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., Mendon&#231;a, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Analysis+of+Viruses+in+the+Cellular+Context+by+Electron+Tomography.">Analysis of Viruses in the Cellular Context by Electron Tomography.</a> Encyclopedia of Virology. 1, 242-247 (2021).</li><li>F&#228;&#223;ler, F., Dimchev, G., Hodirnau, V. -. V., Wan, W., Schur, F. K. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cryo-electron+tomography+structure+of+Arp2/3+complex+in+cells+reveals+new+insights+into+the+branch+junction.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Electron+microscopy+of+frozen+hydrated+sections+of+vitreous+ice+and+vitrified+biological+samples.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Electron+microscopy+of+frozen-hydrated+biological+material.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Electron+tomography+of+cells.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=CorRelator:+Interactive+software+for+real-time+high+precision+cryo-correlative+light+and+electron+microscopy.">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>Mendon&#231;a, L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Correlative+multi-scale+cryo-imaging+unveils+SARS-CoV-2+assembly+and+egress.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+modular+platform+for+automated+cryo-FIB+workflows.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Preparing+samples+from+whole+cells+using+focused-ion-beam+milling+for+cryo-electron+tomography.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=IFITM3+blocks+influenza+virus+entry+by+sorting+lipids+and+stabilizing+hemifusion.">IFITM3 blocks influenza virus entry by sorting lipids and stabilizing hemifusion.</a> Cell Host &amp; 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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Characterization+of+the+rotavirus+assembly+pathway+in+situ+using+cryoelectron+tomography.">Characterization of the rotavirus assembly pathway in situ using cryoelectron tomography.</a> Cell Host &amp; Microbe. 31 (4), 604.e4-615.e4 (2023).</li><li>F&#228;&#223;ler, F., Zens, B., Hauschild, R., Schur, F. K. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=3D+printed+cell+culture+grid+holders+for+improved+cellular+specimen+preparation+in+cryo-electron+microscopy.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Electron+cryo-tomography+reveals+the+subcellular+architecture+of+growing+axons+in+human+brain+organoids.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Tailoring+cryo-electron+microscopy+grids+by+photo-micropatterning+for+in-cell+structural+studies.">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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<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&#160;counting chamber BLAUBRAND&#160;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&#39;s modified eagle&#39;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&#8211;1000 &#181;L, Metal Ejector</td>
			<td>Gilson</td>
			<td>F144059M</td>
			<td></td>
		</tr>
		<tr>
			<td>Pipetman P2, 0.2&#8211;2 &#181;L, Metal Ejector</td>
			<td>Gilson</td>
			<td>F144054M</td>
			<td></td>
		</tr>
		<tr>
			<td>Pipetman P20, 2&#8211;20 &#181;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

JoVE Journal

Biochemistry

2.8K 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 

Sample Preparation for Mass Spectrometry-based Identification of RNA-binding Regions

Noncoding RNAs play important roles in several nuclear processes, including regulating gene expression, chromatin structure, and DNA repair. In most cases, the action of noncoding RNAs is mediated by proteins whose functions are in turn regulated by these interactions with noncoding RNAs. Consistent with this, a growing number of proteins involved in nuclear functions have been reported to bind RNA and in a few cases the RNA-binding regions of these proteins have been mapped, often through laborious, candidate-based methods.
Here, we report a detailed protocol to perform a high-throughput, proteome-wide unbiased identification of RNA-binding proteins and their RNA-binding regions. The methodology relies on the incorporation of a photoreactive uridine analog in the cellular RNA, followed by UV-mediated protein-RNA crosslinking, and mass spectrometry analyses to reveal RNA-crosslinked peptides within the proteome. Although we describe the procedure for mouse embryonic stem cells, the protocol should be easily adapted to a variety of cultured cells.

We describe a protocol to identify RNA-binding proteins and map their RNA-binding regions in live cells using UV-mediated photocrosslinking and mass spectrometry.

Noncoding RNAs play important roles in several nuclear processes, including regulating gene expression, chromatin structure, and DNA repair. In most ...

