The Basic Tools
of Molecular Biology

Molecular Biology is the study of the structure and function of the molecules that make up living cells — primarily nucleic acids (DNA and RNA) and proteins. It asks: how do these molecules work, and how do they carry out the instructions for life?

Genetics is the study of heredity and variation — how traits are passed from parent to offspring and why individuals within a species differ from each other. Molecular biology gives us the tools to study genetics at the DNA level.

Biotechnology is the use of living organisms (microorganisms, cells, or cell components) to make useful products. Traditional biotechnology has existed for thousands of years — bread, beer, cheese all rely on microbial fermentation. Modern biotechnology uses recombinant DNA (rDNA) technology to insert or modify genes in organisms to produce specific desired proteins — insulin, antibiotics, enzymes, vaccines, and more.

This entire topic (Topic 2) is about the individual tools that make rDNA technology possible. Each tool has a specific job in the pipeline. Think of them like instruments in a surgeon's kit — you need to know what each one does, when to use it, and how it works.

The overall workflow of a genetic modification experiment is: isolate a vector, cut it open with restriction enzymes, insert your gene of interest, ligate it shut, transform it into a host cell, screen for successful clones, culture them, and harvest your product. Every tool in this topic fits somewhere in that pipeline.

Figure 1: Overview of a typical genetic modification procedure — from isolating the vector and gene of interest, through insertion and transformation, to cloning and harvesting the product.

A vector is a self-replicating DNA molecule used to carry a gene of interest into a host cell. Think of it as a delivery truck — the gene is the cargo, and the vector is the vehicle that takes it where it needs to go and ensures it gets copied when the cell divides.

Four Essential Properties of a Good Vector

1. Self-replication: The vector must be able to replicate independently inside the host cell. It needs its own origin of replication (ori) — a DNA sequence where replication machinery binds and starts copying. The ori is typically AT-rich (adenine-thymine rich) because A-T base pairs have only 2 hydrogen bonds (vs. 3 for G-C), making them easier to separate and initiate replication.

2. A cloning site (MCS): The vector must have a Multiple Cloning Site (MCS) — a short region containing recognition sequences for many different restriction enzymes. This is where you insert your foreign DNA. Having multiple enzyme sites gives you flexibility to use different restriction enzymes for different inserts.

3. A selectable marker: The vector carries an antibiotic resistance gene (e.g., ampicillin resistance, amp^R). After transformation, you plate bacteria on antibiotic-containing media. Only cells that took up the vector survive — this is how you select for transformed cells.

4. Small size: Smaller vectors are easier to manipulate, easier to get into cells during transformation, and produce higher copy numbers (more copies per cell). This is why plasmids (typically 2-10 kb) are the workhorse of molecular biology.

Types of Vectors

Table 1: Vector types ranked by insert capacity. As insert size increases, the vector system becomes more complex and harder to work with.

The key insight is that insert size determines which vector you use. A plasmid can carry a small gene (~15 kb), but if you need to clone a 200 kb chunk of human genome, you need a BAC or YAC. Think of it like shipping: an envelope (plasmid) for a letter, a box (cosmid) for a book, a shipping container (BAC) for furniture.

Plasmids in detail: In your Week 1 practical, you isolated a plasmid from E. coli using a miniprep. That plasmid is a circular, extrachromosomal DNA molecule that replicates independently of the bacterial chromosome. It carries an antibiotic resistance gene (selective marker), a multiple cloning site (MCS), and an origin of replication (ori).

Figure 2: Structure of a typical plasmid vector showing the origin of replication (ori), antibiotic resistance gene (selective marker), and multiple cloning site (MCS).

Restriction enzymes (also called restriction endonucleases) are proteins that recognise and cut DNA at specific short sequences called recognition sites. They were originally discovered in bacteria, where they evolved as a defence mechanism against bacteriophage (viral) infection — the enzymes cut up invading viral DNA into useless fragments, 'restricting' the virus's ability to replicate.

Scientists co-opted these bacterial enzymes for use in the laboratory. Restriction enzymes are the foundational tool of molecular cloning — without them, we cannot cut DNA precisely.

How Restriction Enzymes Work

Each restriction enzyme recognises a specific DNA sequence (usually 4-8 base pairs long) called its recognition site. These sites are typically palindromic — meaning the sequence reads the same on both strands in the 5' to 3' direction. For example, EcoRI recognises: 5'-GAATTC-3' / 3'-CTTAAG-5'. Notice that both strands read GAATTC when read 5' to 3'.

