Gene cloning, Lecture notes of Biogenetics and Computers

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ASSIGNMENT

ON

GENE CLONING

(ADVANCES IN GENETIC ENGINEERING-UNIT II)

Submitted by:

Evangeline Wills

Reg No. 08HA

I MTech Biotechnology

Sub: AGE BT-

DNA Libraries

In molecular biology, a library is a collection of molecules in a stable form that represents some aspect of an organism. Two common types of libraries are cDNA libraries (formed from Complementary DNA) and genomic libraries. The nucleotide sequences of interest are preserved as inserts to a plasmid or the genome of a bacteriophage that has been used to infect bacterial cells.

Preparation of a DNA Library

DNA library is a collection of cloned DNA fragments. There are two types of DNA library:

• The genomic library contains DNA fragments representing the entire genome of an organism.
• The cDNA library contains only complementary DNA molecules synthesized from mRNA molecules in a cell. Genomic Library A genomic library is a set of clones, packaged in the same vector, that together represents all regions of the genome of the organism in question. The number of clones that constitute a genomic library depends on (1) the size of the genome in question and (2) the insert size tolerated by the particular cloning vector system. The genomic library is normally made by l phage vectors, instead of plasmid vectors, for the following reasons:

The entire human genome is about 3 x 10 9 bp long while a plamid or F 06 C phage vector may carry

up to 20 kb fragment. This would require 1.5 x 10 5 recombinant plasmids or F 06 C phages. When plating E. coli colonies on a 3" petri dish, the maximum number to allow isolation of individual colonies is about 200 colonies per dish. Thus, at least 700 petri dishes are required to construct a human genomic library. By contrast, as many as 5 x 10 4 F 06 C phage plagues can be screened on

a typical petri dish. This requires only 30 petri dishes to construct a human genomic library.

Another advantage of F 06 C phage vector is that its transformation efficiency is about 1000 times higher than the plasmid vector. cDNA Library A cDNA library represents all of the mRNA present in a particular tissue, which has been converted back to a DNA template by the use of the enzyme reverse transcriptase. It represents the genes that are transcribed in particular tissues under particular physiological, developmental, or environmental conditions. cDNA libraries are useful in reverse genetics, but should not be confused with a genomic library, as it does not represent the entire genome, only a very small (less than 1%) portion which is being transcribed.

Usually a cDNA library is created when reproducing eukaryotic genomic material, whereas genomic libraries are often created when working with genomic target material from bacteria and viruses.

partial digests of genomic DNA are subjected to agarose gel electrophoresis for separation from the mixture of fragments of appropriate size.

Partial digestion as a means of isolating longer, overlapping DNA fragments (Fig: Lodish et al, 4th ed)

• Advantages:
  • same set of fragments are obtained from a DNA each time a specific enzyme is used and many of the enzymes produce cohesive ends.
  • Cohesive DNA ends for efficient ligation.
  • Choose enzymes for different average insert sizes (e.g. 4-base cutter, 6- base cutter, longer recognition site)

• Disadvantages:
  • May get non-random distribution of fragment sizes, due to non-random distribution of enzyme sites
  • Enzymes may not recognise methylated DNA.
  • (^) Much eukaryotic DNA is extensively methylated The partial digests of genomic DNA are subjected to agarose gel electrophoresis for separation from the mixture of fragments of appropriate size.

3. (^) These fragments are then inserted into a suitable vector for cloning. In principle, any vector can be used, but λ vectors and cosmids have been the most commonly used for cloning.
4. The vectors containing the inserts are cloned in a suitable bacterial host.

Figure: Preparation of the genomic library using F 06 C phage vectors. It is basically the cloning of all DNA fragments representing the entire genome.

cDNA library

A cDNA library is a population of bacterial transformants or phage lysates in which each mRNA isolated from an organism or tissue is represented as its cDNA insertion in a plasmid or a phage vector.

