Recombinant DNA Technology
The beginning of recombinant DNA techniques has provided scientists with a new tool to address the problems of gene structure, genetic function, and genome regulation. RDT has already been remarkable in providing answers to problems that were a mystery previously. However, the most important reaps achieved are derived from applications to produce biological products that benefit mankind directly. A surplus of additional protein products from recombinant DNA are in various stages of development or research. Undoubtedly, the results will have profound effects on the practice of medicine during the coming decades. Perhaps the next important milestone of pharmaceutical applications will be the production of metabolites such as antibiotics. Essential use of life-saving antibiotics is a cornerstone of modern medicine. Successful development of recombinant DNA methods will provide tools that will help increase the fermentation yields of antibiotics and generate new antibiotic structures that are not available through traditional routes of discovery.
RDT, which is also called gene cloning or molecular cloning, is a general term that encompasses a number of experimental protocols leading to the transfer of genetic information (DNA) from one organism to another. There is no single set of methods that can be used to meet this objective; however, a recombinant DNA experiment often has the following format (Fig. 1).
Step 1: The DNA (cloned DNA, insert DNA, target DNA, or foreign DNA) from a donor organism is extracted, enzymatically cleaved (cut, or digested), and joined (ligated) to another DNA entity (a cloning vector) to form a new, recombined DNA molecule (cloning vector– insert DNA construct, or DNA construct).
Step 2: This cloning vector–insert DNA construct is transferred into and maintained within a host cell. The introduction of DNA into a bacterial host cell is called transformation.
Step 3: Those host cells that take up the DNA construct (transformed cells) are identified and selected (separated, or isolated).
Step 4: If required, a DNA construct can be created so that the protein product encoded by the cloned DNA sequence is produced in the host cell.
Recombinant DNA technology was developed from discoveries inmolecular biology, nucleic acid enzymology, and the molecular genetics of both bacterial viruses (bacteriophages) and bacterial extrachromosomal DNA elements (plasmids). However, recombinant DNA technology would not exist without the availability of enzymes that recognize specific doublestranded DNA sequences and cleave the DNA in both strands at these sequences (restriction enzymes, or restriction endonucleases). Nucleases that cut nucleic acid molecules internally are exonucleases.
The three foremost factors which affect the human life are nutrient deficiency, health risks, and environmental issues. Besides the essential requisite of a clean and riskless environment, food and health fall within the basic demands of every human. The increase in demand for food is prominently observed with an increase in the rate of the world’s population. The provision for availability of nutritious and safe food should be implemented at an affordable price. Numerous deaths resulted globally due to impaired health has been reported which brings a serious concern to the eradication of food deficiency. Every year, an approximate number of 36 million lives go deceased due to various communicable and non-communicable diseases. The list of communicable and non-communicable diseases includes AIDS/HIV, tuberculosis, malaria, cardiovascular diseases, cancer, diabetes and many more. Despite the efforts being made substantially, the ongoing global food production is significantly lower the actual human requisite. It is an inevitable fact that the health facilities fall within the lower range of medical standard in the third-world countries. Human beings are adversely affected directly or indirectly due to the rapid industrialization events which exert a negative impact on the environment. Such industrial growth has led to an uprise of the environmental pollution where the effluents of industries are diverted to the water bodies which inimically affect the aquatic system. In order to circumvent such serious issues, there arises a serious urge to introduce modern technologies.
Various traditional methodologies have been implemented to overcome issues related to agriculture, health, and environment. The methods include breeding techniques, the introduction of conventional medicines, and degradation of pollutants using traditional methods respectively. Unlike the traditional methods, the techniques in the field of genetic engineering employ the most recent approaches, such as molecular cloning and transformation, which consume less time and yield more reliable products. In fact approach of genetic engineering involves the transfers of only a limited group of desired genes to the target via reliable approaches, such as biolistic and Agrobacterium directed transformation. The plant genomes can be transformed via either homologous recombination based genetic targeting or by nuclease-dependent site-specific genome alteration.
Recombinant DNA technology has progressed in the arena of pharmacology in the improvement of the health conditions by giving rise to new vaccines and numerous pharmaceutical products. The strategical improvements in the mode of treatment are also appreciably notable due to the development of diagnostic kits, advanced monitoring devices, and novel therapeutic approaches. Usage of genetically modified bacteria in the production of synthetic human insulin and erythropoietin and generation of novel varieties of experimental mutant mice for research trials are prime examples showing the excelled growth of genetic engineering in the area of health. Likewise, the advanced strategies of genetic engineering have been tackling the serious threats to the environment. The effective turning of harmful wastes into biofuels and bioethanol, eradication of oil spills, carbon, and other toxic waste products are some of the bioconversion methods implemented via recombinant DNA technology. Detection of toxic metals like arsenic and other contaminants in drinking water have been successfully achieved via this technique. Bioremediation technology including biomining have been benefited due to developments in recombinant DNA technology.
Recombinant DNA technology has upgraded the biological field thereby bringing forth remarkable and useful changes. A doorway for innovative manipulation of animals, plants and microorganism for the production of varieties of therapeutically useful products has been generated via this technology. The newly synthesised products are found to bring effective and immediate therapeutic effect in biomedicine and medical genetics. Most of the lethal diseases occurring in a human being can be eradicated by biotechnology based pharmaceutical products which are recombinant in nature. Human life has been found to be transformed due to the development of pharmaceutical products generated via recombinant DNA technology. The FDA (U.S. Food and Drug Administration) have given green signal to more recombinant drugs in the year 1997on the contrary to previous years due to the progression of rDNA technology. The drugs developed was mainly for treatment of life threatening ailments like cancers of different organs, diphtheria, AIDS, hereditary disorders etc. Bearing in mind the fact that the plants are proficient in the multigene transfer, site-specific integration and specifically regulated gene expression are fascinatingly observed as new alternatives. There are serious challenges in plant biotechnology which requires proper focus to be addressed. The challenges include transcriptional regulation of endogenous genes, their effective expression in the new regions, and the specific control over transgene expression.
Human life is under threat due to various factors, like a limitation of food resulting in malnutrition, the rise of deadly diseases, increasing environmental problems due to emergent industrialization and expansion. But such issues can be potentially vanquished by the introduction of genetic engineering which can replace the conventional approaches. Every single facet in the arena of biological sciences has been modernized via recombinant-DNA technology. This brief elucidation is to purposefully impart an outline of the methodologies used for the detection of target genes, isolation of a gene of interest, amplification of necessary gene sequence and to finally clone the genes. The technique of recombinant-DNA mainly aims in detection, isolation, improvisation and successful re-expression of genes derived from the host. Such a cut-and-paste technology aims mainly in achieving few of the listed interests. 1) understand the underlying functions and regulatory mechanism of desired gene products, 2) to process reverse genetics in detection of novel genes whose protein products are yet to be isolated and studied, 3) rectify the genetic defects occurring endogenously such as sickle cell anaemia, 4) transformation of disease prone host with foreign genes followed by its expression. An example includes the transformation of disease susceptible plant with disease resistance genes from another host and 5) wide production of protein product for various use. An example includes the production of antibodies in tobacco plants. Gene being a basic unit of genetic information resides on uninterrupted regions of DNA strand in large number thereby making a mammalian chromosome. Either of DNA strands contains genes where they are aligned in opposite directions on a chromosome. Single gene codes for a single polypeptide.
Gel electrophoresis is a commonly used technique for resolving proteins or nucleic acids. In general, a sample of one particular type of macromolecule (protein, DNA, or RNA) is placed in a well at or near the end of a gel matrix (gel). The composition of an electrophoresis gel is a semisolid open meshwork of interlinked linear strands. A gel is cast as a thin slab with a number of sample wells. After the wells of a gel are loaded with sample, an electric field is applied across the gel, and charged macromolecules of the same size are driven together in the direction of the anode through the gel as discrete invisible bands of material. The distance that a band moves into a gel depends on the mass of its macromolecules and the size of the openings (pore size) of the gel. The smaller macromolecules travel further than the larger ones. The progress of gel electrophoresis is monitored by observing the migration of a visible dye (tracking dye) through the gel. The tracking dye is a charged, low-molecular-weight compound that is loaded into each sample well at the start of a run. When the tracking dye reaches the end of the gel, the run is terminated. The bands, which are aligned in a lane under each well, are visualized by staining the gel with a dye that is specific for protein, DNA, or RNA. Discrete bands are observed when there is enough material present in a band to bind the dye to make the band visible and when the individual macromolecules of a sample have distinctly different sizes. Otherwise, a band is not detected.
