Characterising and cloning genes

Question 1

Goal of gene isolation?

Answer 1

Study, modify or transfer a particular gene or fragment of DNA.

In order to do this, need to identify and replicate the DNA at high copy number in a host cell or in vitro.

Question 2

Problems with gene isolation?

Answer 2

DNA must contain a replication origin site in order to be replicated in a host.

Organisms contain many genes, so how to identify the correct gene?

Question 3

Solutions for gene isolation?

Answer 3

Plasmids and bacteriophages are replicons and so can be used as vectors.

Replicate a single DNA fragment by generating a library of many cells/clones, each containing a different fragment.

PCR can be used to replicate DNA at high copy number in vitro.

Question 4

How does gene cloning happen?

Answer 4

DNA fragment joined with vector to construct a recombinant DNA molecule.

The DNA fragment is cut out by using restriction enzymes from phage resistant bacteria.

Generally a palindromic recognition site. Can generate sticky ends.

Joining cut DNA – Two steps of hydrogen bonding between complementary bases if the fragment had sticky ends.

Then phosphodiester bonds are catalysed by DNA ligase via an ATP-dependent reaction.

Gel electrophoresis – method for separating DNA fragments based on size. DNA fragments migrate towards the anode, smaller fragments move faster. Can label fragments fluorescently

Question 5

What is a genomic library?

Answer 5

A genomic library is a collection of the total genomic DNA from a single organism.

The DNA is stored in a population of identical vectors, each containing a different insert of DNA.

In order to construct a genomic library, the organism’s DNA is extracted from cells and then digested with a restriction enzyme to cut the DNA into fragments of a specific size.

The fragments are then inserted into the vector using DNA ligase.

Next, the vector DNA can be taken up by a host organism - commonly a population of Escherichia coli or yeast - with each cell containing only one vector molecule.

Using a host cell to carry the vector allows for easy amplification and retrieval of specific clones from the library for analysis.

Question 6

When is a cDNA library used?

Answer 6

When only interested in mRNA sequences produced in a cell

Fragmentation isn’t required.

Question 7

How does mRNA get purified for cDNA synthesis?

Answer 7

1. A special column contains short oligo(T) chains linked to cellulose
2. mRNAs have poly(A) tails so the total cellular RNA is isolated from cells and passed through columns
3. The poly(A) tails of mRNA pair with oligo(T) chains and retained in the column
4. mRNA washed from the column by adding a buffer which breaks the H bonds between the poly(A) tails and the oligo(T) chains

Question 8

How does cDNA get synthesised from mRNA?

Answer 8

1. mRNA acts as the template
2. Add and anneal the oligo-dT primer to the poly(A) tail
3. Add dNTPs and reverse transcriptase
4. Generates cDNA
5. RNAase generates nicks in RNA
6. Second strand synthesis begins in the nicks, and has exonuclease activity. The RNA fragments act as primers for DNA polymerase
7. DNA ligase used to close the gaps

Question 9

How do you identify the clone (DNA fragment) of interest in the library?

Answer 9

Requires a way of replicating the plate of clones and a labelled DNA fragment to screen the replica with (a probe)

1. Replicating library plates:

Overlay a nitrocellulose disk to make a replica of the master plate containing genomic library of clones inside E. coli cells.

Remove disk from plate and lyse cells on it and denature DNA. Bake it and treat it with UV light to bind DNA strand to disk.

The replica is needed to have something easy to work with during screening, and to enable preservation of the master plate in viable form for future propagation

2. Labelling a DNA fragment:
   Denature and anneal in presence of mixture of random primers

Add DNA polymerase and dNTPs with some labelled nucleotides. The newly synthesised DNA has incorporated the labelled nucleotide and so is labelled and can be used as a probe (once single stranded)

3. Screening by DNA hybridisation:

Identify the clone of interest by adding the radioactive probe to the replica plate and exposing to X-Rays

4. Polymerase Chain Reaction

Question 10

How is cloned DNA isolated?

Answer 10

Chromosomal DNA can be transformed to single-stranded linear DNA with an ionic detergent and NaOH in pH 12.

Then in pH 7, this becomes a tangled mass of linear DNA, and this can be centrifuged to separate from the chromosomal plasmids.

