Principles of genetic technology

Restriction endonucleases (often just called "restriction enzymes") are bacterial enzymes that cut DNA at specific short sequences. In bacteria, they evolved as a defence against bacteriophage viruses: invading viral DNA gets sliced up at recognition sites that are protected on the bacterium's own DNA by methylation. In the lab, they are the fundamental cutting tools of molecular cloning.

Specific palindromic recognition sites.

Each enzyme recognises a specific palindromic DNA sequence, typically 4-8 base pairs long. A "palindrome" in molecular biology means the sequence reads the same on both strands when read 5'→3'. The most famous example is EcoRI (from E. coli):

5'-G A A T T C-3'
3'-C T T A A G-5'

Read 5'→3' on the top strand: G-A-A-T-T-C. Read 5'→3' on the bottom strand (right to left): G-A-A-T-T-C. The same sequence.

EcoRI cuts between G and A on both strands:

5'-G     A A T T C-3'
3'-C T T A A     G-5'

leaving sticky ends — short single-stranded overhangs of 5'-AATT-3'.

Other named enzymes:

Enzyme	Source	Recognition site	Cut
EcoRI	E. coli	GAATTC	sticky end, 5'-AATT
BamHI	Bacillus amyloliquefaciens	GGATCC	sticky end, 5'-GATC
HindIII	Haemophilus influenzae	AAGCTT	sticky end, 5'-AGCT
SmaI	Serratia marcescens	CCCGGG	blunt end (no overhang)

Sticky ends vs blunt ends.

• Sticky-end cutters (EcoRI, BamHI, HindIII) leave single-stranded overhangs. Two pieces of DNA cut with the same sticky-end enzyme have complementary overhangs that base-pair specifically.
• Blunt-end cutters (SmaI, EcoRV) make clean cuts with no overhang. Blunt-end ligation is possible but much less efficient because there are no base pairs to hold the fragments together while DNA ligase joins them.

For most cloning, sticky-end enzymes are preferred.

Using restriction enzymes in cloning.

1. Cut the source DNA (containing the gene of interest) with a restriction enzyme that has recognition sites flanking the gene → gene released with sticky ends.
2. Cut the vector (plasmid) with the same enzyme → matching sticky ends.
3. Mix gene + vector → sticky ends base-pair.
4. DNA ligase seals the gaps with phosphodiester bonds → recombinant plasmid.

The critical point — the same enzyme must cut both. Different enzymes give different overhangs that won't anneal.

Once the gene of interest has been cut out of the source DNA, it must be inserted into a vector to be carried into the host cell.

Vectors.

A vector is a DNA molecule used to ferry foreign DNA into a host cell. Common vectors:

• Plasmids — small (typically 2-10 kb) circular DNA molecules found naturally in bacteria, replicating independently of the chromosome. The standard vector for bacterial cloning. Examples: pUC19, pBluescript, pBR322.
• Viral vectors — modified viruses (often retroviruses or adenoviruses) used for animal cells. The virus delivers DNA into the host cell using its natural infection machinery.
• Bacterial artificial chromosomes (BACs) — for carrying very large DNA inserts (up to 300 kb).
• Ti plasmid of Agrobacterium tumefaciens — for plant cells.

A useful plasmid vector contains:

• An origin of replication so it can be copied inside the bacterium.
• A multiple cloning site with several restriction enzyme recognition sites for inserting the gene of interest.
• One or more marker genes for selection (see below).

DNA ligase.

After the gene of interest is added to the cut vector and sticky ends base-pair, DNA ligase seals the nicks. Most commonly T4 DNA ligase from a bacteriophage. The enzyme catalyses the formation of phosphodiester bonds between the 3'-OH of one fragment and the 5'-phosphate of the next, closing both strands and producing an intact recombinant plasmid.

Transformation.

The recombinant plasmid is then introduced into a host bacterium — a process called transformation. Two main methods:

(1) Electroporation. A high-voltage electric pulse briefly destabilises the bacterial membrane, allowing the plasmid to enter. High efficiency but requires special equipment.

