Using HBmito Crimson to Observe Mitochondrial Cristae Through STED Microscopy

Materials and reagents

Biological materials

1. COS7 cell (ATCC, CRL-1651)

Reagents

1. Dulbecco’s modified Eagle’s medium (DMEM) (Gibco, catalog number: 11965-092), store at 4 °C

2. Fetal bovine serum (FBS) (Gibco, catalog number: 10091148), store at -20 °C

3. Penicillin/streptomycin solution (Gibco, catalog number: 15140122), store at -20 °C

4. Trypsin-EDTA solution (Gibco, catalog number: 25200056), store at -20 °C

5. Dulbecco’s phosphate buffered saline (PBS) (Gibco, catalog number: 11965-092), store at 4 °C

6. HBmito Crimson, 200 μM DMSO solution, store at -20 °C

The dyes in this article can be synthesized and purified according to the literature [11] or purchased from MedChemExpress (catalog number: HY-D2346).

7. DMSO (analytical reagent grade) (Aladdin, catalog number: D103272), store at room temperature

8. Immersion oil (Olympus, catalog number: IMMOIL-F30CC)

Solutions

1. Complete COS7 cell media (see Recipes)

2. HBmito Crimson solution (see Recipes)

3. HBmito Crimson DMSO solution (see Recipes)

Recipes

1. Complete COS7 cell media

The solution should be stored at 4 °C and used within one month. Ensure that it is sterile each time it is used. FBS should be filtered through a 0.2 μm filter membrane.

2. HBmito Crimson solution

Prepare the solution before each use.

3. HBmito Crimson DMSO solution

Prepare the solution before each use.

Laboratory supplies

1. Sterile 1.5 mL polypropylene centrifuge tubes (any vendor)

2. Sterile 50 mL polypropylene centrifuge tubes (any vendor)

3. Sterile nuclease-free filter tips (10, 200, and 1000 μL) (any vendor)

4. 35 mm glass bottom dishes, 170 μm ± 5 μm (Standard Imaging, catalog number: STGBD-035-1; Cellvis, catalog number: C35-20-1.5H)

5. Pipette tips, sterile (any vendor)

6. Cell culture flasks, vented flask surface area 25 cm² (any vendor)

7. Pipettor (Eppendorf Research^® plus) with volume ranges of 0.5–10 μL, 10–100 μL and 100–1000 μL

8. 0.2 μm filter membrane (Millex-GP, catalog number: SLGPR33RB)

Equipment

1. Abberior Facility Line (Abberior Instruments GmbH, Germany) with a 60× oil immersion objective (N.A. 1.42, Olympus, Japan)

2. Common lab equipment: cell culture incubator, safety cabinet, -20°C freezer, fridge (any vendor)

3. Microscope incubator (OKOlab, model: H301-T-UNIT-BL-PLUS)

Software and datasets

1. Abberior Imspector (16.3.14280)

2. Huygens SVI (23.10)

3. ImageJ Fiji, version 2.1.0. with Java 1.8.0_172

4. Origin (2019b)

Procedure

A. STED saturation power measurement

STED saturation power refers to the laser power required to effectively deplete the fluorescence of dye molecules. Dyes with low saturation power improve the overall quality of STED imaging, especially in high-resolution, long-term experiments. Therefore, saturation power is a key parameter for determining whether a dye is suitable for high-resolution imaging. Additionally, measuring saturation power can help estimate the optimal depletion laser intensity needed for imaging, such as in section C for setting the depletion power of HBmito Crimson. The resolution in STED microscopy can be described by the following formula:

d ≈ λ 2 N . A . 1 + I S T E D I s a t

λ is the excitation wavelength, N.A. is the numerical aperture of the microscope objective, I_STED is the intensity of the depletion laser, and I_sat is the saturation intensity of the fluorophore. From the formula, it is evident that when I_STED remains constant, decreasing the saturation power I_sat increases I_STED/I_sat, resulting in higher resolution. Therefore, low-saturation-power probes can achieve high resolution at lower laser powers, effectively reducing photobleaching and phototoxicity.

1. Add 1 mL of HBmito Crimson DMSO solution to a 35 mm glass bottom dish.

2. Set the spatial light modulator (775 nm) mode to “none” in the background interface of the commercial STED system, transforming the depletion beam of the donut distribution into a Gaussian distribution. Set the excitation intensity to 3%, which can be calibrated according to the dye's brightness. Adjust the excitation intensity to 3%, calibrated based on the brightness of the dye. Gradually increase the depletion laser intensity from 0%, in increments of 1%–3% (approximately 2–5 mW), e.g., 0%, 2%, 4%, and so on, up to 50%. At each depletion laser intensity level, capture two images—one with the excitation laser on and one with it off. Record the average fluorescence intensity for each image.

