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9.12: Making Recombinant DNAs - Biology

9.12: Making Recombinant DNAs - Biology


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Molecular biologists often create recombinant DNAs by joining together DNA fragments from different sources. Plasmids used in recombinant DNA methods

  1. replicate in high numbers in the host cell;
  2. carry markers that allow researchers to identify cells carrying them (antibiotic resistance, for example) and
  3. contain sequences (such as a promoter and Shine Dalgarno sequence) necessary for expression of the desired protein in the target cell. A plasmid that has all of these features is referred to as an expression vector (see an example in the figure at left).

Plasmids may be extracted from the host, and any gene of interest may be inserted into them, before returning them to the host cell. Making such recombinant plasmids is a relatively simple process. It involves

  1. cutting the gene of interest with a restriction enzyme (endonucleases which cut at specific DNA sequences);
  2. cutting the expression plasmid DNA with restriction enzyme, to generate ends that are compatible with the ends of the gene of interest;
  3. joining the gene of interest to the plasmid DNA using DNA ligase;
  4. introducing the recombinant plasmit into a bacterial cell; and
  5. growing cells that contain the plasmid. The bacterial cells bearing the recombinant plasmid may then be induced to express the inserted gene and produce large quantities of the protein encoded by it.

7 Main Stages of Recombinant DNA Technology

The following points highlight the seven main stages of recombinant DNA technology. The stages are: 1. Isolation of the Genetic Material (DNA) 2. Cutting of DNA at Specific Locations 3. Isolation of Desired DNA Fragment 4. Amplification of Gene of Interest using PCR 5. Ligation of DNA Fragment into a Vector 6. Insertion of Recombinant DNA into the Host Cell/Organisms 7. Obtaining or Culturing the Foreign Gene Product.

Stage # 1. Isolation of the Genetic Material (DNA):

Nucleic acid is the genetic material, which is present in all living organisms. In majority of organisms, this is present in the form of deoxyribonucleic acid (DNA). DNA must be present into pure form, i.e., free from other macro-molecules (like proteins, RNA, enzymes, etc.) in order to cut the DNA with restrictor enzymes.

Isolation of genetic material (DNA) is carried out in the following steps:

(a) Since the DNA is enclosed within the membranes, so, in order to release DNA along with other macro-molecules such as proteins, polysaccharides and lipids, bacterial cells/plant or animal tissues are treated with the enzyme lysozyme (bacteria), cellulose (plant cells), chitinase (fungus), respectively.

(b) RNA can be removed by the treatment with ribonuclease, whereas proteins can be removed by the treatment with protease.

(c) Other molecules can be removed by appropriate treatments and ultimately purified DNA will precipitate out, after the addition of chilled ethanol. This can be seen as collection of fine threads in the suspension.

Stage # 2. Cutting of DNA at Specific Locations:

Restriction enzyme digestions are performed by incubating purified DNA molecules with the restriction enzyme. This is done at the optimal conditions for that specific enzyme.

Stage # 3. Isolation of Desired DNA Fragment:

Using agarose gel electrophoresis, the activity of the restriction enzymes can be checked. Since, the DNA is negatively charged, it moves towards the positive electrode or anode and DNA tends to separates out in this process. After that the desired DNA fragment is eluted out.

Stage # 4. Amplification of Gene of Interest using PCR:

Polymerase Chain Reaction (PCR) is best defined as the DNA replication in vitro. This techniques was developed by Kary Mullis in 1985 and received Nobel Prize for chemistry in 1993. PCR is used for the amplification of gene of interest using two set of primers.

The basic requirements of a PCR reaction are the following:

The double-stranded DNA that needed to be amplified.

These are chemically synthesised oligonucleotides (short segment of DNA) that are complementary to a region of DNA template.

Two enzymes are commonly used.

It is isolated from a thermophilic bacterium, i.e., Thermus aquaticus. It has a property to remain active during the high temperature which have induced denaturation of double-stranded DNA.

ii. It also helps in the amplification of a segment of DNA to approximately billion times, i.e., I billion copies are made it the process of replication of DNA is repeated many times.

iii. Vent Polymerase (isolated from Thermococcus litoralis).

Three main steps involved in PCR technique are:

The double-stranded DNA is denatured by using high temperature of 95°C for 15 seconds. Now each separated single-strand acts as a template for DNA synthesis.

Two sets of oligonucleotide primers are used to anneal (hybridise). This step is carried out at a slightly lower temperature (40-60°C) using Mg 2+ and dNTPs (deoxynucleoside triphosphates), depending on the length and sequence of the primers.

The thermo-stable enzyme Taq DNA polymerase is used in this reaction, which can tolerate the high temperature of the reaction that extends the primers by adding nucleotides complementary of the template.

Mg 2+ is required as a cofactor for thermo-stable DNA polymerase, e.g., Taq polymerase.

These steps are repeated many times in order to obtain several copies of desired DNA.

Stage # 5. Ligation of DNA Fragment into a Vector:

This process requires a vector DNA and a source DNA. In order to obtain sticky ends, both of these should cut with the same endonuclease. After which both are ligated by mixing vector DNA, gene of interest and enzyme DNA ligase to form the recombinant DNA/hybrid DNA.

Stage # 6. Insertion of Recombinant DNA into the Host Cell/Organisms:

This can occur by several methods, before which the recipient cells are made competent to receive the DNA. If a recombinant DNA bearing gene for resistance to an antibiotic (e.g., ampicillin) is transferred into E. coli cells, the host cells become transformed into ampicillin resistant cells.

The ampicillin resistance gene in this case is called a selectable marker. When transformed cells are grown on agar plates containing ampicillin, only transformants will grow and others will die.

Stage # 7. Obtaining or Culturing the Foreign Gene Product:

When you insert a piece of alien DNA into a cloning vector and transfer it into a bacterial cell, the alien DNA gets multiplied. The ultimate aim is to produce a desirable protein expression. The expressions of the foreign gene or genes in host cells involve understanding of many technical details.

If the protein encoded gene is expressed in the heterologous host, it is called recombinant protein. The cells harbouring cloned genes of interest are grown on small scale in the laboratory. These cell cultures are used for extracting the desired protein using various separation techniques.


Recombinant DNA Technology | Genetics

Recombinant DNA technology has revolutionised life sciences, opening new vistas for research in molecular biology. It allows genetic manipulation using techniques for synthesizing, amplifying and purifying individual genes from any type of cell.

Genomics has emerged as an extension of recombinant DNA technology for high resolution mapping and characterization of whole genomes and gene products on a large scale, referred to as global analysis.

These rapidly advancing disciplines promise new insights into sequence data, organisation, expression, and regulation of the genetic material. Genetic engineering is a natural fallout of these techniques, exploring application in medical, biotechnological industry and agricultural fields.

