Label-Free Imaging Sample Prep: Step-by-Step

Label-free imaging techniques like quantitative phase microscopy, holographic microscopy, and autofluorescence imaging let you observe living cells without dyes or fluorescent labels. This preserves native cell behavior and avoids phototoxicity, but it demands clean, healthy, and properly mounted samples. This guide walks you through preparing living cells for label-free imaging, from choosing the right dish to maintaining optimal conditions during the session.

What You’ll Need

• Living cells (adherent or suspension) in culture
• Appropriate imaging dish or chambered coverglass (glass-bottom dishes, μ-Slides, or coverslips)
• Complete culture medium (with serum and supplements as needed)
• Phenol red–free medium (to reduce background absorbance)
• HEPES buffer (1 M stock, pH 7.4) or other buffer for pH stability
• Phosphate-buffered saline (PBS), sterile
• Trypsin-EDTA or cell dissociation buffer (for adherent cells)
• Hemocytometer or automated cell counter
• Centrifuge and sterile tubes
• 37°C water bath or bead bath
• COâ‚‚-independent medium (if imaging without COâ‚‚ control)
• Optional: fibronectin, poly-L-lysine, or collagen for coating
• Optional: anti-fade or immersion oil (if using oil-immersion objective)

Step-by-Step

1. Prepare the imaging surface. If using glass-bottom dishes or coverslips, coat them with an extracellular matrix protein (e.g., fibronectin at 5–10 µg/cm², poly-L-lysine at 0.01%, or collagen) to promote cell attachment. Incubate at 37°C for at least 30 minutes, then rinse gently with sterile PBS.
2. Warm all media and reagents. Pre-warm complete culture medium, PBS, and trypsin to 37°C in a water bath. This prevents cold shock to cells.
3. Prepare cells. For adherent cells: remove old medium, rinse with PBS, and add trypsin-EDTA (just enough to cover the monolayer). Incubate at 37°C until cells detach (usually 2–5 minutes). Tap the flask gently to dislodge cells. Add warm complete medium (at least 2× the trypsin volume) to neutralize trypsin. For suspension cells: simply collect the culture medium containing cells.
4. Count and assess viability. Take a small aliquot and count cells using a hemocytometer or automated counter. Use trypan blue exclusion to check viability; aim for >90% viability. If viability is low, consider a gentler dissociation method.
5. Centrifuge and resuspend. Transfer the cell suspension to a sterile tube and centrifuge at 200–300 × g for 5 minutes. Discard the supernatant and gently resuspend the pellet in pre-warmed, phenol red–free complete medium. Adjust to the desired density (typically 1–5 × 10� cells/mL for imaging).
6. Seed cells into the imaging dish. Add the cell suspension to the prepared dish. Use a volume that covers the glass surface (e.g., 200–500 µL for a 35 mm dish). Ensure even distribution by gently rocking the dish.
7. Allow cells to attach. Incubate at 37°C in a CO₂ incubator for at least 4 hours, or overnight, to allow firm attachment and spreading. For suspension cells, you may proceed directly to imaging, but consider using a dish coated with poly-L-lysine to immobilize them.
8. Switch to imaging medium. Just before imaging, replace the culture medium with pre-warmed, phenol red–free medium supplemented with 10–25 mM HEPES (if not using CO₂ control). This maintains pH without the need for a CO₂ supply. If you have CO₂ control on the microscope, you can use standard medium without HEPES.
9. Check cell morphology. Under a standard microscope, verify that cells are well-attached, spread, and show normal morphology (e.g., polygonal shape for epithelial cells, spindle shape for fibroblasts). Avoid cells that look rounded or detached.
10. Mount the dish on the microscope. Place the dish securely on the stage. For oil-immersion objectives, apply a small drop of immersion oil to the objective and raise it until contact with the dish bottom. Ensure the dish is level and the cells are in focus.
11. Maintain temperature. Use a stage-top incubator or heated stage set to 37°C. If not available, use a heated objective and pre-warmed medium, but be aware of temperature drift.
12. Acquire images. Start with low light intensity to minimize photodamage. For label-free techniques like phase contrast or quantitative phase, adjust the condenser and focus to achieve optimal contrast. Take images at appropriate intervals if doing time-lapse.