The linear DNA molecules will become a pellet at the bottom of the centrifuge.

Question 11

How is DNA sequenced?

Answer 11

Sanger sequencing method:

Template to be sequenced gets a primer added, then dNTPs and ddNTPs (each with different fluorescent tags) and DNA Polymerase.

This results in sequences of different lengths. The ddNTPs lack 3’ hydroxyl so block chain extension.

Gel electrophoresis separates them due to length, and the fluorescent tags show which ddNTP was added at the end.

Automated sequencers use capillary electrophoresis and laser detection of products.

Comparisons of cDNA and genomic clone sequences enables positions of promoters, introns and exons to be determined.

Question 12

What is southern blot analysis?

Answer 12

Method for determining the number of sites in a genome that have similarity to DNA of interest

1. Genomic DNA cut with restriction enzyme
2. DNA fragments separated by gel electrophoresis
3. Separated DNA fragments blotted onto nitrocellulose paper after denaturation
4. Remove nitrocellulose paper with tightly bound DNA
5. Labelled DNA probe hybridised to the separated DNA on the nitrocellulose

Question 13

How do you determine the chromosomal localisation of gene copies?

Answer 13

1. Drop cells onto a glass slide. Typically use metaphase cells to enable visualisation of individual condensed chromosomes.
2. Sample is fixed and permeabilised, and DNA is denatured.
3. Add hybridisation probes labelled with fluorescent dye. Radioactive probe may not provide sufficient resolution, or may damage the chromosomes.
4. Visualise by fluorescence microscopy. Can see both sister chromatids.

Question 14

How do you analyse gene expression patterns?

Answer 14

Where and when is a gene expressed? (Western blotting but with RNA)

1. Purify RNA from different tissues
2. Load RNA samples into wells of a gel
3. Separate RNA samples by gel electrophoresis. Blot onto filter. Expose filter to labelled hybridisation probe

4 Wash away any un-hybridised probe. Make autoradiograph.
Provides information on mRNA levels in different tissue samples and on mRNA size

Which cells accumulate the mRNA of interest? (in situ hybridisation)

1. Make a single stranded probe – the antisense RNA probe will hybridise to sense mRNA in sample
2. Detection of probe using enzyme. Forms insoluble purple precipitate at sites of alkaline phosphatase activity (where the probe occurs). Can be visualised with a microscope
3. In situ hybridisation. Each cell with purple stain contains mRNA of interest

Question 15

How do you analyse where proteins accumulate?

Answer 15

An antibody against the protein of interest is needed. Using an antibody to detect protein.

1. Polyacrylamide gel provides better resolution than agarose gel.
2. Run replica gel and stain all proteins
3. Blot proteins to membrane
4. Probe blot with primary then secondary antibodies

Question 16

How do you use a cloned gene to synthesise a protein?

Answer 16

Proteins encoded by cloned genes can be expressed in bacteria, and then used to raise antibodies

Question 17

What are chromosome maps?

Answer 17

Different types of map with different levels of resolution. Sometimes the lower resolution maps are needed to generate the higher resolution maps.

Karyotypic map: From microscopic observation of chromosomal spreads

Linkage map: Derived from monitoring recombination between markers, in cM

Physical map: Measured in bp

Sequence map: Sequence of all bases along the chromosome

Question 18

What does 1cM mean?

Answer 18

1% recombination in a single generation

Question 19

Describe the original human genome project?

Answer 19

Why, How and Funding? Some thought that sequencing the introns was unnecessary, why bother sequencing the “junk”.

Completed in 2003, work officially began in 1990.

Identity of those sequenced not known in order to protect those involved. Both male and female donors were use. 70% of sequence came from one person.

Sequenced using Sanger technology.

Question 20

Describe hierarchical shotgun genome sequencing?

Answer 20

Approach employed by the public project.

Split into several sequences.

These then get all split up. Make sure the fragments are small enough for sanger sequencing.

Sequence each fragment using sanger.

As the DNA has been broken down many different ways, the sequences can be analysed to find overlapping regions and put it back in order.

Question 21

Describe whole genome shotgun sequencing?

Answer 21

Used by private company.

Aim to sequence the genome more quickly, control access and patent genes.

Fragment entire genome and clone pieces directly into plasmid vector.

