, and micro slides. This results in less mechanical stress on the grids and a more intact carbon film. Make sure all required materials are placed in the biosafety cabinet prior to beginning, as this reduces the possibility for grid contamination.</li>
	</ol>
	</li>
	<li>Attaching grids
	<ol>
		<li>Once treated with bovine fibronectin, the grid will become sticky. Using 5/15 tweezers, gently touch the non-carbon surface of the grid to the bottom of an empty dish/plate and open the tweezers. The grid should easily attach to the new surface (Figure 1A2). 
		NOTE: Try to avoid placing grids in the direct center or the edges of the dish/plate. When adding cells, there is a tendency for cell density to accumulate in the center of the plate. This can result in grids with excessive cellular density. Alternatively, 3D-printed grid holders may be used here instead of attaching cells directly to the surface of the dish/plate12.</li>
	</ol>
	</li>
	<li>Incubation
	<ol>
		<li>Incubate the grids for 30 min in the fibronectin adherent solution (20 &#181;g/mL). To ensure that the grids do not become dry during this process, micropipette 1-2 drops of more adherent solution directly on top of grids every 15 min (Figure 1A3). 
		NOTE: The incubation period will vary depending on the adherent solution used. For bovine fibronectin, the recommended time is 30 min.</li>
	</ol>
	</li>
	<li>Washing
	<ol>
		<li>Following the incubation period, remove excess adherent solution via micropipetting. Wash the grids by micropipetting drops of PBS directly on top of the grids. Repeat this 2-3 times. (Figure 1A4).</li>
	</ol>
	</li>
	<li>Sterilization
	<ol>
		<li>Use UV light to sterilize the grids in the biosafety cabinet for 1 h. Place the grids as close to the UV source as possible to maximize sterilization (Figure 1A5). To ensure that the grids do not become dry during this process, micropipette 1-2 drops of PBS directly on top of grids every 10 min. 
		&#8203;NOTE : Steps 1.4-1.6 can be performed concurrently.</li>
	</ol>
	</li>
</ol>
2. Seeding grids
<ol>
	<li>Count cells:
	<ol>
		<li>Detach and count the cells using mechanical or enzymatic methods. Determine the optimum number of cells to use for seeding prior to counting. For U2OS seeded in a 6-well plate, use anywhere between 6 x 104 to 1.6 x 105 cells per well (Figure 1B1). 
		NOTE: The number of cells seeded will vary according to the cell type, the surface area of the well, dish or microslide used, the time between seeding and plunge freezing, and the experimental design. Test a range of cell numbers to identify conditions that will result in 0.25-1 cells per grid square at the time of plunge freezing. It is important to note that high cell density reduces the speed of heat transfer and may impact the vitrification process. To combat this, some labs have found success using cell strainers, which act to prevent cell clumps from forming13. Finally, this protocol results in the random attachment of cells to the carbon layer. If a targeted attachment is necessary, photo-micropatterning may be used (see step 1.3)14.</li>
	</ol>
	</li>
	<li>Adding cells and growth medium : Add cells via micropipette to the wells, avoiding bubbles (Figure 1B2). In a 6-well plate, a total well volume of 1.5-2.0 mL is recommended. 
	NOTE: It is recommended to wet around the grids carefully before adding the entire volume. This will help prevent grids from detaching and floating in the solution. Gently move the plate side-to-side to facilitate homogeneous cell distribution on the well. Do not swirl the plate in a circular motion, as this will result in excessive cell density allocating in the center of the well.</li>
	<li>Incubation: Incubate the slides at 37 &#176;C for an appropriate time according to the experimental design, which will depend on downstream applications. For this example (transfection of HIV molecular clones), incubate between 16-24 h (Figure 1B3). 
	&#8203;NOTE: Longer incubation times will result in more cell division cycles. Therefore, adjust the starting number of cells for seeding accordingly.</li>
</ol>
3. Transfection
<ol>
	<li>Preparing transfection mixture:
	<ol>
		<li>Prepare an appropriate cationic liposome reagent for plasmid transfection into the selected cell line.</li>
		<li>In this transfection example,&#160;use 1 &#181;g of total plasmid DNA per well (in a 6 well plate) to be transfected at a 1:3 ratio of HIVi&#916;Env plasmid to psPAX2 plasmid. For each well, dilute 1 &#181;g of DNA into 50 &#181;L of serum-free DMEM. Homogenize the mixture via repeatedly micropipetting or gentle swirling of the plate.</li>