If there is little or no difference among the sizes of the macromolecules in a concentrated sample, a smear of stained material is observed. The intensity of a stained band reflects the frequency of occurrence of a macromolecule in a sample. The molecular mass (molecular weight) of a gel-fractionated macromolecule (band) is determined from a standard curve that is based on a set of macromolecules of known molecular mass (size markers) that covers the separation range of the gel system and is run in one or both of the outside lanes (calibrator lanes) of the same gel as the samples. The logarithm of the molecular mass of a size marker is related to its relative mobility (Rf) through a gel. The value of Rf is defined as the distance traveled by a band divided by the distance traveled by the tracking dye (ion front). The relationship between the logarithm of the molecular mass of each size marker and its Rf value is plotted. Then, with this standard curve, a molecular mass is calculated for each band in a lane. The units of molecular mass for proteins and double-stranded and single-stranded nucleic acids are daltons, base pairs, and bases, respectively. The size markers are included in the same gel as the samples because the extent of mobility of a macromolecule(s) varies from one electrophoretic run to the next. Polyacrylamide is the preferred gel system for separating proteins. Copolymerization of monomeric acrylamide and the cross-linker bisacrylamide forms a lattice of crosslinked, linear polyacrylamide strands. The pore size of a polyacrylamide gel is determined by the concentration of acrylamide and the ratio of acrylamide to bisacrylamide. For many applications, a protein sample is treated with the anionic detergent sodium dodecyl sulfate (SDS) before electrophoresis.
The SDS binds to proteins and dissociates most multichain proteins. Each SDS-coated protein chain has a similar charge-to-mass ratio. Consequently, during electrophoresis, the separation of the SDS– protein chains is based primarily on size, and the effect of conformation is eliminated. SDS–polyacrylamide gel electrophoresis with a 10% polyacrylamide gel resolves proteins that range from 20 to 200 kilodaltons (kDa). Agarose, which is a polysaccharide from seaweed, is used routinely as the gel matrix for the electrophoretic separation of medium-size nucleic acid molecules. A 1.0% agarose gel can resolve duplex DNA chains that range from about 600 to 10,000 bp. Specialized agarose gel electrophoresis systems are available for fractionating DNA molecules with millions of base pairs, denatured DNA, and denatured RNA. In addition, for specific purposes, polyacrylamide gels are used for separating DNA molecules. For example, DNA chains that are as small as 6 bases and that differ from each other by 1 nucleotide can be resolved with a 20% polyacrylamide gel.
The chief progression in the arena of recombinant DNA technology is the identification and confirmation of enzymes which are efficient in specific cleavage of duplex DNA into distinct reproducible fragments. Many of the prokaryotes are confirmed to be the resource where restriction enzymes can be identified. Restriction enzymes specifically recognize and cleave both strands of DNA at a definite base pair sequence. An axis of symmetry exists at most of the target cleavage sites. There can be a symmetrical cleavage occurring at the axis on either side thereby yielding overlying 5’ or 3’ single-stranded termini. The naming of restriction endonucleases is carried out as per the standard nomenclature. The first three letters represent the genus and species of the host organism. It is followed by a strain identification or a letter which indicates if the enzyme is encoded by an extrachromosomal bacteriophage or plasmid. If there exists more than one restriction endonuclease in the same host, it is denoted by a Roman numeral.
In addition to EcoRI, more than 3,700 type II restriction endonucleases with about 250 different recognition sites have been isolated from various bacteria. The naming protocol for these enzymes is the same as that for EcoRI; the genus is the capitalized letter, and the first two letters of the species name are in lowercase letters. The strain designation is occasionallyadded to the name, such as R in EcoRI, or the serotype of the source bacteriumis sometimes noted, such as d in HindIII. The Roman numerals are used to designate the order of characterization of different restriction endonucleases from the same organism. For example, HpaI and HpaII are the first and second type II restriction endonucleases that were isolated from Haemophilus parainfluenzae.
The palindromic sequences where most type II restriction endonucleases bind and cut a DNA molecule are within the recognition sites. Some restriction endonucleases digest (cleave) DNA, leaving 5′ phosphate extensions (protruding ends, or sticky ends) with recessed 3′ hydroxyl ends; some leave 3′ hydroxyl extensions with recessed 5′ phosphate ends; and some cut the backbones of both strands within a recognition site to produce blunt-ended (flush-ended) DNA molecules. The lengths of the recognition sites for different enzymes can be four, five, six, eight, or more nucleotide pairs. Because of the frequency with which their recognition sites occur in DNA, restriction endonucleases that cleave within sites of four (four-cutters) and six (six-cutters) nucleotide pairs are used for most of the common molecular-cloning protocols.
In addition, ready access to a variety of restriction endonucleases adds versatility to gene-cloning strategies. Type IIS restriction endonucleases form a subgroup of the type II category of restriction enzymes and are occasionally used for cloning and other molecular studies, such as multiplex polony sequencing. These enzymes have the fascinating feature of cutting DNA, usually in both strands, a fixed number of nucleotides away from one end of the recognition site. Moreover, any particular sequence of nucleotides may be present between the binding sequence and the cut sites. The cleavages for most type IIS restriction enzymes are staggered. For example, the FokI restriction endonuclease binds to GGATG CCTAC and cuts 9 nucleotidesdownstream on the upper strand and 13 nucleotides downstream on the lower strand, producing a recessed 3′hydroxyl end and a 4-nucleotide extension at the 5′phosphate end.
To successfully fulfil most of the aim via recombinant DNA technology, there is a requisite for the isolation of individual genes from genomic DNA of reference host. Major four steps are involved in the process of gene cloning and since there are multiple numbers of methods for each step, an investigator can opt one out of many to perform the cloning. To describe in brief, gene cloning is achieved by the restriction endonucleases mediated digesting of host DNA followed by the fusion of desired fragments into any one of four basic vector systems. The vector, in fact, is introduced into an alternative host usually bacteria in which the genes are propagated successfully. Plasmids are known to be one of the most suitable propagating vectors for DNA fragments. These cloning vehicles are extrachromosomally replicating double-stranded DNA molecules which are circular in form, and reside in bacteria. Plasmid vectors used in laboratory scale possess common features such as the origin of replication, the potency to exist in high copy number per bacterial cell, phenotype conferring gene (eg. antibiotic resistance) that permits the selection of plasmid incorporated host bacteria.
The vector and the DNA both that is required to be cloned are processed with a similar restriction endonuclease, producing molecules with well-matched ends. Commercially available plasmid vectors harbour polylinker sequences with a number of cleavage sites of restriction endonuclease and thereby these vectors will suitably receive DNA produced by digestion with multiple enzymes. The hanging end of the vector as well as the insert are capable to join one another via complementary base pairing. The DNA ligase enzyme produces covalent bonds amongst DNA strands at these positions giving rise to a closed circular recombinant plasmid. The competent bacterial cells and the plasmid are then incubated together so that the cells are able to uptake the exogenous DNA possible by multiple methods. The bacterial cells that have taken up the plasmid can be subjected to selection via plating on an agar plate having an antibiotic. The wild type bacterial host is generally vulnerable to the antibiotic and thereby only those cells that have the plasmid obtain the gene that coding for antibiotic resistance and hence are able to grow on the agar media. This type of selection distinguishes only bacteria that are able to uptake circular plasmids where as it does not differentiate plasmids inserted with recombinant DNA from recircularized vector DNA missing inserts.
Plasmids are self-replicating, double-stranded, circular DNA molecules that are maintained in bacteria as independent extrachromosomal entities. Virtually all bacterial genera have natural plasmids. Some plasmids carry information for their own transfer from one cell to another (e.g., F plasmids), others encode resistance to antibiotics (R plasmids), others carry specific sets of genes for the utilization of unusual metabolites (degradative plasmids), and some have no apparent functional coding genes (cryptic plasmids). Although they are not typically essential for bacterial cell survival under laboratory conditions, plasmids often carry genes that are advantageous under particular conditions. Plasmids can range in size from less than 1 kb to more than 500 kb. Each plasmid has a sequence that functions as an origin of DNA replication; without this site, it cannot replicate in a host cell.
