Category Archives: Protocols

Preparation of 1 mM EDTA solution from stock solution of EDTA (0.5M, pH 8.0)

Overview:

  • EDTA (Ethylenediaminetetraacetic acid) is a chelating agent. Due to its ability to sequester metal ions such as Mn2+, Ca2+, Mg2+ and Fe3+, EDTA is widely used in number of cell and molecular biology experiments.
  • Metal ions are necessary for the activity of many enzymes (e.g., DNases and DNA modifying enzymes) as well as interactions of biomolecules (e.g., receptor-ligand interaction). Sequestration of metal ions disrupts metal ion-dependent interactions (e.g., cell-cell and cell substratum interaction) as well as inhibits the action of metal ion dependent enzymes.
  • Here we describe steps to prepare 1 mM EDTA solution from stock EDTA solution.

Requirement

  • Reagents
    • 0.5 M EDTA solution, pH 8.0 at 25°C
    • Deionized / Milli-Q water
  • Equipment and disposables
    • Measuring cylinder

Composition:

1 mM  EDTA, pH 8.0 at 25°C

Objective:

Preparation of 100 ml of 1 mM EDTA solution (pH 8.0) from 0.5 M EDTA solution (pH 8.0)

Preparation:

Step 1: Calculate amount of 0.5M EDTA required for the preparation of 100 ml of 1 mM EDTA.

You need to add 0.2 ml of 0.5 M EDTA in 99.8 ml water to achieve 100 ml of 1 mm EDTA solution. (See calculation)

Step 2: Take ≈50 ml water in a measuring cylinder and add 0.2 ml of 0.5M EDTA in it.

Step 3: Adjust the final volume 100 ml with deionized/milli Q water. Mix it.

Tips: To mix solution, you can cover the measuring cylinder with parafilm and mix the solution by inverting the measuring cylinder 3 – 4 times. Take care no solution leaks out during this process. Alternatively, you can transfer solution to storage bottle and mix solution by swirling the bottle or using magnetic stirrer.

Storage:

Transfer solution to a storage bottle. You can store the solution at room temperature. Solution is stable at room temperature for long time.

 

Follow the table to prepare EDTA solution of specific concentration and volume from 0.5 M EDTA solution
Conc. / Volume 100 ml 250 ml 500 ml 1000 ml
1 mM 0.2 ml 0.5 ml 1.0 ml 2.0 ml
5 mM 1.0 ml 2.5 ml 5.0 ml 10.0 ml
10 mM 2.0 ml 5.0 ml 10.0 ml 20.0 ml
25 mM 5.0 ml 12.5 ml 25.0 ml 50.0 ml

 

Protocol – Subculturing suspension cells

Protocol – Subculturing suspension cells

Overview:

  • Suspension cell cultures are passaged by diluting the existing culture.
  • Since cells float in the medium in suspension culture, they are not treated with trypsin-EDTA solution unless there are some special requirements. For example, if you need to plate estimated number of cells from a culture which forms tight clumps in suspension. In this case, cell clumps are treated with trypsin-EDTA to obtain single cell suspension to count the cell number accurately.
  • To subculture suspension cells, a small amount of cell suspension from the existing culture is transferred to a culture dish containing fresh growth medium.

Requirements:

  • Reagents and solutions:
    • Complete growth medium (room temperature / 37°C)
  • Equipment and disposables
    • T25 flask/Tissue culture dishes
    • Pipettes and pipette aid
    • Laminar flow hood
    • Beaker to discard the waste
Starting material:

Suspension cell culture (high density) ready to subculture

Prior to start:
  • Place complete medium in 37°C water bath for warming.
  • Clean and wipe workspace in the laminar flow hood with 70% ethanol, turn on UV light for 20 – 30 min. After 20 min, turn off the UV light and start the air flow. Let it flow for 10 min.

Objective:

Subculturing of suspension culture growing

Note:
  • Check cells under the microscope to make sure cells are healthy and are not contaminated.
  • Use aseptic techniques while operating cell culture.

Procedure:

Step 1: Transfer all reagent bottles and disposables to the laminar flow hood
  • Spray and wipe all bottles and packets containing disposables (culture dish packets) with 70% ethanol and place them in the laminar flow hood.
  • Take out the estimated number of culture dishes from its packet and label them with the date of subculture, passage number, cell name and your name.
Step 2: Dilute cell suspension at recommended cell density (or split ratio)
  • Take out culture dish from the incubator and place inside laminar flow hood.
  • Pipet suspensension culture to suspend cells homogeneously and transfer an aliquot of cells into fresh culture flask/dish.
  • Add appropriate amount of growth medium (for T25 flask/ 60 mm dish, 4 – 5 ml medium is recommended).
  • Pipette up and down 2 – 3 times to suspend cells homogeneously.
  • Wright all details on culture dish including your name, cell line name, passage number and subculture date.
  • Discard remaining cell suspension if you don’t need them.
Note:
  • Cell counting is not necessary for the regular maintenance of cell in culture. Most researchers use split ratio to seed cells in a fresh culture dish.
  • Split ratio is a rough estimation of how many culture dishes can be prepared from the existing cell culture. For e.g., you can prepare 5 to 10 flask/dish from the cell culture (90% confluent) if the recommended split ratio is 1:5 to 1:10.
Tips:
  • If recommended split ratio is 1:5, transfer ⅕th culture volume to the fresh culture dish. Alternatively, cells can be counted using haemocytometer or any other appropriate method and recommended number of cells can be transferred to the fresh dish. Often counting is performed to seed exact number of cells as needed for the experiment.
  • To count cells accurately, cell clumps should to broken into the single cells. To make single cell suspension, try several rounds of gentle pipetting. If still cell clumps are seen, collect cells by centrifugation, wash the pellet with PBS and briefly treat cells with trypsin-EDTA.
  • If you need to make many culture dishes/flasks, prepare master mix before transferring suspension to the fresh flask.