		<li>For each well, dilute 3 &#181;L of transfection reagent in serum-free DMEM, then homogenize the mixture again by micropipetting or gentle swirling. Add the entire diluted transfection reagent mixture to the diluted DNA mixture (step 3.1.2) and incubate at room temperature (RT) for 15 min (Figure 1C1). 
		NOTE: While this mixture is incubating, replace the media in the wells with 1.5 mL of fresh DMEM. Be careful not to dislodge the grids from the bottom of the well.</li>
	</ol>
	</li>
	<li>To apply the transfection mixture, pipette 100 &#181;L of the mixture from step 3.1.3 dropwise to each well, concentrating the drops over the grids to ensure contact (Figure 1C2). Change the growth medium 16-24 h after transfection (Figure 1C3).</li>
</ol>
4. Plunge freezing
NOTE: Keep cells at 37 &#176;C until needed for blotting. Additionally, be sure to make a note of the carbon side of the grids throughout the process.
<ol>
	<li>Cell density
	<ol>
		<li>Check the grids in an inverted optical microscope and note cell densities on each grid. 
		NOTE: Grids with less than 0.25 cells per grid square (or one cell every 4 grid squares), are considered &#39;empty&#39;. Grids with 0.25-1 cells per grid square are considered &#39;normal&#39;. Grids with more than 1 cell per grid square are considered &#39;full&#39;.</li>
		<li>Take note of the cell density on each grid to allow the dialing of blotting times later during plunge-freezing. Add 2-3 mL of PBS to an empty plate/dish and warm to 37 &#176;C.</li>
	</ol>
	</li>
	<li>Preparing fiducials
	<ol>
		<li>To prepare gold fiducials for downstream cryotomography applications, pipette 50 &#181;L of 10 nm colloidal gold bead solution into a clean 1.5 mL tube. Add 2 &#181;L of 1 mg/mL BSA to the tube and homogenize by micropipetting repeatedly.</li>
		<li>Centrifuge the tube at 15,000-20,000 x g for 15 min. Micropipette to remove as much of the supernatant as possible without disturbing the pellet. 
		NOTE: The pellet will be very loose.</li>
		<li>Add 50 &#181;L of PBS to the pellet and resuspend. Centrifuge the tube again at 15,000-20,000 x g for 15 min.</li>
		<li>Using a 10 &#181;L tip, directly aspirate 4 &#181;L of the pellet, and transfer it to a clean 1.5 mL tube (Figure 1D1). Use 1 &#181;L of the washed and concentrated gold per grid. 
		NOTE: The type of fiducials used depends on downstream applications. For TEM tomography, use 10 nm gold beads. For soft X-ray tomography, use 200 nm gold beads. For correlation microscopy, use fluorescent latex beads. Fiducials are normally not necessary for experiments involving cryoFIB milling, as the milling process will destroy them.</li>
	</ol>
	</li>
	<li>Blotting
	<ol>
		<li>Set up the environmental chamber to 95% humidity and 30&#176;C (Figure 1D1). Select a grid using the 5/15 tweezers.</li>
		<li>Wash the grids with PBS and transfer them to plunging tweezers (Figure 1D2). Once secure, remove the 5/15 tweezers and slide the clamp on the plunging tweezers.</li>
		<li>Insert the grid into the plunge freezer, keeping the clamp secure (Figure 1D3). Add 1 &#181;L of gold fiducials to the back side of the grid (Figure 1D4). Add 2-3 &#181;L of PBS to the carbon side of the grid. Use automated blotting to blot the back side of the grid and plunge freeze into liquid ethane (Figure 1D5). 
		NOTE: It is recommended to have between 3-4 &#181;L of volume per grid for optimal blotting. Grids may retain a variable amount of PBS after washing. Adjust the added volume of PBS to account for pre-existing liquid retention on the grid. It is recommended that the gold fiducials be added to the back of the grid. Addition to the front can result in pools of gold fiducials near the point of insertion, whereas addition from the back results in a more homogenized solution. Blotting duration depends on the cell dentistry of the grid. 2 s for empty, 3 s for normal, and 4 s for full grids, respectively. The type of plunge freezer available may determine how blotting is approached: The Leica GP2 plunge freezer is the one used in this protocol and is capable of automated single-sided blotting by default. Thermo Fisher Vitrobot does automated double-sided blotting, though this often damages the cells attached to the carbon side. Manual blotting in the Vitrobot is also possible, as well as manual gravity plungers.</li>
	</ol>
	</li>
</ol>

