Recombinant Proteins Flashcards
List the key features needed to express a protein from a vector in a bacteria. Why are each of these features necessary for expression?
Vectors require a bacterial promoter and terminator sequence flanking the protein-coding DNA, in order to recruit the RNA Pol for transcription and for transcription of the correct part of the vector. It also requires a ribosome binding site that is specific to the bacteria included at the start of the gene for translation.
Describe a biological switch, using the features of the trp operon, that is often used to regulate expression in bacteria. How does it help to prevent inclusion bodies?
Inducible promoters are used when expressing proteins in bacteria because then the researchers can control whether the transcription is on or off at any point, using biological switches. In the instance of the trp operon, this is a hybrid of the lac and the trc operons because lac is a weak promoter. It uses the trp promoter, which binds the RNA Pol, but still has a lac operator that binds lac repressor. The switch for this operon is the molecule IPTG - this binds to lac repressors to inhibit their binding to DNA. By controlling the IPTG concentration, you can control the level of expression.
Inclusion bodies are formed when the concentration of a foreign protein becomes too high, due to high expression, and forces all the proteins into precipitate into inclusion bodies. This is because large amounts of proteins can be toxic to the bacteria because it will impact bacterial growth. Regulating the rate of transcription using inducible promoters can reduce this effect.
What are the advantages and disadvantages of expressing recombinant proteins in bacteria?
Advantages of expressing recombinant proteins in bacteria would be the relative ease of transformation of bacterial cells such as E.coli. They are cheap, quick and don’t need much space to reproduce in large quantities.
Disadvantages include the potential for inclusion bodies to form because of insoluble proteins, which can mean the proteins are not properly folded. This can also occur because bacteria lack chaperones and machinery to properly fold heterologous proteins. There are different mechanisms for post translational modifications in different species, e.g. bacterial cells cannot glycosylate proteins as it does not have the machinery to.
What are the advantages and disadvantages of expressing recombinant proteins in mammalian cells?
The advantages of expressing recombinant proteins in mammalian cells is that because they are eukaryotic, the intracellular machinery is likely to be similar and more complex than in bacteria, and therefore can carry out proper folding and PTMs. Proteins can also have sub-cellular targeting that guides them towards particular organelles. The proteins also are more biologically active within the cells, exhibiting their normal behaviour.
The disadvantages to using mammalian cells is that it is very expensive - the nutrients, space and time needed to produce large amounts of proteins is much larger than with bacteria.
What is the difference between transient and stable transfection? Describe two methods of transfection that can be used to insert plasmids into mammalian cells.
Transient transfection means that the plasmid vector does not integrate into the genome and therefore, stays in the cell for about 2-3 days. Stable transfection is where the DNA does integrate into the genome and is hugely beneficial for the pharmaceutical industry. This means that recombinant vectors don’t need to keep being produced to make the recombinant proteins - all you need to be is culture these cells that have been transfected.
Electroporation is a method of using a small electric shock, increasing the permeability of the membrane, in order to allow the plasmid to pass through. Lipofection is also a method of transfection, in which the DNA is covered in a lipid treatment and therefore, can pass through the phospholipid membrane.
What are the key features needed on a donor plasmid for transposition of a cloned DNA into a bacmid?
A donor plasmid needs different features for transposition because it uses a transposon-technique to transfer the DNA insert from the plasmid into the baculovirus genome (bacmid).
The DNA insert contains a polyhedrin promoter, which is needed for expression inside insect cells. The insert is flanked by transposon sequences, e.g. Tn7L and Tn7R (for right and left). This means that everything between these two sequences is transposed into the bacmid.
What are the key features of a bacmid that allows transposition to occur?
Bacmid (baculovirus DNA) contains a lacz gene (gene for beta-galactosidase), which contains within that a site called the mini att Tn7 site. This site is where the transposon will insert into. There is also a helper plasmid present in the cell, which contains all the proteins required for the transposition. All of this is done inside an E.coli cell in order to protect the bacmid from damage.
What key steps are needed in order to prepare the baculovirus so that it will produce the recombinant protein in insect cells?
To produce the baculoviruses in preparation for infection of insect cells, first the recombinant bacmid must be produced. This is done by transposition of the desired DNA insert from a donor plasmid into a bacmid inside of E. coli cells. Once the recombinant bacmid is produced, it is then isolated and used to infect insect cells. The cells will eventually lyse and release the baculoviruses. Insect cells are then infected again, but this time with the actual baculoviruses. The bacmid inserts itself into the insect cell genome and insect cell machinery is used to transcribe it, including the recombinant protein.
Describe the contributions of recombinant protein production technology to society.
