[Lecture Notes] Fundamentals of Biotechnology - L3: Recombinant DNA Technology

Lab techniques

DNA isolation and purification

DNA isolation from bacteria is easy because there is little more to a bacterium’s structure other than cell membrane and cell wall. The size difference between genomic DNA and plasmid DNA allows them to be separated.

Lysozyme digests the peptidoglycan which makes up the cell wall of a bacterium. Detergent such as SDS disrupts the phospholipid-bilayer and causes the cell to lyse. This cannot be dome for animal and plant cells which instead need to be ground up to release their contents. Plant cells can be put into a blender to break their thick cell walls made up of lignin and cellulose. An enzyme can then be added to break down these cell wall components into monomers. Mouse tail cells can have their DNA extracted by using proteinase K and SDS to degrade the tissue and dissolve the cell membranes. Cell culture dishes can be disrupted using only detergent as there is no structure outside the cell membrane.

Centrifugation allows one to separate insoluble components from soluble ones - chemical extraction can also be used. After cell lysis for example, the relatively insoluble DNA forms a pellet at the bottom of the tube whereas the cell wall components remain in solution. Phenol can also be used to remove proteins from DNA, this is because phenol is especially good at dissolving proteins and because it’s not very soluble in water, a phase separation occurs where the dissolved protein remains in the phenol phase, while nucleic acids remain in the aqueous phase.

Protein removal does not do anything about the RNA which is still mixed with the DNA. Ribonuclease digests RNA into its contituent ribonucleotides but does not digest DNA. If concentrated ethanol is added in equal parts with the water, long DNA (which should be genomic) leaves the aqueous phase and enters the ethanol, this DNA can then be isolated by centrifugation. The small RNA fragments remain dissolved in water.

Electrophoresis

Gel electrophoresis separates DNA fragments by their size. Agarose powder which is isolated from seaweed is dissolved in hot water and forms a gel when cooled. A rectangular piece of agarose gel can have a comb inserted into it while it cools which makes small wells in the gel. DNA is negatively charged, and when loaded into these wells (along with ethidium bromide to visualise the DNA), and a potential difference applied across the gel, the DNA migrates away from the negative electrode (located at the wells) and downards towards the positive electrode. Agarose acts like a sieve with small holes and the smaller the DNA fragment the faster it will pass through this gel. Ethidium bromide intercalates between DNA bases, it has a higher affinity for DNA than RNA because it is double stranded. Under UV light the DNA-ethidium bromide complex fluoresces orange.

Molecular weight standards or “Ladders” can be run in a single well to determine the molecular weight of each fragment. For small DNA fragments of 50-1000 base pairs one needs to use polyacrylamide gel to separate them out. This gel can resolve DNA fragments which differ by only a single base pair and are critical for sangar sequencing. For giant DNA fragments 10,000-10,000,000 base pairs, pulsed field gel electrophoresis (PFGE) is used. Two angles of electric current are used in this scenario.

Restriction enzymes and DNA ligase

Restriction enzymes (restriction endonucleases) cut DNA into fragments. These are bacterial enzymes that cut DNA at specific recognition sites. They are sensitive to methylation and therefore do cut up the DNA inside their host cells. Type I restriction enzymes cut the DNA 1000 or more base pairs away from the recognition site and type II restriction enzymes cut in the middle of the recognition site itself. They can cut either both strands in the same point, or each strand at a different point. This leaves blunt ends and sticky ends respectively. These type II restriction enzymes recognise inverted repeats so that the same base is cut on both strands. Restriction enzymes recognise a specific nucleic sequence and therefore can be used to compare the genomes of different organisms.

The probability of a cut can be modulated by the number of base pairs in a recognition sequence. For example a specific sequence of 4 base pairs is much more likely to be found than a specific sequence of 5 base pairs. To generate fewer and longer fragments, restriction enzymes which recognise 6-bp sequences are used, and for shorter fragments you can use restriction enzymes that recognise 4-bp sequences.

