GATE Biotechnology Recombinant DNA technology and Other Tools in Biotechnology Syllabus

What is an expression vector and what features distinguish it from a cloning vector?

An expression vector is designed to produce protein from a cloned gene. In addition to cloning-vector features, it contains expression signals: a strong promoter, a ribosome binding site (Shine-Dalgarno sequence in prokaryotes / Kozak in eukaryotes), a start codon, a transcription terminator, and often an inducible regulatory element and an affinity tag for purification.

Name common strong/inducible promoters used in bacterial expression vectors.

lac/lacUV5, tac and trc (hybrid trp-lac, IPTG-inducible), trp (tryptophan-regulated), $\lambda$ $p_L$/$p_R$ (temperature-regulated via cI857), araBAD (arabinose-inducible, BAD promoter), and the T7 promoter (requires T7 RNA polymerase, as in the pET system).

How does the T7/pET expression system work?

The gene of interest is placed under a T7 promoter on a pET vector. Expression requires T7 RNA polymerase, supplied by a host strain (e.g., E. coli BL21(DE3)) carrying the T7 polymerase gene under control of the IPTG-inducible lacUV5 promoter. Adding IPTG induces T7 polymerase, which then drives high-level, selective transcription of the target gene.

What is a cDNA library and how is it constructed?

A cDNA library is a collection of clones derived from the mRNA of a cell/tissue. mRNA is isolated, reverse transcribed into first-strand cDNA using reverse transcriptase (primed by oligo-dT), the second strand is synthesized, and the double-stranded cDNA is ligated into a vector. It represents only the expressed (exon-only) genes of that cell type and contains no introns.

What is a genomic DNA library and how does it differ from a cDNA library?

A genomic library contains clones representing the entire genome, made by fragmenting total genomic DNA (e.g., by partial restriction digestion) and cloning the fragments. Unlike a cDNA library, it includes introns, regulatory/intergenic regions, and non-transcribed sequences; it is identical for all tissues of an organism, whereas a cDNA library is tissue/condition-specific and reflects gene expression.

How is the number of clones needed for a complete genomic library calculated?

The number of clones $N$ required to have a given probability $P$ that any sequence is present is given by the Clarke-Carbon formula: $$N = \frac{\ln(1-P)}{\ln\left(1-\frac{i}{g}\right)}$$ where $i$ is the average insert size and $g$ is the genome size (so $i/g$ is the fraction of the genome per clone).

What does reverse transcriptase do and from which sources is it derived?

Reverse transcriptase is an RNA-dependent DNA polymerase that synthesizes a complementary DNA (cDNA) strand using an RNA template; it also has RNase H and DNA-dependent DNA polymerase activities. It is derived from retroviruses, commonly AMV (Avian Myeloblastosis Virus) and M-MLV (Moloney Murine Leukemia Virus).

Outline the basic steps of gene cloning.

(1) Isolate the DNA fragment of interest and cut it (and the vector) with restriction enzymes; (2) ligate the insert into the vector using DNA ligase to form recombinant DNA; (3) introduce the recombinant DNA into a host cell (transformation/transfection); (4) select and screen for recombinant clones (e.g., antibiotic resistance, blue-white screening); (5) grow the host to propagate (clone) the recombinant DNA.

What is the function of DNA ligase in cloning?

DNA ligase catalyzes the formation of a phosphodiester bond between the $3'$-hydroxyl and $5'$-phosphate ends of DNA, sealing nicks and joining insert to vector. T4 DNA ligase can join both cohesive (sticky) and blunt ends and uses ATP as cofactor; E. coli ligase uses NAD$^{+}$ and joins sticky ends only.

What is alkaline phosphatase used for in cloning, and why?

Alkaline phosphatase (e.g., CIP or SAP) removes the $5'$-phosphate groups from the linearized vector ends. Without $5'$-phosphates, the vector cannot self-ligate (recircularize), so it greatly reduces vector background and favors ligation of insert (which still carries phosphates) into the vector.

What is TA cloning?

TA cloning exploits the template-independent addition of a single $3'$ deoxyadenosine (A) overhang to PCR products by Taq polymerase. These products are ligated directly into a linearized vector bearing complementary single $3'$ thymidine (T) overhangs, allowing efficient cloning of PCR amplicons without restriction digestion.