Step 3: Place the flask in the incubator. Open the lid of the flask slightly for air exchange if you are using sodium bicarbonate containing culture medium. Tighten the lid if you are using vented cap flask.

Protocol – Subculturing adherent cells growing in the serum-containing medium using Trypsin-EDTA

Overview:

  • Trypsin-EDTA method also referred to as trypsinization, is a most commonly used method for passaging/subculturing adherent cells.
  • Cells to be subcultured are first washed with Ca2+-free and Mg2+-free PBS and subsequently treated with Trypsin-EDTA solution.
  • Brief incubation with Trypsin-EDTA disrupts cell-cell and cell-substratum interactions, resulting in the single cell suspension.
  • Trypsin, a proteolytic enzyme, disrupts these interactions by proteolysis and EDTA by chelating divalent cations.
  • Single cell suspension can be used for the determination of cell number and preparation of fresh culture.

Requirements:

  • Reagents and solutions:
    • 1 X Trypsin-EDTA solution (Room Temperature / 37°C)
    • PBS (Ca2+-free and Mg2+-free) (Room Temperature / 37°C)
    • Complete growth medium (Room Temperature / 37°C)
  • Equipment and disposables
    • T25 flask/Tissue culture dishes
    • Pipette and pipette aid (Glass Pipettes)
    • Laminar flow hood
    • Beaker to discard the waste

Starting material: 90% confluent T25 flask/60 mm dish

Prior to start:
  • Place all reagent bottles – Trypsin EDTA, PBS and complete medium in 37°C water bath.
  • Wipe laminar flow hood with 70% ethanol, turn on UV light for 20 – 30 min. After 20 min, turn off the UV light and start the air flow. Let the air flow for 10 min before use.
  • Now wipe the surface of bottles of all reagents and place them in the laminar flow hood.
  • Properly label all dishes/flask with the date of subculture, passage number, cell name and your name.
  • Check cells under the microscope to make sure cells are healthy and are not contaminated.

Objective:

Subculturing of subconfluent (90% confluent) culture growing in a T25 flask/60 mm culture dish

Precautions:
  • Do all operations aseptically.
  • All transfer of medium should be done inside the laminar flow hood.
  • Wear lab coat and disposable latex gloves at all times.
  • Do all steps which involve changing medium from cells quickly to avoid any risk of drying of cells.
  • Don’t decant medium from the flask/dish. This can increase the risk of contamination.

Procedure:

