In earlier posts I've talked about how to replicate and amplify DNA sequences so that you can make shitloads of proteins. Well, I've realised that I haven't actually spoken about how to extract and purify all of those proteins that you just made (or bullied a cell into making for you).
A lot of these purification processes actually require that you've made a purifiable protein to begin with. For example, with a His tag, you need to add some DNA coding for a bunch of histidine residues to the beginning or end of the sequence. I've actually spoken about His tags in an earlier post: Introduction to Cloning. Essentially the proteins are washed down a column, and the ones with His tags bind to nickel beads in the column. Eventually all that's left in the column are the His-tagged proteins bound to metal beads. These can then be eluted from the column by lowering the pH or by adding extra histidine or imidazole.
A second type of purification process is called GST-Fusion, which of course has absolutely nothing to do with Goods and Services Taxes. GST stands for glutathione-S-transferase. DNA coding for glutathione-S-transferase is positioned right before the DNA for the protein of interest, so that the protein is attached to glutathione-S-transferase. Glutathione-S-transferase attaches to glutathione (a.k.a. GSH), rather than nickel beads. It can be eluted with free glutathione. The GST part can then be removed by cleavage with Xa protease at a site which, like the GST enzyme itself, has also been engineered into the protein.
Saturday, March 19, 2016
DNA Sequencing and Synthesis
Last post on recombinant DNA technology for this unit! (That went quickly...)
Understand the principle of dideoxy sequencing.
Dideoxy sequencing, also known as the Sanger method after the guy who invented it, is a pretty reliable technique for sequencing DNA.
In traditional dideoxy sequencing, you need the following materials:
Understand how PCR can be used in the dideoxy sequencing reaction and how the sequencing is automated.
PCR can be used in the first part of the sequencing reaction (i.e. the bit with the synthesising of fragments ending in dideoxynucleotides) in a technique known as "cycle sequencing," or "linear amplification sequencing." In this process, only one strand is primed so that a linear (rather than exponential) amplification of products is achieved- hence "linear amplification sequencing."
In automated sequencing, the process is carried out in a single tube (rather than four tubes for the four different dideoxynucleotides). This is achieved by fluorescently labelling the ddNTPs (sorry, got sick of typing out "dideoxynucleotides") with different colours. After running the fragments through electrophoresis (only one lane required this time), a machine can detect the different wavelengths of emitted light to give a reading of the sequence.
Be familiar with new and rapid “next generation sequencing”.
Nowadays, there are faster and cheaper methods of sequencing DNA, though they are not necessarily as accurate as the dideoxy/Sanger method outlined above. Most forms of "next generation sequencing" involve creating billions of tiny fragments of DNA (30-70 base pairs long) which are immobilised to beads or chips. This immobilisation is done by adding an oligo-dA tail to the fragments, which bind to oligo-dT molecules in the beads or chips.
Fluorescently labelled dNTPs (not ddNTPs) are used for "next generation sequencing." They will bind to oligo-dT anchors if the fragment next to it has a complementary base. This will then cause that strand to fluoresce. The fluorescent tag is then removed, and a different fluorescently labelled dNTP is added. The fluorescence can be picked up by a computer to sequence the DNA.
Be familiar with how DNA can be chemically synthesized and the uses of synthetic DNA.
One thing you might have wondered throughout all of this is how primers and so forth are synthesised. One method of synthesising oligonucleotides is the phosphoramidite procedure, which synthesises oligonucleotides from 3' to 5' (opposite of the normal direction).
The phosphoramidite procedure starts by anchoring a nucleoside to a glass support at the 3' end. This nucleoside also has a blocking group (usually dimethyoxytrityl, a.k.a. DMTr) attached to the 5' end to prevent spontaneous reaction. Throughout this procedure, most of the bases added also have blocking groups added to them until they are washed off at the end.
The first step of the main part of the procedure is washing off the DMTr group by using trichloroacetic acid. Next the second base is added, except it's not added in the form of a nucleotide: it's added in the form of a nucleoside phosphoramidite derivative. This nucleoside phosphoramidite derivative is essentially just a nucleoside with a DMTr blocking group attached to the 5' end and a phosphoramidite group attached to the 3' end. Phosphoramidite groups centre around a trivalent phosphorus (i.e. a phosphorus atom that only forms 3 bonds rather than 5 like in phosphate). They also have other crap attached to them but I'm not going to go into that because it's not important. What's important is that this phosphoramidite group can be activated by a weak acid, such as tetrazole, allowing it to rapidly react with the 5' end of the first nucleotide. Aqueous iodine is then added to form a stable phosphate group. These steps are then repeated until the desired chain length is reached.
At the end of the process, the oligonucleotide is cleaved from the support by using ammonium hydroxide, which also removes blocking groups from the bases.
We're now done for this topic! Whew! (I might make a short mention on His tagging and GST-Fusion tagging in a later post, since that was covered in an earlier lecture but not in my posts here.)
Understand the principle of dideoxy sequencing.
Dideoxy sequencing, also known as the Sanger method after the guy who invented it, is a pretty reliable technique for sequencing DNA.