Some plasmids are represented by 10 to 100 copies per host cell; these are called high-copy-number plasmids. Others maintain one to four copies per cell and are called low-copy-number plasmids. Seldom does the population of plasmids in a bacterium make up more than approximately 0.1 to 5.0% of the total DNA. When two or more different plasmids cannot coexist in the same host cell, they are said to belong to the same incompatibility group, but plasmids from different incompatibility groups can be maintained together in the same cell. This coexistence is independent of the copy numbers of the individual plasmids. Some microorganisms have been found to contain as many as 8 to 10 different plasmids. In these instances, each plasmid can carry out different functions and have its own unique copy number, and each belongs to a different incompatibility group. Some plasmids, because of the specificity of their origin of replication, can replicate in only one species of host cell. Other plasmids have less specific origins of replication and can replicate in a number of bacterial species. These plasmids are called narrow- and broad-host-range plasmids, respectively. As autonomous, self-replicating genetic elements, plasmids have the basic attributes to make them potential vectors for carrying cloned DNA. However, naturally occurring (unmodified, or non-engineered) plasmids often lack several important features that are required for a high-quality cloning vector.
The more important features are
(1) A choice of unique (single) restriction endonuclease recognition sites into which the insert
DNA can be cloned and
(2) One or more selectable genetic markers for identifying recipient cells that carry the cloning vector–insert DNA construct. In other words, plasmid cloning vectors have to be genetically engineered.
An assortment of vector systems is now accessible to meet the demands of a specific experimental procedure. Individually each has precise pros and cons. Bacteriophage vectors have served a predominantly vital role in molecular biology. Bacteriophages are viruses with a liking towards bacteria whose genome (prototype A-bacteriophage vector) is a double-stranded, linear DNA molecule having 50 kilobases (kb) length. Each end is made up of 12 bp complementary DNA that is single-stranded in nature. Once taken up by the bacteria the virus undertakes a circular form at the time when the termini are combined and sealed by the action of host ligase. Two mechanisms of replication are likely to occur. In one of the cases, the circular DNA is multiplied into a high copy number and the host cell ultimately experiences lysis releasing the infectious viral particles. Otherwise, the bacteriophage DNA may integrate with the host bacterial DNA commonly named as the lysogenic pathway. In this scenario the viral DNA remains dormant, expressing only a handful of genes. Bacteriophages have been adapted to receive and proliferate exogenous DNA. The selection of a specific vector is dependent on the fragment size that is required to be cloned, the restriction enzyme applied in the generation of the fragment and if the cloned fragment is required to express into protein in the bacterial host. In this last context, numerous expression vectors permit the cloning of foreign inserts following the location of a strong bacterial promoter required to let the proficient transcription in the bacterial host.
Plasmid Cloning Vector pBR322
In the 1980s, one of the best-studied and most often used “general-purpose” plasmid cloning vectors was pBR322. In general, plasmid cloning vectors are designated by a lowercase p, which stands for plasmid, and some abbreviation that may be descriptive or, as is the case with pBR322, anecdotal. The “BR” of pBR322 recognizes the work of the researchers F. Bolivar and R. Rodriguez, who created the plasmid, and 322 is a numerical designation that has relevance to these workers. Plasmid pBR322 contains 4,361 bp. As shown in Fig. 3.9, pBR322 carries two antibiotic resistance genes. One confers resistance to ampicillin (Ampr), and the other confers resistance to tetracycline (Tetr). This plasmid also has unique BamHI, HindIII, and SalI recognition sites within the Tetr gene; a unique PstI site in the Ampr gene; a unique EcoRI site that is not within any coding DNA.
Cloning vectors called cosmids can carry about 45 kb of cloned DNA and are maintained as plasmids in E. coli. Cosmids combine the properties of plasmids and bacteriophage vectors. For example, the commonly used cosmid pLFR-5 has two cos sites (cos ends) from bacteriophage flanking a ScaI restriction endonuclease site, a multiple cloning site with six unique recognition sites (HindIII, PstI, SalI, BamHI, SmaI, and EcoRI), an origin of DNA replication, and a Tetr gene. Pieces of DNA that are approximately 45 kb in length are purified by sucrose density gradient centrifugation from a partial BamHI digestion of the source DNA. The pLFR-5 DNA (~6 kb) is cleaved initially with ScaI and then with BamHI.
The final two DNA samples are mixed and ligated. Some of the ligated products have an ~45-kb DNA piece inserted between the two fragments that are derived from the digestions of the pLFR-5 DNA. These molecules are about 50 kb long and have cos sequences that are about 50 kb apart.
Consequently, these DNA constructs are successfully packaged into bacteriophageheads in vitro. Reconstituted pLFR-5 without inserted DNA is not packaged. After the assembly of bacteriophage particles, the DNA is delivered by infection into E. coli. Once inside the host cell, the cos ends, which were cleaved during the in vitro packaging, base pair and enable the linear DNA to circularize. This circular form is stable, and the cloned DNA is maintained as a plasmid–insert DNA construct because the vector DNA contains a complete set of plasmid functions. Moreover, the Tetr gene allows colonies that carry the cosmid to grow in the presence of tetracycline. Nontransformed cells are sensitive to tetracycline and die.
The plasmid-based vectors used for cloning DNA molecules generally carry up to 10 kb of inserted DNA. However, for the formation of a library, it is often helpful to be able to maintain larger pieces of DNA. To this end, various high-capacity cloning systems have been developed. The E. coli virus (bacteriophage, or phage) has been engineered to be a vector for inserts in the range of 15 to 20 kb.
The plasmid, pBR322, has been used extensively for cloning DNA in Escherichia coli K12 (E. coli). It is a small DNA plasmid of approximately 4.4 kilobases (Kb) that contains very little extraneous sequence that does not contribute to its utility as a cloning vector. Three functional regions define the important features of the plasmid.
The replication region (Rep) contains all of the information that is needed for replication and maintenance of the plasmid in a suitable E. coli host. Two regions encode resistance to antibiotics. Ap’ designates resistance to ampicillin which is mediated by the enzyme, p-lactamase, and Tc’ designates resistance to tetracycline.
Antibiotic resistance markers are important because they provide positive selection for transformed hosts that carry the plasmid. pBR325 is a useful cloning vector that was derived from pBR322. The marker, Cm’, designates resistance to chloramphenicol that is mediated by chloramphenicol acetyl transferase. DNA molecules are cut by restriction endonucleases at specific recognition sites. Two linear fragments of DNA can be joined together to form a circular molecule by the enzyme, DNA ligase, if the linear fragments have homologous termini. Useful cloning vectors should contain single recognition sites for restriction enzymes so that they can be converted to linear molecules without losing essential functions of the vector. Only a portion of the transformed hosts contain recombinant DNA molecules in a typical cloning experiment. The remainder of the transformed population will contain the vector that has been self-ligated without insertion of a new DNA fragment. One of the antibiotic resistance markers is needed to select transformed cells from those that do not contain plasmid; however, the second resistance marker can be used to identify cells that contain a recombinant plasmid by the phenomenon of insertional inactivation. The cloned fragment may interrupt expression of antibiotic resistance if insertion occurs in the gene that encodes the resistance. pBR322 contains a single recognition site for Pstl, which occurs in the gene for Ap’. Recombinants of pBR322 will be recognized as Ap’ (ampicillin sensitive) and Tc’ transformants if DNA is cloned into the PstI site. Similarly, recombinants of pBR322 will be recognized as Tc5 (tetracycline sensitive) and Ap’ transformants if DNA is cloned into the SalI or BumHI site. Recombinants of pBR325 will be recognized as Cm’ (chloramphenicol sensitive), Ap’, and Tc’ if DNA is cloned into the EcoRI site.
Transformation and Selection
The next step in a recombinant DNA experiment requires the uptake of the cloned plasmid DNA by a bacterial cell, usually E. coli. The process of introducing purified DNA into a bacterial cell is called transformation, and a cell that is capable of taking up DNA is said to be competent. Competence occurs naturally in many bacteria. In different bacterial species, usually when cell density is high or starvation is impending, a set of proteins is produced that facilitates the uptake of DNA molecules. This phenomenon allows genes to be transferred between different bacteria.
A natural transformation process often entails
(1) The binding of double-stranded DNA to components of the cell wall
(2) Entry of the DNA into an inner compartment (periplasm), where it is protected from enzymes that degrade nucleic acids (nucleases)
(3) Transmission of one strand into the cytoplasm while the other one is degraded; and
(4) If the DNA is a linear molecule, integration into the host chromosome. If the introduced DNA is a plasmid, it is maintained in the cytoplasm after the second strand is synthesized.
