Several methods have been developed for passaging cells. Each method has its own advantages and drawbacks. Always check the cell line instruction manual and relevant literature for the optimal procedure.
Most passaging methods aim to obtain single cell suspension by breaking cell-substratum and cell-cell contacts. The single cell suspension is further diluted by addition of fresh growth medium and allowed to grow in its optimal growth environment.
Depending on the experimental requirement, type and growth mode of cell culture, and nature of cell line, one should choose a particular method.
Depending on growth mode, cell culture can be adherent, semi-adherent and non-adherent. Since nonadherent cells grow in suspension, such culture can be passaged essentially by diluting the existing cell culture. In contrast, semi-adherent and firmly adherent cells need to be detached from the substratum for passaging.
Several methods have been developed for passaging cell culture. These methods can be broadly classified into three types based on the nature of treatment used to dissociate cells.
Enzymatic methods are routinely used in most cell culture laboratories for passaging adherent cells.
A brief treatment of cells with enzyme solution and subsequent gentle pipetting results in single cell suspension.
Enzymatic methods are very efficient at disrupting both cell-substratum as well as cell-cell contacts, leading to single cell suspension. Cell number can be determined easily in single cell suspension, therefore, enzymatic methods are valuable for those experiments where accurate cell number is one of the requirements of experimental procedure. However, enzymatic treatments results in digestion and cleavage of cell surface proteins (e.g., receptors), therefore, not suitable for experiments which aims to analyze cell surface proteins.
The sensitivity of cell lines for a particular enzymatic method may differ. Therefore, one needs to standardize treatment condition. Overtreatment with enzyme solution can lead to cell death, whereas, under treatment can cause inefficient detachment of cells and lead to cell clumps.
Often inhibitors (e.g., serum, soybean trypsin inhibitor etc.) are added to stop the enzyme action.
Mechanical means (Scraping by rubber policeman, Dislodging semi-adherent cells by vigorous shaking):
Mechanical means involved disrupting cell-substratum interaction by mechanical force like scraping using rubber policeman or vigorously tapping the culture dish or extensively pipetting culture medium over cell layer.
This method often leads to cell death and is not recommended for tightly adherent cells.
Cell clumps are often observed therefore determining cell number using haemocytometer counting is difficult. However, this method is suitable for semi-adherent cells.
Treatment with chelators (e.g., EDTA or EGTA treatment)
Chelators sequester the divalent cations, e.g., Ca2+ or Mg2+, which are required for adhesion of cell to substratum as well as cell-cell contacts.
This method takes long time incubation with chelator solution. Most cell lines partially loses their contact with substratum and subsequently extensive pipetting or tapping of culture dishes detaches cells from the substratum.
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/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.
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.
Phenol-chloroform extraction can be used to isolate and purify DNA and RNA. It is extremely good at removing protein and lipid from the nucleic acid.
It is an example of liquid – liquid extraction which is based on differential solubilities of biomolecules (e.g., nucleic acids, proteins, carbohydrate, and lipids) in water and phenol/chloroform.
Since most proteins and lipids are highly soluble in phenol, they move to organic phase, whereas nucleic acid stay in aqueous phase due to its high solubility in water.
Phenol has slightly higher density (1.07 g/cm3) than water, but when chloroform is added which has comparatively very high density (1.47g/cm3), the density of the phenol-chloroform solution increases considerably. Therefore during extraction, organic phase is always present at the bottom, allowing aqueous phase to be transferred efficiently.
Solution of phenol and chloroform is not only more efficient at denaturing proteins but also reduces the partitioning of poly(A)+ mRNA into the the organic phase.
Generally a 1:1 ratio of phenol:chloroform is used for the extraction of nucleic acid. Sometime a small quantity of isoamyl alcohol is added in this mixture (25:24:1 ratio of Phenol: Chloroform: Isoamyl Alcohol). Isoamyl alcohol is a anti-foaming agent, thus prevents form-formation during the extraction process.
In this procedure, approximately equal volume of phenol:chloroform solution is added to the sample (aqueous solution DNA/RNA or cell lysate or tissue lysate) and mixed. When this mixture is centrifuged, solution is separated into two phases: upper aqueous phase and bottom organic phase (Phenol/Chloroform). At interface, insoluble material which is nothing but the cell debris and denatured proteins are collected. Aqueous phase which contains nucleic acid is then transferred to another fresh vial and subjected to ethanol or isopropanol precipitation to concentrate nucleic acid.