Plate-based Large-scale Cultivation of Caenorhabditis elegans: Sample Preparation for the Study of Metabolic Alterations in Diabetes

Culturing Caenorhabditis elegans (C. elegans) in a large-scale manner on agar plates can be time-consuming and difficult. This protocol describes a simple and inexpensive method to obtain a large number of animals for the isolation of proteins to proceed with a western blot, mass spectrometry, or further proteomics analyses. Furthermore, an increase of nematode numbers for immunostainings and the integration of multiple analyses under the same culturing conditions can easily be achieved. Additionally, a transfer between plates with different experimental conditions is facilitated. Common techniques in plate culture involve the transfer of a single C. elegans using a platinum wire and the transfer of populated agar chunks using a scalpel. However, with increasing nematode numbers, these techniques become overly time-consuming. This protocol describes the large-scale culture of C. elegans including numerous steps to minimize the impact of the sample preparation on the physiology of the worm. Fluid and shear stress can alter the lifespan of and metabolic processes in C. elegans, thus requiring a detailed description of the critical steps in order to retrieve reliable and reproducible results. C. elegans is a model organism, consisting of neuronal cells for up to one-third, but lacking blood vessels, thus providing the possibility to investigate solely neuronal alterations independent of vascular control. Recently, early neurodegeneration in diabetic retinopathy was found prior to vascular alterations. Thus, C. elegans is of special interest for studying general mechanisms of diabetic complications. For example, an increased formation of advanced glycation end products (AGEs) and reactive oxygen species (ROS) is observed, which are reproducibly found in C. elegans. Protocols to handle samples of adequate size for a broader spectrum of investigations are presented here, exemplified by the study of diabetes-induced biochemical alterations. In general, this protocol can be useful for studies requiring large C. elegans numbers and in which liquid culture is not suitable.

Plate-based Large-scale Cultivation of Caenorhabditis elegans: Sample Preparation for the Study of Metabolic Alterations in Diabetes

This protocol describes a method for the large-scale cultivation of Caenorhabditis elegans on solid media. As an alternative to liquid culture, this protocol allows obtaining parameters of different scales under plate-based cultivation. This increases the comparability of results by omitting the morphological and metabolic differences between liquid and solid media culture.

Culturing Caenorhabditis elegans (C. elegans) in a large-scale manner on agar plates can be time-consuming and difficult. This protocol describes a simple ...

An All-in-one Sample Holder for Macromolecular X-ray Crystallography with Minimal Background Scattering

Macromolecular X-ray crystallography (MX) is the most prominent method to obtain high-resolution three-dimensional knowledge of biological macromolecules. A prerequisite for the method is that highly ordered crystalline specimen need to be grown from the macromolecule to be studied, which then need to be prepared for the diffraction experiment. This preparation procedure typically involves removal of the crystal from the solution, in which it was grown, soaking of the crystal in ligand solution or cryo-protectant solution and then immobilizing the crystal on a mount suitable for the experiment. A serious problem for this procedure is that macromolecular crystals are often mechanically unstable and rather fragile. Consequently, the handling of such fragile crystals can easily become a bottleneck in a structure determination attempt. Any mechanical force applied to such delicate crystals may disturb the regular packing of the molecules and may lead to a loss of diffraction power of the crystals. Here, we present a novel all-in-one sample holder, which has been developed in order to minimize the handling steps of crystals and hence to maximize the success rate of the structure determination experiment. The sample holder supports the setup of crystal drops by replacing the commonly used microscope cover slips. Further, it allows in-place crystal manipulation such as ligand soaking, cryo-protection and complex formation without any opening of the crystallization cavity and without crystal handling. Finally, the sample holder has been designed in order to enable the collection of in situ X-ray diffraction data at both, ambient and cryogenic temperature. By using this sample holder, the chances to damage the crystal on its way from crystallization to diffraction data collection are considerably reduced since direct crystal handling is no longer required.