Compare the major strategies/hosts for producing recombinant proteins.

E. coli: cheap, fast, high yield, but no post-translational modifications (PTMs) and risk of inclusion bodies. Yeast (S. cerevisiae, P. pastoris): eukaryotic, can secrete and glycosylate (though hyperglycosylation occurs). Insect cells (baculovirus): good for complex proteins with most PTMs. Mammalian cells (CHO, HEK293): authentic human-like glycosylation and folding, used for therapeutic proteins, but slow and expensive.

What are inclusion bodies and why are they a problem in recombinant protein production?

Inclusion bodies are dense aggregates of misfolded, insoluble recombinant protein that form in the cytoplasm (often in E. coli) during high-level overexpression. The protein is usually inactive and must be solubilized (e.g., with urea/guanidinium) and refolded in vitro to recover activity, complicating purification.

What are fusion tags and give examples used for purification.

Fusion tags are peptide/protein sequences fused to a recombinant protein to aid purification (and sometimes solubility/detection). Examples: His-tag (6xHis, binds Ni-NTA / IMAC), GST (binds glutathione resin), MBP (maltose-binding protein, also improves solubility), FLAG and c-myc epitope tags. Tags can be removed with site-specific proteases (e.g., thrombin, TEV).

What are transposons (transposable elements)?

Transposons are mobile DNA segments that can move (transpose) from one location to another within a genome, often flanked by inverted repeats and encoding transposase. They can cause insertional mutations and genome rearrangements. They are classified as DNA transposons (move via DNA 'cut-and-paste' or 'copy-and-paste') and retrotransposons (move via an RNA intermediate using reverse transcriptase, 'copy-and-paste').

Differentiate replicative (copy-and-paste) and non-replicative (cut-and-paste) transposition.

Non-replicative (conservative, cut-and-paste): the transposon is excised from the donor site and inserted at a new target site; copy number stays the same. Replicative (copy-and-paste): the transposon is copied and one copy moves to a new site while the original remains, increasing copy number (often via a cointegrate intermediate resolved by resolvase).

What is gene targeting and what mechanism does it rely on?

Gene targeting is the precise modification (knockout, knock-in, or replacement) of a specific endogenous gene using homologous recombination between an introduced DNA construct and the chromosomal locus. The targeting vector carries arms homologous to the target gene flanking a selectable marker, enabling site-specific integration (used to make knockout mice via ES cells).

State the principle of the Polymerase Chain Reaction (PCR).

PCR is an in vitro method to exponentially amplify a specific DNA region using a thermostable DNA polymerase, two flanking primers, dNTPs, and $\text{Mg}^{2+}$. Repeated thermal cycles each double the target, so after $n$ cycles the amount of product is approximately $2^{n}$ times the starting target copies.

List the three steps of one PCR cycle with their typical temperatures.

(1) Denaturation: ~94-95 $^\circ$C, separates the double-stranded template. (2) Annealing: ~50-65 $^\circ$C, primers hybridize to complementary sequences. (3) Extension (elongation): ~72 $^\circ$C, Taq polymerase synthesizes new strands $5' \to 3'$. An initial denaturation and final extension flank the cycles.

How is PCR primer melting temperature (Tm) approximately estimated?

For short primers, the Wallace rule estimates: $$T_m = 2(A+T) + 4(G+C)\ ^\circ C$$ where A, T, G, C are the counts of each base. Annealing temperature is usually set ~3-5 $^\circ$C below the lower primer $T_m$.

What is the advantage of using a proofreading polymerase (e.g., Pfu) over Taq in PCR?

Pfu polymerase has $3' \to 5'$ exonuclease (proofreading) activity, giving much higher fidelity (lower error rate) than Taq, which lacks proofreading. Pfu produces blunt-ended products and is preferred for cloning and applications requiring sequence accuracy, though it is slower; Taq is faster and adds 3'-A overhangs.

What is RT-PCR and how does it differ from qPCR (real-time PCR)?