When the enzyme finds its recognition site, it cuts both strands of the DNA. The cut can produce two types of ends:

1. Sticky ends (cohesive ends): The enzyme cuts the two strands at different positions within the recognition site, leaving short single-stranded overhangs. Example: EcoRI cuts between G and AATTC on each strand, producing 4-nucleotide overhangs (5'-AATT). These sticky ends are incredibly useful because they can base-pair with complementary sticky ends from ANY DNA cut with the same enzyme — this is the basis of recombinant DNA.

2. Blunt ends: The enzyme cuts both strands at exactly the same position, leaving no overhangs. Example: SmaI cuts in the middle of CCCGGG. Blunt ends can be ligated together, but less efficiently than sticky ends because there is no complementary base-pairing to hold the fragments in place.

Figure 3: Restriction enzyme EcoRI cutting DNA at its recognition site (GAATTC), producing sticky ends that base-pair with complementary ends from another DNA source to form recombinant DNA.

Table 2: Common restriction enzymes. Note that all recognition sites are palindromic. The enzyme name comes from the organism it was isolated from.

Why This Matters for Cloning

To make recombinant DNA, you cut BOTH the vector AND the DNA containing your gene of interest with the same restriction enzyme. This produces complementary sticky ends on both pieces. When you mix them together, the sticky ends base-pair, and DNA ligase seals the joins. Result: your gene is now inside the vector.

Ligation is the process of joining two pieces of DNA together using the enzyme DNA ligase. After restriction enzyme digestion has produced compatible ends on your vector and insert, ligase catalyses the formation of phosphodiester bonds between the 3'-hydroxyl and 5'-phosphate groups of adjacent nucleotides, sealing the sugar-phosphate backbone.

Restriction enzymes cut; ligase joins. These are the two essential enzymatic tools. The restriction enzyme creates the compatible ends; ligase makes the join permanent. The resulting recombinant DNA molecule can then be used for protein expression in a host cell.

Sticky ends vs. blunt ends in ligation: Ligation of sticky ends is much more efficient than blunt-end ligation. Sticky ends provide transient base-pairing that holds the fragments together, positioning them correctly for ligase. Blunt-end ligation requires higher concentrations of ligase and longer incubation because the fragments have no 'grip' on each other — they must collide in exactly the right orientation by chance.

After ligation and transformation, you have a mixed population: some cells got the recombinant plasmid (with your insert), some got re-ligated empty vector, and some got no plasmid at all. You need methods to figure out which cells have your gene. There are three main approaches covered in this topic:

Method 1: Blue/White Screening

This method uses insertional inactivation of the lacZ gene. The vector has the MCS (where you insert your gene) located inside the lacZ gene, which encodes beta-galactosidase. If an insert is present, it disrupts lacZ and the cell cannot produce functional beta-galactosidase.

Plates contain X-gal (a substrate) and IPTG (an inducer). Functional beta-galactosidase cleaves X-gal to produce a blue colour. So: Blue colonies = no insert (lacZ intact, enzyme works). White colonies = insert present (lacZ disrupted, no enzyme, no blue). You pick the white colonies.

Figure 4: Blue/white screening — blue colonies have functional lacZ (no insert); white colonies have disrupted lacZ (insert present).

Method 2: Restriction Enzyme Digestion + Electrophoresis

Extract the plasmid from candidate colonies, digest it with the same restriction enzyme used for cloning, and run the fragments on a gel. If the insert is present, you will see the expected fragment sizes — one band for the vector backbone and one for the insert. If there is no insert, you only see the vector band.

Method 3: Sanger Sequencing

The gold standard for confirming your insert. Sequencing reads the actual nucleotide sequence of the DNA in your plasmid, proving that the correct gene was inserted in the correct orientation and reading frame. This is covered in detail in Section 7.

Table 3: Screening methods compared. In practice, labs use all three in sequence — blue/white first (fast, cheap), then digest + gel, then sequencing to confirm.

Gel electrophoresis is a technique for separating DNA fragments based on their size. It is used constantly in molecular biology — to check restriction digests, verify PCR products, assess miniprep quality, and more. If restriction enzymes are the scissors and ligase is the glue, the gel is the quality control inspector — it lets you see what you have.

How It Works — Step by Step

1. The gel matrix: Agarose (a polysaccharide from seaweed) is dissolved in buffer, heated, poured into a mould, and allowed to solidify. It forms a mesh of tiny pores — like a molecular obstacle course. The concentration of agarose determines pore size: higher concentration = smaller pores = better separation of small fragments.

2. Loading: DNA samples are mixed with a loading dye (which adds colour and density so the sample sinks into the well) and pipetted into wells at one end of the gel. A DNA ladder (molecular weight marker) containing fragments of known sizes is loaded alongside — this is your ruler for measuring fragment sizes.

