Cell Culture Protocol for Mammalian Cells: Complete Guide

LabProtocol.co Team·2026-03-22·7 min read
cell-culturemammalian-cellsprotocolstissue-culture

Cell Culture Protocol for Mammalian Cells: Complete Guide

Mammalian cell culture is foundational to modern biomedical research — from drug screening and protein production to gene editing and cancer biology. Whether you are working with HEK293T, HeLa, CHO, Jurkat, or primary cells, the fundamentals of sterile technique, passaging, and maintenance remain the same.

This protocol covers everything you need to culture mammalian cells reliably: setting up your workspace, thawing frozen stocks, routine passaging, cryopreservation, and preventing the contamination that can destroy months of work.

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Materials and Equipment

Reagents

  • Complete growth medium (cell-line-specific):
    • DMEM (Gibco 11965092) + 10% FBS + 1% penicillin-streptomycin (100 U/mL pen, 100 ug/mL strep) — for HEK293, HeLa, NIH 3T3
    • RPMI 1640 (Gibco 11875093) + 10% FBS + 1% pen/strep — for Jurkat, THP-1, K562, most lymphoid/myeloid lines
    • F-12K (Gibco 21127022) + 10% FBS + 1% pen/strep — for A549, CHO
  • Fetal bovine serum (FBS), heat-inactivated (56 degrees C for 30 minutes)
  • Trypsin-EDTA (0.05% or 0.25%, Gibco 25200056)
  • Dulbecco's phosphate-buffered saline (DPBS), without calcium and magnesium (Gibco 14190144)
  • Trypan blue (0.4% solution)
  • Dimethyl sulfoxide (DMSO), cell culture grade
  • Freezing medium: 90% FBS + 10% DMSO (or commercial CryoStor CS10)

Equipment

  • Class II biological safety cabinet (BSC)
  • CO2 incubator (37 degrees C, 5% CO2, >95% humidity)
  • Inverted phase-contrast microscope
  • Hemocytometer or automated cell counter
  • Water bath (37 degrees C)
  • Centrifuge with swinging-bucket rotor
  • Mr. Frosty freezing container (or controlled-rate freezer)
  • Liquid nitrogen storage tank
  • Tissue culture-treated flasks (T25, T75, T175) or plates

Step-by-Step Cell Culture Protocol

Step 1: Prepare Your Workspace

  1. Turn on the BSC at least 15 minutes before starting to establish laminar airflow.
  2. Wipe down all surfaces inside the BSC with 70% ethanol.
  3. Spray all items (media bottles, pipettes, flasks) with 70% ethanol before placing them in the BSC.
  4. Pre-warm complete medium and trypsin-EDTA to 37 degrees C in the water bath (15-20 minutes).

Aseptic technique is non-negotiable. Every culture contamination traces back to a lapse in sterile practice.

Step 2: Thawing Frozen Cells

  1. Remove the cryovial from liquid nitrogen storage and immediately place it in a 37 degrees C water bath.
  2. Gently swirl the vial until only a small ice crystal remains (~1-2 minutes). Do not let the vial fully warm.
  3. Spray the vial with 70% ethanol and transfer to the BSC.
  4. Gently transfer the cell suspension to a 15 mL conical tube containing 9 mL pre-warmed complete medium.
  5. Centrifuge at 200 x g for 5 minutes at room temperature.
  6. Aspirate the supernatant carefully (this removes DMSO).
  7. Resuspend the pellet in 5-10 mL complete medium and transfer to an appropriately sized flask (T25 for small stocks, T75 for larger).
  8. Place in the incubator at 37 degrees C, 5% CO2.
  9. Check cells under the microscope after 24 hours. Replace medium to remove dead cells and residual DMSO.

Step 3: Routine Passaging (Adherent Cells)

Passage cells when they reach 70-90% confluency. Over-confluent cells undergo contact inhibition, altered gene expression, and senescence.