Different DNA cut with the same restriction enzyme will have the same sticky ends, this means that DNA from two different organisms can be ligated together using DNA ligase. The most common DNA ligase is from a virus: the T4 bacteriophage.

Detecting nucleic acids

One can detect the amount of DNA in solution by measuring the absorption of light at 260nm. The looser the structure of nucleic acid chains the more light is absorbed. So RNA absorbs more UV light than DNA. A second absorbance reading at 280nm is used to determine the purity of DNA. It shows the DNA/RNA ratio. Pure DNA has a 260/280 ratio of 1.8, whereas pure RNA has a ratio of 2. This is the basis of the Nanodrop which measures DNA concentration and purity.

DNA can also be radioactively marked and seen with photographic film. Autoradiography identifies the DNA in a gel by placing it under photographic film which will reveal the location of the DNA.

The same can be done but with fluorescent tags instead of radioactive markers. Different tags can bind to different nucleotides and this is the basis of modern sequencing.

DNA can be heated to melt the two strands apart, and then the two strands will reanneal upon cooling. This is because the heat breaks the hydrogen bonds which are quite weak. The GC content of DNA determines how much heat is required to fully melt the DNA as the CG base pair has 3 hydrogen bonds which are stronger than the AT base pair and so requires more heat.

Southern and northern blots

Southern blots are used to determine how closely related two single stranded DNA molecules are from one another if they have come from different sources. Related DNA molecules can anneal to one another in a process called hybridisation. A probe sequence (which is a gene from one organism) is isolated. Another sequence called the target sequence (which is DNA from antoher organism) is digested with a restiction enzyme to produce fragments 500-10,000 bp long. It is run under gel electrophoresis under strong acidic conditions so that the DNA separates into single stranded DNA. This DNA is then transferred to a membrane where it remains single stranded. The probe is then prepared: this sequence must be known and is then prepared with radioactive markers, biotin or digoxigenin. It is then denatured to make it single stranded. The single stranded probe is then incubated with the target DNA and hybrid double stranded DNA then forms. The mismatch amount between the probe and the target can be modulated by changing the temperature (at high temperature there is a low tolerance for mismatch). Photographic film then shows locations where the target and probe have annealed.

Northern blots are based on the same principle as the southern blot but RNA is used instead of DNA. mRNA is typically used in northern blots and with greater success as introns typically interfere with probe binding in southern blots. Moreover strong acid is not needed as RNA is already single stranded.

In situ hybridisation

Fluorescence in situ hybridisation (FISH) can be used to probe DNA or RNA directly within a chromosome. A small strand of DNA can be fluorescently tagged (the probe). The target DNA is the DNA already within the cell. When injected into the cell the fluorescent DNA probe anneals to the in situ DNA, the chromosome can be imaged and a specific gene from the probe DNA can be located physically on the chromosome.

Cloning vectors

Plasmids can take on any piece of foreign DNA for study, and can be used as cloning vectors. Other DNA elements such as viruses and synthetic chromosomes can also be used. Cloning vectors should be small in order to be easy to manipulate. They should be easy to move from cell to cell, easy to isolate, easy to detect and select for, and should have the ability to be copied multiple times to obtain lots of DNA. They should have clustered restriction sites and a way to determine if the DNA has been inserted.

Antibiotic resistance can be found on plasmids meaning that once a gene has been inserted into a plasmid, and that plasmid has been inserted into a bacterial colony, one can screen for successful transformations by growing the bacteria on a plate with that antibiotic on it. Cells which survive and grow successfully took up the plasmid. The 2u plasmid contains genes for synthesising essential amino acids so another screening technique can be to grow bacteria in media which doesn’t contain certain nutrients.

The higher the copy number of a plasmid (the more times it can be found in a cell) the easier it is to isolate that plasmid. The copy number should not be so high as to burden the cell during division. The origin of replication determines the DNA polymerase binding affinity and therefore determines the copy number.