Preparation of Electron Microscopy Grids for In Situ Cellular Cryotomography

<ol><li>Zhang, P., Mendonça, L. Analysis of Viruses in the Cellular Context by Electron Tomography. <a target="_blank" href="http://scholar.google.com/scholar?hl=en&safe=off&q=author%3aZhang%2C+P+%22Encyclopedia+of+Virology%22">Encyclopedia of Virology</a>. 1, Elsevier, Academic Press. 242-247 (2021).</li>
<li>Fäßler, F., Dimchev, G., Hodirnau, V. -V., Wan, W., Schur, F. K. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Cryo-electron+tomography+structure+of+Arp2%2F3+complex+in+cells+reveals+new+insights+into+the+branch+junction%5BTitle%5D)+AND+%22Nature+Communications%22%5BJournal%5D)">Cryo-electron tomography structure of Arp2/3 complex in cells reveals new insights into the branch junction.</a> Nature Communications. 11, 6437(2020).</li>
<li>McDowall, A. W., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Electron+microscopy+of+frozen+hydrated+sections+of+vitreous+ice+and+vitrified+biological+samples%5BTitle%5D)+AND+%22Journal+of+Microscopy%22%5BJournal%5D)">Electron microscopy of frozen hydrated sections of vitreous ice and vitrified biological samples.</a> Journal of Microscopy. 131 (Pt 1), 1-9 (1983).</li>
<li>Stewart, M., Vigers, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Electron+microscopy+of+frozen-hydrated+biological+material%5BTitle%5D)+AND+%22Nature%22%5BJournal%5D)">Electron microscopy of frozen-hydrated biological material.</a> Nature. 319 (6055), 631-636 (1986).</li>
<li>Gan, L., Jensen, G. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Electron+tomography+of+cells%5BTitle%5D)+AND+%22Quarterly+Reviews+of+Biophysics%22%5BJournal%5D)">Electron tomography of cells.</a> Quarterly Reviews of Biophysics. 45 (1), 27-56 (2012).</li>
<li>Yang, J. E., Larson, M. R., Sibert, B. S., Shrum, S., Wright, E. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((CorRelator%3A+Interactive+software+for+real-time+high+precision+cryo-correlative+light+and+electron+microscopy%5BTitle%5D)+AND+%22Journal+of+Structural+Biology%22%5BJournal%5D)">CorRelator: Interactive software for real-time high precision cryo-correlative light and electron microscopy.</a> Journal of Structural Biology. 213 (2), 107709(2021).</li>
<li>Mendonça, L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Correlative+multi-scale+cryo-imaging+unveils+SARS-CoV-2+assembly+and+egress%5BTitle%5D)+AND+%22Nature+Communications%22%5BJournal%5D)">Correlative multi-scale cryo-imaging unveils SARS-CoV-2 assembly and egress.</a> Nature Communications. 12 (1), 4629(2021).</li>
<li>Klumpe, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((A+modular+platform+for+automated+cryo-FIB+workflows%5BTitle%5D)+AND+%22eLife%22%5BJournal%5D)">A modular platform for automated cryo-FIB workflows.</a> eLife. 10, e70506(2021).</li>
<li>Wagner, F. R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Preparing+samples+from+whole+cells+using+focused-ion-beam+milling+for+cryo-electron+tomography%5BTitle%5D)+AND+%22Nature+Protocols%22%5BJournal%5D)">Preparing samples from whole cells using focused-ion-beam milling for cryo-electron tomography.</a> Nature Protocols. 15 (6), 2041-2070 (2020).</li>
<li>Klein, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((IFITM3+blocks+influenza+virus+entry+by+sorting+lipids+and+stabilizing+hemifusion%5BTitle%5D)+AND+%22Cell+Host+%26+Microbe%22%5BJournal%5D)">IFITM3 blocks influenza virus entry by sorting lipids and stabilizing hemifusion.</a> Cell Host & Microbe. 31 (4), 616.e20-633.e20 (2023).</li>
<li>Shah, P. N. M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Characterization+of+the+rotavirus+assembly+pathway+in+situ+using+cryoelectron+tomography%5BTitle%5D)+AND+%22Cell+Host+%26+Microbe%22%5BJournal%5D)">Characterization of the rotavirus assembly pathway in situ using cryoelectron tomography.</a> Cell Host & Microbe. 31 (4), 604.e4-615.e4 (2023).</li>
<li>Fäßler, F., Zens, B., Hauschild, R., Schur, F. K. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((3D+printed+cell+culture+grid+holders+for+improved+cellular+specimen+preparation+in+cryo-electron+microscopy%5BTitle%5D)+AND+%22Journal+of+Structural+Biology%22%5BJournal%5D)">3D printed cell culture grid holders for improved cellular specimen preparation in cryo-electron microscopy.</a> Journal of Structural Biology. 212 (3), 107633(2020).</li>
<li>Hoffmann, P. C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Electron+cryo-tomography+reveals+the+subcellular+architecture+of+growing+axons+in+human+brain+organoids%5BTitle%5D)+AND+%22eLife%22%5BJournal%5D)">Electron cryo-tomography reveals the subcellular architecture of growing axons in human brain organoids.</a> eLife. 10, e70269(2021).</li>
<li>Toro-Nahuelpan, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Tailoring+cryo-electron+microscopy+grids+by+photo-micropatterning+for+in-cell+structural+studies%5BTitle%5D)+AND+%22Nature+Methods%22%5BJournal%5D)">Tailoring cryo-electron microscopy grids by photo-micropatterning for in-cell structural studies.</a> Nature Methods. 17 (1), 50-54 (2020).</li>
</ol>

