Optimizing Sample Preparation for Cryogenic Electron Microscopy

This protocol presents a practical method using Methanocaldococcus jannaschii (MjsHSP16.5) as an example to address uneven particle distribution through sample preparation optimization, providing a reference for researchers to efficiently elucidate macromolecular structures using cryogenic electron microscopy (cryo-EM).

Cryogenic electron microscopy (cryo-EM) has revolutionized structural biology by enabling the study of macromolecular structures in near-native conditions, suspended in vitreous ice. This technique allows for the high-resolution visualization of proteins and other biomolecules without the need for crystallization, offering significant insights into their function and mechanism. Recent advancements in single-particle analysis, coupled with improved computational data processing, have made cryo-EM an indispensable tool in modern structural biology. Despite its growing adoption, cryo-EM faces persistent challenges that can limit its effectiveness, particularly uneven particle distribution. This issue often leads to poor resolution and reduced accuracy in reconstructed protein structures. This article outlines a simple, practical approach to address this challenge, using the small heat-shock protein from Methanocaldococcus jannaschii (MjsHSP16.5) as an example. The method optimizes sample preparation to minimize preferential adsorption, ensuring more homogeneous particle distribution and higher-quality protein cryo-EM structures. This technique offers valuable guidance for researchers aiming to overcome similar challenges in structural studies.

With recent advancements in both instrument hardware^1,2 and image processing software^3,4,5, cryogenic electron microscopy (cryo-EM) has emerged as a popular and powerful tool in modern structural biology. Despite these breakthroughs, bottlenecks persist in achieving high-resolution macromolecular structures using cryo-EM. One such significant challenge is the uneven particle distribution, including the orientation preference phenomenon, which is predominantly observed at the air-water interface^6,7,8,9.

During sample vitrification, some molecules exhibit a tendency to align themselves along specific axes on the grid. This leads to an uneven distribution of particle views in the final dataset. Certain orientations may be overrepresented while others are underrepresented or completely absent, resulting in incomplete sampling of the total protein architecture. Regions of the protein that are preferentially oriented towards the electron beam will appear more prominent in the density map, while regions oriented away from the beam may be poorly resolved or completely missing^10,11. Consequently, uneven particle distribution introduces potential biases and artifacts into the final reconstructed three-dimensional (3D) structure. Notably, key structural elements such as alpha helices and beta sheets may become skewed, amino acid or nucleotide chains may appear fragmented, and densities of specific protein or nucleic acid segments may exhibit distortion¹². Ultimately, these misrepresentations pose a major challenge to accurately unraveling the structure and function of biological molecules.

Various experimental approaches are currently used to overcome such challenges, including sample preparation optimization^13,14,15, grid treatment^{16,17,18,19,20,21,22}, and data collection strategy²³. Notably, it is advised to address the challenge at the sample preparation stage whenever feasible⁷. Common optimizations in sample preparation include modifying buffer composition, introducing small-molecular or macromolecular binding partners, generating intramolecular crosslinks, and varying detergents. This is also true for membrane proteins^24,25, although detergents must be used specifically for purification and stabilization purposes. Among these, the customizability, cost-effectiveness, and widespread accessibility of protein buffer optimization make it a preferred strategy in most laboratories. This approach allows precise and immediate adjustments of the various parameters to match the specific requirement of each protein sample. Through iterative refinement, researchers can systematically test diverse buffer conditions and adjust various parameters aimed at minimizing preferred orientations and improving the overall quality of cryo-EM data. Simply varying protein buffer components and adjusting their concentrations has demonstrated efficacy in influencing protein stability by modulating surface charge, consequently impacting protein behavior within vitreous ice²⁵. Therefore, optimizing protein buffer composition is considered one of the most convenient and straightforward approaches for addressing common challenges in cryo-EM.

Here, a protocol is suggested for addressing a common obstacle in cryo-EM—overcoming uneven particle distribution. In this protocol, key procedures for protein preparation and buffer screening, complemented by grid preparation, are outlined using a small heat shock protein from Methanocaldococcus jannaschii (MjsHSP16.5)²⁶ as a case study (Figure 1). This sHSP is natively stable, has a molecular mass of 16.5 kDa per monomer, and assembles into a 24-mer octahedral cage^26,27, making it an attractive candidate for structural analysis by cryo-EM. However, the observation of an uneven particle distribution during cryo-EM data collection was not anticipated, and it emerged as a significant challenge during the experiments. Furthermore, potential approaches beneficial for researchers tackling similar challenges are discussed, thus facilitating the efficient elucidation of macromolecular structures using cryo-EM.

The details of the reagents and the equipment used in this study are listed in the Table of Materials.

