If their first video wasn't awesome enough, this one is:
I also think it's funny how they made a 70's-esque video about a technique that was not developed until the mid-80's (unless you coun't Kleppe's paper as describing PCR1).
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1. Kleppe K, Ohtsuka E, Kleppe R, Molineux I, Khorana HG "Studies on polynucleotides. XCVI. Repair replications of short synthetic DNA's as catalyzed by DNA polymerases." J. Molec. Biol. vol. 56, pp. 341-61 (1971)
Showing posts with label technique. Show all posts
Showing posts with label technique. Show all posts
Sunday, 14 June 2009
Monday, 16 March 2009
The Kunkel Method, or How You - Yes, You! - Can Intorduce Any Mutation you Please into any Gene you Want!
In molecular biology, there are often occasions where you might want to introduce a specific mutation into a particular gene. Let's say you are examining a particular protein, and you want to find out how crucial a certain cystine residue is. You might want to change that cystine residue into, perhaps, a tyrosine residue, and observe how the protein behaves. You could do this by random mutagenesis and sifting through thousands of mutants until you find the one that you need, or you could do it a much quicker way.
The Kunkel method is one such way, and it is actually quite simple, both practically and theoretically. The first step is to clone the gene you want to mutate into whatever plasmid you choose to use. You can use whatever cloning procedure you wish. It doesn't really matter how you clone, just as long as you have a plasmid with your gene in it.
Next, the plasmid must be transformed into an E.coli strain that is ung- dut-. The dut gene encodes for dUTPase, an enzyme that prevents the bacteria from incorporating uracil during DNA replication (remember that uracil pairs with adenine but only in RNA. It is dUTPase that prevents uracil from being used in DNA by destroying all the cell's reserves of uracil during replication). A strain that lacks ung (dut-) will then randomly add uracil to your plasmid when it replicates. E.coli, however, have a backup mechanism in case dUTPase fails. This is uracil deglycosidase, encoded by the ung gene. Uracil deglycosidase cleaves out any uracil that has been added to DNA if dUTPase has been slacking on the job. Transforming into a strain that is also dut- will make sure the uracil in your plasmid stays there.
The next step is to design a primer that contains the region of the gene which you wish to mutate, along with the mutation you want to introduce. If you wanted to change a cystine to a tyrosine, then your primer would span the codon for cystine, but contain the necessary base pair changes to turn that cystine into a tyrosine. This primer will anneal to your plasmid when it denatures, even if the primer does not match it's target 100%. Once you isolate your uracil-containing plasmid, you can do PCR using your mutated primers to create hybrid plasmids: each plasmid will now contain one strand without the mutation and uracil bases, and another strand with the mutation and lacking uracil.
The final step is to isolate this hybrid plasmid and transform it into a different strain that does contain the ung gene. The uracil deglycosidase will destroy the strands that contain uracil, leaving only the strands with your mutation. When the bacteria replicate, the resulting plasmids will contain your mutation on both strands. In essence, you have completely replaced the original gene with your mutated version! You are then free to do as you wish with your new mutated gene.
The Kunkel method, however, is a bit outdated. Many bioscience companies offer kits that allow you to do site-directed mutagenesis even easier. Stratagene, for example, provides a kit that works as follows:
1) Transform your plasmid with gene of interest into any regular laboratory strain. Most strains will be dam+. The dam gene allows the bacteria to methylate the DNA in the plasmid (that is, they will add methyl groups to spots on the DNA. This is a mechanism that the bacteria has to allow it to determine what DNA is its own, and what DNA might be from an invading virus, since viral DNA will not be methylated).
2) Isolate the methylated plasmid, and do PCR with primers containing the mutation you wish to introduce. This will create hemimethylated plasmids: one strand (the one with the original gene) will be methylated and one (the one with your mutated gene) will not be methylated.
3) Add a small amount of the enzyme DpnI to the reaction mix. DpnI actively cleaves up methylated DNA. This will destroy the strands from the original plasmid and leave only your mutated strands.
4) PCR the remaining fragments to produce complete plasmids that contain your mutated gene.
And there you have it. Now you can mutate any gene you want in any manner you wish, and only take a day to do it. No more screening thousands of mutants! Huzzah!
The Kunkel method is one such way, and it is actually quite simple, both practically and theoretically. The first step is to clone the gene you want to mutate into whatever plasmid you choose to use. You can use whatever cloning procedure you wish. It doesn't really matter how you clone, just as long as you have a plasmid with your gene in it.
Next, the plasmid must be transformed into an E.coli strain that is ung- dut-. The dut gene encodes for dUTPase, an enzyme that prevents the bacteria from incorporating uracil during DNA replication (remember that uracil pairs with adenine but only in RNA. It is dUTPase that prevents uracil from being used in DNA by destroying all the cell's reserves of uracil during replication). A strain that lacks ung (dut-) will then randomly add uracil to your plasmid when it replicates. E.coli, however, have a backup mechanism in case dUTPase fails. This is uracil deglycosidase, encoded by the ung gene. Uracil deglycosidase cleaves out any uracil that has been added to DNA if dUTPase has been slacking on the job. Transforming into a strain that is also dut- will make sure the uracil in your plasmid stays there.