The authors declare no competing interests.

Here, we have provided an accessible, flexible, and reproducible protocol to seed cells on electron microscopy grids for in situ cryoelectron tomography applications. This method can be easily adapted to fit the needs of downstream applications and/or experimental requirements. In addition to great flexibility, we have described a workflow that optimizes and reduces common pitfalls in grid seeding, notably extensive damage to the carbon layer, low cell density, and poor structural integrity of thin cell projections.
Although the protocol described here does provide several alternatives, there are some critical steps that should be followed to optimize general outcomes. One of the biggest issues with grid cell seeding is the detachment and floating of grids from the well or microslide. Therefore, it is important to fully wet the grid with an adherent solution on both sides and prevent it from drying during the incubation period. If using 3D-printed grid holders, be aware that multiple changes of media to these holders have the potential to produce floating grids since the air trapped under the grid can force it out of the holder. 
Our choice of tweezers also improves grid quality in the way of providing a geometrically favorable way of manipulating the grids without extensive grid bending that would damage the carbon layer. Keeping the cells at 37 &#176;C for as long as possible before plunging reduces cell suffering and improves the number of thin imageable cells on the grid. Finally, blotting from the gold side will protect the cells from harsh mechanical forces that could lead to damage to fragile cellular structures.
While not included in this protocol, grid photo-micropatterning has been shown to increase the number of imageable cells by optimizing their attachment to the center of grid squares14. Finally, 3D-printed grid holders have recently been used to reduce grid damage by limiting direct grid manipulation12.
It may be important to note that this protocol is optimized for imaging thin edges and protrusions from cells for the application of cryotomography. We suggest troubleshooting a variety of conditions from our recommendations in the protocol to find the best outcome for the downstream applications of choice. Overall, this protocol provides a reliable yet versatile method of seeding cells onto grids that can be tweaked for specific needs.

In situ cellular cryotomography is a powerful technique allowing for the study of biologically-relevant structures in cells without chemical fixation. By attaching cells to EM grids and plunge-freezing the grids in a cryogen, objects of interest are frozen in their natural cellular contexts without the formation of crystalline ice from intracellular water1,2. Both chemical fixation and crystalline ice formation can disrupt the structures of relevant molecules, such as proteins and lipids, reducing the biological accuracy of images obtained using these techniques3,4. In tomography, grids are imaged at incremental angles using electron microscopy, and these images are then used to construct three-dimensional representations of the target region imaged5. In situ cryotomography can be used alongside other microscopy techniques for integrative and correlative imaging, such as cryofluorescence imaging, soft x-ray tomography, and cryoFIB/SEM (cryogenic Focused Ion Beam/Scanning Electron Microscopy)6,7,8,9,10,11. Integration of multiple techniques allows for more information to be obtained about a structure or process than any single microscopy technique could achieve.
Despite all of the benefits of in situ cellular cryotomography, sample preparation can prove to be challenging for a variety of reasons. Due to their fragility, forceful manipulation of electron microscopy grids can lead to damage, with the thin carbon layer in particular being delicate and prone to tearing, reducing the imageable area of the grids. Electron microscopy grids are also difficult to manipulate due to their small size and are prone to becoming detached from the surface of the wells or microslide used to grow cells. Manipulation of the grids within the wells or microslides can prove difficult due to the geometry of these. Improper preparation of the grids (e.g., allowing them to float) can lead to low cell density and reducing the number of potential imaging areas, especially when cells are not prone to attach to the grids themselves. For direct cellular cryotomography, cells must&#160;spread very thin, which can be disrupted for many reasons, including improper temperatures or rough handling of the grids.
Through a variety of optimizations, the techniques presented in this article are meant to handle these most common pitfalls which arise during the preparation of electron microscopy grids for cryotomography. The use of 5/15 angled tweezers allows for the manipulation of grids within well plates or microslides. A fibronectin solution applied to both sides of the grids prior to plating makes floating grids less likely, which is beneficial in ensuring that grids have adequate cell density and that the grids are less likely to be damaged due to manipulation. By keeping the grids incubated at 37&#176;C until just before plunge freezing, we also ensure that the cells are kept in a comfortable environment to prevent the cells from retracting their thin edges. Blotting the grids from the back side also prevents damage to the cells from mechanical force. Altogether, these measures increase the success rate of sample preparation for in situ cellular cryotomography studies, increasing the accessibility of this imaging approach.

