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BIOC1007 Simona > Applications of molecular biology > Flashcards

Applications of molecular biology Flashcards

1
Q

What is DNA cloning?

A

It is essentially making many copies of DNA. Isolation and amplification of a specific gene or DNA sequence

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2
Q

What does DNA cloning involve?

A

Separating a specific gene or DNA segment from a larger chromosome, attaching it to a small molecule of carrier DNA, and then replicating this modified DNA many times through a) increase in host cell number and b) the creation of multiple copies of the cloned DNA in each cell.

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3
Q

What are the five general procedures entailed in cloning DNA from any organism?

A
  1. Cutting DNA at precise locations
  2. Selecting a cloning vector, a small molecule of DNA capable of self-replication. (plasmid, phage, etc.)
  3. Joining the two DNA fragments covalently. Composite DNA molecules comprising covalently linked segments from two or more sources are called recombinant DNAs.
  4. Moving recombinant DNA from the test tube to a host cell, which will provide the enzymatic machinery for DNA replication. (e.g E.Coli)
  5. Selecting or identifying host cells that contain the DNA.
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4
Q

What is the result of DNA cloning?

A

The result is the selective amplification of a particular gene or DNA segment.

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5
Q

What is the main organism used in recombinant DNA work, and why?

A
  • E.Coli* , as it has many advantages:
  • well understood
  • many naturally-occurring cloning vectors (plasmids, phages) are associated with it
  • techniques are available for moving DNA from one cell to another
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6
Q

Why can’t DNA be isolated biochemically, like proteins?

A

Different DNA molecules do not differ in chemical properties like proteins do, and thus there are no differences to be exploited for separation methods.

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7
Q

Which two enzymes lie at the heart of the classic recombinant DNA approach?

A
  • Restriction endonucleases
  • DNA ligases
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8
Q

What do RE do?

A

They recognise and cleave DNA at specific sequences (restriction sites/recognition sequences) to generate a set of smaller fragments.

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9
Q

What does DNA ligase do?

A

Joins the DNA fragment to be cloned and the suitable cloning vector covalently.

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10
Q

What is the natural biological function of RE?

A

It is to recognise and cleave foreign DNA, e.g viral genetic material. They are naturally occurring bacterial enzymes that serve as defense mechanisms.

The host cell’s DNA is protected from RE digestion by methylation of DNA, catalysed by DNA methylase.

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11
Q

What do type II RE do?

A

Cleave DNA within specific recognition sequences.

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12
Q

Why are type I and type III RE not used?

A

Type I: cleaves at approx. 1000 bp from recognition sequence

Type III: cleaves at approx. 25bp from recognition sequence

So they are less reliable/predictable, more complicated and require ATP for movement along the DNA molecule.

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13
Q

Give two features of recognition sequences

A
  • 4-6 bp long
  • palindromic
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14
Q

What does DNA pol I do?

A

Fills gaps in duplexes by stepwise addition of nucleotides to 3’ end

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15
Q

What does reverse transcriptase do?

A

Makes a DNA copy of an RNA molecule

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16
Q

What is the function of polynucleotide kinase and why is it useful?

A

Adds a phosphate to the 5’ OH end of a polynucleotide to label it or permit ligation. It is useful as this enzyme enables to label the polynucleotide with a radioactive phosphate.

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17
Q

What does terminal transferase do?

A

Adds homopolymer tails to 3’OH of a linear duplex

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18
Q

What does exonuclease III do?

A

Removes nucleotides from 3’ end

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19
Q

What does bacteriophage lambda exonuclease do?

A

Removes nucleotides from the 5’ ends of a duplex to expose single stranded 3’ ends

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20
Q

What does alkaline phosphatase do?

A

Removes terminal phosphates from either or both ends.

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21
Q

What does ribonuclease H do?

