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

Summary

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

Abstract

Cryo‑electron tomography (cryo‑ET) enables 3D visualization of native cellular ultrastructure, but its application to thick tissues is limited by the lack of reliable vitrification and low‑throughput lamella preparation. Here, an end‑to‑end workflow is presented that integrates high‑pressure freezing (HPF; ~2100 bar; dissection‑to‑freezing time ≤90 s) using a “waffle” assembly with copper foil to pre‑thin tissue, cryo‑FIB lift‑out, and controlled thinning to 120–200 nm lamellae with over‑tilt compensation. Conductive coating is standardized (sputter‑Pt 30 mA/15 s; GIS cap 0.5–1.0 µm) to minimize curtaining. Automated, dose‑symmetric data acquisition (0°, ±3°, ±6°… to ±60–66°; step size of 2–3°; total dose 100–150 e⁻/Ų) is implemented via PACEtomo scripts, with the starting tilt angle derived from the milling angle and sample orientation. Processing with AreTomo 3 provides functions for motion correction, CTF estimation, alignment, and WBP reconstruction in near‑real time; IsoNet may optionally reduce missing‑wedge anisotropy. Subtomogram averaging (RELION 4.0) follows a pseudo‑subtomogram strategy with resolution reporting based on an FSC cutoff of 0.143. Demonstrated on mouse hippocampus, the workflow routinely produces well‑vitrified samples and uniform lamellae (target thickness 150 ± 40 nm) and achieves high‑throughput tilt‑series collection (typically 5–8 min/tilt series). This protocol offers a reproducible route for studying in situ structural biology directly from complex tissue environments.

Introduction

Cryo-electron tomography (cryo-ET) has emerged as a pivotal technique for visualizing macromolecular complexes and cellular architectures in their native, hydrated state^1,2,3,4,5. By reconstructing three-dimensional volumes from tilt-series of projection images, cryo-ET bridges the gap between cellular context and near-atomic resolution, offering unprecedented insights into structural biology in situ⁶. While cryo‑electron tomography workflows for cellular specimens⁷ (including cryo‑FIB‑thinned cells in suspensions and adherent cells) are now well‑established, their direct extension to thick, architecturally intact tissues remains highly challenging^8,9,10. This is especially true for high‑water‑content tissues such as brain¹¹, where reliable vitrification and the preparation of electron‑transparent lamellae (typically <200 nm) continue to pose persistent bottlenecks¹².

Achieving vitrification in tissues is challenging due to their high water content and limited heat transfer. While plunge freezing is effective for thin samples (≤20 µm)¹³, it fails for most tissues. High-pressure freezing (HPF) suppresses ice nucleation by applying high pressure, enabling vitrification of samples up to ~200 µm in theory. However, the efficiency of conventional HPF is often suboptimal for vitrifying tissues. To improve outcomes, recent efforts have employed strategies such as adding cryoprotectants¹⁴ and performing pre-sectioning^15,16 (e.g., using a vibratome) to reduce the sample thickness mechanically prior to freezing^11,17. While these approaches can enhance vitrification quality, they often entail prolonged ex vivo handling and the use of chemical additives that risk altering the native structures. To address these limitations, this study modified the “waffle” carrier strategy originally developed by Kelley et al.¹⁸ by incorporating a critical optimization: the brain tissue is physically compressed using two polished copper carriers prior to freezing, which pre-thins the sample while simultaneously securing it between a filmed grid and a copper foil. This design not only promotes rapid and uniform vitrification but also significantly enhances mechanical stability, thus effectively preventing grid breakage and sample loss during subsequent handling, establishing a robust foundation for obtaining intact, high-quality frozen tissue specimens.

