Frequently Asked Questions about Cell Culture

Q1: How to passage adherent cells?

Remove the old medium from the T25 flask, add 3–4 mL PBS and gently pipette to wash 1–2 times, then discard the PBS. Add about 1.5 mL Trypsin-EDTA (1X) digestion solution, return the flask to the incubator or digest at room temperature while monitoring under a microscope. When cells become rounded, intercellular spaces enlarge, and some cells begin to detach, immediately add about 4 mL complete medium and mix gently to stop digestion. Gently pipette to detach cells fully from the flask surface, collect the cell suspension, and centrifuge at 1000 rpm for 5 min at room temperature. Discard the supernatant, resuspend the cell pellet in an appropriate volume of complete medium, split into new T25 flasks at a ratio of 1:2–1:3, 1:3–1:6, or 1:6–1:8 depending on cell density, and bring the final volume in each flask to 5–6 mL. Finally, place the flasks in a 37°C, 5% CO₂ incubator for continued culture.

Q2: How should suspension cells be passaged?

In general, simply add fresh medium to the original culture flask to dilute the cell density. When the culture volume becomes excessive, transfer the suspension to a centrifuge tube, centrifuge at 1000 rpm for 5 min at room temperature, discard the supernatant, and resuspend the pellet in complete medium. Split into new culture bottle at about a 1:4–1:8 ratio according to cell density, add complete medium to 4–5 mL per flask, and continue culture at 37°C, 5% CO₂. Some suspension cells tend to grow in aggregates, which often indicates good growth status; during medium addition, avoid excessive or repeated pipetting to prevent disrupting cell clumps.

Q3: How to use trypsin for passaging adherent cells?

Common trypsin–EDTA formulations for adherent cell passaging include 0.25% trypsin–0.53 mM EDTA·2Na or 0.05% trypsin–0.02 mM EDTA·2Na. Overly high digestion-solution concentrations can easily lead to increased cell debris and black impurities in the medium; therefore, 0.05% trypsin is recommended for routine passaging, while 0.25% trypsin can be used for cells that are difficult to detach. If cell confluence exceeds 80%, stepwise digestion is recommended to avoid overdigestion. Trypsin is generally stored at –20°C and thawed at 4°C. After the first thaw, aliquot immediately into sterile centrifuge tubes or cryovials and store at –20°C to avoid repeated freeze–thaw cycles that reduce trypsin activity, and to lower contamination risk.

Q4: How to control trypsin digestion time?

The extent of trypsinization is a critical step in cell passaging. Overdigestion can cause increased cell debris and black particles, massive cell detachment, reduced viability, and loss of cells together with discarded trypsin; insufficient digestion makes cells hard to detach and requires repeated pipetting, which also damages viability. Different cell types vary in trypsin sensitivity, so there is no fixed digestion time; it is influenced by multiple factors, including trypsin concentration, presence of EDTA, storage time and temperature, repeated freeze–thaw, volume of trypsin added, digestion temperature, and cell density. In practice, digestion time should be adjusted flexibly by dynamically observing morphological changes under a microscope.

Q5: What centrifugation speed should be used to pellet cells?

When cells need to be collected during passaging or cryopreservation, centrifugation at 800–1000 rpm for 5–8 min at room temperature is generally sufficient. Excessively high speed or prolonged centrifugation can damage cell membranes and cause cell rupture, thereby reducing survival; thus, overly high centrifugal force is not recommended.

Q6: How to distinguish live and dead cells?

Under a microscope, live cells are usually plump with clear outlines, slightly translucent centers, and a glossy appearance; dead cells are often dim, shrunken, and have poor refractivity. Trypan Blue staining can also be used for counting: live cells exclude the dye, whereas dead cells take up Trypan Blue and appear blue, allowing calculation of viability.

Q7: Why is EDTA added to trypsin?

The main role of EDTA in trypsin is to chelate free Ca²⁺ and Mg²⁺, removing divalent ions that maintain cell adhesion and inhibit trypsin activity, thereby enhancing digestion efficiency. It is usually recommended to gently rinse cells with Ca²⁺/Mg²⁺-free, EDTA-containing buffer before trypsinization to remove divalent ions in the medium, helping cells detach more smoothly.

Q8: Why do cells grow unevenly?

If flasks are not mixed thoroughly before being placed into the incubator after passaging, or if flasks are frequently moved before cells fully adhere, cells can distribute unevenly at the bottom, leading to uneven growth. In addition, frequent opening/closing of the incubator causing vibration, insufficient medium volume, or an unlevel incubator shelf can also hinder uniform spreading and ultimately cause uneven growth.

Q9: Should 5% or 10% CO₂ be used for cell culture?

Most culture media use the HCO₃⁻/CO₃²⁻/H⁺ buffering system for pH control, and the NaHCO₃ content determines the required CO₂ concentration. In general: if NaHCO₃ is ~3.7 g/L, 10% CO₂ is recommended; if NaHCO₃ is ~1.5 g/L, 5% CO₂ is sufficient. Therefore, select 5% or 10% CO₂ according to the NaHCO₃ formulation of the medium used.

Q10: Are black dots commonly seen during culture contamination?

First, check whether the medium appears turbid to the naked eye. If it is turbid, contamination is highly likely. If the medium is not visibly turbid, examine the black dots under a microscope for size, shape regularity, and motility—whether they show Brownian motion or rapid linear movement. If the dots are irregular in size and show Brownian motion, they may be cell debris (due to poor cell status or overdigestion), protein precipitates from repeated serum freeze–thaw, or cellular metabolites. If the dots are uniform in size and move rapidly, bacterial contamination is very likely.

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