The success of recombinant DNA technology is based on some key discoveries: restriction enzymes that can cut and join DNA fragments (molecular scissors) for manipulations in a test tube use of plasmids and bacteriophage DNA as vectors (vehicles) for foreign DNA that can replicate into identical copies and produce clones containing recombinant DNA introduction of Southern blotting technique and development of polymerase chain reaction for amplification of a specific sequence.

Recombinant DNA molecules can be purified and investigated for understanding gene structure and sequence, and can be inserted into another genome.

Recombinant DNA molecules are constructed artificially by incorporating DNA from two different sources into a single recombinant molecule. The process is distinct from recombination which is a natural process in sexually reproducing organisms, whereby a single individual gets a combination of genes from two parental organisms.

Natural recombination involves the coming together of similar nucleotide sequences in chromosomal DNA, breakage and exchange of corresponding segments and rejoining. This type of recombination, notably, produces new arrangements of alleles and usually occurs in closely related species. It does not take place in unrelated organisms due to natural barriers.

Restriction Endonucleases:

Restriction endonucleases are a class of enzymes that can recognise and bind to specific DNA sequences of four to eight nucleotides, then cleave the sugar-phosphate backbone of each of the two strands at the site of binding. All restriction enzymes cut DNA between the 3′ carbon and the phosphate moiety of the phosphodiester backbone.

Therefore, fragments produced by restriction enzyme digestion have 5′ phosphates and 3′ hydroxyls. Most restriction enzymes are present in bacteria, only one has been found in the green alga Chlorella. In bacteria, restriction enzymes protect the bacterium against foreign DNA such as that in viruses, by cutting up the invading viral DNA. Thus they restrict entry of foreign DNA into the bacterial cell.

The bacterium modifies its own restriction sites by methylation, so that its own DNA is protected from the restriction enzyme it makes. Three scientists who discovered restriction enzymes and their applications namely, Adams, Nathan and Smith were awarded Nobel Prize in 1978.

More than 400 different restriction enzymes have been isolated. The restriction enzyme EcoRI (from E. coli) recognises the following six nucleotide base pair sequence in DNA of any organism: 5′-GAATTC-3′ and 3′-CTTAAG-5′.

This type of a nucleotide sequence is said to be symmetric, called a palindrome because both strands have the same nucleotide sequence in antiparallel orientation. Several different restriction enzymes recognise specific palindromes. Many restriction enzymes including EcoRI cut the sequence producing staggered ends. EcoRI cuts between G and A (Fig. 23.1).

The staggered cuts give rise to a pair of identical five nucleotide long single-stranded “sticky ends”. The ends consist of an overhanging piece of single-stranded DNA that are said to be sticky because they can base pair by hydrogen bonding to a complementary sequence. Sticky ends produced by restriction enzymes are desirable in recombinant DNA technology.

If two different DNA molecules are cut with the same restriction enzyme, the fragments of each will have the same sticky ends that are complementary, enabling them to hybridise with each other under appropriate conditions. Some restriction enzymes such as small cut the target sequence in the middle producing blunt ends which lack sticky ends.

These can also be used for making recombinant DNA molecules with the help of enzymes that join blunt ends, or other special enzymes that synthesise short single-stranded sticky ends on the exposed 3′ strand of the blunt end.

Digestion of DNA by a restriction enzyme yields fragments of different lengths. For example, bacteriophage λ DNA cut with restriction enzyme EcoRI produces six fragments ranging in size from 3.6 to 21.2 kilo bases in length (one kilo-base or kb = 1,000 base pairs). These fragments can be separated according to size by gel electrophoresis (Figs. 23.2, 3) or by other methods.

The fragments together provide a map of the EcoRI sites in phage λ DNA. If multiple different restriction endonucleases are used, the locations of their cleavage sites can be used to generate detailed restriction maps of the DNA molecule.

Gel Electrophoresis for Separation of DNA Fragments:

The cells are homogenized and nuclei isolated for extraction of DNA. To lyse nuclei and release DNA, a negatively charged detergent sodium dodecyl sulphate (SDS) is used. The next step involves purification of DNA from contaminants such as RNA and proteins. To remove proteins, the mixture is shaken with phenol or chloroform.

Phenol makes the proteins insoluble and precipitates them out of solution. Because phenol and buffered saline are immiscible, they form two separate phases. Centrifugation gives DNA or RNA in the upper aqueous phase, while protein precipitate forms the boundary between the two phases.

The aqueous phase containing the nucleic acid is removed from the tube and shaken with phenol, followed by centrifugation. The process is repeated until no further protein can be removed from solution. Cold ethanol is then layered on top of the aqueous DNA solution, and the DNA is taken out with a glass rod at the interface between the ethanol and saline. RNA forms a precipitate that settles at the bottom of the vessel.

To make the DNA preparation free from RNA, the DNA is treated with ribonuclease. The ribonuclease is destroyed by treatment with protease, and the protease is removed by using phenol. Purified DNA is then re-precipitated with ethanol.

Electrophoresis depends on the ability of charged molecules to migrate in an electric field. The small DNA fragments of a few hundred nucleotides or less, produced by a restriction endonuclease, can be separated by polyacrylamide gel electrophoresis (PAGE). The larger DNA fragments ranging in size from a few hundred base pairs to about 20 kb can be separated on agarose gels.

The mechanism for separation of DNA fragments involves migration of DNA molecules through the pores of the matrical gel. Agarose consists of a complex network of polymerised molecules, the pore size of which is determined by the composition of the buffer and concentration of agar used.

Visualisation by fluorescence microscopy has revealed that during electrophoresis DNA molecules display stretching in the direction of applied field and then contract into dense balls. The size of the DNA ball must be smaller than pore size in the gel to pass through. If the volume of the DNA ball exceeds that of pore size in gel, then the DNA molecule can only pass through by a serpentine motion resembling that of a snake.

DNA molecules up to 20 kb can migrate through pores in gel by this mechanism. Studies have shown that both fragment length and molecular weight determine rate of migration of DNA molecules in a gel. For accurate size determination of a DNA molecule, marker DNA samples of known size are electrophoresed in the same gel.

The procedure for polyacrylamide gel electrophoresis is as follows. A mixture of linear DNA fragments of different sizes are driven by the current through the gel composed of organic molecules of acrylamide. Cross linking of acrylamide molecules forms a molecular sieve in a thin slab of the polyacrylamide gel between two glass plates. The gel slab is suspended between two compartments containing buffer in which the positive and negative electrodes are immersed (Fig. 23.2).

The sample containing linear DNA molecules (fragments) is placed in wells along the top of the gel, near the cathode of the electric field. Voltage applied between the buffer compartments allows current to flow across the slab. Because of its negatively charged phosphate groups, DNA migrates towards the positive electrode (anode) at speeds inversely dependent on their size. That is, the smaller fragments migrate most rapidly.

