In the world of cell biology, cell passaging (also known as subculturing or splitting cells) is the bread and butter of laboratory work. Whether you are working with HEK293, HeLa, or primary fibroblast lines, maintaining a healthy, proliferating population requires a consistent and sterile subculturing routine.
This article provides a comprehensive, 1000-word deep dive into the standard protocol for adherent cell passaging. We will cover everything from hood preparation to the enzymatic detachment of cells, ensuring your cultures remain viable for downstream applications like transfection, drug screening, or CRISPR editing.
1. Why Do We Passage Cells?
Adherent cells grow until they cover the available surface area of a culture vessel—a state known as confluency. Once cells reach 80–90% confluency, several physiological changes occur:
Nutrient Depletion: The media becomes acidic (turning yellow) as nutrients are consumed.
Contact Inhibition: Many cell lines stop dividing once they touch neighboring cells.
Metabolic Waste: The accumulation of lactic acid and other byproducts can become toxic.
Passaging involves transferring a small portion of these cells into a new vessel with fresh growth media, effectively “resetting” the growth clock and allowing for continued expansion.
2. Aseptic Technique: Preparing the Workspace
Contamination is the greatest enemy of the cell biologist. Before a single bottle is opened, the environment must be controlled.
Hood Preparation
Airflow: Open the cell culture hood and turn on the blower and light. Allow the hood to run for at least 10–15 minutes to establish a laminar flow that keeps contaminants out.
Sterilization: Spray the work surface generously with 70% ethanol. Wipe the surface thoroughly from back to front.
Material Entry: Every item placed inside the hood—pipette tip boxes, media bottles, and the culture plates themselves—must be wiped down with 70% ethanol.
3. Step-by-Step Subculturing Protocol
Step 1: Media Removal
Remove your cell culture plate from the incubator and examine it under a microscope to confirm confluency and the absence of contamination. Once verified, place it in the hood. Use a sterile glass Pasteur pipette connected to a vacuum pump to vacuum out the spent media from the corner of the plate.
Step 2: The PBS Wash
Residual serum in the growth media contains trypsin inhibitors. If you do not wash the cells, the trypsin added in later steps will be neutralized and fail to detach the cells.
Add 1–2 ml of sterile PBS (Phosphate-Buffered Saline).
Technical Tip: Always add PBS gently against the side wall of the plate rather than directly onto the cell monolayer. Adding liquid directly to the cells can cause them to peel off prematurely.
Gently rotate the plate to ensure the PBS covers the entire surface, then pipette or vacuum out the PBS.
Step 3: Enzymatic Detachment (Trypsinization)
To transition cells from an adherent state to a suspension state, we use Trypsin-EDTA. Trypsin is a protease that breaks down the proteins adhering the cells to the plastic.
Add 1 ml of Trypsin. Unlike PBS, you can add this more directly to ensure it reaches all areas of the monolayer.
Rock the plate to spread the liquid evenly.
Incubation: Place the plate back into the $CO_2$ incubator (usually at 37°C).
Timing: Incubate for approximately 10 minutes. However, this is cell-line dependent. Check the cells under the microscope after 2–3 minutes. If they appear rounded and are floating when the plate is tapped, they are ready.
Step 4: Neutralization and Trituration
Once the cells are detached, you must stop the enzymatic reaction to prevent damage to the cell membranes.
Add 1–2 ml of complete culture medium (which contains serum). The serum contains alpha-1-antitrypsin, which neutralizes the trypsin.
Trituration: Use a pipette to gently suck up and expel the media several times. This mechanical action breaks up cell clumps, ensuring a single-cell suspension. This is vital for accurate downstream counting and even distribution in the new plate.
Step 5: Seeding the New Vessel
Concentration Adjustment: Discard the excess medium, keeping roughly 0.3 – 0.5 ml of the concentrated cell suspension in the plate (or transfer the suspension to a centrifuge tube if a full media change is required).
Dilution: Add fresh growth medium to reach a final volume of 8–10 ml (for a standard 10cm dish).
Distribution: Rotate the plate in a “figure-eight” motion on the benchtop. This ensures the cells are spread evenly across the surface. If you simply place the plate in the incubator without rotating, the cells will gravitate toward the center, leading to uneven growth.
4. Post-Passaging Care and Incubation
Place the newly seeded plates into the $CO_2$ incubator. The standard environment is:
Temperature: 37°C
$CO_2$ Concentration: 5% (to maintain the bicarbonate buffer system in the media)
Humidity: High humidity to prevent evaporation of the media.
Check your cells 24 hours later to ensure they have re-attached and are beginning to proliferate.
Creating Stable Cell Lines: Step-by-Step Protocol, Applications, and FAQs
5. Troubleshooting Common Issues
| Issue | Potential Cause | Solution |
| Cells won’t detach | Trypsin was neutralized by residual serum. | Ensure a thorough PBS wash before adding Trypsin. |
| High cell death after split | Trypsin incubation was too long. | Monitor cells every 2 minutes; remove once they round up. |
| Uneven cell distribution | Poor mixing/rotation after seeding. | Use a crosswise or figure-eight motion before incubating. |
| Contamination | Poor aseptic technique. | Re-evaluate hood cleaning and ethanol wiping protocols. |
6. Summary Checklist for Success
[ ] Hood cleaned with 70% ethanol.
[ ] Cells washed with PBS to remove serum inhibitors.
[ ] Trypsin added and incubated at 37°C.
[ ] Reaction neutralized with serum-containing media.
[ ] Single-cell suspension achieved via pipetting (trituration).
[ ] Cells seeded and rotated for even distribution.
By mastering these fundamental steps, you ensure the longevity and reliability of your cell lines, providing a solid foundation for all your biological experiments.
FAQ: Frequently Asked Questions
How often should I passage my cells?
Most immortalized lines (like HEK293) require passaging every 2–3 days. If the media turns orange or yellow, or if the cells look crowded under the microscope, it is time to split.
Can I passage cells without Trypsin?
Some sensitive cells require “cell scrapers” or non-enzymatic dissociation buffers. However, for most robust adherent lines, Trypsin-EDTA is the standard choice.
What is the ideal “split ratio”?
A common ratio is 1:5 or 1:10. This means if you have 1 ml of cell suspension, you put 0.2 ml into a new plate and add 9.8 ml of fresh media.
References
ATCC – Basic Techniques in Mammalian Cell Culture.
Thermo Fisher Scientific – Cell Culture Basics Handbook.
NCBI – Maintenance of Cell Lines: Principles and Practice.