In traditional dideoxy sequencing, you need the following materials:
- Template strands for the DNA that you want to sequence
- A primer
- DNA Polymerase I
- All four nucleotides- at least one of these should be attached to a radioactive phosphate
- Dideoxynucleotides. These are nucleotides with two -H groups as opposed to -OH groups (hence di- and deoxy-). The other H group is on the 3' carbon. Yup, that means that there's no 3'-OH, which means that new nucleotides cannot be added after the addition of a dideoxynucleotide. This is pretty important, as we're about to see.
Four reactions need to be carried out, each with a different dideoxynucleotide (i.e. one reaction needs to be carried out with dideoxyadenosine, one with dideoxythymidine etc.). Essentially, in each reaction, DNA Polymerase I is used to synthesise a new strand, but it will terminate whenever a dideoxynucleotide is added. (The concentration of dideoxynucleotides is kept reasonably low so that the chain won't always stop at the first incidence of whatever base you're looking at.)
The fragments created from the previous step are then run through an electrophoresis process. Fragments from each reaction are run through different lanes so that you can keep track of which fragments end with dideoxyadenosine, which fragments end with dideoxycytidine etc. The gel used is usually 4-6% polyacrylamide and 6M urea (the urea is used in order to denature the DNA so that its secondary structure won't interfere with electrophoresis). Like in normal electrophoresis, smaller fragments migrate through the gel faster. Thus by reading "up" the gel you can see what the sequence of bases are.
Understand how PCR can be used in the dideoxy sequencing reaction and how the sequencing is automated.
PCR can be used in the first part of the sequencing reaction (i.e. the bit with the synthesising of fragments ending in dideoxynucleotides) in a technique known as "cycle sequencing," or "linear amplification sequencing." In this process, only one strand is primed so that a linear (rather than exponential) amplification of products is achieved- hence "linear amplification sequencing."
In automated sequencing, the process is carried out in a single tube (rather than four tubes for the four different dideoxynucleotides). This is achieved by fluorescently labelling the ddNTPs (sorry, got sick of typing out "dideoxynucleotides") with different colours. After running the fragments through electrophoresis (only one lane required this time), a machine can detect the different wavelengths of emitted light to give a reading of the sequence.
Be familiar with new and rapid “next generation sequencing”.
Nowadays, there are faster and cheaper methods of sequencing DNA, though they are not necessarily as accurate as the dideoxy/Sanger method outlined above. Most forms of "next generation sequencing" involve creating billions of tiny fragments of DNA (30-70 base pairs long) which are immobilised to beads or chips. This immobilisation is done by adding an oligo-dA tail to the fragments, which bind to oligo-dT molecules in the beads or chips.
Fluorescently labelled dNTPs (not ddNTPs) are used for "next generation sequencing." They will bind to oligo-dT anchors if the fragment next to it has a complementary base. This will then cause that strand to fluoresce. The fluorescent tag is then removed, and a different fluorescently labelled dNTP is added. The fluorescence can be picked up by a computer to sequence the DNA.
Be familiar with how DNA can be chemically synthesized and the uses of synthetic DNA.
One thing you might have wondered throughout all of this is how primers and so forth are synthesised. One method of synthesising oligonucleotides is the phosphoramidite procedure, which synthesises oligonucleotides from 3' to 5' (opposite of the normal direction).
The phosphoramidite procedure starts by anchoring a nucleoside to a glass support at the 3' end. This nucleoside also has a blocking group (usually dimethyoxytrityl, a.k.a. DMTr) attached to the 5' end to prevent spontaneous reaction. Throughout this procedure, most of the bases added also have blocking groups added to them until they are washed off at the end.
The first step of the main part of the procedure is washing off the DMTr group by using trichloroacetic acid. Next the second base is added, except it's not added in the form of a nucleotide: it's added in the form of a nucleoside phosphoramidite derivative. This nucleoside phosphoramidite derivative is essentially just a nucleoside with a DMTr blocking group attached to the 5' end and a phosphoramidite group attached to the 3' end. Phosphoramidite groups centre around a trivalent phosphorus (i.e. a phosphorus atom that only forms 3 bonds rather than 5 like in phosphate). They also have other crap attached to them but I'm not going to go into that because it's not important. What's important is that this phosphoramidite group can be activated by a weak acid, such as tetrazole, allowing it to rapidly react with the 5' end of the first nucleotide. Aqueous iodine is then added to form a stable phosphate group. These steps are then repeated until the desired chain length is reached.
At the end of the process, the oligonucleotide is cleaved from the support by using ammonium hydroxide, which also removes blocking groups from the bases.
We're now done for this topic! Whew! (I might make a short mention on His tagging and GST-Fusion tagging in a later post, since that was covered in an earlier lecture but not in my posts here.)
Friday, March 18, 2016
Genetic Recombination and the Use of Recombinase in Cloning
I must admit that my friend and I spent part of the lecture playing Hangman, but hopefully the main details are fine :)
Be familiar with the ability of DNA ligase to join DNA molecules and how to overcome vector self-ligation.
Okay, you should be reasonably familiar with the idea that DNA ligase joins DNA molecules. If not, have another look at my post on enzymes involved in DNA replication and repair.
Now let's have a look at the idea of self-ligation. Self-ligation occurs when, instead of a fragment of DNA being inserted into a plasmid, the two cut ends of the plasmid simply snap back together (i.e. self-ligate). This can be easily prevented by using alkaline phosphatase to remove phosphate groups on the plasmids. Without these phosphate groups, the plasmid cannot self-ligate, but fragments can still be inserted. This does leave single-stranded nicks where the plasmid lacks phosphates, but the effect of this is negligible due to the size of the inserted fragment and the distance between the two nicks.