<table><tbody><tr><td>10 nm colloidal gold bead solution</td><td>Sigma-Aldrich</td><td>741957</td><td></td></tr><tr><td>6 well multidish, 100/CS</td><td>Fisher Scientific</td><td>FB012927</td><td></td></tr><tr><td>Allegra V-15R Benchtop Centrifuge, IVD 120 V 60 Hz</td><td>Beckman-Coulter</td><td>C63125</td><td></td></tr><tr><td>Au G300F1 with R2/2 Quantifoil carbon</td><td>Quantifoil</td><td>TEM-G300F1-AU</td><td></td></tr><tr><td>Bovine serum albumin</td><td>MilliporeSigma</td><td>A9647</td><td></td></tr><tr><td>BRAND&amp;nbsp;counting chamber BLAUBRAND&amp;nbsp;Neubauer improved</td><td>Sigma-Aldrich</td><td>BR717805-1EA</td><td></td></tr><tr><td>DMi1 Inverted Microscope</td><td>Leica</td><td>22A00G119</td><td></td></tr><tr><td>Dulbecco&#039;s modified eagle&#039;s medium - high glucose, no glutamine</td><td>Gibco</td><td>11-960-044</td><td></td></tr><tr><td>Dumont 5/15 tweezer</td><td>Electron Microscopy Sciences</td><td>0103-5/15-PO</td><td></td></tr><tr><td>EM GP2</td><td>Leica</td><td>587085</td><td>Automated plunge freezer</td></tr><tr><td>Fetal Bovine Serum</td><td>Gibco</td><td>A5209</td><td></td></tr><tr><td>Fibronectin from bovine plasma, cell culture grade</td><td>MilliporeSigma</td><td>F1141</td><td></td></tr><tr><td>GenJet version II in vitro DNA transfection reagent</td><td>SignaGen Laboratories</td><td>SL100489</td><td></td></tr><tr><td>GlutaMAX I 100x</td><td>Fisher Scientific</td><td>35050061</td><td>Media supplement</td></tr><tr><td>Neslab EX-211 Heating Circulator</td><td>Neslab</td><td>Out of production</td><td>Water bath for media warming</td></tr><tr><td>Original Portable Pipet-Aid Pipette Controller</td><td>Drummond Scientific</td><td>4-000-100</td><td></td></tr><tr><td>PBS, pH 7.4</td><td>Gibco</td><td>10010023</td><td></td></tr><tr><td>Pelco easyGlow device</td><td>Pelco</td><td>91000S</td><td>Glow discharge device</td></tr><tr><td>Penicillin-Streptomycin</td><td>Sigma-Aldrich</td><td>P0781</td><td>Media supplement</td></tr><tr><td>Pipetman P1000, 100&amp;ndash;1000 &amp;micro;L, Metal Ejector</td><td>Gilson</td><td>F144059M</td><td></td></tr><tr><td>Pipetman P2, 0.2&amp;ndash;2 &amp;micro;L, Metal Ejector</td><td>Gilson</td><td>F144054M</td><td></td></tr><tr><td>Pipetman P20, 2&amp;ndash;20 &amp;micro;L, Metal Ejector</td><td>Gilson</td><td>F144056M</td><td></td></tr><tr><td>Whatman number 2 filter paper, 55 mm</td><td>Whatman</td><td>28455-041</td><td>Blotting paper</td></tr></tbody></table>