3. Running: An electric field is applied across the gel. DNA is negatively charged (due to phosphate groups in the backbone), so it migrates toward the positive electrode (anode). Smaller fragments move faster through the pores; larger fragments move slower. After sufficient time, fragments are separated by size.

4. Visualisation: The gel is stained with a DNA-binding fluorescent dye (e.g., ethidium bromide or SYBR Safe) and viewed under UV light. DNA fragments appear as bands. Each band represents fragments of a particular size. By comparing band positions to the ladder, you can determine the size of your fragments in base pairs (bp) or kilobases (kb).

Figure 5: Gel electrophoresis apparatus — DNA samples are loaded into wells at the negative electrode end and migrate toward the positive electrode through the agarose gel matrix.

Reading a DNA Gel — Practical Skill

On a gel image: the wells are at the top, the positive electrode is at the bottom. Bands near the top are large fragments; bands near the bottom are small fragments. The ladder (usually lane 1) provides reference points. To estimate an unknown fragment's size, find where its band falls relative to the ladder bands and interpolate.

Figure 6: A DNA gel showing fragments separated by size. The ladder (marker) in the left lane contains fragments of known sizes. Compare unknown band positions to the ladder to estimate fragment sizes.

Sanger sequencing (also called chain-termination sequencing or dideoxy sequencing) is the method for determining the exact order of nucleotides in a piece of DNA. It was developed by Frederick Sanger in 1977 and remained the gold standard for decades.

How It Works

1. The DNA to be sequenced is mixed with a primer (short DNA sequence that tells DNA polymerase where to start), DNA polymerase, normal deoxynucleotides (dNTPs: dATP, dTTP, dCTP, dGTP), and small amounts of dideoxynucleotides (ddNTPs).

2. ddNTPs are the key trick. They lack a 3'-hydroxyl group, which means once a ddNTP is incorporated into a growing DNA chain, no further nucleotides can be added — the chain terminates. Because ddNTPs are present at low concentrations mixed with normal dNTPs, chain termination happens randomly at different positions in different copies of the template.

3. The result is a collection of DNA fragments of every possible length, each ending with a fluorescently labelled ddNTP (each base — A, T, C, G — has a different colour label).

4. These fragments are separated by size using capillary electrophoresis (like gel electrophoresis but in a thin tube). A laser reads the fluorescent colour of each fragment as it passes a detector, and the sequence is read as a series of coloured peaks on a chromatogram.

Figure 7: Sanger sequencing chromatogram — each coloured peak represents a nucleotide base (A, T, C, G). The sequence is read from left to right, shortest fragments first.

Blotting techniques transfer separated molecules from a gel onto a membrane, where they can be detected using specific probes. The three main types are named after their inventor (Edwin Southern) and by humorous analogy to his name:

Table 4: The three blotting techniques. The naming convention is a scientific inside joke — Southern was the inventor's actual name; Northern and Western were named to continue the compass theme.

The General Blotting Workflow

1. Separate: Run your sample (DNA, RNA, or protein) on a gel to separate molecules by size.

2. Transfer: Transfer the separated molecules from the gel onto a membrane (nitrocellulose or PVDF) — this is the actual 'blotting' step, often done by capillary action or electric transfer.

3. Probe: Incubate the membrane with a probe that specifically binds your target. For Southern/Northern blots, the probe is a labelled nucleic acid that hybridises to the complementary sequence. For Western blots, the probe is an antibody that binds the target protein.

4. Detect: Visualise where the probe bound — this tells you the size and abundance of your target molecule.

PCR is a technique for making millions of copies of a specific DNA sequence from a tiny starting amount. It is one of the most important inventions in molecular biology — it earned Kary Mullis the Nobel Prize in 1993. Think of it as a molecular photocopier: you put in one copy of a page and get billions out.

The Three Steps of Each PCR Cycle

PCR works by repeating a cycle of three temperature-controlled steps, typically 25-35 times:

Table 5: The three steps of a PCR cycle. Each cycle doubles the amount of target DNA, so after 30 cycles you have approximately 2³⁰ = ~1 billion copies.

Figure 8: The three steps of a PCR cycle — (1) Denaturing at 95°C separates strands, (2) Annealing at 55°C allows primers to bind, (3) Extension at 72°C as Taq polymerase synthesises new strands. Each cycle doubles the number of DNA molecules.