Preparation of cDNA

1. cDNA is the copy or complementary DNA produced by using mRNA as a template and enzyme reverse transcriptase. This enzyme performs similar reactions as DNA polymerase and requires a primer with a free 3’—OH. A poly-T oligonucleotide is conveniently used as the primer since these mRNAs have a poly-A tail at their 3’ ends.
2. The appropriate oligonucleotide primer is annealed with the mRNA: this primer will base pair to the 3’ end of mRNA. Reverse transcriptase extends the 3’ end of the primer using mRNA molecule as a template. This produces a RNA-DNA hybrid molecule.
3. The RNA strand is digested either by RNase H or by alkaline hydrolysis; this frees the single stranded cDNA.
4. The end of this cDNA serves as its own primer and provides the free 3’—OH required for the synthesis of its complementary strand. Therefore, a primer is not required for this step.

5. The complementary strand of cDNA single strand is synthesized either by the reverse transcriptase itself or by E. coli DNA polymerase I.
6. Since the 3’ end of the cDNA single strand is used as primer for this reaction a short hairpin loop is generated at this end. This loop is cleaved by single strand specific nuclease to yield a regular DNA duplex.

cDNA Library Construction

cDNA is created from a mature mRNA from an eukaryotic cell with the use of an enzyme known as reverse transcriptase. In eukaryotes, a poly-(A) tail (consisting of a long sequence of adenine nucleotides) distinguishes mRNA from tRNA and rRNA and can therefore be used as a primer site for reverse transcription

Need for construction of cDNA library

Use of cDNA is absolutely essential when the expression of a eukaryotic gene is required in a prokaryote e.g. a bacterium. This is because eukaryotic genes have introns, which must be removed from their transcripts to yield mature mRNAs. Bacteria do not possess the enzymes necessary for removal of introns. In contrast, functional mRNA molecules do not have introns; hence, their cDNA is also free of introns and can be cloned and expressed in bacteria. For e.g. cDNA for interferon, blood clotting factor VIII C and several other mRNAs have been expressed in bacteria.

cDNA Library uses

cDNA libraries are commonly used when reproducing eukaryotic genomes, as the amount of information is reduced to remove the large numbers of non-coding regions from the library. cDNA libraries are most useful in reverse genetics where the additional genomic information is of less use.

cDNA Library vs. Genomic DNA Library

As previously mentioned, a cDNA library lacks the non-coding and regulatory elements found in genomic DNA. Genomic DNA libraries provide much more detailed information about the organism, but are much more resource-intensive to generate and maintain.

Figure: Screening of a specific DNA fragment. After recombinant F 06 C virions form plaques on the lawn of E. coli , the nitrocellulose filter (membrane) is placed on the surface of the petri dish to pick up F 06 C phages from each plaque.^ Then, the filter is incubated in an alkaline solution to disrupt the virions and release the encapsulated DNA, which is subsequently denatured. Next, the probe is added to hybridize with the target DNA fragment, whose position may be displayed by autoradiography.

Chromosome jumping

Chromosome jumping is a technique of molecular biology that is used as a tool in the physical mapping of genomes. It is related to several other tools used for the same purpose, including chromosome walking.

Chromosome jumping is used to bypass regions difficult to clone, such as those containing repetitive DNA, that cannot be easily mapped by chromosome walking, and is useful in moving along a chromosome rapidly in search of a particular gene.

In chromosome jumping, the DNA of interest is identified, cut into fragments with restriction enzymes, and circularised (the beginning and end of each fragment are joined together to form a circular loop). From a known sequence a primer is designed to sequence across the circularised junction. This primer is used to jump 100 kb-300 kb intervals: a sequence 100 kb away would have come near the known sequence on circularisation. Thus, sequences not reachable by chromosome walking can be sequenced. Chromosome walking can be used from the new jump position (in either direction) to look for gene-like sequences, or additional jumps can be used to progress further along the chromosome.

Chromosome walking

A technique which produces sets of overlapping DNA clones for studying segments of DNA larger than can be cloned individually. A method for the analysis of large regions of DNA, in which a each end of a large single cloned DNA fragment is used separately to screen recombinant DNA genome library for other clones containing neighbouring sequences.