Step 1: Remove and discard culture medium from the flask and wash cells with PBS free of Ca2+ and Mg2+.
  • Take out the flask from the incubator and place inside laminar flow hood.
  • Slightly tilt the flask/dish. All medium will be collected at tilted side. Now remove and discard all medium.
  • Add 3 – 4 ml PBS. Swirl or tilt the dish/flask in opposite direction 2-3 times. Remove and discard PBS.
Note:
  • Serum in the culture medium has trypsin inactivating activity.
  • PBS washing will remove dead cells, cell debris and remaining growth medium.
Tips:
  • To remove liquid from the flask/dish, you can use vacuum aspirator. Vacuum aspirator is very quick and convenient way to remove liquid from the dish. If you don’t have Vacuum aspirator, use pipette and pipette aid to remove liquids from the flask.
  • Always keep the flow of PBS on side walls of the culture vessel. This will minimize the cell detachment due to flow of the liquid.
Precautions:
  • While pipetting PBS into the flask, take care that the flow of PBS should not disturb cell monolayer.
  • Some cell lines are loosely adherent. In such case, care must be taken to avoid any loss while removing culture medium and washing with PBS.
Step 2: Add Trypsin – EDTA solution and incubate 1 – 5 min at 37°C.
  • Add 1 ml Trypsin-EDTA solution and gently spread trypsin solution all over the cell surface by swirling/tilting the vessel . Incubate at 37°C (in incubator) until cells appear rounded and start detaching from the substratum (it can take 1-5 min for most of the cell lines).
  • Gently tap the flask to remove all cells from the surface.
Notes:
  • 0.5 – 1 ml Trypsin-EDTA solution is sufficient for T25 flask/60 mm dish. Depending on the convenience, one can add more trypsin-EDTA solution. The amount of Trypsin-EDTA should be sufficient to cover all the cell surface of the culture vessel.
  • For each cell line, one should optimize trypsin-EDTA treatment condition (incubation time and concentration). Some cell types are strongly adherent and require a higher concentration of the trypsin-EDTA solution as well as longer incubation time.
Tip:
  • Examine the cell morphology under the microscope every minute if you don’t have any idea how long trypsin-EDTA treatment is required for your specific cell line.
Precautions:
  • Make sure trypsin – EDTA solution covers all cell surface. Each cell should come in contact with trypsin solution. Incomplete trypsin treatment may result in cell clumps.
  • Right trypsin-EDTA treatment condition is crucial for successful trypsinization process. While overtreatment will cause cell death, undertreatment results in cell clump and undetached cells in the culture dish.
Step 3: Inactivate trypsin by adding fresh serum containing medium
  • Wash out all the cells from the surface by pipetting the fresh culture medium (4 ml) all over the surface.
  • Disperse all cell clumps by pipetting 2 – 3 times.
Note:
  • Serum has trypsin inactivating activity. If you are using serum-free medium, inactivate trypsin by adding other trypsin inhibitors e.g., soybean trypsin inhibitor.
Tips:
  • While transferring the medium in the vessel, keep the flow of medium toward the surface where cells were attached.
  • If there are cell clumps, disperse them by several rounds of pipetting. More pipetting can cause cell death!
Step 4 (optional): Determine cell number
  • Transfer all cell suspension to a centrifuge tube. Collect cells by centrifugation.
  • Resuspend cells an appropriate amount of complete medium (5 ml) and determine cell number using haemocytometer or any other method.
Notes:
  • Cell counting is not necessary for the regular maintenance of cell culture. Most researchers use split ratio to seed cells in a fresh culture dish.
  • Split ratio is a rough estimation of how many culture dishes can be prepared from the existing cell culture. For e.g., you can prepare 5 to 10 flask/dish from the cell culture (90% confluent) if the split ratio is 1:5 to 1:10.
Step 5: Prepare fresh culture dish from the cell suspension
  • Transfer an aliquot of cells into fresh culture flask/dish.
  • Add fresh growth medium. Pipette cells to resuspend cells again. Wright all details including your name, cell line name, passage number and subculture date.
  • Discard remaining cell suspension if you don’t need them.
  • Keep the flask in the incubator. Open the lid of the flask slightly for air exchange if you are using sodium bicarbonate containing culture medium. Tighten the lid if you are using vented cap flask.
Note:
  • Use recommended seeding density or split ratio to decide how many cells should be transferred to fresh dish/flask. Generally 1:5 – 1:10 split ratio works well for fast-growing cell lines.
Tip:
  • If you need to make many culture dishes/flasks, prepare master mix before transferring suspension to the fresh flask.

Preparation of Tris saturated Phenol

Overview:

  • Phenol is a colorless crystalline solid. Upon exposure to air and light, phenol gradually turns pink to brownish color due to oxidation.
  • Oxidation products (e.g., quinones) of phenol can cause DNA damage (breakdown of phosphodiester bonds, cross-linking of nucleic acids), therefore, should be removed from the phenol by redistillation.
  • Redistillation of phenol at 182°C under nitrogen removes oxidized products from the phenol. Redistilled phenol should be frozen and kept in the dark bottle.
  • In order to utilize phenol for extraction of the nucleic acids (e.g., DNA and RNA), phenol is first equilibrated with buffer (Tris. Cl, pH 8.0) or water.
  • Tris-saturated phenol which has pH ≈8.0 is utilized for the isolation and purification of DNA.
  • To prepare Tris-saturated phenol, phenol is equilibrated with Tris.Cl (pH 8.0) solution until the pH of phenol reaches ≈8.0.
  • Often a small amount of 8-hydroxyquinoline (0.1%) is added in the phenol. 
  • 8-hydroxyquinoline is an antioxidant, which prevents oxidation of phenol. Its yellowish color also helps to identify phenolic phase from aqueous phase during the extraction process.
  • When Tris-saturated phenol or Phenol:chloroform (1:1) solution is used to extract biological samples or DNA solution, both DNA and RNA partitions into the aqueous phase, leaving most of impurities like protein and lipids in phenolic (organic) phase or at the interface.
  • If required, RNA contamination from the extracted DNA can be removed by RNase A digestion.

Requirements

  • Reagents
    • Redistilled Phenol, molecular biology grade: Stored in aliquots at -20℃.
    • 8-hydroxyquinoline
    • 0.5 M Tris.Cl buffer (pH 8.0)
    • 0.1 M Tris.Cl buffer (pH 8.0)
  • Equipment and disposables
    • Fume Hood
    • Magnetic stirrer and Magnetic stir bar
    • Glass Bottle/Beaker
    • Measuring cylinder

Objective

Preparation of Tris.Cl (pH 8.0) saturated Phenol

Precautions:
  • Phenol is volatile and caustic. Care must always be taken when handling phenol (wear lab coat, gloves and eye protection). Do all operations in fume hood.
  • Discard the waste according to your institution’s waste-disposal guidelines.
  • Avoid exposure of phenol to light. Cover the bottle/beaker containing phenol with aluminium foil.
Prior to start:

Set the water bath at 50°C in a fume hood.

Step 1: Thaw the frozen phenol by placing the bottle in 50°C water bath.

Step 2: Transfer 100 ml phenol to a beaker / bottle. (RG)

Step 3: Add ≈0.1 gram 8-hydroxyquinoline [final conc ≈0.1% (w/v)]. Mix to dissolve it.