sample preparation for in situ cryotomography of mammalian cells

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

Following the cotransfection of HIViGFP&#916;Env and psPAX2, all of the grids had minimal tearing in the carbon layer. Grids were imaged using phase light microscopy and fluorescent light microscopy 24 h after incubation with the transfection reagent (Figure 2). Cells on both the mock grids and the co-transfected grids contained viable cells in multiple grid squares.
psPAX2 codes for all structural and enzymatic proteins of HIV-1 without any fluorescence tagging. HIViGFP&#916;Env is similar to psPAX2 but with codes for a GFP-tagged HIV Gag protein. Both plasmids are &#916;Envelope. The cotransfection results in native-like assembly and budding of fluorescent HIV-1 particles, making this a great system for CLEM studies of HIV in Biosafety Level 1 condition. The co-transfected grids showed a subset of cells exhibiting green fluorescence, indicating successful cotransfection. No cells on the mock grid exhibited fluorescence, further validating the cotransfection using HIViGFP&#916;Env and psPAX2. After viewing the grids using light-based microscopy, grids were plunge frozen and moved to long-term storage in liquid nitrogen.
Figure 3&#160;depicts results from grids produced using the same experimental method but utilizing slightly different plasmid constructs. U2OS-containing grids were co-transfected using different HIV clones (HIVmCherry&#916;Env, and NL4-3&#916;EnvGFP at a 1:6 ratio). Since a higher mass of fluorescently tagged plasmids were used, these grids enabled the observation of a larger number of transfected cells, providing an advantage while capturing images using cryoCLEM and cryoET. Using cryoCLEM, full grid atlases were generated for each grid using cryofluorescence microscopy to record the locations of all co-transfected cells. With the locations of cells known, cryoET was performed. A full low-magnification grid atlas was collected and overlaid with the fluorescent atlas collected at cryofluorescence (Figure 3A). Cryotomograms were collected at cellular sites capturing intricate details of the viral life cycle, including the assembly and budding of HIV from cells (Figure 3B).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/65697/65697fig01.jpg" /> 
Figure 1: Cell seeding on grids workflow. A schematic depicting the overall procedure to seed cells on cryoEM grids. The process is divided into four major steps, including (A) preparing grids in the wells for seeding, (B) adding the appropriate amount of cells to each well, (C) the optional transfection of cells for fluorescent imaging, and (D) the plunge freezing of grids to allow for vitrification of the sample. <a href="https://www.jove.com/files/ftp_upload/65697/65697fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/65697/65697fig02.jpg" /> 
Figure 2: Cotransfection of U2OS using HIViGFP&#916;Env and psPAX2. U2OS cells were co-transfected with GFP-containing HIViGFP&#916;Env and psPAX2 in a 1:3 ratio. Grids were imaged by phase contrast and fluorescence microscopy. Cells that are shown to have GFP expression indicate successful cotransfection. Scale bar: 500 &#181;m. Scale bar (insets): 100 &#181;m. <a href="https://www.jove.com/files/ftp_upload/65697/65697fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/65697/65697fig03.jpg" /> 
Figure 3: Potential downstream cryo-based methods. (A) A cryoCLEM image with co-transfected U2OS cells. Cells in green represent HIV-producing cells and are used to measure cotransfection success. Red puncta represent mCherry tagged HIV-1 Gag. Scale bar: 250 &#181;m. Scale bar (inset): 25 &#181;m (B) A cryoET image of multiple HIV particles budding from the plasma membrane of U2OS cells. Scale bar: 50 nm <a href="https://www.jove.com/files/ftp_upload/65697/65697fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Optimizing Grid Preparation for Enhanced Cryoelectron Tomography at JoVE.com

Author Spotlight: Optimizing Grid Preparation for Enhanced Cryoelectron Tomography 

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

Sample Preparation for In Situ Cryotomography of Mammalian Cells

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

sample-preparation-for-in-situ-cryotomography-of-mammalian-cells

Research

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Biochemistry

3.0K Views.  University of Minnesota. This method provides an accessible and flexible protocol for the preparation of electron microscopy (EM) grids for in situ cellular cryotomography and correlative light and electron microscopy (CLEM).

Video: Author Spotlight: Optimizing Grid Preparation for Enhanced Cryoelectron Tomography 

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

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

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

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

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 ...

Electron Cryotomography of Bacterial Cells

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

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

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

Biology

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 ...