Be familiar with the types of site-specific recombination.
There are three main types of site-specific recombination: insertion, deletion or inversion. They're pretty much self-explanatory: insertion involves the insertion of a new DNA fragment, deletion involves its deletion, and inversion involves re-inserting it backwards (i.e. inverting it). These reactions are catalysed by recombinases, which are part of a larger family of enzymes known as integrases. There are two essential parts of the DNA itself that facilitate site-specific recombination: recognition sites for the recombinases, and a crossover region where cutting and rejoining occurs. The crossover region also confers directionality, which is important in the case of inversion.
Understand the mechanism of action of Cre recombinase.
Cre recombinase is a recombinase (duh) taken from Phage P1. Its original function is to circularise the phage genome when it infects bacteria. Cre recombinase recognises a certain site called the LoxP site. The LoxP site contains an 8 base pair core sequence which is not palindromic and therefore confers directionality. This 8 base pair core sequence is flanked by two palindromic 13 base pair sequences. (By palindromic, I mean that the one before the core sequence reads the same as the one after, but backwards.)
Cre recombinase is made up of four identical subunits, each of which has a tyrosine in the active site. This tyrosine breaks the linkage between the 5'-OH and the phosphate of a nucleotide, forming a covalent intermediate until another nucleotide comes along. There are other recombinases that have serine in the active site- these ones break the linkage between the 3'-OH and the phosphate of the next nucleotide. The formation of these intermediates eliminates the need for ATP or other energy sources.
Cre recombinase works by cleaving and rejoining two strands simultaneously. Subunits R1 and R2 hold onto one DNA molecule at the palindromic sequences while R3 and R4 hold onto the other (again, at the palindromic sequences).
One of the benefits of using Cre recombinase is that its recognition sequence, LoxP, doesn't occur naturally in plants or animals. Hence, engineering a LoxP site into the genome and then using Cre recombinase will not cause unwanted cuts. (Of course, there's probably a tiny chance that there could be the exact same site somewhere in the genome by chance, but that chance is really, really negligible at best.)
Be familiar with applications of Cre recombinase and LoxP sites in cloning and conditional gene targeting.
As I just mentioned, Cre recombinase and LoxP are good for "cutting and pasting" (similar to restriction enzymes) as LoxP sites don't occur naturally in most plants and animals. Hence they are good for modifying and moving around genes. LoxP sites can also be used for conditional gene targeting- the so-called "knocking out" of genes.
I'm now going attempt to explain conditional gene targeting, using mice as an example. To create mice that have a gene "knocked out," you first need to start with two kinds of transgenic mice. One has the gene for Cre recombinase tied to an inducible and tissue-specific promoter, whereas the other kind has loxP sites flanking one of the exons of the gene in question. Following breeding, some of the mice should have both characteristics: loxP sites and the gene for Cre recombinase. The gene that you are studying can then be "knocked out" by inducing the production of Cre recombinase. This allows more control over timing of gene expression, and also allows you to see the effects of genes that could have been lethal during embryonic development.
Be familiar with the ability of DNA ligase to join DNA molecules and how to overcome vector self-ligation.
Okay, you should be reasonably familiar with the idea that DNA ligase joins DNA molecules. If not, have another look at my post on enzymes involved in DNA replication and repair.
Now let's have a look at the idea of self-ligation. Self-ligation occurs when, instead of a fragment of DNA being inserted into a plasmid, the two cut ends of the plasmid simply snap back together (i.e. self-ligate). This can be easily prevented by using alkaline phosphatase to remove phosphate groups on the plasmids. Without these phosphate groups, the plasmid cannot self-ligate, but fragments can still be inserted. This does leave single-stranded nicks where the plasmid lacks phosphates, but the effect of this is negligible due to the size of the inserted fragment and the distance between the two nicks.
Be familiar with the types of site-specific recombination.
There are three main types of site-specific recombination: insertion, deletion or inversion. They're pretty much self-explanatory: insertion involves the insertion of a new DNA fragment, deletion involves its deletion, and inversion involves re-inserting it backwards (i.e. inverting it). These reactions are catalysed by recombinases, which are part of a larger family of enzymes known as integrases. There are two essential parts of the DNA itself that facilitate site-specific recombination: recognition sites for the recombinases, and a crossover region where cutting and rejoining occurs. The crossover region also confers directionality, which is important in the case of inversion.
Understand the mechanism of action of Cre recombinase.
Cre recombinase is a recombinase (duh) taken from Phage P1. Its original function is to circularise the phage genome when it infects bacteria. Cre recombinase recognises a certain site called the LoxP site. The LoxP site contains an 8 base pair core sequence which is not palindromic and therefore confers directionality. This 8 base pair core sequence is flanked by two palindromic 13 base pair sequences. (By palindromic, I mean that the one before the core sequence reads the same as the one after, but backwards.)
Cre recombinase is made up of four identical subunits, each of which has a tyrosine in the active site. This tyrosine breaks the linkage between the 5'-OH and the phosphate of a nucleotide, forming a covalent intermediate until another nucleotide comes along. There are other recombinases that have serine in the active site- these ones break the linkage between the 3'-OH and the phosphate of the next nucleotide. The formation of these intermediates eliminates the need for ATP or other energy sources.