Step 4. Add 100 ml of 0.5 M Tris.Cl (pH 8.0) to the phenol. Stir the mixture for 15 – 30 min on a magnetic stirrer at room temperature. Place the bottle/beaker in 50°C water bath and allow phase separation. Discard the aqueous phase. Repeat this step (3 – 4 times) until the pH of the aqueous phase is >7.8.

Step 5: Add 100 ml of 0.1 M Tris.Cl (pH 8.0) to the phenol.  Stir the mixture for 15 – 30 min as described in step 4. Allow phase separation and discard aqueous phase.

Step 6: Now transfer the tris saturated phenol to a glass bottle (100 ml) and add ≈20 ml 0.1M Tris.Cl over the phenol. Tris.Cl will form a thin upper layer (0.5 -1 cm).

Storage:

The Tris.Cl saturated phenol can be stored in dark at 4°C for 3 – 6 month. Periodically check the pH of phenol during storage. Discard it if the pH of the phenol is <7.5.

 

Protocol – Plasmid isolation by boiling method (miniprep)

Overview

  • The boiling lysis method of plasmid isolation is quick and is recommended for isolation of small plasmids (up to 10 kb). Plasmids larger than 10 kb should be isolated by other methods (e.g., alkaline lysis method)
  • The quality of plasmid, isolated by this method, is not as good as the plasmid isolated by alkaline lysis method. However, the quality is good enough for restriction digestion analysis. It’s rapidity together with the plasmid quality, suitable for restriction digestion analysis, makes this a method of choice for screening of large number of clones during cloning experiments.
  • This method is not suitable for isolating plasmids from E. coli endA + strains (e.g., HB101, JM100).
  • In this method, the bacterial cells are given brief heat treatment in boiling water bath in presence of lysozyme and triton X-100. Plasmid DNA, due to its small size, comes out from the bacterial cell, whereas, genomic DNA remains trapped inside the cell.
  • Subsequent high speed centrifugation separates the plasmid DNA from rest of the cell debris, which form pellet. Pellet is removed and plasmid DNA is recovered by ethanol or isopropanol precipitation method.

Requirements

  • Reagents and solutions
    • STE solution [8% (w/v) sucrose, 50 mM Tris-HCl (pH 8.0), 50 mM EDTA (pH 8.0)]
    • STET solution [8% (w/v) sucrose, 50 mM Tris-HCl (pH 8.0), 50 mM EDTA (pH 8.0), 5% (w/v) Triton X-100)]
    • Lysozyme stock solution [10 mg/mL Lysozyme in 10 mM Tris-HCl (pH 8.0)]
    • Phenol : Chloroform : Isoamyl alcohol (25 : 24 : 1) solution (optional)
    • 70% Ethanol
    • Isopropanol
    • Tris – EDTA (TE) (100 mM Tris, 10 mM EDTA, pH 8.0)
    • DNase free RNase A (10 mg/ml)
  • Equipment and disposables
    • Boiling water bath
    • Microcentrifuge tubes
    • Micropipette and tips
    • Ice
    • Gloves

Objective

  • Isolation of plasmid DNA from 1-3 ml of bacterial culture (E. coli DH5α) by boiling lysis method.
Starting material:
  • 3 ml overnight grown culture of E. coli DH5α containing plasmid of interest.
Prior to start:
  • Make sure that STE and STET solutions are chilled
  • Set the centrifuge for cooling (4°C)

Protocol:

Step 1: Harvest bacterial cells from 1.5 ml culture
  • Pour 1.5 ml overnight grown culture in a microcentrifuge tube.
  • Centrifuge at room temperature (or 4°C) for 60 seconds at 12,000 rpm (or 5,000 rpm for 5 min).
  • Remove the supernatant from the tube completely, leaving the bacterial pellet as dry as possible.
Notes:
  • The yield of plasmid DNA is dependent mainly on the copy number of the plasmid. For high copy number plasmid, 1.5 ml culture is sufficient to get a good yield of plasmid DNA. However, more culture is required for good yield of low-copy-number plasmid.
Tips:
  • To remove the medium completely, decant the medium from the microcentrifuge tube after centrifugation. Invert microcentrifuge tube upside down on a paper towel to remove residual liquid. Tap the tube gently on the paper towel to remove liquid sticking on the sides of the tube.
  • To take more bacterial culture (more than 1.5 ml) for plasmid isolation, repeat the above process by adding more culture in the same microcentrifuge tube. Microcentrifuge tube with 2 ml capacity can also be used.
  • Chloramphenicol treatment can be used to amplify low-copy number plasmid.
Precautions:
  • While harvesting the bacteria, the speed of centrifugation and time should be optimized in such a way that the pellet after centrifugation should be loose and at the same time supernatant should be clear. If the pellet is tight, it would be difficult to make the suspension of the pellet. Generally, above mention condition works well.
  • Try to remove medium from the pellet completely. Traces of medium may inhibit some of the sensitive restriction enzymes action.
Step 2 (Optional): Wash the bacterial cells with STE solution
  • Add 500 μl ice cold STE solution.
  • Resuspend the bacterial pellet properly by vortexing or by slow rounds of pipetting.
  • Centrifuge at 4°C for 60 seconds at 12,000 rpm (or 5,000 rpm for 5 min).
  • Remove the supernatant from the tube completely.
Notes:
  • The purpose of this step is to remove traces of culture medium from the bacterial cells, which otherwise can cause inhibition of some sensitive restriction enzyme reaction.
Tips:
  • Tris – EDTA solution (100 mM Tris, 10 mM EDTA, pH 8.0) can also be used in place of STE solution to wash the pellet.
Step 3: Resuspend bacterial pellet in STET solution
  • Add 350 μl of STET solution and resuspend the bacterial pellet properly by vortexing or by slow rounds of pipetting.
Precautions:
  • Make sure that the bacterial pellet is completely dispersed in STET solution. No cell clumps should be visible before the boiling in the water bath. Clumps can cause low yield of plasmid.
Step 4: Treat bacterial cells with lysozyme.
  • Add 25 μl of freshly prepared solution of lysozyme and mix immediately by vortexing for 5 seconds.
Precautions:
  • Lysozyme will not work efficiently if the solution pH is less than 8.0.