A

They are endonucleases that catalyse the cleavage of RNA via a hydrolytic mechanism. In DNA replication, RNase H is responsible for removing the RNA primer, allowing completion of the newly synthesized DNA.

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22
Q

What is a cDNA library?

A

A library that includes only those genes that are expressed, i.e that are transcribed into RNA, in a given organism or even in certain cells or tissues. Such a library is designed to focus on those portions of a genome relevant to the function of a tissue or cell type.

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23
Q

How is a cDNA library constructed?

A
  1. Choose tissue of interest (as genes follow tissue-specific gene expression).
  2. Isolate mRNA (mRNA expressed << total DNA in cell), using affinity chromatography column linked to thymine (polyA tail)
  3. Copy mRNA into cDNA in a multistep reaction catalysed by reverse transcriptase.
  4. Introduce cDNA into a cloning vector and clone.

Result = creation of a population of clones called a cDNA library.

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24
Q

Why is a cDNA library useful?

A

By focusing on a CDNA library generated from the mRNAs of a cell known to expressa certain gene, the search for a particular gene is made easier.

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25
Q

Outline the 5 steps of the synthesis of duplex cDNA from mRNA

A
  1. mRNA template is annealed to a synthetic oligo dT DNA primer.
  2. Complementary strand yielded by reverse transcriptase, giving a DNA-RNA hybrid
  3. mRNA strand is degraded with alkali
  4. A DNA oligonucleotide of known sequence is annealed to the 3’ end of the cDNA to prime synthesis of second DNA strand.
  5. DNA pol I extends the primer to yield double-stranded DNA.
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26
Q

What happens after this?

A
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27
Q

What are the three main features of a plasmid cloning vector?

A
  • Multiple cloning site (MCS, polylinker): contains lots of restriction sites.
    2. Origin of replication (ori)
    3. Selectable marker (e.g Amp R)
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28
Q

Where is the foreign DNA inserted in the plasmid cloning vector?

A

Into one of the sites within the MCS, depending on the RE with which it has been digested.

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29
Q

How can a plasmid be taken up by a host cell (E.Coli)?

A

Two ways of making membrane permeable to large molecules:

  • Transformation: treatment with calcium chloride followed by rapid heat shock
  • Electroporation: cells subjected to high voltage pulse, which transiently renders the bacterial membrane permeable to large molecules
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30
Q

How are host cells that have taken up the plasmid selected for?

A
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31
Q

After a cDNA library has been constructed, what is the next step?

A

Screening the library with an appropriate probe, to detect clones with a particular DNA segment.

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32
Q

What is a probe?

A

It is a labeled (e.g radioactive) DNA or RNA fragment complementary to the DNA being sought.

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33
Q

Describe one classic approach to screen a DNA library, 5 steps

A
  1. Take agar plate with transformed bacterial colonies, and press nitrocellulose paper onto the agar plate. In this way, some cells from each colony stick to the paper.
  2. Treat the nitrocellulose paper with the cells on it with alkali in order to disrupt cells and expose denatured DNA. The result is DNA bound to the nitrocellulose paper.
  3. Incubate the paper with radiolabeled DNA probes, then wash. The probes will hybridise with the complementary sequence on the paper and thus will be bound to the colonies of interest.
  4. Expose X-Ray film to paper and obtain an autoradiograph to detect colonies of interest.
34
Q

What comes after this?

A
35
Q

How are oligonucleotide probes created?

A

They are designed from a protein sequence. However, due to the degenerate nature of the genetic code, the DNA sequence for a known amino acid sequence cannot be known in advance. Thus, the probe is designed to be complementary to a region of the gene with minimal degeneracy, that is, a region with the fewest possible codons for the amino acids (e.g choose a region of aa for which there are two codons at most for each aa).

Synthetic probes are created with selectivley randomised sequences, so that they contain either of the n possible sequences at each position of the potential degeneracy.

36
Q

How is the possible number of oligonucletoide sequences calculated?

