UNIT IGene analysis techniques – Isolation of DNA and RNA – Handling and Quantification ofnucleic acids – Radiolabelling of nucleic acids – Gel Electrophoresis – Probing for a specific gene – Southern blotting, Northern blotting. Dot blotting, Western blotting. Chromosome walking – Heteroduplex analysis.

Section A

1.The lysis of animal cells is usually performed using anionic detergents such as SDS (sodium deodecyl sulfate) or Sarcosyl (sodium deodecyl sarcosinate).

2.

Isolation of Nucleic Acid

There are many methods available for purifying nucleic acids. The choice of method depends on the type, source, size and amount, and quality required for the procedure in which the nucleic acid is to be used. The advantages and disadvantages of each purification method are presented in Table 1 (See notes/conditions & reference protocols).

Table 1Nucleic Acid Purification Method Pierce

Prod. #Benefit Disadvantage

Phenol/alcohol (Guanidine isothiocyanate)

17906 17908 17909 17912 17914

Methods for nucleic acids of all sizes and strands; removes proteins; large or small scale

Phenol must be buffered and free of oxidative products; poison & caustic; dilute purified product may require ethanol precipitation to concentrate

Ethanol 51102, TE 17890

Fast, easy and efficient; works with many salt combinations

Pellets are difficult to see and may detach; ethanol may carry over and damage enzyme activity; over-drying the pellet

make re-dissolving difficult

DEAE, DE52 Column or paper

Purification of modified nucleic acids

Eluate often contains high salt; dilute purified product may require ethanol precipitation to concentrate

Size Exclusion Chromatography

- Fast and easy method to remove small molecules

Sample loss from large surface area; separation is not always 100%; nucleic acid dilution

Cesium Chloride - Cleanest method for DNA; choice method for >15kb and closed circular plasmids for biophysical measurement

Time consuming; expensive; EtBr is a mutagen; DNA must be purified away from CsCl for subsequent use

Extraction from agarose

- Simple to perform Recovery may be low (50%); increased size decreases recovery; poor recovery for < 50 ng; may require extra purification; agarose carryover may inhibit enzymes

Silica Resin - Fast and easy to perform; safe alternative to phenol/chloroform; clean nucleic acids; >80% recovery

Different reagents are required for different sized nucleic acids; dilution of nucleic acids; ethanol may carry over and damage enzyme activity

DNA extraction methods. After discussing briefly the structure of DNA. DNA extraction procedures are given below. For every genetic analysis, we require a good quality DNA. Nowadays readymade kits are available in the market by which the DNA can be extracted but some time it is difficult for the laboratories to use the commercial kits because of the cost.There are various manual protocols about the DNA extraction.However, in each protocol several steps are involved in the preparation of DNA. These are - (i) cell breakage (ii) removal of protein (iii) removal of RNA (iv) concentration of DNA (v) determination of the purity and quantity of DNA.1. Cell breakageCell breakage is one of the most important steps in the purification of DNA. The usual means of cell disruption, such as sonication, grinding, blending, or high pressure, cannot be used in DNA isolation. These procedures apply strong force for the disruption of cells which get sheared, as a result the DNA is fragmented. The best procedure for opening cells and obtaining intact DNA is through application ofchemical (detergents) and / or enzymatic procedures. Detergents can solubilize protein as well as lipids in cell membranes resulting into gentle cell lysis. In addition, detergents have an inhibitory effect on all

cellular DNAses and can denature proteins, thereby aiding in the removal of proteins from the solutions. The lysis of animal cells is usually performed using anionic detergents such as SDS (sodium deodecyl sulfate) or Sarcosyl (sodium deodecyl sarcosinate).2. Removal of proteinThe second step in purification involves removal of major contaminant,namely protein, from the cell lysate. This procedure is called deproteinization. Removal of proteins from the DNA solution depends on differences in the physical properties between nucleic acids and proteins. These differences are in solubility, in partial specific volume, and in sensitivity to digestive enzymes. (i) Deproteinization using organic solventsThe most frequently used methods for removing proteins explore the solubility differences between proteins and nucleic acids in organic solvents. Nucleic acids are hydrophilic molecules and are easily soluble in water. Proteins, on the other hand, contain many hydrophobic re si dues making them partially soluble in organic solvents. There are several methods of deproteinization based on this difference and they vary by the choice of the organic solvent introduced:1. The use of ionic detergents: These detergents, by unfolding the protein, help to expose hydrophobic regions of the polypeptide chains to phenol micelles, thereby aiding partitioning of proteins into the phenol phase.2. Enzymatic removal of proteins before phenol extraction: This reduces the number of extractions needed, thus, limiting the loss and shearing of DNA.3. Addition of 8HQ (8-Hydroxy-Quinoline) to the phenol: This increases the solubility of phenol in water. In the presence of this compound phenol remains liquefied at room temperature with only 5 percent water. In addition, BHQ is easily oxidized and, therefore, it plays the role of an anti-oxidant, protecting phenol against oxidation. Since the reduced form of SHQ is yellow and the oxidized form is colorless, the presence or absence of yellow color is an excellent visual indicator of the oxidation state of phenol.4. Removal of oxidation products from phenol and prevention of oxidation upon storage or during phenol extraction: Because water-saturated phenol undergoes oxidation rather easily, particularly in the presence of buffers such as Tris, Phenol used for DNA purification is twice distilled, equilibrated with water, and stored in the presence of 0.1 percent SHQ. 5. Adjusting the pH of water-saturated phenol solution to above pHS by equilibration of the liquefied phenol with a strong buffer or sodium borate. DNA obtained by this method is usually of high molecular weight, but contains approximately 0.5 percent protein impurities that can be removed by another method.

The application of chloroform: isoamyl alcohol (CIA) mixture can also be used which is also known as the deproteinization method. This is based on a characteristic of this organic solvent that differs from phenol. The chloroform is not miscible with water and, therefore, even numerous extractions do not result in DNA loss into the organic phase. The deproteinization action of chloroform is based on the ability of denatured polypeptide chains which partially enter or be immobilized at the water-chloroform interphase. The resulting high concentration of protein at the interphase causes protein to precipitate. Since the deproteinization action of chloroform occurs at the chloroform-water interphase, efficient deproteinization depends on the formation of a large interphase area. To achieve this, one has to form an emulsion of water and chloroform. Chloroform does not mix with water. This can only be done by vigorous shaking. An emulsifier, isoamyl alcohol, is added to chloroform to help to form the emulsion and to increase the water- chloroform surface area.

The characteristics of enzymatic removal of proteins make enzymatic deproteinization an ideal and indispensable first step in nucleic acid purification. This treatment is used when a large amount of protein ispresent, i.e. right after cell lysis. The remaining proteins can be removed with a single extraction using organic solvent.(ii) Removal of RNAThe removal of RNA from DNA preparations is usually carried out using an enzymatic procedure. Consequently, this procedure does not remove all RNA and, therefore, yield DNA preparations with avery small amount of RNA contamination. Two ribonucleases that can be easily and cheaply prepared free of DNase contamination are ribonucleases A and ribonucleases Tt.

(a) Ribonuclease A (RNase A) is an endoribonuclease that cleaves RNA after C and U residues. The reaction generates 2': 3' –cyclic phosphate which is hydrolyzed to 3' -nucleoside phosphate producingoligonuc1eotides ending with 3' -phosphorylated pyrimidine nucleotide.(b) Ribonuclease Tt (RNase Tt) is an endoribonuclease that is very similar to RNase A in a reaction conditions and stability. The enzyme cleaves double-stranded and single-stranded RNA after G residues,generating oligonucleotides ending in a 3' -phosphorylated guanosine nucleotide.(iii) Concentrating the DNAPrecipitating with alcohol is usually performed for concentration of DNA from the aqueous phase of the deproteinization step. Two types of alcohols can be used for DNA precipitation: ethanol and isopropanol.Alcohol precipitation is based on the phenomenon of decreasing the solubility of nucleic acids in water. Polar water molecules surround the DNA molecules in aqueous solutions. The positively chargeddipoles of water interact strongly with the negative charges on the phosphodiester groups of DNA. This interaction promotes the solubility of DNA in water. Ethanol is completely miscible with water, yet it is far less polar than water. Ethanol molecules cannot interact with the polar groups of nucleic acids as strongly as water, making ethanol a very poor solvent for nucleic acids..(iv) Determination of purity and quality of DNAThe last step in DNA isolation is the quality of the DNA being isolated UV spectrometry is used for determining the DNA concentration. The DNA has maximum absorbance at 260nm and minimumabsorbance at 234 nm. This can get affected by the PH of the medium in which the DNA is dissolved.DNA concentration (N) =A260/E260; E260 is the DNA extinction coefficient. This is 0.02 Ilg -1 cm -1 when measured at neutral or little basic PH for double helical DNA. Thus, an absorbance of 1.0 at 260 nm gives a DNA concentration of 50 Ilg ml-l (1/0.02 =50 mg ml-l)" However, this may differ because of GC percent. If the DNA is found in small concentrations then dust particle scattering may effect the measurements. This can be checked by taking one reading at 320 nm, DNA is not absorbed at 320 nm, hence, if there is any recorded reading, it is because of dust contamination. If there is no contamination then at 320 the reading should be 5% less than the absorbance reading at 260. At 280 nm the protein concentration is measured as the protein is absorbed maximally at 280 nm. This is due to tyrosine, phenylalanine and tryptophan. For DNA purity the ratio at A260:A280 is taken. The best purity is 1.8 to 2.PrecautionsAll the chemicals should be handled with care specially phenol; if skin comes in contact with phenol flush off with large amount of water and then apply polyethylene glycol, never apply ethanol. If using blood or any human tissue precaution should be taken that it is pre-tested and free from HIV. Further, always wear gloves.Nucleic acid blotting These techniques may be applied in the isolation and quantification of specific nucleic acid sequences and in the study of their organization, intracellular localization, expression and regulation. A variety of specific applications includes the diagnosis of infectious and inherited disease. Each of these topics is covered in depth in subsequent chapters. An overview of the steps involved in nucleic acid blotting and membrane hybridization procedures is shown in Fig. 2.4. Blotting describes the immobilization of sample nucleic acids on to a solid support, generally nylon or nitrocellulose membranes. The blotted nucleic acids are then used as ‘targets’ in subsequent hybridization experiments. The main blotting procedures are:Southern blotting

The original method of blotting was developed by Southern (1975, 1979b) for detecting fragments in an agarose gel that are complementary to a given RNA or DNA sequence. In this procedure, referred to as Southern blotting, the agarose gel is mounted on a filter-paper wick which dips into a reservoir containing transfer buffer (Fig. 2.5).

The hybridization membrane is sandwiched between the gel and a stack of paper towels (or other absorbent material), which serves to draw the transfer buffer through the gel by capillary action. The DNA molecules are carried out of the gel by the buffer flow and immobilized on the membrane. Initially, the membrane material used was nitrocellulose. The main drawback with this membrane is its fragile nature.

Supported nylon membranes have since been developed which have greater binding capacity for nucleic acids in addition to high tensile strength. For efficient Southern blotting, gel pretreatment is important. Large DNA fragments (10 kb) require a longer transfer time than short fragments.

To allow uniform transfer of a wide range of DNA fragment sizes, the electrophoresed DNA is exposed to a short depurination treatment (0.25 mol/l HCl) followed by alkali. This shortens the DNA fragments by alkaline hydrolysis at depurinated sites.

It also denatures the fragments prior to transfer, ensuring that they are in the single-stranded state and accessible for probing. Finally, the gel is equilibrated in neutralizing solution prior to blotting. An alternative method uses positively charged nylon membranes, which remove the need for extended gel pretreatment.

With them the DNA is transferred in native (non-denatured) form and then alkali-denatured in situ on the membrane. After transfer, the nucleic acid needs to be fixed to the membrane and a number of methods are available.

Oven baking at 80°C is the recommended method for nitrocellulose membranes and this can also be used with nylon membranes. Due to the flammable nature of nitrocellulose, it is important that it is baked in a vacuum oven.

An alternative fixation method utilizes ultraviolet cross-linking. It is based on the formation of cross-links between a small fraction of the thymine residues in the DNA and positively charged amino groups on the surface of nylon membranes. A calibration experiment must be performed to determine the optimal fixation period.

