SBC351 - Genetic Engineering and Functional Genomics

INTRODUCTION

A RECAPITULATION ON GENES AND GENOMES

Gene organizationThe gene is the basic unit of genetic information. Genes are located on chromosomes at a particular genetic locus. Different forms of the same gene are known as alleles.

Before the advent of molecular biology and the realization that genes were made of DNA, study of the gene was largely indirect; the effects of genes were observed in terms of their output (phenotypes). Despite the apparent limitations of this approach, a vast amount of information about how genes functioned was obtained, and the basic tenets of transmission genetics were formulated. As the gene was studied in greater detail, the terminology associated with this area of genetics became more extensive, and the ideas about genes were modified to take developments into account.The term ‘gene’ is usually taken to represent the genetic information transcribed into a single RNA molecule, which is in turn translated into a single protein. Exceptions are genes for RNA molecules (such as rRNA and tRNA), which are not translated. In addition, the nomenclature used for prokaryotic cells is slightly different because of the way that their genes are organized. In eukaryotes, genes are located on chromosomes, and the region of the chromosome where a particular gene is found is called the locus of that gene. In diploid organisms, which have their chromosomes arranged as homologous pairs, different forms of the same gene are known as alleles.

The anatomy of a geneThere are certain basic requirements for any gene to function. The most prominent is that the gene encodes the information for a particular protein (or RNA molecule). The double-stranded DNA molecule has the potential to store genetic information in either strand, although in most organisms only one strand is used to encode any particular gene. There is the potential for confusion with the nomenclature of the two DNA strands, which may be called coding/non-coding, sense/antisense, plus/minus, transcribed/non-transcribed, or template/non-template. However, the standard name is the coding strand which is eventually copied into an mRNA of a corresponding sequence. Thus, genetic information is expressed by transcription of the non-coding strand of DNA, which produces an mRNA molecule that has the same sequence as the coding strand of DNA (although the RNA has uracil substituted for thymine). In addition to the sequence of bases that specifies the amino acids in a protein-coding gene, there are regions which act as regulatory sequences. These regulate specific genes. In every gene, a

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site called the transcription starting point is required. This covers the region to which the RNA polymerase binds to during transcription and is known as the promoter (P). There is also the start point for transcription (TC). A stop site for transcription (tC) is also required. From TC start to tC stop is sometimes called the transcriptional unit, that is, the DNA region that is copied into RNA. Within this transcriptional unit there may be regulatory sites for translation, namely a start site (TL) and a stop signal (tL) for translation. Other sequences involved in the control of gene expression may be present either upstream or downstream from the gene itself.

Gene organization: The transcriptional unit produces the RNA molecule and is defined by the transcription start site (TC) and stop site (tC). Within the transcriptional unit lies within the coding sequence, from the translation start site (TL) to the stop site (tL). The upstream regulatory region may have controlling elements such as enhancers or operators in addition to the promoter (P), which is the RNA polymerase binding site.

Gene structure in prokaryotesIn prokaryotic cells such as bacteria, genes are usually found grouped together in operons. An operon is a cluster of genes that are related (often coding for enzymes in a specific metabolic pathway) and that are under the control of a single promoter/regulatory region.The best known example of this arrangement is the lac operon, which codes for the enzymes responsible for lactose catabolism. Within the operon there are three genes that code for proteins (termed structural genes) and an upstream control region encompassing the promoter and a regulatory site called the operator. In this control region there is also a site that binds a complex of cAMP (cyclic adenosine monophosphate) and CRP (cAMP receptor protein which is important in positive regulation/stimulation of transcription).Lying outside the operon itself is the repressor gene which codes for a protein (the Lac repressor) that binds to the operator site and is responsible for negative control of the operon by blocking the binding of RNA polymerase. The fact that structural genes in prokaryotes are often grouped

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together means that the transcribed mRNA may contain information for more than one protein. Such a molecule is known as a polycistronic mRNA, Thus, much of the genetic information in bacteria is expressed via polycistronic mRNAs whose synthesis is regulated in accordance with the needs of the cell at any given time. This system is flexible and efficient, and it enables the cell to adapt quickly to changing environmental conditions.

Gene structure in eukaryotesA major feature that affects the processing of genetic material in eukaryotic cells is the presence of a nucleus within which the DNA is stored in the form of chromosomes. The nucleus is bound by the nucleus membrane. Transcription occurs within the nucleus and is therefore separated from the site of translation, which is in the cytoplasm. Eukaryotes also mitochondria (plant and animal cells) and chloroplasts (plant cells only) which also contain DNA i.e. they have their own separate genomes that specify many of the components required by these organelles. This compartmentalization has important consequences for regulation both genetic and metabolic, and thus gene structure and function in eukaryotes are more complex than in prokaryotes.Another unique feature of eukaryotic cells is the presence of segments of DNA along the chromosome that do not code for any genes but interrupt coding sequences. These intervening sequences are called introns. They are eventually cut off in a process called “splicing” to leave only regions that will eventually be copied to the mRNA sequence. The parts that will make up the mRNA are known as exons. In many cases the number and cumulative length of the introns exceeds that of the exons. For example, in the chicken ovalbumin gene has a total of seven introns making up more than 75% of the gene. Introns must be removed before the mRNA can be translated. This is carried out in the nucleus, where the introns are spliced out of the primary transcript. Further intranuclear modification includes the addition of a ‘cap’ at the 5΄-terminus and a ‘tail’ of adenine residues at the 3΄-terminus (the poly A tail). These modifications are part of what is known as RNA processing, and the end product is a fully functional mRNA that is ready for export to the cytoplasm for translation.

The central dogmaPut simply, it means “DNA makes RNA that makes protein". The central dogma of molecular biology deals with the detailed residue-by-residue transfer of sequential information. It states that such information cannot be transferred from protein to either protein or nucleic acid. The more complete form of the central dogma is stated in the following figure:

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Fig. The central dogma.

Gene expressionWhilst a detailed knowledge of gene expression is not required in order to understand the principles of genetic engineering, it is useful to be familiar with the main features of transcription and translation and to have some knowledge of how gene expression is controlled.

Transcription: Transcription is synthesis of RNA from the DNA template. Note that the non-coding strand of a gene is the one that is transcribed. The enzyme responsible is RNA polymerase (DNA-dependent RNA polymerase). Prokaryotes have a single RNA polymerase enzyme, while eukaryotes possess three types of RNA polymerases (I, II, and III). These synthesize ribosomal, messenger, and transfer/5s ribosomal RNAs, respectively. All RNA polymerases are large multi-subunit proteins with relative molecular masses of around 500000.Transcription has several component stages: DNA/RNA polymerase binding Chain initiation Chain elongation Chain termination and release of the RNA.

When the RNA molecule is released, it may be immediately available for translation (if it is prokaryotes) or it may be processed further and exported to the cytoplasm (as in eukaryotes) before translation occurs.

Translation: Translation requires an mRNA molecule, a supply of charged tRNAs (tRNA molecules with their associated amino acid residues), and ribosomes (composed of rRNA and

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ribosomal proteins). Protein synthesis occurs on the ribosomes. In prokaryotes, the codon/anticodon recognition event marks the link between nucleic acid and protein.Ribosomes are composed of three rRNAs and some 52 different ribosomal proteins. The ribosome is a complex structure that essentially acts as a platform that holds the mRNA in place so that the codons may be matched up with the appropriate anticodon on the tRNA, thus ensuring that the correct amino acid is inserted into the growing polypeptide chain. The mRNA molecule is translated in a 5΄→3΄direction which leads to polypeptide elongation from N terminus to C terminus direction.

Regulation of gene expression: Transcription and translation provide the mechanisms by which genes are expressed. However, it is vital that gene expression is controlled so that the correct gene products are produced in the cell at the right time and in the right amounts.Gene can be adaptively regulated to enable the adaptation of an organism to changing environmental conditions for example. In eukaryotes, some genes are developmentally regulated so that they are expressed only at the right stage in an organism. There are genes which are constantly being expressed. Examples include those which code metabolic enzymes that are always present in the cell. Such genes are said to be constitutively expressed and are called housekeeping or constitutive genes. They are essentially unregulated.As brief example of gene regulation, we know that there will always be fluctuations in the availability of nutrients. If the cell is to survive, it must conserve energy resources, which means that wasteful synthesis of non-required proteins should be prevented. Thus, bacterial cells have mechanisms that enable operons to be controlled with a high degree of sensitivity. An operon that encodes proteins involved in a catabolic pathway (one that breaks down materials to release energy) is often regulated by being switched ‘on’ only when the substance becomes available in the extracellular medium. Thus, when the substance is absent, there are systems that keep catabolic operons switched ‘off ’. These are said to be inducible operons and are usually controlled by a negative control mechanism involving a repressor protein that prevents access to the promoter by RNA polymerase.The classic example of a catabolic operon is the lac operon. When lactose is absent, the repressor protein binds to the operator and the system is ‘off ’. The system is a little ‘leaky’, however, and thus the proteins encoded by the operon (β-galactosidase, permease, and transacetylase) will be present in the cell at low levels. When lactose becomes available, it is transported into the cell by the permease and binds to the repressor protein, causing a conformational (shape) change so that the repressor is unable to bind to the operator. Thus, the negative control is removed, and the operon is accessible by RNA polymerase.A second level of control, based on the level of cAMP, ensures that full activity is only attained when lactose is present and energy levels are low. This dual-control mechanism is a very effective way of regulating gene expression, enabling a range of levels of expression that is a bit like a dimmer switch rather than an on/off switch. In the case of catabolic operons like the lac system, this ensures that the enzymes are only synthesized at maximum rate when they are really required.

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Genome organization: The C-value paradox refers to the strange observation that eukaryotic genomes are quite large yet only a small fraction of the genome codes for structural genes. Viral and bacterial genomes on the other hand tend to show very efficient use of DNA for encoding their genes as most of the DNA is coding sequence. In the human genome, only about 3% of the total amount of DNA is actually coding sequence. Even when the introns and control sequences are factored, the majority of the DNA has no obvious function. This is sometimes termed ‘junk’ DNA, although this is perhaps the wrong way to think about this apparently redundant DNA.Many genes in eukaryotes are single copy genes, and tend to be dispersed across the multiple chromosomes found in eukaryotic cell nuclei. Other genes may be part of multigene families, and may be grouped at a particular chromosomal location or may be dispersed. Some genes also exist as tandemly repeated copies.When studying gene organization in the context of the genome itself, features such as gene density, gene size, mRNA size, intergenic distance, and intron/exon sizes are important indicators. Early analysis of human DNA indicated that the ‘average’ size of a coding region is around 1500 base pairs, and the average size of a gene is 10-15 kbp. Gene density is about one gene per 40-45 kbp, and the intergenic distance is around 25--30 kbp. However, as we have already seen, gene structure in eukaryotes can be very complex, and thus using ‘average’ estimates is a little misleading. The advancements in genome sequencing have generated more data and we enter what is sometimes called the ‘post-genomic era’.