All DNA molecules regardless of their length, have a similar charge density, that is, the number of negative charges per unit of mass, and therefore, all have equivalent potential for migration in the electric field. The greater the molecular mass of the DNA fragment, the more slowly it moves through the gel. Hence, the smaller the fragment, the farther it migrates in the gel.

Therefore, if there are distinct size classes in the mixture of DNA fragments, these classes will form distinct bands on the gel. The fragments of different sizes thus get separated from each other (Fig. 23.3). The bands can be visualised by staining the DNA with ethidium bromide.

The migration of the DNA fragments can be compared with a set of control size standards that were loaded in the same gel, to determine the exact size of each fragment in the mixture. Moreover, if the bands are well separated, a single band can be cut from the gel and the DNA sample can be extracted and purified.

Larger DNA fragments that do not pass through the pores of polyacrylamide, are fractionated on agarose gels which have greater porosity. Agarose is a sea weed polysaccharide, it is dissolved in hot buffer and poured into the mould and a comb placed in the molten agarose. Lowering the temperature solidifies agarose as a gel. The comb is removed producing wells in the gel. DNA samples are loaded in the wells.

Gels having 0.3% concentration of agarose are used to separate larger DNA fragments. After running the gel, DNA fragments can be identified by using a labeled probe, for a specific fragment. Alternatively, the gel can be stained with a solution of ethidium bromide which intercalates into the bases in the double helix. The DNA bands can be viewed under ultraviolet light using a trans-illuminator.

Pulse-Field Gel Electrophoresis:

The separation of DNA molecules greater than 20 kb, up to 10 Mb can be accomplished by use of pulse-field gel electrophoresis (PFGE). In PFGE, the orientation of the electric field with respect to the gel is changed in a regular manner, which causes the DNA to periodically alter its direction of migration on the gel.

Each time there is a change in the electric-field orientation, the axis of the DNA must realign before it starts migration in the new direction. The difference between the direction of migration of DNA induced by successive electric fields determines the angle through which DNA must turn in order to change its direction of migration. To make the sample run in straight lines, improved methods for alternating the electric field have been devised. PFGE is used extensively in molecular biology laboratories.

Southern Blotting:

When genomic DNA is subjected to restriction enzyme digestion it results in a very large number of fragments. A stained gel containing numerous fragments separated by electrophoresis shows a continuous smear of DNA instead of distinct bands. The technique of Southern blotting developed by E. M. Southern allows a single DNA fragment or a specific gene to be identified in this mixture.

This is a widely used technique that is based on labelling of DNA and hybridisation on membranes. It is referred to as a blotting technique because it involves transfer of nucleic acids from gels to a solid support of a membrane and immobilisation of DNA on to the membrane. Detection of the DNA fragment is done by using a complementary strand of DNA or RNA that is called probe (something that detects is a probe) which hybridizes with the DNA fragment.

The transfer of DNA from gel to membrane is accomplished by the flow of buffer through the wick, gel, membrane, and above onto adsorbent paper layers. The gel is overlaid on a filter paper wick (3 to 4 sheets) which dips into the buffer contained in a vessel (Fig. 23.4). The hybridisation membrane is placed above the gel.

A stack of paper towels comes above the membrane. The adsorbent paper towels serve to draw the buffer through the gel by capillary action. The flow of buffer carries the DNA molecules out of the gel on to the membrane (nitrocellulose or nylon membrane). Nylon membranes are considered superior in having greater binding capacity for nucleic acids and being stronger than membranes consisting of nitrocellulose.

Large DNA fragments require longer time to transfer out of gel than shorter fragments. To overcome such time differences, the electrophoresed DNA on the gel is pretreated for depurination, which also denatures the fragments into single strands, making them accessible for hybridisation with the probe.

After transfer from the gel, the DNA fragments are attached (permanently fixed) to the membrane by, heating at 80°C in the oven or by cross-linking using UV radiation. The DNA fragments become fixed or imprinted to the membrane at the same locations as in the gel (replica).

In other words, the spatial arrangement of DNA in the gel is preserved in the sheet. Following fixation, the membrane is incubated in a solution containing labelled single-stranded DNA or RNA probe that is complementary to the blot transferred DNA sequence to be detected. Under appropriate conditions, the labelled nucleic acid probe hybridizes with the DNA on the membrane.

After hybridisation the membrane is washed to remove unbound probes, so that the only labelled molecules left are those that hybridised with target DNA. The membrane is then placed in contact with X-ray film for autoradiography that will reveal the position of the desired DNA fragment.

By using size-calibration controls, the size of any fragment from the mixture of fragments can be determined. Variations of Southern blotting are called Dot and Slot blots. The sample is blotted directly on the nylon or nitrocellulose sheet without prior separation on the gel.

This technique is similar to the Southern blotting technique and is used for identifying a specific RNA molecule from a mixture of RNAs separated on a gel. The separated RNA molecules on a gel are blotted on to a membrane and probed in the same way as DNA is blotted and probed in Southern blotting. A cloned gene having complementary sequence to the RNA being searched, can be used as a probe.

Stated briefly, it is a combination of the recent remarkable techniques that are finding widespread application. Generation of restriction fragments, gel electrophoresis, blotting techniques, cloning, together with nucleic acid hybridisation allow a specific DNA sequence to be analysed in a whole genome.

To identify an individual gene, the DNA from the organism is purified and cleaved by restriction enzymes. The restriction fragments are separated by gel electrophoresis. The fragments are denatured to get single strands, then labelled with a radioactive probe.

The probe consisting of single-stranded DNA or RNA must have a sequence that can base pair with the gene of interest. The probe hybridises with its complementary sequence under appropriate conditions of temperature, salt and pH. Exposure to X-ray film indicates the binding of the probe through the presence of the radioactive signal.

Recombinant DNA Molecules:

The generation of recombinant DNA molecules is a process of inserting a gene of interest or donor DNA into a DNA molecule from a different source that acts as a vector or vehicle for the donor DNA. The first step is to obtain donor DNA, that is, isolate the gene of interest. Genomic DNA from the donor organism is isolated, purified and cut with a restriction endonuclease that makes staggered cuts with sticky ends in the donor DNA.

The sticky ends consist of overhanging or cohesive single-stranded tails. Thus donor DNA will be cut into a set of restriction fragments according to the locations of the restriction sites. The vector DNA, which could be a bacterial plasmid or genome of bacteriophage λ (described later), is also cut with the same restriction enzyme that was used for donor DNA.

The fragments thus produced would have the same complementary sticky or cohesive ends, enabling them to hybridise with sticky ends in the donor DNA fragments. The hybridised molecules produced do not have complete covalently joined sugar phosphate backbones. To seal the backbones, the enzyme DNA ligase is used which makes phosphodiester bonds and links the two covalently to form a recombinant DNA molecule (Fig. 23.5).

