Mutagenesis, CRISPR-Cas, biosensors and interaction tools

Common methods

1. Oligonucleotide-directed (Kunkel) mutagenesis: a single-stranded, uracil-containing template is produced in a dut⁻ ung⁻ E. coli host (dut⁻ raises the dUTP pool so uracil is incorporated; ung⁻ stops it being removed). A synthetic mutagenic primer carrying the desired mismatch is annealed, extended in vitro by a non-strand-displacing DNA polymerase and sealed by DNA ligase. On transformation into a wild-type (ung⁺) host, uracil-DNA glycosylase removes the uracils from the parental strand, leaving abasic sites that inactivate it as a template, so the uracil-free mutant strand is strongly enriched.

2. PCR-based overlap-extension mutagenesis: two complementary internal primers, each carrying the desired mutation, are used with two flanking primers in two separate first-round PCRs to generate two fragments that overlap at the mutation. The fragments are mixed, denatured and re-annealed so their overlapping (mutant) ends prime each other, and a final PCR with only the flanking primers amplifies the full-length mutant product, which is then cloned.

3. Whole-plasmid PCR mutagenesis (single-tube, DpnI-based): a pair of complementary primers, each carrying the mutation, anneal at the same site on opposite strands of a double-stranded plasmid template and are extended around the whole plasmid by a high-fidelity, non-strand-displacing polymerase, giving nicked circular mutant plasmids. The parental template, isolated from a dam⁺ E. coli strain, carries methylated adenine in its GATC sites; DpnI cuts GATC only when the adenine is methylated (Gm6ATC), so it destroys the parental DNA, while the strands synthesised in vitro are unmethylated and survive. The nicks are sealed after transformation into E. coli.

4. Cassette mutagenesis: the wild-type restriction fragment spanning the site of interest is excised using two flanking restriction sites and replaced by a chemically synthesised double-stranded oligonucleotide 'cassette' carrying the desired mutation(s), then ligated back into the vector.

Comparing the methods

Design and verification points

1. The mismatch is usually placed near the middle of the mutagenic primer, with enough correctly paired bases on each side for stable annealing.

2. PCR-based methods use a high-fidelity (proofreading) polymerase to limit unintended mutations elsewhere in the amplified DNA.

3. Every mutant clone is confirmed by sequencing the whole mutagenised region (or the whole insert), not just the target codon.

4. Alanine-scanning mutagenesis replaces residues one at a time with alanine to find which side chains a function depends on.

Core components and their roles

1. Guide RNA (sgRNA): a ~20-nt spacer complementary to the genomic target, fused to a scaffold that binds Cas9.

2. PAM (protospacer-adjacent motif): a short sequence immediately 3' of the protospacer on the non-target (PAM-containing) strand, recognised directly by the Cas9 protein, not by the guide RNA. For wild-type SpCas9 it is 5'-NGG-3' (N = any base); 5'-NAG-3' is recognised only weakly. Cleavage requires a compatible PAM, even when the guide matches the target perfectly.

3. Cas9 protein: an RNA-guided DNA endonuclease with two nuclease domains. HNH cleaves the target strand (the strand base-paired to the guide) and RuvC cleaves the non-target strand. Together they produce a blunt-ended double-strand break (DSB), typically 3 bp upstream (5') of the PAM.

Mechanism, in order

1. The sgRNA-Cas9 ribonucleoprotein complex scans DNA, pausing and locally unwinding wherever it finds a PAM.

2. If the adjacent sequence base-pairs with the guide spacer, starting from the PAM-proximal end, an R-loop forms and Cas9's nuclease domains are activated.

3. HNH and RuvC cut the target and non-target strands respectively, generating a blunt DSB.

4. The cell's own repair machinery resolves the break. NHEJ is error-prone and uses no template; it often introduces small indels that can disrupt the reading frame (the basis of gene knockout). HDR uses a donor template with homology arms and enables a precise correction or insertion, but it is largely restricted to S/G2 phase and is generally less efficient than NHEJ.

Cas variants and derived tools

Classification by bioreceptor and by transducer

General working sequence of a biosensor

1. The analyte in the sample diffuses to, and binds or reacts with, the immobilised bioreceptor.

2. The recognition event produces a physicochemical change (charge, mass, heat, light) at the transducer surface.

3. The transducer converts this change into an electrical signal.

4. The signal is amplified and processed by the associated electronics or software into a readable, usually quantitative, output.

Electrochemical transducers and the glucose biosensor

1. Amperometric: current measured at a fixed applied potential; the current is proportional to the rate of the electroactive reaction, and hence to analyte concentration within the linear range.