Step 5: Now place the tube in a boiling water bath for approximately 1 min (40 sec – 60 sec).

Step 6: Clear the lysate by high-speed centrifugation
  • Centrifuge the tube at maximum speed (14,000 rpm) in a microcentrifuge for 10 min at 4°C or room temperature.
  • Transfer the supernatant containing plasmid promptly in new microcentrifuge tube. Alternatively, one can remove the viscous pellet with a sterile toothpick.
Notes:
  • One can centrifuge the tube at room temperature. Centrifugation at 4°C generates tight pellet than centrifugation at room temperature. The tight pellet can be removed easily.
Precautions:
  • While transferring the supernatant or removing the pellet with a sterile toothpick, take care that debris should not come along with the supernatant. Supernatant should be centrifuged again if it contains any suspended particle.
Step 7 (Optional): Extract the supernatant with Phenol : Chloroform : isoamylalcohol solution.
  • This step will remove impurities including protein and lipid contamination from the plasmid preparation.
  • Add equal volume of Phenol:Chloroform:Isoamylalcohol (25:24:1) in the supernatant. Mix by vortexing for 10 sec. Centrifuge at maximum speed at 4°C. Transfer the supernatant to fresh microcentrifuge tube.
Precautions:
  • While transferring the supernatant, take care that no traces of phenol come along with supernatant. Traces of phenol is sufficient to inhibit most enzymatic reactions.
  • Phenol and chloroform are toxic. Follow the safety rules while handling phenol.
Step 8: Recover plasmid from supernatant by isopropanol precipitation.
  • Add equal volume of isopropanol in the supernatant.
  • Mix it by inverting the tube 4 – 6 times. C
  • entrifuge at maximum speed (14,000 rpm) for 30 min at 25°C.
  • Remove the supernatant completely.
Precautions:
  • Incubation of the above mix at room temperature or on ice increases plasmid yield but also causes salt precipitation. Above mentioned condition generally gives good quality of plasmid DNA without much salt contamination.
  • While removing the supernatant, care should be taken as isopropanol precipitated plasmid pellet is loosely attached to the surface and is invisible in most cases. Careless removal of supernatant can result in loss of plasmid pellet.
Step 9: Wash the pellet with 70% ethanol.
  • Add 500 μl of 70% ethanol to the pellet. Close the tube and invert several times.
  • Centrifuge at 14,000 rpm (maximum speed) for 10 min at 25°C.
  • Remove the supernatant completely.
Tips:
  • To remove the supernatant, one can decant the supernatant after first centrifugation. Remains of liquid will be sticking on the wall of microcentrifuge tube. A second flash spin is sufficient to collect all the liquid at the bottom which can be removed by pipetting. Air dry the pellet for 5 min.
Precautions:
  • Take care with this step, as the pellet sometimes does not adhere tightly to the tube and lost while removing the supernatant.
  • Do not overdry the pellet. Overdried pellet is difficult to dissolve.
  • Remove the traces of ethanol as it may inhibit some enzyme reactions.

Step 10: Dissolve the pellet in 25 μl sterile double distilled water or TE (pH 8.0).

Tips:
  • To dissolve the pellet, one can vortex the solution gently for a brief period and also can incubate at 37° for ∼20 minutes.

Storage

  • Solution can be stored at 4°C for few days. Store at -20°C for years.
Precautions:
  • Don’t thaw the plasmid repeatedly. This can cause reduction of the supercoiled form of the plasmid.

Applications

  • The isolated plasmid is suitable for most of our cloning experiments. Often the amount of supercoiled plasmid is comparatively less, therefore, is not suitable for transfection experiments.

Preparation of Neutralization solution (solution III) for the isolation of plasmid by alkaline lysis method

Overview

  • Neutralization solution (solution III) is used for the isolation of plasmid DNA by alkaline lysis method.
  • Neutralization solution is nothing but a potassium acetate solution which has pH 4.8.
  • Addition of neutralization solution in lysed bacterial cells brings the pH back, resulting in precipitation of protein and genomic DNA.
  • Both plasmid and genomic DNA renatures upon addition of neutralization buffer. While plasmid DNA renatures in correct conformation due to its circular and covalent nature, therefore, remains in the solution, genomic DNA precipitates due to random association of both the strands.
  • Sodium dodecyl sulfate (SDS) of the lysis buffer reacts with Potassium acetate and form insoluble Potassium dodecyl sulfate (KDS).