Following vitrification, thinning remains a major barrier to both throughput and quality. As an early and widely adopted technique, cryo-ultramicrotomy enables numerous foundational in situ structural studies. Nevertheless, it frequently introduces compression and cutting artifacts^19,20 that compromise structural integrity. In contrast, cryo-focused ion beam (cryo-FIB) milling²¹ offers superior precision and minimal distortion^7,22. Initially, cryo‑FIB milling was applied directly to vitrified samples on grids to produce lamellae²³; however, this in situ approach is primarily effective for thin specimens^24,25 (e.g., cultured cells or peripheral tissue regions) and cannot access deeper structures within thick, bulky tissues. To overcome this depth limitation, lift-out strategies²⁶ were developed, enabling the site-specific extraction of regions of interest (ROIs) from the interior of samples. Subsequent methodological advances, including serial lift-out²⁷ and on-grid serialized sectioning (SOLIST)²⁸, further enhanced throughput, lamellar stability, and precision of 3D correlative targeting in complex tissues. More recently, innovations have continued to advance the field, including the six‑sided attachment method by Tang et al.²⁹, which enhances mechanical stabilization during lift‑out and thinning, as well as novel on‑grid milling approaches by Benjamin C. Creekmore et al.¹¹ that enable deep‑structure access while preserving native tissue context. Despite these advances, protocols still vary in efficiency, reproducibility, and adaptability across different tissue types. A critical gap persists in the establishment of a standardized, easily replicable, and broadly applicable workflow that integrates these optimizations into a coherent pipeline for diverse thick tissues.

This gap is addressed here by synthesizing key technical details from established methodologies and refining them into a generalized, end-to-end pipeline. This protocol not only employs the enhanced HPF-waffle method for robust vitrification but also incorporates a series of optimized steps in cryo-FIB lift-out and thinning, such as systematic over-tilt compensation, standardized conductive coating, and calibrated milling sequences, that collectively enhance lamella uniformity, reduce artifacts (e.g., curtaining), and improve throughput. Unlike protocols tailored to specific cell types or model organisms, this workflow is designed with generalizability at its core, providing a versatile framework adaptable to a wide range of thick, complex tissues, including those that are relatively soft and prone to compression, beyond the demonstrated mouse hippocampus.

Demonstrated in the challenging model of the mouse hippocampus, notable for its susceptibility to ice damage and structural complexity, this integrated and generalized workflow reliably produces high-quality lamellae (120–200 nm) and supports efficient tilt-series acquisition (5–8 min/series). This consolidation of best practices into a coherent, accessible, and widely applicable protocol provides a reproducible framework that lowers the technical barrier to implementing cryo-ET in thick tissue research across diverse biological and biomedical contexts.

Protocol

All procedures involving animal tissue complied with SUSTech IACUC (Protocol SUSTech‑JY202501019). The regents and the equipment used are listed in the Table of Materials.

1. Sample support preparation

1. HPF carrier preparation (see Figure 1A)
   1. Obtain a pair of 6 mm copper HPF carriers (type B). Ensure one of the carriers has a flat surface.
   2. Wet the flat surface of the carrier and proceed to grind. First, use 8000-grit sandpaper until the original machining marks are completely removed. Then, switch to 12000-grit sandpaper and continue grinding until scratches from the previous step are largely eliminated, resulting in a uniformly smooth finish. Figure 2 shows typical images of the carrier surface during the grinding and polishing process.
   3. Polish the ground surface using a soft metallographic cloth and W0.5 diamond paste: first, apply the paste to the cloth, then polish the surface in a circular motion for 1–2 min until a smooth, highly polished finish is achieved. Rinse the carrier thoroughly with ethanol, followed by distilled water to remove all abrasive residues.
   4. Submerge the polished carriers in 1-hexadecene for a minimum of 4 h (or overnight) prior to their use.
      NOTE: Polished and treated carriers can be stored submerged in 1-hexadecene.
      CAUTION: 1‑Hexadecene is flammable and harmful if inhaled; work in a fume hood with PPE.
2. TEM support grid preparation (see Figure 1B)
   1. Use 100-mesh copper grids with a continuous Formvar/carbon support film.
   2. Sputter-coat both sides of the grids with gold using a sputter coater: set the current to 50 mA and coat each side for 120 s to achieve a layer of approximately 6 nm per side.
3. Copper foil preparation (see Figure 1C)
   1. Obtain high-purity copper foil with a thickness of 1 µm.
      NOTE: The thermal conductivity of copper varies with purity, and lower purity results in lower thermal conductivity. Therefore, high-purity copper foil (Cu ≥ 99.9%) is required.
   2. Punch the copper foil into discs with a diameter of 3 mm using a circular hole punch.

