Practical Exam Molecular

P1 — What is the purpose of H&E staining?

To make tissue structures visible so stained non-diseased and diseased intestinal tissue can be examined and compared.

P1 — What does haematoxylin stain?

Acidic structures, particularly the nucleus (DNA) and RNA-rich cytoplasmic regions such as ribosomes and rough ER; they appear purplish-blue.

P1 — What does eosin stain?

Basic structures such as cytoplasm; these appear red/pink.

P1 — Why do tissue sections need staining?

Most cells are colourless and transparent, so staining makes cellular and tissue structures visible.

P1 — What is H&E?

Haematoxylin and eosin, a widely used histological stain for examining tissue structure.

P1 — What is the purpose of the H&E practical?

To stain non-diseased mouse intestinal tissue and compare tissue morphology with diseased intestinal tissue.

P1 — Why is tissue fixed before sectioning/staining?

The manual states tissue is fixed in paraformaldehyde before being embedded in paraffin and sectioned.

P1 — What is DAB staining used for?

To visualise the location of a target protein in tissue through immunohistochemical staining.

P1 — What does IHC stand for?

Immunohistochemistry.

P1 — What does the primary antibody do in IHC?

It specifically binds the target protein/antigen of interest.

P1 — What does the secondary antibody do in the DAB/IHC method?

It binds the primary antibody and is linked to an enzyme that enables detection.

P1 — What enzyme is involved in the DAB detection system?

Horseradish peroxidase (HRP).

P1 — What produces the brown DAB signal?

HRP catalyses the oxidation of DAB in the presence of hydrogen peroxide, producing an insoluble brown precipitate.

P1 — What does a brown DAB signal indicate?

The location where the target antigen/protein has been detected.

P1 — H&E vs IHC/DAB: what is the key difference?

H&E broadly shows tissue/cellular morphology, while IHC/DAB selectively detects a particular target protein.

P1 — What was the disease-tissue analysis task?

To analyse pre-stained tissue subtypes of diseased intestinal tissue and compare them with non-diseased tissue.

P2 — What were the two DNA sources compared?

Buccal cells from the cheek and epithelial cells from the palm.

P2 — What are buccal cells?

Squamous epithelial cells lining the inside of the mouth.

P2 — Why can palm epithelial cells be useful for DNA analysis?

They can provide low-level 'touch DNA' deposited by skin cells.

P2 — What is the purpose of DNA isolation?

To obtain genomic DNA suitable for downstream analysis such as NanoDrop measurement and PCR.

P2 — What is Buccalyse used for?

Rapid preparation/release of DNA from human buccal swab samples for PCR assays.

P2 — Buccal collection: approximately how many times was the swab rolled on each cheek?

Approximately 10 times on each side.

P2 — After placing the buccal swab into Buccalyse, how long was it vortexed?

10 seconds.

P2 — Buccal DNA: first heat-block condition?

70°C for 15 minutes.

P2 — Buccal DNA: second heat-block condition?

95°C for 2 minutes.

P2 — After the 95°C incubation, what was done to the DNA sample?

It was vortexed briefly and stored on ice.

P2 — What is DNAzol?

A guanidine-based reagent used for DNA extraction/purification from epithelial cells.

P2 — What does a NanoDrop measure?

Absorbance across the UV-visible spectrum to assess nucleic-acid concentration and purity.

P2 — At what wavelength does DNA absorb maximally according to the manual?

260 nm.

P2 — At what wavelengths do proteins absorb UV according to the manual?

230 nm and 280 nm, with 230 nm described as the maximum.

P2 — What does A260 measure in the NanoDrop context?

Absorbance associated with nucleic acids and used to estimate DNA concentration.

P2 — What A260/A280 ratio indicates highly purified DNA?

Approximately 1.8–2.1.

P2 — What is the DNA concentration relationship given in the manual?

An A260 of 1.0 is approximately equivalent to 50 ng/mL double-stranded DNA.

P2 — Why is a blank used on the NanoDrop?

To calibrate the instrument using the appropriate blank solution before measuring the DNA sample.

P2 — What is the purpose of PCR?

To amplify a specific DNA sequence.

P2 — What are the three repeating stages of PCR?

Denaturation, annealing and extension.

P2 — What happens during PCR denaturation?

The DNA strands separate.

P2 — What happens during PCR annealing?

Primers bind to complementary sequences on the template DNA.

P2 — What happens during PCR extension?

DNA polymerase extends the primers by adding nucleotides to form new DNA strands.

P2 — What enzyme is used in PCR?

Taq DNA polymerase.

P2 — What is the purpose of PCR primers?

They define the DNA region to be amplified and provide starting points for DNA synthesis.

P2 — What is the PCR control mentioned in the manual?

A control containing no DNA, provided by technical staff.

P2 — What formula is provided for dilution calculations?

Cs × Vs = Cf × Vf.

P2 — What do Cs, Vs, Cf and Vf mean?

Cs = stock concentration; Vs = stock volume; Cf = final concentration; Vf = final volume.

P3 — What is recombinant DNA technology doing in this practical?

The PCR product is inserted into a plasmid vector, introduced into bacteria, and then screened for recombinant colonies.

P3 — What is a plasmid vector?

A DNA molecule used as a vehicle to carry the inserted DNA into a host cell.

P3 — What is gene ligation?

Joining DNA fragments to form a recombinant DNA molecule.

P3 — What is transformation?

Introducing recombinant DNA/plasmid DNA into bacterial cells.

P3 — What is TOPO cloning?

A cloning method that uses DNA topoisomerase I activity and the A-overhang of PCR products to insert DNA into a compatible vector.

