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

This protocol outlines the preparation of vitreous lamellae of Plasmodium-falciparum-infected red blood cells for cryo-electron tomography. This enables direct observation of detailed structural information about how the parasite causes malaria. Cryo-electron tomography records high resolution information about the insides of cells.

It's powerful enough to observe single protein complexes in situ, which can be computationally extracted and averaged to reveal three-dimensional structural information about proteins. This protocol can be easily adapted for different cell types and can be a starting point for someone new to producing vitreous lamellae for cryo-electron tomography. This is Alejandra Carbajal, who will be carrying out the protocol today.

To begin, screen the frozen copper grids using a CryoStage for a light microscope with particular attention to the gradient of ice across the grid. Check individual grid squares for cell coverage in the thinner areas of ice. Use a black indelible marker to mark the front and the opposite side of Cryo-FIB-SEM-specific Autogrid rims.

Load the clipped grid into the Cryo-FIB-SEM shuttle with the carbon side up, and apply a four nanometer thick carbon or platinum e-beam rotary coat to the surface of the cells. Load the shuttle into the Cryo-FIB-SEM and assess the cell distribution on each grid by SEM at five kilovolts. Take a low magnification overview at 100 power to look at ice gradients across the grid.

Then take higher magnification images at 5, 000 power to look at individual grid squares. Apply a two micrometer organoplatinum coat to the surface of each grid by inserting the gas injection system needle into the chamber above the grid and warming the organoplatinum source to a set temperature for a set time to produce a flow of vapor. To observe the sample, tilt the sample so that the plane of the grid is approximately 10 degrees from the incidence angle of the ion beam, and move the center of a suitable grid square to be seen in both the SEM and FIB images.

using the ion beam at low current, Survey the grid at a high magnification to visualize the cells at the center of a grid square. Mark out two rectangular patterns to mill, one above and one below a three-micrometer-thick protected region. Then choose the beam settings and correct astigmatism by adjusting the brightness and contrast.

after the setup is complete, start milling at a current of 300 picoamperes while monitoring the live progress in the ion beam view and intermittently by SEM. The sample is checked In the SEM view, and after the first milling, If the ion beam has not broken through the sample above and below the protected region, remove more material by increasing the height of the rectangular patterns. Stop milling when the surface above and below the lamella is completely smooth in the ion beam view.

Repeat the milling process stepwise, reducing the ion beam current each time until a thickness of 300 nanometers is reached. Record the X Y Z position of each lamella. To polish the lamellae at the end of the experiment, use a low magnification SEM overview to select a group of lamella to be polished and mark the polishing root.

Once the polishing root is marked, reduce the space between the two rectangular patterns to 100 to 200 nanometers and begin the polishing at an ion beam current of 30 picoamperes. Monitor the progress by SEM at two to three kilovolts. Stop polishing when the contrast in the lamella is lost by SEM or when the organoplatinum coat or the lamella itself begins to lose integrity.

Acquire and save low-magnification SEM images of each lamella. After completion of the experiment, remove the shuttle from the microscope. Remove the grids with lamellae and store them under liquid nitrogen.

Handle the grids with care. To study morphology, the schizonts were stalled at different stages of egress using compound two in E-64 inhibitors. In the presence of compound two, the boundary between the parasitophorous membranes and the surrounding hemoglobin in the host red blood cell was well-defined.

The intact parasitophorous vacuole, or PVs, were packed with merozoites in a single cluster of hemozoin crystals. When compound two synchronized schizonts were washed into E-64, the PV membrane was ruptured and the merozoites spread out within the RBC. A typical example of good cell distribution on a copper grid shows large red blood cells with a well-defined periphery and an intact vacuole membrane.

Within a partially collapsed host red blood cell, the individual merozoites were visualized as small cell clusters. Each schizont had a black spot at its center indicating the position of the hemozoin crystals. After milling, the grids were loaded into the TEM to identify the positions of the lamellae and find suitable areas to target by tomography.

In the sample micrograph, an RBC that had recently been invaded by a merozoite can be seen. The merozoite surrounded by the host-cell-derived membrane could be seen within the boundary of the RBC. At the apical end of the merozoite, the two membranes surrounding the cell and four membranes stacked in the apex of the merozoite associated with an electron-dense mushroom-shaped feature were clearly visible.

Within the cytoplasm of the merozoite, a fusion event between the merozoite plasma membrane and a multilayered vesicle was identified. In an additional TEM analysis of a 230-nanometer-thick lamella, the typical morphology of a mature merozoite treated with E-64 was observed. Within the red blood cell membrane, the merozoite plasma membrane can be seen with organelles like the inner membrane complex, polar rings, micronemes, rhoptries, and the nuclear envelope.

The most important part of this technique is optimizing the grids to enable reproducible and efficient production of thin lamellae. Further development of this technique will enable researchers to visualize single-protein molecules within malarial parasites and determine their role in malarial egress.