2. High-pressure freezing

1. Place a 10 µL droplet of 10% sucrose solution on a clean glass surface.
2. Rapidly dissect the hippocampus from a freshly euthanized mouse. Excise small tissue pieces of approximately 0.5–1 mm³ using a sharp razor blade (see Figure 1D).
3. Transfer the tissue onto the copper support TEM Grids (100 mesh) within the HPF carrier using a toothpick or fine brush. Use 2-methylpentane to gently maneuver the tissue into place while minimizing mechanical stress on the grid.
   CAUTION: 2-Methylpentane is extremely flammable. Handle it in a fume hood with no ignition sources.
4. Cover the tissue and grid with a 3-mm-diameter copper foil disc. Complete the assembly by placing the second HPF carrier (flat face down) on top to form the sealed “waffle” sandwich (see Figure 1E).
   NOTE: The copper foil serves a critical mechanical function in the waffle assembly. It sandwiches the tissue between the TEM grid and the foil itself. During assembly, this configuration applies gentle pressure, helping to form a thinner, more uniform tissue slice ideal for vitrification. Furthermore, the foil prevents tissue loss during handling and helps distribute pressure during high-pressure freezing, reducing the risk of grid breakage. After freezing, the foil also facilitates easier separation of the carriers from the vitrified sample grid. However, this preparation method can introduce mechanical compression damage and may not be suitable for samples sensitive to pressure-induced artifacts. For such samples, alternative cutting techniques that achieve appropriate thickness without compression, such as vibratome sectioning or sample punching, should be considered.
5. Immediately initiate the high-pressure freezing procedure in accordance with the manufacturer's operational protocol. Set the target pressure to ~2100 bar and ensure the total time from tissue dissection to freezing does not exceed 90 s.
   ​NOTE: The entire procedure from dissection to freezing should be completed within 90 s to minimize structural alterations associated with prolonged ex vivo time. The tissue thickness range for brains is about 20–40 µm.

3. FIB milling and lift-out

NOTE: Perform the following steps using a dual-beam microscope equipped with a cryo-stage. Maintain the sample temperature below -170 °C throughout.