Competence and transformation are not intrinsic properties of E. coli. However, competence can be induced in E. coli by various special treatments, such as cold calcium chloride, which in turn enhance the acquisition of DNA by the cell.
A brief heat shock facilitates the uptake of exogenous DNA molecules. After the transformation step, it is necessary to identify, as easily as possible, those cells that contain plasmids with cloned DNA. In a pBR322 system in which the target DNA was inserted into the BamHI site, this specific identification is accomplished using the two antibiotic resistance markers that are carried on the plasmid. Following transformation, the cells are incubated in medium without antibiotics to allow the antibiotic resistance genes to be expressed, and then the transformation mixture is plated onto medium that contains the antibiotic ampicillin. Cells that carry pBR322 with or without insert DNA can grow under these conditions because the Ampr gene on pBR322 is intact. The nontransformed cells are sensitive to ampicillin.
Creating and Screening a Library
Making a Genomic Library
One of the fundamental objectives of molecular biotechnology is the isolation of genes that encode proteins for industrial, agricultural, and medical applications. In prokaryotic organisms, structural genes form a continuous coding domain in the genomic DNA, whereas in eukaryotes, the coding regions (exons) of structural genes are separated by noncoding regions (introns). Consequently, different cloning strategies have to be used for cloning prokaryotic and eukaryotic genes.
In a prokaryote, the desired sequence (target DNA, or gene of interest) is typically a minuscule portion (about 0.02%) of the total chromosomal DNA. The problem, then, is how to clone and select the targeted DNA sequence. To do this, the complete DNA of an organism, i.e., the genome, is cut with a restriction endonuclease, and each fragment is inserted into a vector. Then, the specific clone that carries the target DNA sequence must be identified, isolated, and characterized. The process of subdividing genomic DNA into clonable elements and inserting them into host cells is called creating a library (clone bank, gene bank, or genomic library). A complete library, by definition, contains all of the genomic DNA of the source organism. One way to create a genomic library is to first treat the DNA from a source organism with a four-cutter restriction endonuclease, e.g., Sau3AI, which theoretically cleaves the DNA approximately once in every 256 bp. The conditions of the digestion reaction are set to give a partial, not a complete, digestion. In this way, all possible fragment sizes are generated.
Autoradiography is used to detect the location of a radiolabeled entity in a cell or sample of fractionated macromolecules. In principle, autoradiography consists of placing a radioactive source next to a radiosensitive photographic film that contains silver bromide. The energy from the decay of the radioisotope hits the photographic emulsion and produces electrons that are trapped by specks of silver bromide crystals in the emulsion. The negatively charged specks attract silver ions, and metallic silver is formed. The grains of metallic silver are visualized by developing the photographic film. Thus, an exposed dark region on a developed film indicates that the underlying material was radiolabeled.
Parenthetically, fluorography is the term used for the exposure of light-sensitive photographic film to molecules that directly or indirectly generate light as the source of energy that reduces silver in the photographic emulsion. Proteins and nucleic acids that are radiolabeled and separated by gel electrophoresis can be visualized by placing an X-ray film on a dried gel and developing the film after a suitable exposure time. All autoradiographic steps are carried out in the dark to avoid inadvertent exposure of the X-ray film to light. A number of autoradiographic techniques have been devised for the quantitative and qualitative analysis of proteins and nucleic acids.
One of the major applications of autoradiography is the detection of the hybridization of a radiolabeled DNA probe to a DNA molecule that has been electrophoretically fractionated. However, DNA molecules in a gel are not accessible to hybridization with a DNA probe. Consequently, the DNA molecules in the gel are transferred by blotting or electrotransfer to a nitrocellulose or nylon membrane.
The transfer process retains the same positions on the membrane as the DNA molecules had in the gel. The DNA molecules that are transferred to a membrane are denatured, bound to the membrane, and hybridized with a radiolabeled DNA probe. Autoradiography of the membrane reveals whether the probe hybridized to a particular DNA band(s).
The transfer of DNA from a gel to a membrane is called Southern blotting (Southern DNA blotting) after Edwin Southern, who devised the original DNA blotting strategy. Northern blotting and Western blotting are methods for the transfer of RNA and protein, respectively, from a gel to a membrane. The terms “Northern” and “Western” have nothing to do with direction and were coined by molecular biologists both to give Edwin Southern further credit for developing the notion of blotting macromolecules from a gel to a membrane and to distinguish the macromolecules that are transferred. The designations “Northern” and “Western” are also examples of molecular biology humor There are at least two possible sources of probes for screening a genomic library. First, cloned DNA from a closely related organism (a heterologous probe) can be used. In this case, the conditions of the hybridization reaction can be adjusted to permit considerable mismatch between the probe and the target DNA to compensate for the natural differences between the two sequences. Second, a probe can be produced by chemical synthesis. The nucleotide sequence of a synthetic probe is based on the probable nucleotide sequence that is deduced from the known amino acid sequence of the protein encoded by the target gene. Genomic libraries are often screened by plating out the transformed cells on the growth medium of a master plate and then transferring samples of each colony to a solid matrix, such as a nitrocellulose or nylon membrane. The cells on the membrane are broken open (lysed), the protein is removed, and the DNA is bound to the membrane. At this stage, a labeled probe is added, and if hybridization occurs, signals are observed on an autoradiograph. The colonies from the master plate that correspond to samples containing hybridized DNA are then isolated and cultured. Because most libraries are created from partial digestions of genomic DNA, a number of colonies may give a positive response to the probe. The next task is to determine which clone, if any, contains the complete sequence of the target gene. Preliminary analyses that use the results of gel electrophoresis and restriction endonuclease mapping reveal the length of each insert and identify those inserts that are the same and those that share overlapping sequences. If an insert in any one of the clones is large enough to include the full gene, then the complete gene can be recognized after DNA sequencing because it will have start and stop codons and a contiguous set of nucleotides that code for the target protein. Alternatively, a gene can be assembled by using overlapping sequences from different clones. Unfortunately, there is no guarantee that the complete sequence of a target gene will be present in a particular library. If the search for an intact gene fails, then another library can be created with a different restriction endonuclease and screened with either the original probe or probes derived from the first library. On the other hand, as discussed below, libraries that contain DNA fragments larger than the average prokaryotic gene can be created with specialized vectors to increase the chance that some members of the library will carry a complete version of the target gene.
Screening by Immunological Assay
Alternative methods are used to screen a library when a DNA probe is not available. For example, if a cloned DNA sequence is transcribed and translated, the presence of the protein, or even part of it, can be determined by an immunological assay. Technically, this procedure has much in common with a DNA hybridization assay. All the clones of the library are grown on several master plates. A sample of each colony is transferred to a known position on a matrix, where the cells are lysed and the released proteins are attached to the matrix. The matrix with the bound proteins is treated with an antibody (primary antibody) that specifically binds to the protein encoded by the target gene. Following the interaction of the primary antibody with the target protein (antigen), any unbound antibody is washed away, and the matrix is treated with a second antibody (secondary antibody) that is specific for the primary antibody. In many assay systems, the secondary antibody has an enzyme, such as alkaline phosphatase, attached to it. After the matrix is washed, a colorless substrate is added. If the secondary antibody has bound to the primary antibody, the colorless substrate is hydrolyzed by the attached enzyme and produces a colored compound that accumulates at the site of the reaction.
Screening by Protein Activity
DNA hybridization and immunological assays work well for many kinds of genes and gene products. If the target gene produces an enzyme that is not normally made by the host cell, a direct (in situ) plate assay can be devised to identify members of a library that carry the particular gene encoding that enzyme. The genes for amylase, endoglucanase, glucosidase, and many other enzymes from various organisms have been isolated in this way. This approach has proven effective for isolating genes encoding biotechnologically useful enzymes from microorganisms present in environmental samples. Most of the organisms contained in these samples be grown in the laboratory, outside of their natural environment.
However, the total genomic DNA from these organisms can be extracted directly from the sample, for example, a soil sample, and used to prepare a metagenomic library that can be expressed in a host bacterium, such as E. coli, and screened for target protein activity. This technique has enabled the isolation of many novel proteins with interesting properties without the need to first culture the natural host microorganism.