A

For a minimal degeneracy region with k amino acids:

Nº possible sequences = Nº codon aa1 + Nº codons for aa2 + …….+ Nº codons for aaK

37
Q

What are expression vectors?

A

Cloning vectors with transcription and translation signals needed for the regulated expresion of a cloned gene.

Used because sometimes it is the product of the cloned gene, rather than the gene itself, that is of primary interest, particularly if the gene product (protein) has commercial, therapeutic or research value.

38
Q

What elements does a typical expression vector have? (7)

A
  • Polylinker: contains the gene to be expressed.
  • Promoter (P): allows efficient transcription of the inserted gene.
  • Transcription termination sequence: improves amount and stability of mRNA produced.
  • Operator (O): permits regulation by means of a repressor that binds to it.
  • Ribosome binding site: provides sequence signals needed for efficient translation of the mRNA derived from the gene.
  • Selectable marker: allows selection of cells containing recombinant DNA.
  • ori
  • gene encoding repressor that binds O and regulates P
39
Q

What is an alternative way of screening a cDNA library, particularly when interseted in a gene product?

A

Even if DNA is present (known froms screening with DNA probes), it doesn’t mean that the cell is expressing the protein, so in some cases we are better off screening directly for that by using antibodies.

40
Q

How is a cDNA library screened with antibodies for a particular gene product? (4)

A
  1. Press nitrocellulose paper onto plate with transformed bacterial colonies. The proteins will bind to the nitrocellulose paper.
  2. Incubate the paper with primary antibody, made specifically for the protein that we’re trying to express using an organism such as a mouse.
  3. Incubate with secondary antibody which recognises the primay antibody (commercially available, produced in mass) and is enzyme-linked.
  4. Add enzyme substrate, identify product of enzymatic reaction and identify which colonies are expressing the gene.
41
Q

What is an important element to have when carrying out expression of foreign protein in bacteria?

A

A “switch” to turn expression on/off to not slow down growth, or in case the protein we’re trying to express is toxic to the host cell.

42
Q

What is an example of this expression switch?

A

IPTG: turns on promoter, looks like lactose.

Lac promoter: when there is hi [glucose], or no lactose, transcription of lac genes doesn’t occur.

So when IPTG is present (looks like lactose), transcription of lac genes occur. If we put our cDNA in the lacZ gene’s place, we can regulate its expression by adding/removing IPTG.

43
Q

What is blue white selection?

A

It is a screening technique that allows for the rapid and convenient detection of recombinant bacteria in vector-based molecular cloning experiments. DNA of interest is ligated into a vector. The insert disrupts the lacZ gene, meaning that no alpha-peptide is made, and by extension no functional B-galactosidase is formed. The vector is then inserted into a competent host cell viable for transformation, which are then grown in the presence of X-gal. X-gal is hydrolysed by B-galactosidase to yield an insoluble blue compound. Cells transformed with vectors containing recombinant DNA will produce white colonies; cells transformed with non-recombinant plasmids (i.e. only the vector) grow into blue colonies.

44
Q

What are markers and what are they used for?

A

It is a gene that researchers attach to a regulatory sequence of another gene of interest in bacteria, cell culture, animals or plants. A cDNA library can be made even more specialised by cloning cDNAs fragments into a vector that fuses each cDNA sequence with the sequence for a marker, or reporter gene; the fused genes form a “reporter construct”.

45
Q

What is GFP?

A

Green fluorescent protein, used as useful marker: the gene for GFP is fused with the target gene, which generates a fusion protein that is highly fluorescent and thus can be easily visualised microscopically, allowing the study of its location and movement in a cell.

46
Q

What is an epitope tag?

A

Epitope = part of antigen recognised by immune system

A short protein sequence that is bound tightly by a well-characterised monoclonal antibody. This allows for the tagged protein to be specifically precipitated from a crude protein extract by interaction with the antibody. The precipitated proteins can then be separated by means of gel electrophoresis, and new proteins in the preicpitate can then be identified (see below).

