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Restriction Digestion of DNA

Cleaving DNA at Specific Recognition Sequences ("Molecular Scissors")

1 Aim

To cleave DNA at specific recognition sites using restriction endonucleases and analyze the resulting fragments using agarose gel electrophoresis.

2 Principle

Restriction Digestion is a fundamental molecular biology technique used to cut DNA molecules at highly specific sequences using enzymes known as Restriction Endonucleases. These enzymes act as "molecular scissors."

Restriction enzymes naturally evolved in bacteria as a defense mechanism to chop up invading viral (bacteriophage) DNA. They recognize short palindromic DNA sequences (sequences that read the same forward and backward on complementary strands) and cleave the DNA backbone.

Types of Cleavage:

• Sticky Ends (Cohesive Ends): The enzyme makes a staggered cut, leaving short, single-stranded DNA overhangs. These are highly desirable for DNA cloning because they easily pair with complementary sticky ends. (e.g., EcoRI, HindIII)
• Blunt Ends: The enzyme cuts straight down the middle of the recognition sequence, leaving no overhangs. (e.g., SmaI, HaeIII)

Fig 1: EcoRI recognizes the palindromic sequence GAATTC and makes a staggered cut, generating 5' sticky ends.

3 Materials Required

Chemicals and Reagents

• DNA sample (Plasmid, Lambda phage, or Genomic DNA)
• Restriction enzyme (e.g., EcoRI, HindIII, BamHI)
• Specific 10X Restriction Enzyme Buffer
• Nuclease-free water
• 6X DNA Loading Dye

Equipment

• Dry bath incubator or Water bath (set to 37°C)
• Microcentrifuge
• Micropipettes and sterile tips
• Agarose gel electrophoresis apparatus
• UV Transilluminator / Gel Doc

4 Preparation of Reaction Mixture

All reagents must be kept on ice during preparation to prevent premature enzymatic activity. Typical reaction volume is 20 µl.

Component	Volume (20 µl Reaction)
DNA sample (~1 µg)	5 µl
10X Restriction Buffer	2 µl
Restriction Enzyme (e.g., EcoRI)	1 µl (10 Units)
Nuclease-free water	12 µl

*Rule of thumb: The volume of the enzyme should never exceed 10% of the total reaction volume to prevent Star Activity from glycerol.

5 Procedure

1. Label sterile microcentrifuge tubes properly (Control, EcoRI digest, etc.).
2. Add nuclease-free water, followed by the 10X buffer, and then the DNA sample.
3. Add the restriction enzyme last. Always use a fresh tip to avoid contaminating the enzyme stock.
4. Mix gently by flicking the tube or pipetting up and down slowly. Do not vortex heavily, as it may denature the enzyme.
5. Centrifuge briefly to collect all contents at the bottom.
6. Incubate the reaction mixture at 37°C for 1 hour in a water bath or dry bath incubator. (Note: Some enzymes require different temperatures, always check the manufacturer protocol).
7. Terminate the reaction by heat inactivation (e.g., 65°C for 20 mins) or by adding EDTA/Loading Dye.

6. Analysis & Result

To confirm that the DNA was successfully cleaved, we run the products on a gel.

1. Prepare a 1% agarose gel containing Ethidium Bromide.
2. Add 6X Loading Dye to your digested samples.
3. Load an Undigested Control DNA in Lane 1, the DNA Ladder in Lane 2, and the Digested Samples in subsequent lanes.
4. Run electrophoresis at 80–100 V until the dye front is 75% down the gel.
5. Visualize under a UV Transilluminator.

Result: The Undigested control will show a single high-molecular-weight band (or multiple bands if it is a supercoiled plasmid). The Digested sample will show distinct, separate bands corresponding to the cleaved fragments. The size of these fragments can be estimated using the DNA ladder.

7. Troubleshooting: Common Digestion Errors

Observation	Likely Cause & Solution
Partial Digestion (Extra unexpected larger bands)	The enzyme didn't cut all the sites. Caused by too little enzyme, too much DNA, insufficient incubation time, or inactive enzyme. Increase time or enzyme units.
Star Activity (Unexpected smaller fragments)	The enzyme lost its specificity and cut at incorrect sites. Caused by too much glycerol (>5%), wrong buffer, or incubating for too long.
Smeared DNA on gel	DNA degradation caused by nuclease contamination. Ensure tips and water are strictly nuclease-free.

8. Advantages & Applications

Key Advantages

• Highly specific and predictable DNA cleavage.
• Essential first step for Recombinant DNA Technology.
• Simple, reproducible protocol.