Other Sources of Donor DNA:

Besides genomic DNA from a donor organism, it is possible to use RNA sequences as donors. The first step is to synthesise a DNA copy of the RNA using the enzyme reverse transcriptase. The complementary DNA or cDNA obtained can be used as donor DNA by incorporating it into vector DNA and making a recombinant DNA molecule.

In studies that require characterisation of the gene transcript, that is mRNA, a cDNA is prepared by using mRNA as a template for reverse transcriptase. This is particularly useful because mRNA is short-lived and techniques for isolating individual mRNA molecules are not available. cDNAs can be used to learn about the variety of mRNAs in a cell, and the number of copies of different mRNAs present in a cell.

If the sequence to be inserted for making a recombinant DNA molecule cannot be isolated from natural genomic DNA, nor as cDNA, then chemically synthesised DNA can be used. Chemical synthesis of oligonucleotides, that is, DNA fragments 15 to 100 nucleotides in length can be developed by highly automated techniques.

Vectors for Generating Recombinant DNA:

A vector must be a small molecule capable of independent replication in a living host cell must have convenient restriction sites that can be used for insertion of the DNA to be cloned must permit easy identification and recovery of the recombinant molecule. Vectors are also referred to as cloning vehicles or replicons. The basic vector systems use bacterial plasmid or bacteriophage λ DNA as vectors.

Plasmids are small circular DNA molecules that replicate independently of the bacterial chromosome. Plasmid molecules are partitioned accurately to daughter cells. Most plasmids exist as double-stranded DNA molecules.

If both strands of the DNA are intact circles, the molecule is described as a covalently closed circle DNA (CCC DNA), if only one strand is intact, then open circle DNA (OC DNA). Not all plasmids are circular, some exist as linear molecules such as those in Streptomyces and Borrelia.

Plasmid DNA is 2 to 4 kb in length and has a sequence which is origin of replication, that signals the host cell DNA polymerase to replicate the DNA molecule. Plasmids have the advantage of carrying genes for resistance to antibiotics, so bacteria carrying antibiotic-resistance phenotype (plasmids) can be selected.

Suppose we want to insert gene X in human genome into a plasmid vector. Fragments of human DNA are prepared by cutting with a restriction enzyme, for example EcoRI, and the same restriction enzyme is used to cut plasmid DNA (Fig. 23.3).

The sticky ends will have complementary single-stranded cohesive ends that would hybridise with each other. Among the large number of fragments produced from human DNA, only a very small fraction would have gene X.

To isolate the fragment with gene X, plasmid and human DNA restriction digests are incubated together, along with DNA ligase. During incubation, the sticky ends of the two types of DNA become hydrogen-bonded to each other, their broken ends sealed by ligase to form circular recombinant DNA molecules.

Now there would be a large number of different recombinant DNA molecules, each containing a bacterial plasmid with a human DNA fragment. To isolate those plasmids that have human gene X, the process of DNA cloning has to be done.

Bacteriophage λ vectors can carry foreign DNA inserts as large as 15 kb. There are two basic types of phage λ vectors, insertional vectors and replacement vectors. The wild type phage λ DNA contains several target sites for most of the commonly used restriction endonucleases, hence is not suitable as a vector.

Derivatives of the wild type phage have, therefore, been produced that either have a single target site at which foreign DNA can be inserted (insertional vectors), or have a pair of sites defining a fragment that can be removed and replaced by foreign DNA (replacement vectors).

Since phage DNA can accommodate only about 5% more than its normal complement of DNA, vectors are constructed with deletions to increase space within the genome. The shortest λ DNA molecules generally used are 25% deleted.

Foreign DNA is first ligated to phage DNA (Fig. 23.6). The recombinant molecules are then introduced into E. coli cells. DNA replication produces numerous phage progeny containing the foreign DNA fragment. This fragment can be isolated from the rest of phage DNA by restriction endonuclease digestion and gel electrophoresis.

Phagemids are plasmid vectors that carry the origin of replication from the genome of a single-stranded filamentous bacteriophage such as M13 or f1. Phagemids combine the best features of plasmids and single-stranded bacteriophage vectors.

They have two separate modes of replication: as a double-stranded DNA plasmid, and as a template to produce single-stranded copies of one of the phagemid strands. A phagemid can therefore, be used in the same way as a plasmid vector, or it can be used to produce filamentous bacteriophage particles that contain single-stranded copies of cloned segments of DNA.

Some studies require large fragments of foreign DNA to be inserted for which phage λ vectors are not suitable. Artificially prepared cosmid and yeast artificial chromosome (YAC) vectors are useful for this purpose. Foreign DNA up to 45 kb in length can be accommodated in cosmid vectors. Cosmid vectors can exist as plasmids but they also contain the complementary overhanging single-stranded ends of phage λ.

The presence of bacteriophage λ sequences in cosmid vectors permit packaging of the recombinant DNA into phage particles. The λ phage then introduces these large sized recombinant DNA molecules into recipient E. coli cells. Cosmids also contain origins of replication and genes for drug-resistance, so that they can replicate as plasmids in bacterial cells (Fig. 23.7).

However, because cosmid vectors lack the essential phage sequences necessary to form progeny phage particles, the recombinant DNA molecule depends on the plasmid sequences in the cosmid. Once inside the recipient E. coli cell, the recombinant cosmids form circular molecules that replicate extra-chromosomally in the same manner as plasmids.

The YAC vectors can accommodate still larger fragments, from a hundred to a thousand kilo-bases in length, that is one million base pairs. As the name implies, YACs are artificial versions of normal yeast chromosomes and replicate as chromosomes in yeast cells.

They contain all the elements of a yeast chromosome that are required for replication during S phase, one or more origins of replication, and segregation to daughter cells during mitosis, telomeres at the ends of the chromosome, as well as centromere to which spindle fibres can attach during chromosome separation.

In addition to these elements, YACs are constructed to contain the following: a gene whose encoded product allows those cells containing the YAC to be selected from those that lack the element the DNA fragment to be cloned. Yeasts can take up DNA from the medium, allowing YACs to be introduced into host yeast cells.

For introducing such large-sized DNA fragments (100 to 1000 kb in length), it is necessary to use restriction enzymes that recognise particularly long nucleotide sequences, 7 to 8 nucleotides containing CG di-nucleotides.

For example, the restriction enzyme NotI recognises the 8 nucleotide sequence GCGGCCGC, which cleaves mammalian DNA into fragments approximately one million base pairs long. These fragments can then be incorporated into YACs and cloned within host yeast cells. YAC technology has been used extensively in the Human Genome Project.

Bacterial artificial chromosomes (BACs) are based on the F factor of E. coli, and are among the most widely used vectors for very large DNA fragments of up to 300 kb. BAC vectors are 6 to 8 kb in length and include genes for essential functions such as replication (genes repE, and oriS), genes for regulating copy number (parA and parB), and genes for resistance to the antibiotic chloramphenicol.