Recombinant protein production is becoming a massive industry, especially in pharmaceuticals. It can be used to make proteins without the need for animal sources, e.g. insulin is now made by recombinant bacteria. These proteins are considered safer and with fewer off-target side effects, because they can be catered towards safety. It has opened the door for protein therapies e.g. protein replacement therapies for those who have dysfunctional proteins leading to disease. The cells themselves are also being used as vaccines for delivery of pathogenic proteins into bodies in order to stimulate an immune response.
However, there is still issues with the cost of production, as well as the delivery of the treatments in terms of protein therapies.
What is the principle of site-directed mutagenesis? Explain the main steps of site-directed mutagenesis using recombinant plasmids.
Site-directed mutagenesis is where a mutation is introduced into the base sequence of DNA to study whether this has an effect on the protein produced from the gene, or to study whether the mutation affects protein-DNA interactions.
The process of site directed mutagenesis requires: DNA template, DNA polymerase, dNTPs and primers. The plasmid is formed, containing the gene to be mutated, and isolated out of E.coli. It is important that these plasmids were isolated from E.coli that is able to methylated DNA. The plasmids should be methylated on both strands at the two Dpn1 sites - Dpn1 is an endonuclease that cleaves a blunt end at its recognition site.
The primer used to initiate DNA replication must contain the mutation that you want to introduce into the gene. The primer is usually a 25mer oligonucleotide, and so will still bind to the gene despite the mutation.
The plasmids are denatured to form the two mother template strands. The primers anneal to the complementary sequence of the plasmid. The DNA Pol will work to add dNTPs to form the new copy strand. This will form two hemimethylated plasmids. The sample is then heated so that the plasmids denature again. This allows the single strands to bind to different strands. The aim is for the two mutated strands to anneal and produce a mutated unmethylated plasmid. There will still be some hemimethylated and fully methylated plamids that have anneal, but these are degraded because Dpn1 digests methylated and hemimethylated DNA. The mutated plasmid is then isolated, transformed into bacteria, and can replicate. This is then used for further experimentation.
How can insertions and deletions be introduced through this site-directed mutagenesis method?
Insertions can be introduced using primers that have complementary sequences at its ends, and a new sequence in the centre. When it anneals, the new sequence part will form a loop structure while the complementary sequence hybridises to the plasmid. Once the whole new strand is encoded, heated to separate the strands, and hybridises to another mutated strand, it will incorporate the insertion into the sequence.
Deletions work in the opposite manner. The primer will contain a sequence that has part of it deleted in the centre. When it anneals to the plasmid, the plasmid DNA will form a looped structure in order to bind to the primer complementarily.
What is meant by a GFP fusion protein?
GFP fusion proteins include the protein that is being studied with a GFP protein attached to either its N or C terminus. When it is encoded, the GFP gene is added to the end of the desired protein gene, so that when it is transcribed and translated, it produces this fusion (or chimeric) protein.
How did GFP help establish that glut4 moves through a fat cell when stimulated by insulin?
Fluorescence tagging can be used for in situ visualisation of proteins. Fat cells were analysed by using a fluorescent GLUT4 protein and studying the movement of GLUT4:GFP in the cells when insulin was introduced. When there is no insulin, the GLUT4 are seen in the centre of the cell, where they are stored in intracellular storage units. When insulin is introduced, the GLUT4 can be seen to all move to the surface membrane of the cell. This is because GLUT4 is a transporter for glucose and so insulin stimulates them to start transporting glucose into the cells for metabolism.
Why was it necessary to mutate the wild-type GFP into the enhanced GFP? How was this enhanced GFP produced?
Wild-type GFP was found in jellyfish that lives at an optimal temperature of about -28C, where GFP gave off its maximum fluorescence, whereas mammalian cells have an optimal of 37C. Therefore, site-directed mutagenesis was needed to optimise the fluorescence of GFP at the temperature it would be used at (~37C).
This enhanced GFP was produced by creating a mutant GFP library, where single mutations were introduced one at a time to every residue between 55-74 to see which mutations had what effect. This was done using many primers, each with one mutation. This then was turned into a clone library as the plasmids were transformed into E. coli.
Why was it necessary to mutate the wild-type GFP into the enhanced GFP? How was this enhanced GFP produced?
Wild-type GFP was found in jellyfish that lives at an optimal temperature of about -28C, where GFP gave off its maximum fluorescence, whereas mammalian cells have an optimal of 37C. Therefore, site-directed mutagenesis was needed to optimise the fluorescence of GFP at the temperature it would be used at (~37C).
This enhanced GFP was produced by creating a mutant GFP library, where single mutations were introduced one at a time to every residue between 55-74 to see which mutations had what effect. This was done using many primers, each with one mutation. This then was turned into a clone library as the plasmids were transformed into E. coli. The plasmids were expressed within these bacterial cells. The cells underwent fluorescence cell sorting (FACS), which separated the cells out by the level of fluorescence. The highest fluorescence cells had their plasmids sequenced and these mutated GFP were used in research as their fluorescence was ~30 fold higher than wild-type GFP.