2. Potentiometric: potential measured at (near) zero current; it varies linearly with the logarithm of the ion activity (see formula).

3. Conductometric: change in solution conductance as the reaction produces or consumes ions.

4. Glucose sensor, first generation: glucose oxidase with measurement of O₂ consumed or H₂O₂ produced at the electrode (dependent on dissolved O₂).

5. Second generation: an artificial redox mediator shuttles electrons from the enzyme's reduced FAD to the electrode, reducing the O₂ dependence.

6. Third generation: direct electron transfer between the enzyme and the electrode, with no mediator.

Performance characteristics

1. Sensitivity: slope of the calibration curve (signal per unit concentration).

2. Selectivity: response to the analyte relative to interfering species.

3. Limit of detection (LOD): the lowest concentration distinguishable from the blank (see formula for a common convention).

4. Linear (dynamic) range, response time, stability (shelf and operational) and reproducibility.

EMSA (electrophoretic mobility shift assay, or gel-shift assay), in order

1. A short, labelled (radioactive or fluorescent) double-stranded DNA probe containing the putative binding site is incubated with the candidate protein (a purified protein, or a nuclear or cell extract).

2. The mixture is run on a native (non-denaturing) polyacrylamide gel alongside free probe alone.

3. Free DNA migrates fast. DNA bound by protein forms a larger complex that migrates more slowly and is seen as a retarded ('shifted') band.

4. Specificity is tested by competition: excess unlabelled probe of the same sequence (specific competitor) abolishes the shifted band, while an unrelated unlabelled DNA (non-specific competitor) does not.

5. Identity is tested with a specific antibody against the suspected protein. It either produces a further-retarded 'supershifted' band or, if it blocks the DNA-binding surface, abolishes the shifted band. Either result implicates that protein.

DNase I footprinting, in order

1. A DNA fragment labelled at one end only (on one strand) is incubated with and without the DNA-binding protein.

2. Both samples are treated briefly with a limiting amount of DNase I, so that each molecule is cut on average no more than once (single-hit conditions) at a random accessible position.

3. The products are run on a high-resolution denaturing sequencing gel alongside a sequencing ladder of the same fragment (e.g., a G+A ladder).

4. In the protein-bound lane, the region physically covered by the protein is protected from cleavage and appears as a gap (the 'footprint') in the otherwise continuous ladder of bands. This maps the protected nucleotides. Enhanced (hypersensitive) bands sometimes flank the footprint where binding distorts the DNA. Smaller probes such as hydroxyl radicals, or base-modifying agents such as dimethyl sulfate (DMS), give finer contact maps.

Chromatin immunoprecipitation (ChIP), in order

1. Intact cells are treated with a cross-linking agent (commonly formaldehyde) to covalently fix protein-DNA contacts in place.

2. Chromatin is sheared (by sonication or nuclease digestion) into small fragments, typically about 200-1000 bp.

3. An antibody specific to the protein of interest immunoprecipitates the protein together with its cross-linked DNA. A non-specific IgG (or no-antibody) sample and an 'input' sample of sheared chromatin taken before immunoprecipitation serve as controls.

4. Cross-links are reversed and the DNA is purified. The recovered sequences are identified by PCR (or qPCR), microarray (ChIP-chip) or sequencing (ChIP-seq), revealing where the protein binds in the native genome.

Other DNA-protein tools

1. Yeast one-hybrid: a DNA sequence (the 'bait') is placed upstream of a reporter gene in yeast, and a library of proteins fused to a transcriptional activation domain is screened. Reporter expression identifies proteins that bind the bait sequence.

2. Southwestern blot: proteins separated by SDS-PAGE and blotted onto a membrane are probed with a labelled DNA sequence.

3. DNA-affinity chromatography: a binding-site DNA immobilised on a matrix purifies the proteins that bind it.

4. Reporter-gene assays: the effect of a protein, or of mutating a binding site, on transcription from a promoter-reporter construct tests the functional consequence of binding.

Comparing the three core techniques

Yeast two-hybrid (Y2H) assay, in order

1. Protein X (the 'bait') is fused to the DNA-binding domain (DBD) of a transcription factor, commonly GAL4. Protein Y (the 'prey') is fused to the activation domain (AD) of the same transcription factor.

2. Both fusion constructs are co-expressed in a yeast reporter strain carrying reporter genes (e.g., HIS3, lacZ) downstream of the transcription factor's binding site (UAS).

3. If X and Y interact, the DBD and AD are brought together, reconstituting a functional transcriptional activator.

4. This switches on the reporter genes, giving a detectable phenotype: growth on medium lacking histidine, or blue colonies on X-gal.