Genomic DNA Libraries:

Collections of DNA fragments obtained by DNA cloning from the entire genome of an organism constitute a DNA library. Basically, the genomic library is a collection of bacteria, typically E. coli, each carrying a fragment of DNA from the genome. In one approach for making a DNA library, all the DNA from an organism is cut into fragments with a restriction enzyme.

Each segment is inserted into a different copy of the vector, thereby creating a collection of recombinant DNA molecules, which collectively represent the entire genome. These are then used to transform separate recipient bacterial cells, where they are amplified. The resulting collection of recombinant DNA-bearing bacteria is called a genomic library.

If the cloning vector used could accommodate an average insert size of 10 kb, and if the entire genome size is 100,000 kb, then we can expect 10,000 independent recombinant clones to represent the library of the whole genome.

In another approach, genomic DNA is cleaved by one or two restriction enzymes that recognize very short nucleotide sequences such as Sau3A (recognises GATC) and HaeIII (recognises GGCC). The enzymes are used in low concentration so that only a small percentage of target sites are actually cleaved.

One can expect that the small-sized tetra-nucleotide sequence would occur by chance at very high frequency, so that every portion of the DNA could yield fragments. After partial digestion of DNA, the fragments are separated by gel electrophoresis. Fragments about 20 kb in length are incorporated into phage λ heads from which about half a million plaques are generated, to ensure that every portion of the genome is represented (Fig. 23.8).

The important point in this method is that, due to low concentration of the restriction enzyme, the DNA is randomly fragmented, and each and every target site is not cleaved. The phage recombinants produced constitute a permanent collection of all the DNA sequences in the genome.

Whenever a particular sequence is required for isolation from the library, phage can be grown in bacteria. Each of the plaques produced would have originated from infection of a single recombinant phage, and can be screened for the presence of a particular sequence.

Positional Cloning or Chromosome Walking:

When there is no biological information about a gene, but its position can be mapped relative to other genes or markers, it is called positional cloning. The approach involves cloning a gene from its known closest markers. It requires only the mapped position of the gene. On the basis of this information, researchers can locate the nearest physical markers.

The closest linked marker is used to probe the genomic library. DNA cleaved randomly, as described above, generates overlapping fragments that can be used in the analysis of regions of the chromosome extending out in both directions from a particular sequence, the gene of interest. These extended regions or linked markers serve as starting points for the process of chromosome walking (Fig. 23.11).

Using a small sequence at the end of the linked marker, let us call it M1, as a labelled probe, find a clone in the library. The positive clone that is found will lead to identification of an adjacent gene segment which is now a second marker. Isolate the end sequence of second marker (M2) and use it to probe library and find another adjacent genomic segment which becomes the third marker.

Isolate end of third marker (M3) and use it to again probe for the next adjacent segment, and so on. Using the new fragments as labelled probes the process is repeated in successive screening steps, leading to isolation of more and more of the original DNA molecule. Because this process consists of steps, hence the name chromosome walking. This approach allows study of the organisation of linked sequences over a considerable length of the chromosome.

Chromosome jumping is a variation of chromosome walking using larger high capacity vectors to bridge un-clonable gaps. Whereas in chromosome walking each step is an overlapping DNA clone, in chromosome jumping each jump is from one chromosome location to another without “touching down” on the intervening DNA.

The gene for cystic fibrosis (CF), a severe autosomal recessive disorder in humans, was identified by chromosome walking and jumping. Cloning of the CF gene was a breakthrough for studying the biochemistry of the disorder (abnormal chloride channel function), for designing probes for prenatal diagnosis, and for potential treatment by somatic gene therapy or other means.

The method described above for making genomic DNA libraries can also be used for generating cDNA libraries. A cDNA library would contain hundreds of thousands of independent cDNA clones, representating collections of cDNA inserts. It may be recalled that cDNA is produced from the mRNA by using reverse transcriptase.

If a specific gene that is being actively transcribed in a particular tissue is desired for study, then it is considered useful to convert its mRNA into cDNA and make a cDNA library from that sample. A cDNA library represents a subset of the transcribed regions of the genome, hence the cDNA library would be smaller than a complete genomic library.

Screening DNA Library for a Specific Clone:

Any sequence for which a probe is available can be isolated from a recombinant library. Two types of probe can be used, those that recognise a specific DNA sequence and those that recognise part of a specific protein.

Probes for DNA Sequences:

A probe that consists of a single strand of DNA would be able to find and bind to other complementary denatured (single-stranded) DNAs in the library and specifically hybridise with it. The procedure for identification of a specific clone in a library is carried out in two steps. First, the recombinant phages are plated on E.coli, and each phage replicates to produce a plaque on the lawn of bacteria.

The pattern of plaques of the library on the petri dish are transferred to an absorbent nitrocellulose membrane by laying the membrane directly on the surface of the medium. When the membrane is peeled off, the plaques remain clinging to its surface, are lysed in situ and DNA is denatured. The membrane is incubated in a solution containing radiolabeled probe that is specific for the sequence being searched (Fig. 23.9).

Generally, the probe is a cloned piece of DNA that has a sequence homologous to that of the desired sequence. The single-stranded probe will bind to the DNA of the clone being searched. To determine the position of the positive clone, the position of the radioactive label can be found out by placing the membrane on the X-ray film.

Emissions from the decay of radioactive label will reduce the grains in X-ray film, seen as a dark spot after developing the film. The procedure is called autoradiography, the exposed film an autoradiogram. The probe used for finding sequence of interest in a library can also be labelled with a fluorescent dye.

In that case the membrane is exposed to a particular wavelength of light that would excite fluorescence and a photograph of the membrane is taken. The position of the spot of label (radioactive or fluorescent) indicates the location of the DNA segments containing sequences complementary to the probe.

Probes for Protein Products of Genes:

The protein product of a gene can be used to find the clone of its corresponding gene in a library. This can be accomplished if the amino acid sequence of the protein product is known, and the protein can be isolated in a purified form. Antibodies that bind specifically with unique protein molecules are used as probes to screen an expression library. These libraries are special cDNA libraries generated by using expression vectors.

An expression vector is a cloning vector containing the regulatory sequences necessary to allow transcription and translation of a cloned gene. Expression vector produces the protein encoded by a cloned gene in the transformed host. Expression vectors are essentially derivatives of a phage or plasmid cloning vector that has been modified by addition of a promoter specific to the host.

The cloned gene in such an expression vector is placed under the control of the promoter that ensures transcription of the cloned gene in the appropriate host cell. Expression vectors designed in this manner can yield high levels of recombinant protein in host cells that can be purified for structural and functional characterisation.

To make the cDNA library, the cDNA to be cloned is inserted into the special phage vector downstream from the bacterial promoter, in the correct triplet reading frame which ensures that the foreign DNA is transcribed and translated during infection.

Phages that have incorporated the gene of interest form plaques that contain the protein encoded by that gene. A membrane is laid over the surface of the medium and removed with some cells of each colony attached to the membrane.