The transcriptome and proteome: We finish this look at molecular biology by introducing two more”-omes” to complement the ‘genome’. These terms have become widely used as researchers begin to delve into the bioinformatics of cells. The transcriptome refers to the population of transcripts at any given point in a cell’s life. This expressed subset of the genomic information will be determined by many factors affecting the status of the cell. There will be general ‘housekeeping’ genes for basic maintenance of cell function, but there may also be tissue-specific genes being expressed, or perhaps developmentally regulated genes will be ‘on’ atthat particular point. Analysis of the transcriptome therefore gives a good snapshot of what the cell is engaged in at that point in time.The proteome is a logical extension to the genome and transcriptome in that it represents the population of proteins in the cell. The proteome the transcriptome reflects the genome, although there will be some transcripts that may not be translated efficiently, and there may be proteins that persist in the cell when their transcripts have been removed from circulation. Many biologists now accept that an understanding of the proteome is critical in developing a full understanding of how cells work. Some even consider the proteome as the ‘holy grail’ of cell biology, comparing it with the search for the unifying theory in physics. The argument is that, if we understand how all the proteins of a cell work, then surely we have a complete understanding of cell structure and function? As with most things in biology, this is unlikely to be a simple

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process, although the next few years will provide much excitement for biologists as the secrets of gene expression are revealed in more detail

RECOMBINANT DNA TECHNOLOGY

The field of modern biotechnology started when recombinant human insulin produced by bacteria was first marketed in the United States in 1982. The effort leading up to this landmark event began in the early 1970s when research scientists developed protocols to construct new types of bacterial plasmids or (vectors), by cutting out and pasting pieces of DNA together to create a new piece of DNA (recombinant DNA) that could be inserted into host bacterium such as E. coli. We have also observed that yeasts can be made to produce vaccines such as hepatitis B, plants having special properties such as resistance to certain diseases, pests, and herbicides and plants with superior nutritive qualities can be generated very efficiently. These achievements of genetic engineering came as a result of recombinant DNA technology. Rec DNA technology is one of the few techniques that made conventional biotechnology into “Modern Biotechnology.” Paul Berg, Herbert Boyer, Annie Change, and Stanley Cohen are Some of the pioneers of recombinant DNA technology and hence biotechnology.

Rec DNA (rec DNA or rDNA although rDNA can also refer to ribosomal DNA) technology covers a number of techniques that enable the construction of new combinations of DNA in the lab for different purposes. The rDNA molecule thus constructed can be introduced into an appropriate host cell, where it can be multiplied and generate many copies. This is what is known as gene cloning or DNA cloning. In this chapter we will examine the basic tools, methodologies, and applications of recombinant DNA techniques in various fields of biological research.Gene cloning can generate unlimited copies of a DNA molecule (e.g., recombinant DNA) by replication in a host cell. It was first developed in 1970. In addition, there are techniques such as Polymerase chain reaction (PCR) which can also be used to amplify DNA without necessarily growing them in a cell.

The following are some of the major applications of rec DNA technique: Genetic mapping DNA sequencing Mutation studies Transformation Genetic engineering Recombinant DNA libraries Restriction enzyme site analysis Analysis of gene transcripts

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The goal of rDNA is gene cloning to generate large amounts of pure DNA that can be manipulated and studied. The following are the basic steps involved in the process of the rec DNA technique for gene cloning:

Isolate DNA from organism (e.g. extraction) Cut DNA with restriction enzymes to generate DNA fragments of desired size, which may

have a specific type of DNA sequence or gene that has to be cloned. Ligate or splice each piece of DNA into a cloning vector to create a recombinant DNA

molecule. A cloning vector is an artificial DNA molecule, capable of replicating in a host organism (e.g., bacteria).

Transform recombinant DNA (cloning vector + DNA fragment) into a host that will replicate and transfer copies to progeny

Fig. A diagram showing the basic steps of molecular cloning

Basically, if one understands the answers to following questions, one will understand what cloning is:1 How is the DNA isolated from the cells?2 How is the DNA cut into pieces?3 How are the pieces of DNA put back together?4 How do we monitor each of these steps?

DNA isolation from the cell:

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Recombinant DNA technology and Cloning are usually treated as synonymous terms although this may not be entirely true. We henceforth talk about Cloning:The first step in cloning is to isolate a sufficient amount of the DNA (vector or chromosomal). The piece of DNA to be cloned is called an insert, while the carrier is called a vector, which is usually plasmid (small circular non-chromosomal DNA found in bacteria).

Isolation of plasmid DNA will be examined first.In laboratory jargon, it is called the mini-prep preparation. In the general scheme, cells containing the plasmid are grown to a high cell density, lysed gently to release the plasmid DNA which is then isolated and concentrated. When the cells are growing, the antibiotic for which the plasmid contains an antibiotic resistance gene is added in the growth medium. This ensures that the cells containing the plasmid will thrive. Without the antibiotic selection, an unstable plasmid (i.e. one without a par function) can be lost from the cell population in a few generations. Cells can be lysed by several different methods depending on the size of the plasmid, the strain of E. coli the plasmid will be isolated from, and how the plasmid DNA will be purified. Most procedures use EDTA to chelate the Mg++ associated with the outer membrane and destabilize the outer membrane. Lysozyme is added to digest the peptidoglycan and detergents are frequently used to solubilize the membranes. RNases are added to degrade the large amount of RNA found in actively growing E. coli cells. The RNase gains access to the RNA after the EDTA and lysozyme treatments. This mixture is centrifuged to pellet intact cells and large pieces of cell debris. The supernatant contains a mixture of soluble components, including the plasmid, and is known as a lysate. The methods used to purify the plasmid DNA from the cell lysate rely on the small size and abundance of the plasmid DNA relative to the chromosome, and the covalently closed circular nature of plasmid DNA. Most plasmids exist in the cytoplasm of the cell as highly supercoiled circular DNA molecules. The lysate is treated with sodium hydroxide to denature all of the DNA, and with detergent, sodium dodecyl sulphate (SDS). The pH is then abruptly lowered, causing the SDS to precipitate along with denatured chromosomal DNA, membrane fragments, and other cell debris. Most of the plasmid DNA re-anneals to form dsDNA because each strand is a covalently closed molecule and the two strands are not physically separated from each other. The small size of the plasmid allows the plasmid molecules to remain in suspension. The supernatant, which contains plasmid DNA, proteins, and other small molecules, is then subjected to purification. The most common protocol uses a column resin that binds DNA. A small amount of the resin is mixed with the plasmid-containing supernatant and the plasmid-bound resin is collected in a small column. The remaining cell components are washed away and the plasmid is eluted from the resin. This procedure is quick, simple, and reliable and can be easily carried out on a large number of samples. Many modifications of this procedure have been devised.

Chromosomal DNA isolation

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For chromosomal DNA, cells are lysed in much the same way as for plasmid DNA isolation. The cell lysate is phenol-extracted or otherwise treated to remove all of the proteins. The chromosomal DNA is precipitated as long threads. The chromosomal DNA is very fragile and breaks easily. Column based kits exist for chromosomal DNA isolation although care must be taken to prevent breakage of the DNA during isolation because of its length.

Tools of recombinant DNA technologyGeneration of recombinant DNA molecules require (i) the DNA fragment to be cloned (known as the insert DNA) and (ii) a vehicle DNA (vector) to carry the insert into a host cell for multiplication, The process therefore involves cutting and stitching together of DNA fragments to make a construct. The following major enzymes are important in that regard.

Enzyme Type # 1.

Restriction Endonucleases:

A commonly used tool in molecular biology is restriction endonucleases. Restriction endonu-cleases, otherwise known as restriction enzymes, are molecular scissors that can cut double-stranded DNA at a specific base-pair sequence.

Each type of restriction enzyme recognizes a characteristic sequence of nucleotides that is known as its recognition site. Researchers can use these enzymes to cut DNA in a predictable and precise manner.

Discovery of Restriction Endonucleases:

Scientific discoveries often have their origin in seemingly unimportant observation that receive little attention by researchers before their general significance is appreciated. In case of genetic engineering, the original observation was that bacteria use enzymes to defend themselves against viruses.

Most organisms eventually evolve means of defending themselves from predators and parasites, and bacteria are no exception. Among the natural enemies of bacteria are bacteriophages, viruses that infect bacteria and multiply within them.

At some point, they cause the bacterial cells to burst, releasing thousands more viruses. Through natural selection, some types of bacteria have acquired powerful weapons against these viruses; they contain enzymes called restriction endonucleases that fragment the viral DNA as soon as it enters the bacterial cell.

Many restriction endonucleases recognize specific nucleotide sequences in a DNA strand, bind to the DNA at those sequences, and cleave the DNA at a particular place within the recognition sequence. Why don’t restriction endonucleases cleave the bacterial cells’ own DNA as well as that of the viruses?

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The answer to this question is that bacteria modify their own DNA, using other enzymes known as methylases to add methyl (—CH3) groups to some of the nucleotides in the bacterial DNA. When nucleotides within a restriction endonuclease’s recognition sequence have been methylated, the endonuclease cannot bind to that sequence.

Consequently, the bacterial DNA is protected from being degraded at that site. Viral DNA, on the other hand, has not been methylated and, therefore, is not protected from enzymatic cleavage.

Types of Restriction Endonucleases:

All restriction enzymes fall into one of three classes, basing upon their molecular structure and need for specific co-factors.

I. Class I Endonucleases:

These have a molecular weight around 300,000 Daltons, Eire composed of non-identical subunits, and require Mg2+, ATP (adenosine triphosphate), and SAM (S-adenosylmethionine) as cofactors for activity. Not used in RDT experiments.

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II. Class II Endonucleases:

These are much smaller, with molecular weights in the range of 20,000 to 100,000 Daltons. They have identical sub-units and require only Mg2+ as a cofactor (Nathans and Smith, 1975). Only class II endonucleases are used in RDT experiments due to their site specific cleavage action.

III. Class III Endonucleases:

These are large molecules, with a molecular weight of around 200,000 Daltons, composed of non- identical sub-units. These enzymes differ from enzymes of other two classes in that they require both Mg2+ and ATP but not SAM as co-factors. Class III endonucleases are the rarest of three types. Not used in RDT experiments.

Nomenclature:

As a large number of restriction enzymes have been discovered, a uniform nomenclature system is adopted to avoid confusion.

This nomenclature was first proposed by Smith and Nattens in 1973.

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1. The first letter of restriction enzymes (RE) should be from first letter of the species name of organism from which the enzyme is isolated.

The letter should be written in capitals and italics, e.g., RE from E. coli will have E as starting letter.

2. The second and third letters of RE should be from the first and second letters of genus name of the organism. The letter should be written in lower case and should be in italics, e.g., RE from E coli will have Eco as starting words.

3. If the RE is isolated from particular strain of an organism, then that should be written as fourth letter. It should be in capitals and not in italics. For example, RE from E. coli R strain will be written as Eco R.

4. If the RE isolated is the first of its kind from that particular organism, then the number I should be given. If already two REs are isolated, then number III should be given for new restriction enzymes. The number should be written in roman, e.g., the first E. coli RE should be written as Eco RI whereas the third restriction enzyme isolated from E. coli R strains should be written as Eco RIII.

Recognition Sequences:

The recognition sequences for class II endonucleases form palindromes with rotational symmetry. In a palindrome, the base sequence in the second half of a DNA strand is the mirror image of sequence in its first half (Fig. 4.2). But in a palindrome with rotational symmetry, the base sequence in the first half of one strand of a DNA double helix is the mirror image of second half of its complementary strand (Fig 4.3).