A novel sample holder for macromolecular X-ray crystallography along with a suitable handling protocol is presented. The system allows crystal growth, crystal soaking and in situ diffraction data collection at both, ambient and cryogenic temperature without the need of any crystal manipulation or mounting.

Macromolecular X-ray crystallography (MX) is the most prominent method to obtain high-resolution three-dimensional knowledge of biological macromolecules. ...

A Plasma Sample Preparation for Mass Spectrometry using an Automated Workstation

Sample preparation for mass spectrometry analysis in proteomics requires enzymatic cleavage of proteins into a peptide mixture. This process involves numerous incubation and liquid transfer steps in order to achieve denaturation, reduction, alkylation, and cleavage. Adapting this workflow onto an automated workstation can increase efficiency and reduce coefficients of variance, thereby providing more reliable data for statistical comparisons between sample types. We previously described an automated proteomic sample preparation workflow1. Here, we report the development of a more efficient and better controlled workflow with the following advantages: 1) The number of liquid transfer steps is reduced from nine to six by combining reagents; 2) Pipetting time is reduced by selective tip pipetting using a 96-position pipetting head with multiple channels; 3) Potential throughput is increased by the availability of up to 45 deck positions; 4) Complete enclosure of the system provides improved temperature and environmental control and reduces the potential for contamination of samples or reagents; and 5) The addition of stable isotope labeled peptides, as well as &#946;-galactosidase protein, to each sample makes monitoring and quality control possible throughout the entire process. These hardware and process improvements provide good reproducibility and improve intra-assay and inter-assay precision (CV of less than 20%) for LC-MS based protein and peptide quantification. The entire workflow for digesting 96 samples in a 96-well plate can be completed in approximately 5 hours.

Here, we present an automated plasma protein digestion method for mass spectrometry-based quantitative proteomic analysis. In this protocol, the liquid transferring and incubation steps for protein denaturation, reduction, alkylation, and trypsin digestion reactions are streamlined and automated. It takes approximately five hours to prepare a 96-well plate with desired precision.

Sample preparation for mass spectrometry analysis in proteomics requires enzymatic cleavage of proteins into a peptide mixture. This process involves ...

Sample Preparation for Probe Electrospray Ionization Mass Spectrometry

Mass spectrometry (MS) is a powerful tool in analytical chemistry because it provides very accurate information about molecules, such as mass-to-charge ratios (m/z), which are useful to deduce molecular weights and structures. While it is essentially a destructive analytical method, recent advancements in the ambient ionization technique have enabled us to acquire data while leaving tissue in a relatively intact state in terms of integrity. Probe electrospray ionization (PESI) is a so-called direct method because it does not require complex and time-consuming pretreatment of samples. A fine needle serves as a sample picker, as well as an ionization emitter. Based on the very sharp and fine property of the probe tip, destruction of the samples is minimal, allowing us to acquire the real-time molecular information from living things in situ. Herein, we introduce three applications of PESI-MS technique that will be useful for biomedical research and development. One involves the application to solid tissue, which is the basic application of this technique for the medical diagnosis. As this technique requires only 10 mg of the sample, it may be very useful in the routine clinical settings. The second application is for in vitro medical diagnostics where human blood serum is measured. The ability to measure fluid samples is also valuable in various biological experiments where a sufficient volume of sample for conventional analytical techniques cannot be provided. The third application leans toward the direct application of probe needles in living animals, where we can obtain real-time dynamics of metabolites or drugs in specific organs. In each application, we can infer the molecules that have been detected by MS or use artificial intelligence to obtain a medical diagnosis.

This article introduces sample preparation methods for a unique real-time analytical method based on the ambient mass spectrometry. This method lets us perform real-time analysis of the biological molecules in vivo without any special pretreatments.

Mass spectrometry (MS) is a powerful tool in analytical chemistry because it provides very accurate information about molecules, such as mass-to-charge ...