The genes isolation can be implemented via using two fundamental approaches. The gene can be isolated as it occurs in the genome, harbouring the multifaceted intron-exon structure as well as the promoter elements. However, higher eukaryotic genes comprise of many introns and are generally too huge to be proliferated as a single entity. Frequently the gene is cloned with much difficulty as overlapping fragments. Nevertheless, using the enzyme reverse transcriptase it is likely to develop a double-stranded copy of the transcript of mRNA, as it lacks introns containing only the sequence responsible for protein-coding with portions that are untranslated restricted to the 5’ and 3’ ends. Total genomic DNA digested signifies the cells’ total mRNA that is possible to be inserted into a range of vector systems generating a library whose constituents comprises a host of a variety of genes. When one desires to isolate a specific gene of interest from these libraries, it is obligatory to screen the library to recognize the species of interest. In one of the example, cDNA is cloned in a bacteriophage vector that is further bundled into infectious viral particles. Each virus comprises of a dissimilar phage with an exclusive gene insert. These viruses are combined with host bacteria and grown on nutrient agar media. Bacteria that have been infested with a single phage ultimately lead to lysis or diminished growth of the host which looks like a hole on the grown bacteria colonies. A sample of each phage can be multiplied onto a section of nitrocellulose filter paper. The filter paper takes up the viral nucleic acids as well as proteins produced by the virus. The filters are then subjected to screening using a radioactive probe. The probe might be a part of the gene itself, purified beforehand from a different source having similar nature. Once the filter is incubated with the radioactive probe since the sequence on the capture and the target stands are complementary the probe will attach the to samples having alike sequences. After showing the filter to x-ray film, selected samples having the gene of interest can be recognized and selected from the initial master plate.
The DNA is inserted into a eukaryotic cell by an assortment of procedures, comprising of injection, transformation, viral infection, or ballistic gene gun tactic. When externally introduced DNA that is initially obtained from that organism is inserted into the genome, there is a possibility either the resident gene will be replaced or insertion will be ectopically. If it is a transgenic DNA obtained from another species, it generally adds ectopically. Vectors with the capacity to replicate independently in eukaryotic systems are occasionally found, so in the majority of the cases, chromosomal addition route is preferably adopted.
The likelihood of transgenic alteration of eukaryotes like plants and animals unlocks many novel tactics for investigation because genotypes can be genetically modified to render them appropriate for the certain precise experiment. An instance in fundamental research is in the application of reporter genes. Occasionally in the tissue environment, it is tough to notice the action of a particular gene where it generally functions. This difficulty can be avoided by splicing the gene promoter accountable for the coding portion of a gene called reporter gene, the product of which can be effortlessly diagnosed. In the case when the gene is functioning the reporter gene will affirm that action in the applicable tissue. Moreover, because fungi, animals and plants make the foundation for a huge part of the economy, transgenic or genetically modified genotypes are widely applied in practical research. A predominantly stimulating application of transgenesis is in gene therapy in humans via the introduction of a routinely functional transgene that can substitute or recompense for an existing malfunctioning gene.
Genetic Engineering in Baker’s Yeast
The Saccharomyces cerevisiae yeast has developed to be the most refined eukaryotic model for recombinant DNA technology. One of the chief reasons is that the transmission genetics of yeast is particularly well defined and the accumulation of thousands of mutants influencing hundreds of dissimilar phenotypes is a valued reserve when applying yeast as a molecular system. In case of yeast, another significant benefit is the accessibility of natural yeast plasmid of circular 6.3-kb. It makes the foundation for numerous well equipped cloning vectors. This plasmid is transferred to the cellular meiosis and mitosis products. The most naive yeast vectors are by-products of bacterial plasmids into which the locus of interest of yeast has been introduced. When transmuted into yeast cells, these plasmids merge into yeast chromosomes usually by homologous recombination with the existing gene via a double or a single crossover. As a consequence of this, the whole plasmid is introduced or else the targeted allele is substituted by the allele present in the plasmid. Such additions can be distinguished by plating cells on a selective medium that picks for the allele present on the plasmid. Since bacterial plasmids do not duplicate in yeast, incorporation is the lone technique to produce a stable altered genotype by application of these vectors.
If the plasmid of 2-μm is applied as the fundamental vector and other bacterial, as well as yeast sections, are merged into it thereby producing a construct harbouring numerous valuable properties. Initially, the 2-μm part in the yeast cell delivers the potential to duplicate independently, and addition is not essential for a stable transformation. Thereafter, the genes can be inserted into yeast and their activity can be investigated in that organism following which the plasmid can be inserted back into E. coli after recovery, as long as a bacterial origin of replication and a selectable bacterial marker are existing on the plasmid. Such vectors known as shuttle vectors are very beneficial in the repetitive cloning and manipulation of yeast genes.
With some plasmid autonomously replicating there is a likelihood that an offspring cell will not receive a copy as the segregation of plasmid copies to offsprings is fundamentally a casual process reliant on where the plasmids are located in the cell when the cell wall of the daughter cell is formed. Nevertheless, if the segment of yeast DNA harbouring a centromere is inserted into the plasmid, then the proper separation of chromosomes is ensured and will consider the plasmid to be a part of the cell and divide it into daughter cells during cell division. The insertion of a centromere is a single step in the direction of the conception of an artificial chromosome. Further endeavours have been adopted by linearizing a plasmid comprising of a centromere and thereafter addition of the DNA from yeast telomeres to the ends. In case this construct encloses yeast origins of replication and thereafter a yeast artificial chromosome (YAC) is established, which performs in many functions like a small yeast chromosome during cell division.
Plasmids that are centromeric can be applied to understand the regulatory elements present upstream of a gene. The applicable coding area and its upstream area can be merged into a plasmid, which can be chosen by a distinct yeast marker. The upstream portion can be influenced by making a sequence of deletions, which are attained by chopping the DNA employing a special exonuclease to remove the DNA in a particular direction to dissimilar extents followed by joining it again. The investigational aim is then to decide which of these removals still allows for the usual functioning of the gene. By transformation of the plasmid, the appropriate function is evaluated into a receiver where in the locus of the chromosome harbours a mutant allele that is defective in nature and then checking is done for the reversal of the gene function in the recipient.
The consequences normally describe a precise region that is essential for usual function and gene regulation. In such studies, it is frequently more suitable to apply a reporter gene as an alternative of the gene of interest. lacZ gene of bacteria has been comprehensively applied as a reporter in yeast as it encodes the β-galactosidase enzyme which is accountable for breakdown of lactose and is well known for its activity towards an analog of lactose known as X-Gal producing 5-bromo-4-chloroindigo that is blue in colour and it is conveyed as a blue colony of yeast when it is activated. Yeast artificial chromosomes have been widely applied as cloning vectors for huge segments of eukaryotic DNA. Moreover, the huge size of mammalian genomes over-all means that the libraries prepared in bacterial vectors would be also respectively huge however Yeast artificial chromosomes in contrast harbour much longer inserts approximately 1000 kb even though the library is smaller.
Genetic Engineering in Plants
Genetic analysis of plants has been extensively explored to enroot enriched varieties because of their commercial as well as ecological importance. New pathways to this effort have been outstretched by the limitless genome modifications made possible by recombinant DNA technology. However.Breeding has no longer been restrained to selecting breeds within the specified species but DNA from alternative species of animals, plants or even bacteria can be introduced.
The transgenic plants are typically produced only from the vectors derived /imitated from the soil bacterium called Agrobacterium tumefaciens. However, this bacterium infects the plant causing uncontrolled growth (tumours, or galls), commonly at the base(crown) of the plant which is popularly known as crown gall disease. The circular DNA plasmid- the Ti (tumour-inducing) plasmid which is about 200kb in size is majorly responsible for tumour production. The Ti plasmid containing T-DNA portion is transported and combined into the genome of the host plant in an arbitrary fashion when the plant cell is infested by the bacterium. And this transfer is enabled through a region outside the T-DNA on the Ti plasmid. Amongst numerous interesting functions, the T-DNA produces a tumour and and is also accountable for leading the synthesis of opines in the host plant. Nopaline and octopine are the significant opines formed by two dissimilar Ti- plasmids. The opines are taken up by the bacterium using the opine- dependent genes on the Ti plasmids for its individual purposes.
The character of plant vectors supports the natural behaviour of the Ti plasmid. By implementing the essential amendments, if the DNA of interest can be merged into T DNA in that scenario the entire system could be introduced for establishing a consistent state into a plant chromosome. Additionally, alongwith their larger size, the Ti plasmids are not easy to be manipulated neither their size can be reduced owing to their few exclusive restriction sites. Originally, the insert of interest and the several other genes as well as segments essential for recombination, replication, and antibiotic resistance are accepted by the smaller transitional vector. Ti plasmid can then be introduced with the transitional vector after being modified with the element of a gene of interest. This assembled plasmid can then be put into a plant cell by means of transformation.