Thus in such palindromes, the base sequence in both the strands of a DNA duplex reads the same when read from the same end (either 5′ or 3′) of both the strands.

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Mechanism of Action of Restriction Endonucleases:

EcoRI can be taken as an example of class II endonucleases and its works can be seen. When the restriction endonuclease encounters its respective restriction site sequence (5′ GAATTC 3′), it cleaves each backbone between the G and the closest A base residues.

Once the cuts have been made, the resulting fragments are held together only by relatively weak hydrogen bonds that hold the complementary bases to each other. The weakness of these bonds allows the DNA fragments to separate from each other. Each resulting fragment has a protruding 5′ end composed of unpaired bases (Fig. 4.4).

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Other enzymes also create cuts in the DNA backbone in the same manner, which results in protruding 3′ ends. Protruding ends—both 3′ and 5’—are sometimes called ‘sticky ends’ because they tend to bond with complementary sequences of bases.

In other words, if an unpaired length of bases (5′ A A T T 3′) encounters another unpaired length with the sequence (3′ T T A A 5′) they will bond to each other—they are ‘sticky’ for each other. Ligase enzymes are then used to join the phosphate backbones of two molecules.

The cellular origin, or even the species origin, of the sticky ends does not affect their stickiness. Any pair of complementary sequences will tend to bond, even if one of the sequences comes from a length of human DNA, and the other comes from a length of bacterial DNA.

In fact, it is this quality of stickiness that allows the production of recombinant DNA molecules (molecules which are composed of DNA from different sources and have given birth to a powerful technology and industry) the genetic engineering or the recombinant DNA.

Examples of some other restriction enzymes, their mode of cutting and generation of 5′ overhangs and 3′ overhangs, are illustrated in (Fig. 4.5). Sticky ends (also called cohesive ends or overhanging heads) are useful for DNA cloning because complementary sequences anneal and can be ligated directly by DNA ligase.

If two different DNA samples cleaved with the same type of restriction enzymes are mixed together in the presence of DNA ligase, a recombinant DNA molecule can be generated. This is possible because of the presence of same type of sticky ends.

The complementary sequences of sticky ends from the unrelated DNA samples will anneal to-gether and are finally joined by the DNA ligase enzyme to form the recombinant DNA molecule.

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Some restriction enzymes, on the other hand, cut both the strands of a DNA molecule at the same site so that the resulting termini or ends have blunt or flush ends (Fig. 4.6) in which the two strands end at the same point. The blunt ends also can effectively be utilized as recombinants followed by some end point modifications.

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As the list of restriction enzymes grew and their recognition sequences were identified, it was found in some cases that more than one enzyme could recognize the same sequence. RJ Roberts conferred the term isoschizomer (same cutter) on restriction enzymes that recognized the same DNA sequence.

 

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Star Activity:

Sometimes restriction enzymes recognize and cleave the DNA strand at the recognition site with asymmetrical palindromic sequence; for example, Bam HI cuts at the sequence GA TCC, but under extreme conditions, as in low ionic strength it will cleave in any of the following sequences NGA TCC, GPOA TCC, GGNTCC. Such an activity of the restriction endonucleases is called star activity.

Difficulties Associated with Restriction Digestion:

There are certain limitations for restriction endonucleases.

Those are as follows:

1. Different enzymes can generate the same ends. For example, Sau3A1 and BamHI produce GATC−. If these two enzymes are used then they will form the same ends which will not ligate later.

2. Restriction endonuclease preparation should be free from nucleases, otherwise the ends produced by theses enzymes can be degraded by exonucleases.

3. Most restriction endonucleases are very stable when stored at -20°C in the recommended storage buffer. Exposure to temperature above -20°C can decrease the activity of these enzymes.

4. The specificity of some restriction endonuclease is affected by the type of buffer used.

5. Secondary structures in DNA often interfere with recognition or cleavage by endonuclease.

Works of Reference for Restriction Digestion:

Nowadays we get a complete online database for every available restriction endonuclease. REBASE provides a current review of restriction enzymes, whether and where they can be obtained, a list of publications concerning the enzymes, and much more. At this site, you can search DNA sequences for their open reading frames and cleavage sites can be searched out.

Uses of Restriction Endonucleases:

Restriction enzymes have been used for sequence analysis, cloning and amplifying DNA. DNA from animal viruses bacteriophages contains 5,000 to 50,000 base pairs. It is important to know the primary structure of DNA, i.e., the sequence of bases, for decoding the information stored in genes, for understanding gene structure and regulation at molecular level.

The discovery of restriction enzymes was a major breakthrough in sequence analysis of DNA. By using combinations of different restriction enzymes it is possible to hydrolyse large DNA molecules into fragments less than 300 base pairs in length.

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These fragments can then be used for sequence analysis and are arranged into a physical map of the chromosome. This is a slow and laborious process. The mapping of entire 5,000 base pair DNA of the virus SV40 into some 100 fragments has taken several years. Complete sequence analysis of the fragments would take much longer.

The DNA fragments produced by restriction endonucleases can covalently be linked in vitro to linear plasmid DNA or to lambda phage DNA. The recombinant DNA species produced can be inserted into E. coli by transformation.

Each transformed cell can then be grown as a separate clone. By this method a complex genome can be broken down into thousands or millions of pieces, and each piece is isolated to form a separate clone.

Enzyme Type # 2.

Alkaline Phosphatases:

Alkaline phosphatase is a glycoprotein with two identical subunits. The cohesive ends of broken plasmids, instead of joining with foreign DNA, join the cohesive end of the same DNA molecules and get re-circularized. To overcome this problem the restricted plasmid is treated with an enzyme, alkaline phosphatase, that digests the terminal phosphoryl group.

The restriction fragments of the foreign DNA to be cloned are not treated with alkaline phosphatase.

Therefore, the 5′ end of foreign DNA fragment can covalently join to 3′ end of the plasmid. The recombinant DNA thus obtained has a nick with 3′ and 5′ P hydroxy ends. Ligase will only join 3′ and 5′ ends of recombinant DNA together if the 5′ end is phosphorylated.

Thus, alkaline phosphatase and ligase prevent re-circularization of the vector and increase the frequency of production of recombinant DNA molecules. The nicks between two 3′ ends fragment and vector DNA are repaired inside the bacterial cells during the transformation.

Enzyme Type # 3.

Reverse Transcriptase:

Many times we do not get our gene of interest rather its mRNA. In this case reverse transcriptase enzyme can be used to prepare a double stranded DNA (our gene of interest) from the available single-stranded mRNA (template) by a process called reverse transcription.

Reverse transcriptase enzyme is also called RNA dependent DNA polymerase. These enzymes are present in most of the RNA tumour viruses and retroviruses.

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Enzyme Type # 4.

Polynucleotide Kinase:

Kinase is the group of enzyme, which adds a free pyrophosphate (PO4) to a wide variety of substrates like proteins, DNA and RNA. It uses ATP as cofactor and adds a phosphate by breaking the ATP into ADP and pyrophosphate. It is widely used in molecular biology and genetic engineering to add radio-labelled phosphates. In RDT experiments mostly T4

polynucleotide kinase is used.

Enzyme Type # 5.

DNA Ligase:

Recombinant DNA experiments require the joining of two different DNA segments or fragments in vitro. The ends generated by some RE will be either cohesive (sticky) or blunt. The cohesive ends will anneal (join) themselves by forming hydrogen bonds. But the segments annealed thus are weak and do not withstand experimental conditions.

To get a stable joining, the DNA should be joined by using an enzyme called ligase. In the case of blunt ends we use linker or adaptors for successful ligation.

There are two types of DNA ligases:

(a) T4 DNA Ligase:

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Naturally coded by T4 bacteriophage. The catalytic activity of the enzyme requires the presence of ATP as cofactor and Mg++. This is predominantly used in RDT experiments.

(b) NAD+ dependent DNA Ligase:

Naturally found in E. coli. Uses NAD+ as a co-factor and only found in bacteria.

Mechanism of Action:

The cofactor is first spited (ATP→ AMP + 2Pi) and then AMP binds to the enzyme to form the enzyme-AMP complex. This complex then binds to the nick or break (with 5′ −PO4 and 3′ −OH) and makes a covalent bond in the phosphodiester chain. The ligase reaction is carried out at 4°C for better results.

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Enzyme Type # 6.

Nucleases:

Nucleases are group of enzymes which cleave or cut the genetic material (DNA or RNA). These enzymes are further classified into two types based upon the substrate on which they act. Nucleases which act on or cut the DNA are classified as DNases, whereas those which act on the RNA are called RNases.

DNases are further classified into two types based upon the position where they act. DNases that act on the ends or terminal regions of DNA are called exonucleases and those that act at a non-specific region in the centre of the DNA are called endonucleases.

Exonucleases require a DNA strand with at least two 5′ and 3′ ends. They cannot act on DNA which is circular. Endonucleases can act on circular DNA and do not require any free DNA ends (i.e., 5 or 3 end). Exonucleases release nucleotides, whereas endonucleases release short segments of DNA. The frequently used nucleases in the experiments of RDT are Exonuclease III and bacteriophage exonuclease.

Enzyme Type # 7.

Terminal Deoxynucleotide Transferases (TDNT):

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This is a polymerase which adds nucleotides at 3′-OH end (like Klenow fragment) but does not require any complementary sequence and does not copy any DNA sequence (unlike Klenow fragment). Terminal deoxynucleotide transferase (TDNT) adds nucleotide whatever comes into its active site and it does not show any preference for any nucleotide.

Enzyme Type # 8.

DNA Polymerase:

These are mostly used when we are carrying out the cloning of the recombinant DNA in the prokaryotic host cells like E. coli. Then we fill the gaps in duplexes by stepwise addition of nucleotide to 3′ ends.

Visualizing the cloning process

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At each step of the cloning process, what is happening to the DNA molecules in the test-tube can be monitored using a technique called gel electrophoresis. In this technique, a gel (Fig. below) containing small wells is cast. The DNA is loaded into the wells and the gel is placed in an elec-tric current. Because DNA is negatively charged, it will move in the gel towards the positive pole. The DNA migrates or moves in the electric current based on size and shape. The larger a DNA molecule, the slower it moves. The more compact, or supercoiled a piece of DNA, the faster it moves.The gel can be made from several different polymers, depending on the specifics of the experi-ment. Agarose forms a matrix that will separate DNA molecules from ~500bp up to entire chro-mosomes (several million base pairs). If an electric current is constantly applied to an agarose gel from only one direction, agarose gels will separate DNA from ~500bp to ~25,000bp. An alterna-tive polymer, polyacrylamide, can be used to separate molecules a few base pairs in length to ap-proximately 1000 bp.Once the DNA has been separated in the gel, the gel is immersed in a solution containing ethid-ium bromide. If ultraviolet light is used to illuminate the gel, the ethidium bromide that is bound to the DNA will fluoresce, indicating the presence of bands of DNA. Each band is composed of DNA molecules that are similar in size and shape. For example, when plasmid DNA is extracted from the cell, the majority of it is supercoiled. Supercoiled DNA migrates very fast in an agarose gel (Fig. below).

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CLONING VECTORS

Common features

Most cloning vectors found in laboratories are synthetic substances (manufactured as opposed to being isolated from microorganisms). They have key features that have made their use in molecular biology successful.