3. For each depletion laser power, subtract the average fluorescence intensity of the excitation-off image from that of the excitation-on image. Plot the normalized fluorescence intensity against the depletion laser intensity. Fit the data to a curve and identify the point at which the fluorescence intensity decreases to 50% of its initial value. The corresponding depletion laser intensity at this point is the saturation power of the dye.

Note: Repeat the imaging and recording at each laser intensity multiple times to ensure data accuracy and reliability. Monitor the sample for photobleaching to prevent damage from excessive excitation and adjust the laser settings as needed to minimize photobleaching.

B. Sample preparation

1. In our work, COS7 cells were cultured in a cell culture flask with COS7 medium. The cells were maintained at 37 °C in a humidified atmosphere containing 5% CO₂ for 2–3 days.

2. When the cells reach 80% density, remove the original COS7 medium and wash the cells with 3 mL of PBS to eliminate the residual medium. Add 1 mL of trypsin-EDTA solution for digestion for 3 min. After digestion, add 1 mL of COS7 cell medium to stop the digestion process.

3. Centrifuge the digested cells at 200× g for 3 min to collect the cell pellet.

4. Seed approximately 6 × 10⁴–9 × 10⁴ cells into 35 mm glass-bottom dishes. For optimal imaging results, ensure that the cells are evenly distributed and not overcrowded after seeding.

5. Allow the cells to grow in the incubator for 24 h. A confluence between 40%–80% is ideal for subsequent labeling and imaging procedures. If cell washing is required, the optimal imaging time is within 1 h after washing.

6. Remove the original medium and add 1 mL of HBmito Crimson solution, incubating for 10 min in a 37 °C, 5% CO₂ environment. Without washing, proceed directly to STED imaging, making the necessary adjustments to meet specific experimental requirements.

Note: When preparing cells for imaging, several key factors must be considered:

Cell density: Maintain an appropriate cell density (between 40% and 80%) to ensure cell health while providing sufficient space for single-cell imaging. High cell density may lead to cell aggregation, adversely affecting image quality.

Culture medium selection: Choose an appropriate basal medium and serum for different cell lines to ensure optimal growth conditions.

Contamination control: Cells must be free of any contamination to ensure reliable imaging results. The inclusion of antibiotics in the culture medium can help prevent contamination.

C. Image acquisition

This protocol is specifically designed for use with the Abberior Facility Line, but the method can be adapted for use with other STED microscopes as well.

When acquiring live-cell imaging with fluorophores, it is crucial to select the appropriate wavelength range. Optimal excitation power, depletion laser power, pixel size, line accumulations, and dwell time should be empirically determined to maximize the signal-to-noise ratio (SNR) while minimizing photobleaching, particularly during extended time-lapse imaging. We also recommend positioning the cells in the center of the field of view (FOV), as the imaging quality is best at the center of the FOV.

Excitation power: Select an appropriate excitation wavelength based on the dye properties and experimental requirements. For HBmito Crimson, use an excitation wavelength of 640 nm. Start with a low excitation intensity (1%–5%) and gradually increase the excitation intensity in increments of 1%–3%. Determine an appropriate excitation power value based on the signal intensity observed in the image. Be cautious to avoid over-excitation, which can lead to photobleaching of the sample and prevent signal oversaturation.

Depletion laser power: Select an appropriate depletion wavelength. For HBmito Crimson, use a STED depletion laser wavelength of 775 nm. Start with a depletion laser power of 15%–20% (approximately 20–30 mW) and gradually increase the intensity in increments of 1%–3% (approximately 2–5 mW). Adjust the depletion light intensity based on the observed resolution and SNR in the images.

Pixel size: In STED microscopy, setting an appropriate pixel size is crucial to satisfy the Nyquist sampling theorem and ensure that the acquired images accurately reflect the sample's details and resolution. Pixel size should be at least 1/2 of the resolution, i.e., pixel size ≤ d/2.

Dwell time: Dwell time determines the exposure time for each pixel, thereby influencing the SNR and resolution of the image. For most samples, the setting of dwell time is typically between 1 and 10 μs. This range ensures that the image maintains an adequate SNR. If the sample signal is relatively weak, the exposure time can be increased to 10–20 μs to enhance the signal collection per pixel. A longer dwell time will improve the SNR, but it also increases the overall imaging time and the risk of photobleaching.