The MCS (multiple cloning site) is an area on the vector with many restriction enzyme sites. This high density of restriction enzyme sites allows the plasmid to be cut in one location without disrupting other genes on the plasmid to do with replication. Insertional inactivation can be used to detect if a vector contains an insert, the vector may have two antibiotic resistance genes and the insert can be put into one these AB resistance genes, thus breaking it up and removing the resistance to that AB.

Another method is alpha complementation. the gene for lacZ-alpha (part of the lacZ) gene is on the plasmid. An insert is placed into this gene, breaking it up meaning that beta-galactosidase isn’t produced. This means that bactera cannot break down X-gal and therefore do not turn blue.

Most cloning vectors used are based on E. coli plasmids as this organism is studied so much. The MCS is placed between the promoter and terminator - some vectors also include a ribosome binding site so that inserted genes get made into protein. The 2u circle of yeast has been used a cloning vector. A shuttle vector based on the 2u circle has an origin of replication for two organisms and genes to survive in two organisms. Therefore 2u shuttle vectors can survive in yeast and bacteria. Selecting for yeast which have taken up the shuttle vector is done using a gene that codes for leucine production, so that one can grow a yeast culture in leucine deficient media and have a pure sample of 2u shuttle vector incorporated yeast.

Bacteriophage vectors are modified viruses that can deliver non-viral DNA into a host organism. DNA within the viral particle is kept linear using a protein coat but when injected the DNA circularises and becomes the replicative form (RF). The lambda bacteriophage can be modified to be used as a cloning vector, it can take an insert of 37-52kb in length. Genes necessary for virus packaging creation are deleted so that the virus doesn’t kill the entire colony when it infects it. To create fully formed bacteriophages, helper virses are used to supply the genes for the coat proteins which allow for the assembly of fully formed bacteriophages as the lysates of the helper virus are mixed with the recombinant DNA to be put into the lambda virus. We call this in-vitro packaging

Artificial chromosomes can be yeast artificial chromosomes or bacterial artificial chromosomes or bacteriophage artificial chromosomes. They can contain DNA up to 2000kb (YAC are the biggest).

Transformation

Competant E. coli cells are mixed with loning vectors in a process called transformation. The cell wall is opened up by mixing the cells with calcium ions on ice and then shocking them at high temperatures - this opens up the membrane to allow the cells to take in the plasmid. Most cells die during this process but a few will take up the plasmid. Another method uses high voltages shocks instead of heat shock to disrupt the cell membrane. This is called electroporation and is more versatile and faster and can be used with both bacteria and yeast.

A bacterium can have multiple plasmids within it but they must have different origins of replication, otherwise when the bacteria divides it will lose one of the plasmids because only one will be copied. This is called plasmid incompatibility.

Gene libraries

A gene library is created when the entire genome of an organism is digested into fragments and inserted into a cloning vector and transformed into a host. The length of sequence recognised by a restriction enzyme will affect the size of the library and the length of the inserts into the plasmids. We want to find a gene within our DNA library. The DNA library can be screened by hybridisation. Gene libraries are stored as E. coli cell cultures each with a unique plasmid. These cells are grown up on agar plates (so that they are spaced apart) and then lysed to release all their DNA (using detergent). The probe is synthesised and labelled and mixed with the library of DNA fragments created from the lysed cells - this probe will hybridise with matching DNA. The location where the DNA hybridises is matched up with its corresponding cell culture (which in theory should have grown from a single cell) and this cell can then be rescreened to ensure that a single transformant was isolated.