This method is used to move systematically along a chromosome from a known location and to clone overlapping genomic clones that represent progressively longer parts of a particular chromosome. Chromosome walking is used as a means of finding adjacent genes ( positional cloning ), or parts of a gene which are missing in the original clone as well as to analyse long stretchs of eukaryotic DNA

A small segment of DNA from one end of the genomic clone is used as a probe to isolate clones containing this sequence and adjacent sequences encoding the next portion of the genome. The end sequence of the second clone is used to isolate a third clone and so forth until a series of overlapping clones are isolated. It is necessary to use DNA probes whose sequences are single- copy, otherwise if the probe used is a repeated sequence, then several unrelated recombinants could be identified.

banding pattern, or electropherogram, much like a bar code, that can identify a species or individual (some genes will be vary at the species level and others at the individual level).

The simplest application of this technique is to assess whether a given protein binds to a region of interest within a DNA molecule. The wet lab methodology is summarized, with appropriate selection of reagents discussed, below.

1. Polymerase chain reaction (PCR) amplify and label region of interest that contains a potential protein-binding site, ideally amplicon is between 50 to 200 bit/s in length.
2. Add protein of interest to a portion of the labeled template DNA; a portion should remain separate without protein, for later comparison
3. Add a cleavage agent to both portions of DNA template. The cleavage agent is a chemical or enzyme that will cut at random locations in a sequence independent manner. The reaction should occur just long enough to cut each DNA molecule in only one location. A protein that specifically binds a region within the DNA template will protect the DNA it is bound to from the cleavage agent.
4. (^) Run both samples side by side on a polyacrylamide gel electrophoresis. The portion of DNA template without protein will be cut at random locations, and thus when it is run on a gel, will produce a ladder-like distribution. The DNA template with the protein will result in ladder distribution with a break in it, the "footprint", where the DNA has been protected from the cleavage agent.

Note: Maxam-Gilbert chemical DNA sequencing can be run alongside the samples on the polyacrylamide gel to allow the prediction of the exact location of ligand binding site.

Labeling

The DNA template can be labeled at the 3' or 5' end, depending on the location of the binding site(s). Labels that can be used are:

• (^) Radioactivity has been traditionally used to label DNA fragments for footprinting analysis, as the method was originally developed from the Maxam-Gilbert chemical sequencing technique. Radioactive labeling is very sensitive and is optimal for visualizing small amounts of DNA.
• Fluorescence is a desirable advancement due to the hazards of using radio-chemicals. However, it has been more difficult to optimize because it is not always sensitive enough to detect the low concentrations of the target DNA strands used in DNA footprinting experiments. Electrophoretic sequencing gels or capillary electrophoresis have been successful in analyzing footprinting of fluorescently tagged fragments.

Cleavage agent

A variety of cleavage agents can be chosen. Ideally a desirable agent is one that is sequence neutral, easy to use, and is easy to control. Unfortunately none available meet all these all of these standards, so an appropriate agent can be chosen, depending on your DNA sequence and ligand of interest. The following cleavage agents are described in detail:

• (^) DNase I : a large protein that functions as a double-strand endonuclease. It binds the minor groove of DNA and cleaves the phosphodiester backbone. It is a good cleavage agent for footprinting because its size makes it easily physically hindered. Thus is more likely to have its action blocked by a bound protein on a DNA sequence. In addition, the DNase I enzyme is easily controlled by adding EDTA to stop the reaction. There are however some limitations in using DNase I. The enzyme does not cut DNA randomly; its activity is affected by local DNA structure and sequence and therefore results in an uneven ladder. This can limit the precision of predicting a protein’s binding site on the DNA molecule.
• Hydroxyl radicals : are created from the Fenton reaction, which involves reducing Fe 2+ with H 2O 2 to form free hydroxyl molecules. These hydroxyl molecules react with the DNA backbone, resulting in a break. Due to their small size, the resulting DNA footprint has high resolution. Unlike DNase I they have no sequence dependence and result in a much more evenly distributed ladder. The negative aspect of using hydroxyl radicals is that they are more time consuming to use, due to a slower reaction and digestion time.
• Ultraviolet irradiation : can be used to excite nucleic acids and create photoreactions, which results in damaged bases in the DNA strand. Photoreactions can include: single strand breaks, interactions between or within DNA strands, reactions with solvents, or crosslinks with proteins. - The workflow for this method has an additional step, once both your protected and unprotected DNA have been treated, there is subsequent primer extension of the cleaved products. The extension will terminate upon reaching a damaged base, and thus when the PCR products are run side-by-side on a gel; the protected sample will show an additional band where the DNA was crosslinked with a bound protein. - Advantages of using UV are that it reacts very quickly and can therefore capture interactions that are only momentary. Additionally it can be applied to in vivo experiments, because UV can penetrate cell membranes. A disadvantage is that the gel can be difficult to interpret, as the bound protein does not protect the DNA, it merely alters the photoreactions in the vicinity.