Cre recombinase works by cleaving and rejoining two strands simultaneously. Subunits R1 and R2 hold onto one DNA molecule at the palindromic sequences while R3 and R4 hold onto the other (again, at the palindromic sequences).
One of the benefits of using Cre recombinase is that its recognition sequence, LoxP, doesn't occur naturally in plants or animals. Hence, engineering a LoxP site into the genome and then using Cre recombinase will not cause unwanted cuts. (Of course, there's probably a tiny chance that there could be the exact same site somewhere in the genome by chance, but that chance is really, really negligible at best.)
Be familiar with applications of Cre recombinase and LoxP sites in cloning and conditional gene targeting.
As I just mentioned, Cre recombinase and LoxP are good for "cutting and pasting" (similar to restriction enzymes) as LoxP sites don't occur naturally in most plants and animals. Hence they are good for modifying and moving around genes. LoxP sites can also be used for conditional gene targeting- the so-called "knocking out" of genes.
I'm now going attempt to explain conditional gene targeting, using mice as an example. To create mice that have a gene "knocked out," you first need to start with two kinds of transgenic mice. One has the gene for Cre recombinase tied to an inducible and tissue-specific promoter, whereas the other kind has loxP sites flanking one of the exons of the gene in question. Following breeding, some of the mice should have both characteristics: loxP sites and the gene for Cre recombinase. The gene that you are studying can then be "knocked out" by inducing the production of Cre recombinase. This allows more control over timing of gene expression, and also allows you to see the effects of genes that could have been lethal during embryonic development.
Cloning by PCR and Mutagenesis
Appreciate that there are different
approaches to the cloning of specific
genes.
A moment of silence here while we appreciate all the different approaches...
Okay, back to being serious. There's really just two main approaches (well, at least only two that we learned about during the lecture). They both start off with figuring out what protein you want to clone, determining some of the N-terminal amino acid sequence and from there taking a stab at what the potential DNA sequence would be (presumably taking into account that some amino acids can be coded for by more than one codon).
From there you can take one of two approaches, depending on what technology is available to you. Since several different genomes have already been sequenced, the easiest approach nowadays would generally be to search the database for the gene of interest. You might also be able to find similar genes coding for other proteins, and from there you might be able to get some kind of idea of what your protein does. To be a bit more certain, though, you can amplify the gene (and therefore protein production), allowing you to study the protein more and find out more about it. To amplify the gene, use the database to help you design some oligonucleotide primers so that you can use PCR (more on PCR in an earlier post). This amplified DNA can then be cloned into an expression vector (see the end of my most recent post) to create lots of protein.
An alternative approach can be used if you don't have access to a database for searching for genes. After working out a potential DNA sequence, you can make a DNA probe using this sequence data. This probe can then be used to find the gene of interest in a gene library (more on gene libraries in a bit). Once you've found your gene, you can clone it and so forth.
Be familiar with cDNA library construction and traditional screening by hybridisation.
cDNA libraries can be bought commercially, but just in case you wanted to make your own, I'm gonna tell you how (or rather, the theory behind it anyway). *Do not try this at home*
cDNA is essentially the complement of mRNA. To create cDNA, simply use the enzyme reverse transcriptase to transcribe the mRNA into DNA. Remove the mRNA with alkali and add a poly-G tail (this is so you can add a poly-C primer for synthesis of the other strand). Synthesise the other strand using DNA polymerase.
Next you have to do a couple more fancy things with the cDNA. Firstly, you have to methylate it so that restriction enzymes won't cut it where you don't want it to be cut. Secondly, you have to add EcoRI linkers to either side (essentially the "cutting sites" for EcoRI). Cleave them with EcoRI to form sticky ends.
Now for the recombination part! Take some bacteriophage lambda (a bacteriophage is essentially a virus that infects bacteria) and cleave its DNA with EcoRI. Ligate this to the cDNA. Now you can package this cDNA into bacteriophage lambda so that it can go ahead and infect EcoRI with some shiny new cDNA!
cDNA libraries can be used to screen by hybridisation of a probe, as alluded to in the previous section. Firstly, the library has to be plated out and a nitrocellulose membrane placed on top. The colonies or plaques will be transferred to this membrane due to the binding of DNA. Place the nitrocellulose membrane in a plastic bag along with a solution containing a radioactive probe (I don't know how long for, sorry :P). Later wash and radiograph the nitrocellulose paper to find the location of the radioactive probes, which should be located in the same place as colonies containing the gene of interest. These colonies can then be removed from the agar plate and cultured in nutrient broths.
Understand the principle of PCR.
Know how PCR can be used for the cloning of specific genes.
See my previous post on PCR.
Be familiar with how PCR can be used for site-directed mutagenesis.
One way of testing out how a mutation affects the function of a protein is to create a primer with a mutation in it. The mutation can be located almost anywhere in a primer, except for the 3' end, because that has to fit to the DNA well for synthesis to proceed.
A moment of silence here while we appreciate all the different approaches...
Okay, back to being serious. There's really just two main approaches (well, at least only two that we learned about during the lecture). They both start off with figuring out what protein you want to clone, determining some of the N-terminal amino acid sequence and from there taking a stab at what the potential DNA sequence would be (presumably taking into account that some amino acids can be coded for by more than one codon).