Following the fixation step, the membrane is placed in a solution of labelled (radioactive or non-radioactive) RNA, single-stranded DNA or oligodeoxynucleotide which is complementary in sequence to the blottransferred DNA band or bands to be detected. Conditions are chosen so that the labelled nucleic acid hybridizes with the DNA on the membrane.

Since this labelled nucleic acid is used to detect and locate the complementary sequence, it is called the probe. Conditions are chosen which maximize the rate of hybridization, compatible with a low background of non-specific binding on the membrane .

After the hybridization reaction has been carried out, the membrane is washed to remove unbound radioactivity and regions of hybridization are detected autoradiographically by placing the membrane in contact with X-ray film

Northern blotting Southern’s technique has been of enormous value,but it was thought that it could not be

applied directly to the blot-transfer of RNAs separated by gel electrophoresis, since RNA was found not to bind to nitrocellulose.

Alwine et al. (1979) therefore devised a procedure in which RNA bands are blot-transferred from the gel on to chemically reactive paper, where they are bound covalently.

The reactive paper is prepared by diazotization of aminobenzyloxymethyl paper (creating diazobenzyloxymethyl (DBM) paper), which itself can be prepared from Whatman 540 paper by a series of uncomplicated reactions.

Once covalently bound, the RNA is available for hybridization with radiolabelled DNA probes. As before, hybridizing bands are located by autoradiography.

Alwine et al.’s method thus extends that of Southern and for this reason it has acquired the jargon term northern blotting. Subsequently it was found that RNA bands can indeed be blotted on to nitrocellulose membranes under appropriate conditions (Thomas 1980) and suitable nylon membranes have been developed. Because of the convenience of these more recent methods, which do not require freshly activated paper, the use of DBM paper has been superseded.

Western blotting The term ‘western’ blotting (Burnette 1981) refers to a procedure which does not directly

involve nucleic acids, but which is of importance in gene manipulation.It involves the transfer of electrophoresed protein bands from a polyacrylamide gel on to a membrane of nitrocellulose or nylon, to which they bind strongly (Gershoni & Palade 1982, Renart & Sandoval 1984).

The bound proteins are then available for analysis by a variety of specific protein–ligand interactions. Most commonly, antibodies are used to detect specific antigens. Lectins have been used to identify glycoproteins. In these cases the probe may itself be labelled with radioactivity, or some other ‘tag’ may be employed. Often, however, the probe is unlabelled and is itself detected in a ‘sandwich’ reaction, using a second molecule which is labelled, for instance a species-specific second antibody, or protein A of Staphylococcus aureus (which binds to certain subclasses of IgG antibodies), or streptavidin (which binds to antibody probes that have been biotinylated).

These second molecules may be labelled in a variety of ways with radioactive, enzyme or fluorescent tags. An advantage of the sandwich approach is that a single preparation of labelled second molecule can be employed as a general detector for different probes. For example, an antiserum may be raised in rabbits which reacts with a range of mouse immunoglobins.

Such a rabbit anti-mouse (RAM) antiserum may be radiolabelled and used in a number of different applications

to identify polypeptide bands probed with different, specific, monoclonal antibodies, each monoclonal antibody being of mouse origin. The sandwich method may also give a substantial increase in sensitivity, owing to the multivalent binding of antibody molecules.

Chromosome walkingEarlier in this chapter, we discussed the advantages of making genomic libraries from random DNA fragments. One of these advantages is that the resulting fragments overlap, which allows genes to be cloned by chromosome walking. The principle of chromosome walking is that overlapping clones will hybridize to each other, allowing them to be assembled into a contiguous sequence. This can be used to isolate genes whose function is unknown but whose genetic location is known, a technique known as positional cloning.To begin a chromosome walk, it is necessary to have in hand a genomic clone that is known to lie very close to the suspected location of the target gene. In humans, for example, this could be a restriction fragment length polymorphism that has been genetically mapped to the same region. This clone is then used to screen a genomic library by hybridization, which should reveal any overlapping clones.These overlapping clones are then isolated, labeled and used in a second round of screening to identify further overlapping clones, and the process is repeated to build up a contiguous map. If the same library is used for each round of screening, previously identified clones can be distinguished from new ones, so that walking back and forth along the same section of DNA is prevented. Furthermore, modern vectors, such as DASH and FIX, allow probes to be generated from the end-points of a given genomic clone by in vitro transcription (see Fig. 6.4), which makes it possible to walk specifically in one direction.In Drosophila, the progress of a walk can also be monitored by using such probes for in situ hybridization against polytene chromosomes. Monitoring is necessary due to the dangers posed by repetitive DNA. Certain DNA sequences are highly repetitive and are dispersed throughout the genome. Hybridization with such a sequence could disrupt the orderly progress of a walk, in the worst cases causing a ‘warp’ to another chromosome. The probe used for stepping from one genomic clone to the next must be a unique sequence clone, or a subclone that has been shown to contain only a unique sequence.Chromosome walking is simple in principle, but technically demanding. For large distances, it is advisable to use libraries based on high-capacity vectors, such as BACs and YACs, to reduce the number of steps involved. Before such libraries were available, some ingenious strategies were used to reduce the number of steps needed in a walk. In one of the first applications of this technology, Hogness and his co-workers (Bender et al. 1983) cloned DNA from the Ace and rosy loci and the homoerotic Bithorax gene complex in Drosophila. The number of steps was minimized by exploiting the numerous strains carrying well-characterized inversions andtranslocations of specific chromosome regions. A different strategy, called chromosome jumping, has been used for human DNA (Collins & Weissman 1984, Poustka & Lehrach 1986). This involves

the circularization of very large genomic fragments generated by digestion with endonucleases, such as NotI, which cut at very rare target sites. This is followed by subcloning of the region covering the closure of the fragment, thus bringing together sequences that were located a considerable distance apart. In this way a jumping library is constructed, which can be used for long-distance chromosome walks (Collins et al. 1987, Richards et al. 1988). The application of chromosome walking and jumping to the cloning of the human cystic fibrosis gene.

UNIT IIEnzymes – Nucleases: Restriction endonucleases – DNA cloning – Hybrid vectors –Restriction cloning – selection for hybrid vectores – Methods of cloning – Synthesis of cDNA – Clonning from genomic DNA – Genomic libraries – Selection and screening methods.

Endonucleases are enzymes that cleave the phosphodiester bond within a polynucleotide chain, in contrast to exonucleases, which cleave phosphodiester bonds at the end of a polynucleotide chain. Restriction endonucleases (Restriction enzymes) cleave DNA at specific sites, and are divided into three categories, Type I, Type II, and Type III, according to their mechanism of action. These enzymes are often used in genetic engineering to make recombinant DNA for introduction into bacterial, plant, or animal cells.

Restriction endonucleases are enzymes that cleave the sugar-phosphate backbone of DNA. In most practical settings, a given enzyme cuts both strands of duplex DNA within a stretch of just a few bases. Several thousand different restriction endonucleases have been isolated, which collectively exhibit a few hundred different sequence (substrate) specificities.

Nomenclature and Classification

Restriction enzymes are named based on the organism in which they were discovered. For example, the enzyme Hind III was isolated from Haemophilus influenzae, strain Rd. The first three letters of the name are italicized because they abbreviate the genus and species names of the organism. The fourth letter typically comes from the bacterial strain

designation. The Roman numerals are used to identify specific enzymes from bacteria that contain multiple restriction enzymes. Typically, the Roman numeral indicates the order in which restriction enzymes were discovered in a particular strain.

There are three classes of restriction enzymes, labeled types I, II, and III. Type I restriction systems consist of a single enzyme that performs both modification (methylation) and restriction activities. These enzymes recognize specific DNA sequences, but cleave the DNA strand randomly, at least 1,000 base pairs (bp) away from the recognition site. Type III restriction systems have separate enzymes for restriction and methylation, but these enzymes share a common subunit. These enzymes recognize specific DNA sequences, but cleave DNA at random sequences approximately twenty-five bp from the recognition sequence. Neither type I nor type III restriction systems have found much application in recombinant DNA techniques.

Type II restriction enzymes, in contrast, are heavily used in recombinant DNA techniques. Type II enzymes consist of single, separate proteins for restriction and modification. One enzyme recognizes and cuts DNA, the other enzyme recognizes and methylates the DNA. Type II restriction enzymes cleave the DNA sequence at the same site at which they recognize it. The only exception are type IIs (shifted) restriction enzymes, which cleave DNA on one side of the recognition sequence, within twenty nucleotides of the recognition site. Type II restriction enzymes discovered to date collectively recognize over 200 different DNA sequences.

Type II restriction enzymes can cleave DNA in one of three possible ways. In one case, these enzymes cleave both DNA strands in the middle of a recognition sequence, generating blunt ends. For example: (The notations 5′ and 3′ are used to indicate the orientation of a DNA molecule. The numbers 5 and 3 refer to specific carbon atoms in the deoxyribose sugar in DNA.)

These blunt ended fragments can be joined to any other DNA fragment with blunt ends, making these enzymes useful for certain types of DNA cloning experiments.

Type II restriction enzymes can also cleave DNA to leave a 3′ ("three prime") overhang. (An overhang means that the restriction enzyme leaves a short single-stranded "tail" of DNA at the site where the DNA was cut.) These 3′ overhanging ends can only join to another compatible 3′ overhanging end (that is, an end with the same sequence in the overhang). Finally, some type II enzymes can generate 5′ overhanging DNA ends, which can only be joined to a compatible 5′ end.

In the type II restriction enzymes discovered to date, the recognition sequences range

from 4 bp to 9 bp long. Cleavage will not occur unless the full length of the recognition sequence is encountered. Enzymes with a short recognition sequence cut DNA frequently; restriction enzymes with 8 or 9 bp sequences typically cut DNA very infrequently, because these longer sequences are less common in the target DNA.

1. Explain about the Restriction-Modification Systems and Recognition Sequences.

A large majority of restriction enzymes have been isolated from bacteria, where they appear to serve a host-defense role.

The idea is that foreign DNA, for example from an infecting virus, will be chopped up and inactivated ("restricted") within the bacterium by the restriction enzyme.

The presence of restriction enzymes immediately begs the question of why they do not chew up the genomic DNA of their host. In almost all cases, a bacterium that makes a particular restriction endonuclease also synthesizes a companion DNA methyltransferase, which methylates the DNA target sequence for that restriction enzyme, thereby protecting it from cleavage.

This combination of restriction endonuclease and methylase is referred to as a restriction-modification system.

By convention, restriction enzymes are named after their host of origin.

For example, EcoRI was isolated from Escherichia coli (strain RY13), Hind II and Hind III from Haemophilus influenzae, and XhoI from Xanthomonas holcicola.

Restriction Enzyme Recognition Sequences

The substrates for restriction enzymes are more-or-less specific sequences of double-stranded DNA called recognition sequences. Examining the following table will illustrate some important points (recognition sites are shown as double stranded DNA).

The length of restriction recognition sites varies: The enzymes EcoRI, SacI and SstI each recognize a 6 base-pair (bp) sequence of DNA, whereas NotI recognizes a sequence 8 bp in length, and the recognition site for Sau3AI is only 4 bp in length. Length of the recognition sequence dictates how frequently the enzyme will cut in a random sequence of DNA. Enzymes with a 6 bp recognition site will cut, on average, every 46 or 4096 bp; a 4 bp recognition site will occur roughly every 256 bp.

Different restriction enzymes can have the same recognition site - such enzymes are called isoschizomers: Look at the recognition sites for SacI and SstI - they are identical. In some cases isoschizomers cut identically within their recognition site, but sometimes they do not. Isoschizomers often have different optimum reaction conditions, stabilities and costs, which may influence the decision of which to purchase.

Restriction recognitions sites can be unambiguous or ambiguous: The enzyme BamHI recognizes the sequence GGATCC and no others - this is what is meant by unambiguous. In contrast, HinfI recognizes a 5 bp sequence starting with GA, ending in TC, and having any base between (in the table, "N" stands for any nucleotide) - HinfI has an ambiguous recognition site. XhoII also has an ambiguous recognition site: Py stands for pyrimidine (T or C) and Pu for purine (A or G), so XhoII will recognize and cut sequences of AGATCT, AGATCC, GGATCT and GGATCC.

The recognition site for one enzyme may contain the restriction site for another: For example, note that a BamHI recognition site contains the recognition site for Sau3AI. Consequently, all BamHI sites will cut with Sau3AI. Similarly, one of the four possible XhoII sites will also be a recognition site for BamHI and all four will cut with Sau3AI.