The locations of these cells are identical to their positions in the original petri dish (replica plating). The membrane is dried, and immersed in a solution containing antibody. The antibody will bind only to the protein product of the gene of interest.

For detection of positive clones, a second antibody is prepared that is specifically against the bound antibody and labeled radioactively or with a fluorochrome. The plaque having the gene of interest is thus located on the replica plate through detection of bound antibody.

It is frequently considered useful to express high levels of a cloned gene in eukaryotic cells rather than in bacteria. The reason is that post-translational modifications of the protein (such as addition of carbohydrates or lipids) that take place in eukaryotic cells would take place normally.

One system frequently used for protein expression in eukaryotic cells involves infection of insect cells by baculovirus vectors, which yield very high levels of protein product of a gene. High levels of protein expression can also be achieved by using appropriate vectors in mammalian cells.


Examples of Recombinant DNA Technology

Recombinant DNA technology is used in a number of applications including vaccines, food products, pharmaceutical products, diagnostic testing, and genetically engineered crops.

Vaccines

Vaccines with viral proteins produced by bacteria or yeast from recombined viral genes are considered safer than those created by more traditional methods and containing viral particles.

Other Pharmaceutical Products

As mentioned earlier, insulin is another example of the use of recombinant DNA technology. Previously, insulin was obtained from animals, primarily from the pancreas of pigs and cows, but using recombinant DNA technology to insert the human insulin gene into bacteria or yeast makes it simpler to produce larger quantities.

A number of other pharmaceutical products, like antibiotics and human protein replacements, are produced by similar methods.

Food Products

A number of food products are produced using recombinant DNA technology. One common example is the chymosin enzyme, an enzyme used in making cheese. Traditionally, it is found in rennet which is prepared from the stomachs of calves, but producing chymosin through genetic engineering is much easier and faster (and does not require the killing of young animals). Today, a majority of the cheese produced in the United States is made with genetically modified chymosin.

Diagnostic Testing

Recombinant DNA technology is also used in the diagnostic testing field. Genetic testing for a wide range of conditions, like cystic fibrosis and muscular dystrophy, have benefited from the use of rDNA technology.

Crops

Recombinant DNA technology has been used to produce both insect- and herbicide-resistant crops. The most common herbicide-resistant crops are resistant to the application of glyphosate, a common weed killer. Such crop production is not without issue as many question the long term safety of such genetically engineered crops.


Steps in Recombinent DNA Technology

Isolate DNA then cutting the DNA with restriction enzymes after that ligate into cloning vector then transform recombinant DNA molecule into host cell so that each transformed cell will divide many, many times to form a colony of millions of cells, each of which carries the recombinant DNA molecule (DNA clone).

  1. Isolation of Desired Gene
  2. Preparation of Vector
  3. Ligation
  4. Introduction of Recombinant DNA into Host cell
  5. Selection of recombinant host cell
  6. Expression of cloned gene

Isolation of desired DNA

For making recombinant organisms, a desired DNA fragment has to be introduced into the host cell, these desired DNAs can be obtain from the total genome of the cell either by restriction digestion (with restriction endonucleases or by mechanical shearing). The desired gene can be clone is located in the cell DNA of the source organisms along with several genes so firstly, it should be isolated from other genes of the cell DNA. For this purpose two methods are follows to isolate desired DNA from the genome of the cell. They are restriction digestion and mechanical shearing.

DNA Extraction

  1. Preheat 5ml CTAB (add 10μl mercaptoethanol to each 5ml CTAB) in a blue-topped 50ml centrifuge tube at 60-65°C. Remove and discard midribs and wrap laminae in aluminum foil and freeze in liquid nitrogen. 0.5 – 1.0 gm tissue/5ml CTAB (Can store leaf material after liquid Nitrogen – 1-2 days at –20°C or –80°C for longer periods)
  2. Gently crumble leaf tissue over cold pestle of liquid nitrogen. Grind frozen leaves with one spatula of fine sand add 0.5 spatula of PVPP powder after grinding.
  3. Scrape powder into dry tube and add pre-heated buffer and mix gently. Avoid leaving dry material around rim of tube. Adjust CTAB volume to give a slurry-like consistency, mix occasionally.
  4. Incubate for 60 min at 60°C
  5. Add equal volume of chloroform/iso-amyl alcohol (24:1), Mix for about 3min, then transfer contents to narrow bore centrifuge tubes. Balance by adding extra chlor/iso. Spin 5,000rpm for 10min (ensure correct tubes used), brake off. (For extra pure DNA isolation – spin and retain supernatant before chloroform extraction).
  6. Remove supernatant with wide-bore pastette (cut off blue tip) to clean tube, repeat chloroform extraction once. Supernatant should be clear, though may be coloured.
  7. Precipitate DNA with 0.66 vol. of cold isopropanol – can leave overnight. Spool out or spin down DNA, 2min at 2,000rpm.
  8. Transfer to 5ml wash buffer for 20min.
  9. Dry briefly and resuspend in 1ml T.E. (can be left overnight)
  10. Add 1μl 10mg/ml RNase to each 1ml T.E./DNA mixture and incubate for 60min at 37 °C. (If RNase in the sample doesn’t matter – stages 11 and 12 may be omitted)
  11. Dilute with 2 volumes TE and add 0.3vol 3M Sodium acetate (pH 8) +2.5 vol. cold 100% ethanol
  12. Spool DNA out. Air dry and resuspend in 0.5 to 1ml TE or water (takes time) and freeze until required.

Cutting DNA

Restriction Digestion

Restriction digestion is the cutting of DNA into fragments by restriction enzymes.The total DNA of an organism is treated with restriction enzymes. These enzymes cut the cell DNA into many fragments each fragment is differing in their size and molecular weight. These restriction enzymes cut the DNA into blunt end and cohesive end depending upon the mixture of these DNA fragments is electrophoresed on an agarose gel for a particular time By electrophoresis the mixture of this DNA fragments get separated on the basis of their size and molecular weight. Each DNA fragment obtained from electrophoresis band is cloned into a suitable vector.

The restriction enzyme may cut the DNA at the middle of desired gene and make it useless.

Mechanical Shearing

Genomic DNA is subjected to mechanical forces to cut genomic DNA into small fragments. Sonication with ultrasound cut this DNA randomly with the size of about 300-325 bp.

Limitations

Each time it produces new DNA fragments This method cannot be applied for Eukaryotic organism because of presence of introns.

Joining DNA

Recombinant DNA is a hybrid DNA formed by joining a desired foreign DNA and a vector DNA, It is indicated by rDNA. The vector DNA and genomic DNA fragments are mixed together. Cohesive end of the vector DNA anneals with genomic DNA fragment by complimentary base pairing and there is nick between these two DNA ends.