Requirements

  • Reagents
    • 5 M Potassium acetate (CH3CO2K) solution
    • Glacial acetic acid
    • Deionized / Milli-Q water
  • Equipment and disposables
    • Measuring cylinder
    • Conical flask / Beaker
    • Magnetic stirrer (optional)

Composition

  • 3 M Potassium
  • 5 M Acetate

Objective:

Preparation of 100 ml of Neutralization solution (solution III)

Preparation:

Step 1: To prepare, 100 ml of Neutralization solution, take 28.5 ml of Deionized / Milli-Q water in a 100 ml measuring cylinder.

Step 2: Add 60 ml of 5 M Potassium acetate and 11.5 ml of glacial acetic acid. Mix the solution.

Storage
  • Solution can be stored at room temperature in a tightly closed bottle for 1 year.
Applications
  • Plasmid isolation by alkaline lysis method
Follow the table To prepare Neutralization solution of various volume (10 ml, 25 ml, 50 ml and 1,00 ml).
Reagents / Volume 10 ml 25 ml 50 ml 100 ml
5 M Potassium acetate 6.0 ml 15 ml 30 ml 60 ml
Glacial acetate acid 1.15 ml 2.875 ml 5.75 ml 11.5 ml
Water 2.85 ml 7.13 ml 14.25 ml 28.5 ml

Preparation of Lysis solution (solution II) for the isolation of plasmid by alkaline lysis method

Overview

  • Lysis solution (solution II) is used for the isolation of plasmid DNA by alkaline lysis method.
  • The plasmid-containing bacterial cells are lysed by treatment with the lysis solution.
  • Lysis solution contains sodium hydroxide (NaOH) and sodium dodecyl sulfate (SDS).
  • SDS is a detergent which solubilizes the phospholipid and denatures the protein, leading to lysis and release of the cell contents. Denaturing action of SDS also releases protein from DNA, leaving the DNA (both genomic and plasmid DNA) free from proteins.
  • High alkaline condition due to NaOH denatures the plasmid and genomic DNA.

Requirements

  • Reagents and solutions

    • 10 N Sodium hydroxide (NaOH) solution
    • 10% sodium dodecyl sulfate (SDS)
    • Deionized / Milli-Q water
  • Equipment and disposables
    • Measuring cylinder
    • Conical flask / Beaker
    • Magnetic stirrer (optional)
Composition
  • 0.2 N Sodium hydroxide (NaOH)
  • 1% (wt/vol) Sodium Dodecyl Sulfate (SDS)

Objective

  • Preparation of 10 ml of lysis solution (solution II)

Preparation

Step 1: To prepare, 10 ml of lysis solution, take 8 ml of Deionized / Milli-Q water in a 10 ml measuring cylinder.

Step 2: Add 0.2 ml of 10 N NaOH solution and 1.0 ml of 10% sodium dodecyl sulfate (SDS).

Tip:
  • You may see some white precipitate when you add SDS. Dissolve it by mixing.
Precaution:
  • Do not mix concentrated stock solutions together. This will cause precipitation.

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

Storage
  • Solution can be stored at room temperature for a week. It is recommended to prepare fresh lysis solution for optimal lysis.
Applications
  • Preparation of plasmid DNA by alkaline lysis method
Follow the table to prepare lysis solution of various volume (10 ml, 25 ml, 50 ml and 1,00 ml).
Reagents / Volume 10 ml 25 ml 50 ml 100 ml
10 N Sodium hydroxide (NaOH) 0.2 ml 0.5 ml 1 ml 2 ml
10% sodium dodecyl sulfate (SDS) 1.00 ml 2.5 ml 5 ml 10 ml
Water 8.8 ml 22 ml 44 ml 88 ml

Protocol – Growing large volume of E. coli culture for large scale plasmid isolation

Overview:

  • Large scale isolation of plasmid requires large volume of E. coli culture. In DNA cloning experiments, a large amount of plasmid is prepared after confirming the presence of right sequence by restriction digestion and sequencing.
  • Large scale plasmid isolation procedures are termed, midiprep (25 – 50 ml starting culture volume) and maxiprep (100 – 500 ml starting culture volume).
  • A starter culture is initially prepared by inoculating a colony in a small volume (2 – 10 ml) of culture medium.
  • Large culture volume is prepared by diluting starter culture in a ratio of 1: 100 to 1: 1000 in the growth medium.
Note:
  • Here we have taken an example of preparing liquid culture from a colony of E. coli DH5α, transformed with the pcDNA plasmid. The pcDNA plasmid carries ampicillin resistance gene, therefore, requires ampicillin for selection of plasmid-containing bacteria.
  • If your plasmid carries another antibiotic resistant gene, add the respective antibiotic in the culture medium.