(2) Heat-shock. Bacteria are first treated with calcium chloride to make their membranes permeable ("chemically competent" cells). The cells + plasmid mixture is held on ice, then briefly heated to 42 °C for ~45 seconds, then returned to ice. The heat shock drives some plasmid molecules through the membrane.

Even after optimised transformation, only a small fraction of cells take up the plasmid — typically 0.0001%-0.1%. The transformed cells must be identified.

Marker genes — identifying transformed cells.

The plasmid is engineered to carry a marker gene alongside the gene of interest. The marker provides a distinguishing feature that allows transformed cells to be picked out:

(a) Antibiotic-resistance markers (the classic approach).

• The plasmid carries an antibiotic-resistance gene (commonly ampR, ampicillin resistance, encoding β-lactamase).
• Bacteria are spread on agar plates containing the antibiotic.
• Only transformed cells (with the plasmid + resistance gene) survive and form colonies. Untransformed cells are killed.

(b) Fluorescent protein markers (modern alternative).

• Plasmid carries the gene for green fluorescent protein (GFP) from jellyfish Aequorea victoria (or RFP from coral).
• Transformed colonies glow green under UV illumination.

(c) Blue-white screening (to identify cells with the actual insert).

• The gene-of-interest insertion site sits within a lacZα gene encoding β-galactosidase.
• Plates contain X-gal (colourless artificial substrate of β-galactosidase) and IPTG (inducer of lacZ).
• If lacZ is intact (empty plasmid — no insert), bacteria produce β-galactosidase, X-gal is cleaved to a blue product → blue colonies.
• If the insert is present, it disrupts lacZα, no β-galactosidase produced, X-gal stays colourless → white colonies.
• Pick white colonies → contain the recombinant plasmid with the desired insert.

Combining antibiotic selection (plasmid is present) with blue-white screening (insert is present) gives a reliable two-step selection for correctly assembled recombinants.

When cloning a eukaryotic gene into bacteria, the gene must be in a form that bacteria can transcribe and translate correctly. Genomic eukaryotic DNA contains introns — non-coding sequences that must be spliced out of pre-mRNA before translation. Bacteria lack the spliceosome and cannot remove introns. So researchers don't clone the genomic gene; they clone cDNA made from the mature mRNA.

What is reverse transcriptase?

Reverse transcriptase is an enzyme from retroviruses (notably HIV and avian myeloblastosis virus). It synthesises DNA from an RNA template — the reverse of the central dogma's normal transcription direction, and the reason for the name "reverse" transcriptase.

In nature, retroviruses use it to integrate their RNA genome into the host's DNA. In molecular biology labs, researchers exploit the same enzyme to make DNA copies of mRNA molecules.

How cDNA is made — step by step.

Step 1 — Isolate mRNA. Total mRNA is extracted from a tissue or cell type that expresses the gene of interest at high level. For example:

• Insulin gene → mRNA from pancreatic β-cells.
• Haemoglobin gene → mRNA from reticulocytes (immature red blood cells).
• Crystallin (eye lens) genes → mRNA from lens fibre cells.

Isolation typically uses oligo-dT magnetic beads — strands of T residues that bind to the poly-A tail of mature eukaryotic mRNA. Other RNAs (rRNA, tRNA) lack poly-A tails and are washed away.

Step 2 — First-strand synthesis. A short DNA primer (often oligo-dT, complementary to the poly-A tail) anneals to the mRNA. Reverse transcriptase then uses the mRNA as template and free dNTPs to synthesise a complementary DNA strand in the 5'→3' direction. Product: a hybrid RNA-DNA double-stranded molecule.

Step 3 — Second-strand synthesis. The mRNA template is degraded by RNase H (which specifically digests RNA in RNA-DNA duplexes), leaving short RNA fragments that act as primers. DNA polymerase I uses these to synthesise the second strand of cDNA, producing double-stranded cDNA.