Eukaryotic expression libraries are typically made from complementary DNA rather than normal DNA to avoid introns. Complementary DNA is just the DNA copy of mRNA and is made using reverse transcriptase. To make this cDNA library mRNA is first isolated from a host organism by running it through a column containing poly(T), therefore mRNA will anneal to the poly(T) column as it always has its poly(A) tail attached. This mRNA is purified and using reverse transcriptase converted into cDNA. These cDNAs are ligated into expression vectors with transcription and translation initiation sequences. Due to DNA being translated using the triplet code, if a cDNA does not have its own transcription and translation start site, there are three possible open reading frames that can be read, but only one will produce the correct protein. Therefore all three ORFs are cloned which significantly increases the number of transformations needed. The cloned genes are then transformed into bacteria which express the DNA. They are again transferred to a nylon membrane and subsequently lysed. Proteins that were expressed then attach to the nylon membrane and are screened. Usually we are interested in an antibody and we can screen using a secondary anti-body that conjgates with the first with a detection system that either fluoresces or leaves a colour on the plate. Even though prokaryotes do not perform post-translational modification to the proteins, secondary antibodies can still identify proteins of itnerest.

Protein production needs to be modulated so that not too much foreign protein is made which ends up killing the host cell. Expression vectors have promoters which can have their on/off switches controlled. Therefore bacteria can be grown to a high enough cell density and the promoter can then be switched on to begin protiein synthesis. An example is lacUV, a mutant version of the lac promoter. The lacI repressor is also present downstream, this means that the gene is not expressed due to the lacI repressor. When IPTG is added however the lacI is released and RNA polymerase can transcribe the cloned gene.

Another promoter is the lambda left promoter which has a binding site at the lambda repressor, this gene is not expressed unless the repressor is removed and a mutant version has bee created so that the repressor falls off at 42 degrees, initiating transcription. Certain tags can also be detect expressed proteins if the protein of interest has already been cloned.

Recombineering

Recombineering is the process of creating large recombinant DNA vectors. This is useful when we have long genes we want to clone, it is difficult to find compatible restriction enzymes that do not cut the gene itself up with the polylinker. We can engineer RED enzymes from the lambda phage to be expressed by a host bacteria, the proteins can recognise a 45 base pair region of homology with the BAC to initiate recombination. The gene is electroporated into the bacteria expressing the RED proteins within the cytoplasm. The homologous sections to the gene are found on the BAC, and the enzymes splice the gene into the BAC. See figure 3.25 in the book for more information. One needs different screening techniques to identify if the recombineering was successful, because the bacteria already has the RED vector before the recombineering began. Instead selection/counterselection is used. The initial vector used contains a galK gene, which codes for galactose kinase which allows the bacteria to grow in galactose media. GalK can also convert 2-deoxygalactose kinase into a toxic substance so if grown on 2-DOG media they die. The galK gene is replaced during recombineering and this means that successful recombinants will not die when grown on 2-DOG media.

Proprietary GATEWAY(R) cloning vectors

The lambda bacteriophate phage DNA can integrate into the E. coli chromosome at the attB site to form a prophate. The phate DNA has an attB site and an enzyme called integrase makes staggered cuts on the centre of both attB and attP sites, the ends then connect and the prophage is formed. After integration the attB and attP sites cease to exist as the cuts were made at the centre of each site. Each site is now half an attP and half an attB. We call these attL and attR sites now. The reaction can be reverse but needs a different enzyme called excisionase. GATEWAY needs to get the gene of interest inbetween two attL sites. This can be done by first cloning the gene into an MCS in an “entry clone”. The entry clone has a gene called ccdB in the MCS which odes for a toxin that kills the host. When a gene is spliced into the ccdB gene, E. coli are able to grow. (The entry clone is created using E.coli which contain another vector with the antitoxin to ccdB.) Once the entry vector has the gene of interest in it, it is moved around by two reactions: the LR reaction positions the gene between an attR1 and attR2 site, so the gene is flanked by these two sites. Now ccdB is able to be expressed again and the bacteria dies. The BP reaction removes the gene from teh vector and puts it into a vector with attL sites. See figure 3.29 for a better explanation than this.

References: my notes are made from, and follow the structure of my course textbook which is Biotechnology 2nd edition by David P. Clark, which can be found for purchase here.