1. Preparation of the receiver grid
   1. Obtain a new 300-mesh gold grid to serve as the final receiver grid for the lift-out procedure.
   2. Under a stereomicroscope at room temperature, gently clip the grid into a compatible cryo-FIB AutoGrid ring.
   3. Store the assembled AutoGrid directly in a dry oven maintained at approximately 40 °C to minimize frost accumulation prior to its use.
      NOTE: During clipping ensure the grid lies flat and centered to prevent bending or misalignment.
2. Retrieval and mounting of the vitrified tissue grid
   1. While continuously immersing the assembly in liquid nitrogen (LN₂), carefully separate the vitrified TEM grid (containing the tissue and the copper foil) from the surrounding “waffle” assembly components.
      NOTE: The copper foil spacer typically makes the grid easy to separate from the carriers.
   2. On a pre-cooled cryo-loading workstation, mount the isolated tissue grid into a second cryo-FIB AutoGrid ring under LN₂ protection.
3. Co-loading grids onto the cryo-transfer shuttle
   1. Load both AutoGrids into the designated slots of the cryo-transfer shuttle.
   2. Orient each AutoGrid so that the milling notch faces outward and is vertical.
   3. Keep the loaded shuttle under LN₂ immersion or pre-cooled until transfer into the FIB/SEM microscope.
4. Localization and conductive coating in the FIB/SEM
   1. After cryo-transfer into the microscope, navigate to the tissue grid at low SEM magnification.
   2. Apply a conductive and protective coating to the entire grid surface.
      1. Sputter-coat the grid surface with platinum (Pt) at 30 mA for 15 s.
      2. Deposit a 0.5–1.0 µm thick organometallic Pt cap using the Gas Injection System (GIS).
      3. Apply a final sputter-coated Pt layer to the grid surface at 30 mA for 15 s.
         NOTE: The GIS cap is critical for reducing curtaining artifacts.
5. Tissue block isolation and lift-out (see Figure 3)
   1. Site selection and orientation
      1. Navigate to the selected region of interest (ROI) on the tissue grid using the SEM electron beam image.
      2. Orient the sample for front-side milling. A typical stage position is set to 110° rotation and 7° tilt, and this position presents the sample surface at an angle suitable for ion beam access.
         NOTE: In the absence of fluorescent markers a pre-screening strategy can be employed. This strategy aims to locate the target tissue region and exclude areas containing cryoprotectant or other non-tissue components. After selecting a region apparently free of surface ice, mill a small test trench using a higher beam current (e.g., 15 nA at 30 kV) at the standard front-side orientation and polish using a lower beam current (e.g., 1 nA at 30 kV). Switch to the electron beam to examine the cross section: the tissue exhibits excellent contrast under the electron beam, a characteristic that is particularly pronounced in membranous structures and regions rich in heavy elements; in contrast, the cryoprotectant typically shows uniform contrast under the electron beam. This allows for rapid verification before proceeding with the full lift-out procedure.
   2. Trench milling to isolate the block (Figure 3A)
      1. Define a rectangular trench surrounding the target tissue block using the ion beam operated at 30 kV.
      2. Begin milling at a certain distance from the block using a relatively high current of 15 nA to achieve rapid material removal.
      3. As the trench walls approach the block to within a few micrometers, reduce the ion beam current to 7 nA. Reducing the current achieves more precise control and minimizes ion beam damage and heating of the biological material in the region of interest.
      4. Leave one side of the block attached to the bulk material as a “tab” to hold it in place temporarily.
   3. Optional undercut milling for thick blocks (Figure 3B)
      1. For tissue blocks with an estimated thickness greater than 20 µm, perform an undercut milling step from the reverse side to facilitate the final lift-out.
      2. Rotate the stage (e.g., to -70°) and adjust the stage tilt (e.g., to 23°) to access the bottom of the block.
      3. Mill the underside connection using the FIB at 30 kV and a current of 3 nA.
   4. Attachment to the micromanipulator needle (Figure 3C)
      1. To attach the sample to the needle perform sequential cross-section milling patterns.
         NOTE: The gold block can be extracted from the bar of a 300-mesh gold grid.
   5. Lift-out of the tissue block (Figure 3D)
      1. Sever the remaining “tab” connecting the block to the bulk sample using the FIB (30 kV, 1 nA).
      2. Carefully maneuver the micromanipulator needle to lift the now-isolated tissue block so that it is completely clear of the trench and the surrounding original grid. Then, retract the needle and transport the block to the receiver grid.
   6. Block transfer, welding, and re-coating
      1. Navigate the needle to a pre-cooled 300-mesh gold receiver grid.
      2. Lower the block onto a chosen grid square. Bring the tissue block into contact with the film on the grid, then lift the block vertically to break the contact. Mill a ~3–4 µm thick slice from the block onto the grid film using the ion beam (30 kV, 1 nA) (Figure 3E).
         NOTE: If ice debris is present, pre-clean the attachment site with a rectangular pattern (30 kV, 50 pA). Make sure the block makes contact with the grid film to reduce electrostatic charge, which may randomly displace the slice.
      3. Weld the slice securely by milling using cleaning cross-section patterns on both sides of the junction (30 kV, 30 pA) (Figure 3F).
      4. Reapply the conductive/protective coating as described in Step 3.4.2 (Figure 3G).
         NOTE: For this GIS plating step the duration can be reduced depending on block size.
   7. Lamella thinning
      1. Rotate the stage (e.g., to -70°) and set an initial stage tilt (e.g., 15°) (Figure 3H).
         NOTE: The sample holder features a pre‑tilt of 45°.
      2. Define the final lamella position within a smooth area of the tissue block.
         NOTE: Confirm that the selected area is free of ice contamination.
      3. Perform systematic milling using the ion beam currents specified in Table 1. Progress is made from higher to lower currents as the target plane is approached (Figure 3I,J). Optionally, when within 1–2 µm of the lamella, a stress-relief trench may be milled approximately 1.5 µm away to prevent bending or curling of the lamella^30,31.
         ​NOTE: When the remaining material is within ~1 µm of the target plane limit the thickness removed per pass.