Applications

• DNA Cloning: Splicing genes into plasmids.
• Restriction Fragment Length Polymorphism (RFLP): Used in genetic fingerprinting and paternity testing.
• Gene Mapping: Creating restriction maps of unknown genomes.

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Important Viva Questions

Q1. What is a "Palindromic Sequence"?

A palindromic sequence in DNA reads the same in the 5' to 3' direction on one strand as it does in the 5' to 3' direction on the complementary strand. Examples include GAATTC (EcoRI) and AAGCTT (HindIII).

Q2. What is "Star Activity"?

Star activity is the relaxation or alteration of the specificity of a restriction enzyme. Instead of cutting at its exact recognition sequence, it cuts at similar, incorrect sequences, resulting in a smear or unexpected extra bands. It occurs under sub-optimal conditions like high glycerol concentration (>5%), incorrect pH, or over-incubation.

Q3. Why are restriction enzymes shipped and stored in 50% Glycerol, and why must they be kept on ice?

Glycerol acts as a cryoprotectant, preventing the enzyme from freezing solid and denaturing at -20°C storage. They must be kept on ice on the benchtop because they are highly temperature-sensitive proteins; leaving them at room temperature degrades their enzymatic activity quickly.

Q4. What are Isoschizomers?

Isoschizomers are pairs of restriction enzymes isolated from different bacterial strains that recognize and cut at the exact same recognition sequence.

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🧠 Interactive Concept Quiz

Test your knowledge! Click on the questions below to reveal the correct answers.

1. What does the "Eco" and "R" stand for in the enzyme EcoRI?

✅ Answer: Escherichia coli, strain RY13

Restriction enzymes are named after the bacteria they are isolated from. "Eco" represents the genus and species (Escherichia coli), "R" represents the strain, and "I" indicates it was the first enzyme isolated from that strain.

2. Why doesn't a bacteria's own restriction enzyme cut its own DNA?

✅ Answer: DNA Methylation

Bacteria protect their own DNA by adding methyl groups (-CH3) to specific bases within the recognition sequence. The restriction enzyme cannot bind to or cut methylated DNA.

3. Why must you keep the volume of the restriction enzyme under 10% of the total reaction volume?

✅ Answer: To prevent Star Activity caused by Glycerol.

Enzymes are stored in 50% glycerol. If the enzyme makes up more than 10% of your reaction volume, the glycerol concentration in the tube will exceed 5%, which alters the enzyme's structure and causes it to cut the DNA randomly.

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Polymerase Chain Reaction (PCR)

In Vitro Amplification of Specific Target DNA Sequences

1 Aim

To amplify a specific DNA fragment in vitro using the Polymerase Chain Reaction (PCR) technique with Taq DNA Polymerase in a Thermal Cycler.

2 Principle

The Polymerase Chain Reaction is a revolutionary molecular biology technique used to exponentially amplify a specific region of DNA. It relies on thermal cycling, consisting of repeated cycles of heating and cooling of the reaction for DNA melting and enzymatic replication.

The 3 Major Steps of PCR:

1. Denaturation (94–95°C): The intense heat breaks the hydrogen bonds between the double-stranded DNA, yielding two single strands.
2. Annealing (50–65°C): The temperature is lowered to allow short, target-specific DNA primers to bind (anneal) to their complementary sequences on the single-stranded template.
3. Extension (72°C): The temperature is raised to the optimal working temperature of Taq DNA Polymerase, which synthesizes a new DNA strand by adding dNTPs (nucleotides).

Fig 1: The three fundamental stages of a single PCR cycle.

Each cycle doubles the DNA amount, resulting in exponential amplification (2ⁿ copies, where n is the number of cycles). After 25–35 cycles, millions of copies of the specific DNA fragment are produced.

3 Materials Required

Chemicals and Reagents

• Template DNA
• Forward & Reverse Primers
• dNTP mixture (dATP, dTTP, dGTP, dCTP)
• 10X PCR buffer
• MgCl₂ (Cofactor for Taq)
• Taq DNA Polymerase
• Nuclease-free water

Equipment

• Thermal Cycler (PCR Machine)
• Micropipettes and sterile tips
• 0.2 ml PCR tubes
• Microcentrifuge
• Vortex mixer
• Gel electrophoresis apparatus

4 Preparation of PCR Reaction Mixture

All reagents should be kept on ice while preparing the master mix. Typical PCR reaction volume is 25 µl.