1. Protein purification

1. Induce expression of recombinant MjsHSP16.5 in E. coli BL21(DE3) and purify the protein using a nickel-chelating chromatography column as described previously²⁶. Following purification on a size-exclusion chromatography (SEC) column²⁶, examine protein purity by loading 5 µL of peak fractions onto a 12.5% SDS-PAGE gel and visualizing by standard Coomassie blue staining²⁸ (Figure 2).
2. Concentrate the pool fractions containing MjsHSP16.5 at 4 °C to approximately 65 mg/mL using centrifugal filters with a 50-kDa molecular weight cutoff, according to the manufacturer's instructions (see Table of Materials).
   NOTE: Gently pipette protein solution every 5 min during centrifugation to prevent protein aggregation.
3. After concentration, centrifuge the sample at 16,000 x g for 10 min at 4 °C to remove aggregates, aliquot the supernatant into 50 μL volumes in 0.2-mL thin-wall PCR tubes, flash freeze the tubes in liquid nitrogen, and store the samples at -80 °C until further use.

2. Protein preparation for transmission electron microscopy (TEM) imaging

1. Thaw two frozen protein tubes on ice. Once thawed, gently mix the solution by flicking the tube several times to ensure homogeneity and centrifuge the protein solution at 16,000 × g for 10 min at 4 °C to remove aggregates.
2. Inject the supernatant onto a SEC column and collect 0.5 mL elution fractions.
3. Collect the elution fractions corresponding to the peak exhibiting the highest UV absorbance at 280 nm and measure the protein concentration in these fractions using the Bradford assay²⁹.
   NOTE: To ensure sufficient protein concentration in a single peak fraction for the downstream applications, loading a high amount of protein sample onto the SEC column while staying within the column's recommended capacity is necessary.

3. Buffer exchange

1. Prepare the desired buffer solution (9 mM of MOPS-Tris (pH 7.2), 50 mM of NaCl, and 0.1 mM of EDTA), aliquot the buffers into 50-mL beakers, and keep the buffer beakers at 4 °C.
   NOTE: Based on prior knowledge of the target protein biochemistry, prepare a variety of buffer solutions with different compositions (buffer types, pH, and ionic strength), such as 20 mM of HEPES (pH 7.5), 100 mM of NaCl, and 0-5 mM of DTT, to identify the conditions that promote optimal protein solubility, homogeneity, and stability in downstream grid preparation.
2. Prepare the dialysis membrane following the manufacturer's instructions (see Table of Materials).
3. Cut the dialysis membrane into 30 mm x 30 mm pieces using scissors. Equilibrate these membranes by incubating them in distilled water or buffer solutions (Figure 3A).
4. Add 55 μL of the protein solution (approximately 2.5 mg/mL) to the chamber of 50-μL microdialysis buttons (Figure 3B).
5. Hold the dialysis membrane vertically using forceps and gently touch one edge of the membrane against tissue paper to drain excess liquid (Figure 3C). Carefully cover the microdialysis button with the membrane, ensuring no air bubbles are introduced into the microdialysis chamber (Figure 3D). Place the O-ring on top of the dialysis membrane. Gently roll the O-ring into the groove on the button's edge.
6. Alternative method: Place the O-ring along the axis of a golf tee and position the golf tee upside down on the membrane (Figure 3E). Gently slide the O-ring down the golf tee until it slips off onto the button's rim (Figure 3F). Carefully guide the O-ring into the groove on the button's edge (Figure 3G).
7. Submerge each dialysis button in a separate beaker containing the destination buffer with the membrane side facing upwards (Figure 3H).
8. Keep the beakers at 4 °C during the dialysis process. Change the dialysis buffer after 2 h and continue dialysis overnight.
9. Use a syringe with a fine needle (e.g., Hamilton syringe or insulin syringe) to carefully punch the membrane and recover the protein solution from the microdialysis chamber (Figure 3I). Store the protein sample in a microcentrifuge tube on ice until further use. Measure the protein concentration using the Bradford assay²⁹.

4. Grid preparation

1. Choose the appropriate grid type.
   NOTE: Amorphous holey carbon film on a copper support is commonly used. The carbon film thickness, hole size, and mesh number should be chosen based on the desired ice thickness. Holey carbon-coated grids were used for sample preparation of MjsHSP16.5.
2. Grip the grids gently at the edges using tweezers to avoid damaging the grid holes. Position the grids on a grid holder with the carbon side facing upwards. Place the grid holder into the chamber of the glow discharge system.
3. Perform glow discharge for 60 s at 15 mA to render the grid hydrophilic. Applying a negative charge is recommended during this process.
   NOTE: The recommended parameters serve as a starting point for the glow discharge system used in this study. Optimization might be necessary depending on the glow discharge instrument type and to achieve the desired level of hydrophilicity for specific grid materials. For plasma cleaning, consider testing alternative gases or gas mixtures such as hydrogen or argon-oxygen mixtures.
4. Optional step: introduce additional chemical solutions, such as 99% amylamine in an open glass bottle, into a chamber adjacent to the grid holder to specifically produce a net positive charge on the grid.
5. Use the discharged grids within 0.5-1 h to ensure optimal performance.

