Kulture Scientific · Technical Protocol
Passage Like a Pro
Standard subculturing protocol for adherent mammalian cell lines
Kulture Scientific · Technical Protocol
Passage Like a Pro
Standard subculturing protocol for adherent mammalian cell lines
This standard passaging protocol covers everything you need to reliably subculture adherent mammalian cell lines — from pre-warming reagents to seeding the new vessel. Whether you are working with T-flasks, multi-well plates, or dishes, the principles of aseptic technique, confluency assessment, and trypsin-based dissociation apply across your culture system. Use this protocol as your lab standard or as a training reference for new team members.
Purpose and Scope
This protocol establishes aseptic technique standards for routine subculturing of adherent mammalian cell lines. It applies to standard culture vessels including T-flasks, multi-well plates, and dishes. Its purpose is twofold: to prevent contamination from bacteria, fungi, mycoplasma, and other cell lines, and to protect anyone who may come into contact with culture materials or shared lab surfaces.
Precautions
Microbial contamination is one of the most common and consequential risks in cell culture work. Bacteria, mycoplasma, yeast, and fungal spores can be introduced through the operator, the surrounding atmosphere, work surfaces, and reagent solutions. Strict aseptic technique is the primary defense.
Contamination can spread from a single flask to an entire incubator. Early signs include turbidity in the medium or an unexpected pH shift. Always confirm suspected contamination by microscopy before discarding cultures.
The following practices significantly reduce contamination risk:
Materials Needed
Preparing Complete Growth Medium
Before beginning any cell culture procedure, confirm the specific media requirements for your cell line: basal media formulation, required supplements, and recommended working volumes for your culture vessel.
Basal media is typically supplemented with 5–20% bovine serum depending on cell line requirements, and may include antibiotics such as penicillin/streptomycin, growth factors, or amino acid supplements. Note that free L-glutamine in liquid medium degrades rapidly — significant loss can occur within 1–2 weeks at 4°C. If your formulation contains L-glutamine, consider a stable dipeptide alternative (such as GlutaMAX or GlutaPlus) that maintains activity throughout the shelf life of your medium. Complete growth medium is generally stable for up to 6 weeks when stored at 4°C.
Examining Your Cells
Cells should be examined by inverted microscopy regularly to confirm they are healthy and growing as expected. Healthy adherent cells are predominantly attached to the culture surface, appearing round and plump or elongated depending on cell type, with visible light refraction at the cell membrane.
At each observation, record the following:
Identifying contamination
Contamination should be confirmed through visual inspection, microscopic examination, and where appropriate, biochemical testing. Key signs include:
| Sign | What to look for |
|---|---|
| Turbidity | Cloudy or opaque medium — often the first visible indicator of bacterial or yeast contamination |
| pH shift | Rapid color change of phenol red toward yellow (acidic) or pink/purple (alkaline) without a corresponding increase in cell density |
| Microscopic | Visible rods, cocci, or budding yeast cells; unusual granularity or vacuolation in cells |
| Mycoplasma | No visible signs — requires PCR, ELISA, or fluorescence-based testing. Suspect if cells show unexplained growth changes or reduced proliferation. |
| Cell morphology | Cells appearing shriveled, dark, grainy, or detaching in large numbers without mechanical cause |
Routine Maintenance: Split Ratios
Split ratios for routine maintenance passaging are cell-line dependent and should always be determined by consulting the guidelines specific to your cell line. Slow-growing lines may fail to recover at high split ratios, while fast-growing lines may require higher ratios to prevent overgrowth between passages.
As a general principle, passage adherent cells when they reach 70–90% confluency. If subculturing is necessary before that threshold, use a lower split ratio to ensure the seeding density is sufficient for survival and recovery.
Sub-culturing and Passaging
Prepare your workspace and pre-warm reagents
Pre-warm calcium- and magnesium-free PBS, dissociation reagent, and complete growth medium in a 37°C water bath for at least 15 minutes. Spray the biosafety cabinet with 70% ethanol and allow to dry before beginning.
Check confluency
Passage cells when they reach 70–90% confluency. Always verify under the inverted microscope before proceeding — never go by time or schedule alone. Not sure what 70–90% looks like for your cell type? See the Visual Guide to Confluency.
Remove spent media
Aspirate and discard the spent cell culture media from the culture vessel.
Wash with calcium- and magnesium-free PBS
Add approximately 2 mL of PBS per 10 cm² of cultured surface area. Gently add the wash solution to the side of the vessel opposite the cell layer to avoid disturbing the monolayer. Rock gently to distribute, then aspirate and discard. Adherent cells may be washed twice.
Add dissociation reagent
Add pre-warmed dissociation reagent (e.g., 0.25% Trypsin-EDTA) to the side of the vessel — approximately 0.5 mL per 10 cm². Rock gently to achieve complete coverage of the cell layer.