In addition, any other proteins that interact with the tagged protein also precipitate, helping to elucidate protein-protein interactions.

47
Q

How can recombinant DNA-technology be applied to the production of proteins important for medicine or industry?

A

Aim: synthesise a human protein in bacteria, as these are difficult to isolate from naturally-occurring sources (humans).

How is it done?

  • clone human gene into bacteria in a vector that has all the elements that the host cell needs to express the protein (promoter, operator, ribosome binding site, etc.) and then trick the bacteria into thinking they are producing their own protein.
48
Q

Nam 3 proteins made in bacteria for medical use

A
  • Insulin: recombinant insulin has many advantages, one of which is that it can be used instantly after injection, generally more effective
  • Growth hormone
  • Factors VIII and IX
49
Q

What is oligonucleotide site-directed mutagenesis?

A

Cloning techniques can be used not only to overproduce proteins but also to produce protein products slightly altered from their native forms. Site-directed mutagenesis is a technique used to replace specific amino acids in order to make gene product more efficient.

It is used when a gene is not flanked by appropriate restriction sites and thus the DNA segment cannot simply be removed and replaced with a synthetic one that is identical to the original except for the desired change.

50
Q

What is (a), what is (b)?

A

(a) A synthetic DNA segment replaces a DNA fragment that has been removed by cleavage with an RE.
(b) Oligonucleotide site-directed mutagenesis: a synthetic oligonucleotide with a desired sequence change at one position is hybridised to a single-stranded copy of the gene to be altered. This acts as a primer for synthesis of a duplex DNA which is then used to transform cells. Cellular DNA repair systems will convert about 50% of the mismatches to reflect the desired sequence change.

51
Q

In OSDM, under which conditions is annealing possible?

A
  • If a single bp is altered
  • If T is appropriate
52
Q

What are the 3 steps/requirements of OSDM?

A
  1. Decide which codons to change (site-specific)
  2. Synthesise a primer (oligonucleotide) for DNA synthesis which includes the change
  3. Prepare a single-strand DNA template.
53
Q

How is the single-stranded template for OSDM prepared?

A
  1. Recombinant M13 vector (double stranded DNA) is transfected into E.Coli
  2. Phage are releasesd, a recombinant M13 phage has a DNA core wrapped in a protein coat.
  3. Protein coat is removed: single stranded DNA is obtained, which can be used in the experiment.
54
Q

What is a trick used in OSDM?

A

Used of modified deoxynucleotides to synthesise the new strand. The sulphur-substituted DNA strand cannot be cut by RE, but these enzymes can cut the strand on the outside. So a RE with a RS downstream from the region we want to change is used, followed by exonuclease III which removes nucleotides to an extent dependent on enzyme and incubation time, then DNA is synthesised with appropriate change.

55
Q

Which aa is subsituted in Subtilisin by means of OSDM?

A

AUG (met222) —-> GCG (ala222)

56
Q

What is the most efficient way of purifying recombinant proteins?

A

Affinity chromatography- requires a specificity tag to be incorporated into cloning strategy.

57
Q

What is the advantage of adding a (His)6 tag, and what is its immobilised ligand?

A
  • Doesn’t require correct folding
  • Ni2+: the imidazole group has a great affinity for chelated metals.
58
Q

What is the immobilised ligand for GST?

A

Glutathione (GSH). GST catalyses the conjugation of electrophilic substrates to glutathione (GSH).