Component	Volume (per 25 µl rxn)
Template DNA	1–2 µl
Forward Primer (10 µM)	1 µl
Reverse Primer (10 µM)	1 µl
dNTP Mix (10 mM)	1 µl
10X PCR Buffer	2.5 µl
MgCl₂ (25 mM)	1.5 µl
Taq DNA Polymerase	0.5 µl
Nuclease Free Water	Up to 25 µl

5 PCR Amplification Program

Place the sealed PCR tubes into the Thermal Cycler and program the following temperature cycles:

Step	Temperature	Time	Cycles
Initial Denaturation	95°C	3–5 min	30 - 35 Cycles
Denaturation	94°C	30 sec
Annealing	50–65°C*	30 sec
Extension	72°C	1 min/kb
Final Extension	72°C	5–10 min	1
Hold	4°C	∞	-

*Annealing temperature depends on the melting temperature (Tm) of the specific primers used.

6. Procedure

1. Label sterile 0.2 ml PCR tubes properly.
2. Prepare the PCR master mix on ice according to the reaction table (multiply volumes by number of reactions + 1 extra).
3. Aliquot the master mix into individual tubes.
4. Add the template DNA carefully to the respective tubes.
5. Mix the reaction gently using a vortex mixer and briefly centrifuge to bring contents to the bottom.
6. Place the tubes inside the Thermal Cycler block.
7. Program the PCR cycling conditions and start the run.
8. After completion, proceed to analysis or store PCR products at 4°C or −20°C.

7. Troubleshooting PCR

Problem	Likely Cause & Solution
No Bands (No Amplification)	Missing reagent (e.g., Taq or Taq buffer), annealing temp too high, or degraded template DNA.
Non-specific Bands (Multiple Bands)	Annealing temperature is too low (primers binding randomly), or MgCl₂ concentration is too high.
Primer Dimers (Fuzzy band at bottom)	Primers are binding to each other. Use less primer or redesign primers to avoid self-complementarity.

8. Advantages & Applications

Key Advantages

• Highly sensitive: Can amplify from a single DNA molecule.
• Rapid: Yields millions of copies in just 2 hours.
• Specific: Primers ensure only the target region is copied.

Applications

• Gene cloning and sequencing
• Detection of infectious diseases (e.g., COVID-19, HIV)
• Forensic DNA fingerprinting
• Detection of GMOs

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Important Viva Questions

Q1. Why is Taq Polymerase used in PCR instead of normal human DNA polymerase?

Normal polymerases denature and are destroyed at 95°C. Taq polymerase is isolated from the thermophilic bacterium Thermus aquaticus (found in hot springs). It is thermostable and survives the extreme heat of the denaturation step without needing to be replenished every cycle.

Q2. What is the role of MgCl₂ in the PCR reaction?

Magnesium ions (Mg²⁺) act as a crucial cofactor for the Taq DNA polymerase. Without it, the enzyme cannot function. Furthermore, Mg²⁺ helps stabilize the interaction between the primers and the DNA template.

Q3. How do you determine the correct Annealing Temperature?

The annealing temperature is usually set 3°C to 5°C below the lowest Melting Temperature (Tm) of the primers being used. If it is too high, primers won't bind. If it's too low, they will bind randomly, creating non-specific products.

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🧠 Interactive Concept Quiz

Test your knowledge! Click on the questions below to reveal the correct answers.

1. At what temperature does the Denaturation step occur?

✅ Answer: 94°C – 95°C

At this extreme temperature, the hydrogen bonds connecting the double-stranded DNA break apart.

2. What does "dNTP" stand for in the PCR mixture?

✅ Answer: Deoxynucleotide triphosphate

These are the free nucleotides (A, T, C, G) that the polymerase uses as building blocks to create the new DNA strand.

3. Why must we cool the reaction down to 50–65°C during the Annealing step?

✅ Answer: To allow the primers to attach.

Cooling the reaction provides the ideal conditions for the short DNA primers to hydrogen-bond to their complementary target sequences on the template DNA.

4. If you run 30 cycles of PCR starting from a single DNA molecule, approximately how many copies will you have?

✅ Answer: Over 1 Billion (2³⁰)

PCR results in exponential growth. Each cycle doubles the amount of target DNA.

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AGAROSE GEL ELECTROPHORESIS

Separation, Sizing, and Visualization of DNA Fragments

1. Aim

To separate, analyze, and visualize DNA fragments using agarose gel electrophoresis and staining with Ethidium Bromide under UV light.

2. Principle

Agarose gel electrophoresis is a widely used technique in molecular biology to separate DNA fragments based on their size.

DNA molecules carry a negative charge because of the phosphate groups in their backbone. When an electric field is applied across an agarose gel, DNA molecules migrate toward the positive electrode (anode).

Fig 1: Schematic representation of DNA migrating from the negative electrode to the positive electrode.

The agarose gel acts as a molecular sieve, allowing smaller DNA fragments to move faster than larger fragments.

After electrophoresis, DNA fragments are visualized using Ethidium Bromide, which intercalates between DNA bases and fluoresces under a UV Transilluminator.

3. Materials Required

Chemicals and Reagents

• Agarose powder
• 1X TAE or TBE buffer
• DNA sample
• DNA loading dye (contains glycerol to add density and a tracking dye)
• DNA ladder (molecular marker)
• Ethidium Bromide
• Distilled water

Equipment

• Electrophoresis tank & Power supply
• Gel casting tray & Comb
• Micropipette and tips
• Gel documentation system & UV Transilluminator

4. Preparation of Agarose Gel

Expert Note: Ensure the ends of the gel casting tray are properly sealed before pouring the gel to prevent leaks.

1. Weigh 1 g agarose powder.
2. Add agarose to 100 ml 1X TAE buffer to prepare a 1% agarose gel.
3. Heat the mixture in a microwave oven until the agarose dissolves completely.
4. Allow the solution to cool to about 50–60°C.
5. Add Ethidium Bromide (0.5 µg/ml) carefully and mix gently.
6. Seal the ends of the casting tray and pour the gel.
7. Insert the comb to create wells.
8. Allow the gel to solidify for 20–30 minutes.

5. Procedure

1. Place the solidified gel into the electrophoresis tank.
2. Add 1X TAE buffer until the gel is completely submerged.
3. Carefully remove the comb from the gel.
4. Load the samples into wells using a micropipette.
5. Connect the electrophoresis unit to the power supply.
6. Run electrophoresis at 80–120 V for 30–45 minutes.
7. When the dye front reaches about 75% of the gel, stop the electrophoresis.

6. Visualization & Results

To measure the exact size of your DNA fragments, you compare your sample bands against the DNA Ladder lane. The ladder acts like a ruler.

Fig 2: Simulated view of glowing DNA bands. Notice how smaller fragments migrate further down the gel.

7. Troubleshooting Common Errors

Observation	Possible Cause
Smeared DNA Bands	DNA degradation by nucleases, too much DNA loaded, or voltage too high.
No Bands Visible	Forgot to add Ethidium Bromide, DNA concentration too low, or DNA ran off the end of the gel.
Gel Melting During Run	Voltage applied is too high or incorrect running buffer was used.

8. Advantages & Limitations

Advantages

• Simple to perform and cost-effective.
• Non-destructive (DNA can be extracted later).
• Easily handles a wide range of DNA sizes.

Limitations

• Poor resolution for very tiny DNA fragments (use PAGE instead).
• Ethidium Bromide is highly toxic/mutagenic.
• Low quantitative accuracy.

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9. Important Viva Questions & Answers

Q1. Why does DNA migrate towards the anode during electrophoresis?

DNA has a sugar-phosphate backbone. The phosphate groups give DNA a net negative charge. Because opposite charges attract, the negatively charged DNA moves towards the positive electrode (anode).

Q2. What is the role of Ethidium Bromide (EtBr)?

EtBr acts as an intercalating agent. It slides between the nitrogenous base pairs of the DNA double helix. When exposed to UV light, it fluoresces strongly, making the invisible DNA bands visible to the naked eye.

Q3. Why is glycerol added to the DNA loading dye?

Glycerol increases the density of the DNA sample. This ensures that when the sample is pipetted into the well, it sinks heavily to the bottom rather than floating away and dissolving into the running buffer.

Q4. What is the purpose of the TAE or TBE running buffer?

The buffer serves two main purposes: it provides the necessary ions to conduct the electrical current through the gel, and it maintains a stable pH so the DNA does not denature during the run.

Q5. How does the concentration of agarose affect DNA separation?

Agarose creates a porous molecular matrix. A higher concentration of agarose (e.g., 2%) creates smaller pores, which is ideal for separating very small DNA fragments. A lower concentration (e.g., 0.8%) creates larger pores, ideal for large DNA fragments.

Q6. What is a "DNA Ladder" and why is it loaded in the first well?

A DNA ladder is a mixture of DNA fragments of known sizes (measured in base pairs, or bp). It acts as a ruler. By comparing the distance your sample traveled against the ladder, you can accurately estimate the size of your DNA fragments.

Q7. Why do we add tracking dyes like Bromophenol Blue?

Because DNA is invisible to the naked eye, tracking dyes are added to monitor the progress of the electrophoresis. The dye migrates ahead of the DNA, letting the researcher know exactly when to turn off the power supply before the DNA runs off the edge of the gel.

Plasmid DNA Isolation from Escherichia coli by Alkaline Lysis Method

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Plasmid DNA Isolation

Alkaline Lysis Method from Escherichia coli

Aim

To isolate and purify recombinant plasmid DNA from Escherichia coli culture using the alkaline lysis method.

Principle

Plasmid DNA is a small, circular, double-stranded DNA molecule that replicates independently of chromosomal DNA in bacteria. The alkaline lysis method is widely used to isolate plasmid DNA because it relies on the selective denaturation and renaturation of plasmid DNA versus chromosomal DNA.

• Cell Lysis: Bacterial cells are lysed using an alkaline solution containing NaOH and SDS. SDS disrupts the cell membrane and denatures proteins, while the high pH (from NaOH) denatures both chromosomal and plasmid DNA.
• Neutralization: The addition of acidic potassium acetate drops the pH. The large, bulky chromosomal DNA tangles and precipitates out of solution along with denatured proteins. However, the small, circular plasmid DNA rapidly renatures and remains dissolved in the solution.
• Centrifugation: The precipitated cellular debris, proteins, and genomic DNA are spun down into a pellet, leaving pure plasmid DNA in the supernatant.
• Precipitation & Purification: The plasmid DNA is precipitated using cold alcohol and washed to remove excess salts.

1. Harvest E. coli Cells

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2. Resuspend (Solution I)

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3. Lyse (Solution II)

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4. Neutralize (Solution III)

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5. Centrifuge & Keep Supernatant

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6. Isopropanol Precipitation

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7. 70% Wash & Dry

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8. Dissolve in TE

Materials & Reagents Required

The 3 Core Solutions (Added Detail)

• Solution I (Resuspension Buffer): 50 mM Glucose (maintains osmolarity), 25 mM Tris-HCl pH 8.0 (buffer), 10 mM EDTA (chelates Mg²⁺ to destabilize the cell wall and inhibit DNases).
• Solution II (Lysis Buffer): 0.2 N NaOH (denatures DNA), 1% SDS (dissolves lipid membranes).
• Solution III (Neutralization Buffer): 3 M Potassium acetate pH 5.5 (neutralizes pH to allow plasmid renaturation; potassium precipitates the SDS and proteins).

Other Requirements

• E. coli culture containing recombinant plasmid
• RNase A (often pre-added to Solution I)
• Cold Isopropanol & 70% Ethanol
• TE buffer or nuclease-free water
• Microcentrifuge, 37°C Incubator, Micropipettes

Procedure

1. Culture Preparation: Inoculate a single colony of E. coli containing the plasmid into 5 mL LB broth with the appropriate antibiotic. Incubate overnight at 37°C with shaking.
2. Cell Harvesting: Transfer 1.5 mL bacterial culture into a microcentrifuge tube. Centrifuge at 10,000 rpm for 5 minutes. Discard the supernatant and retain the pellet.
3. Resuspension: Add 100 µL Solution I (Resuspension Buffer containing RNase A). Resuspend the pellet completely by gentle pipetting until no clumps remain.
4. Alkaline Lysis: Add 200 µL Solution II (Lysis Buffer). Mix gently by inversion 4-6 times (DO NOT vortex, as this shears genomic DNA). Incubate for 5 minutes at room temperature until the solution turns clear and viscous.
5. Neutralization: Add 150 µL Solution III (Neutralization Buffer). Mix gently by inversion. A white, cloudy precipitate (genomic DNA, proteins, and KDS) will immediately form. Incubate on ice for 5–10 minutes.
6. Centrifugation: Centrifuge at 12,000 rpm for 10 minutes. Carefully transfer the clear supernatant (containing the plasmid DNA) into a fresh tube, leaving the white pellet behind.
7. DNA Precipitation: Add an equal volume of cold isopropanol to the supernatant. Mix gently and incubate at −20°C for 20 minutes.
8. Pellet Collection: Centrifuge at 12,000 rpm for 10 minutes. A small white plasmid DNA pellet will appear at the bottom of the tube.
9. Washing: Carefully discard the supernatant. Wash the pellet with 500 µL of 70% ethanol. Centrifuge for 5 minutes, discard the ethanol, and let the pellet air dry completely.
10. DNA Dissolution: Dissolve the purified plasmid DNA in 30–50 µL of TE buffer or nuclease-free water. Store at −20°C.