Step 4 — Cloning the cDNA. The double-stranded cDNA is now ready for insertion into a vector. Linkers carrying restriction enzyme sites are often ligated to the cDNA ends to allow standard cutting and ligation into a plasmid.

Why cDNA is preferred over genomic DNA for cloning a eukaryotic gene in bacteria.

(1) No introns. Bacteria lack the spliceosome. cDNA is made from mature mRNA in which introns are already removed, so bacteria can transcribe and translate it directly into the correct protein.

(2) Enriched for the desired gene. Choosing a cell type that highly expresses the gene of interest gives a starting mRNA pool dominated by that gene. Far easier than picking the gene out of total genomic DNA.

(3) Smaller and easier to handle. Eukaryotic genes with introns can be huge (the dystrophin gene is 2.4 megabases). cDNA contains only exons — much shorter, fits comfortably in plasmids.

Applications of cDNA cloning.

• Recombinant human insulin for diabetes treatment.
• Recombinant growth hormone for dwarfism.
• Factor VIII for haemophilia A.
• Vaccines like recombinant hepatitis B surface antigen.
• cDNA libraries for tissue-specific gene discovery.

The polymerase chain reaction (PCR) is one of the most transformative techniques in biology. Invented by Kary Mullis in 1983 (Nobel Prize 1993), it allows a specific region of DNA to be amplified exponentially in a test tube — billions of copies of the target sequence in 2-3 hours.

The reaction mixture (in a small plastic tube):

• Template DNA — the source DNA containing the target region (can be as little as a single molecule).
• Two primers — short DNA oligonucleotides (~20 nucleotides each) complementary to the sequences flanking the target. One for each strand.
• Free deoxynucleotides (dNTPs) — A, T, G, C, the building blocks of new DNA.
• Taq DNA polymerase — a heat-stable polymerase from Thermus aquaticus, a hot-springs bacterium.
• Reaction buffer containing Mg²⁺ (a cofactor for the polymerase).

The reaction takes place in a thermal cycler — a programmable heating block that rapidly cycles temperatures between the three values below.

The three steps of one PCR cycle:

---

Step 1 — Denaturation at 95 °C (~90 s).

Heat to 95 °C. The hydrogen bonds holding the two DNA strands together break; the double helix separates into two single strands. This provides single-stranded template for the polymerase.

Why 95 °C? — to fully denature the DNA. Why doesn't this destroy the polymerase? — because Taq polymerase is heat-stable. It evolved in a hot-springs bacterium adapted to ~70 °C, so it survives 95 °C without inactivation. Before Taq's discovery, PCR required fresh polymerase to be added every cycle — impractical.

---

Step 2 — Annealing at ~55 °C (~30 s).

Cool to about 55 °C (the exact value depends on primer sequence — usually 5 °C below the primers' melting temperature, $T_m$).

The two primers in the reaction anneal (base-pair) to complementary sequences on the flanking regions of the target — one primer on each strand. The primers must be designed in advance with sequences complementary to known regions just outside the target.

Why this temperature? — low enough for the short primers to base-pair specifically, high enough to prevent the long template strands from re-annealing to each other (which would block the primers).

---

Step 3 — Extension at 72 °C (~60 s).

Heat to 72 °C — the optimum temperature for Taq polymerase.

Taq polymerase extends each primer in the 5' → 3' direction, using the single-stranded template as a guide and free dNTPs as substrate. Each extension takes ~30 seconds for ~1 kb of product.

By the end of this step, each original DNA molecule has become two double-stranded copies of the target region.

---

Repeating the cycle → exponential amplification.

The three steps are repeated 30 to 40 times in the thermal cycler. Each cycle doubles the target DNA:

$\text{copies after } n \text{ cycles} = 2^n$

Cycle	Copies
0	1
1	2
5	32
10	~1,024
20	~10⁶ (1 million)
30	~10⁹ (1 billion)

So 30 cycles starting from a single template molecule can produce a billion copies in about 2-3 hours — extraordinary amplification.