Line accumulations: The setting of line accumulations affects the SNR and resolution of the image. For common samples, the default line accumulations are typically set between 2 and 4 times. This range significantly enhances the SNR while avoiding excessive exposure that could lead to photobleaching. If the SNR is weak, the number of line accumulations can be increased, ensuring the sample remains stable and motionless during the acquisition process and does not shift due to prolonged acquisition times. For unstable samples, it is advisable to reduce the number of line accumulations. For experiments requiring high resolution and high SNR, consider setting line accumulations to higher values (e.g., 8 or more). Conversely, for rapid imaging needs or samples prone to photobleaching, lower line accumulations (e.g., 1–2 times) can be selected to reduce photobleaching and imaging time.

To determine the optimal imaging parameters, we recommend using the method of controlled variables, systematically altering one parameter at a time to observe its effect on image quality. This approach allows for achieving the best image quality and highest resolution.

1. Power on the microscope system, ensuring proper communication between the software and hardware, and open the Abberior Facility Line software.

2. Select the 63× oil immersion objective and apply immersion oil to the lens.

3. Place the standard beads sample on the sample holder, open the software, select the alignment function, and wait for the system dialog to display the message "alignment successful," indicating the program is complete. Ensure both excitation and depletion lasers are properly aligned.

4. Connect the live-cell incubation chamber to the microscope system. Once the environmental conditions in the incubation chamber stabilize at 37 °C, 5% CO₂, and 75% humidity, add the prepared sample.

5. Use the microscope’s autofocus function to locate the sample’s focal plane. The Abberior autofocus actively stabilizes the Z-position of the sample, with continuous Z-drift compensation ensuring that focal drift does not occur during confocal and STED imaging.

6. Identify a suitable imaging area in confocal mode.

7. Adjust the imaging parameters accordingly.

As shown in Figure 2, λ_Ex 640 nm: 15.7 μW; λ_Dep 775 nm: 71.2 mW; pixel size: 20 nm; line accumulations: 3; dwell time: 6.5 μs; ROI: 44.5 μm × 50.74 μm; detection wavelength: 650–750 nm.

Figure 2. Comparison of confocal and STED imaging results in living COS7 cell mitochondria labeled with HBmito Crimson. Scale bar in the original image is 5 μm. Scale bar in the enlarged image is 1 μm.

8. Acquire STED images, ensuring that confocal and STED images are collected simultaneously.

9. For long-term imaging, appropriately reduce the excitation power, depletion power, line accumulations, and dwell time based on empirical observations. Depletion laser intensity can be reduced to approximately half of that used for single-frame imaging to meet the imaging requirements. Set the time intervals and number of imaging frames, then acquire images.

As shown in Figure 3, λ_Ex 640 nm: 12.75 μW; λ_Dep 775 nm: 34.7 mW; pixel size: 25 nm; line accumulations: 3; dwell time: 5 μs; frame time: 8.22 s; ROI: 4.75 μm × 7.025 μm; detection wavelength: 650–750 nm.

Figure 3. Long-term imaging of mitochondria in COS7 cells labeled with HBmito Crimson; white arrow indicates fusion event. Scale bar: 1 μm.

Note: When determining parameters for long-term imaging, ensure the minimal necessary laser intensity and exposure time to achieve only the required resolution. For mitochondrial cristae imaging, the minimum resolution requirement is defined as the clear visualization of cristae structures at the given power level. By utilizing these optimized settings, long-term imaging can minimize photobleaching effects, allowing for the acquisition of more frames and extending the imaging duration.

10. Save the data.

D. Extended image processing

Process single-frame images directly using Huygens software, setting post-processing parameters for each channel separately. For long-term imaging, it is recommended to first perform intensity correction, followed by batch deconvolution in Huygens software.

1. Open the Huygens Professional software. Import the image files that need deconvolution by selecting File > Open.

2. Access the image properties by selecting the image in the main window. This option can be found under the Edit menu or by right-clicking the image and selecting Properties. Set the microscope parameters accordingly:

Objective lens: Enter the numerical aperture (N.A.) and magnification of the objective lens used to acquire the images.

Excitation wavelength (λ_Ex): Specify the excitation wavelength used during imaging.

Emission wavelength (λ_Em): Enter the emission wavelength corresponding to the fluorescence channel.

Voxel size: Input the pixel size (in x, y, and z dimensions) used during image acquisition.

Refractive index: Enter the refractive index of the immersion medium used (e.g., oil, water, glycerol).

Pinhole radius: If using a confocal microscope, specify the pinhole radius or size.

3. Select point spread function (PSF): In the Restoration menu, select the PSF option. If the software does not automatically detect the PSF, you can manually select or load the appropriate PSF file. Select Deconvolution Wizard from the Restoration menu. Deconvolution is typically performed using the classic maximum likelihood estimation (CMLE) algorithm, where the software automatically calculates parameters like SNR and background. These default settings are optimized to handle most images. However, parameters such as iteration count, SNR, and background can be manually adjusted to enhance image quality, especially for more complex or low-SNR images.

