Super-Resolution Imaging and Shared Management: A Protocol for Confocal Microscopy with Multiplex Detection

Summary

This protocol details the advanced imaging applications, and shared-resource management of a laser-scanning confocal microscope system integrated with a multiplexed array detector for high-resolution cellular and subcellular imaging. The workflow enables multimodal imaging from confocal to super-resolution (~120 nm) within a single platform, making nanoscale visualization more accessible.

Abstract

Super-resolution microscopy (SRM) overcomes the diffraction limit of conventional light microscopy, enabling nanoscale visualization of subcellular structures. However, the widespread adoption of techniques such as stimulated emission depletion (STED) microscopy, photoactivated localization microscopy (PALM) / stochastic optical reconstruction microscopy (STORM), and structured illumination microscopy (SIM) can be hindered by factors such as high phototoxicity, specialized sample preparation, or complex operation. Here, we present a comprehensive protocol that leverages a laser-scanning confocal platform integrated with a multiplexed super-resolution detector. This system, combined with computational processing, enables a straightforward transition between confocal and super-resolution imaging within the same workflow. The protocol provides a user-friendly approach for fast, high-resolution, and low-phototoxicity imaging. The detailed method encompasses multimodal imaging capabilities, including multicolor fluorescence, differential interference contrast (DIC), three-dimensional Z-stack reconstruction, time-lapse live-cell imaging, large-area tiling, and various super-resolution modes offered by the multiplexed array detector. Systematic comparisons demonstrate that the super-resolution modes achieve a lateral resolution of approximately 120 nm, representing about a two-fold improvement over the diffraction limit of conventional wide-field microscopy and a significant enhancement in both resolution and signal-to-noise ratio (4-8 fold) compared to standard confocal operation on the same platform. The discussion also covers key points of a shared resource management model essential for supporting the sustainable operation of this technology. This integrated guide serves as a valuable resource for both new and experienced researchers and facility managers aiming to adopt and maximize the potential of this accessible form of advanced imaging in life sciences research.

Introduction

The visualization of dynamic subcellular structures is fundamental to advancing life sciences research. However, the resolution of traditional optical microscopy is limited by diffraction to approximately 200 nm laterally, restricting its ability to resolve dynamic subcellular processes. To overcome this barrier, various super-resolution techniques have been developed. However, techniques such as STED microscopy, PALM/ STORM, and SIM often present barriers to widespread adoption in multi-user environments, including high phototoxicity, the need for specialized fluorophores or sample preparation, complex alignment procedures, or intensive computational processing^1,2,3.

In this context, confocal microscope systems equipped with a multiplexed array detector offer a compelling alternative by enabling super-resolution imaging through multi-pixel detection and computational reassignment, significantly lowering the technical threshold⁴. This approach leverages a 32-element gallium arsenide phosphide (GaAsP) detector array to collect more photons from each scanning point. Subsequent intelligent deconvolution algorithms reassign this information, effectively reducing the detection volume. This process achieves spatial resolutions of approximately 120 nm laterally and 350 nm axially, representing about a 2x improvement over the diffraction limit of wide-field microscopy and yielding a 4-8x increase in signal-to-noise ratio compared to standard confocal mode on the same instrument^4,5,6. This unique combination of enhanced resolution, superior signal quality, and compatibility with common fluorescent labels and protocols constitutes its primary advantage for broad application.

Consequently, this form of accessible super-resolution microscopy has found broad application in life sciences research, enabling detailed studies of organelle morphology, cytoskeleton dynamics, virus-host interactions, and molecular co-localization in both fixed and living cells^7,8. Its relative gentleness makes it particularly suitable for extended time-lapse observations of live samples.

The integration of such advanced systems into core facilities is crucial for democratizing access, but their sustainable and effective operation presents distinct challenges. In institutional core facilities, such as at the Zhejiang University School of Medicine, optimal performance and impact depend critically on three pillars: (1) standardized and reproducible imaging protocols, (2) rigorous routine maintenance, and (3) an effective management framework for shared access. While valuable resources exist for specific aspects, such as imaging methods for earlier systems⁹or general facility management principles^10,11, a comprehensive guide that integrates detailed, reproducible protocols for modern multiplexed array detector platforms with a proven shared-resource management model is currently lacking. This protocol article aims to bridge this gap.