Incubate and monitor detachment
Incubate at room temperature or 37°C for approximately 2–5 minutes (cell-line dependent). Check by inverted microscopy; cells should appear rounded and begin detaching. If less than 90% have detached, extend incubation by a few minutes, checking every 30 seconds.
Neutralize and collect cell suspension
Aspirate the cell suspension and transfer to a conical tube. Add an equivalent volume of pre-warmed complete growth medium to the culture vessel, disperse by pipetting over the cell layer surface several times, then transfer to the same conical tube. Complete media neutralizes the dissociation reagent and ensures full cell recovery.
Centrifuge
If removal of the dissociation reagent is required for your cell line, centrifuge at 300–500 × g for 5–10 minutes. Centrifuge speed and time are cell-type dependent.
Resuspend and count
Carefully aspirate the supernatant without disturbing the pellet. Resuspend the cell pellet in a minimal volume of pre-warmed complete growth medium. Remove a sample for counting using a hemocytometer or automated cell counter with Trypan blue exclusion to assess viability.
Dilute, seed, and label
Dilute the cell suspension to the seeding density recommended for your cell line. Pipette the appropriate volume into the new culture vessel. Label immediately with cell line, passage number, date, and initials. Return to the incubator.
Changing Media
If cells are growing well but have not yet reached passaging confluency, a media change may be needed to replenish nutrients. Indicators include a high number of cells in suspension or a visible acidic shift in medium color (phenol red indicator turning yellow).
Pre-warm complete media to 37°C before use. Carefully aspirate spent media and discard. Replace with fresh pre-warmed complete media and return the vessel to the incubator.
Troubleshooting
| Issue | Likely Cause | Corrective Action |
|---|---|---|
| Cells won’t detach | Dissociation reagent too cold or incubation too short | Ensure reagent is fully pre-warmed; extend incubation and check every 30 seconds |
| Cells detaching in clumps | Culture was over-confluent at time of passage | Passage earlier next time; pipette more vigorously to break clumps into single-cell suspension |
| Low viability after passage | Over-exposure to dissociation reagent or rough pipetting | Reduce incubation time; pipette gently to minimize shear damage |
| Cells not attaching | Dissociation reagent not fully neutralized | Use more complete media to neutralize; ensure centrifuge spin is complete before resuspending |
| Turbid media or pH shift | Microbial contamination | Remove from incubator immediately; confirm by microscopy; discard if confirmed |
| Slow or no growth | Seeding density too low, media issue, or mycoplasma | Verify seeding density; check media preparation; test for mycoplasma if problem persists |
| Cells look grainy or dark | Cell stress, high passage number, or contamination | Check passage number against line limits; verify media and CO₂ levels; test for mycoplasma |
Notes
Frequently Asked Questions
This standard passaging protocol covers everything you need to reliably subculture adherent mammalian cell lines — from pre-warming reagents to seeding the new vessel. Whether you are working with T-flasks, multi-well plates, or dishes, the principles of aseptic technique, confluency assessment, and trypsin-based dissociation apply across your culture system. Use this protocol as your lab standard or as a training reference for new team members.
Purpose and Scope
This protocol establishes aseptic technique standards for routine subculturing of adherent mammalian cell lines. It applies to standard culture vessels including T-flasks, multi-well plates, and dishes. Its purpose is twofold: to prevent contamination from bacteria, fungi, mycoplasma, and other cell lines, and to protect anyone who may come into contact with culture materials or shared lab surfaces.
Precautions
Microbial contamination is one of the most common and consequential risks in cell culture work. Bacteria, mycoplasma, yeast, and fungal spores can be introduced through the operator, the surrounding atmosphere, work surfaces, and reagent solutions. Strict aseptic technique is the primary defense.
Contamination can spread from a single flask to an entire incubator. Early signs include turbidity in the medium or an unexpected pH shift. Always confirm suspected contamination by microscopy before discarding cultures.
The following practices significantly reduce contamination risk:
Materials Needed
Preparing Complete Growth Medium
Before beginning any cell culture procedure, confirm the specific media requirements for your cell line: basal media formulation, required supplements, and recommended working volumes for your culture vessel.
Basal media is typically supplemented with 5–20% bovine serum depending on cell line requirements, and may include antibiotics such as penicillin/streptomycin, growth factors, or amino acid supplements. Note that free L-glutamine in liquid medium degrades rapidly — significant loss can occur within 1–2 weeks at 4°C. If your formulation contains L-glutamine, consider a stable dipeptide alternative (such as GlutaMAX or GlutaPlus) that maintains activity throughout the shelf life of your medium. Complete growth medium is generally stable for up to 6 weeks when stored at 4°C.
Examining Your Cells
Cells should be examined by inverted microscopy regularly to confirm they are healthy and growing as expected. Healthy adherent cells are predominantly attached to the culture surface, appearing round and plump or elongated depending on cell type, with visible light refraction at the cell membrane.
