Molecular cloning, mutagenesis and transfection

What is random mutagenesis and what does it result in?

introduction of random mutations throughout the DNA sequence without targeting specific sites

generate a broad range of mutations across the entire genome, resulting in a large pool of diverse mutations

How is random mutagenesis usually applied for research purposes?

• a stock of fruit flies is grown

• the flies are exposed to ionising radiation or mutagens

• flies are bred and they lay eggs

• surviving offspring is isolated

• visually identifiable mutations are assessed

• genomic DNA is isolated

• genes possessing a mutation are identifies

• genes of interest are sequenced

Why are fruit flies used for random mutagenesis experiments?

• only have 4 chromosomes

• their genome has a degree of similarity to humans

• easy to keep and breed

• small

What are the disadvantages of random mutagenesis experiments?

it is a lengthy, expensive and wasteful process

• some of the organisms won’t survive or reproduce

• mutations that are fatal for the offspring may be the most interesting ones

• there is a lot of redundancy is the genome (80% of DNA is non-coding, the 3rd position in each codon is silent)

What is site-directed mutagenesis?

targeted alteration of a single gene for research purposes

What are the advantages of site-directed mutagenesis?

• cheap and fast (commercially available kits)

• provides data to understand the functional properties of a protein (e.g. finding the binding site or active site)

• provides information on cell signalling pathways (e.g. post-translational modification sites)

What is required for site-directed mutagenesis?

• DNA template

• mutagenic primers (12-18 bases should anneal to the template on each side of the mutagenesis site)

• nucleotides (dNTPs)

• high fidelity DNA polymerase

• sequential thermal cycler (PCR machine)

• Dpn1 restriction enzyme (cleaves the methylated parental DNA)

• ultracompetent cells

What are the steps of a site-directed mutagenesis?

1) the reaction mix is made

2) double-stranded plasmid is separated

3) primer binds to the antisense of plasmid strand, DNA polymerase creates the new sense strand

4) the new plasmid is introduced into a competent bacterial cell

5) the plasmid is replicated by the bacterial cell (when the bacterium divides, one of the daughter cells will have a plasmid with mutation, the other will have the original plasmid)

6) bacterial cells containing the mutated plasmid are selected (e.g using antibiotic resistance)

What could be the reasons for negative results from a site-directed mutagenesis experiment?

• the mutation makes the protein non-functional

• the protein is not being expressed

• incorrect protein trafficking

• incorrect protein folding

How can you check if the protein from a mutated gene is being expressed?

Western blotting

How can you check if the protein from a mutated gene is being trafficked to the correct part of the cell?

tag it with GFP

What elements are usually present in a plasmid used for gene cloning?

• origin of replication

• a tag (gets attached to the protein to track it)

• multiple cloning site (for inserting the gene of interest)

• bacterial selection marker (e.g ampicillin resistance gene)

• mammalian selection marker (e.g. gentamicin resistance gene)

what are the steps to clone a gene into a plasmid?

1) adding sticky ends to the gene of interest that will match the sticking ends of the multiple cloning site (by designing PCR primers with sticky ends hanging over)

2) cut the plasmid with a restriction enzyme to create sticky ends

3) ligate the plasmid and PCR product

What is the criteria for choosing a restriction enzyme for cloning a gene into a plasmid?

• cuts the plasmid only once (in the multiple cloning site)

• doesn't cut the gene of interest

How can you make sure the gene of interest is cloned into a plasmid the right way around?

by using different restriction enzymes at each end

If using the same restriction enzyme for both ends of gene when cloning it into a plasmid, how can you test if the plasmid was inserted in the correct orientation?

Cut the plasmid with a restriction enzyme that has a restriction site within the gene of interest and somewhere else in the plasmid. The fragments will be of different sizes, depending on the orientation of the gene.

What could be the reasons for no bacterial colonies growing after cloning?

• plasmid not circularised (ligase not working)

• using the wrong antibiotic for selection

• bacteria not competent enough

What is transfection?

the insertion of a gene or group of genes into an organism for the purpose of expressing those genes

What are the two types of transfection?

• stable

• transient

What is stable transfection?

the DNA of interest is inserted into the parent genome of the cell

What is transient transfection?

genetic material of interest (DNA or RNA) is introduced into a cell for a short period of tike

What are the advantages and disadvantages of stable transfection?

+ the cells will always express the protein, they can be frozen and then thawed and used any time they are needed

- a complicated, time-consuming process that is often unsuccessful

What are the advantages and disadvantages of transient transfection?

+ cheap, simple, quick

+ genes can be easily switched off by inserting a mirror image of the mRNA

- inserted construct can be spontaneously be lost from the cell

- repeated cell division "dilutes the presence of the inserted genetic material

What are the different methods of transfection?

1) physical methods

• microinjection (direct injection of nucleic acid into the nucleus or cytoplasm)

• optical (creating temporary nanoscale pores in the cell membrane using a high-intensity, tightly focused laser beam)

• acoustic (ultrasound creates transient membrane pores)

• electroporation (short electrical pulses create temporary pores in the membrane)

• osmotic shock (cells are briefly exposed to hypotonic conditions, temporary membrane disruption allows DNA or RNA entry)

2) biological methods

• retroviral and lentiviral vectors

• adenoviral vectors

• adeno-associated virus

3) chemical methods

• calcium phosphate (cationic material)

• polymer-mediated (cationic material)

• lipid-mediated

4) magnetofection

What is the principal behind chemical transfection methods?

overcoming the repulsive force between negatively charged nucleic acids and the cell membrane

What are the advantages and disadvantages of transfection using cationic material?

+ low immunoreactivity

+ relatively cheap

+ theoretically don’t limit construct size

- high concentration of material required to create a concentration gradient

- high concentration may be toxic for the cells

What are the advantages and disadvantages of lipid-based transfection?

+ not toxic

- the mechanical/heat energy required to create liposomes can damage the genetic material

- if the liposomes are too big, they end up being degraded

How does magnetofection work?

• utilised positively-charged iron non-particles

• DNA/RNA binds to the nano-particles (this overcomes the negative charge of nucleic acids and concentrates the genetic material)

• the complex is taken up into the cell endosomally

What are the advantages of magnetofection?

• low toxicity

• allows selection of successfully-transfected cells with a magnet (suspension cells)

• allows to titrate the amount of DNA added (by changing the strength of the magnet)

What are the differences between lentiviral and adenoviral transfection?

lentiviral: stable transfection, acts on dividing and non-dividing cell, the main safety concern is risk of oncogene insertion

adenoviral: transient transfection, acts mainly on dividing cells, the main safety concern is immunoreactivity

What are the advantages of viral transfection methods?

• hijacking pre-made biological machinery

• very high transfection efficacy

• suitable for DNA and RNS

• easy to scale up

What are the disadvantages of viral transfection methods?

• size on construct is limited by virus size

• risk of random insertion

• potential immunoreactivity

• too much virus can kill the cell

Why are chemical methods not an option for transfection is whole animals?

toxic for cells

What are the issues with lentiviral transfection in whole animals?

• can trigger an immune response

• potential for insertion of oncogenic genes (as derived from HIV-1)

• potential unintentional insertion of genetic material

• production of lentiviral vectors involves multiple plasmids, which could complicated their use as gene therapy

• need for adjuvants to activate the virus

• variable efficiency (different cell types show different susceptibility to transfection)

• high viral load can damage the cells

• expensive as a lot of DNA and virus needed

How can transfection efficiency be measured?

by know much of the gene product is made

What is the issue with using electrophoresis or western blotting to measure transfection efficiency?

destroys the cells

How can transfection efficiency be measured without destroying the cells?

by added a reporter gene (e.g. GFP) after the GOI to confirm complete transcription

How can transfected cells be selected?

• using an antibiotic degradation gene

• fluorescence-activated sorting based on a fluorescent marker