Monthly Archives: February 2018

Passaging/subculturing cells

  • Cell culture is not static. Cells acquire changes when maintained for a long time in culture. Stressful condition accelerates such changes, which results in inconsistency in the experimental outcome. Therefore, it is important to maintain culture under specified culture condition.
  • Cells in culture grow and divide in presence of nutrients and proper physiological condition. As they grow, cell density in culture increases and culture becomes confluent. With the increase in cell density, cell-cell contact becomes more prominent, which affect the cell physiology in various ways, leading to transient (inhibition of cell division by contact inhibition in untransformed cells) or sometimes even permanent changes in cell characteristics (e.g., may induce differentiation in cells).
  • Moreover, highly confluent culture consumes nutrients from the medium quickly, causing nutrient deprived state. Nutrient deprived state results in poor proliferation and cell death, leading to accumulation of toxic products in the medium. Accumulation of toxic products further enhances cell death.
  • In order to avoid complications and attain reproducibility in cell culture-based experiments, cell culture must be maintained under a condition, which allows exponential growth. Therefore, it is necessary that culture must not reach 100% confluency (adherent cell line) or gets overcrowded (suspension culture). To reduce the confluency/cell density, cells are regularly diluted by transferring a small amount of cells from the existing culture to another culture dish containing fresh culture medium, where cells further grow. The process of transferring a small proportion of cell to another fresh tissue culture dish is called passaging or subculturing.
  • The procedure of passaging is dependent on growth mode of cells. Adherent cells need to be detached from the substratum for passaging. Many methods of detaching cells from the substratum have been developed. Enzymatic treatment (Trypsin, Collagenase) of cell detachment is the most common methods used in most laboratories for routine passaging of adherent cells. However, other methods, like mechanical means (using rubber policeman or vigorous shaking for semi-adherent cells) or treatment with chelators can also be used, which depend on the cell type or experimental requirement.
  • Enzymatic methods often aim to make single cell suspension of cells which require both disrupting the cell-substratum as well as cell-cell contacts. However, use of only mechanical means often results in small cell clumps.
  • Detached cells are further diluted with the fresh culture medium and placed into new culture dishes.
  • Non-adherent cell culture (suspension cell culture) is simply diluted with culture medium and placed in new culture dishes. However, sometimes cells are treated with the enzymatic solution to make single cells suspension for the experiment.

Subculturing adherent cells using trypsin-EDTA

  • Subculturing/passaging can be defined as preparation of fresh culture by transferring cells from an existing culture.
  • Subculturing is done by transferring a small amount of cells (usually 1/3 to 1/10 cells of the existing semi confluent culture) from an existing culture dish to a new culture dish containing fresh growth medium.
  • Trypsin-EDTA method, also referred to as  trypsinization, is a most commonly used method for passaging adherent cells.
  • Trypsin-EDTA method of subculturing of a cell culture involves following steps.
    • Washing of cells with Ca2+- free  and Mg2+ – free PBS
    • Trypsin – EDTA treatment
    • Inactivation of trypsin
    • Preparation of fresh culture dish from the cell suspension
Washing of cells with Ca2+– free  and Mg2+ – free PBS
  • This step involves removing old culture medium from the culture dish, followed by washing with PBS which is free of Ca2+- free  and Mg2+ ions.
  • This step is intended to remove divalent cations and serum-containing medium from the cell culture. Serum in culture medium has trypsin inactivating activity (trypsin inhibitors) and divalent cations strengthen the cell-cell and cell-substratum interaction by stabilizing them.
Trypsin – EDTA treatment
  • This step involves brief incubation (few minutes, varies from 1 – 5 min for most cell lines) of adherent cell culture with Trypsin EDTA solution at 37°C.
  • This step is intended to disrupt both cell-cell and cell-substratum interactions. These interactions are mediated by various proteins (cadherins, integrins, extracellular matrix proteins like fibronectin, vitronectin) and their interactions are strengthen by divalent cations (e.g., Fibronectin-integrin interactions is promoted by Ca2+).
  • Trypsin, a serine protease, cleaves the polypeptide at C-terminal of lysine or arginine amino acid, except when either is followed by proline. Trypsin shows optimal activity at 37°C and pH 8.0.
  • EDTA, a chelating agents, sequesters divalent cations (e.g., Ca2+, Mg2+).
  • Trypsin disrupts cell-cell and cell substratum interactions by digesting proteins and EDTA weakened these interaction by chelating divalent cations.
Inactivation of trypsin
  • Since trypsin digests proteins, excessive trypsin treatment can cause high cell death by disrupting the plasma membrane. Therefore, inactivation of trypsin is an essential step in this method.
  • Usually trypsin is inactivated by adding serum-containing growth medium. In specific conditions where serum can not be added, other trypsin inhibitors, e.g., soybean trypsin inhibitor, are used.
Preparation of fresh culture dish from the cell suspension
  • This step aimed to distribute cell suspension into fresh culture dishes. Usually when the purpose is to maintain a culture in a healthy state, a rough estimation of cells called split ratio is used to distribute cells to fresh culture dishes. Split ratio suggest that how many culture dishes can be prepared from the existing culture dish. For example you can prepare 4 – 6 culture dishes if a recommended split ratio is 1:4 to 1:6  for a specific cell line. Alternatively calls can be counted and a specified number of cells can be transferred to another fresh flask containing  medium.