At each observation, record the following:
Identifying contamination
Contamination should be confirmed through visual inspection, microscopic examination, and where appropriate, biochemical testing. Key signs include:
| Sign | What to look for |
|---|---|
| Turbidity | Cloudy or opaque medium — often the first visible indicator of bacterial or yeast contamination |
| pH shift | Rapid color change of phenol red toward yellow (acidic) or pink/purple (alkaline) without a corresponding increase in cell density |
| Microscopic | Visible rods, cocci, or budding yeast cells; unusual granularity or vacuolation in cells |
| Mycoplasma | No visible signs — requires PCR, ELISA, or fluorescence-based testing. Suspect if cells show unexplained growth changes or reduced proliferation. |
| Cell morphology | Cells appearing shriveled, dark, grainy, or detaching in large numbers without mechanical cause |
Routine Maintenance: Split Ratios
Split ratios for routine maintenance passaging are cell-line dependent and should always be determined by consulting the guidelines specific to your cell line. Slow-growing lines may fail to recover at high split ratios, while fast-growing lines may require higher ratios to prevent overgrowth between passages.
As a general principle, passage adherent cells when they reach 70–90% confluency. If subculturing is necessary before that threshold, use a lower split ratio to ensure the seeding density is sufficient for survival and recovery.
Sub-culturing and Passaging
Prepare your workspace and pre-warm reagents
Pre-warm calcium- and magnesium-free PBS, dissociation reagent, and complete growth medium in a 37°C water bath for at least 15 minutes. Spray the biosafety cabinet with 70% ethanol and allow to dry before beginning.
Check confluency
Passage cells when they reach 70–90% confluency. Always verify under the inverted microscope before proceeding — never go by time or schedule alone. Not sure what 70–90% looks like for your cell type? See the Visual Guide to Confluency.
Remove spent media
Aspirate and discard the spent cell culture media from the culture vessel.
Wash with calcium- and magnesium-free PBS
Add approximately 2 mL of PBS per 10 cm² of cultured surface area. Gently add the wash solution to the side of the vessel opposite the cell layer to avoid disturbing the monolayer. Rock gently to distribute, then aspirate and discard. Adherent cells may be washed twice.
Add dissociation reagent
Add pre-warmed dissociation reagent (e.g., 0.25% Trypsin-EDTA) to the side of the vessel — approximately 0.5 mL per 10 cm². Rock gently to achieve complete coverage of the cell layer.
Incubate and monitor detachment
Incubate at room temperature or 37°C for approximately 2–5 minutes (cell-line dependent). Check by inverted microscopy; cells should appear rounded and begin detaching. If less than 90% have detached, extend incubation by a few minutes, checking every 30 seconds.
Neutralize and collect cell suspension
Aspirate the cell suspension and transfer to a conical tube. Add an equivalent volume of pre-warmed complete growth medium to the culture vessel, disperse by pipetting over the cell layer surface several times, then transfer to the same conical tube. Complete media neutralizes the dissociation reagent and ensures full cell recovery.
Centrifuge
If removal of the dissociation reagent is required for your cell line, centrifuge at 300–500 × g for 5–10 minutes. Centrifuge speed and time are cell-type dependent.
Resuspend and count
Carefully aspirate the supernatant without disturbing the pellet. Resuspend the cell pellet in a minimal volume of pre-warmed complete growth medium. Remove a sample for counting using a hemocytometer or automated cell counter with Trypan blue exclusion to assess viability.
Dilute, seed, and label
Dilute the cell suspension to the seeding density recommended for your cell line. Pipette the appropriate volume into the new culture vessel. Label immediately with cell line, passage number, date, and initials. Return to the incubator.
Changing Media
If cells are growing well but have not yet reached passaging confluency, a media change may be needed to replenish nutrients. Indicators include a high number of cells in suspension or a visible acidic shift in medium color (phenol red indicator turning yellow).
Pre-warm complete media to 37°C before use. Carefully aspirate spent media and discard. Replace with fresh pre-warmed complete media and return the vessel to the incubator.
Troubleshooting
| Issue | Likely Cause | Corrective Action |
|---|---|---|
| Cells won’t detach | Dissociation reagent too cold or incubation too short | Ensure reagent is fully pre-warmed; extend incubation and check every 30 seconds |
| Cells detaching in clumps | Culture was over-confluent at time of passage | Passage earlier next time; pipette more vigorously to break clumps into single-cell suspension |
| Low viability after passage | Over-exposure to dissociation reagent or rough pipetting | Reduce incubation time; pipette gently to minimize shear damage |
| Cells not attaching | Dissociation reagent not fully neutralized | Use more complete media to neutralize; ensure centrifuge spin is complete before resuspending |
| Turbid media or pH shift | Microbial contamination | Remove from incubator immediately; confirm by microscopy; discard if confirmed |
| Slow or no growth | Seeding density too low, media issue, or mycoplasma | Verify seeding density; check media preparation; test for mycoplasma if problem persists |
| Cells look grainy or dark | Cell stress, high passage number, or contamination | Check passage number against line limits; verify media and CO₂ levels; test for mycoplasma |