This nick is sealed by an enzyme called DNA ligase, It makes the Phosphodiester bond in between these two DNA fragments. Sometimes adapters, linkers and homopolymer tail are also used to join blunt ended DNA molecules.

Transformation

The recombinant DNA can be introduced into the host cell by direct transformation and indirect transformation. In direct method, bacterial cell intake the recombinant DNAs in the medium like microinjection, liposomes, electroporation and particle bombardment method. In indirect method the pathogenic agents such as bacteriophage and Agrobacterium pick up the recombinant DNA and introduced it into plant cell.

DNA clone is a section of DNA that has been inserted into a vector molecule and then replicate in a host cell to form several copies.


DNA Structure, Replication, and Technology - Recombinant DNA

In the most recent Spider-Man movie, a mutant spider bit Peter Parker and passed its spider genes to him, giving him super strength, vision, and web-slinging action. Therefore, in this case, Spider-Man exists because recombinant DNA from spider genes entered the human genome of Peter Parker.

If you are a Spider-Man fan, you will know the original story had Peter Parker bitten by a radioactive spider. Unfortunately, that method of transferring spider powers is highly unlikely, knowing what we know about DNA. Making recombinant DNA involves taking a gene, usually from an eukaryote, and putting it into a bacterial plasmid.

Many people fear that manipulating recombinant DNA is akin to "playing God," and some non-Spider-Man films play off those fears with the evil scientist character. Movies like Rise of the Planet of the Apes, The Island of Doctor Moreau, and Jurassic Park all feature evil scientists doing evil science with evil recombinant DNA. However, this technology has far more advantages than disadvantages, and the chances of creating super apes that will wipe out humanity are very small. Hey, we at Shmoop admit that super apes are super scary.

We will discuss the advantages and disadvantages of recombinant DNA technology in the "Biotechnology" section. For now, let's pretend we're scientists making recombinant DNA. And just for fun, let's be evil about it.

Yes! Even You Can Make a Super Ape!

So you want to make an army of super apes, but where do you start? While the following section is completely hypothetical, it shows how scientists make genetically recombinant organisms.

Disclaimer: do not tell your friends after reading this that you know how to make an army of super apes. That secret will always remain in the vaults at Shmoop. We can neither confirm nor deny that Shmoop has an army of super apes.

Let's make some hypothetical super apes. First, you start off with a vector, or a specific plasmid that is used for generating recombinant DNA. Vectors come in a variety of forms, but they must have three sequences:

  1. They need an origin of replication (oriR) site, which is a sequence that signals bacteria to replicate this plasmid whenever they replicate.
  2. There must be an antibiotic resistance gene, which is important for selecting bacteria that have the recombinant plasmid. When a scientist grows bacteria, he will add an antibiotic to kill all bacteria that do not have the plasmid. The antibiotic resistance gene will allow the bacteria that have the plasmid to stay alive. Score.
  3. A multiple cloning site (MCS) is important for facilitating insertion of the target gene into the vector. Putting the MCS into a lacZ or ccdB gene allows selection for bacteria that have the target gene inserted into the vector. This is because the target gene will disrupt the lacZ or ccdB gene, and one can easily test the activity of either of these genes by looking for colonies on a plate that lack the relevant gene activity.

A multiple cloning site is a DNA sequence that contains many restriction enzyme sites. Restriction enzymes cleave DNA at specific palindromic sequences, such as GGATCC, GAATTC, and GTTAAC. There are thousands of different restriction enzymes that each cleave a specific sequence. MCS sequences have sites for common restriction enzymes, and these enzymes were designed to cut the DNA only in the MCS and nowhere else in the vector.

Restriction enzymes cut in a way that either leaves sticky ends, where each strand is cut at a different position, leaving 2–5 bases of single-stranded DNA, or blunt ends, where the same spot on both strands is cleaved. Sticky ends are generally easier to handle, and different restriction enzymes often leave the same overhanging sequences, so you can cut your vector with one enzyme and your DNA insert with another enzyme, and the two sticky ends are then called compatible. Check out this link. Match.com could learn something from restriction enzymes.

Once you have cut your vector in the MCS, or elsewhere in the vector that has a unique restriction site, you need to join it together. How do you join the two sequences? Naturally, you join them in the same way you join everything else: glue!

Wait, not actual glue. Elmer's does not make bottles that small. Instead, we use a "molecular glue" called DNA ligase. DNA ligase is an enzyme that takes a free phosphate from the 5' end of one piece of DNA and attaches it to the free –OH at the 3' end of the other piece of DNA.

Above is a schematic of how most scientists make recombinant DNA plasmids. The target gene here is our SUPER APE gene. The only concern with making a plasmid is that there is a DNA sequence limitation. You cannot insert a DNA sequence into a traditional vector if it is over 20 kilobases (kb) in length. What if your hypothetical SUPER APE gene is huge (> 300 kb)? That will put a damper on your plot of world domination, right? Not at all, my fiendish friend!

Our scientist minions, er, friends have developed cosmids, or plasmids that can handle large pieces of DNA and contain cos sequences from viruses that infect bacteria, called bacteriophages. Cosmids can handle 50-kb inserts because they integrate into the bacterial genome rather than hanging outside of the bacterial genome, like plasmids.

If you want even more space for DNA, you can use bacterial artificial chromosomes (BACs), which handle 150–350 kb sequence inserts. BACs are similar in size to most other plasmids (5–7 kb in length), though they contain parA and parB genes, as well as oriS sites and the repE and repF genes for regulating the BAC (and its insert) so that it is properly replicated, maintained, and segregated into progeny cells. They basically act like a second bacterial genome in bacteria, and those sequences make sure your giant SUPER APE gene is replicated in the bacteria. Stick your SUPER APE gene into the BAC, and you are all set for world domination.

Brain Snack

I Have Made a Plasmid: Now What?

All those months working diligently in the lab to make your SUPER APE plasmid have come to fruition (Heaven help us all…). You're ready to lead an army of super apes. But, how do you go from plasmid to super ape?

There are two strategies, and it depends on the biology of your SUPER APE gene. Sorry, you need to do more research on how SUPER APE makes apes super. There are a few possibilities on why current apes are not super: either they lack the enzyme that makes them super, or they have the SUPER APE gene already but it is defective.

Let's say that apes have a dysfunctional SUPER APE gene. We can put a functional SUPER APE gene into ape embryos, and they will make the super ape gene. How does one insert a gene into an embryo? One way is to apply an electric current to the embryo, which opens the cells to take in our SUPER APE BAC. Alternatively, you can take a tiny syringe and inject the DNA into the embryo. However, both methods have several problems and are incredibly difficult to do.

There is potentially a major biological concern if we add functional SUPER APE genes to an embryo. Inserting DNA into an embryo will lead to every cell in the body having that DNA. Perhaps not every part of the body needs a functional SUPER APE gene to make the ape super. What if SUPER APE kills apes when expressed in the heart or lungs? This might be whythe gene is nonfunctional in all apes.