The overarching goal is to provide a comprehensive, step-by-step guide for effectively implementing and utilizing a confocal system with a multiplexed array detector for super-resolution imaging within a shared-resource setting. It is designed for two primary audiences: researchers requiring resolution beyond conventional confocal limits (~250‬-300 nm) but for whom live-cell compatibility, ease of use, and throughput are also critical; and facility managers establishing or optimizing such a service. Herein, we systematically detail the system configuration, multimodal imaging protocols (covering multicolor, 3D, live-cell, tiling, and super-resolution modes), and a proven shared management model. This integrated guide serves as a resource to maximize the potential and accessibility of this powerful imaging modality.

Protocol

All experiments involving biological samples were conducted in accordance with the guidelines and regulations of Core Facilities, Zhejiang University School of Medicine and were approved by the Institutional Biosafety Committee (Approval Certificate No. BSL20235710079).

NOTE: All procedures involving cell cultures must be performed under sterile conditions in a Class II biological safety cabinet unless otherwise specified. This protocol assumes the microscope system is installed, and laser powers are calibrated. Detailed specifications and recommended vendors for key reagents, consumables, and equipment (e.g., imaging dishes, cover glass, mounting medium) are listed in the Table of materials.

1. Sample preparation

NOTE: Successful super-resolution imaging begins with optimal sample preparation. The use of high-quality, 1.5 thickness (0.17 mm) cover glass-based imaging dishes is essential to match the correction collar of high-NA oil immersion objectives and avoid spherical aberration. All centrifugation steps should be performed at 300 × g for 5 min unless noted.

1. Fixed cell sample preparation for multicolor super-resolution imaging (e.g., COS7 cells)
   NOTE: This protocol describes immunostaining for microtubules (Alexa Fluor 568) and mitochondria (Alexa Fluor 488), with 4',6-diamidino-2-phenylindole (DAPI) nuclear counterstain, as shown in Figure 1.
   1. Culture COS7 cells in Dulbecco's modified Eagle medium (DMEM) supplemented with 10% fetal bovine serum (FBS) at 37°C in a 5% CO₂ humidified incubator.
   2. Plate cells onto 35 mm glass-bottom imaging dishes at a density of 1.0 × 10⁵ cells/dish. Allow cells to adhere and grow to 60-70% confluency (typically 24 h).
   3. Aspirate the culture medium and wash cells once gently with pre-warmed (37°C) phosphate-buffered saline (PBS). Fix the cells by adding 1 mL of 4% paraformaldehyde (PFA) in PBS and incubate for 15 min at room temperature (RT).
   4. Aspirate the PFA solution (treat as hazardous waste). Wash the cells three times with 2 mL of PBS, incubating for 5 min per wash with gentle rocking.
   5. Permeabilization and blocking: Incubate the cells with 1 mL of permeabilization/blocking buffer (0.1% v/v) Triton X-100, 1% (w/v) bovine serum albumin (BSA) in PBS) for 30 min at RT.
   6. Primary antibody incubation: Incubate cells with a mixture of primary antibodies against α-tubulin (mouse monoclonal) and TOMM20 (rabbit polyclonal) in antibody dilution buffer (1% BSA in PBS) at 4°C overnight (or 1-2 h at RT). Use 300 µL per dish to cover the glass surface.
   7. Washing: Carefully aspirate the primary antibody solution. Wash the cells three times with 2 mL of PBS containing 0.05% Tween-20 (PBST), 5 min per wash.
   8. Secondary antibody and nuclear staining: Prepare a mixture of fluorescent secondary antibodies and DAPI in antibody dilution buffer (1% BSA/ PBS), protected from light.
      1. Use donkey anti-mouse IgG conjugated to Alexa Fluor 568 (1:500) and donkey anti-rabbit IgG conjugated to Alexa Fluor 488 (1:500).
      2. Include DAPI at a final concentration of 1 µg/mL for nuclear staining. Apply 300 µL of this mixture to the cells and incubate for 1 h at RT in the dark.
   9. Final washing: Aspirate the staining solution and wash the cells three times with 2 mL of PBS, 5 min per wash.
   10. Apply mounting medium: Carefully aspirate the PBS from the imaging dish, leaving the sample slightly moist. Immediately apply a small drop (~20 µL) of hardened anti-fade mounting medium (e.g., ProLong Gold) directly onto the glass-bottom region.
   11. Cover and cure: Gently lower a #1.5 high-precision cover glass onto the droplet, avoiding bubbles. Allow the mountant to cure overnight at RT in the dark, or for 2-4 hours at 37°C if faster curing is required.
2. Live cell sample preparation for time-lapse imaging (e.g., Hep3B cells expressing a fluorescent protein tag).
   NOTE: This protocol describes imaging of Hep3B cells stably expressing a fluorescent protein (e.g., EGFP) targeted to mitochondria.
   1. Culture Hep3B cells in minimum essential medium (MEM) supplemented with 10% FBS. Plate cells onto 35 mm glass-bottom imaging dishes (1.5 cover glass) at a density of 0.8 × 10⁵cells/dish. Allow cells to grow to approximately 70-80% confluency for optimal health and imaging density (typically 18-24 h).
   2. Preparation for imaging: On the day of imaging, gently replace the standard culture medium with 2 mL of pre-equilibrated (37°C, 5% CO₂ for at least 30 min), phenol-red-free, low-fluorescence imaging medium supplemented with 10% FBS and, if required, 25 mM HEPES buffer for pH stabilization outside a CO₂ incubator. This minimizes background fluorescence and maintains physiological pH during imaging.
      NOTE: Allow the dish to acclimate on the pre-warmed microscope stage inside the environmental chamber for at least 30-45 min before starting time-lapse acquisition to ensure temperature, focus, and cell health stabilization.