Important details:

• Primers define what is amplified. Only the region flanked by the two primers gets exponentially amplified. Off-target sequences are amplified at most linearly.
• PCR product length is the distance between the primer binding sites.
• Negative controls (no template) are run alongside to detect contamination — even a single contaminating molecule will be amplified.

Applications.

PCR underlies many of modern biology's techniques:

• DNA cloning — amplify a gene before inserting it into a vector.
• Forensic DNA profiling — amplify STR (short tandem repeat) loci from minute crime-scene samples.
• Medical diagnostics — detect pathogens (COVID-19 RT-PCR; HIV viral load; Mycobacterium tuberculosis).
• Prenatal genetic testing — amplify DNA from amniocentesis or CVS samples.
• Paleogenetics — amplify DNA from ancient bones (Neanderthals, woolly mammoths).
• Genetic engineering — site-directed mutagenesis to introduce specific changes.

Gel electrophoresis separates DNA fragments (or RNA, or proteins) by size so that they can be visualised, identified or extracted. It is a routine step in essentially every molecular biology lab.

The apparatus.

• A horizontal tank containing buffer solution (typically TAE — Tris-acetate-EDTA — buffer) that conducts electricity.
• An agarose gel cast in the tank. Agarose is a polysaccharide from seaweed; it forms a porous gel resembling a microscopic mesh. Typical gel concentration:
  • 0.5-0.8% agarose — separating large fragments (5-20 kb).
  • 1-2% agarose — typical PCR product sizes (200-5000 bp).
  • 2-3% agarose — small fragments (50-500 bp).
• Wells at one end of the gel, formed by a comb during casting.
• Electrodes — cathode (−ve) at the wells end, anode (+ve) at the far end.
• A power supply (typically 50-150 V).

Loading and running.

DNA samples are mixed with loading dye — a glycerol-based solution containing tracking dyes (bromophenol blue, xylene cyanol). The glycerol makes the sample dense enough to sink into the well; the tracking dyes allow the run progress to be visualised.

Samples are pipetted into the wells. A DNA size marker (ladder) containing fragments of known sizes is run in a separate well. Voltage is applied, and the run continues for 30-90 minutes.

The physics of separation — TWO independent properties.

(1) Why DNA migrates: charge. Each phosphate group in DNA's sugar-phosphate backbone carries a negative charge at neutral pH. DNA is therefore uniformly negatively charged along its length. In the electric field, all DNA fragments migrate towards the anode (+ve electrode).

Important: charge causes migration but does not separate by size — all fragments have the same charge-to-mass ratio.

(2) Why DNA separates by size: the gel as a molecular sieve. The agarose mesh has pores roughly the size of medium DNA fragments. Smaller fragments find it easier to weave through the pores and migrate faster; larger fragments are physically held back by the mesh.

The relationship between size and migration distance is approximately logarithmic — small fragments are much faster than predicted by a linear relationship.

After the run, fragments have separated into bands at different positions along the gel: smallest at the leading edge (furthest from wells), largest closest to the wells.

Visualisation.

The gel is stained with a DNA-binding fluorescent dye that intercalates between DNA base pairs and emits visible light under UV illumination:

• Ethidium bromide — historically standard; effective but a known mutagen, now being phased out.
• SYBR Safe and GelRed — safer modern alternatives.

The stained gel is photographed under UV light; DNA bands appear as bright stripes against a dark background.

Size determination using the ladder.

By running a DNA size marker (ladder) alongside the samples, the size of unknown bands can be estimated. The ladder contains fragments of known sizes (e.g. 100 bp, 200 bp, 500 bp, 1 kb, 2 kb). A plot of log(size) vs migration distance gives an approximately straight line that can be used to interpolate unknown sizes.

Applications of gel electrophoresis.

• Checking the success of restriction digests — expected number and size of fragments?
• Checking the specificity of PCR — single band of correct size, or multiple bands suggesting non-specific amplification?
• Estimating fragment sizes.
• Purifying DNA bands of interest — the band can be cut out of the gel and the DNA extracted.
• DNA profiling in forensic science — comparing STR fragment patterns between samples.
• Quality control for cloning, sequencing, library preparation.