What You Need for PCR

• Template DNA — the DNA containing the sequence you want to amplify (even a tiny amount works)
• Two primers — short synthetic single-stranded DNA sequences (~18-25 nucleotides) that flank the target region. One binds the forward strand, one binds the reverse strand.
• Taq DNA polymerase — a heat-stable DNA polymerase isolated from Thermus aquaticus, a bacterium that lives in hot springs. It survives the 94-98°C denaturation step.
• dNTPs — the four deoxynucleotide building blocks (dATP, dTTP, dCTP, dGTP)
• Buffer + MgCl₂ — provides optimal conditions for polymerase activity

Applications of PCR

PCR is used everywhere in modern biology and medicine: cloning DNA for recombination, amplifying DNA to detectable levels from trace samples (forensics, ancient DNA), sequencing DNA, diagnosing genetic diseases, detecting pathogens (e.g., COVID-19 PCR tests), paternity testing, and identifying organisms from environmental samples.

A miniprep (miniature preparation) is a laboratory method for extracting plasmid DNA from a small volume of bacterial culture. This was the procedure you performed in your Week 1 practical. The principle is to selectively isolate the small, circular plasmid DNA while removing the much larger chromosomal DNA, RNA, proteins, and cell debris.

The Three Key Steps (Alkaline Lysis Method)

Step 1 — Resuspension (P1 buffer): Bacterial pellet is resuspended in a buffer containing Tris (pH buffer), EDTA (chelates Mg²⁺ to inhibit DNases), and RNase A (degrades RNA). This creates a uniform suspension.

Step 2 — Lysis (P2 buffer): NaOH + SDS (sodium dodecyl sulfate) is added. NaOH denatures both chromosomal and plasmid DNA and proteins. SDS (a detergent) dissolves the cell membrane. The solution becomes clear and viscous as cells lyse. Critical: do not mix too vigorously or incubate too long — this can shear chromosomal DNA into small fragments that co-purify with the plasmid.

Step 3 — Neutralisation (N3 buffer): Potassium acetate at acidic pH is added. This does three things: (a) neutralises the NaOH, causing the large chromosomal DNA to re-anneal into an insoluble tangled mass; (b) SDS precipitates with potassium, trapping proteins and chromosomal DNA; (c) the small circular plasmid DNA, being supercoiled and small, re-anneals correctly and stays in solution. Centrifugation pellets the debris; the plasmid remains in the clear supernatant.

The supernatant is then passed through a silica column (plasmid DNA binds to silica in high-salt conditions), washed to remove contaminants, and eluted with low-salt buffer or water. The result is purified plasmid DNA ready for downstream applications (restriction digest, ligation, sequencing, transformation).

Now that you understand each tool individually, here is how they all fit together in a typical recombinant DNA experiment. Each step uses one or more of the tools from this topic:

Table 6: The complete cloning workflow showing where each tool from Topic 2 is used. Every tool has a specific job — none works in isolation.

Sticky Ends vs. Blunt Ends

Cutting Tools vs. Joining Tools

All Tools — One-Line Summary

Vector Essentials: ROSS

Mnemonic: R-O-S-S: Replication, Opening (MCS), Selection marker, Small size
Meaning: The four properties every good vector needs.

Blotting: SNoW DRoP

Mnemonic: Southern=DNA, Northern=RNA, Western=Protein
Meaning: Read the capital initials: S-N-W matches D-R-P.

PCR Steps: DAE

Mnemonic: D-A-E: Denature, Anneal, Extend
Meaning: The three steps of every PCR cycle, in order. Hot-warm-medium.

Gel Direction

Mnemonic: DNA is Negative → runs to the Positive (red) electrode
Meaning: Remember: 'Run to Red' — DNA migrates toward the anode.

Blue/White Screening

Mnemonic: Blue = Bad (no insert), White = Winner (insert present)
Meaning: Insert disrupts lacZ → no enzyme → no blue → white colony.

Restriction Enzyme Origin

Mnemonic: Eco = E. coli, Hind = H. influenzae, Bam = B. amyloliquefaciens
Meaning: First 3 letters = genus + species of source organism.

Miniprep Steps

Mnemonic: R-L-N: Resuspend, Lyse, Neutralise
Meaning: The three buffers (P1, P2, N3) in order. Plasmid survives because it is small and circular.

Sanger Sequencing Key

Mnemonic: ddNTPs = Dead-end NTPs (no 3'-OH = chain terminates)
Meaning: dideoxy = missing the 3'-OH group needed for the next nucleotide to attach.

Sticky vs Blunt

Mnemonic: Sticky = Staggered cut = overhangs = efficient ligation
Meaning: Blunt = straight cut = no overhangs = less efficient ligation.