Quality Analysis & Results

To check the quality of the isolated plasmid, perform Agarose Gel Electrophoresis (1% agarose gel run at 80–100 V).

Result: Recombinant plasmid DNA is successfully isolated. On the gel, high-quality intact plasmid will primarily show a bright, fast-migrating band representing Supercoiled plasmid DNA, along with fainter bands for nicked (relaxed circle) or linear forms.

⚠️ Precautions

• Never vortex after adding Solution II or III; shearing the chromosomal DNA will contaminate your plasmid yield.
• Do not leave cells in Solution II for longer than 5 minutes, or the plasmid DNA may become permanently denatured.
• Ensure the final ethanol wash is completely dried off; residual ethanol inhibits downstream enzymatic reactions.

🔬 Applications

• Gene cloning and sub-cloning
• Restriction enzyme digestion mapping
• Automated DNA sequencing
• PCR amplification of target genes
• Bacterial transformation and mammalian cell transfection

References: Sambrook, J., & Russell, D. W. (2001). Molecular Cloning. | Brown, T. A. (2016). Gene Cloning and DNA Analysis.

🧠 Top 5 Viva Voce Questions

Q1. What is the specific role of Glucose in Solution I?
A: Glucose maintains the osmolarity of the solution, preventing the bacterial cells from bursting prematurely before the addition of the lysis buffer.

Q2. Why do we see a thick white precipitate after adding Solution III?
A: The white precipitate is a complex of Potassium Dodecyl Sulfate (KDS), denatured cellular proteins, and tangled chromosomal DNA. SDS is soluble in sodium salt but insoluble as a potassium salt.

Q3. Why is it strictly advised not to vortex after adding Solution II?
A: Vigorous mixing or vortexing generates physical shear forces that will break the large, fragile genomic DNA into smaller fragments. These small genomic fragments will renature and contaminate your plasmid DNA preparation.

Q4. How does plasmid DNA separate from chromosomal DNA in this protocol?
A: Both are denatured by NaOH. However, because plasmid DNA is small and supercoiled, its two strands stay physically close together. When the pH is neutralized by Solution III, the plasmid strands snap back together (renature) and stay in solution, while the massive chromosomal DNA strands tangle and precipitate out.

Q5. Why might you see three bands for a single plasmid on an agarose gel?
A: Plasmids can exist in three conformations: Supercoiled (intact and most compact, travels fastest), Linear (cut on both strands, travels at its true size), and Nicked/Relaxed Circular (cut on one strand, bulky, travels the slowest).

GENOMIC DNA ISOLATION

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The Ultimate Guide to Genomic DNA Isolation

Complete Laboratory Protocols for Bacteria, Plant Tissues, and Animal Cell Lines

Welcome to the comprehensive laboratory guide for genomic DNA extraction. Whether you are working with bacterial cultures, tough plant cell walls, or fragile animal cell lines, this guide covers the principles, precise reagent compositions, and step-by-step procedures you need for pure, high-yield DNA.

Experiment 1: Isolation of Genomic DNA from Bacterial Cells

Aim

To isolate and purify high-molecular-weight genomic DNA from bacterial cells.

Principle

Bacterial genomic DNA isolation involves cell lysis, removal of proteins and contaminants, and DNA precipitation. Bacterial cells are lysed using detergents such as SDS, which disrupt the cell membrane and release cellular components. Proteinase K digests proteins, while RNase A removes RNA contamination. Proteins are removed by phenol–chloroform extraction, and DNA is precipitated using cold alcohol.

1. Overnight Culture

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2. Harvest Pellet

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3. Lysis (SDS + Prot. K)

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4. RNase A Treatment

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5. Phenol-Chloroform

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6. Ethanol Precipitation

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7. 70% Wash & Dry

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8. Dissolve in TE

Materials Required

• Sample: Overnight bacterial culture (e.g., Escherichia coli)
• Reagents: TE buffer (10 mM Tris-HCl, 1 mM EDTA, pH 8.0), SDS (10%), Proteinase K, RNase A, Phenol:Chloroform:Isoamyl alcohol (25:24:1), Sodium acetate (3 M), Absolute ethanol (cold), 70% ethanol
• Equipment: Microcentrifuge, Water bath (37°C), Micropipettes, Microcentrifuge tubes