Vectors can be used (i) to amplify copies of a particular DNA of interest (i) to simply transfer/ shuttle DNA of interest between cells (sub-cloning) (iii) to express a particular gene to yield a protein or some specific result.

In the case of expression vectors, the main purpose of these vehicles is the controlled expression of a particular gene inside a convenient host organism (e.g. E. coli). It is necessary to insert the target DNA into a site under the control of a particular promoter. Some commonly used promoters are T7 promoters, lac promoters (bla promoter) and cauliflower mosaic virus's 35s promoter (for plant vectors).

To allow for convenient and favorable insertions, most cloning vectors have nearly all of their naturally occurring restriction sites engineered out of them and a synthetic multiple cloning site (MCS) inserted that contains many restriction sites. MCSs allow for insertions of DNA into the vector to be targeted and possibly directed in a chosen orientation. A selectable marker, such as an antibiotic resistance [e.g. beta-lactamase] is often carried by the vector to allow the selection of positively transformed cells (Screening). All plasmids must carry a functional origin of replication (ORI).Some other possible features present in cloning vectors are: vir genes for plant transformation, integrase sites for chromosomal insertion, lacZα fragment for α complementation and blue-white selection, and/or reporter genes in frame with and flanking the MCS to facilitate the production of recombinant proteins [e.g. fused to the Green fluorescent protein (GFP) or to the glutathione S-transferase].

Fig: Diagram of a simple cloning vector derived from a plasmid, a circular, double-stranded DNA molecule that can replicate within an E. coli cell. Plasmid vectors are ~1.2–3 kb in length and contain a replication origin (ORI) sequence and a gene that permits selection, usually by conferring resistance to a particular drug. Here the selective gene is ampr; it encodes the enzyme β-lactamase, which inactivates ampicillin. Exogenous DNA can be inserted into the

bracketed region without disturbing the ability of the plasmid to replicate or express the ampr gene.

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Fig: A map of a commercially available plasmid pBR322 (4Kb) showing a number of restriction sites and regions encoding for resistance to ampicillin (ampr) and tetracy-cline (tetr), and origin of replication (ori)

Other components of cloning vectors:

Promoter: used to drive transcription of the vector's transgene. The promoter is the most impor-tant component of an expression vector. This is because the promoter controls the very first stage of gene expression (attachment of an RNA polymerase enzyme to the DNA) and determines the rate at which mRNA is synthesized. The amount of recombinant protein obtained therefore de-pends to a great extent on the nature of the promoter carried by the expression vector. Although most E. coli promoters do not differ much from these consensus sequences (e.g., TT-TACA instead of TTGACA), a small variation may have a major effect on the efficiency with which the promoter can direct transcription. Strong promoters are those that can sustain a high rate of transcription; strong promoters usually control genes whose translation products are re-quired in large amounts by the cell (Figure 13.6a). In contrast, weak promoters, which are rela-tively inefficient, direct transcription of genes whose products are needed in only small amounts. Clearly an expression vector should carry a strong promoter, so that the cloned gene is tran-scribed at the highest possible rate.A second factor to be considered when constructing an expression vector is whether it will be possible to regulate the promoter in any way. Two major types of gene regulation are recognized in E. coli—induction and repression. An inducible gene is one whose transcription is switched on by addition of a chemical to the growth medium; often this chemical is one of the substrates for

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the enzyme coded by the inducible gene. In contrast, a repressible gene is switched off by addi-tion of the regulatory chemical.Gene regulation is a complex process that only indirectly involves the promoter itself. However, many of the sequences important for induction and repression lie in the region surrounding the promoter and are therefore present in an expression vector. It may therefore be possible to extend the regulation to the expression vector, so that the chemical that induces or represses the gene normally controlled by the promoter is also able to regulate expression of the cloned gene. This can be a distinct advantage in the production of recombinant protein. For example, if the recom-binant protein has a

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Examples of promoters used in expression vectorsSeveral E. coli promoters combine the desired features of strength and ease of regulation. Those most frequently used in expression vectors are as follows:l The lac promoter (Figure 13.8a) is the sequence that controls transcription of the lacZ gene coding for b-galactosidase. The lac promoter is induced by isopropylthiogalactoside (IPTG, p. 80), so addition of this chemical into the growth medium switches on transcription of a gene in-serted downstream of the lac promoter carried by an expression vector.The trp promoter (Figure 13.8b) is normally upstream of the cluster of genes coding for several of the enzymes involved in biosynthesis of the amino acid tryptophan. The trp promoter is re-pressed by tryptophan, but is more easily induced by 3-b-indoleacrylic acid.The tac promoter (Figure 13.8c) is a hybrid between the trp and lac promoters. It is stronger than either, but still induced by IPTG.The EPL promoter (Figure 13.8d) is one of the promoters responsible for transcription of the e DNA molecule. ePL is a very strong promoter that is recognized by the E. coli RNA polymerase, which is subverted by e into transcribing the bacteriophage DNA. The promoter is repressed by the product of the ecI gene. Expression vectors that carry the ePL promoter are used with a mu-tant E. coli host that synthesizes a temperature-sensitive form of the cI protein (p. 40). At a low temperature (less than 30°C) this mutant cI protein is able to PThe T7 promoter (Figure 13.8e) is specific for the RNA polymerase coded by T7 bacteriophage. This RNA polymerase is much more active than the E. coli RNA polymerase, which means that a gene inserted downstream of the T7 promoter will be expressed at a high level. The gene for the T7 RNA polymerase isnot normally present in the E. coli genome, so a special strain of E. coli is needed, one which is lysogenic for T7 phage. Remember that a lysogen contains an inserted copy of the phage DNA in its genome (p. 19). In this particular strain of E. coli, the phage DNA has been altered by placing a copy of the lac promoter upstream of its gene for the T7 RNA polymerase. Addition of IPTG to the growth medium therefore switches on synthesis of the T7 RNA polymerase, which in turn leads to activation of the gene carried by the T7 expression vector.

Genetic markers: Genetic markers for viral vectors allow for confirmation that the vector has in-tegrated with the host genomic DNA.

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Antibiotic resistance: Vectors with antibiotic-resistance open reading frames allow for survival of cells that have taken up the vector in growth media containing antibiotics through antibiotic selection.

Polylinker: is a region within a cloning vector that contains several restriction enzyme recogni-tion sites. Each restriction site occurring in the polylinker is unique i.e it occurs only once in the plasmid. Polylinker is also called a multiple cloning site or MCS.

Epitope: Vector contains a sequence for a specific epitope that is incorporated into the expressed protein. Allows for antibody identification of cells expressing the target protein.

Reporter genes: Some vectors may contain a reporter gene that allow for identification of plas-mid that contains inserted DNA sequence. An example is lacZ-α which codes for the N-terminus fragment of β-galactosidase, an enzyme that digests galactose. This operon forms the basis of

Blue/white screening:

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The concept of alpha-complementation is important because it is a quick, easy, 1-step process of determining whether a transformed bacterial colony has plasmid+insert or not (The tetracycline resistance of pBR322 works, it is just time consuming). The key to alpha-complementation is the fact that the lac-Z gene product (B-galactosidase) is a tetramer, and each monomer is made of two parts - lacZ-alpha, and lacZ-omega. Researchers determined that if the alpha fragment was deleted, the omega fragment is non-functional; however, alpha fragment functionality can be restored in-trans via plasmid. Hence, then name alpha-complementation. What is needed for this to work as a cloning technique is a strain of E. coli that has the deletion of the lac Z-alpha (lacZ DM15 works well as a genotype), and a plasmid with the lacZ-alpha fragment as the scorable marker (such as pBluescript or pCR2.1). If plain plasmid is successfully transformed into a cell, then the cell will express functional B-galactosidase. However, if the plasmid+insert is transformed into a cell, then it will express non-functional B-galactosidase (the lac Z-alpha will be disrupted with the insert gene product). Plate the cells out onto selection media based on the selectable marker, IPTG (induces lac repressor to disengage), and X-gal (chromogenic substrate that yields blue product when cleaved by B-galactosidase) and the white colonies (non-functional B-galactosidase) are the ones with plasmid + insert; the blue ones have plain plasmid. Another method of accomplishing the same task is to completely delete the lac operon from the chromosome, but introduce lac-Z-omega fragment on an F’ fertility factor. This accomplishes the same task as above, with a plasmid with lac-Z-alpha scorable marker.

1. The lacZ’ gene from E. coli is present in the plasmid DNA, not in the chromosomal DNA2. Multiple cloning site (MCS) is adjacent to lacZ’ gene and encodes β-galactosidase (β-gal)

protein3. Any fragment of DNA, when inserted into the MCS, will disrupt the lacZ’ gene and β-gal

expression4. β-gal protein is normally detectable by adding X-gal substrate and a Blue dye is produced

(turns cells blue)5. Colonies turn blue only if β-gal protein present. Otherwise they remain white. So cells

without insert will form blue colonies (blue spots on the agar plate)

Where does the blue come from? Normally lactose sugar is catabolized into galactose + glucose (cell energy) When clear substrate X-gal is added to cells it is cleaved by β-gal enzyme Cleavage of X-gal generates blue/indigo precipitation in the cells

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“Empty” Vector Lacks a DNA Insert and no insert means the lacZ’ gene is intact and expresses β-gal protein Cells turn blue

Other commonly used reporters include green fluorescent protein and luciferase.

Targeting sequence: Expression vectors may include encoding for a targeting sequence in the finished protein that directs the expressed protein to a specific organelle in the cell or specific lo-cation such as the periplasmic space of bacteria.

Protein purification tags: Some expression vectors include proteins or peptide sequences that al-lows for easier purification of the expressed protein. Examples include polyhistidine-tag, glu-tathione-S-transferase, and maltose binding protein. Some of these tags may also allow for in-creased solubility of the target protein. The target protein is fused to the protein tag, but a pro-tease cleavage site positioned in the polypeptide linker region between the protein and the tag al-lows the tag to be removed later.

Common DNA Vector (Commercially available): Plasmids – accepts <10 kb of DNA λ-Phage – accepts of DNA ~20 kb of DNA Cosmid – accepts ~45 kb of DNA BAC – accepts ~200 kb of DNA YAC – accepts ~1000 kb of DNA

Many more ….

Select the Vector that best fits your sub-cloning! Large DNA inserts are hard to work with Small inserts are limited by size

Plasmids

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Figure. 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.  

Fig. pBR322

Plasmids are circular, double-stranded DNA molecules that naturally exist in bacteria and in the nuclei of some eukaryotic cells such as yeast cells (e.g the 2µ plasmid).  They can replicate inde-pendently of the host cell.  The size of plasmids ranges from a few kb to near 100 kb.A plasmid vector is for commercial applications is made from natural plasmids by removing un-necessary segments and adding essential sequences.

The term 'plasmid' was introduced by American molecular biologist Joshua Lederberg. Plasmids are considered as transferrable genetic elements or 'replicons'. They are actually naked DNA. Plasmids are important tools in genetic and biotechnology labs where they are commonly used to

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multiply or express particular genes. Plasmids are also used to make large amounts of proteins.Plasmids encoding Zinc Finger Nucleases are used to deliver therapeutic genes to a preselected chromosomal site with a frequency higher than that of random integration. Mainly there are two types of plasmids: conjugative and non conjugative. Conjugative plasmids have tra-genes (tra-transfer) and can perform conjugation. Non conjugative plasmids cannot perform conjugation. There is an intermediate class of plasmid called mobilizable plasmid. Mobilizable plasmid can carry only a subset of genes required for transfer. They can parasitize a conjugative plasmid transferring at high frequency only in its presence.