GST can be added to a protein of interest to purify it from solution. This is accomplished by inserting the GST DNA coding sequence next to that which codes for the protein of interest. Thus, after transcription and translation, the GST protein and the protein of interest will be expressed together as a fusion protein. Because the GST protein has a strong binding affinity for GSH, beads coated with the compound can be added to the protein mixture; as a result, the protein of interest attached to the GST will stick to the beads, isolating the protein from the rest of those in solution. The beads are recovered and washed with free GSH to detach the protein of interest from the beads, resulting in a purified protein. This technique can be used to elucidate direct protein-protein interactions

59
Q

Give the name of two tags used in affinity chromtography

A
  • Maltose binding protein (IL: maltose)
  • Chitin binding domain (IL: chitin)
60
Q

How can GST be cleaved off the target protein after affinity chromatography?

A

Protease cleavage site between protein of interest and GST. This is done to ensure that the observed properties are of the protein and not of the tag.

61
Q

How can each step in the purification procedure be monitored?

A

By means of gel electrophoresis.

62
Q

What is

a) Southern blotting
b) Northern blotting
c) Western blotting

A
63
Q

How does Southern blotting work?

A
64
Q

How can Southern blotting be used in forensic medicine?

A
65
Q

What is shown in the image?

A

The Southern blot procedure, as applied to RFLP DNA fingerprinting.

66
Q

What is PCR?

A

If we know the sequence of at least the flanking parts of a DNA segment to be cloned, we can hugely amplify the number of copies of that DNA segment, using the polymerase chain reaction (PCR). The amplified DNA can be cloned directly or used in a variety of analytical procedures.

67
Q

What are the requirements for PCR? (5)

A
  1. Two primers, complimentary to the sequene to be amplified. In order to synthesise these primers, the sequence of the DNA that we need to amplify must be known.
  2. Template DNA (that which is going to get copied)
  3. Thermostable DNA polymerase (TAC polymerase)
  4. dNTPs
  5. Thermal cycler
68
Q

What is the annealing temperature?

A

Tm = 4* (G+C) + 2* (A+T) (ºC)

69
Q

What are the three steps in the PCR procedure?

A
  1. Heat to separate strands
  2. Anneal primers containing noncomplementary regions with cleavage site for restriction endonuclease.
  3. DNA synthesis catalysed by TAC polymerase.

Repeat for 25-30 cycles, after which the target sequence will have been amplified a million fold

70
Q

In what procedure are fluorescent dye-labeled PCR primers used?

A

In PCR analysis of an STR locus.

  1. Conduct PCR of STR locus.
  2. Run PCR fragments on a capillary gel (very thin, polyacrylamide)
  3. Scan of PCR product bands derived from locus, generates a series of peaks (as each allele in the locus might have different number of repeats, the PCR can generate two products of slightly different size).
  4. Combining the scans for different loci gives a multiplex pattern.
71
Q

What is a DNA microarray and what is it used for?

A

A DNA microarray is a collection of microscopic DNA spots attached to a solid surface. Scientists use DNA microarrays to measure the expression levels of large numbers of genes simultaneously or to genotype multiple regions of a genome.

Essentially, it is used to analyse gene expression in 2 different places (e.g 2 different points of development, or differences between 2 people to look at disease)

72
Q

What are the 3 steps for a DNA microarray?

A
  1. Isolate mRNAs from cells (each mRNA represents all the genes expressed in the cells at that particular stage)
  2. Convert inot cDNAs, using fluorescently labeled dNTPs.
  3. Add cDNAs to a microarray; fluorescent cDNAs anneal to complementary sequences on the microarray.

More detailed explanation:

A microarray can be prepared from any known DNA sequence, from any source, generated by chemical synthesis or by PCR. The DNA is positioned on a solid surface (usually specially treated glass slides) with the aid of a robotic device capable of depositing very small (nanoliter) drops in precise patterns. UV light cross-links the DNA to the glass slides. Once the DNA is attached to the surface, the microarray can be probed with other fluorescently la- beled nucleic acids. Here, mRNA samples are collected from cells at two different stages in the development of a frog. The cDNA probes are made with nucleotides that fluoresce in different colors for each sam- ple; a mixture of the cDNAs is used to probe the microarray. Green spots represent mRNAs more abundant at the single-cell stage; red spots, sequences more abundant later in development. The yellow spots indicate approximately equal abundance at both stages

73
Q

What are the implications of cloning in plants?