  1. Aspirate spent culture medium.
  2. Wash the monolayer once with pre-warmed DPBS (use 5 mL for T75, 10 mL for T175).
  3. Add pre-warmed trypsin-EDTA (1 mL for T25, 2 mL for T75, 4 mL for T175).
  4. Incubate at 37 degrees C for 2-5 minutes. Check under the microscope — cells should round up and detach. Tap the flask gently to dislodge remaining cells.
  5. Neutralize trypsin by adding 2-3x volume of complete medium (FBS inactivates trypsin).
  6. Transfer the cell suspension to a 15 mL conical tube.
  7. Centrifuge at 200 x g for 5 minutes.
  8. Aspirate the supernatant and resuspend the pellet in fresh complete medium.
  9. Count cells using trypan blue exclusion (mix 10 uL cells + 10 uL 0.4% trypan blue; count in hemocytometer). Viability should be >95%.
  10. Seed cells at the recommended density:
    • HEK293T: 2-4 x 10^4 cells/cm2
    • HeLa: 1-3 x 10^4 cells/cm2
    • CHO: 2-5 x 10^4 cells/cm2
  11. Add complete medium to the appropriate volume and place in the incubator.

Typical passage ratios: 1:3 to 1:10 depending on growth rate. Passage every 2-4 days.

Step 4: Routine Passaging (Suspension Cells)

  1. Transfer the cell suspension to a conical tube.
  2. Count cells and assess viability.
  3. Centrifuge at 200 x g for 5 minutes.
  4. Resuspend in fresh medium at the recommended seeding density:
    • Jurkat: 1-3 x 10^5 cells/mL
    • K562: 2-5 x 10^5 cells/mL
    • THP-1: 2-4 x 10^5 cells/mL
  5. Transfer to a new flask and return to the incubator.

Step 5: Cryopreservation

Always freeze cells at a low passage number and high viability (>90%).

  1. Harvest cells as in passaging steps above.
  2. Count cells and centrifuge at 200 x g for 5 minutes.
  3. Resuspend at 1-5 x 10^6 cells/mL in freezing medium (90% FBS + 10% DMSO).
  4. Aliquot 1 mL per cryovial.
  5. Place vials in a Mr. Frosty container (pre-chilled to 4 degrees C with isopropanol) and transfer to -80 degrees C overnight. This achieves a cooling rate of -1 degree C per minute, which is critical for cell viability.
  6. Transfer to liquid nitrogen the next day for long-term storage.

Contamination Prevention

Bacterial/Fungal Contamination

  • Signs: Turbid medium, pH shift (yellow medium), visible particles or filaments under microscope.
  • Prevention: Strict aseptic technique. Use pen/strep in routine culture. Wipe incubator shelves with 70% ethanol monthly.
  • Treatment: Discard contaminated cultures. Decontaminate the incubator if contamination is widespread.

Mycoplasma Contamination

  • Signs: Often invisible — no turbidity, no pH change. Causes subtle changes in growth rate, gene expression, and transfection efficiency.
  • Prevention: Test all new cell lines with a mycoplasma detection kit (PCR-based, e.g., Lonza MycoAlert) before introducing them to your incubator. Test every 1-3 months.
  • Treatment: BM-Cyclin (Roche) or Plasmocin (InvivoGen) can sometimes eliminate mycoplasma, but the safest approach is to discard and thaw a clean stock.

Cross-Contamination

  • The problem: HeLa cells are notorious for contaminating other lines. Studies show ~15-20% of cell lines in repositories are misidentified.
  • Prevention: Authenticate cell lines by STR profiling. Work with only one cell line at a time in the BSC. Never share media bottles between cell lines.

Common Mistakes to Avoid

  1. Over-trypsinizing — extended trypsin exposure damages surface proteins and reduces viability. Check cells at 2-minute intervals.
  2. Letting cells overgrow — cells at 100% confluency behave differently than cells at 80%. Many experiments require log-phase growth.
  3. Forgetting to passage on weekends — fast-growing cells like HEK293T can overgrow in 3 days. Set weekend passage schedules.
  4. Using non-heat-inactivated FBS — complement proteins in FBS can kill sensitive cells. Always heat-inactivate at 56 degrees C for 30 minutes.
  5. Not recording passage number — cells drift genetically over passages. Most experiments should use cells within 5-20 passages of thawing.

Pro Tips

  • Batch-test FBS — request samples from vendors and test several lots side-by-side for growth rate and viability before committing to a large purchase.
  • Photograph confluency — take microscope photos at each passage to calibrate your eye for 70%, 80%, and 90% confluency.
  • Color of medium matters — phenol red in DMEM/RPMI indicates pH. Red-orange = healthy (pH ~7.4). Yellow = acidic (overgrown or contaminated). Purple = basic (CO2 too low or cap sealed too tight).
  • Gentle pipetting — avoid foaming when resuspending cells. Foam means shear stress which means dead cells.

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