One other point to notice from the table above is that most recognition sequences are palindromes - they read the same forward (5' to 3' on the top strand) and backward (5' to 3' on the bottom strand). Most, but certainly not all recognition sites for commonly-used restriction enzymes are palindromes. Most restriction enzymes bind to their recognition site as dimers (pairs), as depicted for the enzyme PvuII in the figure to the right.

2. What are the Patterns of DNA Cutting by Restriction Enzymes?

Restriction enzymes hydrolyze the backbone of DNA between deoxyribose and phosphate groups. This leaves a phosphate group on the 5' ends and a hydroxyl on the 3' ends of both strands. A few restriction enzymes will cleave single stranded DNA, although usually at low efficiency.

The restriction enzymes most used in molecular biology labs cut within their recognition sites and generate one of three different types of ends. In the diagrams below, the recognition site is boxed in yellow and the cut sites indicated by red triangles.

5' overhangs: The enzyme cuts asymmetrically within the recognition site such that a short single-stranded segment extends from the 5' ends. BamHI cuts in this manner.

3' overhangs: Again, we see asymmetrical cutting within the recognition site, but the result is a single-stranded overhang from the two 3' ends. KpnI cuts in this manner.

Blunts: Enzymes that cut at precisely opposite sites in the two strands of DNA

generate blunt ends without overhangs. SmaI is an example of an enzyme that generates blunt ends.

The 5' or 3' overhangs generated by enzymes that cut asymmetrically are called sticky ends or cohesive ends, because they will readily stick or anneal with their partner by base pairing.

3. What is Exonuclease? And its mode of action.

Exonucleases are enzymes that work by cleaving nucleotides one at a time from the end of a polynucleotide chain. A hydrolyzing reaction that breaks phosphodiester bonds at either the 3’ or the 5’ end occurs. Its close relative is the endonuclease, which cleaves phosphodiester bonds in the middle of a polynucleotide chain. Eukaryotes and prokaryotes have three types of exonucleases involved in the normal turnover of mRNA: 5’ to 3’ exonuclease, which is a dependent decapping protein, 3’ to 5’ exonuclease, an independent protein, and poly(A)-specific 3’ to 5’ exonuclease.

In both archaebacteria and eukaryotes, one of the main routes of RNA degradation is performed by the multi-protein exosome complex, which consists largely of 3' to 5' exoribonucleases.

Significance to polymerase

RNA polymerase II is known to be in effect during transcriptional termination; it works with a 5’ exonuclease (human gene Xrn2) to degrade the newly formed transcript downstream, leaving the polyadenylation site and simultaneously shooting the polymerase. This process involves the exonuclease's catching up to the pol II and terminating the transcription.

Pol I then synthesizes DNA nucleotides in place of the RNA primer it had just removed. DNA polymerase I also has 5' to 3' exonuclease activity, which is used in editing and proofreading DNA for errors.

E. coli types

WRN Exonuclease with active sites in yellow

In 1971, Lehman IR discovered exonuclease I in E. coli. Since that time, there have been numerous discoveries including: exonuclease, II, III, IV, V, VI, VII, and VIII. Each type of exonuclease has a specific type of function or requirement.

Exonuclease I breaks apart single-stranded DNA in a 3'=>5' direction, releasing deoxyribonucleoside 5'-monophosphates one after another. It does not cleave DNA strands with terminal 3'-OH groups because they are blocked by phosphoryl or acetyl groups.

Exonuclease II is associated with DNA polymerase I, which contains a 5' exonuclease that clips off the RNA primer contained immediately upstream from the site of DNA synthesis in a 5' → 3' manner.

Exonuclease III has four catalytic activities:

3’ to 5’ exodeoxyribonuclease activity, which is specific for double-stranded DNA RNase activity

3’ phosphate activity

AP endonuclease activity (later found to be called endonuclease II).

Exonuclease IV adds a water molecule, so it can break the bond of an oligonucleotide to nucleoside 5’ monophosphate. This exonuclease requires Mg 2+ in order to function and works at higher temperatures then exonuclease I.

Exonuclease V is a 3’ to 5’ hydrolyzing enzyme that catalyzes linear double-stranded DNA and single-stranded DNA, which requires Ca2+.

Exonuclease VIII is 5’ to 3’ dimeric protein that does not require ATP or any gaps or nicks in the strand, but requires a free 5’ OH group to carry out its function.

Discoveries in humans

The 3’ to 5’ human type endonuclease is known to be essential for the proper processing of histone pre-mRNA, in which U7 snRNP directs the single cleavage process. Following the removal of the downstream cleavage product (DCP) 5’ to 3’ exonuclease continues to further breakdown the product until it is completely degraded. This allows the nucleotides to be recycled. 5’ To 3’ exonuclease is linked to a co-transcriptional cleavage (CoTC) activity that acts as a precursor to develop a free 5’ unprotected end, so the exonuclease can remove and degrade the downstream cleavage product (DCP). This initiates transcriptional termination because one does not want DNA or RNA strands building up in their bodies.

Discoveries in yeast

CCR4-NOT is a general transcription regulatory complex in yeast that is found to be associated with mRNA metabolism, transcription initiation, and mRNA degradation. CCR4 has been found to contain RNA and single-stranded DNA 3’to 5’ exonuclease activities. Another component associated with the CCR4 complex is CAF1 protein, which has been found to contain 3’to 5’ or 5’ to 3’ exonuclease domains in Mus musculus and Caenorhabditis elegans. This protein has not been found in yeast, which suggests that it is likely to have an abnormal exonuclease domain like the one seen in a metazoan. Yeast contains Rat1 and Xrn1 exonuclease. The Rat1 works just like the human type (Xrn2) and Xrn1 function in the cytoplasm is in the 5’ to 3’ direction to degrade RNAs (pre-5.8s and 25s rRNAs) in the absence of Rat1.

Hybride vector

1. Explain about the Phagemid.

A phagemid or phasmid is a type of cloning vector developed as a hybrid of the filamentous phage M13 and plasmids to produce a vector that can grow as a plasmid, and also be packaged as single stranded DNA in viral particles.

Phagemids contain an origin of replication (ori) for double stranded replication, as well as an f1 ori to enable single stranded replication and packaging into phage particles.

Many commonly used plasmids contain an f1 ori and are thus phagemids. Similarly to a plasmid, a phagemid can be used to clone DNA fragments and be introduced into a

 

bacterial host by a range of techniques (transformation, electroporation).

However, infection of a bacterial host containing a phagemid with a 'helper' phage, for example VCSM13 or M13K07, provides the necessary viral components to enable single stranded DNA replication and packaging of the phagemid DNA into phage particles.

These are secreted through the cell wall and released into the medium. Filamentous phage retard bacterial growth but, in contrast to lambda and T7 phage, are not generally lytic.

Helper phage are usually engineered to package less efficiently than the phagemid so that the resultant phage particles contain predominantly phagemid DNA.

F1 Filamentous phage infection requires the presence of a pilus so only bacterial hosts containing the F-plasmid or its derivatives can be used to generate phage particles.

Prior to the development of cycle sequencing, phagemids were used to generate single stranded DNA template for sequencing purposes.

Today phagemids are still useful for generating templates for site-directed mutagenesis. Detailed characterisation of the filamentous phage life cycle and structural features lead to the development of phage display technology, in which a range of peptides and proteins can be expressed as fusions to phage coat proteins and displayed on the viral surface.

The displayed peptides and polypeptides are associated with the corresponding coding DNA within the phage particle and so this technique lends itself to the study of protein-protein interactions and other ligand/receptor combinations.

[hide]v · d · eTypes of nucleic acids

Constituents Nucleobases · Nucleosides · Nucleotides · Deoxynucleotides

Ribonucleic acids(coding and non-coding)

translation: mRNA (pre-mRNA/hnRNA) · tRNA · rRNA · tmRNA

regulatory: miRNA · siRNA · piRNA · aRNA

RNA processing: snRNA · snoRNA

other/ungrouped: gRNA · shRNA · stRNA · ta-siRNA

Deoxyribonucleic acids cDNA · cpDNA · gDNA · msDNA · mtDNA

Nucleic acid analogues GNA · LNA · BNA · PNA · TNA · morpholino

Cloning vectorsphagemid · plasmid · lambda phage · cosmid · fosmid · PAC · BAC · YAC · HAC

 

2. What is Cosmid Vectors?and its features.

They have been developed in the late 1970s and have been improved significantly since. (Basic features of a cosmid).

They are predominantly plasmids with a bacterial oriV, an antibiotic selection marker and a cloning site, but they carry one, or more recently two cos sites derived from bacteriophage lambda.

Depending on the particular aim of the experiment broad host range cosmids, shuttle cosmids or 'mammalian' cosmids (linked to SV40 oriV and mammalian selection markers) are available.

The loading capacity of cosmids varies depending on the size of the vector itself but usually lies around 40-45 kb.

The cloning procedure involves the generation of two vector arms which are then joined to the foreign DNA. Selection against wildtype cosmid DNA is simply done via size exclusion! Remember however that cosmids always form colonies and not plaques! Also clone density is much lower with around 105 - 106 cfu per ug of ligated DNA.

After the construction of recombinant lambda or cosmid libraries the total DNA is transfered into an appropriate E.coli host via a technique called in vitro packaging.

The necessary packaging extracts are derived from E.coli cI857 lysogens (red- gam- Sam and Dam (head assembly) and Eam (tail assembly) respectively).

These extracts will recognize and package the recombinant molecules in vitro, generating either mature phage particles (lambda-based vectors) or recombinant plasmids contained in phage shells (cosmids).

These differences are reflected in the different infection frequencies seen in favour of lambda-replacement vectors. This compensates for their slightly lower loading capacity. Phage library are also stored and screened easier than cosmid (colonies!) libraries.

Target DNA: the genomic DNA to be cloned has to be cut into the appropriate size range of restriction fragments.

This is usually done by partial restriction followed by either size fractionation or dephosphorylation (using calf-intestine phosphatase ) in order to avoid chromosome scrambling, ie the ligation of physically unlinked fragments.

Cosmid features and uses

Cosmids are predominantly plasmids with a bacterial oriV, an antibiotic selection marker and a cloning site, but they carry one, or more recently two cos sites derived from bacteriophage lambda.

Depending on the particular aim of the experiment broad host range cosmids, shuttle cosmids or 'mammalian' cosmids (linked to SV40 oriV and mammalian selection markers) are available.

The loading capacity of cosmids varies depending on the size of the vector itself but usually lies around 40–45 kb.

The cloning procedure involves the generation of two vector arms which are then joined to the foreign DNA. Selection against wildtype cosmid DNA is simply done via size exclusion. Cosmids, however, always form colonies and not plaques. Also the clone density is much lower with around 105 - 106 CFU per µg of ligated DNA.

After the construction of recombinant lambda or cosmid libraries the total DNA is transferred into an appropriate E.coli host via a technique called in vitro packaging.

The necessary packaging extracts are derived from E.coli cI857 lysogens (red- gam- Sam and Dam (head assembly) and Eam (tail assembly) respectively).

These extracts will recognize and package the recombinant molecules in vitro, generating either mature phage particles (lambda-based vectors) or recombinant plasmids contained in phage shells (cosmids). These differences are reflected in the different infection frequencies seen in favour of lambda-replacement vectors.

This compensates for their slightly lower loading capacity. Phage library are also stored and screened easier than cosmid (colonies!) libraries.

Target DNA: the genomic DNA to be cloned has to be cut into the appropriate size range of restriction fragments.

This is usually done by partial restriction followed by either size fractionation or dephosphorylation (using calf-intestine phosphatase) to avoid chromosome scrambling, i.e. the ligation of physically unlinked fragments.

The cDNA synthesis reactionThe synthesis of double-stranded cDNA suitable for insertion into a cloning vector involves three major steps: (i) first-strand DNA synthesis on the mRNA template, carried out with a reverse transcriptase; (ii) removal of the RNA template; and (iii) secondstrand DNA synthesis using the first DNA strand as a template, carried out with a DNA-dependent DNApolymerase, such as E. coli DNA polymerase I. All DNA polymerases, whether they use RNA or DNA as the template, require a primer to initiate strand synthesis.

Development of cDNA cloning strategiesThe first reports of cDNA cloning were published in the mid-1970s and were all based on the homopolymer tailing technique, Of several alternative methods, the onethat became the most popular was that of Maniatis et al. (1976). This involved the use of an oligo-dT primer annealing at the polyadenylate tail of the mRNA to prime first-strand cDNA synthesis, and took advantage of the fact that the first cDNA strand has the tendency to transiently fold back on itself, forming a hairpin loop, resulting in self-priming of the second strand (Efstratiadis et al. 1976). After the synthesis of the second DNA strand, this loop must be cleaved with a single-strand-specific nuclease,e.g. S1 nuclease, to allow insertion into the cloning vector (Fig. 6.5). A serious disadvantage of the hairpin method isthat cleavage with S1 nuclease results in the loss of a certain amount of sequence at the 5′end of the clone. This strategy has therefore been superseded by other methods in which the second strand is primed in a separate reaction. One of the simplest strategies (Land et al. 1981). After first-strand synthesis, which is primed with an oligo-dT primer as usual, the cDNA is tailed with a string of cytidine residues using the enzyme terminal transferase. This artificial oligo-dC tail is then used as an annealing site for a synthetic oligo-dG primer, allowing synthesis of the second strand.Using this method, Land et al. (1981) were able to isolate a full-length cDNA corresponding to the chicken lysozyme gene. However, the efficiency can be lower for other cDNAs (e.g. Cooke et al. 1980). For cDNA expression libraries, it is advantageous if the cDNA can be inserted into the vector in the correct orientation. With the self-priming method,this can be achieved by adding a synthetic linker to the double-stranded cDNA molecule before the hairpin loop is cleaved (e.g. Kurtz & Nicodemus 1981; Fig. 6.7a). Where the second strand is primed separately, direction cloning can be achieved using an oligo-dT primer containing a linker sequence (e.g. Coleclough & Erlitz 1985; Fig. 6.7b). An alternative

is to use primers for cDNA synthesis that are already linked to a plasmid (Fig. 6.7c). This strategy was devised by Okayama and Berg (1982) and has two further notable characteristics. First, full-length cDNAs are preferentially obtained because an RNA– DNA hybrid molecule, the result of first-strand synthesis, is the substrate for a terminal transferasereaction. A cDNA that does not extend to the end of the mRNA will present a shielded 3-hydroxyl group, which is a poor substrate for tailing. Secondly, the second-strand synthesis step is primed by nicking the RNA at multiple sites with RNase H. Second-strand synthesis therefore occurs by a nick-translation type of reaction, which is highly efficient. SimplercDNA cloning strategies incorporating replacement synthesis of the second strand are widely used (e.g. Gubler & Hoffman 1983, Lapeyre & Amalric 1985). The Gubler–Hoffman reaction.

Genomic Library A genomic library is a population of host bacteria, each of which carries a DNA

molecule that was inserted into a cloning vector, such that the collection of cloned DNA molecules represents the entire genome of the source organism. This term also represents the collection of all of the vector molecules, each carrying a piece of the chromosomal DNA of the organism, prior to the insertion of these molecules into the host cells

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 109 bp long while a plamid or l phage vector may carry up to 20 kb fragment.  This would require 1.5 x 105 recombinant plasmids or l 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 104 l 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 l phage vector is that its transformation efficiency is about 1000 times higher than the plasmid vector.

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. 

Figure 9-B-1.  Preparation of the genomic library using l phage vectors.  It is basically the cloning of all DNA fragments representing the entire genome

cDNA Library

The advantage of cDNA library is that it contains only the coding region of a genome.  To prepare a cDNA library, the first step is to isolate the total mRNA from the cell type of interest.  Because eukaryotic mRNAs consist of a poly-A tail, they can easily be separated.  Then the enzyme reverse transcriptase is used to synthesize a DNA strand complementary to each mRNA mlecule.  After the single-stranded DNA molecules are converted into double-stranded DNA molecules by DNA polymerase, they are inserted into vectors and cloned.

Probes 

A probe is a piece of DNA or RNA used to detect specific nucleic acid sequences by hybridization (binding of two nucleic acid chains by base pairing) .  They are radioactively labeled so that the hybridized nucleic acid can be identified by autoradiography.

The size of probes ranges from a few nucleotides to hundreds of kilobases.  Long probes are usually made by cloning.  Originally they may be double-stranded, but the working probes must be single-stranded.  Short probes (oligonucleotide probes) can be made by chemical synthesis. They are single-stranded.

Suppose we have cloned a specific gene in yeast and want to find its homologous gene in human, then we may use the specific yeast gene as a probe to detect its homologous gene from the human genomic library.  On the other hand, if we know the conserved sequence in the specific gene between yeast and human, we may use oligonucleotide probes containing only the conserved sequence.  Typically, an oligonucleotide about 20 nucleotides long is sufficient to screen a library.

In some cases, we have known the partial sequence of a protein and want to detect its gene in the library.  Then we may synthesize oligonucleotide probes based on the known peptide sequence.  Since an amino acid may be encoded by several DNA triplets, many different oligonucleotide probes are often needed.  

 

Figure 9-B-2.  The relationship between a peptide and all possible DNA sequences.  In this example, the peptide sequence Leu-Phe-Tyr-Met-His-Asp corresponds to 96 (= 6 x 2 x 2 x 1 x 2 x 2) possible DNA sequences.

 

Screening

Once a particular DNA fragment is identified, it can be isolated and amplified to determine its sequence.  If we know the partial sequence of a gene and want to determine its entire sequence, the probe should contain the known sequence so that the detected DNA fragment may contain the gene of interest.  

Figure 9-B-3.  Screening of a specific DNA fragment.  After recombinant l 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 l 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.

UNIT IIIBiology of genetic engineering – Plasmids used for E,coli vectors, based onbacteriophage and M-13 phage vectors. Eukaryotic vectors – Yeast vectors, animal vectors, plant vectors. Prokaryotic and Eukaryotic hosts.

1. Give general an account on prokaryotic vector. A plasmid is a DNA molecule that is separate from, and can replicate independently of,

the chromosomal DNA They are double-stranded and, in many cases, circular.

Plasmids usually occur naturally in bacteria, but are sometimes found in eukaryotic organisms (e.g., the 2-micrometre-ring in Saccharomyces cerevisiae).

Plasmid sizes vary from 1 to over 1,000 kilobase pairs (kbp). The number of identical plasmids in a single cell can range anywhere from one to even thousands under some circumstances. Plasmids can be considered part of the mobilome because they are often associated with conjugation, a mechanism of horizontal gene transfer.

The term plasmid was first introduced by the American molecular biologist Joshua Lederberg in 1952.

Plasmids are considered transferable genetic elements, or "replicons", capable of autonomous replication within a suitable host.

Plasmids can be found in all three major domains: Archea, Bacteria and Eukarya. Similar to viruses, plasmids are not considered a form of "life" as it is currently defined. Unlike viruses, plasmids are "naked" DNA and do not encode genes necessary to encase the genetic material for transfer to a new host, though some classes of plasmids encode the sex pilus necessary for their own transfer.

Plasmid host-to-host transfer requires direct, mechanical transfer by conjugation or changes in host gene expression allowing the intentional uptake of the genetic element by transformation.

Microbial transformation with plasmid DNA is neither parasitic nor symbiotic in nature, because each implies the presence of an independent species living in a commensal or detrimental state with the host organism. Rather, plasmids provide a mechanism for horizontal gene transfer within a population of microbes and typically provide a selective advantage under a given environmental state.

Plasmids may carry genes that provide resistance to naturally occurring antibiotics in a competitive environmental niche, or alternatively the proteins produced may act as toxins under similar circumstances.

Plasmids also can provide bacteria with an ability to fix elemental nitrogen or to degrade recalcitrant organic compounds which provide an advantage when nutrients are scarce.

There are two types of plasmid integration into a host bacteria: Non-integrating plasmids replicate as with the top instance; whereas episomes, the lower example, integrate into the host chromosome.

Plasmids used in genetic engineering are called vectors. Plasmids serve as important tools in genetics and biotechnology labs, where they are commonly used to multiply (make many copies of) or express particular genes.

Many plasmids are commercially available for such uses. The gene to be replicated is inserted into copies of a plasmid containing genes that make cells resistant to particular antibiotics and a multiple cloning site (MCS, or polylinker), which is a short region containing several commonly used restriction sites allowing the easy insertion of DNA fragments at this location.

Next, the plasmids are inserted into bacteria by a process called transformation. Then, the bacteria are exposed to the particular antibiotics. Only bacteria which take up copies of the plasmid survive, since the plasmid makes them resistant.

In particular, the protecting genes are expressed (used to make a protein) and the expressed protein breaks down the antibiotics.

In this way the antibiotics act as a filter to select only the modified bacteria. Now these bacteria can be grown in large amounts, harvested and lysed (often using the alkaline lysis method) to isolate the plasmid of interest.

Another major use of plasmids is to make large amounts of proteins. In this case, researchers grow bacteria containing a plasmid harboring the gene of interest.

Just as the bacteria produces proteins to confer its antibiotic resistance, it can also be induced to produce large amounts of proteins from the inserted gene.

This is a cheap and easy way of mass-producing a gene or the protein it then codes for, for example, insulin or even antibiotics.

However, a plasmid can only contain inserts of about 1–10 kbp. To clone longer lengths of DNA, lambda phage with lysogeny genes deleted, cosmids, bacterial artificial chromosomes or yeast artificial chromosomes could be used.

2. What are the Applications of vector?

Disease Models

Plasmids were historically used to genetically engineer the embryonic stem cells of rats in order to create rat genetic disease models.

The limited efficiency of plasmid based techniques precluded their use in the creation of more accurate human cell models.

Fortunately, developments in Adeno-associated virus recombination techniques, and Zinc finger nucleases, have enabled the creation of a new generation of isogenic human disease models.

Gene therapy

The success of some strategies of gene therapy depends on the efficient insertion of therapeutic genes at the appropriate chromosomal target sites within the human genome, without causing cell injury, oncogenic mutations (cancer) or an immune response.

Plasmid vectors are one of many approaches that could be used for this purpose. Zinc finger nucleases (ZFNs) offer a way to cause a site-specific double strand break to the DNA genome and cause homologous recombination.

This makes targeted gene correction a possibility in human cells. Plasmids encoding ZFN could be used to deliver a therapeutic gene to a pre-selected chromosomal site with a frequency higher than that of random integration.

Although the practicality of this approach to gene therapy has yet to be proven, some aspects of it could be less problematic than the alternative viral-based delivery of therapeutic genes.

3. Define Episomes.and its role in the genetics.

An episome is a portion of genetic material that can exist independent of the main body of genetic material (called the chromosome) at some times, while at other times is able to integrate into the chromosome.

Examples of episomes include insertion sequences and transposons. Viruses are another example of an episome. Viruses that integrate their genetic material into the host chromosome enable the viral nucleic acid to be produced along with the host genetic material in a nondestructive manner. As an autonomous unit (i.e., existing outside of the chromosome) however, the viral episome destroys the host cell as it commandeers the host's replication apparatuses to make new copies of itself.

Another example of an episome is called the F factor. The F factor determines whether genetic material in the chromosome of one organism is transferred into another organism. The F factor can exist in three states that are designated as FPLUS, Hfr, and F prime.

FPLUS refers to the F factor that exists independently of the chromosome. Hfr stands for high frequency of recombination, and refers to a factor that has integrated into the host chromosome. The F prime factor exists outside the chromosome, but has a portion of chromosomal DNA attached to it.

An episome is distinguished from other pieces of DNA that are independent of the chromosome (i.e.,plasmids) by their large size.

Plasmids are different from episomes, as plasmid DNA cannot link up with chromosomal DNA. The plasmid carries all the information necessary for its independent replication. While not necessary for bacterial survival, plasmids can be advantageous to a bacterium. For example, plasmids can carry genes that confer resistance to antibiotics or toxic metals, genes that allow the bacterium to degrade compounds that it otherwise could not use as food, and even genes that allow the bacterium to infect an animal or plant cell. Such traits can be passed on to another bacterium.

Transposons and insertion sequences are episomes. These are also known as mobile genetic elements. They are capable of existing outside of the chromosome. They are also designed to integrate into the chromosome following their movement from one cell to another. Like plasmids, transposons can carry other genetic material with them, and so pass on resistance to the cells they enter. Class 1 transposons, for example, contain drug resistance genes. Insertion sequences do not carry extra genetic material. They code for only the functions involved in their insertion into chromosomal DNA.

Transposons and insertion sequences are useful tools to generate changes in the DNA sequence of host cells. These genetic changes that result from the integration and the exit of the mobile elements from DNA, are generically referred to as mutations. Analysis of the mobile element can determine what host DNA is present, and the analysis of the mutated host cell can determine whether the extra or missing DNA is important for the functioning of the cell.

4. What is Yeast Plasmid and its types?

Other types of plasmids are often related to yeast cloning vectors that include:

Yeast integrative plasmid (YIp), yeast vectors that rely on integration into the host chromosome for survival and replication, and are usually used when studying the functionality of a solo gene or when the gene is toxic. Also connected with the gene URA3, that codes an enzyme related to the biosynthesis of pyrimidine nucleotides (T, C);

Yeast Replicative Plasmid (YRp), which transport a sequence of chromosomal DNA that includes an origin of replication. These plasmids are less stable, as they can "get lost" during the budding.

5. Describe the pUC vector.

o Plasmids are circular, double-stranded DNA molecules that exist in bacteria and in the nuclei of some eukaryotic cells. 

o They can replicate independently of the host cell.  The size of plasmids ranges from a few kb to near 100 kb.

Figure 9-A-3.  A typical plasmid vector.  It contains a polylinker which can recognize several different restriction enzymes, an ampicillin-resistance gene (ampr) for selective amplification, and a replication origin (ORI) for proliferation in the host cell.  

A plasmid vector is made from natural plasmids by removing unnecessary segments and adding essential sequences. 

To clone a DNA sample, the same restriction enzyme must be used to cut both the vector and the DNA sample. 

Therefore, a vector usually contains a sequence (polylinker) which can recognize several restriction enzymes so that the vector can be used for cloning a variety of DNA samples.

A plasmid vector must also contain a drug-resistance gene for selective amplification.  After the vector enters into a host cell, it may proliferate with the host cell. 

However, since the transformation efficiency of plasmids in E. coli is very low, most E. coli cells that proliferate in the medium would not contain the plasmids. 

Therefore, we must find a way to allow only the transformed E. coli to proliferate.  Typically, antibiotics are used to kill E. coli cells which do not contain the vectors. 

The transformed E. coli cells are protected by the ampicillin-resistance gene (ampr) which can express the enzyme, lactamase, to inactivate the antibiotic ampicillin.

1. Write about the Lambda phage vector.

phages are viruses that can infect bacteria. 

The major advantage of the phage vector is its high transformation efficiency, about 1000 times more efficient than the plasmid vector.

Figure 9-A-4.  Schematic drawing of the DNA cloning using

phages as vectors.  The DNA to be cloned is first inserted into the DNA, replacing a nonessential region. 

hen, by an in vitro assembly system (described below), the virion carrying the recombinant DNA can be formed. 

The genome is 49 kb in length which can carry up to 25 kb foreign DNA.

Lambda Phage Vectors types. Plasmid vectors described in the previous section are often used for cloning DNA

segments of small size (upto 10 kilobases). However, while preparing a genomic library in a eukaryote, the cloned fragments should be large enough to contain a whole gene.

This will also allow cloning of the whole genome into a number, which will not be unreasonably large and therefore can be screened without serious difficulty

The above properties and other requirements of cloning whole genome In eukaryotes are fulfilled by the phage lambda and cosmid vectors, the former permitting cloning of segments upto 20-25kb long (kb = kilobases) and latter accommodating segments upto 45kb long. Phage lambda (A).

However, is easier and more efficient for making genomic and cDNA Libraries

(a) λgt10 and λgt11. λgt10 and λgt11 are modified lambda phages designed to clone cDNA fragments. The major difference between these two Vectors is that λgt11 is an expression vector, where inserted DNA is expressed as β galactosidase fusion protein.

λgt10 is a 43 kb double stranded DNA for cloning fragments that are only 7 kb in length. The insertion of DNA inactivates c+ (repressor) gene generating a cl- bacteriophage. Non recombinant λgt10 is cl+ and forms cloudy plaques on appropriate E. coli host, while recombinant cl- λgt10 forms clear plaques permitting screening of recombinant plaque

Further, in an E. coli strain carrying hf lA 150 mutation (high frequency lysogeny mutation) only cl- phage will form plaques, because cl+ will form lysogens (integrate with bacterial genome) and will not undergo lysis to form any plaques. Recombinant λgt10 plaques can thus be easily selected

λgt11 is a 43.7 kb double stranded A phage for cloning DNA segments, which are less than 6 kb in length (usually for cDNA). Foreign DNA can be expressed as β galactosidase fusion proteins. Recombinant λgt11 can be screened using either).

The recombinant λgt11 becomes gar, while non recombinant λgt11 remains gal+, so that an appropriate E. coli host, with recombinant phage (gar) will form white or clear colonies and that with non recombinant phage (gal+) will form blue colonies permitting screening in the presence of IPTG (inducer) and Xgal (substrate).

Section C

3. How did types the plasmid vector?

Overview of bacterial conjugation

One way of grouping plasmids is by their ability to transfer to other bacteria. Conjugative plasmids contain so-called tra-genes, which perform the complex process of conjugation, the transfer of plasmids to another bacterium .

Non-conjugative plasmids are incapable of initiating conjugation, hence they can only be transferred with the assistance of conjugative plasmids, by 'accident'.

An intermediate class of plasmids are mobilizable, and carry only a subset of the genes required for transfer. They can 'parasitize' a conjugative plasmid, transferring at high frequency only in its presence. Plasmids are now being used to manipulate DNA and may possibly be a tool for curing many diseases.

It is possible for plasmids of different types to coexist in a single cell. Several different plasmids have been found in E. coli. But related plasmids are often incompatible, in the sense that only one of them survives in the cell line, due to the regulation of vital plasmid functions. Therefore, plasmids can be assigned into compatibility groups.

Another way to classify plasmids is by function. There are five main classes:

Fertility-F-plasmids, which contain tra-genes. They are capable of conjugation (transfer of genetic material between bacteria which are touching).

Resistance-(R)plasmids, which contain genes that can build a resistance against antibiotics or poisons and help bacteria produce pili. Historically known as R-factors, before the nature of plasmids was understood.

Col-plasmids, which contain genes that code for (determine the production of) bacteriocins, proteins that can kill other bacteria.

Degradative plasmids, which enable the digestion of unusual substances, e.g., toluene or salicylic acid.

Virulence plasmids, which turn the bacterium into a pathogen (one that causes disease).

Plasmids can belong to more than one of these functional groups.

Plasmids that exist only as one or a few copies in each bacterium are, upon cell division, in danger of being lost in one of the segregating bacteria. Such single-copy plasmids have systems which attempt to actively distribute a copy to both daughter cells.

Some plasmids or microbial hosts include an addiction system or "postsegregational killing system (PSK)", such as the hok/sok (host killing/suppressor of killing) system of plasmid R1 in Escherichia coli.

This variant produces both a long-lived poison and a short-lived antidote. Several types of plasmid addiction systems (toxin/ antitoxin, metabolism-based, ORT systems) were described in the literature and used in biotechnical (fermentation) or biomedical (vaccine therapy) applications.

Daughter cells that retain a copy of the plasmid survive, while a daughter cell that fails to inherit the plasmid dies or suffers a reduced growth-rate because of the lingering poison from the parent cell. Finally, the overall productivity could be enhanced.

PLASMID VECTORS

Plasmids are:

Circular, autonomous molecules of DNA. Found naturally in most bacterial (and some other) species.

Size: 1.5 - 300 kilobases.

Function: carry non-essential (dispensable) genes, e.g. antibiotic resistance, toxin production.

But "cryptic" plasmids have no known function!

Plasmids can be conjugative or non-conjugative (conjugation is generally not required in GM).

Plasmids can be mobilizable or non-mobilizable (non-mobilizable plasmids are preferred as they are less likely to "escape" from host cells).

Plasmids can be relaxed (multiple copies per host cell) or stringent (1-3 copies per host cell).

For GM work we want: small, relaxed, non-conjugative, non-mobilizable plasmids with good markers and unique restriction sites.

THREE EXAMPLES OF NATURAL PLASMIDS

PLASMID

SIZE

(kb)

RELAXED

(AMPLIFIED)

SINGLE SITES FOR

RESTRICTION ENZYMES

MARKER GENES FOR SELECTING TRANSFORMAN

TS

ADDITIONAL MARKER

GENES SHOWING

INSERTIONAL INACTIVATIO

N

pSC101 6.5 NO XhoI, EcoRI,

PvuII, HincII

Tetracycline resistance

-

HindIII, BamHI, SalI

- Tetracycline resistance

ColE1 8.0 YES EcoRI Immunity to

colicin E1

Colicin E1 production

RSF2124 11.0 YES EcoRI, BamHI Ampicillin resistance

Vector" is an agent that can carry a DNA fragment into a host cell.  If it is used for reproducing the DNA fragment, it is called a "cloning vector".   If it is used for expressing certain gene in the DNA fragment, it is called an "expression vector".

Commonly used vectors include plasmid, Lambda phage, cosmid and yeast artificial chromosome (YAC).

 

4. Explain the PBR322.draw the diagram.

pBR322 is a plasmid and for a time was one of the most commonly used E. coli cloning vectors. Created in 1977, it was named eponymously after its Mexican creators, p standing for plasmid, and BR for Bolivar and Rodriguez.

pBR322 is 4361 base pairs in length and contains a replicon region (source plasmid pMB1), the ampR gene, encoding the ampicillin resistance protein (source plasmid RSF2124) and the tetR gene, encoding the tetracycline resistance protein (source plasmid pSC101).

The plasmid has unique restriction sites for more than forty restriction enzymes. 11 of these 40 sites lie within the tetR gene.

There are 2 sites for restriction enzymes HindIII and ClaI within the promoter of the tetR gene. There are 6 key restriction sites inside the ampR gene.

The origin of replication or ori site in this plasmid is pMB1 (a close relative of ColE1). The ori encodes two RNAs (RNAI and RNAII) and one protein (called Rom or Rop).

The circular sequence is numbered such that 0 is the middle of the unique EcoRI site and the count increases through the tet genes.

The ampicillin resistance gene is a penicillin beta-lactamase. Promoters P1 and P3 are for the beta-lactamase gene. P3 is the natural promoter, and P1 is artificially created by the ligation of two different DNA fragments to create pBR322.

P2 is in the same region as P1, but it is on the opposite strand and initiates transcription in the direction of the tetracycline resistance gene.

Bits of the pBR322 sequence were used to create the "dinosaur" DNA in the Novel Jurassic Park

5. Give details an account on pUC19 vector.

pUC19 is a plasmid cloning vector created by Messing and co-workers in the University of California. p in the name stands for plasmid and UC represents the University in which it was created. It is a circular double stranded DNA and has 2686 base pairs.

pUC19 is one of the most widely used vector molecules as the recombinants, or the cells into which foreign DNA has been introduced, can be easily distinguished from the non-recombinants based on colour differences of colonies on growth media.

pUC18 is similar to pUC19, but the MCS region is reversed.

Components

It has one ampR gene (ampicillin resistance gene), and an N-terminal fragment of β-galactosidase (lac Z) gene of E. coli.

The multiple cloning site (MCS) region is split into the lac Z gene (codons 6–7 of lac Z are replaced by MCS), where various restriction sites for many restriction endonucleases are present.

The ori site or replicon, rep is derived from pMB1 vector. pUC vector is small but has a high copy number.

The high copy number of pUC plasmids is a result of the lack of the rop gene and a single point mutation in rep of pMB1. The lac Z gene codes for β-galactosidase.

Function

This plasmid is introduced into a bacterial cell by a process called "transformation", where it can multiply and express itself. However due to the presence of MCS and several restriction sites, a foreign piece of DNA of choice can be introduced into it by inserting it into place in MCS region.

The cells which have taken up the plasmid can be differentiated from cells which have not taken up the plasmid by growing it on media with Ampicillin. Only the cells with the plasmid containing the ampicillin resistance (ampR) gene will survive.

Further more, the transformed cells containing the plasmid with the gene of our interest can be distinguished from cell with the plasmid but without the gene of interest, just by looking at the colour of the colony they make on agar media. Recombinants are white, whereas non-recombinants are blue in colour. This is the most notable feature of pUC19.