The Ti plasmid containing the entire right hand portion of its T-DNA, including the tumour genes as well as the genes accountable for nopaline synthesis is initially mitigated after the transitional vector is being accepted making it incompetent to develop a tumour considered as an undesirable feature of the T-DNA function. The left-hand portion of T-DNA is retained which is applied in the inoculation of the transitional vector as a crossover site. The transitional vector contains a suitable cloning section having the diversity of exclusive restriction sites joined in. Also for acquiring the spectinomycin resistance a selectable marker bacterial gene (spcR), for plant expression an adapted bacterial kanamycin-resistance gene (kanR) and two sections of T-DNA are merged into the vector. Gene for Nopaline synthesis (nos) and the right-hand T-DNA border sequence are supported by one portion. Recombinant plasmids can be then carefully chosen by growing in presence of spectinomycin, followed by the transitional vectors implantation into Agrobacterium cells possesing the disarmed Ti plasmids (by conjugation with E. coli). The carefully chosen bacterial colonies will harbour the Ti plasmid only because the transitional vector is devoid of the capacity to replicate in Agrobacterium. Thereafter, once the spectinomycin selection is done, the bacteria comprising of the recombinant double or cointegrant plasmid are then applied to infest expurgated parts of plant tissue, like perforated leaf disks.
If bacterial contamination of plant cells occurs, any genetic cargo between the left and right T-DNA border sequences can be introduced into the chromosomes of the plant. In the case where the leaf disks are located on a kanamycin containing medium, the plant cells that specifically experience cell division are the ones that have attained the kanamycin resistance gene from the transfer of T-DNA. The development of such cells consequences in the formation of a clump/callus, which signals successful transformation. These calli can be made to generate shoots and roots when they are transported to soil and finally mature into transgenic plants. Frequently only one T-DNA insertion is evident in such plants, where it separates at meiosis in a conventional pattern. The addition can be distinguished by a probe specific for T-DNA in Southern blot or can be established by the nopaline estimation in the transgenic tissue.
Expression of cloned DNA
The cloned DNA incorporated into the T-DNA can be to some extent the DNA that the researcher needs to add into the plant of interest. Mostly, foreign DNA that has been incorporated with the help of T-DNA is the gene coding for the luciferase enzyme extracted from fireflies. The reaction of a chemical called luciferin with ATP is catalysed by the enzyme and in this procedure, light emission is observed which clarifies the glow of fireflies in the dark. Likewise in case of a transgenic technology mediated established tobacco plant when drenched with luciferin will lit in the dark. This kind of influence appears like an effort to advance a technology for production of Christmas trees without decorating with artificial lights. Nevertheless, the luciferase gene is also beneficial to monitor any gene function during development by acting as a reporter i.e. the promoter sequences upstream of any gene that is of interest can be merged with the luciferase gene and introduced by T-DNA into a plant. Thereafter, the luciferase gene will undergo the same evolving pattern as the usually controlled gene does but will proclaim its action importantly by shining at several times or in different tissues subjected to the regulatory sequence.
Other genes applied in plants as reporters are the bacterial lac (β-galactosidase) gene, which acts on X-Gal to convert it into blue colour and bacterial GUS (β-glucuronidase) gene, which changes the compound X-Gluc to blue. These reporter expressing cells which turn blue in colour can be effortlessly observed either under the microscope or by the naked eye.
Transgenic plants harbouring an assortment of exogenous genes are existing in use, and several more are in the process of development. Like the properties of plants themselves are being influenced by the microorganisms as well are also applied as a suitable source to harvest proteins programmed by foreign genes.
Materials and Equipment
Commercial suppliers for plasmid transformation systems may be purchased in kits. These plasmids should contain the gene for ampicillin resistance (pBLU), as experimental procedures typically use ampicillin to select transformed cells. In addition, plasmids with colored marker genes like beta-GAL and fluorescence markers like green fluorescent protein (GFP) and its cousins make it possible to measure gene expression directly, to follow cell populations as they grow or move, and to find cells that have taken up a second plasmid that we cannot see easily.
The following materials are included in a typical eight-station ampicillin-resistant plasmid system.
- coli (1 vial or slant)
- Plasmid (pBLU), hydrated (20 μg)
- Ampicillin, lyophilized (30 μg)
- Transformation solution (50mM CaCl2, pH 6), sterile (15 mL)
- LB nutrient agar powder, sterile (to make 500 mL) (20 g) or prepared agar
- LB nutrient broth, sterile (10 mL)
- Pipettes, sterile (50)
- Inoculation loops, sterile (10 μL, packs of 10 loops)
- Petri dishes, sterile, 60 mm (packs of 20)
- Multicolor 2.0 mL microcentrifuge tubes (60)
- Microcentrifuge tube holders
- Clock or watch to time 50 seconds
- Microwave oven/water bath
- Thermometer that reads 42°C
- 1 L flask
- 500 mL graduated cylinder
- Distilled water
- Crushed ice and containers
- 10% solution household bleach
- Permanent marker pens
- Masking tape
- Biohazardous waste disposal bags or plastic trash bags
- Micropipettes, adjustable volume, 2–20 μL (and pipette tips)
- Parafilm laboratory sealing film
- 37°C incubator oven*
Advance Preparation Quick Guide for Teachers
Step Objective Time Required When
Step 1 Prepare agar plates. 1 hr. 3–7 days prior
Step 2 Rehydrate E. coli. 2 min 24–36 hours prior
Streak starter plates. 2 min 24–36 hours prior
Rehydrate plasmid DNA 15 min. 24–36 hours prior.
Step 3 Aliquot solutions. 10 min. Immediately prior
Advance Preparation for Step 1: 3–7 Days before the Transformation
- Prepare nutrient agar (autoclave-free).
The agar plates should be prepared at least three days before the investigation(s) are performed. Plates should be left out at room temperature for two days and then refrigerated until use. (Two days at room temperature allows the agar to cure, or dry, sufficiently to readily take up the liquid transformation solution.) If time is short, incubate the plates at 37°C overnight. This will dry them out as well, but it shortens their shelf life. Refrigerated plates are good for up to 30 days.
To prepare the agar, add 500 mL of distilled water to a one liter or larger Erlenmeyer flask. Add the entire content of the LB nutrient agar packet. Swirl the flask to dissolve the agar and heat to boiling in a microwave or water bath or by using a hot plate with stir bar. Heat and swirl until all the agar is dissolved.
When all the agar is dissolved, allow the LB nutrient agar to cool so that the outside of the flask is just comfortable to hold (approximately 50°C.). While the agar is cooling, you can label the plates and prepare the ampicillin as outlined below in Step 3.
Pre-prepared nutrient agar also can be purchased. However, it will have to be melted before it can be poured into plates. To do this, the plastic bottles containing solid agar can be microwaved at a low temperature (such as using the “poultry defrost” option) for several minutes. Be sure to loosen the cap slightly to expel any air. At high microwave temperatures, the agar can boil over. Another option is to place the bottles in a hot water bath; however, this will take up to 45 minutes or so to melt the agar.
- Prepare ampicillin.
Ampicillin is either shipped dry in a small vial or already hydrated. If shipped dry, you need to hydrate the ampicillin. Do this by adding 3 mL of transformation solution to the vial to rehydrate the antibiotic. Use a sterile pipette. The nutrient agar solidifies at 27°C, so you must be careful to monitor the cooling of the agar and then pour the plates from start to finish without interruption. Keeping the flask with liquid agar in a water bath set to 45–50°C can help prevent the agar from cooling too quickly. Before adding ampicillin to the flask of agar, make sure you can hold the flask in your bare hand (approximately 50°C). If your hand tolerates the temperature of the flask, so will the antibiotic!
- Label plates.
While the agar is cooling, reduce preparation time by labeling the plates. Label with a permanent marker on the bottom of each plate close to the edge. For each class using an eight-station kit, label 16 plates LB and 16 plates LB/amp.
- Pour nutrient agar plates. First, pour LB nutrient agar into the 16 plates that are labeled LB. If you do not do this and add ampicillin to the flask with agar, you will not be able to make control plates containing just nutrient agar. Fill each plate to about one-third to one-half (approximately 12 mL) with agar and replace the lid. You may want to stack the plates and let them cool in the stacked configuration. Add the hydrated ampicillin to the remaining LB nutrient agar. Swirl briefly to mix. Pour into the 16 plates labeled LB/amp using the same technique. Plates should set within 30 minutes.