5. Negative stain grid preparation

1. Load 5 µL of the sample (50-300 nM or 20-120 ng/mL) onto the glow-discharged grid. Allow the sample to rest undisturbed on the grid surface for 1 min to facilitate the settling of proteins.
2. Prepare three water drops of 50 µL each on a paraffin film sheet. Gently wash the sample on the grid by rubbing the grid surface against the surface of the first water drop. Repeat the washing step with the remaining two water drops.
   NOTE: Washing times may be optimized according to the specific requirements of each experiment.
3. Load 5 µL of the filtered 1% (w/v) uranyl acetate solution onto the grid and immediately pipette away the uranyl acetate solution.
4. Load another 5 µL of the 1% (w/v) uranyl acetate solution onto the grid. Allow the grid to incubate in the uranyl acetate solution for 90 s for staining.
   NOTE: Staining time can be adjusted and optimized according to specific samples.
5. Gently touch the tweezer side of the grid with filter paper to remove excess staining solution. Allow the grid to air dry completely at room temperature. Store the dried grid in a grid container at room temperature until observation.

6. Sample vitrification

1. Dilute the protein sample to the desired concentration.
   NOTE: The protein concentration should be high enough to maximize the number of particles in each hole, but low enough to ensure the distribution of single particles. Cryo-EM protein concentrations typically range from 0.5 to 2 mg/mL. However, certain proteins, depending on their molecular weight and specific sample preparation conditions, may require concentrations outside this range.
2. Centrifuge the sample at 16,000 × g for 10 min at 4 °C to remove any aggregates.
3. Turn on the plunge-freezing instrument. Fill the humidifier reservoir with distilled water. Mount filter papers with rings on blot pads. Set the parameters for vitrification (temperature: 4 °C, humidity: 100%, blot time: 6 s, blot force: 5 units, wait time: 10 s).
   NOTE: Blot time and force are important parameters for the vitrification device optimization, as water removal from the grid and particle exposure to the air-water interface can induce dissociation or orientation bias.
4. Assemble the cryogen container components, including the brass cup, grid box holder, spider unit, and anti-contamination ring, into the base tray. Cool the container assembly by filling the tray with liquid nitrogen. Fill the brass cup with cryogen (liquid ethane). Remove the spider unit once the cryogen reaches melting temperature.
5. Mount the tweezers and grid assembly onto the instrument, load the sample (typically 4 µL) onto the carbon side of the grid, and initiate the plunge-freezing process.
6. Carefully undock the tweezers from the instrument, place the grid onto a storage grid box, and seal the box with the lid. Store the grid box containing the vitrified samples in liquid nitrogen.

7. Loading the grids to TEM

1. Assemble the autogrid assembly station and cool it with liquid nitrogen.
2. Place the grid box containing vitrified grids and unscrew the lid carefully to access the grids. Position an empty autogrid storage box nearby for easy transfer of assembled grids.
3. Place the autogrid ring in the designated cut-out space for the ring at the assembly station. Carefully move the vitrified grid from the grid box and fit it inside the autogrid ring. Ensure the grid and ring are aligned.
4. Align the disc to position the circle opening on top of the autogrid ring and grid assembly. To fix the grid into the autogrid ring, place the C-clip insertion tool on top of the grid and press down gently to click the C-clip inside.
5. Rotate the disc back and move the assembled grids into the autogrid storage box.
6. Assemble the cassette-loading station and cool the loading station with liquid nitrogen.
7. Place the autogrid storage box in the loading station. Move the grid assembly from the autogrid storage box to the cassette and tap the grid gently to ensure proper placement.
8. Use the handle to place the cassette into the autoloader capsule. Check the pin in the autoloader to confirm its free movement and ensure it is not frozen.
9. Dock the autoloader capsule to the TEM for grid observation.

To identify optimal grid conditions for MjsHSP16.5, an initial cryo-EM screening was conducted, primarily focusing on examining various protein buffer conditions: (1) the final purification buffer, which ensures the stability and homogeneity of MjsHSP16.5 and is important for its crystallization³⁰; (2) buffers adapted from conditions necessary for the growth of high-diffraction-quality MjsHSP16.5 crystals³⁰; and (3) buffers previously employed in electron microscopy (EM) st...

All protein structural studies begin with protein purification, an iterative process to achieve a balance between isolating protein targets with high purity and homogeneity while preserving their native functionality. Even though the buffer compositions to purify and preserve protein samples are carefully selected during the purification process, these buffers frequently pose challenges during subsequent cryo-EM sample preparation and imaging⁶. This problem arises from a mismatch between the buffe...

The authors have nothing to disclose.

We thank the Cooperative Center for Research Facilities (CCRF) (Sungkyunkwan University, Korea) for generously granting us access to their cryo-EM facility. This work was supported by the National Research Foundation of Korea (NRF) grant funded by the Korea government (MSIT) to K.K.K. (No. 2021M3A9I4022936). Use of cryo-EM facilities of NEXUS consortium was supported by a National Research Foundation of Korea grant RS-2024-00440289.