Requirements

  • Reagents
    • LB medium
    • Ampicillin (Stock conc 100 mg/ml)
    • A 25-ml conical flask with cotton plug (autoclaved)/Falcon polypropylene tubes (Cat No. #352059)
    • A 500-ml conical flask with cotton plug (autoclaved)
  • Equipment and disposables
    • Bunsen burner
    • Clean workbench
    • Autoclaved toothpicks/Pipette tips/Inoculation loop

Objective:

Growing large volume of culture (100 ml) of E. coli harboring pcDNA plasmid for large scale plasmid isolation

Starting material: Grown bacterial colony on ampicillin-containing LB-Agar plate
Prior to start: Set the shaking incubator at 37°C.
  • Perform all microbiological operations close to the flame of Bunsen burner in a clean place, wiped with 70% ethanol.
  • Do all operations aseptically and use sterile material and reagents. All operation which involves opening of media bottle should be done quickly to reduce the risk of contamination. Before starting your work, clean your hands with soap.

Procedure:

A. Preparation of Starter Culture
Step 1: Prepare LB medium with antibiotic for starter culture
  • Transfer 5 ml LB medium aseptically to a 25-ml conical flask. You can use sterile pipette to transfer liquid medium into the tube.
  • Add 5 µl of ampicillin antibiotic stock solution (100mg/ml). Swirl the flask. The final concentration of kanamycin will be 100 µg/ml.
Precautions:
  • Whenever you open media bottle, show the mouth of the bottle to the flame.
Step 2: Inoculate culture medium with bacterial colony
  • Touch the surface of a bacterial colony with a sterile toothpick or pipette tip
  • Drop it into the antibiotic-containing LB medium.
Precaution:
  • Don’t inoculate culture medium directly from glycerol stock. This can cause low yield and unpredictable result.
  • Make sure that at least some bacterial cells stick to toothpick/pipette tip while picking up the colony from the LB-Agar plate.
Step 3: Grow the culture for 8 – 12 h at 37°C with vigorous shaking.
  • Set the flask in shaker incubator.
  • Set the speed 200 – 300 rpm and start the shaker. Incubate for 8 – 12 h.
Note:
  • Incubation for 8 h is sufficient to see turbidity. At this growth stage, the culture will be in log phase of the growth curve, which represents exponential growing cells. When these cells are diluted, they will maintain their exponential growth.
Tip:
  • A good way to prepare starter culture is to inoculate colony in the morning. Starter culture will be ready in the evening.

B. Preparation of large volume of culture by diluting starter culture in a ration of 1:100 to 1:1000 in growth medium

Step 5: Prepare 100 ml LB medium with antibiotics
  • Transfer 100 ml LB medium aseptically to 500-ml conical flask.
  • Add 100 µl of ampicillin stock solution. Swirl the flask. The final concentration of ampicillin will be 10 µg/ml.

Step 6: Transfer 1 ml starter culture aseptically to 100 ml LB medium.

Step 7: Grow the culture overnight (12 – 16 h) at 37°C with vigorous shaking.
  • Set the flask in the shaker incubator.
  • Set the speed 200 – 300 rpm.
  • Incubate for 12 – 16 h.

Step 8: Take out the culture next moning. Culture is ready for plasmid isolation.

Protocol – Growing liquid culture of E. coli for plasmid miniprep

Overview:

  • Small-scale plasmid isolation procedure, the miniprep, yields sufficient amount of plasmid for the screening of clones and DNA sequencing. Once a clone is confirmed for the presence of insert with right sequence, a large amount of plasmid can be prepared by midiprep or maxiprep.
  • Miniprep requires a small amount of culture of the plasmid-containing bacterial cells. Most often a single colony from the LB-agar plate is inoculated in a liquid medium. Culture is grown at the 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. At this stage, the grown culture has a density of 3 – 4 × 109 cells/ml.
  • Sometimes, a well-grown colony from the LB-agar plate can directly be utilized for plasmid isolation. A well-grown colony on LB-agar plate is prepared by streaking a colony in a small area (0.5 – 1 cm long). This is more convenient when you need to screen a large number of colonies.
  • An antibiotic should be present at all stages of culture growth. The choice of antibiotic depends on the antibiotic resistant gene carried by the plasmid. In absence of antibiotic, dividing cells can lose the plasmid, resulting in low plasmid yield.
Note:
  • Here we have taken an example of preparing liquid culture from the a colony of E. coli DH5α, transformed with the pEGFP plasmid. The pEGFP plasmid contains kanamycin resistance gene, therefore, requires kanamycin for selection of plasmid-containing bacteria.
  • If your plasmid carries another antibiotic resistant gene, add the respective antibiotic in the culture medium.

Requirements

  • Reagents
    • LB medium
    • Kanamycin (Stock conc. 50 mg/ml)
    • Falcon® 14mL Round Bottom polypropylene tube with Snap Cap (Cat No. #352059)/ 25-ml conical flask with cotton plug (autoclaved)
  • Equipment and disposables
    • Bunsen burner
    • Clean workbench
    • Autoclaved toothpick/Pipette tips/Inoculation loop

Objective:

Growing liquid culture of E. coli DH5α harboring pEGFP plasmid for miniprep

Starting material: Bacterial colony on antibiotic containing LB-Agar plate
Prior to start: Set the shaking incubator at 37°C.
  • Perform all microbiological operations close to the flame of bunsen burner in a clean place, wiped with 70% ethanol.
  • Do all operations aseptically and use sterile material and reagents. All operation which involves opening of media bottle should be done quickly to reduce the risk of contamination. Before starting your work, clean your hands with soap.