- Store the plates.
After the plates have cured for two days at room temperature, they may be either used or stored by stacking them in a plastic sleeve bag slipped back down over them. The stack is then inverted, the bag taped closed, and the plates stored upside down at 4°C until used. (The plates are inverted to prevent condensation on the lid, which may drip onto the agar.)
Advance Preparation for Step 2: 24–36 Hours before the Transformation
- Rehydrate bacteria.
Some E. coli cultures come prepared in a slant and will not have to be rehydrated. For bacteria that must be rehydrated, use a sterile pipette to add 250 μL of transformation solution directly to the vial. Recap the vial and allow the cell suspension to stand at room temperature for 5 minutes. Then shake the mix before streaking on the LB starter plates. Store the rehydrated bacteria in the refrigerator until used (within 24 hours for best results and no longer than three days).
- Streak starter plates.
Starter plates are needed to produce bacterial colonies of E. coli on agar plates. Each lab team will need its own starter plate as a source of cells for transformation. LB plates should be streaked for single colonies and incubated at 37°C for 24–26 hours before the transformation investigation begins.
Using E. coli and LB agar plates, streak one starter plate to generate single colonies from a concentrated suspension of bacteria. A small amount of the bacterial suspension goes a long way. Under favorable conditions, one cell multiples to become millions of genetically identical cells in just 24 hours. There are millions of individual bacteria in a single millimeter of a bacterial colony.
- Insert a sterile inoculation loop straight into the vial of rehydrated bacterial culture. Remove the loop and streak the plates. Streaking takes place sequentially in four quadrants. The first streak spreads out the cells. Go back and forth with the loop about a dozen times in each of the small areas shown. In subsequent quadrants, the cells become more and more dilute, thus increasing the likelihood of producing single colonies.
- For subsequent streaks, use as much of the surface area of the plate as possible. After the initial streak, rotate the plate approximately 45 degrees and start the second streak. Do not dip into the rehydrated bacteria a second time. Go into the previous streak about two times and then back and forth for a total of about 10 times.
- Rotate the plate again and repeat streaking.
- Rotate the plate for the final time and make the final streak. Repeat steps a–c with the remaining LB plates for each student workstation. Although you can use the same inoculation loop for all starter plates, it is recommended that you use a new, sterile loop for each plate if you have enough. When you are finished with each plate, cover it immediately to avoid contamination.
- Place the plates upside down inside the incubator overnight at 37°C or at room temperature for 2–3 days if an incubator is unavailable. Use for transformation within 24–36 hours because bacteria must be actively growing to achieve high transformation efficiency. (Remember, bacterial growth is exponential.) Do not refrigerate before use. This will slow bacterial growth.
- coli forms off-white colonies that are uniformly circular with smooth edges. Avoid using plates with contaminant colonies such as mold.
- Prepare plasmid.
The quantity of DNA is so small that the vial may appear empty. Tap the vial or spin it in a microcentrifuge to ensure that the DNA is not sticking to the cap. If the plasmid is not hydrated, refer to instructions that come with the sample. Store the vial of hydrated DNA in a refrigerator. Rehydrated plasmid should be used within 24 hours.
Advance Preparation for Step 3: Immediately Before Transformation Investigation
- Aliquot solutions.
- Each student workstation will need 1 mL of transformation solution and 1 mL of LB nutrient broth. You might have to aliquot these solutions into separate color-coded 2 mL microtubes. If the LB nutrient broth is aliquoted one day prior to the lab, it should be refrigerated. Make sure to label the tubes with permanent marker.
- Set up the workstations. See the list of materials required. If the plasmid goes through multiple freeze-thaw cycles in a frost-free freezer, the DNA in the plasmid can degrade. It is recommended that you check the shelf life of materials with the commercial vendor.
The plasmid likely will contain the gene for resistance to ampicillin (pBLU) antibiotic that is lethal to many bacteria, including E. coli cells. This transformation procedure involves the following three main steps to introduce the plasmid DNA into the E. coli cells and to provide an environment for the cells to express their newly acquired genes:
- Adding CaCl2
- “Heat shocking” the cells
- Incubating the cells in nutrient broth for a short time before plating them on agar
- coli starter plate prepared.
- Poured agar plates prepared.
- 2 LB agar plates
- 2 LB/amp agar (LB agar containing ampicillin) plates
- Transformation solution (CaCl2, pH 6.1) kept ice cold
- LB nutrient broth
- Sterile inoculation loops
- 100–1000 μL sterile bulb pipettes
- 1–10 μL micropipettes with sterile tips
- Microcentrifuge tubes
- Microcentrifuge tube holder/float
- Container full of crushed ice
- Marking pen
- DNA plasmid (0.005 μg/μL)
- 42°C water bath and thermometer
- 37°C incubator
- 20 μL adjustable-volume micropipettes and tips (optional)
- 10% household bleach
- Biohazardous waste disposal bags
- Masking or lab tape
Label one closed microcentrifuge tube (micro test tube) “+ plasmid” and one tube “-plasmid.” Label both tubes, and place them in the microcentrifuge tube holder/float.
Carefully open the tubes and, using a 100–1000 μL bulb pipette with a sterile tip, transfer 250 μL of the ice cold transformation solution (CaCl2) into each tube.
Place both tubes on (into) the ice.
Use a sterile inoculation loop to pick up a single colony of bacteria from your starter plate. Be careful not to scrape off any agar from the plate. Pick up the “+ plasmid” tube and immerse the loop into the CaCl2 solution (transforming solution) at the bottom of the tube. Spin the loop between your index finger and thumb until the entire colony is dispersed in the solution.
Use a new sterile 100–1,000 μL micropipette to repeatedly pulse the cells in solution to thoroughly resuspend the cells. Place the tube back on the ice.
Using a new sterile inoculation loop, repeat Steps 5 and 6 for the “- plasmid” tube.
Using a 1–10 μL micropipette with a sterile tip, transfer 10 μL of the plasmid solution directly into the E. coli suspension in the “+ plasmid” tube. Tap tube with a finger to mix, but avoid making bubbles in the suspension or splashing the suspension up the sides of the tube. Do not add the plasmid solution into the “- plasmid” tube.
Incubate both tubes (“+ plasmid” and “- plasmid”) on ice for 10 minutes. Make sure the bottom of the tubes make contact with the ice.
While the tubes are sitting on ice, label each of your agar plates on the bottom.
Following the 10-minute incubation at 0°C, remove the tubes from the ice and “heat shock” the cells in the tubes. It is critical that the cells receive a sharp and distinct shock.Make sure the tubes are closed tightly! Place the tubes into a test tube holder/ float, and dunk the tubes into the water bath, set at 42°C, for exactly 50 seconds. Make sure to push the tubes all the way down in the holder so that the bottom of the tubes with the suspension makes contact with the warm water.
When the 50 seconds have passed, place both tubes back on ice. For best transformation results, the change from 0°C to 42°C and then back to 0°C must be rapid. Incubate the tubes on ice for an additional two minutes.
Remove the holder containing the tubes from the ice and place on the lab counter. Using a 100–1,000 μL micropipette with sterile tip, transfer 250 μL of LB nutrient broth to the “+ plasmid” tube. Close the tube and gently tap with your finger to mix. Repeat with a new sterile micropipette for the “- plasmid” tube.
Incubate each tube for 10 minutes at room temperature.
Use a 10–1,000 μL micropipette with sterile tip to transfer 100 μL of the transformation (“+ plasmid”) and control (“- plasmid”) suspensions onto the appropriate LB and LB/Amp plates. Be sure to use a separate pipette for each of the four transfers.
Using a new sterile inoculation loop for each plate, spread the suspensions evenly around the surface of the agar by quickly “skating” the flat surface of the sterile loop back and forth across the plate surface. Do not poke or make gashes in the agar. Allow the plates to set for 10 minutes.
Stack your plates and tape them together. Place the stack upside down in the 37°C incubator for 24 hours
By calculating transformation efficiency, you can measure the success of your transformation quantitatively.
Genetic Engineering in Animals
The animals that are utmost widely applied as model systems for genetic manipulation are Drosophila, Caenorhabditis elegans, and mice. Types of most of the practices adopted for gene manipulation so far can also be functional in these animal models.