Sequence plasmid clones at random, meaning the assembly is computationally intensive, but could use info from the public project.

Question 22

Types of Next Gen Sequencing?

Answer 22

454 pyrosequencing

Illumina sequencing

Question 23

Describe 454 pyrosequencing?

Answer 23

Makes use of the pyrophosphate (PPi) released upon nucleotide incorporation.

PPi is used to fuel a downstream set of reactions that ultimately produces light by the action of luciferase on luciferin.

Amplification is by PCR not vector based. Emulsion PCR has everything necessary for PCR in a droplet. Form an emulsion with each water droplet carrying a single bead, each droplet functions as a discrete micro-reactor, eliminating cross talk during PCR.

PCR amplifies the unique sequence on the surface of each bead, release the bead and add them to a sequencing plate.

Pyrosequencing involves smaller enzyme beads being added to each well, surrounding the DNA carrying beads. Sequencing primer, DNA polymerase and two substrates are added.

Different dNTPs are added sequentially to the wells in repeated cycles. The incorporation of a nucleotide results in light emission, intensity of which is recorded.

Question 24

Describe illumina sequencing?

Answer 24

Reversible terminator sequencing, similar in principle to Sanger sequencing, but the terminator can be removed.

Bridge amplification. Library fragments are bound to the flow cell by hybridisation to the oligos (which are attached to the flow cell).

PCR bridge amplification using only the bound oligos constrains the distribution of the products, producing clusters comprising numerous identical fragments.

Sequencing occurs with the clusters being supplied with polymerase and all four nucleotides (all tagged with a different fluorescence).

As the nucleotides have their 3’-OH chemically blocked, only one is incorporated per cycle.

After each cycle, the cell is imaged to identify the new nucleotide incorporated at each cluster.

Then a chemical step removes the fluorescent tag and the 3’ block.

Question 25

What are third generation technologies?

Answer 25

They can sequence single molecules.

PacBio is a single molecule real time sequencing.

Question 26

Why is resequencing faster?

Answer 26

Can be aligned to the previously derived sequence. Can be used to detect polymorphisms and mutations.

Question 27

What is bioinformatics?

Answer 27

Annotation – dependent on researchers for entry so can contain errors. Contains origins, background information, important regions of the sequence and links to protein sequence.

How to annotate?
Identify Open Reading Frames (ORFs - part of a reading frame that has the ability to be translated).

Search databases using ORFs as queries, may identify related genes, conserved functional domains, protein targeting sequences, etc.

Find related genes using BLAST, nucleotide query sequence searched against a nucleotide database.

Question 28

Stable vs transient transformation?

Answer 28

Transient
Introduced DNA is not integrated into host genome and so is diluted with every host cell cycle. Suitable for short term expression or use with non-rapidly dividing cells.

Stable
Introduced DNA is integrated into the host genome, and so is replicated as cells divide. Often non-targeted, but in some species targeted integration is possible via homologous recombination.

Question 29

Methods of gene transfer?

Answer 29

Calcium phosphate precipitation.

Electroporation.

Microinjection.

Micro-projectile bombardment.

Viral infection of mammalian cells.

Agrobacterium infection of plants.

Genome editing.

Question 30

Outline calcium phosphate precipitation?

Answer 30

Plasmid DNA precipitates with CaPO4, and when the DNA is added to cells.

The precipitate binds to cell surface when added to mammalian culture cells and is taken up by endocytosis.

After waiting for 4-16 hours at 37 degrees, fresh medium is added to the cells and the DNA solution is removed.

Quick, cheap & simple, not vector dependent and can assay transient expression or score for stable integration. Essentially only used for mammalian cell lines.

Amount incorporated into nucleus is low (1-2%).

Question 31

Outline electroporation?

Answer 31

Electric field causes polarisation of cell membrane.

Pores form in the membrane (pore channels) allowing DNA to enter.

The membrane heals with the gene inside.

Quick and not vector dependent. Used for bacteria, yeast, plant protoplasts and mammalian cells.

Can assay transient expression or score for stable integration.

Equipment is quite expensive and can only be used for single cells.

Question 32

Outline microinjection?

Answer 32

Can be used to transform various organisms including nematodes, insects, fish, amphibians and mammals.

Inject gene into the male pronucleus.

Frequently used to make transgenic mice.