Procedure

1. Culture Preparation: Inoculate bacterial cells into LB broth medium and incubate overnight at 37°C with shaking.
2. Cell Harvesting: Transfer 1.5 mL bacterial culture to a microcentrifuge tube. Centrifuge at 10,000 rpm for 5 minutes. Discard the supernatant and retain the pellet.
3. Cell Lysis: Add 500 µL TE buffer to the pellet and resuspend. Add 50 µL SDS (10%) and 10 µL Proteinase K. Incubate at 37°C for 30 minutes.
4. RNA Removal: Add 5 µL RNase A. Incubate at 37°C for 15 minutes.
5. Phenol–Chloroform Extraction: Add an equal volume of phenol:chloroform:isoamyl alcohol. Mix gently by inversion. Centrifuge at 12,000 rpm for 10 minutes. Transfer the upper aqueous layer containing DNA to a new tube.
6. DNA Precipitation: Add 0.1 volume sodium acetate and 2 volumes cold ethanol. Mix gently and incubate at −20°C for 30 minutes.
7. Pellet Collection: Centrifuge at 12,000 rpm for 10 minutes. Discard supernatant carefully.
8. Washing: Wash DNA pellet with 500 µL of 70% ethanol. Centrifuge for 5 minutes and discard ethanol. Air dry the pellet.
9. DNA Dissolution: Dissolve DNA pellet in 50 µL TE buffer. Store at −20°C.

Result: High-molecular-weight genomic DNA is obtained from bacterial cells.
References: Sambrook, J., & Russell, D. W. (2001). Molecular Cloning. | Wilson, K., & Walker, J. (2010). Principles and Techniques.

Experiment 2: Isolation of DNA from Plant Tissue (CTAB Method)

Aim

To isolate genomic DNA from plant leaf tissue using the CTAB extraction method.

Principle

Plant cells possess cell walls and high levels of polysaccharides and phenolic compounds, which can interfere with DNA extraction. CTAB (Cetyltrimethylammonium bromide) is a cationic detergent that lyses cells and removes polysaccharides. After cell lysis, proteins and other contaminants are removed using chloroform–isoamyl alcohol extraction, and DNA is precipitated with isopropanol or ethanol.

1. Grind in Liquid N₂

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2. Add CTAB Buffer

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3. Incubate at 65°C

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4. Chloroform:IAA

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5. Aqueous Phase

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6. Cold Isopropanol

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7. Wash & Dry

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8. Dissolve in TE

Materials Required

• Sample: Fresh plant leaves (100 mg)
• Reagents: CTAB extraction buffer, Chloroform:Isoamyl alcohol (24:1), Isopropanol (cold), 70% ethanol, TE buffer
• Equipment: Mortar and pestle, Liquid nitrogen, Water bath (65°C), Microcentrifuge

CTAB Buffer Composition
CTAB	2%
Tris-HCl	100 mM
EDTA	20 mM
NaCl	1.4 M
β-mercaptoethanol	0.2%

Procedure

1. Tissue Grinding: Take 100 mg fresh leaf tissue. Grind the tissue in liquid nitrogen using mortar and pestle until a fine powder forms.
2. Cell Lysis: Transfer powdered tissue to a tube. Add 700 µL CTAB extraction buffer and mix gently.
3. Incubation: Incubate at 65°C for 30 minutes. Mix occasionally.
4. Organic Extraction: Add equal volume of chloroform:isoamyl alcohol (24:1). Mix gently by inversion. Centrifuge at 12,000 rpm for 10 minutes.
5. Transfer Aqueous Phase: Carefully transfer the upper aqueous phase into a new tube.
6. DNA Precipitation: Add 0.6 volume cold isopropanol. Mix gently. Incubate at −20°C for 30 minutes.
7. Pellet Collection: Centrifuge at 12,000 rpm for 10 minutes. A DNA pellet will appear.
8. Washing: Wash pellet with 70% ethanol. Air dry pellet.
9. DNA Dissolution: Dissolve DNA pellet in TE buffer.

Result: High-quality plant genomic DNA suitable for PCR and sequencing is obtained.
References: Doyle, J. J., & Doyle, J. L. (1990). Focus. | Wilson, K., & Walker, J. (2010).

Experiment 3: Isolation of DNA from Animal Cell Lines

Aim

To isolate genomic DNA from cultured animal cells (e.g., HeLa or HEK293).

Principle

Animal cells lack a cell wall, making DNA extraction simpler. Cells are lysed using detergents such as SDS, proteins are digested with Proteinase K, and RNA is removed using RNase A. DNA is purified using phenol–chloroform extraction and precipitated with ethanol.