Mechanism

A schematic representation of the molecular mechanism involved for screening recombinant cells

The lac Z fragment, whose synthesis can be induced by IPTG, is capable of intra-allelic complementation with a defective form of β-galactosidase enzyme encoded by host chromosome (mutation lacZDM15).

In the presence of IPTG in growth medium, bacteria synthesise both fragments of the enzyme. Both the fragments can together hydrolyse X-gal (5-bromo-4-chloro-3-indolyl- beta-D-galactopyranoside) and form blue colonies on media with X-gal.

Insertion of foreign DNA into the MCS located within the lac Z gene causes insertional inactivation of this gene at the N-terminal fragment of beta-galactosidase and abolishes intra-allelic complementation.

Thus bacteria carrying recombinant plasmids in the MCS cannot hydrolyse X-gal, giving rise to white colonies, which can be distinguished on culture media from non-recombinant cells, which are blue. Therefore the media used should contain ampicillin, IPTG, and X-gal.

Sequence

The recognition sites for HindIII, SphI, PstI, SalI, XbaI, BamHI, SmaI, KpnI, SacI and EcoRI restriction enzymes have been derived from the vector M13mp19 and are on the strand complementary to that shown.

6. Write about the M13 Vectors.

M13 is a filamentous bacteriophage composed of circular single stranded DNA (ssDNA)

which is 6407 nucleotides long encapsulated in approximately 2700 copies of the major coat protein P8, and capped with 5 copies of two different minor coat proteins (P9, P6, P3) on the ends. The minor coat protein P3 attaches to the receptor at the tip of the F pilus of the host Escherichia coli. Infection with filamentous phages is not lethal, however the infection causes turbid plaques in E. coli. It is a non-lytic virus. However a decrease in the rate of cell growth is seen in the infected cells. M13 plasmids are used for many recombinant DNA processes, and the virus has also been studied for its uses in nanostructures and nanotechnology.

Phage particles

The phage coat is primarily assembled from a 50 amino acid protein called pVIII (or p8), which is encoded by gene VIII (or g8) in the phage genome. For a wild type M13 particle, it takes approximately 2700 copies of p8 to make the coat about 900 nm long. The coat's dimensions are flexible though and the number of p8 copies adjusts to accommodate the size of the single stranded genome it packages. For example, when the phage genome was mutated to reduce its number of DNA bases (from 6.4 kb to 221 bp) , then the number of p8 copies was decreased to fewer than 100, causing the p8 coat to shrink in order to fit the reduced genome. The phage appear to be limited at approximately twice the natural DNA content. However, deletion of a phage protein (p3) prevents full escape from the host E. coli, and phage that are

10-20X the normal length with several copies of the phage genome can be seen shedding from the E. coli host.

There are four other proteins on the phage surface, two of which have been extensively studied. At one end of the filament are five copies of the surface exposed pIX (p9) and a more buried companion protein, pVII (p7). If p8 forms the shaft of the phage, p9 and p7 form the "blunt" end that is seen in the micrographs. These proteins are very small, containing only 33 and 32 amino acids respectively, though some additional residues can be added to the N-terminal portion of each which are then presented on the outside of the coat. At the other end of the phage particle are five copies of the surface exposed pIII (p3) and its less exposed accessory protein, pVI (p6). These form the rounded tip of the phage and are the first proteins to interact with the E. coli host during infection. p3 is also the last point of contact with the host as new phage bud from the bacterial surface.

Phage life-cycle

The general stages to a viral life cycle are: infection, replication of the viral genome, assembly of new viral particles and then release of the progeny particles from the host. Filamentous phage use a bacterial structure known as the F pilus to infect E. coli, with the M13 p3 tip contacting the TolA protein on the bacterial pilus. The phage genome is then transferred to the cytoplasm of the bacterial cell where resident proteins convert the single stranded DNA genome to a double stranded replicative form ("RF"). This DNA then serves as a template for expression of the phage genes.

Two phage gene products play critical roles in the next stage of the phage life cycle, namely amplification of the genome. pII (aka p2) nicks the double stranded form of the genome to initiate replication of the + strand. Without p2, no replication of the phage genome can occur. Host enzymes copy the replicated + strand, resulting in more copies of double stranded phage DNA. pV (aka p5) competes with double stranded DNA formation by sequestering copies of the + stranded DNA into a protein/DNA complex destined for packaging into new phage particles. Interestingly there is one additional phage-encoded protein, pX (p10), that is important for regulating the number of double stranded genomes in the bacterial host. Without p10 no + strands can accumulate. What's particularly interesting about p10 is that it's identical to the C-terminal portion of p2 since the gene for p10 is within the gene for p2 and the protein arises from transcription initiation within gene 2. This makes the manipulation of p10 inextricably linked to manipulation of p2 (an engineering headache) but it also makes for a compact and efficient phage in nature.

Phage maturation requires the phage-encoded proteins pIV (p4), pI (p1) and its translational restart product pXI (p11). Multiple copies (on the order of 12 or 14) of p4 assemble in the outer membrane into a stable, i.e. detergent resistant, barrel-shaped structure. Similarly a handful of the p1 and p11 proteins (5 or 6 copies of each) assemble in the bacterial inner membrane, and genetic evidence suggests C-terminal portions of p1 and p11 interact with the N-terminal portion of p4 in the periplasm. Together the p1, p11, p4 complex forms channels

through which mature phage are secreted from the bacterial host.

To initiate phage secretion, two of the minor phage coat proteins, p9 and p7, are thought to interact with the p5-single stranded DNA complex at a region of the DNA called the packaging sequence (aka PS). The p5 proteins covering the single stranded DNA are then replaced by p8 proteins that are embedded in the bacterial membrane and the growing phage filament is threaded through the p1, p11, p4 channel. This replacement of p5 by p8 explains the microphage data presented earlier indicate how the size of the phage particle is determined by the number of bases the phage packages. Once the phage DNA has been fully coated with p8, the secretion terminates by adding the p3/p6 cap, and the new phage detaches from the bacterial surface. How long does all this take? Amazingly, new M13 phage particles are secreted within 10 minutes from a newly infected host and can arise at a rate of 1000/cell within the first hour of infection. The bacterial host can continue to grow and divide, allowing this process to continue indefinitely.

Replication in E. coli

Below are steps involved with replication of M13 in E. coli.

Viral (+) strand DNA enters cytoplasm Complementary (-) strand is synthesized by bacterial enzymes

DNA Gyrase, a type II topoisomerase, acts on double-stranded DNA and catalyzes formation of negative supercoils in double-stranded DNA

Final product is parental replicative form (RF) DNA

A phage protein, pII, nicks the (+) strand in the RF

3'-hydroxyl acts as a primer in the creation of new viral strand

pII circulizes displaced viral (+) strand DNA

Pool of progeny double-stranded RF molecules produced

Negative strand of RF is template of transcription

mRNAs are translated into the phage proteins

Phage proteins in the cytoplasm are pII, pX, and pV, and they are part of the replication process of DNA. The other phage proteins are synthesized and inserted into the cytoplasmic or outer membranes.

pV dimers bind newly synthesized single-stranded DNA and prevent conversion to RF DNA

RF DNA synthesis continues and amount of pV reaches critical concentration

DNA replication switches to synthesis of single-stranded (+) viral DNA

pV-DNA structures from about 800 nm long and 8 nm in diamter

pV-DNA complex is substrate in phage assembly reaction

Research

George Smith showed that fragments of EcoRI endonuclease could fuse to amino-terminal portion of pIII.

In 2006, MIT researchers modified the DNA of M13 phages to produce a protein that would complex with cobalt ions in solution, leading to cobalt oxide, a material with energy storage capacity higher than current carbon-based lithium-ion batteries.

7. Define Ti plasmid. Give detail an account on that.

Ti plasmid is a circular plasmid that often, but not always, is a part of the genetic equipment that Agrobacterium tumefaciens and Agrobacterium rhizogenes use to transduce its genetic material to plants. The Ti plasmid is lost when Agrobacterium is grown above 28°C.

Such cured bacteria do not induce crown galls, i.e. they become avirulent. pTi and pRi share little sequence homology but are functionally rather similar.

The Ti plasmids are classified into different types based on the type of opine produced by their genes. The different opines specified by pTi are octopine, nopaline, succinamopine and leucinopine.

The plasmid has 196 genes that code for 195 proteins. There is no one structural RNA. The plasmid is 206,479 nucleotides long, the GC content is 56% and 81% of the material is coding genes. There are no pseudogenes.

The modification of this plasmid is very important in the creation of transgenic plants, but only in dicotyledon plants.

Virulence Region

Genes in the virulence region are grouped into the operons virABCDEFG, which code for the enzymes responsible for mediating transduction of T-DNA to plant cells.

virA codes for a receptor which reacts to the presence of phenolic compounds such as acetosyringone, syringealdehyde or acetovanillone which leak out of damaged plant tissues.

virB encodes proteins which produce a pore/pilus-like structure.

virC binds the overdrive sequence.

virD1 and virD2 produce endonucleases which target the direct repeat borders of the T-DNA segment, beginning with the right border.

virG activates vir-gene expression after binding to a consensus sequence, once it has been phosphorylated by virA.

UNIT IVRestriction mapping : Restriction map construction – Double digest. RFLP – PCR. Site directed mutagenesis, Protein engineering.

Site-directed mutagenesis

Three different methods of site-directed mutagenesis have been devised: cassette mutagenesis, primer extension and procedures based on the PCR. All three are described below but the reader wishing more detail should consult the review of Ling and Robinson (1998). In some cases, the goal of protein engineering is togenerate a molecule with an improvement in some operating parameter, but it is not known what amino acid changes to make. In this situation,a random mutagenesis strategy provides a route to the desired protein. However, methods based on gene manipulation differ from traditional mutagenesis in that the mutations are restricted to the gene of interest or a small portion of it. Genetic engineering also provides a number of simple methods of generating chimeric proteins where each domain is derived from a different protein. It should not be forgotten that constructing the mutant DNA is only part of the task. The vector for expression, the expression system and strategies for purification and assay must also be consideredbefore embarking on protein mutagenesis.Cassette mutagenesisIn cassette mutagenesis, a synthetic DNA fragment containing the desired mutant sequence is used to replace the corresponding sequence in the wild-type gene. This method was originally used to generate improved variants of the enzyme subtilisin (Wells et al. 1985). It is a simple method for which the efficiency of mutagenesis is close to 100%. The disadvantages are the requirement for unique restriction sites flanking the region of interest and the limitation on the realistic number of different oligonucleotide replacements that can be synthesized. The latter problem can be minimized by the use of doped oligonucleotides ( Reidhaar-Olson and Sauer, 1988). Primer extension:the single-primer methodThe simplest method of site-directed mutagenesis is the single-primer method (Gillam et al. 1980, Zoller & Smith 1983). The method involves priming in vitro DNA synthesis with a chemically synthesized oligonucleotide (7–20 nucleotides long) that carries a base mismatch with the complementary sequence. The method requires that the DNA to be mutated is available in single-stranded form, and cloning the gene in M13-based vectors makes this easy. However, DNA cloned in a plasmid and obtained in duplex form can also be converted to a partially single-stranded molecule that is suitable (Dalbadie-McFarland et al. 1982).The synthetic oligonucleotide primes DNA synthesis and is itself incorporated into the resulting heteroduplex molecule. After transformation of the host E. coli, this heteroduplex gives rise to homoduplexes w hose sequences are either that of the original wild-type DNA or that containing the mutated base. The frequency with which mutated clones arise,compared with wild-type clones, may be low. In order to pick out mutants, the clones can be screened by nucleic acid hybridization (see Chapter 6) with 32P-labelled oligonucleotide as probe. Under suitable conditions of stringency, i.e. temperature and cation concentration, a positive signal will be obtained only with mutant clones. This allows ready detection of the desired mutant (Wallace et al. 1981, Traboni et al. 1983). In order to check that the procedure has not introduced other adventitious changes, it is prudent to check the sequence of the mutant directly by DNA sequencing. This was a particular necessity with early versions of the technique which made use of E. coli DNA polymerase. The more recent use of the high-fidelity DNA polymerases from phages T4 and T7 has minimized the problem of extraneousmutations, as well as shortening the time for copying the second strand. Also, these polymerases do not ‘strand-displace’ the oligomer, a process which would eliminate the original mutant oligonucleotide. A variation of the procedure (Fig. 7.10) outlined above involves oligonucleotides containing inserted or deleted sequences. As long as stable hybrids areformed with single-stranded wild-type DNA, priming of in vitro DNA synthesis can occur, ultimately g ving rise to clones corresponding to the inserted or deleted sequence (Wallace et al. 1980, Norrander et al. 1983). PCR methods of site-directed mutagenesisEarly work on the development of the PCR method of DNA amplification showed its potential for mutagenesis (Scharf et al. 1986). Single bases mismatched between the amplification primer and the template become incorporated into the template sequence as a result of amplification (Fig. 7.11). Higuchi et al. (1988) have described a variation of the basic method which enables a mutation in a PCR-produced DNA fragment to be introduced anywhere along its length. Two primary PCR reactions produce two overlapping DNA fragments, both bearing the same mutation

in the overlap region. The overlap in sequence allows the fragments to hybridize (Fig. 7.11). One of the two possible hybrids is extended by DNA polymerase to produce a duplex fragment. The other hybrid has recessed 5′ends and, since it is not a substrate for the polymerase, is effectively lost from the reaction mixture. As with conventional primerextension mutagenesis, deletions and insertions can also be created.

