Published: December 15th, 2023
This method provides an accessible and flexible protocol for the preparation of electron microscopy (EM) grids for in situ cellular cryotomography and correlative light and electron microscopy (CLEM).
In situ cellular cryotomography is a powerful technique for studying complex objects in their native frozen-hydrated cellular context, making it highly 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.
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 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°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.
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.
2. Seeding grids
4. Plunge freezing
NOTE: Keep cells at 37 °C until needed for blotting. Additionally, be sure to make a note of the carbon side of the grids throughout the process.
Following the cotransfection of HIViGFPΔEnv 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. HIViGFPΔEnv is similar to psPAX2 but with codes for a GFP-tagged HIV Gag protein. Both plasmids are ΔEnvelope. 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 HIViGFPΔEnv 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 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 (HIVmCherryΔEnv, and NL4-3ΔEnvGFP 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).
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. Please click here to view a larger version of this figure.
Figure 2: Cotransfection of U2OS using HIViGFPΔEnv and psPAX2. U2OS cells were co-transfected with GFP-containing HIViGFPΔEnv 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 µm. Scale bar (insets): 100 µm. Please click here to view a larger version of this figure.
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 µm. Scale bar (inset): 25 µm (B) A cryoET image of multiple HIV particles budding from the plasma membrane of U2OS cells. Scale bar: 50 nm Please click here to view a larger version of this figure.
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 °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.
The authors declare no competing interests.
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.
|10 nm colloidal gold bead solution
|6 well multidish, 100/CS
|Allegra V-15R Benchtop Centrifuge, IVD 120 V 60 Hz
|Au G300F1 with R2/2 Quantifoil carbon
|Bovine serum albumin
|BRAND counting chamber BLAUBRAND Neubauer improved
|DMi1 Inverted Microscope
|Dulbecco's modified eagle's medium - high glucose, no glutamine
|Dumont 5/15 tweezer
|Electron Microscopy Sciences
|Automated plunge freezer
|Fetal Bovine Serum
|Fibronectin from bovine plasma, cell culture grade
|GenJet version II in vitro DNA transfection reagent
|GlutaMAX I 100x
|Neslab EX-211 Heating Circulator
|Out of production
|Water bath for media warming
|Original Portable Pipet-Aid Pipette Controller
|PBS, pH 7.4
|Pelco easyGlow device
|Glow discharge device
|Pipetman P1000, 100–1000 µL, Metal Ejector
|Pipetman P2, 0.2–2 µL, Metal Ejector
|Pipetman P20, 2–20 µL, Metal Ejector
|Whatman number 2 filter paper, 55 mm
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