1. Collect Cells

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2. PBS Wash

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3. Add Lysis Buffer (SDS)

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4. Proteinase K (55°C)

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5. RNase A (37°C)

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6. Phenol:Chloroform

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7. Ethanol Precipitation

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8. Dissolve in TE

Lysis Buffer Composition

• ✔ Tris-HCl: 10 mM
• ✔ EDTA: 10 mM
• ✔ NaCl: 100 mM
• ✔ SDS: 0.5%

Procedure

1. Cell Collection: Collect 1–5 × 10⁶ cultured cells in a centrifuge tube. Centrifuge at 3000 rpm for 5 minutes.
2. Washing: Wash the pellet with PBS. Centrifuge again and discard supernatant.
3. Cell Lysis: Add 500 µL lysis buffer. Add 20 µL SDS. Add 10 µL Proteinase K.
4. Incubation: Incubate at 55°C for 1–2 hours until solution clears.
5. RNA Removal: Add RNase A. Incubate at 37°C for 20 minutes.
6. Phenol–Chloroform Extraction: Add equal volume phenol:chloroform:isoamyl alcohol. Centrifuge at 12,000 rpm for 10 minutes. Transfer aqueous layer to a new tube.
7. DNA Precipitation: Add 0.1 volume sodium acetate. Add 2 volumes cold ethanol. Incubate at −20°C for 30 minutes.
8. Pellet Collection: Centrifuge at 12,000 rpm for 10 minutes. Wash pellet with 70% ethanol.
9. DNA Resuspension: Dissolve pellet in TE buffer.

Result: Purified high-molecular-weight genomic DNA is obtained from animal cell lines.
References: Freshney, R. I. (2016). Culture of Animal Cells. | Green, M. R., & Sambrook, J. (2012).

🧠 Top 10 Viva Voce Questions & Answers

Perfect for practical exams, these questions cover the critical "why" behind the steps in all three extraction methods.

Q1. Why is Liquid Nitrogen used specifically in plant DNA extraction?
A: Plant cells have rigid cellulose cell walls. Liquid nitrogen freezes the tissue, making it brittle and easy to grind into a fine powder without degrading the DNA. It also instantly inactivates endogenous nucleases.

Q2. What is the role of CTAB in the plant extraction protocol?
A: CTAB is a cationic detergent. It binds to polysaccharides and phenolic compounds (which are abundant in plants) and helps separate them from DNA during the organic extraction phase.

Q3. Why do we use SDS (Sodium Dodecyl Sulfate) in bacterial and animal cell lysis?
A: SDS is an anionic detergent that dissolves the lipid bilayer of cell membranes and denatures cellular proteins, causing the cell to burst open (lyse) and release its genomic contents.

Q4. What is the function of Proteinase K in these protocols?
A: Proteinase K is a broad-spectrum serine protease. It rapidly degrades cellular proteins, including histones bound to DNA and highly destructive DNase enzymes, protecting the DNA from degradation.

Q5. Why is Phenol-Chloroform used, and why in a specific ratio?
A: Phenol denatures proteins, while chloroform ensures phase separation and removes phenol residues. Isoamyl alcohol reduces foaming. The mixture forces denatured proteins into the organic phase, leaving pure nucleic acids in the upper aqueous phase.

Q6. Why is ice-cold absolute ethanol or isopropanol added at the end?
A: DNA is insoluble in alcohol. Adding cold alcohol alters the dielectric constant of the solution, causing the DNA molecules to aggregate and precipitate out of the aqueous solution so they can be pelleted via centrifugation.

Q7. What is the purpose of washing the final DNA pellet with 70% ethanol?
A: 70% ethanol washes away residual salts (like sodium acetate or NaCl) that precipitated with the DNA. The remaining 30% water dissolves the salts, while the 70% ethanol keeps the DNA precipitated.

Q8. Why do animal cells not require the intense physical grinding used for plant cells?
A: Animal cells lack a rigid cell wall; they only have a flexible plasma membrane. Mild chemical detergents (like SDS) are perfectly sufficient to lyse them.

Q9. What is the role of EDTA in the TE buffer and extraction buffers?
A: EDTA is a chelating agent. It binds divalent cations like Mg²⁺, which are essential cofactors for DNase enzymes. By sequestering Mg²⁺, EDTA inactivates DNases, preventing DNA degradation.

Q10. How do you check the purity and quantity of the isolated DNA?
A: Purity and concentration are checked using a UV spectrophotometer (like a NanoDrop). Pure DNA has an A260/A280 absorption ratio of ~1.8. A lower ratio indicates protein contamination, while a higher ratio suggests RNA contamination.