4. Click the Run button to start deconvolution processing.

5. After processing, use the software's visualization tool to view the deconvolved image.

6. Save the processed image by selecting Save As from the File menu. Save the image in common formats such as TIFF, JPEG, or other available formats.

7. Use the Batch Processor tool to perform batch deconvolution processing on multiple image files. After setting the batch processing parameters and file path, click the Start Batch button to start processing.

Data analysis

The resolution of the STED images is analyzed based on the full width at half maximum (FWHM) of mitochondrial structures. This analysis provides a quantitative measure of the system's resolution, allowing us to accurately assess the spatial resolution achieved during imaging.

1. Launch ImageJ software and open the desired image by selecting File > Open.

2. Duplicate the field of image to be analyzed. Use the Straight Line function in ImageJ to select multiple cristae within this area, as shown in Figure 4, and generate the signal intensity profile at the specified location following the direction indicated by the arrows.

Figure 4. Enlarged STED and STED+ results and fluorescence signal intensity profile correspond to the white arrow. Scale bar: 1 μm. Results were processed with the commercial deconvolution software Huygens (SVI, Netherlands).

3. Utilize the Analyze > Plot Profile function in ImageJ to obtain the signal intensity profile. Click List to open the intensity values, then copy these values into Origin software.

4. In Origin, apply a Gaussian Fit to derive the Gaussian fitting curve, as shown in Figure 5.

5. From the Gaussian fit, calculate the standard deviation (σ). Use the formula FWHM = 2.355 × σ to determine the FWHM of the image.

Figure 5. STED and STED+ results and the fitted fluorescence signal intensity distribution correspond to the white arrows. Scale bar: 1 μm.

Validation of protocol

This protocol or parts of it has been used and validated in the following research article:

1. Ren et al [11]. Visualization of cristae and mtDNA interactions via STED nanoscopy using a low saturation power probe. (Figures 2–6).

General notes and troubleshooting

General notes

1. The dye concentration must be carefully optimized; neither too high nor too low. The recommended dye concentration of HBmito Crimson is 500 nM and the dyeing time is 10 min. Adjustments can be made according to the specific experimental requirements.

2. Select an appropriate laser wavelength for excitation based on the specific dye.

3. Calibrate the excitation and depletion lasers of the STED system to achieve optimal STED performance.

4. Maintaining stable laboratory temperature and humidity is essential to avoid environmental fluctuations that could impact microscope performance.

Troubleshooting

Problem 1: Motion artifacts in STED imaging

Possible cause: Sample jitter or mitochondrial movement.

Solution: It is essential to ensure the sample remains stable and motionless during imaging. In live-cell imaging, slow imaging speeds can lead to mitochondrial movement, resulting in motion artifacts. The imaging speed is limited by pixel dwell time and the line accumulations. Therefore, it is important to minimize line accumulations and dwell time while ensuring sufficient fluorescence signal collection. Additionally, reducing the FOV can help increase imaging speed and reduce the impact of mitochondrial movement.

Problem 2: Insufficient resolution in STED images

Possible cause: Improper alignment of the STED optical path or suboptimal parameter settings.

Solution: Ensure that the system is correctly calibrated. After calibration, image standard 40 nm beads. Perform Gaussian fitting on the signal intensity of individual beads to obtain their FWHM. If the FWHM reaches 40 nm, the system is in good condition. During sample imaging, improper parameter settings can lead to insufficient resolution. Refer to the parameter setting recommendations in the procedure. When adjusting parameters, avoid repeatedly imaging the same FOV to minimize photobleaching effects.

Acknowledgments

This protocol is related to the following paper: Ren et al. Visualization of cristae and mtDNA interactions via STED nanoscopy using a low saturation power probe. DOI: 10.1038/s41377-024-01463-9. This work was supported by the National Key R&D Program of China (2022YFC3401100), National Natural Science Foundation of China (22177024, 62025501, 31971376, 92150301), and Central Guidance Fund for Local Science and Technology Development (246Z1302G). We thank the National Center for Protein Sciences at Peking University in Beijing, China, for assistance with STED super-resolution imaging. We thank Abberior China and Optofem Technology Limited for providing Facility Line STED and Huygens software.

Competing interests

B.G. is an inventor of the awarded patent (ZL202011401937.1) of HBmito Crimson. The other authors declare no competing interests.

Ethical considerations

There are no ethical considerations associated with this protocol.

References

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