 Based on function plasmids can be of five types: 

F/Fertility plasmid for conjugation. R/Resistant plasmid which contains genes that provides resistance to antibiotics. It also

helps bacteria in producing pilus. Col plasmid which contain genes that code for bacteriocin (toxins produced by bacteria to

inhibit the growth of similar or closely related bacterial strains) Degradative plasmid which help in the digestion of unusual substances like toluene. Virulence plasmid which is responsible for pathogenicity.

 Bacteria contain one or more plasmids in them and are present in the form of a number of copies in each cell. Each bacteria have several mechanisms to maintain high copy number of plasmids . 

Relaxed plasmids have high copy number Stringent plasmids have low copy number.

 Plasmid DNA may appear in one of the five conformations, which run at different speeds in a gel during electrophoresis. The different plasmid conformations are listed below in the order of elec-trophoretic mobility . 

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1. Nicked Open-Circular DNA ,which has one strand cut.

 2. Relaxed Circular DNA is fully intact with both strands uncut, but has been enzymati-

cally relaxed.

  

3. Linear DNA has free ends, either because both strands have been cut, or because the DNA was linear in vivo. 

 4. Super coiled (or Covalently Closed-Circular) DNA is fully intact with both strands un-

cut, and with a twist built in, resulting in a compact form. 

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 5. Super coiled Denatured DNA is like super coiled DNA, but has unpaired regions that

make it slightly less compact; this can result from excessive alkalinity during plasmid preparation.

 

The main components and properties of plasmid cloning vectors are:Origin of replication ( ori ) This is the site where DNA replication is initiated. The presence of the ori allows the plasmid to replicate independently of the cell. An example is the ori from plasmid pMB1 which only in E. coli but not other organisms. However, broad-host-range plasmids capable of replicating in more than 1 species exist. Exam-ples include IncQ plasmids which replicate in many Gram negative bacteriaShuttle vectors contain origins of replication derived from different species and integrated into one plasmid e.g. YEp (yeast episomal plasmid) contains the pMB1 ori and the S. cerevisiae au-tonomously replicating sequence (ARS). The plasmid is thus capable of replicating in both species.

Most plasmids replicate at the same rate as genome replication and normally exist at 1 -2 copies per cell only. This are said to be stringently regulated. However, high copy number plas-mids replicate independently of the genomic DNA and are said to have a relaxed control. They can therefore exist at more than 15 copies per cell. The pMB1 ori  is an example of relaxed. The copy numbers of the plasmid can be amplified by chloramphenicol. Chloramphenicol interferes with protein synthesis and thus prevents genomic DNA replication which requires protein syn-thesis.pMB1 replication on the other hand, does not require protein synthesis and the plasmid replica-tion continues yielding 100s to 1000s of plasmid copis per cell. Copy numbers can also be in-creased by deleting regulatory genes – e.g.in pUC plasmids. This leads to > 500 copies per cell.

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plasmid incompatibility: normally different types of plasmids sharing similar ori cannot be present in same cell. This is because copy number regulatory systems will cause plasmids with same ori to interfere with each other’s replication. The copy number for both plasmids will be kept low and its statistically unlikely both will be maintained in the same cell.

Marker genes for selection and/or screeningPlasmids also require a selection marker. This works by killing of cells that lack specific gene for example those lacking antibiotic resistance genes e.g amicillin resistance gene. Only cells containing plasmid with antibiotic resistance genes form colonies. If for instance they contain gene for ampicillin resitance, then they produce β-lactamase that breaks down ampicillin and they can therefore form colonies.

TransmissibilityMany wild type plasmids are transmissible by conjugation (bacterial mating). This requires a se-quence for the tra  genes which codes for pili (the sex stylus of bacteria), the mob gene whose products nick DNA at nic/bom site and starts rolling circle replication. In cloning vectors, this transmission capability is usually disabled by deletion of some or all conjugation functions. This prevents recombinant DNA transfer to wild bacterial strains. For ex-ample the pBR322 has no tra or mob regions and cannot transfer itself. It does have the nicand bom sites and can thus be transferred if tra & mob are present in same cell e.g. on “helper plas-mid. On the other hand, pUC vectors lack nic & bom sites and therefore cannot be transferred, even with tra & mob present.

Promoters for gene expressionCommercial plasmids may also contain controllable promoters before the multiple cloning site. This allows for the expression of cloned genes in E. coli.

Lambda (λ)Phage VectorsLambda (λ)-Phages are viruses that can infect bacteria.  The major advantage of the lambda phage vector is its high transformation efficiency, about 1000 times more efficient than the plasmid vector.• Double-stranded, linear DNA ~50 kb• Packaged into head which transfects E. coli cells• Ends of DNA have cohesive (cos) sites which allow DNA to circularize in E. coli• Contains 25 kb of non-essential DNA which can be deleted. Deleted from lab strains• Accepts DNA fragments ~20 kb (can vary) that replaces the non-essential DNA

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Fig. Schematic drawing of the DNA cloning using λ phages as vectors.  The DNA to be cloned is first inserted into the λ DNA, replacing a nonessential region.  Then, by an in vitro assembly system (described below), the λ virion carrying the recombinant DNA can be formed.  The λ genome is 49 kb in length which can carry up to 25 kb foreign DNA.

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Fig. The assembly process of the lambda virion.

The extreme ends of the λ DNA are known as COS sites, each is single stranded, 12 nucleotides long.  Because their sequences are complementary to each other, one end of λ DNA may base-pair with the other end of a different λ DNA, forming concatamers.  The two ends of a λ DNA may also bind together, forming a circular DNA.  In the host cell, the λ DNA circularizes be-cause ligase may seal the nicks on the COS sites. In the assembly process of virions, two proteins Nu1 and A can recognize the COS site, directing the insertion of the λ DNA between them into an empty head.  The filled head is then attached to the tail, forming a complete λ virion.  The whole process normally takes place in the host cell.  However, to prepare the λ virion carrying recombinant l DNA, the following in vitro assembly system is commonly used. Proteins Nu1 and A are encoded by the genes in the λ genome. If the two genes are mutated, λ DNA cannot be packaged into the pre-assembled head.  Because tails attach only to filled heads, the cell will accumulate separate empty heads and tails, which can then be extracted.  When the extract is mixed with recombinant λ DNA and proteins Nu1 and A, the complete λ virion carry-ing recombinant λ DNA will be assembled.

Cosmid vectorsThe cosmid vector is a combination of the plasmid vector and the COS site which allows the tar-get DNA to be inserted into the λ head.  It has the following advantages:

High transformation efficiency. The cosmid vector can carry up to 45 kb whereas plasmid and l phage vectors are limited

to 25 kb.They contain one or two lambda “cos sites”. (The cos site and the DNA between 2 cos sites is the only requirement for DNA to be packaged into a phage particle).

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Fig.  Cloning by using cosmid vectors.  (a) In addition to ampr, ORI, and polylinker as in the plasmid vector, the cosmid vector also contains  a COS site.  (b) After cosmid vectors are cleaved with restriction enzyme, they are ligated with DNA fragments.  The subsequent assembly and transformation steps are the same as cloning with λ phage

How is DNA cloned into a cosmid vector?1. Use a polylinker.2. Package this DNA into phage particles like phage DNA.3. Propagate the cosmid as a plasmid (a plasmid with a selector gene).4. Purify the cosmids as if they were plasmids.The head of a phage can accept between 40 and 55 kb of DNA and as most cosmids are about 5 kb in length, between 33 and 48 kb of DNA can be cloned in these vectors.

BAC (bacterial artificial chromosome) Vectors:• Is a large, circular double stranded DNA• Has E. coli chromosomal origin of replication (oriC)• Can replicate and are stable in E. coli• Contain antibiotic resistance gene(s)• Accepts DNA fragments ~200 Kb in size

Why are they called artificial chromosomes?

YAC (yeast artificial chromosome) Vectors:The yeast artificial chromosome (YAC) vector is capable of carrying a large DNA fragment (up to 2 Mb), but its transformation efficiency is very low.

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Fig.  Cloning by the yeast artificial chromosome (YAC) vector.

• Large, linear dsDNA• Has a centromere (that ensures that YAC is inherited in progeny)• Has telomeres at ends of chromosomes (stabilize)• Contain antibiotic resistance gene (s)• Accepts DNA fragments ~1000 kb in size

Essential components of YAC vectors Centromers (CEN), telomeres (TEL) and autonomous replicating sequence (ARS) for

proliferation in the host cell.

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ampr for selective amplification and markers such as TRP1 and URA3 for identifying cells containing the YAC vector.

Recognition sites of restriction enzymes (e.g., EcoRI and BamHI)Procedure

1. The target DNA is partially digested by EcoRI and the YAC vector is cleaved by EcoRI and BamHI.

2. Ligate the cleaved vector segments with a digested DNA fragment to form an artificial chromosome.

3. Transform yeast cells to make a large number of copies.

What is a genome? What is genomic sequencing?There are different Vectors to Sub-clone human genome:• A typical human genomic DNA is about 3.3 x 109bp long• The genomic DNA is cut into fragments and ligated into vectors. Below are some of the capacities and number of clones generated per type of vector. So the larger the vector, the fewer the number of clones to analyze.

Why is it necessary to sub-clone genomic DNA piece into Vectors? It is difficult to manipulate large genomic DNA It is easier to study small piece with one geneBut then this has the disadvantage that to know the sequence of the entire genome, one has to have a mechanism of reassembling all these genomic libraries (hence the field bioinformatics).

A typical cloning for genomic sequencing experiment may involve• Cut genomic DNA into fragments to insert into a vector (plasmid, phage, BAC, YAC, etc…)This produces many (1000 - 100,000) different plasmids each with different gDNA insertOne can use X-Gal reporter (blue/white) which on the basis of color one can determine if plasmid contains DNA insert or notOne then eliminates all E. coli colonies with “empty” plasmids vectors (without insert).

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PLANT TRANSFORMATIONIntroduction of exogenous DNA into a plant cell can be transient (whereby there is no incorpora-tion of exogenous DNA into the genome) or stable (there is incorporation of introduced exoge-nous DNA into the genome). Transformation of multicellular organisms cannot directly transform every cell but rather leads to a mosaic of recombinant and non-recombinant cells. To have all the cells of an organism trans-formed, one cell is transformed, which then regenerates an entire organism.

Agrobacterium tumefaciens (updated nomenclature Rhizobium radiobacter):

It is a natural tool for plant transformation. It is a soil resident gram positive bacterium which causes gall tumor in dicotyledonous plants. Agrobacterium tumefaciens (or A. tumefaciens) is an alpha-proteobacterium of the family Rhizobiaceae, which includes the nitrogen fixing legume symbionts. Unlike the nitrogen fixing symbionts, tumor producing Agrobacterium are pathogenic and do not benefit the plant. The wide variety of plants affected by Agrobacterium makes it of great concern to the agriculture industry. It is a serious pathogen of walnuts, grape vines, stone fruits, nut trees, sugar beets, horse radish and rhubarb.