The polymerase chain reaction (PCR)The impact of the PCR upon molecular biology has been profound. The reaction is easily performed, and leads to the amplification of specific DNA sequences by an enormous factor. From a simple basic principle, many variations have been developed with applications throughout gene technology (Erlich 1989, Innis et al. 1990). Very importantly, the PCRhas revolutionized prenatal diagnosis by allowing tests to be performed using small samples of fetal tissue. In forensic science, the enormous sensitivity of PCR-based procedures is exploited in DNA profiling; following the publicity surrounding Jurassic Park, virtually everyone is aware of potential applications in palaeontology and archaeology. Many other processes have been described which should produce equivalent results to a PCR (for review, see Landegran 1996) but as yet none has found widespread use. In many applications of the PCR to gene manipulation,the enormous amplification is secondary to the aim of altering the amplified sequence. This often involves incorporating extra sequences at the ends of the amplified DNA. In this section we shall consider only the amplification process. The applicationsof the PCR will be described in appropriate places.Basic reactionFirst we need to consider the basic PCR. The principle is illustrated in Fig. 2.7. The PCR involves two oligonucleotide primers, 17–30 nucleotides in length, which flank the DNA sequence that is to be amplified. The primers hybridize to opposite strands of the DNA after it has been denatured, and are orientated so that DNA synthesis by the polymerase proceeds through the region between the two primers. The extension reactions create two doublestranded target regions, each of which can again be denatured ready for a second cycle of hybridization and extension. The third cycle produces two doublestranded molecules that comprise precisely the target region in double-stranded form. By repeated cycles of heat denaturation, primer hybridization and extension, there follows a rapid exponential accumulation of the specific target fragment of DNA. After 22 cycles, an amplification of about 106- fold is expected (Fig. 2.8), and amplifications of thisorder are actually attained in practice. In the original description of the PCR method (Mullis & Faloona 1987, Saiki et al. 1988, Mullis 1990), Klenow DNA polymerase was used and, because of the heat-denaturation step, fresh enzymehad to be added during each cycle. A breakthrough came with the introduction of Taq DNA polymerase (Lawyer et al. 1989) from the thermophilic bacterium Thermus aquaticus. The Taq DNA polymerase is resistant to high temperatures and so does not need to be replenished during the PCR (Erlich et al. 1988, Sakai et al. 1988). Furthermore, by enabling theextension reaction to be performed at higher temperatures, the specificity of the primer annealing is not compromised. As a consequence of employing the heat-resistant enzyme, the PCR could be automated very simply by placing the assembled reaction in a heating block with a suitable thermal cycling programme ··

UNIT VDNA sequencing – Dideoxy method, Maxam Gilbert method – Mapping and sequencingthe Human genome.

Sanger Method for DNA Sequencing

DNA sequencing, first devised in 1975, has become a powerful technique in molecular biology, allowing analysis of genes at the nucleotide level. For this reason, this tool has been applied to many areas of research. For example, the polymerase chain reaction (PCR), a method which rapidly produces numerous copies of a desired piece of DNA, requires first knowing the flanking sequences of this piece. Another important use of DNA sequencing is identifying restriction sites in plasmids. Knowing these restriction sites is useful in cloning a foreign gene into the plasmid. Before the advent of DNA sequencing, molecular biologists had to sequence proteins directly; now amino acid sequences can be determined more easily by sequencing a piece of cDNA and finding an open reading frame. In eukaryotic gene expression, sequencing has allowed researchers to identify conserved sequence motifs and determine their importance in the promoter region. Furthermore, a molecular biologist can utilize sequencing to identify the site of a point mutation. These are only a few examples illustrating the way in which DNA sequencing has revolutionized molecular biology.

Dideoxynucleotide sequencing represents only one method of sequencing DNA. It is commonly called Sanger sequencing since Sanger devised the method. This technique utilizes 2',3'-dideoxynucleotide triphospates (ddNTPs), molecules that differ from deoxynucleotides by the having a hydrogen atom attached to the 3' carbon rather than an OH group. (Figure 1). These molecules terminate DNA chain elongation because they cannot form a phosphodiester bond with the next deoxynucleotide.

In order to perform the sequencing, one must first convert double stranded DNA into single stranded DNA. This can be done by denaturing the double stranded DNA with NaOH. A Sanger reaction consists of the following: a strand to be sequenced (one of the single strands which was denatured using NaOH), DNA primers (short pieces of DNA that are both complementary to the strand which is to be sequenced and radioactively labelled at the 5' end), a mixture of a particular ddNTP (such as ddATP) with its normal dNTP (dATP in this case), and the other three dNTPs (dCTP, dGTP, and dTTP). The concentration of ddATP should be 1% of the concentration of dATP. The logic behind this ratio is that after DNA polymerase is added, the polymerization will take place and will terminate whenever a ddATP is incorporated into the growing strand. If the ddATP is only 1% of the total concentration of dATP, a whole series of labeled strands will result (Figure 1). Note that the lengths of these strands are dependent on the location of the base relative to the 5' end.

This reaction is performed four times using a different ddNTP for each reaction. When these reactions are completed, a polyacrylamide gel electrophoresis (PAGE) is

performed. One reaction is loaded into one lane for a total of four lanes (Figure 2). The gel is transferred to a nitrocellulose filter and autoradiography is performed so that only the bands with the radioactive label on the 5' end will appear. In PAGE, the shortest fragments will migrate the farthest. Therefore, the bottom-most band indicates that its particular dideoxynucleotide was added first to the labeled primer. In Figure 2, for example, the band that migrated the farthest was in the ddATP reaction mixture. Therefore, ddATP must have been added first to the primer, and its complementary base, thymine, must have been the base present on the 3' end of the sequenced strand. One can continue reading in this fashion. Note in Figure 2 that if one reads the bases from the bottom up, one is reading the 5' to 3' sequence of the strand complementary to the sequenced strand. The sequenced strand can be read 5' to 3' by reading top to bottom the bases complementary to the those on the gel.

Figure 1. This figure shows the structure of a dideoxynucleotide (notice the H atom attached to the 3' carbon). Also depicted in this figure are the ingredients for a Sanger reaction. Notice the different lengths of labeled strands produced in this reaction.

Figure 2. This figure is a representation of an acrylamide sequencing gel. Notice that the sequence of the strand of DNA complementary to the sequenced strand is 5' to 3' ACGCCCGAGTAGCCCAGATT while the sequence of the sequenced strand, 5' to 3', is AATCTGGGCTACTCGGGCGT

Maxam & Gilbert Sequencing

There are four chemical cleavage reactions at the core of the Maxam and Gilbert sequencing system. The figure below left shows an example from these reactions, the reaction cleaving specifically at guanine. The other three reactions cleave at G+A, C+T, or C. Guanine and cytosine, therefore, give bands in 2 lanes, adenine and thymine in only one. An example of the gel pattern produced is presented below right.

Maxam and Gilbert DNA sequencing reaction specific for Guanidine residues. The Guanine base is first modified with Dimethyl Sulfate (DMS), which makes the chain susceptible to cleavage by piperidine, destroying the Guanidine residue and releasing a labeled fragment for electrophoresis.

In a Maxam and Gilbert gel, the identity of guanine or cytosine in the sequence can be assigned most easily because two of the four reaction sets cleave at those bases alone. Adenine or thymine are slightly more difficult, being represented by those bands in the G+A or C+T lanes which do not appear, respectively, in the G or C lanes.

The DNA to be sequenced must first be end labeled, at one end only. This is accomplished by kinase treatment with 32P ATP, which labels both ends, followed by restriction digestion and isolation of the two labeled fragments. Alternatively, digestion of a plasmid containing a clone of the DNA of interest with an appropriate enzyme can yield a unique labeling site. Plasmid vectors containing the rare site for Tth111I, which leaves a single 5' base overlap, have been generated for this purpose. Cleavage with Tth111I leaves a G at one end and a C at the other in these vectors. By filling in the gap with Klenow polymerase fragment in the presence of dGTP or dCTP, one end or the other can be labeled specifically. Labeled DNA is first precipitated to remove any salts which might interfere in the cleavage reactions. It is then modified, cleaved and run on a denaturing gel for analysis. NB: THE HYDRAZINE AND DMS USED IN THESE PROTOCOLS ARE TOXIC AND VOLATILE. KEEP TUBES SEALED AND WORK IN A HOOD.

Maxam and Gilbert Sequencing Reactions 1. Precipitate the substrate: To the 32P labeled DNA, add 0.1 vol. 3M Sodium Acetate

and 1 vol. Isopropanol. Precipitate at -70°C for 10 minutes, and centrifuge at max RPM in a microcentrifuge for 5 minutes to collect the DNA. Wash the pellet twice with 1 ml cold 70% ethanol to remove all salt. Redissolve the DNA in 45 µl of sterile water. Count one microliter of the solution in scintillation cocktail to confirm >5x103 cpm total counts.

2. Aliquot 10 µl of the DNA solution into each of 4 tubes. Label the tubes C, G, C+T, G+A.

3. Reactions:

C: Add 10µl 2.5M NaCl and mix well. Add 30µl of Hydrazine (toxic!) and incubate at 25°C for 7-9 minutes.

G: Add 200µl of: 50mM sodium cacodylate, pH 8, 1mM EDTA. Mix well and add 1µl Dimethyl Sulfate (DMS) (Toxic!) and incubate at 25°C for 4-5 minutes.

C+T: Add 10µl H2O and mix well. Add 30µl Hydrazine and incubate at 25°C for 7-9 minutes.

G+A: Add 25µl of formic acid, mix well and incubate at 25°C for 4-5 minutes.

4. Stop the reactions:

Stop buffers:

G reaction: Add 50µl of:1.5M sodium acetate pH 7, 1M mercaptoethanol, 100µg/ml tRNA.

All other reactions: Add 200µl of 0.3M sodium acetate, pH 7, 0.1mM EDTA, 25µg/ml tRNA.

Ethanol precipitation:Add 750µl of Ethanol, and transfer reactions to a -70°C bath for 5 minutes. Collect DNA by microcentrifugation for 5 minutes. Discard the supernatants as appropriate for DMS or Hydrazine waste. Rinse the pellets twice with 70% ethanol. Redissolve the pellets in 300µl of water, add 30µl of 3M sodium acetate and 1ml of ethanol. Pellet DNA and wash twice with 70% ethanol. Allow the pellets to air dry.

5. Piperidine cleavage reactions:Resuspend pellets in 75µl of 10% piperidine, and transfer to screw top tubes. It is essential that the tubes used for the piperidine reaction seal well in order to ensure that the reaction goes to completion. Incubate the tubes at 90°C for 30 minutes. Cool the tubes, centrifuge briefly to collect the condensate, and evaporate to dryness in a speedvac. Redissolve the pellet in 40µl of water and dry again. Repeat the rehydration and drying once more to ensure that all of the piperidine has been removed. The samples are now ready for denaturing PAGE.

Mapping and Sequencing the Human Genome

A primary goal of the Human Genome Project is to make a series of descriptive diagrams maps of each human chromosome at increasingly finer resolutions. Mapping involves (1) dividing the chromosomes into smaller fragments that can be propagated and characterized and (2) ordering (mapping) them to correspond to their respective locations on the chromosomes. After mapping is completed, the next step is to determine the base sequence of each of the ordered DNA fragments. The ultimate goal of genome research is to find all the genes in the DNA sequence and to develop tools for using this information in the study of human biology and medicine.