From there you can take one of two approaches, depending on what technology is available to you. Since several different genomes have already been sequenced, the easiest approach nowadays would generally be to search the database for the gene of interest. You might also be able to find similar genes coding for other proteins, and from there you might be able to get some kind of idea of what your protein does. To be a bit more certain, though, you can amplify the gene (and therefore protein production), allowing you to study the protein more and find out more about it. To amplify the gene, use the database to help you design some oligonucleotide primers so that you can use PCR (more on PCR in an earlier post). This amplified DNA can then be cloned into an expression vector (see the end of my most recent post) to create lots of protein.
An alternative approach can be used if you don't have access to a database for searching for genes. After working out a potential DNA sequence, you can make a DNA probe using this sequence data. This probe can then be used to find the gene of interest in a gene library (more on gene libraries in a bit). Once you've found your gene, you can clone it and so forth.
Be familiar with cDNA library construction and traditional screening by hybridisation.
cDNA libraries can be bought commercially, but just in case you wanted to make your own, I'm gonna tell you how (or rather, the theory behind it anyway). *Do not try this at home*
cDNA is essentially the complement of mRNA. To create cDNA, simply use the enzyme reverse transcriptase to transcribe the mRNA into DNA. Remove the mRNA with alkali and add a poly-G tail (this is so you can add a poly-C primer for synthesis of the other strand). Synthesise the other strand using DNA polymerase.
Next you have to do a couple more fancy things with the cDNA. Firstly, you have to methylate it so that restriction enzymes won't cut it where you don't want it to be cut. Secondly, you have to add EcoRI linkers to either side (essentially the "cutting sites" for EcoRI). Cleave them with EcoRI to form sticky ends.
Now for the recombination part! Take some bacteriophage lambda (a bacteriophage is essentially a virus that infects bacteria) and cleave its DNA with EcoRI. Ligate this to the cDNA. Now you can package this cDNA into bacteriophage lambda so that it can go ahead and infect EcoRI with some shiny new cDNA!
cDNA libraries can be used to screen by hybridisation of a probe, as alluded to in the previous section. Firstly, the library has to be plated out and a nitrocellulose membrane placed on top. The colonies or plaques will be transferred to this membrane due to the binding of DNA. Place the nitrocellulose membrane in a plastic bag along with a solution containing a radioactive probe (I don't know how long for, sorry :P). Later wash and radiograph the nitrocellulose paper to find the location of the radioactive probes, which should be located in the same place as colonies containing the gene of interest. These colonies can then be removed from the agar plate and cultured in nutrient broths.
Understand the principle of PCR.
Know how PCR can be used for the cloning of specific genes.
See my previous post on PCR.
Be familiar with how PCR can be used for site-directed mutagenesis.
One way of testing out how a mutation affects the function of a protein is to create a primer with a mutation in it. The mutation can be located almost anywhere in a primer, except for the 3' end, because that has to fit to the DNA well for synthesis to proceed.
Cloning Vectors and Restriction Enzymes
So, it turns out that this online test is actually an in-class test next week. Which means I might actually have to pay attention to details and commit them to memory just in case they ask some horrible questions. Yikes. (I'm one of those people who does better at understanding general concepts rather than remembering details, so this will be fun.)
To understand the types of cuts to DNA made by restriction enzymes.
See my previous post- Introduction to Cloning.
To be familiar with some of the applications of restriction enzymes in recombinant DNA technology.
This is something that's probably going to crop up again and again over the next few posts in more detail. In a nutshell, though: recombinant DNA technology is all about "cutting and pasting" bits of DNA together. Restriction enzymes act as "scissors" to allow the cutting to be done. (DNA ligase acts as the paste.)
To be aware of the essential characteristics of plasmid vectors.
There are several different kinds of plasmid vectors, but they all have some essential features:
To be familiar with other cloning vectors including BACs and YACs.
BACs, or Bacterial Artificial Chromosomes, are plasmids that can be used to copy fairly large genomic fragments (100 000-300 000 base pairs). They are based on the F plasmid of E. coli (yeah, I get the feeling that we're going to be hearing a helluva lot about E. coli for the rest of my degree). Here are some of the essential characteristics in a list, because lists are great:
To know the important characteristics of expression vectors and how they can be used to express the protein encoded by a foreign gene.
Expression vectors are used to stimulate the production of a protein. The most important characteristic of an expression vector is that it has an inducible promoter (i.e. a promoter that you can switch on and off). For example, the lactose operon, which I've written about before, has a promoter which can be induced by allolactose (although in the lab IPTG, a lactose analogue, is more commonly used). In order to get this vector to produce something else, however, the rest of the gene is simply switched out with the gene of interest.
To understand the types of cuts to DNA made by restriction enzymes.
See my previous post- Introduction to Cloning.
To be familiar with some of the applications of restriction enzymes in recombinant DNA technology.
This is something that's probably going to crop up again and again over the next few posts in more detail. In a nutshell, though: recombinant DNA technology is all about "cutting and pasting" bits of DNA together. Restriction enzymes act as "scissors" to allow the cutting to be done. (DNA ligase acts as the paste.)
To be aware of the essential characteristics of plasmid vectors.
There are several different kinds of plasmid vectors, but they all have some essential features:
- An origin of replication (ori) site to allow the plasmid to be replicated.