Tris saturated phenol (pH ≈8.0) is used for purification of DNA, whereas water saturated phenol (pH ≈4.8) is used for purification of RNA.
At slightly alkaline pH, phosphate backbone of both DNA and RNA is negatively charged, therefore, they remains in aqueous phase. As the pH of Phenol/Chloroform solution drops, DNA tend to move to organic phase and at pH 4.8 most DNA is present either at interphase (mostly large DNA fragments) or in the organic phase (smaller DNA fragments). The reason why DNA move to organic phase, but not RNA is because of higher pKa value of DNA, which results in neutralization of negatively charged phosphate backbone of DNA.
Since phenol is slightly soluble in water (≈7-8% in water), aqueous phase can be reextracted with chloroform alone to remove traces of phenol from the final preparation of nucleic acid. Traces of phenol can inhibit downstream enzymatic reactions and interfere with spectroscopic analysis.
Plasmid isolation is a routine task in most cell and molecular biology labs. It is an essential step in many procedures such as gene cloning, DNA sequencing, and transfection.
A good method of plasmid isolation must be rapid, economical and produce plasmid of high quality which can be used for multiple purposes including transfection in cell lines, sequencing, and cloning.
To isolate plasmid from the host bacteria, cells are first lysed, leading to release of plasmid and in subsequent steps plasmid is purified from the lysate.
There are number of methods available to lyse bacterial cells. Most common methods are alkaline lysis, boiling lysis, enzymatic lysis and lysis with detergents.
Purification of plasmid from the lysed cells mostly dependent on the type of lysis method used to release plasmid in solution. For example, alkaline lysis which completely disrupt the bacterial cells leading to release of cell components including both plasmid DNA and genomic DNA in denatured state, rely on selective renaturation of only plasmid DNA in a perfect manner at purification step. On the other hand, boiling lysis selectively releases only plasmid DNA from the bacterial cells.
Purified plasmid can be further purified by number of methods to obtain high quality of plasmids. These methods are selective precipitation in high salt SDS, centrifugation in gradients of CsCl – ethidium bromide (EtBr), extraction with Phenol-chloroform, and hydroxylapatite chromatography.
Following factors can be considered while choosing a method of plasmid isolation isolation……
Plasmid characteristics (copy number and size)
Quality and quantity of plasmid
Complexity, cost and rapidity of the method
The yield of plasmid DNA is governed by two most important parameters – copy number of plasmid and amount of initial culture taken to isolate plasmid DNA. Based on initial culture volume, plasmid isolation method can be termed as…….
Amplification of plasmid is desirable for many applications including gene cloning, DNA sequencing, transfection, and probe preparation. Fastest and routinely used method to amplify plasmid is to introduce plasmid in an appropriate strain of E. coli e.g. DH5α (the process is called transformation), grow them to a suitable culture volume, and finally, extract plasmid from them (the process called plasmid isolation).
Alternatively, E. coli DH5α harboring a plasmid can also be revived from the stored stocks e.g., glycerol stock or stab culture, if available. Cells obtained from stored stock can be either streaked or plated on an antibiotic containing solid LB-agar plate.
Plasmid copy number and culture volume are the two most important parameter which predicts the quantity of the plasmid extracted at the end of the isolation process. Comparatively, large culture volume is required for low copy number plasmid.
Plasmid copy number can be increased by chloramphenicol treatment. Several rich growth media can also be used to grow bacteria. These media support high cell density due to nutrient enrichment.
Depending on the initial culture volume, plasmid isolation methods are called miniprep (1-5 ml culture volume), midiprep (25-50 ml culture volume), and maxiprep (100-500 ml culture volume).
Generally miniprep yields sufficient amount of plasmid for applications like the screening of clones for the presence of insert, DNA sequencing etc. Other applications, like probe preparations, plasmid distribution, transfection, etc., can require a large quantity of plasmid (midiprep or maxiprep).