2. Microscope system startup, initialization, and basic alignment

1. Power on the microscope main unit, the laser launch module (containing solid-state lasers at 405, 488, 561, and 640 nm), the LED light source, and the control computer.
   1. Allow the lasers to warm up and stabilize for at least 15-20 min. This is critical for stable laser power output, especially for quantitative or time-lapse experiments.
2. Launch the microscope operating software.
3. From the main interface, select 'Microscope' > ' Initialize' or follow the software's startup wizard.
4. In the software, navigate to the 'Acquisition' panel. From the objective lens dropdown menu, select 'Plan-Apochromat 63 x/ 1.40 Oil 'or the appropriate objective for your experiment.
5. Apply a drop of immersion oil (refractive index = 1.518) directly onto the front lens of the objective. Avoid introducing bubbles.
6. For live-cell imaging, activate the environmental chamber and the active focus stabilization system (e.g., 'Definite Focus' >) at least 45-60 min prior to imaging. Set the temperature to 37°C, CO₂ to 5%, and humidity to >60%. Allow the system to stabilize. Verify stability using the chamber sensors.
7. The system is now ready for sample loading and imaging parameter setup.

3. Multicolor fluorescence imaging

1. Place the prepared sample on the microscope stage. Carefully raise the stage until the cover glass contacts the immersion oil.
2. In the acquisition software, navigate the 'Acquisition' tab and select Create a new experiment.
3. Add multiple imaging tracks (channels) corresponding to the fluorophores used. For the example in Figure 1, configure:
   1. Track 1 (DAPI): Excitation 405 nm, detection 410-480 nm.
   2. Track 2 (Alexa Fluor 488): Excitation 488 nm, detection 495-550 nm.
   3. Track 3 (Alexa Fluor 568): Excitation 561 nm, detection 570-620 nm.
4. For each track, set the confocal pinhole diameter to 1 Airy Unit (1 AU).
5. Optimize laser power and detector gain for each channel using the following iterative process:
   1. Select the first channel (e.g., DAPI). Switch to 'Live' scanning mode.
   2. Set initial laser power to a low value (e.g., 0.1-0.5%). Set Detector Gain to a moderate level within its linear range (e.g., 700 V for a GaAsP PMT).
   3. Gradually increase the laser power until the target structures are clearly visible above the background in the live image.
   4. Open the Histogram display. Ensure no pixel saturation (histogram not piled at the right edge). Reduce laser power if necessary.
   5. Adjust the Detector Gain to achieve a bright, low-noise image. Avoid gains above 850 V to prevent excessive noise.
   6. Repeat steps 3.5.1-3.5.5 for all remaining fluorescence channels.
      NOTE: Balance laser power, gain, and pixel dwell time to maximize signal-to-noise ratio while minimizing photodamage. For live cells, prioritize lower power and shorter dwell times.
6. Set the image format (number of pixels) and pixel dwell time.
   1. Set the pixel size to satisfy the Nyquist criterion: pixel size ≤(resolution / 2.3). For standard confocal imaging (lateral resolution ~250 nm), set pixel size ≤108 nm.
   2. Use the software's 'Nyquist 'sampling button to calculate this automatically if available.
   3. Set the pixel dwell time. Use a longer dwell time (e.g., 1.0-2.0 µs) for higher signal-to-noise ratio, or a shorter dwell time (e.g., 0.5-1.0 µs) for faster imaging.