TMT Sample Preparation for Proteomics Facility Submission and Subsequent Data Analysis

Proteomic technologies are powerful methodologies that can aid our understanding of mechanisms of action in biological systems by providing a global view of the impact of a disease, treatment, or other condition on the proteome as a whole. This report provides a detailed protocol for the extraction, quantification, precipitation, digestion, labeling, and subsequent data analysis of protein samples. Our optimized TMT labeling protocol requires a lower tag-label concentration and achieves consistently reliable data. We have used this protocol to evaluate protein expression profiles in a variety of mouse tissues (i.e., heart, skeletal muscle, and brain) as well as cells cultured in vitro. In addition, we demonstrate how to evaluate thousands of proteins from the resulting dataset.

We present an optimized tandem mass tag (TMT) labeling protocol that includes detailed information for each of the following steps: protein extraction, quantification, precipitation, digestion, labeling, submission to a proteomics facility, and data analyses.

Proteomic technologies are powerful methodologies that can aid our understanding of mechanisms of action in biological systems by providing a global view ...

Plant Sample Preparation for Nucleoside/Nucleotide Content Measurement with An HPLC-MS/MS

Nucleosides/nucleotides are building blocks of nucleic acids, parts of cosubstrates and coenzymes, cell signaling molecules, and energy carriers, which are involved in many cell activities. Here, we describe a rapid and reliable method for the absolute qualification of nucleoside/nucleotide contents in plants. Briefly, 100 mg of homogenized plant material was extracted with 1 mL of extraction buffer (methanol, acetonitrile, and water at a ratio of 2:2:1). Later, the sample was concentrated five times in a freeze dryer and then injected into an HPLC-MS/MS. Nucleotides were separated on a porous graphitic carbon (PGC) column and nucleosides were separated on a C18 column. The mass transitions of each nucleoside and nucleotide were monitored by mass spectrometry. The contents of the nucleosides and nucleotides were quantified against their external standards (ESTDs). Using this method, therefore, researchers can easily quantify nucleosides/nucleotides in different plants.

A precise and reproducible method for in vivo nucleosides/nucleotides quantification in plants is described here. This method employs an HPLC-MS/MS.

Nucleosides/nucleotides are building blocks of nucleic acids, parts of cosubstrates and coenzymes, cell signaling molecules, and energy carriers, which ...

A Sample Preparation Pipeline for Microcrystals at the VMXm Beamline

The mounting of microcrystals (&#60;10 &#181;m) for single crystal cryo-crystallography presents a non-trivial challenge. Improvements in data quality have been seen for microcrystals with the development of beamline optics, beam stability and variable beam size focusing from submicron to microns, such as at the VMXm beamline at Diamond Light Source1. Further improvements in data quality will be gained through improvements in sample environment and sample preparation. Microcrystals inherently generate weaker diffraction, therefore improving the signal-to-noise is key to collecting quality X-ray diffraction data and will predominantly come from reductions in background noise. Major sources of X-ray background noise in a diffraction experiment are from their interaction with the air path before and after the sample, excess crystallization solution surrounding the sample, the presence of crystalline ice and scatter from any other beamline instrumentation or X-ray windows. The VMXm beamline comprises instrumentation and a sample preparation protocol to reduce all these sources of noise.
Firstly, an in-vacuum sample environment at VMXm removes the air path between X-ray source and sample. Next, sample preparation protocols for macromolecular crystallography at VMXm utilize a number of processes and tools adapted from cryoTEM. These include copper grids with holey carbon support films, automated blotting and plunge cooling robotics making use of liquid ethane. These tools enable the preparation of hundreds of microcrystals on a single cryoTEM grid with minimal surrounding liquid on a low-noise support. They also minimize the formation of crystalline ice from any remaining liquid surrounding the crystals. 
We present the process for preparing and assessing the quality of soluble protein microcrystals using visible light and scanning electron microscopy before mounting the samples on the VMXm beamline for X-ray diffraction experiments. We will also provide examples of good quality samples as well as those which require further optimization and strategies to do so.