For miniprep, a single colony from the LB-agar plate is inoculated into a antibiotic-containing liquid medium. Culture is grown at 37°C in a shaker incubator overnight (12- 16 h). Grown culture corresponds to late log phase/early stationary phase of bacterial growth and is characterized by low content of RNA. Incubating culture for a long time can cause the death of bacteria, which can result in low yield of plasmid. Sometimes, a well-grown colony from the LB-agar plate can directly be utilized for plasmid miniprep.
For a large amount of culture which is required for midiprep and maxiprep, initially, a starter culture is prepared by inoculating a small amount of culture medium (2 – 10 ml) with a single colony. When the culture reaches mid- to late-exponential growth phase (takes 8 – 12 h), culture is diluted in a ratio of 1:100 to 1:1000 to prepare large culture volume for midiprep and maxiprep.
All plasmid vectors carry at least one antibiotic resistance gene, which enables bacteria to survive and grow in presence of a respective antibiotic. Antibiotic functions as a selective marker which allows growth of only plasmid containing E. coli cells. In absence of antibiotic, bacteria will lose the plasmid, which will result in low or no yield.
Synonyms: Pancreatic Ribonuclease, RNase A, Ribonucleate 3′-pyrimidinooligonucleotidohydrolase, Ribonuclease A from bovine pancreas Type I-A
Molecular weight: 13.7 kDa
Optimal temperature : 60°C (activity range of 15–70°C)
Ribonuclease A (RNase A) belongs to an endoribonuclease class of ribonucleases. In contrast to exoribonucleases which cleave/degrade RNA in 3’-5’ direction, endoribonucleases degrade RNA endoribonucleolytically in 5’-3’ direction.
RNase A is a digestive enzyme which is secreted by the pancreas to digest RNA. It is abundantly present in the pancreas, therefore, the pancreas is a valuable source for RNase A.
Mature bovine pancreatic RNase A only has 124 amino acids with molecular weight 13.7 kDa. It lacks tryptophan amino acid.
In contrast to others known members of endoribonuclease, RNase A is not a glycoprotein.
RNase A is active under a wide range of reaction conditions (temperature range 15 – 70°C; pH range 6–10). The optimal temperature for its activity is 60°C and optimal pH is 7.6.
RNase A is quite stable to both heat and detergents.
It cleaves both single-stranded and double-stranded RNA as well the RNA strand in RNA-DNA hybrids at a low salt concentration (0 to 100 mM NaCl). However, it specifically cleaves single-stranded RNA at higher salt concentration (0.3M NaCl or higher)
Preparation of RNase A solution:
Generally, a 10 mg/ml RNase A solution is prepared in 10 mM Tris.Cl (pH 7.5) or TE [(10 mM Tris.Cl (pH 7.6), 1 mM EDTA]. The stock is stable for at least 1 year at -20°C.
Most suppliers provide molecular biology grade RNase A powder which is free from any DNase activity. Sometimes, a low level of DNase activity in RNase stock solution can be detected which can be easily eliminated by incubating the stock solution in boiling water bath for 5 to 10 min (see protocol). Boiling RNase A solution for a short time does not inactivate RNase A, but is sufficient to inactivate DNase activity.
If the RNase A stock is suspected to have high DNase activity, it is recommended to prepare 10 mg/ml stock solution in sodium acetate (pH 5.2), incubate the solution in boiling water bath for 20 – 30 min, then adjust the pH with 1M Tris.Cl (pH 7.5). RNase A is comparatively very stable at low pH (between pH 2.0 – 4.5).
Cell culture is not static. Cells in culture acquire changes which can be either genetically programmed (e.g., senescence in primary culture) or due to accumulation of genetic abnormalities (mutations, gain or loss of whole chromosomes or part of chromosomes). In addition to this, changes in gene expression pattern and epigenetic modifications due to several reasons including fluctuations in culture condition, contamination, mishandling and stressful condition to culture, can also lead to permanent changes in cell behavior (e.g., stem cell culture can differentiate, or lose its ability to differentiate). Therefore, we need a method to preserve cell culture which stop or slow down these processes.
Cryopreservation is an efficient way to preserve cells at ultra-low temperature (below -135°C) which stop all physiological processes and biological aging. It is a routinely used technique in all cell culture laboratories.