Preparation of Ethidium Bromide stock solution (10 mg/ml)


  • Ethidium bromide (EtBr) is a commonly used fluorescent stain to visualize nucleic acid especially DNA in agarose gels.
  • Ethidium bromide intercalates between DNA bases. Upon intercalation, its fluorescence increases several folds (25 fold increase when it binds DNA), much higher than the unbound ethidium bromide fluorescence, therefore, eliminates the need to wash gel to remove unbound ethidium bromide.
  • When exposed to uv light, it appears bright pink/orange colour.
  • Ethidium bromide can detect as little as 1 ng DNA/band in agarose gel.
  • It can also bind to RNA which results in 21 fold increase in its fluorescence intensity.
  • It has been reported to bind to single-stranded DNA.
  • Ethidium bromide powder is quite stable at room temperature but need to be protected from exposure of light.


  • Reagents
    • Ethidium bromide
    • Deionized / Milli-Q water
  • Equipment and disposables
    • Measuring cylinder
    • Conical flask/Beaker/15-ml screw-cap graduated polypropylene centrifuge tube
    • Magnetic stirrer / Tube-Rotator


  • 10 mg/ ml ethidium bromide in water


  • Preparation of 10 ml of 10 mg/ml ethidium bromide solution in water


General precaution:

Wear gloves and lab coat at all times when handling ethidium bromide or ethidium bromide contaminated solutions, glassware, pipette tips, and so forth.

Step 1: Weigh out 100 mg ethidium bromide conical flask / beaker / 15-ml polypropylene centrifuge tube. Add 7 – 8 ml water.

  • We recommend you to use 15-ml screw-cap graduated polypropylene centrifuge tube. Since these tubes have milliliter marks, you can adjust the solution volume without transferring the solution to measuring cylinder. Moreover, you don’t need to contaminate reusable vessels (beaker).

Step 2: Mix until all ethidium bromide dissolves completely. This may take long time.

  • Use tube rotator to mix all content if you 15-ml screw-cap graduated polypropylene centrifuge tube is used.
  • Cover the tube with aluminium foil to protect ethidium Bromide from light exposure.

Step 3: Adjust the volume to 10 ml with Deionized / Milli-Q water.

  • The solution will appear red.

Ethidium bromide solution is ready for use.


  • Ethidium bromide is quite stable and can be stored at 4°C for many years if protected from light.