Genes involved in crown gall disease are not present on the chromosome of A. tumefaciens but on a large extrachromosomal plasmid called the Ti (tumor-inducing) plasmid. The Ti plasmid in-tegrates a segment of its DNA, known as T-DNA, into the chromosomal DNA of its host plant cells. The T-DNA is transferred via an F-pilus. The T-DNA contains genes for encoding en-zymes that cause the plant to create specialized amino acids which the bacteria can metabolize, called opines. Opines are a class of chemicals that serve as a source of nitrogen for A. tumefa-ciens, but not for most other organisms. The specific type of opine produced by A. tumefaciens -infected plants is nopaline

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T-DNA transfer process is activated when Agrobacterium gets in contact with damaged plant tis-sue• T-DNA is nicked at the RB and is replicated to the LB and moved into the plant cell – this is catalyzed by products of vir genes

In details, the T-DNA must be cut out of the circular plasmid. A VirD1/D2 complex nicks the DNA at the left and right border sequences. The VirD2 protein is covalently attached to the 5' end. VirD2 contains a motif that leads to the nucleoprotein complex being targeted to the type IV secretion system (T4SS).

In the cytoplasm of the recipient cell, the T-DNA complex becomes coated with VirE2 proteins, which are exported through the T4SS independently from the T-DNA complex. Nuclear localization signals, or NLS, located on the VirE2 and VirD2 are recognised by the importin alpha protein, which then associates with importin beta and the nuclear pore complex to transfer the T-DNA into the nucleus. VIP1 also appears to be an important protein in the process, possibly acting as an adapter to bring the VirE2 to the importin. Once inside the nucleus, VIP2 may target the T-DNA to areas of chromatin that are being actively transcribed, so that the T-DNA can integrate into the host genome.

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How can we engineer the Ti plasmid to make it useful?• Delete auxin and cytokinin genes• Retain vir genes, LB & RB, ori• Ti plasmid is huge (~120 kb) – need to make it smaller. The vir genes and T-DNA can be on separate plasmids.• Only left and right borders (LB & RB) are required for T-DNA to be transferred

Steps in plant transformation1. Propagate binary vector in E. coli2. Isolate binary vector from E.coli and engineer (introduce a foreign gene)3. Re-introduce engineered binary vector into E. coli to amplify4. Isolate engineered binary vector and introduce into Agrobacterium containing a modified (smaller) Ti plasmid5. Infect plant tissue with engineered Agrobacterium(T-DNA containing the foreign gene gets inserted into a plant cell genome)In each cell T-DNA gets integrated at a different site in the genome• Each cell is hemizygous for the insertion – only one of the homologous chromosomes gets the insertion• Consequences of the insertion:- Foreign DNA is inserted- Insertional mutagenesis(does not kill the cell – the organism is diploid!)

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ReportersTo determine which cells have taken –up insert DNA., there are two types of marker genes: a selectable marker (Antibiotics ) and a marker for screening. A reporter gene (often simply reporter) is a gene that researchers attach to a regulatory sequence of another gene of interest in cell culture, animals or plants. A selectable marker is really only used to kill off all cells that didn't take up the gene (plasmid) while leaving the rest that did, especially when transfection rates are very low. A reporter gene is much more versatile because it is there to 'report' something. This can be whether a gene was taken up, how high the gene is expressed, but also whether a signal pathway was turned on, whether the cell is going into apoptosis, anything you manage to engineer into the gene that tells you something you are interested in. (If you use a FACS you can even isolate all cells expressing a GFP reporter gene making the reporter gene a sort of selectable marker.)A particularly striking foreign DNA that has been used a reporter is the enzyme luciferase, which is originally from fireflies. The enzyme catalyzes the reaction of a chemical called luciferin with ATP; in this process, light is emitted, the same process that occurs in fireflies that glow in the dark. A transgenic tobacco plant expressing the luciferase gene also will glow in the dark when wa-tered with a solution of luciferin. Luciferase gene is useful as a reporter to monitor the function of any gene during development. In other words, the upstream promoter sequences of any gene of interest can be fused to the luciferase gene and put into a plant by T-DNA. Then the luciferase gene will follow the same developmental pattern as the normally regulated gene does, but the lu-ciferase gene will announce its activity prominently by glowing at various times or in various tis-sues, depending on the regulatory sequence.Other genes used as reporters in plants are the bacterial GUS (β-glucuronidase) gene, which turns the compound X-Gluc to blue, and the bacterial lac (β-galactosidase) gene, which turns X-Gal blue. Cells in which these reporters are expressed turn blue, and this blueness can be easily seen either by the naked eye or under the microscope.Transgenic plants carrying any one of a variety of foreign genes are in current use, and many more are in development. Not only are the qualities of plants themselves being manipulated, but, like microorganisms, plants are also being used as convenient “factories” to produce proteins en-coded by foreign genes.

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The Luciferase Reporter Vectors have been specially constructed to report the binding activity of an individual TF. Multiple copies of the cis-acting enhancer element have been inserted into each vector upstream of a minimal TA promoter, the TATA box from the Herpes simplex virus thymi-dine kinase promoter. This promoter sequence drives expression of the luciferase gene (luc). The backbone of the vector contains an antibiotic resistance gene for cloning purposes, an origin of replication, and an f1 origin for single-stranded DNA production.To assess in vivo TF binding activity, the Luciferase Vector is first transfected into cells. If de-sired, an antibiotic resistant vector can be cotransfected to establish a stable cell line. After a set amount of time, the cells are lysed and subjected to the standard luciferase assay. Luminescence is detected and measured by a luminometer or scintillation counter. The resulting data can be used to quantify TF activity.

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MAKING PROTEINS

The synthesis and purification of proteins from cloned genes is one of the most important aspects of genetic manipulation, particularly where valuable therapeutic proteins are concerned. Many such proteins have already been produced by recombinant DNA (rDNA) techniques and are already in widespread use.

In most cases a bacterial host cell is used for the expression of cloned genes. However, there are special cases where eukaryotic host is required. In the cell, eukaryotic proteins are often subjected to post-translational modification (PTM), and therefore this modifications must be achieved in an in vitro expression system to produce a functional protein.

As we already mentioned, to express cloned DNA, the gene must be inserted into a vector that has a suitable promoter and which can be introduced into an appropriate host such as E. coli. Although E. coli is not ideal for expressing eukaryotic genes, many of the problems of using E. coli can be overcome by constructing the recombinant so that the expression signals are recognised by the host cell. Such signals include promoters and terminators for transcription, and ribosome binding sites (Shine--Dalgarno sequences) for translation. Alternatively, eukaryotic host such as the yeast S. cerevisiae, or mammalian cells in tissue culture, may be more suitable for certain proteins. For eukaryotic proteins, the coding sequence is usually derived from a cDNA clone of the mRNA. This is particularly important if the gene contains introns, as these will not be processed out of the primary transcript in a prokaryotic host. When the cDNA has been obtained, a suitable vector must be chosen. Although there is a very wide variety of expression vectors, there are two main categories, those which (i) produce native proteins or (ii) fusion proteins. Native proteins are synthesized directly from the N terminus of the cDNA, whereas fusion proteins contain short, N-terminal amino acid sequences encoded by the vector. In some cases these may be important for protein stability or secretion and are thus not necessarily a problem. However, such sequences can be removed if the recombinant is constructed so that the fusion protein contains a methionine residue at the point of fusion. The chemical cyanogen bromide (CNBr) can be used to cleave the protein at the methionine residue, thus releasing the desired peptide. A major problem with this approach occurs if the protein contains one or more internal methionine residues, as this will result in unwanted cleavage by CNBr. When constructing a recombinant for the synthesis of a fusion protein, it is important that the cDNA sequence is inserted into the vector in a position that maintains the correct reading frame.

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Yeast expression systemsThe yeast Saccharomyces cerevisiae is favoured microbial eukaryote in the development of recombinant DNA technology. Whilst S. cerevisiae is still useful for gene expression studies and protein production, other yeasts may offer advantages in terms of growth characteristics, yield of heterologous protein, and large-scale fermentation characteristics. Other species include Schizosaccharomyces pombe, Pichia pastoris, Hansela polymorpha, Kluyveromyces lactis, and Yarrowia lipolytica. These demonstrate many of the characteristics of bacteria with respect to ease of use -- they grow rapidly on relatively inexpensive media, and a range of different mutant strains and vectors is available for various applications. In some cases scale-up fermentations present some difficulties compared to bacteria, but these can usually be overcome by careful design and process monitoring. Yields of heterologous proteins of around 12 g L−1 (10--100 times more than in S. cerevisiae) have been obtained using P. pastoris, which can be grown on methanol as sole carbon source. In this situation growth is regulated by the enzyme alcohol oxidase, which has a low specific activity and is consequently overproduced in these cells, making up around 30% of total soluble protein. By placing heterologous genes downstream from the alcohol oxidase promoter (AOX1), high levels of expression are achieved. One of the advantages of using yeast as opposed to bacterial hosts is that proteins are subjected to PTMs such as glycosylation. In addition, there is usually a higher degree of ‘authenticity’ with respect to three-dimensional conformation and the immunogenic properties of the protein. Thus, in a situation where the biological properties of the protein are critical, yeasts may provide a better product than prokaryotic hosts.

The baculovirus expression systemBaculoviruses are rod-shaped (baculum in Latin means 'stick') dsDNA viruses found mainly in insects. The most common baculovirus used for expression studies is Autographa californica multiple nuclear polyhedrosis virus (AcMNPV), which relies on the lepidopteran species Spodoptera frugiperda and Trichoplusiani as host insects. AcMNPV particles surround themselves with a protective matrix consisting of the protein polyhedrin, which permits survival in the environment and efficient spread to new hosts.

Under the control of the extremely strong promoter pPolh, polyhedrin is expressed at extremely high levels (up to 50% of all cellular protein) at the end of the baculovirus life cycle. The baculovirus expression system makes use of the fact that in cell culture a polyhedrin coat is not essential for virus propagation and thus heterologous proteins can be expressed under the control of the pPolh promoter .Baculoviruses infect insects and do not appear to infect mammalian cells. Thus, any system based on such viruses is more preferred on the basis of biosafety. During normal infection of insect cells, virus particles are packaged within what is known as polyhedra. These are nuclear inclusion bodies composed mostly of the protein polyhedrin. This is synthesised late in the virus infection cycle and can represent as much as 50% of infected cell protein when fully expressed. In culture, the virus does not require polyhedrin and therefore the polyhedrin gene can be used in

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the construction of an expression vector, as it encodes a late-expression protein that is dispensable and can be synthesized in large amounts. The baculovirus genome is a circular double-stranded DNA molecule. The genome size is from 88 to 200 kb, depending on the particular virus, and the genome is therefore too large for direct manipulation in vitro. Thus, insertion of foreign DNA into the vector has to be accomplished by using an intermediate known as a transfer vector. These are based on E. coli plasmids and carry the promoter for the polyhedrin gene with the essential expression signals. The cloned gene for expression is inserted into the transfer vector, and the recombinant is used to co-transfect insect cells with non-recombinant viral DNA. Homologous recombination between the viral DNA and the transfer vector results in the generation of recombinant viral genomes, which can be selected for and used to produce the protein of interest. Systems based on baculoviruses demonstrate transient gene expression, in which the protein of interest is synthesised as part of the infection cycle of the viral-based vector system. More recently, stable insect-cell expression systems have also been developed, in which insect cells can be used for continuous expression of protein. One disadvantage of using insect cells as opposed to bacteria or yeast is that they require more complex growth media for maintenance and production. The cells are also less robust than the microbial cells and, thus, require careful handling if success is to be achieved.