During preservation at ultra-low temperature, cells die due to many reason including lysis due to ice crystal formation, pH change, dehydration, and alterations in the concentration of electrolytes. Four distinct phases of cell preservation and revival process can cause to damage to cells…………
when temperature reduced to above freezing point (hypothermia)
when temperature reduced to below freezing point
during frozen state
Cryopreservation methods ensure that cells are alive at ultra-low temperature and maintain their features when revived after long term frozen state.
Most cryopreservation methods rely on
To cryopreserve cells, cells are suspended in freezing medium, followed by slow cooling and subsequently storage in liquid nitrogen.
Freezing medium is nothing but growth medium supplemented with cryoprotectant. Serum containing growth medium contains high amount of serum (upto 90%).
Cryoprotectants, the most important component of freezing medium, function by preventing the formation of ice crystals, thus protect cells from lysis.
Polyalcohols (e.g., glycerol, ethylene glycol, 2,3 butanediol) and DMSO can be used as cryoprotectants, often a concentration varies from 5 – 20%. Most cryoprotectants have ability to penetrate the cell membrane and function by replacing part of the water in the cell.
DMSO is most frequently used cryoprotectant. However, some cells lines are sensitive to DMSO. In such situation, glycerol can be used. Glycerol is less toxic than DMSO, however, osmotic problem associated with glycerol at the time of thawing restrict its uses.
High concentration of serum can also be added in freezing medium. High serum concentration correlate with better survival upon thawing.
Serum-free chemically defined freezing medium are also available which are prepared by adding cryoprotectant to serum-free chemically defined medium growth.
Serum-containing freezing mediums are used for cell lines growing in serum-supplemented growth medium whereas serum-free freezing medium is used for those cell lines which are maintained in serum-free chemically defined medium.
Cells are the basic structural and functional unit of life. Cells from all organisms including multicellular organisms can perform all physiological functions independently. Therefore, cells can be maintained under artificial environment if the right condition is provided.
Cell culture can be defined as a process of maintaining cells under the artificially controlled environment in a culture dish, outside their natural environment. This definition can be applied to any organism including prokaryotes, as well as unicellular and multicellular eukaryotes. However, in practice, the term cell culture is used for cells from multicellular organisms, especially multicellular animals. Specific terminology, like bacterial culture (maintaining bacteria in the controlled laboratory environment), yeast culture (maintaining yeast in the controlled laboratory environment), plant culture are used frequently to denote other types of culture.
Cells in culture behave as an independent unit like unicellular organisms and perform all necessary functions including cell division and metabolism in the culture dish.
Classically, the term ‘Tissue culture’ was used to grow plant and animal explant in a controlled artificial environment in the laboratory. The term ‘Animal tissue culture’ refers to cell culture derived from multicellular animals whereas ‘Plant tissue culture’ refers to the culture of plant cells/tissues.
Culture condition must maintain cell’s characteristics as it possesses in its natural environment. Practically, it is very difficult, in part, due to limited knowledge of physiological requirements of specific cell type. However, many different cell types have been maintained in culture and are in use in both basic and applied research. One such example is a successful maintenance of stem cells (embryonic and adult stem cells) in culture.
Cell culture can be classified on the basis of cell’s characteristics including growth mode, lifespan, morphology and cell types.
Based on growth mode of cells, culture can be broadly classified into two types – suspension culture and adherent culture. Semi-adherent culture, which contains loosely adherent cells to the dish surface, also exist.
Based on cell morphology in the culture dish, cell culture can be broadly classified into three types – Fibroblast-like, Epithelial-like, and Lymphoid-like.
Cell culture is not static. Cells in culture acquire changes which can be genetically programmed (e.g., senescence in primary culture) or due to the accumulation of genetic abnormalities (mutations, gain or loss of whole chromosomes or part of chromosomes). Furthermore, in response to fluctuations in culture condition, cells in culture can show altered behavior due to changes in gene expression pattern which sometimes lead to permanent changes in cell behavior (e.g., stem cells can differentiate).
Cell culture technology has found wide application both in basic research and applied research including industry (pharmaceuticals, medical sciences, cancer research, diagnostics, drug and product development, manufacturing of biological compounds, etc.)