P3 — What does Taq polymerase add to PCR products for TOPO cloning?

A single deoxyadenosine (A) to the 3′ end.

P3 — What does the TOPO vector provide for the PCR product?

Compatible thymidine-containing ends that allow the A-tailed PCR product to bind and form the recombinant plasmid.

P3 — What is directional cloning?

A cloning approach where the insert and vector are cut with restriction enzymes to create specific, non-compatible ends that control insert orientation.

P3 — Why can directional cloning be limited?

It requires suitable restriction sites in the DNA sequence/vector.

P3 — Why is antibiotic selection used after bacterial transformation?

Only a small fraction of bacteria take up plasmid DNA, so antibiotic resistance allows transformed cells to be selected.

P3 — What is agarose gel electrophoresis used for in this practical?

To separate PCR products and recombinant plasmid DNA according to size and visualise them.

P3 — In DNA gel electrophoresis, which direction does DNA migrate?

Toward the positive electrode because DNA is negatively charged.

P3 — Which DNA fragments migrate further through agarose?

Smaller fragments.

P3 — What is a DNA ladder used for?

To provide fragments of known sizes so the size of sample DNA bands can be estimated.

P3 — What is blue-white screening used for?

To distinguish bacterial colonies containing recombinant plasmids from colonies containing non-recombinant plasmids.

P3 — What does a blue colony indicate?

Non-recombinant plasmid without the DNA insert.

P3 — What does a white colony indicate?

Recombinant plasmid containing the DNA insert.

P3 — Why are recombinant colonies white?

The inserted DNA disrupts the relevant lacZ function, so X-gal is not broken down to produce the blue colour.

P3 — Why are non-recombinant colonies blue?

The intact lacZ function allows X-gal to be broken down, producing the blue colour.

P3 — How long was the bacterial transformation step before plating?

45 minutes.

P3 — How much transformed bacterial culture was plated?

100 μL.

P3 — At what temperature were plates incubated?

37°C overnight.

P3 — Why were plates incubated inverted?

To prevent condensation from dripping onto colonies and interfering with the results.

P3 — What was determined from the bacterial plates?

Colony counts and transformation efficiency.

P4 — What is recombinant protein expression?

Producing a desired protein from a gene inserted into an expression vector and introduced into a suitable host.

P4 — What happens to recombinant DNA inside the host?

It is transcribed and translated using the host's cellular machinery to produce the desired protein.

P4 — What are examples of expression/purification tags mentioned?

His-tag and GST-tag.

P4 — Why purify recombinant proteins?

To obtain the target protein away from other host-cell components, ideally as a highly pure and biologically active product.

P4 — What purification methods are mentioned?

Affinity chromatography, ion-exchange chromatography and size-exclusion chromatography.

P4 — What is RIPA buffer used for?

Protein extraction/lysis of cells so proteins are released into solution.

P4 — What happens during freeze-thaw lysis?

Cells are alternated between freezing and warming to disrupt the cells and release proteins.

P4 — What were the freeze-thaw conditions for the protein extracts?

Dry ice for 2 minutes followed by a 37°C heat block for 6 minutes, repeated three times.

P4 — After cell lysis, why is the sample centrifuged?

To separate cellular debris from the protein-containing supernatant.

P4 — What were the centrifugation conditions?

10,000 rpm for 30 minutes at 4°C.

P4 — Which fraction contains the proteins after centrifugation?

The supernatant.

P4 — At what temperature were protein extracts stored for later use?

−80°C.

P4 — What is the purpose of the Qubit assay?

To determine protein concentration using a fluorescence-based assay.

P4 — How much protein sample was added to each Qubit tube?

20 μL.

P4 — How much fluorescence assay buffer was added?

150 μL.

P4 — How much fluorescence assay reagent was added?

30 μL.

P4 — What was the final Qubit tube volume?

200 μL.

P4 — How long were Qubit samples incubated?

10 minutes at room temperature.

P4 — What was used as the Qubit blank?

RIPA buffer in place of the protein sample.

P4 — Does the faint yellow colour in the Qubit assay indicate protein concentration?

No. The manual specifically states that the yellow colour does not indicate protein concentration.

P4 — What is a dot blot?

A protein detection method where samples are spotted directly onto a membrane rather than separated by electrophoresis.

P4 — How does dot blot differ from Western blot?

Dot blot does not separate proteins by electrophoresis before antibody detection.

P4 — What is the purpose of blocking in a dot blot?

To reduce non-specific antibody binding to the membrane.

P4 — What blocking buffer was specified?

5% non-fat milk or 2% BSA in TBST.

P5 — What is the purpose of SDS-PAGE?

To separate proteins primarily according to molecular size/weight after denaturation.

P5 — What does SDS do to proteins?

It denatures proteins and gives them an overall net negative charge.

P5 — What does a reducing agent such as 2-β-mercaptoethanol do?

It disrupts disulfide bonds within proteins.

P5 — Why can SDS-PAGE separate proteins by size?

SDS gives proteins a relatively uniform negative charge, so migration through the polyacrylamide gel is primarily related to molecular size.

P5 — Which proteins migrate furthest in SDS-PAGE?

Smaller proteins.

P5 — What is a protein molecular-weight standard/ladder used for?

To estimate the molecular weight of sample proteins by comparison with proteins of known sizes.

P5 — What were the SDS-PAGE sample preparation conditions?

Mix 3 parts sample with 1 part 4× SDS sample buffer, then heat at 95°C for 5 minutes.

P5 — What were the SDS-PAGE run conditions?

200 V for 39 minutes, or until the dye front leaves the gel.