Transgenic animals can be created in a number of ways. Transgenic Drosophila can be created by the incorporation of P elements comprising plasmid vectors into the egg of the fly. Transgenic Drosophila delivers one more sketch of the application of the bacterial gene lacZ as a reporter in the genetic regulation study all through the developmental stages. The gene lacZ is merged with the promoter region of the heat shock gene of Drosophila, which generally is stimulated by elevated temperatures. To generate transgenic flies this construct is then applied. Following heat shock, the flies are executed and submerged in X-Gal. The subsequent design of tissues blue in colour is created and this delivers evidence on the heat shock gene’s chief location of the action. Yet another method for generation of transgenic mammals is by inserting distinct plasmid vectors into a fertilized egg.
In each of these cases, since the egg is originally prepared to be transgenic the foreign DNA can discover its way into germ-line cells and then distributed to the offspring resulting from these cells. Like in the case of plants, animals are subjected to manipulation to expand the potentials of the animal itself as well as to behave as suitable manufacturers of foreign proteins. For instance, mammalian milk is effortlessly acquired and it becomes an appropriate medium for collection of proteins which are more problematic to get hold of otherwise, without killing the animal.
Gene disruptions and replacement in mice
Mice are one of the most important and widely applicable models for the study of mammals. Moreover, most of the wide-ranging tools established in mice can be functional in humans also. Two important procedures are adopted where in either the capacity to interrupt a gene is applied or substitution of one allele with another is performed. Disruptions of genes are occasionally termed as knockouts. An organism harbouring the knockout gene can then be inspected for transformed phenotypes. Knockout mice are priceless models for the research on mutants comparable to those prevalent in humans. For example, knockout mice devoid of key DNA repair enzymes have been produced to examine the effect of these enzymes in controlling rates of cancer.
In the process of the development of a knockout mouse initially, an interrupted, cloned gene is applied to giving rise to embryonic stem (ES) cells harbouring a gene knockout. Even though recombination of the faulty portion of the gene into nonhomologous sites is considerably more common compared to its recombination into homozygous sites, the choice of site-specific recombinants and against ectopic recombinants can be applied. Additionally, the ES cells having a single copy of the interrupted gene of interest are inserted into a primary embryo. The subsequent offsprings are chimeric containing the tissue obtained from the recipient or the transplanted ES cell lines. Chimeric mice produced are then coupled to yield homozygous mice with the copy of the knockout gene in each of them.
Recombinant DNA Technology Applications
Health and Diseases
Recombinant DNA technology has an extensive range of applications in combating diseases and refining health conditions. Some of the areas of application are briefed below
Recombinant DNA technology is considered as an important tool of gene therapy and is the basis of inhibition and cure of many genetic disorders altogether. The development of DNA vaccines is a novel approach to deliver immunity against numerous diseases. In this procedure, the DNA provided has the genes that encode for pathogen specific proteins. Human gene therapy in clinical trials is typically intended for the treatment of cancer. Research has been directed mostly towards the high transfection effectiveness connected to gene delivery system development. Transfection in cancer gene therapy such as for treatment of brain cancer, breast cancer, lung cancer, and prostate cancer with negligible toxicity is still in infancy. Also for transplantation purposes, specially in case of renal problems as well as some other diseases are under investigation for gene therapy applications.
Production of Antibodies
The plant has been lately applied for the expression and expansion of several antibodies as well as their derivatives. Transgenic tobacco plants are utilised for the generation of secretory IgA/G chimeric in nature. Another monoclonal antibody called T84 can proficiently identify carcinoembryonic antigen and a full-length humanized IgG1 for herpes simplex virus (HSV)-2-glycoprotein B, has been produced in Chinese Hamster Ovary (CHO) cells and transgenic soybean. The manufacturing of monoclonal antibody and derivatives was possible in plants via transgenesis and agroinfiltration technologies in tobacco that is transiently transformed.
Examination of the Drug Metabolism
The multifaceted system of drug digesting enzymes indulged in the drug metabolism is vital to be examined for the appropriate effectiveness and properties of drugs. Recombinant DNA tactics have of late contributed in this regard via heterologous expression in which the genetic information of the enzyme is conveyed in vitro or in vivo by the means of the gene transfer.
Improvement of Vaccines and Recombinant Hormones
Reasonably traditional vaccines have inferior effectiveness and specificity than their recombinant counterparts. A user friendly and trouble-free procedure for transmission of adenovirus vectors encrypting pathogen antigens is via nasal transfer. This technique is quick and a defence supporting means against mucosal pathogens. This performs as a drug vaccine where a state of anti-influenza can be brought about through an airway transgene expression.
The human follicle-stimulating hormone’s in vitro manufacturing of is possible due to recombinant DNA technology as it is a significantly complex heterodimeric protein and also dedicated cell line from eukaryotes has been nominated for its expression. Supported reproduction management via stimulation of follicular expansion is an accomplishment of recombinant DNA technology. Most of the patients are subjected to treatment through r-FSH and remarkably r-FSH and recombination of Luteinizing Hormone was completed fruitfully to augment the process of formation/release of eggs as well as pregnancy.
A significant constituent of alternative medicine is the traditional Chinese medicines which have a vital role in diagnostics as well as therapeutics. These medicines are linked with theories which are consistent with gene therapy principles to a certain extent. These drugs offer to be the sources of a system bearing therapeutic genes are coadministrated as drugs. The transgenic root system has valued prospective for the introduction of supplementary genes along with the Ri plasmid and is mostly found with altered genes in A. rhizogenes vector systems to augment properties for precise use. The principles or the culture became a respected tool to examine the biochemical characteristics and the gene expression summary of metabolic pathways. The by-products, as well as the key enzymes produced during the secondary metabolite’s biosynthesis, can be explained by the turned cultures.
Agriculture and Food
Recombinant DNA technology has augmented towards the development of important enzymes for application in food and agriculture industries. Numerous significant enzymes such as lipases and amylases are utilized in large scale productions owing to their specific roles and important applications in food industries. Microbial strains mediated production is one more enormous accomplishment that turned out to be likely with the assistance of recombinant DNA technology. Many of the microbial strains have been advanced for enzyme production through precise engineering in terms of protease production. Fungi have also been modified so that their capacity of production of toxic products could be limited. Lysozymes are known as effective mediators to reduce bacterial contamination in food industries and are deemed appropriate for food products containing fruits, vegetables, dairy products, fish and meat to enhance their shelf life.
Most of the recombinant proteins have been produced in a variety of plant species for their use as enzymes in industries out of which milk proteins are one which has got nutritional value are being applied in industries and medical field. With the development of HBV vaccine in plants, the concept of oral vaccines implemented via edible plants has garnered increased attention. Plants have also been applied to yield several protein based therapeutic products. The recombinant DNA technology has also been applied to enhance characteristics of plants like in case of the Lycopene β-cyclase genes whose insertion into the tomato’s plastid genome advances the lycopene conversion into provitamin A. Immunity to fungal and bacterial contaminations can be boosted by WRKY45 gene in rice and the Kala4 gene is accountable for the black color of rice making it resistant to offensive pathogens.
Several vegetables and many other plants are being established with necessary characteristics like herbicide tolerance, resistance to pests and drought, salt tolerance. Also, nitrogen consumption, ripening, etc and other types of properties have also been improved. Enhancement in nutritional standards of strawberries has been supported through rolC gene and this gene is known to upsurge the sugar content as well as the antioxidant activity. The two types of enzymes namely glycosyl-transferase and transferase are involved in glycosylation of the Anthocyanins. Some nutrition linked genes for a variety of components in strawberry involving proanthocyanidin, l-ascorbate, flavonoid, polyphenols, and flavonoid are significant for refining the constituent of interest through genetic alteration. In the context of raspberry, bHLH and FRUITE4 genes regulate the anthocyanin mechanisms whereas ERubLRSQ072H02 is linked to flavonol. By specific alteration, these genes have the potential to augment the production and progress the value of the berries.
Genetic engineering has extensive applications in resolving the environmental matters. The development of genetically modified microbes, like Pseudomonas fluorescens for bioremediation solutions in the field was originally accomplished by the University of Tennessee in collaboration with Oak Ridge National Laboratory. The modified strain harboured catabolic naphthalene plasmid pUTK21 and lux gene combined within a promoter resulting in enhanced naphthalene breakdown and a simultaneous bioluminescent tracking. HK44 attends as a reporter for bioavailability and biodegradation of naphthalene and its luminescence marker potential helps it to be applied as an important technology for in situ monitoring of bioremediation practices. The generation of the bioluminescent signal is measurable applying optical fibres and photon detectors.