4. Cryo-ET data collection

1. Following successful lamella preparation, carefully load the grid into a transmission electron microscope (TEM) equipped with a cryo-holder, an energy filter, and tomography software (e.g., SerialEM).
2. Acquire an atlas map of the entire grid (e.g., Figure 4A; typical magnification is 155×).
3. Navigate to the lamella position and acquire a montage map (e.g., Figure 4B; typical magnification is 3600× with a defocus of -100 µm).
4. Set the nominal recording magnification and verify that the pixel size is appropriate for the intended resolution; in the present study, a pixel size of 2.109 Å was used. Adjust the corresponding illumination parameters, including spot size, C2 aperture diameter, and illumination area, to achieve an optimized dose rate for the direct electron detector.
5. Configure the focus and trial illumination conditions to match those set for the Record parameter.
6. Conduct data collection using the PaceTomo scripts³², following the steps below:
   1. Pre-select targets on the montage map using the "add points" function in the navigator, ensuring the first feature exhibits sufficient contrast.
   2. Select targets using the PACEtomo_selectTargets.py script³², adjusting the featured area at the preview magnification if necessary. Typical regions of interest are displayed in Figure 4C.
   3. Acquire tilt series using a dose-symmetric scheme (e.g., 0°, +3°, -3°, +6°, -6°... up to ±60°–66°) with an increment of 2° or 3°. Set acquisition parameters in the PACEtomo.py script: the starting tilt angle is determined by the milling angle specified in Step 3.7.1 (i.e., 8°), with its sign dependent on the sample loading orientation in the cryo-EM; the minimum and maximum tilt angles refer to absolute values; and the step size corresponds to the tilt increment (e.g., 3°).
      NOTE: This starting tilt angle is designed to position the lamella horizontally and perpendicular to the electron beam. If the milling angle is unknown, tilt the image to display the lamella at its largest apparent height; this provides a sufficiently accurate alignment.
   4. Maintain a cumulative electron dose of 100–150 e⁻/Ų distributed across all tilts. If available, use an energy filter with a slit width of 10 eV.
   5. Initiate data acquisition. Automated focusing and tracking are implemented at each tilt angle. The typical acquisition time for a single tilt series is 5–8 min.
      NOTE: A troubleshooting guide is provided in Table 2.