- Genes that confer antibiotic resistance. This way they can be "isolated out" by growing a bunch of plasmids on a plate with a particular antibiotic. Only the plasmid vectors with the antibiotic resistance genes will survive.
- A cutting site where the plasmid can be "cut open" and the DNA fragment inserted.
There are many modern engineered plasmids which have a "polylinker" at the cutting site. A "polylinker," in a nutshell, is essentially a section of DNA which contains several cutting sites for several different restriction enzymes. This way, scientists don't have to be limited to just using EcoRI (a restriction enzyme in E. coli).
To be familiar with other cloning vectors including BACs and YACs.
BACs, or Bacterial Artificial Chromosomes, are plasmids that can be used to copy fairly large genomic fragments (100 000-300 000 base pairs). They are based on the F plasmid of E. coli (yeah, I get the feeling that we're going to be hearing a helluva lot about E. coli for the rest of my degree). Here are some of the essential characteristics in a list, because lists are great:
- It has a low copy number (i.e. few copies per cell), which apparently makes the insert more stable. Not sure how this works, though I would assume that fewer replications leads to a lower propensity for mutations.
- It contains par genes which couple plasmid replication to chromosomal replication. This ensures that every daughter cell gets at least one plasmid.
- It has resistance to chloramphenicol, so this can be used as a selectable marker. Unfortunately the plasmid without the inserted DNA also has resistance to chloramphenicol, so this can't be used to tell the recombinant DNA plasmid apart from the plain old normal plasmid. Good thing there's another built-in selectable marker- see my next point:
- The plasmid contains the lacZ gene which codes for beta-galactosidase (see my previous post on the lactose operon) AND the restriction site is located in the middle of the lacZ gene. The implications of this is that when the restriction site is cut and a fragment added, the lacZ gene ceases to function and beta-galactosidase is no longer produced. When colonies are cultured on plates containing X-gal, those colonies with a functioning lacZ gene and thus sufficient amounts of beta-galactosidase will react with X-gal to form a blue product. The recombinant DNA plasmids that do not have a functioning lacZ gene, however, will remain white.
YACs, or Yeast Artificial Chromosomes, are cloning vectors that can be used for eukaryotes. They have two TEL (telomere) sites and a CEN (centromere) site which are required for stability and cell division. YACs can exist in a circular form in bacteria. They can be turned into a linear form by digestion with BamHI- this form is required to insert the YAC into a yeast cell. As for the actual insertion process (which is actually called "transformation")- the yeast cell wall is first digested with enzymes, and then electroporation (the use of an electric current) is used to insert the YAC into the cell.
To know the important characteristics of expression vectors and how they can be used to express the protein encoded by a foreign gene.
Expression vectors are used to stimulate the production of a protein. The most important characteristic of an expression vector is that it has an inducible promoter (i.e. a promoter that you can switch on and off). For example, the lactose operon, which I've written about before, has a promoter which can be induced by allolactose (although in the lab IPTG, a lactose analogue, is more commonly used). In order to get this vector to produce something else, however, the rest of the gene is simply switched out with the gene of interest.
Thursday, March 17, 2016
Enzymes Involved in DNA Replication and Repair
This post should have a lot more new stuff in it, I hope. Or maybe I don't, because that involves more typing and therefore more thinking and more work...
To be familiar with the major enzymes involved in DNA replication in E. coli.
From my previous posts, you should have a good handle on how DNA replication works, so I won't go over that again. Instead, I'm just going to list off a bunch of enzymes and how they aid in this process:
To understand the polymerase and exonuclease activities of the different DNA polymerases.
E. coli has five different DNA polymerases:
To understand how the DNA polymerases replicate DNA with high fidelity.
To be familiar with the different steps in DNA replication that contribute to the overall accuracy of the process.
As mentioned before, DNA polymerase has proofreading functions, eliminating most errors quite quickly. The structure of DNA polymerase also helps to ensure accurate replication. DNA polymerase is shaped like a three-fingered "hand" which closes as bases are added. If an incorrect base is added, the "hand" cannot close as well, allowing the base more time to dissociate. In the event that a base is added, the next base is even slower to associate. The previous mismatched pair is then repositioned into the 3' -> 5' exonuclease site, and the incorrect base is removed.
After replication has occurred, there are still ways of detecting and patching up errors. An incorrect base pairing will cause the width of the helix to change at that point. This can be detected by a protein called MutS. MutS can also recognise the parent strand from its methyl groups: you see, a newly created strand will not have any methyl groups, but will gain them over time. Two more proteins then become involved: a protein called MutL recruits another one called MutH, which nicks the DNA strand near the mutation. Exonucleases then remove a bit more DNA, including the site of the mutation. DNA Polymerase III and DNA Ligase then come in to fix things up.
To be aware that the use of DNA polymerases in the laboratory has been fundamental to the development of Recombinant DNA Technology.
This is something that I'm probably going to end up covering in the next few posts on BIOC2001. PCR is a good example: see an earlier post of mine on this topic.
To be familiar with the major enzymes involved in DNA replication in E. coli.
From my previous posts, you should have a good handle on how DNA replication works, so I won't go over that again. Instead, I'm just going to list off a bunch of enzymes and how they aid in this process:
- Helicase separates the DNA strands so that DNA Polymerase can get in and start replicating.