Advanced Applications

In vivo footprinting

as a reference to engineer plasmids or other relatively short pieces of DNA, and sometimes for longer genomic DNA. There are other ways of mapping features on DNA for longer length DNA molecules, such as mapping by Transduction (Bitner, Kuempel 1981).

One approach in constructing a restriction map of a DNA molecule is to sequence the whole molecule and to run the sequence through a computer program that will find the recognition sites that are present for every restriction enzyme known to man.

Before sequencing was automated, it would have be prohibitively expensive to sequence an entire DNA strand. Even today sequencing is overkill for many applications. To find the relative positions of restriction sites on a plasmid a technique involving single and double restriction digests is used. Based on the sizes of the resultant DNA fragments the positions of the sites can be inferred. Restriction mapping is very useful technique when used for determining the orientation of an insert in a cloning vector, by mapping the position of an off-center restriction site in the insert (Dale, Von Schantz, 2003).

Restriction enzymes:

Restriction enzymes are enzymes that cut DNA at specific recognition sequences called "sites." They probably evolved as a bacterial defense against DNA bacteriophage. DNA invading a bacterial cell defended by these enzymes will be digested into small, non-functional pieces. The name "restriction enzyme" comes from the enzyme's function of restricting access to the cell. A bacterium protects its own DNA from these restriction enzymes by having another enzyme present that modifies these sites by adding a methyl group. For example, E.coli makes the restriction enzyme Eco RI and the methylating enzyme Eco RI methylase. The methylase modifies Eco RI sites in the bacteria's own genome to prevent it from being digested.

Restriction enzymes are endonucleases that recognize specific 4 to 8 base regions of DNA. For example, one restriction enzyme, Eco RI, recognizes the following six base sequence:

5'... G-A-A-T-T-C... 3'

3'... C-T-T-A-A-G... 5'

A piece of DNA incubated with Eco RI in the proper buffer conditions will be cut wherever this sequence appears. As you can see, this site is palindromic; that is, reading the upper strand from 5' to 3' is the same as reading the lower strand from 5' to 3'. As a result, each strand of the DNA can self-anneal and the DNA forms a small cruciform structure:

All restriction enzyme sites are palindromic. This structure may help the enzyme to recognize the sequence that it is designed to cut.

There are hundreds of restriction enzymes that have been isolated and each one recognizes its own specific nucleotide sequence. Sites for each restriction enzyme are distributed randomly throughout a particular DNA stretch. Digestion of DNA by restriction enzymes is very reproducible; every time a specific piece of DNA is cut by a specific enzyme, the same pattern of digestion will occur. Restriction enzymes are commercially available and their use has made manipulating DNA very easy.

Restriction Mapping:

Restriction mapping involves digesting DNA with a series of restriction enzymes and then separating the resultant DNA fragments by agarose gel electrophoresis. The distance between restriction enzyme sites can be determined by the patterns of fragments that are produced by the restriction enzyme digestion. In this way, information about the structure of an unknown piece of DNA can be obtained. An example of how this works is shown below. If we have isolated a clone in pBluescript we know how big the pBluescript portion of the plasmid is (3.0 kilobases) and what restriction enzymes are present in the plasmid. We also know that the insert is 2.0 kb long and that it is inserted the Eco RI site.

At this point, we digest plasmid with an enzyme that we know is in the pBluescript plasmid. For e.g., we know that there is only one Bam HI site in pBluescript, and it is in the multiple cloning site next to the Eco RI site.If we digest this plasmid with Bam HI, there are 2 possibilities: