DNA Molecular Flashcards
Describe transformation of bacterial cells, including state and techniques
In order for a bacterial cell to be capable of being transformed, it must be competent (1). This can be achieved artificially via chemical transformation or electroporation (1).
Chemical transformation involves cells being placed in ice cold transformation solution which contains divalent cations e.g. CaCl2 (1). DNA is added to the cells and the mixture is subjected at a heat shock treatment, involving heating to 42 °C before returning to ice (1). Cells are then allowed to recover before being plated on selective media.
Electroporation involves cells being subjected to a series of high voltage electrical impulses (1). DNA is mixed with the cells, and the electric shock increases permeability of the cell membrane allowing DNA uptake (1). Cells are then allowed to recover before being plated on selective media.
Describe methods of selecting transformed bacteria
If the bacteria have been transformed with a plasmid containing an antibiotic resistance gene, cells can be plated on selective media containing that particular antibiotic (1). This will allow only cells transformed with the plasmid to grow (1).
Alternatively, selection can be performed using blue/white screening. Insertion of DNA into the LacZ gene causes inactivation, meaning X-gal cannot be converted to a blue product, so colonies remain white (1). Cells which contain a plasmid without the DNA insert will have a functioning LacZ gene, so X-gal can be converted to a blue product when induced by IPTG, producing blue colonies (1).
Cloning vectors; features and advantages
Plasmids, bacteriophage, cosmids, BACS, YACS (1).
Advantages include being able to insert larger pieces of DNA (1) and having an increased transformation efficiency (1).
Features:
Presence of ori
Multiple cloning site
Presence of a selectable marker
High transformation efficiency
Easy to purify
High copy number in host cell
Inducible promoter and example
An inducible promoter can be switched on or off to control gene expression (1).
In the lac operon, the absence of lactose (the inducer) results in the repressor protein, produced by the repressor gene, being able to being to the operator which blocks RNA polymerase access to the promoter and preventing gene transcription (1).
When lactose is present, it binds to the repressor protein preventing it from binding the operator. This allows RNA polymerase to bind the promoter, resulting in transcription of structural genes (1).
Dideoxy chain termination for fragment sequencing
This method involves the use of dideoxyribonucleoside triphosphates (ddNTPs), which are nucleotides that have been modified to have their 3’-OH group removed (1).
Each of the four ddNTPs bases are labelled with a different coloured fluorophore (1).
The DNA to be sequenced is incubated with the reagents required for DNA synthesis and new strands are synthesised by elongating specific primer (1).
A ddNTP is incorporated at random which terminates the elongation process, generating DNA strands of different sizes (1).
The DNA strands created can be separated according to size using capillary gel electrophoresis (1).
During the capillary electrophoresis, the fluorophore passes through a detector and the fluorescent signal produced from the ddNTP indicates the nucleotide at the end of the strand (1).
These results can be printed out as an electropherogram, allowing sequence complementary to the template to be read (1).
Human genome sequencing stages
Linkage Mapping (1): a map of genes, showing the relative position of alleles, created based on recombination frequencies (1).
Physical Mapping (1): restriction fragments are generated, and genetic markers are used to identify overlapping fragments, giving a sequential order which corresponds to the order on the chromosome (1).
DNA Sequencing (1): nucleotide sequence of small DNA fragments are determined and assembled into the complete genome sequence (1).
BAC-To-BAC method
Chromosomes to be sequenced are cut into large fragments of approximately 150,000 bp and inserted into bacterial artificial chromosomes (BACs) (1).
Fragments in BAC are fingerprinted to create a physical chromosome map (1).
Fragments are then cut into even smaller pieces using restriction digests and cloned into M13 phage vectors which can be sequenced to identify the chromosome DNA sequence (1).
Whole genome sequencing
It is much quicker since:
* There is not a need to produce linkage maps and no need for BAC libraries to produce physical maps of chromosomes (1).
* Genomic DNA is randomly broken into many small pieces of about 2000 bp and inserted directly into cloning vectors for sequencing (1).
It differs from hierarchical shotgun sequencing as it starts directly with DNA sequencing of restriction fragments from randomly cut genomic DNA, skipping the need for linkage and physical maps.
DNA microarrays
Features:
Thousands of spots in ordered rows and columns on a chip (1).
Each spot contains multiple identical strands of DNA (1).
Each spot represents one gene (1).
Precise locations of each gene/spot is recorded in a database (1).
Hybridisation:
Microarrays involve hybridisation between two DNA strands: the oligonucleotide probe and the sample cDNA (1).
Sequences with high levels of complementarity will bind tightly and remain hybridised after washing (1).
cDNA:
Organisms entire genome is sequenced (1).
DNA fragments containing genes are amplified by PCR (1) using specifically designed primer sets (1).
The resulting double stranded cDNA is denatured and single stranded cDNA is produced (1).
The single stranded DNA copies in solution are spotted onto recorded ‘well’ on a microarray (1).
DNA microarray application
Used in cancer research to compare expression profiles of cancerous and non-cancerous cells (1).
Identifying genes that are up/down regulated in samples can provide insights into genes involved in the development of cancers and can be used as a tool for cancer diagnosis (1).
Process of microinjection
Use of glass micropipette to inject DNA in solution at a microscopic level (1).
For pronuclear injection the target cell is positioned under the microscope and two micromanipulators—one holding the pipette and one holding a microcapillary needle usually between 0.5 and 5 µm in diameter (larger if injecting stem cells into an embryo)—are used to penetrate the cell membrane and/or the nuclear envelope (1).
In this way the process can be used to introduce a vector containing transgene into a single cell (1).
For pronuclear injection to be successful, the genetic material (typically linear DNA) must be injected while the genetic material from the oocyte and sperm are separate (i.e., the pronuclear phase) (1).
Process of chromosomal recombination allows some transgene DNA to be integrated into the host egg chromosomes (1).
The oocyte is then implanted in the oviduct of a pseudo pregnant animal (1).
Transgenic mice can then be selected and bred to create transgenic lines (1).
Stages of stem cell manipulation to make a transgenic mouse
Microinject recombinant DNA (containing the desired gene, driven by high affinity promoter) into embryonic stem cells (1).
Select transformed embryonic stem cells via genetic marker and microinject into mouse blastocyst (1).
Implant transgenic blastocyst into a foster mother mouse (1).
Test offspring for gene – heterozygous founder line (1).
Selectively breed founder line to produce mice that are homozygous for desired gene (1).
How would a retrovirus produce a transgenic animal?
Retrovirus containing transgene used to infect host cells of choice (1).
Transmission of the transgene to offspring is possible only if the retrovirus integrates into germ cells (1).
Retrovirus facilitates integration of the transgene into the host genome (1).
Offspring derived from this method are chimeric, i.e. not all cells carry the retrovirus (1).
Benefit of transgenic animal example
Requirement of aquaculture due to exhausting natural fisheries. Introduction of an alternative growth-hormone regulating gene to Atlantic salmon allows them to grow all year round giving improved size/yield compared to traditionally farmed salmon.
How could a gene of interest be removed from one plasmid and put into another?
First, the gene of interest would be removed using a restriction enzyme (1). These cut unmethylated dsDNA at specific sequences called restriction sites (1).
The gene would then be purified e.g. by using affinity chromatography (1) and then amplified by PCR (1).
The new plasmid would be cut using the same restriction enzyme that was used to isolate the gene (1).
The amplified gene can then be mixed with the cut plasmid (1) and DNA ligase is used to join the DNA fragments together (i.e. gene inserted into new plasmid) (1). DNA ligase works by creating phosphodiester bonds between DNA molecules when there is a free 5’ phosphate residue (1).