- Topoisomerase I (a.k.a. gyrase) stops DNA from overwinding by creating and resealing single-stranded nicks. (There's also a topoisomerase II, which can create double-stranded nicks. Should be easy enough to remember- topoisomerase I nicks one strand, topoisomerase II nicks two.)
- Primase creates short RNA primers. DNA Polymerase cannot do its job without these primers.
- DNA Polymerase does the actual replication stuff. Specifically, DNA Polymerase III is the one that does the chromosomal replication in E. coli.
- RNase H removes the primers created by primase.
- DNA Ligase seals off gaps in the lagging strand.
- A clamp loader uses ATP to load a beta clamp onto the DNA. The beta clamp holds the DNA polymerase in place so that it won't have to dissociate and reassociate between bases (dissociative synthesis); rather, it can stay on and do its job smoothly (processive synthesis). (Processive synthesis is MUCH more efficient and rapid than dissociative synthesis.)
To understand the polymerase and exonuclease activities of the different DNA polymerases.
E. coli has five different DNA polymerases:
- DNA Polymerase I is pretty versatile, but it's mainly used for repairing DNA and processing Okazaki fragments. It has three sites: a 5' -> 3' polymerase site, which is where most of the copying and stuff takes place; a 3' -> 5' exonuclease site, which can remove the last base that was added if there's a problem with it; and a 5' -> 3' exonuclease site, which can remove stuff in front of it and is particularly useful for removing RNA primers. The 5' -> 3' exonuclease site is unique to DNA Polymerase I.
- DNA Polymerase II also has roles in DNA repair.
- DNA Polymerase III is the one that replicates the chromosomal DNA.
- 5. DNA Polymerase IV and V are the only two that do not have the 3' -> 5' exonuclease site, and therefore lack proofreading function. They are instead "translesion" DNA polymerases that can replicate through damaged DNA.
Since the 5' -> 3' site on DNA Polymerase I is pretty unique, I'll just talk about it a bit more. As DNA Polymerase I moves along the lagging strand, eventually it reaches a one-base gap and then the RNA primer of the next Okazaki fragment. This one-base gap is called a nick. DNA Polymerase I can simultaneously remove bases ahead of the nick while adding new bases. This process is known as "nick translation." A nick is left at the end, which can be sealed with DNA ligase.
Also, just a quick note on the DNA polymerase reaction itself. Even though there's only one phosphate group per base in DNA, deoxynucleotriphosphates (dNTPs) are used for the synthesis reaction. A new base is added from the previous 3'-OH nucleophilically attacking (I think that's what it's called...) the second phosphate group on the dNTP. This releases pyrophosphate. Pyrophosphate is then broken down by pyrophosphatase, so that the DNA polymerase reaction cannot be reversed. DNA polymerase also has two Mg2+ ions, attached to highly conserved aspartic acid residues, that help to facilitate this process. One Mg2+ ion deprotonates the 3'-OH group, making it more nucleophilic. The other Mg2+ ion facilitates the departure of the pyrophosphate product. (The magnesium ions also have some roles in the 3' -> 5' exonuclease activity, but apparently we don't get to cover that yet. Or something.)
To understand how the DNA polymerases replicate DNA with high fidelity.
To be familiar with the different steps in DNA replication that contribute to the overall accuracy of the process.
As mentioned before, DNA polymerase has proofreading functions, eliminating most errors quite quickly. The structure of DNA polymerase also helps to ensure accurate replication. DNA polymerase is shaped like a three-fingered "hand" which closes as bases are added. If an incorrect base is added, the "hand" cannot close as well, allowing the base more time to dissociate. In the event that a base is added, the next base is even slower to associate. The previous mismatched pair is then repositioned into the 3' -> 5' exonuclease site, and the incorrect base is removed.
After replication has occurred, there are still ways of detecting and patching up errors. An incorrect base pairing will cause the width of the helix to change at that point. This can be detected by a protein called MutS. MutS can also recognise the parent strand from its methyl groups: you see, a newly created strand will not have any methyl groups, but will gain them over time. Two more proteins then become involved: a protein called MutL recruits another one called MutH, which nicks the DNA strand near the mutation. Exonucleases then remove a bit more DNA, including the site of the mutation. DNA Polymerase III and DNA Ligase then come in to fix things up.
To be aware that the use of DNA polymerases in the laboratory has been fundamental to the development of Recombinant DNA Technology.
This is something that I'm probably going to end up covering in the next few posts on BIOC2001. PCR is a good example: see an earlier post of mine on this topic.
Revision of the "Central Dogma" of Molecular Biology
New year, new content!
Or rather... old content wrapped up with a shiny new "BIOC2001" label in the case of this post.
This semester I'm taking ANHB2212 (Human Structure and Development), PHAR2210 (Foundations of Pharmacology), PHYl2001 (Physiology of Human Body Systems) and BIOC2001 (Biochemistry and Molecular Biology of the Cell). Stay tuned for hearing more on all four units, if time and motivation permit!
Without further ado, let's get stuck into the first lecture for BIOC2001 (simply because that's the first one that I have an online test for... or at least I think I'm meant to be having an online test for but they haven't said anything). Most of this is just revision, so I'll be linking to the relevant posts.
Outline the central dogma
I'm just going to copy-paste an earlier post on this one: "I'm sure you probably know this by now- DNA gets transcribed to mRNA, mRNA gets translated into protein, yada yada yada." (Yup, I'm so lazy that I couldn't even be bothered trawling through to find an actual description.)