A

Allows to enhance natural properties (e.g increase nutritional content), and allows to give other properties: herbicide resistance, pathogen resistance, biofuels, bioremediations, etc.

74
Q

What is one method used for cloning in plants?

A

Using Agrobacterium tumefaciens

Lives in soil, attached to the root of a plant, forms “tumor” on plant via transformation with Ti plasmid (pk to ek DNA transfer).

It can invade plants at the site of a wound, transform nearby cells, induce them to form a tumor called crown gall.

75
Q

What are the main features of the Ti plasmid, found in agrobacterium tumefaciens?

A

vir genes: involved in transfer of genes from bacteria, together with the 25bp repeats that flank DNA. Encode enzymes necessary to introduce TDNA segment of Ti plasmid into genome of neraby cells.

T DNA: transferred from plasmid into plant cell chromosome. It encodes enzymes that convert plant metabolites into 2 compounds that benefit the bacterium: plant growth hormones, to form the crown gall tumor, and opines (aa), which serve as a food source for the bacterium

76
Q

How does AT work?

A
77
Q

How can the natural process mediated by AT be exploited?

A

Require: transfer ability, ability to make recombinant plasmids

We do not need: additional cell growth, production of opines

78
Q

How is a recombinant plant created?

A

One plasmid is a modified Ti plasmid that contains the vir genes but lacks T DNA. The other plasmid contains a segment of DNA that bears both a foreign gene (the gene of interest) and an antibiotic-resistance element (here, kanamycin resistance), flanked by the two 25 bp repeats of T DNA that are required for transfer of the plasmid genes to the plant chromosome. This plasmid also contains the replication origin needed for propagation in Agrobacterium.

When bacteria invade at the site of a wound (the edge of the cut leaf), the vir genes on the first plasmid mediate transfer into the plant genome of the segment of the second plasmid that is flanked by the 25 bp repeats. Leaf segments are placed on an agar dish that contains both kanamycin and appropriate levels of plant growth hormones, and new plants are generated from segments with the transformed cells. Non- transformed cells are killed by the kanamycin. The foreign gene and the antibiotic-resistance element are normally transferred together, so plant cells that grow in this medium generally contain the foreign gene.

79
Q

What is this?

A

A two-plasmid strategy to create a recombinant plant.

80
Q

What are the forward and reverse PCR primers for this sequence? (—– is the gene of interest)

5’ ATGCTGACCGTGCATGTA——–GTCACCTGGTGATTACGT 3’

3’ TACGACTGGCACGTACAT——–CAGTGGACCACTAATGCA 5’

A

In order to amplify the gene of interest, two replication primers must be designed, complementary to sequences on opposite strands of the target DNA at positions defining the ends of the segment to be amplified. In this case, the two PCR primers are:

Forward: 5’ ATGCTGACCGTGCATGTA 3’

Reverse: 5’ ACGTAATCACCAGGTGAC 3’

81
Q

What features should you try and avoid when designing primers?

A

The Tm of both primers needs to be similar enough to enable to set an annealing temperature that is suitable for both. Thus, very dissimilar Tm values between both primers should be avoided.(if too low, allows for non-specific binding, if high enough, E is put into the system, only strongest interactions reisst, i.e the specific ones).

In addition, the sequences of the primers must be designed so as to avoid self- annealing (arises from the fact that it is easier for it to close on itself because it’s closer, even though binding to complementary strand is much more favourable)

The length of the primers is another feature that must be taken into account. Ideally, the primers should be 18-30 nucleotides long. Shorter sequences should be avoided because they largely decrease the specificity of the primer.

Runs of the same nucleotide or dinucleotide repeats should also be avoided (can potentially cause a frameshift)

BIOC1007 Simona (5 decks)

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