Procedure:

Step 1: Prepare LB medium with antibiotics
  • Transfer 3 ml LB medium aseptically to polypropylene tube (Falcon, Cat No. #352059).
  • Add 3 µl of antibiotics stock solution of kanamycin (50 mg/ml). The final concentration of kanamycin will be 50 µg/ml.
Note:
  • A single colony can be inoculated in 2 – 10 ml culture volume. Since miniprep needs 1 – 3 ml culture, inoculating 3 ml culture medium is sufficient.
  • Disposable plastic tubes with Snap Cap is a good choice for culture vessel. These tubes are available in ready to use form (sterile), easy to cap and provide good aeration and are cheap. These tubes can be discarded after use, therefore, no effort is required for cleaning and preparing them for the next use. Conical flasks and glass test tubes can also be used for culturing bacteria.
  • Depending on how many colonies you want to inoculate, prepare the same number of flasks. If you are screening for the presence of an insert in a plasmid, you need to inoculate many colonies in separate polypropylene tubes. For example, if you need to inoculate 10 colonies, you must prepare 10 polypropylene tubes with culture medium. In this case, you need 30 ml culture medium. Take 30 ml culture medium, add 30 µl kanamycin and distribute 3 ml in each polypropylene tube.
Tips: 
  • You can either pour or use sterile pipette to transfer liquid medium into the tube. Since Falcon polypropylene tubes (Cat No. #352059) have markings, pouring is more convenient and quick if you have many colonies to inoculate.
Precautions:
  • Whenever you open media bottle, show the mouth of the bottle to the flame.
Step 2: Inoculate culture medium with bacterial colony
  • Touch the surface of a bacterial colony with a sterile toothpick or pipette tip
  • Drop it into the antibiotic containing LB medium.
Precaution:
  • Don’t inoculate culture medium directly from glycerol stock. This can cause low yield and unpredictable result.
  • Make sure that at least some bacterial cells stick to toothpick/pipette tip while picking up the colony from the LB-Agar plate.
Step 3: Grow the culture overnight (12 – 16 h) at 37°C with vigorous shaking.
  • Set the culture tube in shaker incubator. Use appropriate inclined (30° – 45°) angle if you are using polypropylene tube to ensures good shaking.
  • Set the speed 200 – 300 rpm and start the shaker. Incubate for 12 – 16 h.
Precaution:
  • Don’t grow more than 16h.
Step 4: Take out the culture next moning. Culture is ready for plasmid isolation.
 
Note:
  • It is always good to start the isolation process immediately or in the same day. Culture can be stored for 2 – 3 days at 4°C. Longer storage may cause low plasmid yield.

Preparation of resuspension buffer for the isolation of plasmid by alkaline lysis method

Overview:

  • Resuspension buffer is used to resuspend bacterial cells during plasmid isolation by alkaline lysis method. It provides an optimal starting pH (pH 8.0) and an ideal condition for subsequent lysis.
  • Resuspension buffer containing Tris and EDTA is very common.
  • Tris.Cl acts as a buffering agent and maintains the pH of the resuspension buffer 8.0.
  • EDTA  chelates the divalent cations which are released upon bacterial lysis. Divalent cations are required for many enzymatic reactions. EDTA action results in inactivation of many enzymes which may harm plasmid DNA.
  • Resuspension buffer can be supplemented with RNase A which helps to get rid of RNA contamination from the plasmid preparation.

To know more, please read the article: Resuspension buffer (solution I) for isolation of plasmid by alkaline lysis method.

Requirements

  • Reagents
    • 1M Tris.Cl (pH 8.0) solution, autoclaved
    • 0.5 EDTA (pH 8.0) solution, autoclaved
    • Deionized / Milli-Q water
  • Equipment and disposables
    • Measuring cylinder
    • Conical flask / Beaker
    • Magnetic stirrer (optional)

Composition

  • 25 mM Tris.Cl (pH 8.0)
  • 10 mM EDTA (pH 8.0)

Objective

Preparation of 100 ml of resuspension buffer (solution I)

Preparation

Step 1: To prepare 100 ml of resuspension buffer, take 95.5 ml of deionized / Milli-Q water in a 100 ml measuring cylinder/beaker.
Precaution:
  • Do not mix concentrated stock solutions together. This can cause precipitation.
Step 2: Add 2.5 ml of Tris.Cl (pH 8.0) and 2.0 ml of EDTA (pH 8.0). Mix and transfer to a transparent bottle.
Tip:
  • A transparent bottle can easily be examined for any microbial growth in resuspension buffer.

Storage

The solution can be stored at 4°C for 6 months.

Precaution:
  • Frequently check the presence of any microbial growth in resuspension buffer. Discard if you detect any microbial growth.

Application

Preparation of plasmid DNA by alkaline lysis method

Follow the table to prepare resuspension buffer of various volume.
Reagents / Volume 10 ml 25 ml 50 ml 100 ml
1M Tris.Cl (pH 8.0) 0.25 ml 0.625 ml 1.25 ml 2.5 ml
0.5 M EDTA (pH 8.0) 0.2 ml 0.5 ml 1.0 ml 2.0 ml
Water 9.45 ml 23.625 ml 47.25 ml 95.5 ml