(Actually, I just stumbled across one serendipitously while finding links for the other sections. In my post on DNA structure, I wrote "The central dogma is simply the process in which DNA is transcribed into mRNA, which is then translated into protein." Same shit really.)
Describe the structure of DNA & RNA
See my previous posts:
DNA Structure
Nucleic Acids- Properties of RNA
Outline mechanism of DNA replication
DNA Replication
Compare & contrast transcription & translation in prokaryotes & eukaryotes
I don't appear to have covered this before, so I might actually have to write something new. Ugh. Anyway, most of the general processes are similar between prokaryotes and eukaryotes: RNA polymerase creates mRNA which is translated on ribosomes to form protein. However, there are some important differences.
The main differences involve timing and location of the two processes. In prokaryotes, since DNA is just floating around in the cytosol, both transcription and translation can occur simultaneously in the cytosol (translation occurs as the DNA is transcribed). In eukaryotes, however, transcription occurs in the nucleus, and then the mRNA goes to the cytosol where it is translated. Since the transcript obviously can't be in the nucleus and the cytosol at the same time, the two processes are not simultaneous in eukaryotes.
A more subtle difference between the two is the type of ribosome that is used. Eukaryotic ribosomes are bigger than prokaryotic ribosomes- 80S as opposed to 70S. (Not 100% sure what "S" stands for but it has something to do with sedimentation. Most important thing here is larger S = bigger ribosomes).
Yet another subtle difference is that the initial amino acid in translation is formylmethionine in prokaryotes, as opposed to plain old methionine in eukaryotes. Not sure what the significance of this is though.
Hmm... what else? Oh yes, there's a "Shine Dalgarno" sequence on prokaryotic DNA, which is where ribosomes attach to and begin translating. I don't think there's anything entirely analogous in eukaryotes. Instead there's a "Kozak sequence," which to my understanding isn't really a binding site, but rather acts to enhance the initiation of translation. The Kozak sequence can vary slightly, and its variations can affect how readily translation is initiated.
Describe the genetic code and its characteristic features
See Translation- Part 1.
Describe a general outline for the process of translation and the sequence features that control its initiation and termination
Translation- Part 1
Translation- Part 2
Or rather... old content wrapped up with a shiny new "BIOC2001" label in the case of this post.
This semester I'm taking ANHB2212 (Human Structure and Development), PHAR2210 (Foundations of Pharmacology), PHYl2001 (Physiology of Human Body Systems) and BIOC2001 (Biochemistry and Molecular Biology of the Cell). Stay tuned for hearing more on all four units, if time and motivation permit!
Without further ado, let's get stuck into the first lecture for BIOC2001 (simply because that's the first one that I have an online test for... or at least I think I'm meant to be having an online test for but they haven't said anything). Most of this is just revision, so I'll be linking to the relevant posts.
Outline the central dogma
I'm just going to copy-paste an earlier post on this one: "I'm sure you probably know this by now- DNA gets transcribed to mRNA, mRNA gets translated into protein, yada yada yada." (Yup, I'm so lazy that I couldn't even be bothered trawling through to find an actual description.)
(Actually, I just stumbled across one serendipitously while finding links for the other sections. In my post on DNA structure, I wrote "The central dogma is simply the process in which DNA is transcribed into mRNA, which is then translated into protein." Same shit really.)
Describe the structure of DNA & RNA
See my previous posts:
DNA Structure
Nucleic Acids- Properties of RNA
Outline mechanism of DNA replication
DNA Replication
Compare & contrast transcription & translation in prokaryotes & eukaryotes
I don't appear to have covered this before, so I might actually have to write something new. Ugh. Anyway, most of the general processes are similar between prokaryotes and eukaryotes: RNA polymerase creates mRNA which is translated on ribosomes to form protein. However, there are some important differences.
The main differences involve timing and location of the two processes. In prokaryotes, since DNA is just floating around in the cytosol, both transcription and translation can occur simultaneously in the cytosol (translation occurs as the DNA is transcribed). In eukaryotes, however, transcription occurs in the nucleus, and then the mRNA goes to the cytosol where it is translated. Since the transcript obviously can't be in the nucleus and the cytosol at the same time, the two processes are not simultaneous in eukaryotes.
A more subtle difference between the two is the type of ribosome that is used. Eukaryotic ribosomes are bigger than prokaryotic ribosomes- 80S as opposed to 70S. (Not 100% sure what "S" stands for but it has something to do with sedimentation. Most important thing here is larger S = bigger ribosomes).
Yet another subtle difference is that the initial amino acid in translation is formylmethionine in prokaryotes, as opposed to plain old methionine in eukaryotes. Not sure what the significance of this is though.
Hmm... what else? Oh yes, there's a "Shine Dalgarno" sequence on prokaryotic DNA, which is where ribosomes attach to and begin translating. I don't think there's anything entirely analogous in eukaryotes. Instead there's a "Kozak sequence," which to my understanding isn't really a binding site, but rather acts to enhance the initiation of translation. The Kozak sequence can vary slightly, and its variations can affect how readily translation is initiated.
Describe the genetic code and its characteristic features
See Translation- Part 1.
Describe a general outline for the process of translation and the sequence features that control its initiation and termination
Translation- Part 1
Translation- Part 2
Subscribe to:
Posts (Atom)