Mammalian cell linesWhere the expression of recombinant human proteins is concerned, it might seem obvious that a mammalian host cell would be a better system than bacteria, eukaryotic microbes, or insect cells. However, the use of such cell lines in protein production presents some problems. As with insect cell lines, the media required to sustain growth of mammalian cells are complex and expensive, and the cells are relatively fragile when compared with microbial cells, particularly where large-scale fermentation is involved. Downstream processing (DSP), which refers to the purification of the final product from the host cell after expression, is usually more difficult for mammalian cells. Despite these difficulties, many vectors are now available for protein expression in mammalian cells. They are often based on a viral system, vectors utilise selectable markers (often drug-resistance markers) and have promoters that enable expression of the cloned gene sequence. Common promoters are based on simian virus (SV40) or cytomegalovirus.

Protein engineeringOne of the most exciting applications of gene manipulation lies in the field of protein engineering. This involves altering the structure of proteins via alterations to the gene sequence and has become possible because of the availability of a range of techniques, as well as a deeper understanding of the structural and functional characteristics of proteins. This has enabled workers to pinpoint the essential amino acid residues in a protein sequence; thus, alterations can be carried out at these positions and their effects studied. The desired effect might be alteration of the catalytic activity of an enzyme by modification of the residues around the active site, an improvement in the nutritional status of a storage protein, or an improvement in the stability of a protein used in industry or medicine. Proteins that have been engineered by the incorporation of

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mutational changes have become known as muteins. There are two types of approach that can be used to engineer proteins. These are sometimes called rational design and directed evolution.

Examples of biotechnological applications of rDNA technologyWe will consider some examples of the types of products that can be produced using rDNA tech-nology in biotechnological processes. This is a rapidly developing area for many biotechnology companies, with large-scale investment in both basic research and in development to production status. This aspect of gene manipulation technology is likely to become increasingly important in the future, particularly in medicine and general healthcare, with many diverse products being brought to market.

Production of manufactured enzymesThe commercial production and use of enzymes is already a well established part of the biotech-nology industry. Enzymes are used in brewing, food processing, textile manufacture, the leather industry, washing powders, medical applications, and basic scientific research , to name just a few examples. Previously enzymes used to be prepared from natural sources, but in recent years there has been a move towards the use of enzymes produced by recDNA methods whenever possible. Recombinant enzymes can sometimes be engineered so that their character-istics fit the criteria for a particular process better than the natural enzyme, which increases the fidelity and efficiency of the process. In the food industry, one area that has involved the use of recombinant enzyme is the production of cheese. In cheese manufacture, rennet (also known as rennin, chymase, or chymosin) has been used as part of the process. Chymosin is a protease that is involved in the coagulation of milk casein following fermentation by lactic acid bacteria. It was traditionally prepared from animal (bovine or pig) or fungal sources. In the 1960s the Food and Agriculture Organisation of the United Nations predicted that a shortage of calf rennet would develop as more calves were reared to maturity to satisfy increasing demands for meat and meat products. Today there are six sources for natural chymosin -- veal calves, adult cows and pigs, and the fungi Rhi-zomucor miehei, Endothia parasitica and Rhizomucor pusillus. Chymosin is now also avail-able as a recombinant-derived preparation from E. coli, Kluyveromyces lactis, and Aspergillus niger. Recombinant chymosin was first developed in 1981, approved in 1988, and is now used to prepare around 90% of hard cheeses in the UK. Another example of recombinant-derived proteins in consumer products is the use of enzymes in washing powder. Proteases and lipases are commonly used to assist cleaning by degradation of protein and lipid-based staining. A recombinant lipase was developed in 1988 by Novo Nordisk A/V (now known as Novozymes). The company is the largest supplier of enzymes for commercial use in cleaning applications. Their recombinant lipase was known as LipolaseTM, which was the first commercial enzyme developed using rDNA technology and the first li-pase used in detergents. A further development involved an engineered variant of Lipolase called Lipolase Ultra, which gives enhanced fat removal at low wash temperatures.

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Recombinant bovine somatotropin (rBST)The story of recombinant bovine somatotropin (rBST) illustrates some of the problems that may be encountered once the scientific part of the process has been achieved. BST is also known as bovine growth hormone and is a naturally occurring protein that acts as a growth promoter in cattle. Milk production can be increased substantially by administering BST and, thus, it was an attractive target for cloning and production for use in the dairy industry. The BST gene was one of the first mammalian genes to be cloned and expressed in the 80s, using bacterial cells for production of the protein. Thus, the production of rBST at a commer-cial level, involving the basic science and technology transfer stages, was achieved without too much difficulty. A summary of the process is shown in Fig. 10.4. In the USA, the Food and Drug Administration (FDA) is the central regulatory body, and in 1994 approval was given for the commercial distribution of rBST, marketed by Monsanto under the trade name PosilacTM. At that time the European Union did not approve the product, but this was partly for socioeco-nomic reasons (increasing milk production was not necessary) rather than for any concerns about the science. Evaluation of evidence at that time suggested that milk from rBST-treated cows was identical to normal untreated milk, and it was therefore unlikely that any negative ef-fects would be seen in consumers. The effects of rBST must be considered in three different con-texts -- the effect on milk production, the effects on the animals themselves, and the possible ef-fects on the consumer. Milk production is usually increased by around 10--15% in treated cows, although yield increases of much more than this have been reported. Thus, from a dairy herd management viewpoint, use of rBST would seem to be beneficial. However, as is usually the case with any new development that is aimed at ‘improving’ what we eat or drink, public con-cern grew along with the technology. The concerns fuel a debate that is still ongoing and is at times emotive. One area that is hotly debated is the effect of rBST on the cows themselves. Ad-ministering rBST can produce localised swelling at the site of injection and can exacerbate problems with foot infections, mastitis, and reproduction.

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Fig. 10.4. Production of recombinant bovine growth hormone (rBST). (a) A plasmid vector is prepared from E. coli and cut with a restriction enzyme (RE). (b) The BST gene coding sequence is ligated into the plasmid to generate the recombinant, which produces rBST protein in the cell following transformation. Scale-up to commercial production is shown in (c), and with product approval granted, administration can begin. The whole process from basic science to market usu-ally takes several/many years from start to finish, with a large amount of investment capital re-quired. From Nicholl (2000), Cell and Molecular Biology, Advanced Higher Monograph Series, Learning and Teaching Scotland. Reproduced with permission.

Therapeutic products for use in human healthcareIn addition to the actual treatment of conditions, the area of medical diagnostics is a large and fast-growing sector of the biotechnology market, with rDNA technology involved in many as-

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pects. Recombinant DNA products for use in medical therapy can be used for replacement or supplementation of human proteins that may be absent or ineffective in patients with a particu-lar illness. Second, proteins can be used in specific disease therapy, to alleviate a disease state by intervention. Third, the production of recombinant vaccines is an area that is developing rapidly and that offers great promise. Some examples of therapeutic proteins produced using rDNA technology are listed in Table 10.1. We will consider examples Table 10.1. The widespread condition diabetes mellitus (DM) is usually caused either by -cells in the islets of Langerhans in the pancreas fail to produce adequate amounts of the hormone insulin, or by target cells being non-responsive to the hormone. Many millions of people worldwide are af-fected by DM, and the World Health Organisation estimates that the global incidence will dou-ble by 2025. Some 3-6% of people in the UK and the USA have diabetes, although there is thought to be significant under-diagnosis. Sufferers are classed as having either type I DM (for-merly known as insulin-dependent DM or IDDM) or type II DM (formerly non-insulin-de-pendent DM or NIDDM). Some 10% of patients have type I DM, with around 90% having type II. There are also some other variants of the disease that are much less common. Type I patients obviously require the hormone, but many type II patients also use insulin to manage their condi-tion. Delivery of insulin is achieved by injection (traditional syringe or ‘pen’-type devices), in-fusion using a small pump and catheter, or inhalation of powdered insulin.

The structure of insulin. Amino acids are represented by circles. The A chain (21 amino acids) and B chain (30 amino acids) are held together by disulphide linkages between cysteine residues (filled circles).

Insulin is composed of two amino acid chains, the A-chain (acidic, 21 amino acids) and B-chain (basic, 30 amino acids). When synthesised naturally, these chains are linked by a further 30--amino acid peptide called the C-chain. This 81 amino acid precursor molecule is known as proinsulin. The A- and B-chains are linked together by disulphide bonds between cysteine residues, and the proinsulin is cleaved by a protease to produce the active hormone. Insulin was the first protein to be sequenced by Frederick Sanger in the mid 1950s. As DM is caused by a problem with a normal body constituent (insulin), therapy falls into the category of replacement or supplementation. Banting and Best developed the use of insulin therapy in 1921, and for the next 60 or so years diabetics were dependent on natural sources of insulin, with the attendant problems of supply and quality. In the late 1970s and early 1980srDNA technology enabled sci-entists to synthesize insulin in bacteria, with the first approvals granted by 1982. Recombinant-derived insulin is now available in several forms and has a major impact on diabetes therapy.

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One of the most widely used forms is marketed under the name HumulinTMby the Eli Lilly Com-pany. In an early method for the production of recombinant insulin, the insulin A- and B-chains were synthesized separately in two bacterial strains. The insulin A and B genes were placed under the control of the lac promoter, so that expression of the cloned genes could be switched on by using lactose as the inducer. Following purification of the A- and B-chains, they were linked together by a chemical process to produce the final insulin molecule. The process is shown in Fig. 10.6. A development of this method involves the synthesis of the entire proinsulin polypeptide (shown in Fig. 10.7) from a single gene sequence. The product is converted to in-sulin enzymatically.

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There are many recombinant proteins for use in specific disease therapy. One example of this type of protein is tissue plasminogen activator (TPA). This is a protease that occurs naturally

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and functions in breaking down blood clots. TPA acts on an inactive precursor protease called plasminogen, which is converted to the active form called plasmin. This protease attacks the clot by breaking up fibrin, the protein that is involved in clot formation. TPA is used as a treat-ment for heart attack victims. If administered soon after an attack, it can help reduce the dam-age caused by coronary thrombosis. Recombinant TPA was produced in the early 1980s by the company Genentech using cDNA technology. It was licensed in the USA in 1987, under the trade name Activase, for use in treatment of acute myocardial infarction. It was the first recombinant-derived therapeutic protein to be produced from cultured mammalian cells, which secrete rTPA when grown under appro-priate conditions. The amount of rTPA produced in this way was sufficient for therapeutic use; thus, a major advance in coronary care was achieved. Further uses were approved in 1990 (for acute massive pulmonary embolism) and 1996 (for acute ischaemic stroke). The final group of recombinant-derived products are vaccines. There are now many vaccines available for animals, and the development of human vaccines is also beginning to have an im-pact in healthcare programmes. One vaccine that has been produced by rDNA methods is the hepatitis B vaccine. The yeast S. cerevisiae is used to express the surface antigen of the hep-atitis B virus (HBsAg), under the control of the alcohol dehydrogenase promoter. The protein can then be purified from the fermentation culture and used for inoculation. This removes the possibility of contamination of the vaccine by blood-borne viruses or toxins, which is a risk if natural sources are used for vaccine production. A further development in vaccine technology involves using transgenic plants as a delivery mechanism. This area of research and development has tremendous potential, particularly for vaccine delivery in underdeveloped countries where traditional methods of vaccination may not be fully effective because of cost and distribution problems. The attraction of having a vaccine-containing banana or tomato is clear, and development and trials are currently under way for a variety of plant vaccines. The use of gene manipulation techniques in the biotechnology industry is a major developing area of applied science. In addition to the scientific and engineering as-pects of the work, the financing of biotechnology companies is an area that presents its own risks and potential rewards -- for example, a new drug may take 10--15 years to develop, at a cost of several hundreds of millions of pounds. The stakes are therefore high, and many fledgling companies fail to survive their first few years of operation. Even established and well-financed companies are not immune to the risks associated with the development of a new and untried product. The next few years will certainly be interesting for this sector of the applied science in-dustry.