7. Ensure 'Sequential Scan' mode is selected between tracks to minimize crosstalk.
8. Click 'Start' to acquire a single image. Save the image with relevant metadata.

4. Differential interference contrast (DIC) imaging

1. In the acquisition software's 'Acquisition' tab, select 'Add a new channel ' and configure it for transmitted light detection.
2. Select the appropriate transmitted light detector (often called T-PMT (transmitted-light photomultiplier tube) or ESID (external system interface detector) in the software).
3. DIC prism and polarizer configuration:
   1. From the hardware control panel or software interface, select the DIC prism matched to your objective (e.g., 'DIC III' for a 63x objective).
   2. Insert the selected prism into the light path.
4. Adjust the polarizer and DIC slider for optimal contrast:
   1. Rotate the analyzer (polarizer) until the background in the 'Live' image appears dark or very dim.
   2. While observing the live image, slowly adjust the DIC slider position in small increments.
   3. Optimize until cellular features (e.g., membranes, nuclei) appear with a distinct shadow-cast, pseudo-three-dimensional relief against a neutral gray background.
5. Set the illumination wavelength for DIC to a value in the visible range (e.g., 488 nm or 546 nm).
6. Adjust the transmission intensity (laser or lamp power) and the detector gain for the transmitted light channel to achieve a clear image without saturation.
7. Acquire the DIC image.

5. Z-stack acquisition for three-dimensional reconstruction

NOTE: Before defining the Z-stack, configure all imaging parameters (laser power, gain, pinhole, pixel size) on a single, optimal focal plane as described in Section 3. This ensures consistent settings across all optical sections.

1. In the 'Acquisition' tab, select the 'Z-stack' mode.
2. Define the volume of interest:
   1. Using a fast-speed live scan, focus on the topmost structure of interest.
   2. Click 'Set First' (or 'Top').
   3. Carefully focus down to the bottommost structure of interest.
   4. Click 'Set Last' (or 'Bottom'). The software will display the total depth.
3. Set the Z-step Size to satisfy the Nyquist sampling criterion in the axial direction.(typically d_r/2.3, where d_r is the axial resolution). For a 63x/1.4 NA oil objective, use 0.3-0.4 µm.
4. Alternatively, use the software's 'Optimal' or 'Nyquist' button to calculate the step size automatically.
5. Configure imaging parameters (laser power, gain, pinhole = 1 AU) for each channel as described in Section 3. For weak signals, consider slightly increasing laser power or gain compared to a single 2D image.
6. Start the acquisition to collect the entire image stack.
   NOTE: Total acquisition time = (Frame Time from a single slice) × (Number of slices)
   ​Plan accordingly, especially for live samples.
7. 3D Reconstruction and Visualization:
   1. Use the software's 'Processing' module (e.g., 3D projection function) or open the image stack in dedicated software like ImageJ/Fiji.
   2. Generate maximum intensity projections (MIP) to view all in-focus signals in a single 2D image.
   3. For a true 3D representation, apply volume rendering. Adjust the brightness, opacity, and lighting as needed.
   4. Generate Orthogonal views (XZ and YZ cross-sections) to inspect axial extent and registration.