Cryosectioning Method for Microdissection of Murine Colonic Mucosa

The colonic mucosal tissue provides a vital barrier to luminal antigens. This barrier is composed of a monolayer of simple columnar epithelial cells. The colonic epithelium is dynamically turned over and epithelial cells are generated in the stem cell containing crypts of Lieberk&#252;hn. Progenitor cells produced in the crypt-bases migrate toward the luminal surface, undergoing a process of cellular differentiation before being shed into the gut lumen. In order to study these processes at the molecular level, we have developed a simple method for the microdissection of two spatially distinct regions of the colonic mucosa; the proliferative crypt zone, and the differentiated surface epithelial cells. Our objective is to isolate specific crypt and surface epithelial cell populations from mouse colonic mucosa for the isolation of RNA and protein.

Here we describe a simple method for the isolation of spatially distinct murine colonic epithelial surface cells from the underlying crypt-base cells.

The colonic mucosal tissue provides a vital barrier to luminal antigens. This barrier is composed of a monolayer of simple columnar epithelial cells. The ...

Indirect Immunofluorescence on Frozen Sections of Mouse Mammary Gland

Indirect immunofluorescence is used to detect and locate proteins of interest in a tissue. The protocol presented here describes a complete and simple method for the immune detection of proteins, the mouse lactating mammary gland being taken as an example. A protocol for the preparation of the tissue samples, especially concerning the dissection of mouse mammary gland, tissue fixation and frozen tissue sectioning, are detailed. A standard protocol to perform indirect immunofluorescence, including an optional antigen retrieval step, is also presented. The observation of the labeled tissue sections as well as image acquisition and post-treatments are also stated. This procedure gives a full overview, from the collection of animal tissue to the cellular localization of a protein. Although this general method can be applied to other tissue samples, it should be adapted to each tissue/primary antibody couple studied.

The indirect immunofluorescence protocol described in this article allows the detection and the localization of proteins in the mouse mammary gland. A complete method is given to prepare mammary gland samples, to perform immunohistochemistry, to image the tissue sections by fluorescence microscopy, and to reconstruct images.

Indirect immunofluorescence is used to detect and locate proteins of interest in a tissue. The protocol presented here describes a complete and simple ...

Cryosectioning of Contiguous Regions of a Single Mouse Skeletal Muscle for Gene Expression and Histological Analyses

With this method, consecutive cryosections are collected to enable both microscopy applications for tissue histology and enrichment of RNA for gene expression using adjacent regions from a single mouse skeletal muscle. Typically, it is challenging to achieve adequate homogenization of small skeletal muscle samples because buffer volumes may be too low for efficient grinding applications, yet without sufficient mechanical disruption, the dense tissue architecture of muscle limits penetration of buffer reagents, ultimately causing low RNA yield. By following the protocol reported here, 30 &#956;m sections are collected and pooled allowing cryosectioning and subsequent needle homogenization to mechanically disrupt the muscle, increasing the surface area exposed for buffer penetration. The primary limitations of the technique are that it requires a cryostat, and it is relatively low throughput. However, high-quality RNA can be obtained from small samples of pooled muscle cryosections, making this method accessible for many different skeletal muscles and other tissues. Furthermore, this technique enables matched analyses (e.g., tissue histopathology and gene expression) from adjacent regions of a single skeletal muscle so that measurements can be directly compared across applications to reduce experimental uncertainty and to reduce replicative animal experiments necessary to source a small tissue for multiple applications.

Consecutive cryo-sections are collected to enable histological applications and enrichment of RNA for gene expression measurements using adjacent regions from a single mouse skeletal muscle. High-quality RNA is obtained from 20 - 30 mg of pooled cryosections and measurements are directly compared across applications.

With this method, consecutive cryosections are collected to enable both microscopy applications for tissue histology and enrichment of RNA for gene ...

Cryogenic Sample Loading into a Magic Angle Spinning Nuclear Magnetic Resonance Spectrometer that Preserves Cellular Viability

Dynamic nuclear polarization (DNP) can dramatically increase the sensitivity of magic angle spinning (MAS) nuclear magnetic resonance (NMR) spectroscopy. These sensitivity gains increase as temperatures decrease and are large enough to enable the study of molecules at very low concentrations at the operating temperatures (~100 K) of most commercial DNP-equipped NMR spectrometers. This leads to the possibility of in-cell structural biology on cryopreserved cells for macromolecules at their endogenous levels in their native environments. However, the freezing rates required for cellular cryopreservation are exceeded during typical sample handling for DNP MAS NMR and this results in loss of cellular integrity and viability. This article describes a detailed protocol for the preparation and cryogenic transfer of a frozen sample of mammalian cells into a MAS NMR spectrometer.