You tried adding your plasmid to embryos, and all your apes died or were never born. That is a bummer, and not just because you have a lot of dead apes on your hands now. To avoid this, we could target SUPER APE to specific tissues, such as muscles and blood cells.

There are many tissue-specific markers, or transmembrane proteins, that are expressed in certain cell types. Therefore, we can use viruses that infect cells expressing muscle and blood markers to put our SUPER APE gene in muscle and blood cells. As a disclaimer, this would only be possible if SUPER APE were a small gene (< 10 kb) using current technology. But let's assume that you are way smarter than every scientist in the world and move on. Super apes here we come!

Oops! When we put the SUPER APE gene into blood and muscle cells, it integrates into the genome and is immediately silenced by DNA methylation and tight histone packing. We need to deliver the SUPER APE gene as a protein into blood and muscle cells. Therefore, we need to make SUPER APE protein in bacteria and deliver it to our apes however, we cannot use the same plasmid that we were using before.

"Why not?" you ask. We were using the SUPER APE gene, which includes introns, and if we are making protein, we do not want intron sequences, because bacteria do not know how to splice out introns. You will have a nonfunctional SUPER APE protein.

Instead of the SUPER APE gene, we want to put the cDNA (complementary DNA) of SUPER APE into a plasmid, which will be much smaller than the gene. A cDNA is a DNA made from an RNA using reverse transcriptase from retroviruses.

While every other polymerase you have heard of either makes DNA from a DNA template (DNA polymerase) or RNA from DNA (RNA polymerase), retroviruses can make DNA from an RNA template, though there are viruses that make RNA from RNA templates. Reverse transcriptase is the enzyme that makes DNA from RNA, and we can purify SUPER APE mRNA and add reverse transcriptase to make cDNA.

This cDNA is inserted into a plasmid that has a promoter to drive expression in bacteria. We can then make lots of super ape protein in bacteria. How we get it into muscle and blood cells is a trickier problem, which we will leave to you to solve (and is probably another reason why we do not already have super apes).

One possible way to deliver super ape proteins to blood and muscle cells would be through ingesting food that makes super ape protein. Therefore, we could have plants make super ape protein and feed those plants to our apes.

The most trusted way of delivering genes to plants for biotechnology is using Agrobacterium tumerfaciens. This bacteria infects plants and causes tumors, collectively known as crown-gall disease. Agrobacterium transfers a portion of its tumor inducing (Ti) plasmid DNA, called T-DNA,which causes the plant to make opines, or compounds that the bacterium uses for energy and carbon sources.

Scientists have used this genetic transfer as a tool for gene delivery into plants, much in the same way the super ape protein can be made in bacteria. Putting the SUPER APE cDNA into the Ti plasmid could allow plants to make the super ape protein. Our apes can eat the super-ape-expressing plants, making them super strong for our invincible army. The bonus is that if they get out of control, we stop feeding them super ape protein plants. Hopefully, they do not learn how to grow their own super ape plants. Then, it would be curtains for us all.

Can We Stop Talking About Super Apes?

Much of what we previously described are techniques that real scientists use to actually make proteins and deliver genes for numerous biotechnology applications. What are some real, non-super-ape applications of recombinant DNA technology?

There are so many applications for recombinant DNA technology: medical applications, including gene therapy and vaccines the creation of genetically modified crops that are resistant to pesticides or that make extra vitamins and minerals and bacteria that can clean oil spills and even make "fake snow." While the applications of recombinant DNA technology are numerous, its limitations are its potential effects on our ecosystem. A longer discussion of this can be found in the "Biotechnology" section.

Brain Snack

Scientists use protein-making bacteria that improve the ability of water to form ice crystals. Here is a video of how scientists found those bacteria.


How To: Recombinant Protein Construct Design

Creating recombinant proteins has become much easier over the past few decades. However, those with the skills to do design such constructs are usually limited to the biochemists who study proteins. With the increase of protein therapeutics on the market for the treatment of cancer and other diseases, construct design has become a more relevant skill in the drug discovery field. Even if you are not directly responsible for making these constructs, knowing how they are created could give you important insight into how they work and why they were made. Thankfully the actual cloning process has been optimized to be faster than ever, with DNA synthesis becoming an even more attractive way to get the actual recombinant DNA. The design process can remain a challenge, but these few simple guidelines can help.


The first, and maybe most obvious step of construct design is getting ahold of your protein&rsquos template DNA. This can usually be found easily on an online genomic database. The next step should be to run an alignment with your protein&rsquos amino acid sequence against one or more organisms. The alignment will give you a readout of matching sequences and their percent homology with the original input sequence. This information can help find proteins of the same family, and any homologs that might be present in other organisms. Using this data, you can now begin to decipher your own protein sequence. The basic rules are conserved sequences between proteins usually denote a critical region of the protein important for structure or function. Unique sequences tend to impart a unique function or characteristic or denote a region of the protein that has no critical function. Conserved sequences are usually shared across homologs, or families of proteins and unique sequences, particularly within otherwise conserved regions, and are different between proteins and a consequence of an organism&rsquos evolution in a particular environment. This information can also help you find previous or current research related to your project. If your protein has little to no previous work done on it, it can be hard to find a starting point. Often experiment design and optimization can take months or even years before they can produce consistent data. This background research is critical if you want to keep your project on track and on time.


Sometimes it can be enticing to try everything under the sun and create dozens of construct designs when starting a project. Simplicity, however, is a key factor in designing a good construct. There are many mutations you can make to your protein: from point mutations, to truncations, to complex combinations of protein domains connected by linkers. Too many mutations too quickly can give confusing results, however. A good rule is to keep the construct as close to the native sequence as possible. The addition of a tag is usually required in order to facilitate purification, but other than that, try and limit each construct to one mutation until you have data suggesting you should make more complex mutants. Mutations should also be chosen very carefully as it can take weeks or months to generate and test each set of mutants. Make a note of the charge, size, and relative location of the native residue, and decide which mutant to make based on your hypothesis and project goals. Run a quick check and compare your potential mutants to past mutants from other papers, and make sure each has a specific and expected outcome. his is often the hardest part of the cloning stage, so once it&rsquos done it&rsquos just a matter of doing the leg work in lab to finish up.


Designing your first construct can be an exciting opportunity or even just an interesting exercise of your skills and problem-solving. It&rsquos important to note that even if your cloning succeeds, the resulting construct may not readily purify due to any number of solubility or stability issues. Many proteins can easily be undone by a single mutation, resulting in insoluble protein, non-native oligomeric states, or very little to no yield in the expression system used. The overall challenge tends to be finding the right mutations that can test your hypothesis and produce usable protein for your experiments. I hope this short explanation was enough to help you understand construct design, and maybe even try it out for yourself.


Watch the video: How recombinant DNA technology is used to create HPV vaccine (May 2022).