Medical and forensic applications of gene manipulationThe diagnosis and treatment of human disease is one area in which genetic manipulation is be-ginning to have a considerable effect. As outlined in Chapter 11, many therapeutic proteins are now made by recombinant DNA (rDNA) methods, and the number available is increasing steadily. Thus, the treatment of conditions by recombinant-derived products is already well es-tablished. In this chapter we will look at how the techniques of gene manipulation impact more

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directly on medical diagnosis and treatment, and we will also examine the use of rDNA technol-ogy in forensic science. Recent progress in both of these areas is of course closely linked to our increasing knowledge of the human genome, and new developments in medical and forensic ap-plications will undoubtedly appear as we continue to decipher the genome.

Diagnosis and characterization of medical conditionsGenetically based diseases (often called simply ‘genetic diseases’) represent one of the most im-portant classes of disease, particularly in children. A disorder present at birth is termed a congen-ital abnormality, and around 5% of newborn babies will suffer from a serious medical problem of this type. In most of these cases there will be a significant genetic component in the aetiol-ogy(cause) of the disease. It is estimated that about a third of primary admissions to paediatric hospitals are due to genetically based problems, whilst some 70% of cases presenting more than once are due to genetic defects. In addition to genetic problems appearing at birth or in child-hood, it seems that a large proportion of diseases presenting in later life also have a genetic cause or predisposition. Thus, medical genetics, in its traditional non-recombinant form, has already had a major impact on the diagnosis of disease and abnormality. The development of molecular genetics and rDNA technology has not only broadened the range of techniques available for di-agnosis, but has also opened up the possibility of novel gene-based treatments for certain condi-tions.

Diagnosis of infectionIn addition to genetic conditions that affect the individual, rDNA technology is also important in the diagnosis of certain types of infection. Normally, bacterial infection is relatively simple to di-agnose, once it has taken hold. Thus, the prescription of antibiotics may follow a simple investi-gation by a general practitioner. A more specific characterisation of the infectious agent may be carried out using microbiological culturing techniques, and this is often necessary when the in-fection does not respond well to treatment. Viral infections may be more difficult to diagnose, al-though conditions such as Herpes infections are usually obvious. Despite traditional methods be-ing applied in many cases, there may be times when these methods are not appropriate. Infection by the human immunodeficiency virus (HIV) is one case in point. The virus is the causative agent of acquired immune deficiency syndrome (AIDS). The standard test for HIV infection requires immunological detection of anti-HIV antibodies, using techniques such as ELISA (en-zyme linked inmmunosorbent assay, sometimes known as the enzyme immunoassay), West-ern blot, and IFA (indirect immunofluorescence assay). However, these antibodies may not be detectable in an infected person until weeks after initial infection, by which time others may have been infected. A test such as this, where no positive result is obtained even though the individual is infected, is a false negative. The use of DNA probes and PCR technology circumvents this problem by assaying for nucleic acid of viral origin in the T-lymphocytes of the patient, thus per-mitting a diagnosis before the antibodies are detectable. Other examples of the use of rDNA technology in diagnosing infections include tuberculosis (caused by the bacterium Mycobac-

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terium tuberculosis), human papilloma virus infection, and Lyme disease (caused by the spirochaete Borrelia burgdorferi).

TRANSGENIC ANIMALS

The generation of transgenic animals is one of the most complex aspects of genetic engineering, both in terms of technical difficulty and in the ethical problems that arise. Many people, who accept that the genetic manipulation of bacterial, fungal, and plant species is beneficial, find difficulty in extending this acceptance when animals (particularly mammals) are involved. The need for sympathetic and objective discussion of this topic by the scientific community, the media, and the general public are likely to present one of the great challenges in scientific ethics over the next few years. Why transgenic animals? Genetic engineering has already had an enormous impact on the study of gene structure and expression in animal cells, and this is one area that will continue to develop. Cancer research is one obvious example, and current investigation into the molecular genetics of the disease requires extensive use of gene manipulation technology. In the field of protein production in biotechnology, the synthesis of many mammalian-derived recombinant proteins is often best carried out using cultured mammalian cells, as these are sometimes the only hosts that will ensure the correct expression of such genes. Cell-based applications such as those outlined above are an important part of genetic engineering in animals. However, the term ‘transgenic’ is usually reserved for whole organisms, and the generation of a transgenic animal is much more complex than working with cultured cells. Many of the problems have been overcome using a variety of animals, with early work involving amphibians, fish, mice, pigs, and sheep.Transgenics can be used for a variety of purposes, covering both basic research and biotechnological applications. The study of embryological development has been extended by the ability to introduce genes into eggs or early embryos, and there is scope for the manipulation of farm animals by the incorporation of desirable traits via transgenesis. The use of whole organisms for the production of recombinant protein is a further possibility, and this has already been achieved in some species. The term pharm animal or pharming (from pharmaceutical) is sometimes used when talking about the production of high-value therapeutic proteins using transgenic animal technology. When considered on a global scale, the potential for exploitation of transgenic animals would appear to be almost unlimited. Achieving that potential is likely to be a long and difficult process in many cases, but the rewards are such that a considerable amount of money and effort has already been invested in this area.

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Producing transgenic animals There are several possible routes for the introduction of genes into embryos, each with its own advantages and disadvantages. Some of the methods are (1) direct transfection or retroviral infection of embryonic stem cells followed by nuclear transfer into an embryo at the blastocyst stage of development; (2) retroviral infection of early embryos; (3) direct microinjection of DNA into oocytes, zygotes, or early embryo cells; (4) sperm-mediated transfer; (5) transfer into unfertilised ova; and (6) physical techniques such as biolistics or electrofusion.The technique of nuclear transfer is sometimes associated with a transgenesis protocol. Early success was achieved by injecting DNA into one of the pronucleiof a fertilised egg, just prior to the fusion of the pronuclei (which produces the diploid zygote). This approach led to the production of the celebrated ‘supermouse’ in the early 1980s, which represents one of the milestones of genetic engineering. The experiments that led to the ‘supermouse’ involved placing a copy of the rat growth hormone (GH) gene under the control of the mouse metallothionine (mMT) gene promoter. To create the ‘supermouse’, a linear fragment of the recombinant plasmid carrying the fused gene sequences (MGH) was injected into the male pronuclei of fertilized eggs (linear fragments appear to integrate into the genome more readily than circular sequences). The resulting fertilized eggs were implanted into the uteri of foster mothers, and some of the mice resulting from this expressed the GH gene. Such mice grew some 2-3 times faster than control mice and were up to twice the size of the controls. In generating a transgenic animal, it is desirable that all the cells in the organism receive the transgene. The presence of the transgene in the germ cells of the organism will enable the gene to be passed on to succeeding generations, and this is essential if the organism is to be useful in the long term. Thus, introduction of genes has to be carried out at a very early stage of development, ideally at the single-cell zygote stage. If this cannot be achieved, there is the possibility that a mosaic embryo will develop, in which only some of the cells carry the transgene. Another example of this type of variation is where the embryo is generated from two distinct individuals, as is the case when embryonic stem cells are used. This results in a chimaeric organism. In practice this is not necessarily a problem, as the organism can be crossed to produce offspring that are homozygous for the transgene in all cells. A chimaeric organism that contains the transgene in its germ line cells will pass the gene on to its offspring, which will therefore be heterozygous for the transgene (assuming they have come from a mating with a homozygous non-transgenic). A further cross with a sibling will result in around 25% of the offspring being homozygous for the transgene. In this book so far, we have been considering the topic of molecular cloning, where the aim of an experimental process is to isolate a gene sequence for further analysis and use. In organismal cloning, the aim is to generate an organism from a cell that carries a complete set of genetic instructions. From a wider public perspective, organismal cloning is seen as an issue for concern and, thus, a discussion of the topic is essential even in a book where the primary goal is to illustrate the techniques of gene manipulation. Organismal cloning can be further subdivided according to the purpose of the procedure. Where the function is to generate a ‘copy’ of the original organism, this is termed reproductive cloning. Recent advances in stem cell technology open up the possible use of cloning embryos to enable production of matched tissue types for use

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in research and potentially in the treatment of disease. This aspect of cloning is called therapeutic cloning. In 1938 Spemann published his book Embryonic Development and Induction, detailing his work. In this he proposed what he called ‘a fantastical experiment’, in which the nucleus would be removed from a cell and implanted into an egg from which the nucleus had been removed. Spemann could not see any way of achieving this, hence his caution by using the word ‘fantastic’. However, he had proposed the technique that would later become known as cloning by nuclear transfer (more fully known as somatic cell nuclear transfer or SCNT). Unfortunately, he did not live to see this attempted; the first cloning success was not achieved until 1952, eleven years after his death.

Nuclear totipotencyThe work of Spemann was an important part of the development of modern embryology - indeed, he is often called the ‘father’ of this discipline. He had proposed nuclear transfer and had demonstrated cloning by embryo splitting. The experiments with embryo splitting that had refuted Weismann’s ideas showed that embryo cells retain the capacity to form all cell types. This became known as the concept of nuclear totipotency, which is now a fundamental part of developmental genetics. A cell is said to be totipotent if it can direct the formation of all cells in the organism. If it can direct a more limited number of cell types, it is said to be pluripotent or multipotent. Extending this along the developmental timeline, a cell that is not capable of directing development under appropriate conditions is said to be irreversibly differentiated. Nuclear totipotency is in many ways self-evident, as an adult organism has many different types of cell. The original zygote genome, passed on by successive mitotic divisions, must have the capacity to generate these different cells.

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Plant development is somewhat simpler than animal development, largely because there are fewer types of cell to arrange in the developing structure. However, the concept of nuclear totipotency is just as valid in plants as it is in animals. In fact, one of the early unequivocal

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experimental demonstrations of nuclear totipotency was provided by the humble carrot in the late 1950s. Work by F. C. Steward and his colleagues at Cornell University showed that carrot plants could be regenerated from somatic (body) tissue, as shown in Fig. 14.3. This technique is now often used in the propagation of valuable plants in agriculture. The ability of plants to regenerate has of course been exploited for many years by taking cuttings and grafting -- these are essentially asexual ‘cloning’ procedures.

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