5. Data processing

1. Process Tilt-Series Motion Correction, CTF Estimation, Alignment, and 3D Tomogram Reconstruction images using AreTomo 3³³ to perform three core steps: (1) motion correction of raw image frames, (2) estimation of the contrast transfer function (CTF) for each tilt image, and (3) alignment of the full tilt-series.
   1. Following alignment reconstruct three-dimensional (3D) tomograms via the weighted back-projection algorithm. Required inputs for this step included: raw image frames, mdoc metadata files (which document tilt-series acquisition parameters), and the pre-determined rotation axis of the sample.
2. Missing wedge correction
   1. After reconstruction subject the generated tomograms to missing wedge correction using IsoNet³⁴.
3. Particle picking and curating
   1. Conduct particle picking in EMAN 2.9³⁵ using the template matching method. After automated picking, false positives (i.e., non-target regions or erroneously selected regions) were manually excluded to ensure the quality of the selected particle dataset.
4. Subtomogram averaging
   1. Import the tilt series and their corresponding particle coordinates into RELION 4.0. Perform subtomogram averaging following the standard workflow outlined in the RELION 4.0 documentation (https://relion.readthedocs.io/en/release-4.0/STA_tutorial/Introduction.html), which ensures adherence to established best practices for reproducibility.

Results

Successful application of this protocol enables the production of high-quality, vitrified tissue lamellae and the subsequent reconstruction of tomograms that reveal native cellular ultrastructure. The following outcomes, demonstrated using mouse hippocampal tissue, are representative of a successful experiment.

Tissue blocks processed by high-pressure freezing with the "waffle" method exhibited a uniform and electron-transparent appearance (Figure 4B,C

Discussion

The workflow presented herein establishes a generalized, optimized, and reproducible pipeline for in situ cryo-electron tomography of native, thick tissues. Its primary contribution lies not in the invention of singular techniques, but in the systematic integration, refinement, and detailed standardization of the entire process, from robust vitrification using a modified waffle-HPF method to a precisely calibrated cryo-FIB lift-out and thinning protocol. This synthesis addresses the critical reproducibility and ...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
1-Hexadecene	Sigma-Aldrich	H2131	Chemicals & Reagents
2-Methylpentane	Sigma-Aldrich	M65807	Chemicals & Reagents
Clip-ring	Thermo Fisher Scientific	1131750;1188953	Consumables
Copper foil	Fengcheng Metal Materials	1 µm thickness	Consumables
Copper support TEM Grids (100 mesh)	Beijing Zhongjingkeyi Technology Co., Ltd	BZ10021b	Consumables
Cryo-EM	Thermo Fisher Scientific	Krios G4	Equipment
Cryo-FIB	Thermo Fisher Scientific	Aquilos 2	Equipment
Cryo-FIB Autogrid	Thermo Fisher Scientific	1205101	Consumables
Grids (UltrAufoil R 1.2/1.3 300 mesh)	Quantifoil	AltrAufoil	Consumables
High-Pressure Freezer	Leica Microsystems	EM ICE	Equipment
HPF carrier, copper	Leica Microsystems	type B (6 mm)	Consumables
Ion Sputter Coater	Quorum	Q150RS plus	Equipment
Metallographic Polishing Paste	Lab-testec	W0.5	Consumables
Polishing cloth	GORAL	B00809	Consumables
Sandpaper	Eagle Brand Sandpaper	8000-grit, 12000-grit	Consumables
Stereomicroscope	Leica Microsystems	S9i	Equipment
Sucrose	Sigma-Aldrich	200-334-9	Chemicals & Reagents
Tilt-Series Acquisition Software	https://bio3d.colorado.edu/SerialEM/	SerialEM 4.0.9	Software
Tilt-Series Alignment & Reconstruction Software	https://github.com/czimaginginstitute/AreTomo3	AreTomo 3	Software
Tilt-Series Acquisition Software	https://github.com/eisfabian/PACEtomo	PaceTomo	Software
Tilt-Series Alignment & Reconstruction Software	https://github.com/IsoNet-cryoET/IsoNet	IsoNet	Software
Tilt-Series Alignment & Reconstruction Software	https://github.com/cryoem/eman2	EMAN2	Software
Tilt-Series Alignment & Reconstruction Software	https://relion.readthedocs.io/en/release-4.0/index.html	Relion 4	Software