6. Time-lapse imaging for live cells

1. Prepare live cells as described in Section 1.2. and mount the dish on the pre-warmed microscope stage within the environmental chamber.
2. Allow the sample to acclimate for at least 45-60 min as described in Section 2.6 to ensure stabilization.
3. Navigate to a field of view with healthy, well-spaced cells and acquire a single high-quality snapshot using parameters optimized per Section 3.5.
4. Engage the active focus stabilization system (e.g., Definite focus or similar). This is non-negotiable for long-term experiments to compensate for thermal drift.
5. In the 'Acquisition' tab, select the 'Time Series' mode.
6. Select the appropriate imaging mode based on the required balance between speed, resolution, and phototoxicity:
   1. For maximum spatial resolution in less dynamic processes, select 'SR' (SuperResolution) mode.
   2. For a balance suitable for many organelle dynamics, select 'SR 2 Y ' or 'SR 4 Y ' (accelerated superresolution modes).
   3. For very highspeed imaging without superresolution, select 'Confocal' or 'Co 2 Y' mode.
7. Define the experimental timeline: Set the total duration of the experiment and the time interval (Δt) between consecutive frames. (e.g., 5-30 s for mitochondrial motility)
8. Optimize imaging parameters to minimize photodamage:
   1. Start with the lowest laser power that yields a detectable signal.
   2. Adjust detector gain within its linear range (600-850 V) to achieve a workable signal level, preferring gain over laser power increases.
   3. Use the fastest scan speed (shortest pixel dwell time) compatible with image quality requirements.
9. Start the time-lapse acquisition. Monitor the first few frames for focus stability and cell health.

7. Large-area tile scan imaging

1. In the 'Acquisition' tab, select the 'Tile Scan' mode.
2. Define the tilingRegion using one of two methods:
   1. Layout-based: Specify the number of tiles in the X and Y directions (e.g., 3 x 3).
   2. ROI-based: Manually draw a rectangle over the 'Live' image to define the region of interest. The software will automatically calculate the number of tiles needed.
3. Set the overlap between adjacent tiles to 10-15%.
   NOTE: This overlap provides sufficient information for the software's stitching algorithm to create a seamless composite image.
4. Configure imaging parameters (laser power, gain, resolution) for a single tile as you would for a static image.
   NOTE: For very large tiled areas, consider using a faster scanning mode (e.g., ' Co 2 Y ') to keep total acquisition time practical and minimize photobleaching.
5. Start the tiling acquisition. The software will automatically acquire and stitch the tiles.
6. After acquisition, carefully inspect the stitched image, especially at tile boundaries, for any artifacts. Use the software's manual stitching adjustment tools if necessary.