Y2H limitations

1. A bait that activates transcription on its own (autoactivation) gives false positives and must be tested alone first.

2. The interaction must occur in the yeast nucleus. Membrane proteins, or proteins needing modifications that yeast does not make, can give false negatives (split-ubiquitin variants are used for membrane proteins).

3. Yeast proteins can bridge the bait and prey, so a positive does not by itself prove direct binding.

Co-immunoprecipitation (Co-IP), in order

1. Cells expressing proteins X and Y are lysed under gentle, non-denaturing conditions that preserve native complexes.

2. An antibody specific to protein X is added to the lysate and captured on protein A/G beads.

3. The bead-antibody-X complex is washed to remove unbound material.

4. If Y was bound to X, it co-precipitates and is detected by Western blotting. This shows that X and Y associate in a cellular context, but it does not distinguish direct binding from binding bridged by another protein.

GST/His pull-down assay, in order

1. One purified, tagged protein (e.g., GST-tagged X) is immobilised on an affinity matrix: glutathione beads for GST, or Ni-NTA for a His tag.

2. Purified protein Y (untagged or differently tagged) is added and allowed to bind. GST alone on beads is run in parallel as the control.

3. Unbound protein is washed away.

4. Bound Y is eluted and detected by Western blot or staining. With only purified components and no cell lysate, retention of Y by GST-X but not by GST alone indicates direct binding. Caveats: a contaminant co-purifying with either protein could still bridge them, and proteins expressed in E. coli lack eukaryotic post-translational modifications.

Other protein-protein tools

1. FRET (Förster resonance energy transfer): a donor fluorophore on X transfers energy non-radiatively to an acceptor on Y only when they are very close (roughly 1-10 nm) and the donor's emission spectrum overlaps the acceptor's absorption. Efficiency falls with the sixth power of distance (see formula).

2. BiFC (bimolecular fluorescence complementation): two non-fluorescent halves of a fluorescent protein are fused to X and Y. Interaction brings the halves together and restores fluorescence in living cells (the reconstitution is essentially irreversible).

3. SPR (surface plasmon resonance): one partner is immobilised on a gold sensor surface and the other flows over it. Binding changes the refractive index near the surface, giving a label-free, real-time signal from which association and dissociation rate constants, and hence $K_D$, are obtained.

4. Affinity purification-mass spectrometry (AP-MS), including tandem affinity purification (TAP) with two sequential purification steps under native conditions: identifies the members of a complex that co-purify with a tagged bait.

5. Chemical cross-linking followed by mass spectrometry: covalently links neighbouring proteins to capture transient or weak contacts.

Comparing the core techniques

Representative genomics-based approaches

1. Whole-genome sequencing (WGS): determines an organism's complete DNA sequence, either assembled de novo or by aligning reads to an existing reference genome, to identify variants and provide the coordinates on which other data are mapped.

2. RNA sequencing (RNA-seq): quantifies the transcriptome, revealing which genes are up- or down-regulated genome-wide after a mutation, treatment or CRISPR edit.

3. ChIP-seq: chromatin immunoprecipitation followed by next-generation sequencing, mapping genome-wide the sites a given DNA-binding protein occupies in vivo.

4. CRISPR-based functional screens: pooled libraries of guide RNAs knock out (or, with dCas9 fusions, repress or activate) genes across a cell population. A phenotypic readout (e.g., survival under a drug), followed by sequencing of the guides, identifies the genes that matter for that phenotype.

Representative proteomics-based approaches

1. Two-dimensional gel electrophoresis (2D-PAGE): separates a complex protein mixture first by isoelectric point (isoelectric focusing) and then by molecular mass (SDS-PAGE), resolving up to a few thousand proteins as spots for comparative expression analysis.

2. Mass spectrometry (MS): measures mass-to-charge ratio ($m/z$) of ionised molecules. MALDI (mostly singly charged ions) and ESI (multiply charged ions) are the common soft-ionisation methods. Proteins are identified by peptide mass fingerprinting (matching measured masses of digested peptides against a database) or by tandem MS (MS/MS) fragmentation, which reads peptide sequences, and post-translational modifications are located the same way. MS is the standard way to identify unknown partners recovered by Co-IP, pull-down or AP-MS.

3. Quantitative proteomics: stable-isotope metabolic labelling or isobaric chemical tags, or label-free comparison, measure relative protein amounts between samples.

4. Protein microarrays: thousands of purified proteins or antibodies spotted on a solid support to screen in parallel for binding partners, substrates or biomarkers.

5. Proteome-scale Y2H: one bait (or many baits) screened against an entire cDNA library to build a protein-protein interaction network (interactome).