Not vector dependent and allows stable integration, but usually random.

Expensive, requires skill and very labour intensive.

Question 33

Outline micro-projectile bombardment?

Answer 33

Particularly useful for plant transformation as plant cells have a tough cell wall.

Tungsten or gold particles coated with DNA are fired into tissue using compressed gas as the propellant, under vacuum.

Quick, can in principle be used on any tissue. Not vector dependent.

Can assay transient expression or score for integration, and can deliver DNA to organelles, usually chloroplasts.

Equipment is expensive and is best suited for use with robust cells.

Question 34

Outline viral infection of mammalian cells?

Answer 34

Various mammalian viruses can be adapted for use as vectors (gene delivery vehicles).

Such vectors are typically replication defective, with genes responsible for replication removed.

Can be used to deliver DNA to cells or intact organisms (in gene therapy).

Can use adenovirus which doesn’t integrate into host genome, allows short/medium term expression.

Effective delivery of DNA to cells, and offers high levels of transgene expression (adenovirus) as well as long term, stable integration (lentivirus).

Vector dependent, but integrating vectors may activate cellular oncogenes.

Question 35

Outine agrobacterium infection of plants?

Answer 35

The most common method for transforming plants uses the parasitic bacterium.

Bacteria infects the plant tissue and transfers the T-DNA region of Ti into the plant genome.

T-DNA encodes genes for hormones to promote cell proliferation.

The T-DNA can be disarmed and adapted for use as a vector.

Widely and routinely used, many vector options available. Enables stable integration.

Limited host range, but can be time consuming and site of integration in genome is random.

Question 36

Outline genome editing?

Answer 36

New technologies have emerged that enable the specific editing of a gene of interest.

These employ a sequence-specific guiding mechanism to target a nuclease to a gene of interest.

Once in situ, the nuclease creates a double stranded break within the target gene, with different outcomes.

Question 37

How can you manipulate gene expression?

Answer 37

When transferring DNA, you could repress the endogenous gene expression by RNA interference (need cDNA sequence of endogenous gene), over express an endogenous gene with a constitutive (means that the enzyme is continuously produced in an organism regardless of the cell needs) promoter (need cDNA sequence).

Express a novel gene with a constitutive promoter (need sequence of novel gene) or express a gene in a defined spatial/temporal context (need cDNA sequence of relevant gene and a suitable promoter).

Promoters for constitutive gene expression – choice depends on system. Mammalian cells often use actin.

RNAi mediated suppression of gene expression – native function of RNAi is an antiviral defence mechanism in eukaryotes, recognises dsDNA. siRNAs target RISC complex to the mRNA which it then cleaves. System can be exploited to selectively silence a gene of interest.

Question 38

Applications of gene manipulation in agriculture?

Answer 38

Polygalacturonase RNAi in tomatoes:
an enzyme that degrades the plant cell wall component pectin.
Activity of the enzyme causes tomato fruit softening, leading to the habit of harvesting fruit prematurely.
However, these aren’t as flavourful as those ripened on the vine.
Using RNAi, led to a much higher level of pectin.

Insect resistant cotton express Bt toxin:
toxin derived from soil bacterium Bacillus thuringiensis - affects insect digestive system specifically.
Products prepared from this bacterium have been used as an insecticide since the 1920s, and are used in organic farming.
Gene encoding the toxin can be expressed in plants, and very effectively controls only pests eating the plants (unlike spraying which is indiscriminate).

Question 39

Applications of gene manipulation in medicine?

Answer 39

Insulin in bacteria: treatment of diabetes.

Was isolated from pig pancreas, but some patients made antibodies against the pig form, now use human form produced in bacteria.

Can produce the protein in lactating mammals or in eggs.

Direct production to mammary gland, by injecting DNA into pronucleus.

Question 40

Disease therapy example?

Answer 40

Sickle cell disease:

Patients make less or defective haemoglobin and fewer or defective red blood cells, causing anaemia.

Result from defects in the β-globin gene.

Therapy = deliver normal gene into hematopoietic stem cells from patient (ex vivo) using a lentiviral vector (modified HIV) – ‘LentiGlobin’.

Some patients in trials are becoming transfusion independent.

Commercial venture – Bluebird Bio, Massachusetts, USA