8. Super-resolution imaging using the multiplexed array detector

1. In the 'Acquisition' tab, navigate to the mode selector, click 'Smart Setup' and select 'Airyscan' mode. Then choose 'SuperResolution' as the operating mode for the multiplexed array detector.
2. Choose the specific acquisition mode based on experimental priorities:
   1. 'SR' Mode: For the highest possible spatial resolution (~120 nm lateral). Use for fixed samples.
   2. 'SR 2 Y' Mode: Provides a balance between resolution and speed.
   3. 'SR 4 Y' Mode: Fastest super-resolution mode (up to ~25 fps line rate,theoretically), optimized for live-cell imaging to minimize light dose.
   4. 'Co 2 Y' Mode: Offers confocal-like imaging speed with enhanced signal-to-noise ratio, when super-resolution is not required.
      NOTE: The resolution values (e.g., 120 nm) are system specifications under ideal conditions¹². The actual resolution depends on sample preparation, signal-to-noise ratio, and correct processing.
3. Configure imaging parameters (laser power, detector gain).
   1. For livecell imaging, start with lower laser power than in confocal mode to reduce photodamage.
   2. For dim, fixed samples, higher laser power and/or gain may be needed to achieve sufficient signaltonoise ratio.
   3. Find the minimum laser power that yields a clean, highcontrast image in ' Live' mode.
4. Set pixel size and dwell time.
   1. To capture the enhanced resolution, set pixel size to satisfy the Nyquist criterion: pixel size ≤(desired resolution / 2.3). For SR mode (target 120 nm), set pixel size ≤~52 nm.
   2. Use the software's 'Optimal' or 'Nyquist' button to calculate this automatically if available.
5. Acquire the image. The raw data contains multiplexed signals and is not yet superresolved.
6. Process the acquired data to achieve superresolution:
   1. Navigate to the software's processing module.
   2. Select the dataset and choose the 'SuperResolution Processing' function.
   3. Start with 'Auto' settings. For lowSNR data, manually adjust 'Strength' and 'Noise Filter' to enhance sharpness without introducing artifacts.
   4. (Optional) For highest fidelity, especially in 3D stacks, apply an additional joint deconvolution (jDCV) step.
   5. Execute processing. Compare processed and unprocessed views to verify enhancement of real details.
7. (Optional) For quantitative comparison of signaltonoise ratio (SNR) across modes:
   1. Acquire images of the same sample in different modes.
   2. In analysis software, define a region of interest (ROI) on a uniform structure and measure mean signal intensity.
   3. Define a background ROI and measure the standard deviation of background intensity.
   4. Calculate SNR = (Mean Signal) / (Standard Deviation of Background).
8. (Optional) For empirical systemresolution measurement using fluorescent beads:
   1. Prepare a sample of 100 nm fluorescent beads.
   2. Acquire a Zstack of an isolated bead in SR mode with Nyquist sampling.
   3. Process the stack with superresolution and (optionally) jDCV.
   4. Use the software's 'PSF Measurement' tool to fit a 3D Gaussian and record the fullwidth at halfmaximum (FWHM) values.

Results

Multicolor fluorescence imaging
Multicolor fluorescence imaging of fixed COS7 cells was performed as described in Section 3. Antibodies against α-tubulin (microtubules) and TOMM20 (mitochondria) were used with DAPI nuclear counterstain. As shown in Figure 1, nuclei (blue), mitochondria (green, Alexa Fluor 488), and microtubules (red, Alexa Fluor 568) were clearly delineated with high specificity and minimal spectral crosstalk, validating the effectiveness of the sequenti...