A typical workflow combines PCR (to amplify) + gel electrophoresis (to check / size) + Sanger sequencing (to read).

Two additional technologies — DNA microarrays and DNA sequencing — round out the modern genetic-technology toolkit.

DNA microarrays (DNA chips).

A microarray is a glass slide or silicon chip carrying thousands to millions of different single-stranded DNA probes, each one a known short sequence, immobilised in a known position on a grid. A typical chip can carry probes for every gene in a genome.

How it works:

1. Sample preparation — DNA or RNA from the test sample is extracted. If studying gene expression (transcriptomics), mRNA is converted to cDNA. The sample is labelled with fluorescent dye.
2. Hybridisation — the labelled sample is washed across the chip. Complementary base pairing allows sample fragments to bind specifically to their matching probes; non-matching sequences are washed away.
3. Imaging — the chip is scanned with a laser; positions where labelled DNA has bound fluoresce. The pattern of fluorescence reveals which genes/sequences are present (or expressed) in the sample.

Applications:

• Gene expression profiling — which genes are turned on in tumours, embryos, infected cells?
• Genotyping — SNP arrays for studying genetic variation (e.g. ancestry tests, GWAS).
• Comparative genomic hybridisation — detect deletions and duplications in cancer cells.
• Pathogen detection — diagnostic chips for influenza strains, foodborne pathogens.

Two-colour microarrays label test and control samples with different dyes (red and green) and apply them together; ratios reveal differential expression.

---

DNA sequencing.

DNA sequencing reads the actual nucleotide sequence of a DNA fragment — the ultimate analytical resolution.

(1) Sanger sequencing (chain-termination method).

Developed by Fred Sanger (1977 — earned him a second Nobel Prize). The principle:

• The DNA region to be sequenced is used as a template.
• A primer, DNA polymerase, and free dNTPs are added.
• Crucially, a small amount of dideoxynucleotides (ddNTPs) is included. Each ddNTP lacks the 3'-OH group, so once incorporated, DNA synthesis terminates at that point.
• In the modern version, each of the four ddNTPs (ddA, ddT, ddG, ddC) is labelled with a different fluorescent dye.
• Synthesis produces a mixture of DNA fragments of every possible length, each terminating in a ddNTP whose colour identifies the last base.
• Fragments are separated by capillary electrophoresis — automated, high resolution.
• A laser detects the colour of each fragment as it passes; the sequence is read off the colour ladder.

Sanger sequencing typically reads 500-1000 bp per reaction with high accuracy. Still used for targeted sequencing of small regions (e.g. checking a single PCR product, confirming a clone).

(2) Next-generation sequencing (NGS).

From ~2005 onwards, next-generation sequencing technologies (Illumina, Ion Torrent, PacBio, Oxford Nanopore) revolutionised the field by sequencing millions of fragments in parallel. Key features:

• Massively parallel — many sequencing reactions run simultaneously on a single chip or flowcell.
• Short reads (Illumina: ~150-300 bp per read) or long reads (PacBio, Oxford Nanopore: 10,000+ bp per read).
• Whole genomes in days.
• Cost has fallen dramatically: the first human genome (2003) cost $3 billion and took 13 years; today a human genome can be sequenced for ~$200 in a few days.

Applications of NGS:

• Personal genomics — consumer ancestry tests, medical sequencing for rare diseases.
• Cancer genomics — sequencing tumours to identify mutations for targeted therapy.
• Microbiome studies — sequencing all microbes in a sample (metagenomics).
• Population genetics — sequencing thousands of individuals.
• Evolutionary biology — sequencing extinct species (Neanderthals, woolly mammoths).
• Pandemic surveillance — tracking SARS-CoV-2 variants in real time.

NGS underlies almost all modern genomic research and is increasingly part of routine medical care.