RT-PCR (Reverse Transcription PCR) first converts RNA into cDNA using reverse transcriptase, then amplifies it by PCR; it is used to detect/analyze gene expression. qPCR (real-time/quantitative PCR) monitors product accumulation in real time using fluorescent dyes (SYBR Green) or probes (TaqMan), quantifying DNA by the threshold cycle ($C_t$). The two are often combined as RT-qPCR.

Explain the principle of Sanger (dideoxy / chain-termination) sequencing.

DNA is synthesized by DNA polymerase from a primer in the presence of normal dNTPs plus a small amount of chain-terminating dideoxynucleotides (ddNTPs), which lack the $3'$-OH and stop extension when incorporated. This produces fragments of every possible length ending in a known base; separation by size (capillary/gel electrophoresis, with fluorescently labeled ddNTPs) reveals the sequence.

What is the principle of Maxam-Gilbert sequencing?

Maxam-Gilbert (chemical) sequencing uses base-specific chemical reagents to modify and then cleave end-labeled DNA at particular bases (G; A+G; C; C+T). Partial cleavage generates labeled fragments of varying lengths, which are resolved on a gel to read the sequence. It does not require DNA synthesis but uses hazardous chemicals and is now largely obsolete.

What are common methods of labelling DNA/RNA probes?

Radioactive labels (e.g., $^{32}\text{P}$, $^{35}\text{S}$) and non-radioactive labels (biotin, digoxigenin, fluorophores). Labelling methods include nick translation (DNA Pol I + DNase I incorporates labeled nucleotides), random primer (hexamer) labelling (Klenow fragment, gives high specific activity), end-labelling ($5'$ with T4 polynucleotide kinase and $\gamma$-$^{32}$P-ATP; $3'$ with terminal transferase), and in vitro transcription for RNA probes.

Describe Southern blotting and what it detects.

Southern blotting detects specific DNA sequences. Genomic DNA is digested with restriction enzymes, separated by agarose gel electrophoresis, denatured, and transferred (blotted) onto a nitrocellulose/nylon membrane. The membrane is hybridized with a labelled complementary probe, and the target band is visualized by autoradiography or detection. Used for gene detection, RFLP, and mutation analysis.

Describe Northern blotting and how it differs from Southern blotting.

Northern blotting detects specific RNA (mRNA) molecules to study gene expression and transcript size/abundance. RNA is separated (often under denaturing conditions to prevent secondary structure), transferred to a membrane, and probed with a labelled complementary nucleic acid probe. Difference: Southern analyzes DNA, Northern analyzes RNA (and Northern uses no restriction digestion).

What is a Western blot (for contrast with Southern/Northern)?

A Western blot detects specific proteins. Proteins are separated by SDS-PAGE, transferred to a membrane, and detected using specific primary antibodies followed by enzyme- or fluorophore-conjugated secondary antibodies. Mnemonic: Southern = DNA, Northern = RNA, Western = protein, Eastern = post-translational modifications.

What is in-situ hybridization (ISH) and FISH?

In-situ hybridization detects specific nucleic acid sequences directly within intact cells, tissue sections, or chromosomes using a labelled complementary probe, preserving spatial/positional information. FISH (Fluorescence In-Situ Hybridization) uses fluorescently labelled probes to localize genes/sequences on chromosomes or to detect transcripts, widely used in cytogenetics for mapping and detecting chromosomal abnormalities.

What is DNA fingerprinting and what is its molecular basis?

DNA fingerprinting is a technique to identify individuals based on unique patterns in their DNA. Its basis is variation in the number of tandemly repeated sequences such as VNTRs (minisatellites) and STRs (microsatellites), which differ between individuals. These polymorphic loci are detected by RFLP/Southern blotting or, more commonly, multiplex PCR of STRs, giving an individual-specific banding pattern used in forensics and paternity testing.

What is RAPD and what are its key features?

RAPD (Random Amplified Polymorphic DNA) is a PCR-based marker technique using a single short (~10 bp) primer of arbitrary sequence and low annealing temperature to amplify random genomic regions. Polymorphisms appear as presence/absence of bands due to differences in primer-binding sites. Features: no prior sequence knowledge needed, quick and cheap, but dominant markers with low reproducibility and sensitivity to reaction conditions.