Common Mistakes

• Using phenol red medium: Phenol red absorbs light and can interfere with some label-free techniques, especially those relying on absorbance or fluorescence. Always switch to phenol red–free medium.
• Cells too dense: Overconfluent monolayers cause multiple scattering and make individual cell analysis difficult. Aim for 50–70% confluency.
• Air bubbles: Bubbles in the medium or between the dish and objective cause artifacts. Pipette gently and check for bubbles.
• Temperature fluctuations: Cells respond to temperature changes, affecting morphology and movement. Keep the stage heated and minimize time out of the incubator.
• Using trypsin for too long: Over-trypsinization damages cell membranes and alters cell shape. Watch cells under a microscope and stop as soon as they detach.
• Forgetting to coat the dish: Some cells (e.g., neurons) require specific coatings to attach properly. Skipping this step leads to poor attachment and floating cells.

Tips

• Use low-passage cells (passage number < 20) for more consistent behavior.
• Filter all media through a 0.22 µm filter to remove particulates that could scatter light.
• If using a glass-bottom dish, ensure the glass is #1.5 thickness (0.17 mm) for optimal imaging with high-NA objectives.
• For suspension cells, consider using a concanavalin A–coated dish to immobilize them without affecting viability.
• Test the imaging setup with a blank dish containing only medium to check for background artifacts.
• Keep a separate dish of cells in the incubator as a backup in case the imaging session fails.

Troubleshooting

• Cells are floating or detached: This may be due to insufficient coating, over-trypsinization, or temperature shock. Re-coat the dish, use gentler dissociation, and ensure all reagents are warm.
• Poor contrast or blurry images: Check focus, adjust the condenser aperture, and ensure the dish bottom is clean. Also, check for air bubbles or dirt on the objective.
• Cells look unhealthy (vacuolated or blebbing): This could be due to pH imbalance. Add more HEPES (up to 25 mM) or use a COâ‚‚-controlled chamber. Also check that the medium is fresh and not expired.
• Drifting focus over time: This is often due to thermal expansion. Allow the stage to equilibrate for 15–20 minutes before starting the experiment. Use a focus-hold system if available.
• Phototoxicity even without labels: Some label-free techniques still use light that can cause damage. Reduce illumination intensity or exposure time, and use a sensitive camera.
• Contamination: If you see bacteria or yeast, discard the sample and start over with sterile technique. Always work in a laminar flow hood.

Frequently Asked Questions

Can I use regular culture medium with phenol red?

It’s not recommended. Phenol red absorbs light in the visible range and can interfere with quantitative phase and other label-free techniques. Switch to phenol red–free medium.

How long can I keep cells in the imaging dish?

Typically, you can maintain cells for several hours to a day if temperature and pH are controlled. For longer experiments, use a perfusion system to refresh nutrients and remove waste.

Do I need to fix the cells for label-free imaging?

No, the whole point is to image living cells. Fixation kills cells and alters their morphology, defeating the purpose.

What if my cells don’t attach to the glass?

Try coating the glass with poly-L-lysine or fibronectin. Also, ensure the glass is clean and that the cells are healthy. Some cell types (e.g., suspension cells) may need special treatment.

Can I use a regular culture dish instead of a glass-bottom dish?

Plastic dishes are not suitable for high-resolution imaging because they are not optically flat and may autofluoresce. Use glass-bottom dishes or coverslips.

How do I prevent evaporation during long time-lapse?

Use a stage-top incubator with humidity control, or overlay the medium with mineral oil. Alternatively, seal the dish with parafilm to reduce evaporation.

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