Improving the instrumentation and techniques required for mapping and sequencing a major focus of the genome project will increase efficiency and cost- effectiveness. Goals include automating methods and optimizing techniques to extract the maximum useful information from maps and sequences.

A genome map describes the order of genes or other markers and the spacing between them on each chromosome. Human genome maps are constructed on several different scales or levels of resolution. At the coarsest resolution are genetic linkage maps, which depict the relative chromosomal locations of DNA markers (genes and other identifiable DNA sequences) by their patterns of inheritance. Physical maps describe the chemical characteristics of the DNA molecule itself.

Geneticists have already charted the approximate positions of over 2300 genes, and a start has been made in establishing high- resolution maps of the genome (Fig. 7: Assignment of Genes to Specific Chromosomes). More- precise maps are needed to organize systematic sequencing efforts and plan new research directions.

HUMAN GENOME PROJECT GOALS Resolution

Complete a detailed human genetic map 2 Mb

Complete a physical map 0.1 Mb

Acquire the genome as clones 5 kb

Determine the complete sequence 1 bp

Find all the genes

With the data generated by the project, investigators will determine the functions of the genes and develop tools for biological and medical applications.

Mapping Strategies

Genetic Linkage Maps

A genetic linkage map shows the relative locations of specific DNA markers along the chromosome. Any inherited physical or molecular characteristic that differs among individuals and is easily detectable in the laboratory is a potential genetic marker. Markers can be expressed DNA regions (genes) or DNA segments that have no known coding function but whose inheritance pattern can be followed. DNA sequence differences are especially useful markers because they are plentiful and easy to characterize precisely.

Markers must be polymorphic to be useful in mapping; that is, alternative forms must exist among individuals so that they are detectable among different members in family studies. Polymorphisms are variations in DNA sequence that occur on average once every 300 to 500 bp. Variations within exon sequences can lead to observable changes, such as differences in eye color, blood type, and disease susceptibility. Most variations occur within introns and have little or no effect on an organisms appearance or function, yet they are detectable at the DNA level and can be used as markers. Examples of these types of markers include (1) restriction fragment length polymorphisms (RFLPs), which reflect sequence variations in DNA sites that can be cleaved by DNA restriction enzymes, and (2) variable number of tandem repeat sequences, which are short repeated sequences that vary in the number of repeated units and, therefore, in length (a characteristic easily measured). The human genetic linkage map is constructed by observing how frequently two markers are inherited together.

Two markers located near each other on the same chromosome will tend to be passed together from parent to child. During the normal production of sperm and egg cells, DNA strands occasionally break and rejoin in different places on the same chromosome or on the other copy of the same chromosome (i.e., the homologous chromosome). This process (called meiotic recombination) can result in the separation of two markers originally on the same chromosome (Fig. 8: Constructing a Genetic Linkage Map). The closer the markers are to each other the more tightly linked the less likely a recombination event will fall between and separate them. Recombination frequency thus provides an estimate of the distance between two markers.

On the genetic map, distances between markers are measured in terms of centimorgans (cM), named after the American geneticist Thomas Hunt Morgan. Two markers are said to be 1 cM apart if they are separated by recombination 1% of the time. A genetic distance of 1 cM is roughly equal to a physical distance of 1 million bp (1 Mb). The current resolution of most human genetic map regions is about 10 Mb.

The value of the genetic map is that an inherited disease can be located on the map by following the inheritance of a DNA marker present in affected individuals (but absent in unaffected individuals), even though the molecular basis of the disease may not yet be understood nor the responsible gene identified. Genetic maps have been used to find the exact chromosomal location of several important disease genes, including cystic fibrosis, sickle cell disease, Tay- Sachs disease, fragile X syndrome, and myotonic dystrophy.

One short- term goal of the genome project is to develop a high- resolution genetic map (2 to 5 cM); recent consensus maps of some chromosomes have averaged 7 to 10 cM between genetic markers. Genetic mapping resolution has been increased through the application of recombinant DNA technology, including in vitro radiation- induced chromosome fragmentation and cell fusions (joining human cells with those of other species to form hybrid cells) to create panels of cells with specific and varied human chromosomal components. Assessing the frequency of marker sites remaining together after radiation- induced DNA fragmentation can establish the order and distance between the markers. Because only a single copy of a chromosome is required for analysis, even nonpolymorphic markers are useful in radiation hybrid mapping. [In meiotic mapping (described above), two copies of a chromosome must be distinguished from each other by polymorphic markers.]

Restriction Enzymes: Microscopic Scalpels

Isolated from various bacteria, restriction enzymes recognize short DNA sequences and cut the DNA molecules at those specific sites. (A natural biological function of these enzymes is to protect bacteria by attacking viral and other foreign DNA.) Some restriction enzymes (rare- cutters) cut the DNA very infrequently, generating a small number of very large fragments (several thousand to a million bp). Most enzymes cut DNA more frequently, thus generating a large number of small fragments (less than a hundred to more than a thousand bp).

On average, restriction enzymes with

4-base recognition sites will yield pieces 256 bases long, 6-base recognition sites will yield pieces 4000 bases long, and

8-base recognition sites will yield pieces 64,000 bases long.

Since hundreds of different restriction enzymes have been characterized, DNA can be cut into many different small fragments.

Physical Maps

Different types of physical maps vary in their degree of resolution. The lowest- resolution physical map is the chromosomal (sometimes called cytogenetic) map, which is based on the distinctive banding patterns observed by light microscopy of stained chromosomes. A cDNA map shows the locations of expressed DNA regions (exons) on the chromosomal map. The more detailed cosmid contig map depicts the order of overlapping DNA fragments spanning the genome. A macrorestriction map describes the order and distance between enzyme cutting (cleavage) sites. The highest- resolution physical map is the complete elucidation of the DNA base- pair sequence of each chromosome in the human genome. Physical maps are described in greater detail below.

Low-Resolution Physical Mapping

Chromosomal map. In a chromosomal map, genes or other identifiable DNA fragments are assigned to their respective chromosomes, with distances measured in base pairs. These markers can be physically associated with particular bands (identified by cytogenetic staining) primarily by in situ hybridization, a technique that involves tagging the DNA marker with an observable label (e.g., one that fluoresces or is radioactive). The location of the labeled probe can be detected after it binds to its complementary DNA strand in an intact chromosome.

As with genetic linkage mapping, chromosomal mapping can be used to locate genetic markers defined by traits observable only in whole organisms. Because chromosomal maps are based on estimates of physical distance, they are considered to be physical maps. The number of base pairs within a band can only be estimated.

Until recently, even the best chromosomal maps could be used to locate a DNA fragment only to a region of about 10 Mb, the size of a typical band seen on a chromosome. Improvements in

fluorescence in situ hybridization (FISH) methods allow orientation of DNA sequences that lie as close as 2 to 5 Mb. Modifications to in situ hybridization methods, using chromosomes at a stage in cell division (interphase) when they are less compact, increase map resolution to around 100,000 bp. Further banding refinement might allow chromosomal bands to be associated with specific amplified DNA fragments, an improvement that could be useful in analyzing observable physical traits associated with chromosomal abnormalities.

cDNA map. A cDNA map shows the positions of expressed DNA regions (exons) relative to particular chromosomal regions or bands. (Expressed DNA regions are those transcribed into mRNA.) cDNA is synthesized in the laboratory using the mRNA molecule as a template; base- pairing rules are followed (i.e., an A on the mRNA molecule will pair with a T on the new DNA strand). This cDNA can then be mapped to genomic regions.

Because they represent expressed genomic regions, cDNAs are thought to identify the parts of the genome with the most biological and medical significance. A cDNA map can provide the chromosomal location for genes whose functions are currently unknown. For disease- gene hunters, the map can also suggest a set of candidate genes to test when the approximate location of a disease gene has been mapped by genetic linkage techniques.

High- Resolution Physical Mapping

The two current approaches to high- resolution physical mapping are termed top- down (producing a macrorestriction map) and bottom- up (resulting in a contig map). With either strategy (described below) the maps represent ordered sets of DNA fragments that are generated by cutting genomic DNA with restriction enzymes (see previously discussed Restriction Enzymes). The fragments are then amplified by cloning or by polymerase chain reaction (PCR) methods (see DNA Amplification below). Electrophoretic techniques are used to separate the fragments according to size into different bands, which can be visualized by direct DNA staining or by hybridization with DNA probes of interest. The use of purified chromosomes separated either by flow sorting from human cell lines or in hybrid cell lines allows a single chromosome to be mapped (see Separating Chromosomes below).

A number of strategies can be used to reconstruct the original order of the DNA fragments in the genome. Many approaches make use of the ability of single strands of DNA and/or RNA to hybridize to form double- stranded segments by hydrogen bonding between complementary bases. The extent of sequence homology between the two strands can be inferred from the length of the double- stranded segment. Fingerprinting uses restriction map data to determine which fragments have a specific sequence (fingerprint) in common and therefore overlap. Another approach uses linking clones as probes for hybridization to chromosomal DNA cut with the same restriction enzyme.

Macrorestriction maps: Top- down mapping. In top- down mapping, a single chromosome is cut (with rare- cutter restriction enzymes) into large pieces, which are ordered and subdivided; the smaller pieces are then mapped further. The resulting macro- restriction maps depict the order of and distance between sites at which rare- cutter enzymes cleave (Fig. 9a: Physical Mapping Strategies: Macrorestriction Map). This approach yields maps with more continuity

and fewer gaps between fragments than contig maps, but map resolution is lower and may not be useful in finding particular genes; in addition, this strategy generally does not produce long stretches of mapped sites. Currently, this approach allows DNA pieces to be located in regions measuring about 100,000 bp to 1 Mb.

The development of pulsed- field gel (PFG) electrophoretic methods has improved the mapping and cloning of large DNA molecules. While conventional gel electrophoretic methods separate pieces less than 40 kb (1 kb = 1000 bases) in size, PFG separates molecules up to 10 Mb, allowing the application of both conventional and new mapping methods to larger genomic regions.

Contig maps: Bottom- up mapping. The bottom- up approach involves cutting the chromosome into small pieces, each of which is cloned and ordered. The ordered fragments form contiguous DNA blocks (contigs). Currently, the resulting library of clones varies in size from 10,000 bp to 1 Mb (Fig. 9b: Physical Mapping Strategies: Contig Maps). An advantage of this approach is the accessibility of these stable clones to other researchers. Contig construction can be verified by FISH, which localizes cosmids to specific regions within chromosomal bands.

Contig maps thus consist of a linked library of small overlapping clones representing a complete chromosomal segment. While useful for finding genes localized to a small area (under 2 Mb), contig maps are difficult to extend over large stretches of a chromosome because all regions are not clonable. DNA probe techniques can be used to fill in the gaps, but they are time consuming. Figure 10 is a diagram relating the different types of maps.

Technological improvements now make possible the cloning of large DNA pieces, using artificially constructed chromosome vectors that carry human DNA fragments as large as 1 Mb. These vectors are maintained in yeast cells as artificial chromosomes (YACs). (For more explanation, see DNA Amplification below) Before YACs were developed, the largest cloning vectors (cosmids) carried inserts of only 20 to 40 kb. YAC methodology drastically reduces the number of clones to be ordered; many YACs span entire human genes. A more detailed map of a large YAC insert can be produced by subcloning, a process in which fragments of the original insert are cloned into smaller- insert vectors. Because some YAC regions are unstable, large- capacity bacterial vectors (i.e., those that can accommodate large inserts) are also being developed.

Separating Chromosomes

Flow sorting

Flow sorting employs flow cytometry to separate, according to size, chromosomes isolated from cells during cell division when they are condensed and stable. As the chromosomes flow singly past a laser beam, they are differentiated by analyzing the amount of DNA present, and individual chromosomes are directed to specific collection tubes.

Somatic cell hybridization

In somatic cell hybridization, human cells and rodent tumor cells are fused (hybridized); over time, after the chromosomes mix, human chromosomes are preferentially lost from the hybrid cell until only one or a few remain. Those individual hybrid cells are then propagated and maintained as cell lines containing specific human chromosomes. Improvements to this technique have generated a number of hybrid cell lines, each with a specific single human chromosome