Presented here is a protocol for cryogenic transfer of frozen samples into the dynamic nuclear polarization (DNP) magic angle spinning (MAS) nuclear magnetic resonance (NMR) probe. The protocol includes directions for rotor storage prior to the experiment and directions for viability measurements before and after the experiment.

Dynamic nuclear polarization (DNP) can dramatically increase the sensitivity of magic angle spinning (MAS) nuclear magnetic resonance (NMR) spectroscopy. ...

A 3D Cartographic Description of the Cell by Cryo Soft X-ray Tomography

Imaging techniques are fundamental in order to understand cell organization and machinery in biological research and the related fields. Among these techniques, cryo soft X-ray tomography (SXT) allows imaging whole cryo-preserved cells in the water window X-ray energy range (284-543 eV), in which carbon structures have intrinsically higher absorption than water, allowing the 3D reconstruction of the linear absorption coefficient of the material contained in each voxel. Quantitative structural information at the level of whole cells up to 10 &#181;m thick is then achievable this way, with high throughput and spatial resolution down to 25-30 nm half-pitch. Cryo-SXT has proven itself relevant to current biomedical research, providing 3D information on cellular infection processes (virus, bacteria, or parasites), morphological changes due to diseases (such as recessive genetic diseases) and helping us understand drug action at the cellular level, or locating specific structures in the 3D cellular environment. In addition, by taking advantage of the tunable wavelength at synchrotron facilities, spectro-microscopy or its 3D counterpart, spectro-tomography, can also be used to image and quantify specific elements in the cell, such as calcium in biomineralization processes. Cryo-SXT provides complementary information to other biological imaging techniques such as electron microscopy, X-ray fluorescence or visible light fluorescence, and is generally used as a partner method for 2D or 3D correlative imaging at cryogenic conditions in order to link function, location, and morphology.

Here, a protocol describing the sample preparation and data collection steps required in cryo soft X-ray tomography (SXT) to image the ultrastructure of whole cryo-preserved cells at a resolution of 25 nm half pitch, is presented.

Imaging techniques are fundamental in order to understand cell organization and machinery in biological research and the related fields. Among these ...

Cryo-Structured Illumination Microscopic Data Collection from Cryogenically Preserved Cells

Three-dimensional (3D) structured illumination microscopy (SIM) allows imaging of fluorescently labelled cellular structures at higher resolution than conventional fluorescence microscopy. This super-resolution (SR) technique enables visualization of molecular processes in whole cells and has the potential to be used in conjunction with electron microscopy and X-ray tomography to correlate structural and functional information. A SIM microscope for cryogenically preserved samples (cryoSIM) has recently been commissioned at the correlative cryo-imaging beamline B24 at the UK synchrotron. 
It was designed specifically for 3D imaging of biological samples at cryogenic temperatures in a manner compatible with subsequent imaging of the same samples by X-ray microscopy methods such as cryo-soft X-ray tomography. This video article provides detailed methods and protocols for successful imaging using the cryoSIM. In addition to instructions on the operation of the cryoSIM microscope, recommendations have been included regarding the choice of samples, fluorophores, and parameter settings. The protocol is demonstrated in U2OS cell samples whose mitochondria and tubulin have been fluorescently labelled.

This protocol demonstrates how to image biological cryo-preserved samples using cryo-structured illumination microscopy. We demonstrate the methodology by imaging the cytoskeleton of U2OS cells.

Three-dimensional (3D) structured illumination microscopy (SIM) allows imaging of fluorescently labelled cellular structures at higher resolution than ...

Sample Preparation for In Situ Cryotomography of Mammalian Cells

Summary

Explore More Videos

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

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

Electron Cryotomography of Bacterial Cells

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

Cryosectioning Method for Microdissection of Murine Colonic Mucosa

Indirect Immunofluorescence on Frozen Sections of Mouse Mammary Gland

Cryosectioning of Contiguous Regions of a Single Mouse Skeletal Muscle for Gene Expression and Histological Analyses

Cryogenic Sample Loading into a Magic Angle Spinning Nuclear Magnetic Resonance Spectrometer that Preserves Cellular Viability

A 3D Cartographic Description of the Cell by Cryo Soft X-ray Tomography

Cryo-Structured Illumination Microscopic Data Collection from Cryogenically Preserved Cells