The signal-to-noise ratio of data is one of the most important considerations in performing X-ray diffraction measurements from microcrystals. The VMXm beamline provides a low-noise environment and microbeam for such experiments. Here, we describe sample preparation methods for mounting and cooling microcrystals for VMXm and other microfocus macromolecular crystallography beamlines.

The mounting of microcrystals (&#60;10 &#181;m) for single crystal cryo-crystallography presents a non-trivial challenge. Improvements in data quality ...

RIBO-seq in Bacteria: a Sample Collection and Library Preparation Protocol for NGS Sequencing

The ribosome profiling technique (RIBO-seq) is currently the most effective tool for studying the process of protein synthesis in vivo. The advantage of this method, in comparison to other approaches, is its ability to monitor translation by precisely mapping the position and number of ribosomes on a mRNA transcript.
In this article, we describe the consecutive stages of sample collection and preparation for RIBO-seq method in bacteria, highlighting the details relevant to the planning and execution of the experiment.
Since the RIBO-seq relies on intact ribosomes and related mRNAs, the key step is rapid inhibition of translation and adequate disintegration of cells. Thus, we suggest filtration and flash-freezing in liquid nitrogen for cell harvesting with an optional pretreatment with chloramphenicol to arrest translation in bacteria. For the disintegration, we propose grinding frozen cells with mortar and pestle in the presence of aluminum oxide to mechanically disrupt the cell wall. In this protocol, sucrose cushion or a sucrose gradient ultracentrifugation for monosome purification is not required. Instead, mRNA separation using polyacrylamide gel electrophoresis (PAGE) followed by the ribosomal footprint excision (28-30 nt band) is applied and provides satisfactory results. This largely simplifies the method as well as reduces the time and equipment requirements for the procedure. For library preparation, we recommend using the commercially available small RNA kit for Illumina sequencing from New England Biolabs, following manufacturer&#39;s guidelines with some degree of optimization.
The resulting cDNA libraries present appropriate quantity and quality required for next generation sequencing (NGS). Sequencing of the libraries prepared according to the described protocol results in 2 to 10 mln uniquely mapped reads per sample providing sufficient data for comprehensive bioinformatic analysis. The protocol we present is quick and relatively easy and can be performed with standard laboratory equipment.

Here we describe the stages of sample collection and preparation for RIBO-seq in bacteria. Sequencing of the libraries prepared according to these guidelines results in sufficient data for comprehensive bioinformatic analysis. The protocol we present is simple, uses standard laboratory equipment and takes seven days from lysis to obtaining libraries.

The ribosome profiling technique (RIBO-seq) is currently the most effective tool for studying the process of protein synthesis in vivo. The advantage of ...

Density Gradient Ultracentrifugation for Investigating Endocytic Recycling in Mammalian Cells

Endosomal trafficking is an essential cellular process that regulates a broad range of biological events. Proteins are internalized from the plasma membrane and then transported to the early endosomes. The internalized proteins could be transited to the lysosome for degradation or recycled back to the plasma membrane. A robust endocytic recycling pathway is required to balance the removal of membrane materials from endocytosis. Various proteins are reported to regulate the pathway, including ADP-ribosylation factor 6 (ARF6). Density gradient ultracentrifugation is a classical method for cell fractionation. After the centrifugation, organelles are sedimented at their isopycnic surface. The fractions are collected and used for other downstream applications. Described here is a protocol to obtain a recycling endosome-containing fraction from transfected mammalian cells using density gradient ultracentrifugation. The isolated fractions were subjected to standard Western blotting for analyzing their protein contents. By employing this method, we identified that the plasma membrane targeting of engulfment and cell motility 1 (ELMO1), a Ras-related C3 botulinum toxin substrate 1 (Rac1) guanine nucleotide exchange factor, is through ARF6-mediated endocytic recycling.

This paper aims to present a protocol for preparing recycling endosomes from mammalian cells using sucrose density gradient ultracentrifugation.

Endosomal trafficking is an essential cellular process that regulates a broad range of biological events. Proteins are internalized from the plasma ...