  • Detection of DNA and RNA on agarose gel.
  • Purification of supercoiled DNA using cesium chloride – ethidium bromide gradient centrifugation method

Table: Migration of bromophenol blue and xylene cyanol on agarose gel in TBE and TAE electrophoresis buffer

Position of bromophenol blue and xylene cyanol in agarose gel in relation to the position of double standard DNA fragment in TAE (1x) and TBE (0.5 x) electrophoresis buffer.
For example in 0.5% agarose gel, Bromophenol blue migrates at approximately 750 bp long double standard DNA fragment in TBE buffer and at approximately 1150 bp long double standard DNA fragment in TAE buffer.
Agarose gel, % Bromophenol blue in TBE buffer (0.5x) Xylene cyanol FF) in TBE buffer (0.5x) Bromophenol blue in TAE buffer (1x) Xylene cyanol FF in TAE buffer (1x)
0.5 750 13000 1150 16700
0.6 540 8820 850 11600
0.7 410 6400 660 8500
0.8 320 4830 530 6500
0.9 260 3770 440 5140
1.0 220 3030 370 4160
1.2 160 2070 275 2890
1.5 110 1300 190 1840
2.0 65 710 120 1040

Preparation of 3 M Sodium acetate (CH3COONa), pH 5.2 solution from CH3COOH.3H2O


  • Sodium acetate is a sodium salt of acetic acid.
  • It is water-soluble.
Related Content


  • Reagents
    • Sodium acetate, trihydrate (CH3COONa.3H2O) (Molecular weight = 136.08)
    • Deionized / Milli-Q water
  • Equipment and disposables
    • Measuring cylinder
    • Conical flask / Beaker
    • Magnetic stirrer


  • 3 M Sodium acetate


Preparation of 100 ml of 3M Sodium acetate solution, pH 5.2 in water from sodium acetate trihydrate (CH3COONa.3H2O)


Step 1: To prepare 100 ml aqueous solution of 3M Sodium acetate (CH3COONa), weigh out 40.82 gram CH3COONa.3H2O (Molecular weight = 136.08). Transfer it to 250 ml beaker/conical flask. Add 80 ml deionized / Milli-Q water. Mix until all sodium acetate dissolves completely.

  • One can use manual shaking using a glass pipette to mix the ingredients. Magnetic stirrer makes the dissolving process easy and convenient.
  • Do not dissolve in 100 ml of deionized / Milli-Q water. In most cases, solution volume increases when a large amount of solute dissolves in the solvent.

Step 2: Adjust the pH to 5.2 with glacial acetic acid.

  • Since pH is dependent on temperature, one should adjust pH at room temperature (25°C).

Step 3: Adjust the volume to 100 ml with deionized / Milli-Q water. Mix it again.

  • The solution will appear colorless and transparent.

Step 4: Transfer the solution to autoclavable bottle. Sterilize the solution by autoclaving (20 minutes at 15 lb/ (psi) from 121-124°C on liquid cycle).

  • Depending on the consumption, one can make small aliquots of the solution.
  • One can sterilize a solution by passing through 0.2μ filter unit. Filter sterilization removes all suspended particles with size more than 0.2 μ which includes most bacteria and their spores but not mycoplasma. Moreover, it does not inactivate enzyme activities (e.g., DNases). Autoclaving inactivates most enzymes except some (e.g., RNases) and kills most microorganisms including mycoplasma.


Solution can be stored at room temperature.


  • Precipitation of nucleic acid (DNA and RNA)
  • Buffer solution
  • Protein crystallization
Follow the table, to prepare Sodium acetate (CH3COONa) solution of specific concentration and volume from CH3COONa.3H2O.
Conc. / Volume 10 ml 50 ml 100 ml 250 ml 500 ml 1000 ml
0.1 M 0.136 g 0.68 g 1.36 g 3.40 g 6.80 g 13.61 g
0.5 M 0.68 g 3.40 g 6.80 g 17.01 g 34.02 g 68.04 g
1 M 1.36 g 6.80 g 13.61 g 34.02 g 68.04 g 136.08 g
3 M 4.08 g 20.41 g 40.82 g 102.06 g 204.12 g 408.24 g