Discussion

The integrated protocol presented here provides a comprehensive roadmap for implementing the advanced imaging capabilities of a modern confocal microscope system equipped with a multiplexed array super-resolution detector, within a sustainable shared-resource framework. This section synthesizes the method's critical aspects, practical implementation, and its position in the bioimaging field. Successful execution of this protocol hinges on several pivotal steps. First, meticulous sample preparation is foundational; th...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
COS7 cells	ATCC (or similar)	CRL-1651	African green monkey kidney fibroblast cell line.Used for preparation of fixed-cell samples for multicolor super-resolution imaging
Donkey anti-mouse IgG, Alexa Fluor 568 conjugate	Invitrogen (or similar)	A10037	Secondary antibody for microtubule staining. Used at 1:500 dilution
Dulbecco‘s modified eagle medium (DMEM), high glucose	Gibco (or similar)	11965092	Standard culture medium for COS7 cells, supplemented with 10% FBS.
Fetal Bovine Serum (FBS), heat-inactivated	Gibco (or similar)	10082147	Serum supplement used at 10% (v/v) in all cell culture media.
Glass-bottom Imaging Dish (35 mm, µ-Dish, #1.5 polymer coverslip, 0.17 mm thickness)	ibidi (or similar)	81156	Substrate for cell plating and imaging. Critical for compatibility with high-NA oil immersion objectives.
Image processing & analysis software (ZEN modules)	Carl Zeiss AG	ZEN 3.5 (blue)	Used for 3D reconstruction, super-resolution processing (e.g., Airyscan Processing), PSF measurement, and SNR analysis.
ImageJ / Fiji	Open Source	N/A	Used for additional 3D reconstruction, generating orthogonal views, and general image analysis.
Immersion oil (Refractive Index = 1.518)	Carl Zeiss (or similar)	444960-0000-000	For use with oil immersion objectives (e.g., 63x/1.4 NA).
Laser-scanning confocal microscope system with multiplex array detector (e.g., ZEISS LSM 900 with Airyscan 2)	Carl Zeiss AG	LSM 900 (Airyscan 2)	The primary imaging platform used for all confocal and super-resolution modes described in the protocol.
Minimum Essential Medium (MEM) with Earle‘s Salts	Gibco (or similar)	11095080	Standard culture medium for Hep3B cells, supplemented with 10% FBS.
Mouse anti-α-tubulin monoclonal antibody (Clone DM1A)	Sigma-Aldrich (or similar)	T6199	Primary antibody for labeling microtubules. Used at 1:500 dilution
Paraformaldehyde (PFA), 4% in PBS	Jinpan (or similar)	TBS5083	Fixative for cells. Use at room temperature for 15 min. Handle with appropriate precautions.
Phenol-red-free Imaging Medium	Gibco (or similar)	21063029	Low-autofluorescence medium used for live-cell imaging. Pre-equilibrated with 5% CO2 at 37°C prior to use.
Phosphate-buffered saline (PBS), 1X, pH 7.4	CellWorld (or similar)	C0162-311	Used as washing and dilution buffer throughout the protocol.
ProLong gold antifade mountant	Invitrogen (or similar)	P36930	Hardened mounting medium for preserving fluorescence in fixed samples.
Triton X-100	Sigma-Aldrich (or similar)	T8787	Used at 0.1% (v/v) for cell permeabilization
Tween-20	Sigma-Aldrich (or similar)	P9416	Used at 0.05% (v/v) in PBS (PBST) for washing steps after antibody incubations.
Bovine serum albumin (BSA), Fraction V	Sigma-Aldrich (or similar)	A7906	Used at 1% (w/v) in PBS for blocking and as a component of antibody dilution buffer.
DAPI (4‘,6-diamidino-2-phenylindole)	Sigma-Aldrich (or similar)	D9542	Nuclear counterstain. Used at a final concentration of 1 µg/mL.
Donkey anti-rabbit IgG, Alexa Fluor 488 conjugate	Invitrogen (or similar)	A21206	Secondary antibody for mitochondrial staining. Used at 1:500 dilution
Fluorescent microspheres (Beads), 100 nm diameter	Invitrogen (e.g., TetraSpeck™) or similar	T7279	Used for empirical measurement of the system‘s point spread function (PSF) and resolution.
Hep3B cells stably expressing EGFP-mitochondria	IMMOCELL	IM-H367	Human hepatoma cell line engineered to stably express EGFP targeted to the mitochondrial matrix. Used for live-cell time-lapse imaging.
HEPES buffer solution, 1 M	Gibco (or similar)	15630080	Used at a final concentration of 25 mM in imaging medium for pH stabilization during live imaging outside a CO2 incubator.
High-precision coverglass (#1.5, 0.17 mm thickness)	Marienfeld (or similar)	107052	Used for mounting fixed samples.
Microscope operating software (e.g., ZEN blue edition)	Carl Zeiss AG	ZEN 3.5 (blue)	Used for system control, image acquisition, and basic processing.
Objective lens: Plan-apochromat 63x/1.40 NA Oil DIC	Carl Zeiss AG	421462-9900-000	Primary objective used for high-resolution imaging.
Rabbit anti-TOMM20 polyclonal antibody	Abcam (or similar)	ab186735	Primary antibody for labeling mitochondria. Used at 1:1000 dilution