The next step is to design a primer that contains the region of the gene which you wish to mutate, along with the mutation you want to introduce. If you wanted to change a cystine to a tyrosine, then your primer would span the codon for cystine, but contain the necessary base pair changes to turn that cystine into a tyrosine. This primer will anneal to your plasmid when it denatures, even if the primer does not match it's target 100%. Once you isolate your uracil-containing plasmid, you can do PCR using your mutated primers to create hybrid plasmids: each plasmid will now contain one strand without the mutation and uracil bases, and another strand with the mutation and lacking uracil.
The final step is to isolate this hybrid plasmid and transform it into a different strain that does contain the ung gene. The uracil deglycosidase will destroy the strands that contain uracil, leaving only the strands with your mutation. When the bacteria replicate, the resulting plasmids will contain your mutation on both strands. In essence, you have completely replaced the original gene with your mutated version! You are then free to do as you wish with your new mutated gene.
The Kunkel method, however, is a bit outdated. Many bioscience companies offer kits that allow you to do site-directed mutagenesis even easier. Stratagene, for example, provides a kit that works as follows:
1) Transform your plasmid with gene of interest into any regular laboratory strain. Most strains will be dam+. The dam gene allows the bacteria to methylate the DNA in the plasmid (that is, they will add methyl groups to spots on the DNA. This is a mechanism that the bacteria has to allow it to determine what DNA is its own, and what DNA might be from an invading virus, since viral DNA will not be methylated).
2) Isolate the methylated plasmid, and do PCR with primers containing the mutation you wish to introduce. This will create hemimethylated plasmids: one strand (the one with the original gene) will be methylated and one (the one with your mutated gene) will not be methylated.
3) Add a small amount of the enzyme DpnI to the reaction mix. DpnI actively cleaves up methylated DNA. This will destroy the strands from the original plasmid and leave only your mutated strands.
4) PCR the remaining fragments to produce complete plasmids that contain your mutated gene.
And there you have it. Now you can mutate any gene you want in any manner you wish, and only take a day to do it. No more screening thousands of mutants! Huzzah!
Monday, 11 August 2008
Techniques in Molecular Biology: Western Blotting
Today I thought I'd go into a little detail on one of the most common experimental techniques used in molecular biology - one that I've done far too many times this summer - the western blot.
In general, western blots are a technique that is used to determine the presence of a particular protein from a biological sample. For example, let's say that you've introduced a plasmid carrying the gene for alcohol dehydrogenase into E.coli and you want to check to see if the bacteria are actively transcribing and translating the gene. An easy way to check this is with a western blot. Westerns are also a useful diagnostic tecnhique in medical labs: westerns can tell you whether a protein is the wrong size, or if a particular protein from a virus or bacterium is present in a sample from a patient.
The theory behind Western Blotting is rather simple. First, you need a sample of proteins. These can be obtained in a variety of ways but they way that I'm most familiar with is through the use of sonication. A culture of cells is grown which (supposedly) express the protein you're interested in. By subjecting these cells to very high frequency sound, the cells rupture and spill all of their contents. One can then put the sonicated sample into a centrifuge, so all of the cellular debris clumps together at the bottom of the test tube, and the proteins in the cell are left floating around in the supernatant.
Now that all the proteins have been isolated, you need to seperate all of them. This is done by using electrophoresis on a polyacrylamide gel. Polyacrylamide forms a mesh through which the proteins travel through when an electric current is applied. Bigger proteins travel though the gel slower than smaller proteins; this, then, allows us to seperate all of the proteins based on their size. At this stage, it is possible to use a protein stain to stain the gel. This will allow us to easily visualize the proteins on the gel. Unfortunately, unless you were working with a pure sample of your protein of interest, you'd see a large smear since the gell contains ALL the proteins from the sonicated cells. To determine the presence of your protein of interest, you'd need a way to specifically visualize your protein and not the rest. Western blotting allows us to do this with ease.
We can speficically visualize whichever protein we want, but polyacrylamide gels do not allow us to do this. Thus, the next step is to transfer the protins to a medium which we can use to probe for a specific protein. One commonly used material for this is nitrocellulose membrane. Nitrocellulose has a very useful property - pretty much any protein sticks to it like glue (this also makes it a little tricky to handle, because you dont want proteins from your hands sticking to it and messing up your results. By making a nitrocellulose-polyacrylamide gel sandwich, and applying an electric current, you can force the proteins in the gel to transfer to the nitrocellulose. The membrane should then contain the proteins seperated as they were on the gel.
Now you're ready to detect specific proteins. This can be accomplished by using antibodies against whichever protein you're interested in. Alot of companies sell antibodies for commonly used proteins. Alternatively, you can design your protein so that it contains a "tag" - a short sequence of amino acids - at one end of the protein which is absent in proteins naturally produced in cells. You can buy antibodies which recognize different tags, and thereby bind only to your protein of interest. These antibodies will bind directly to the proteins on the membrane. Next, secondary antibodies are applied. These will bind specifically to the first antibodies.
The secondary antibodies are the key to detection of your protein. Attached to the secondary antibodies is an enzyme. Which enzyme is attached is a matter of the method of detection used. There are multiple ways of detection, but the most commonly used methods are chemiluminescent detection and pigment production. Both methods work in a similar way; attached to the antibody is an enzyme which will convert a substrate into a product when applied to the membrane. In the case of pigment production, the product is a pigment which can be direcly visualized on the membrane. In the case of chemiluminescent detection, the conversion from substrate to product produces light, which can be detected on a piece of X-ray film. In both cases, the result is a dark band representing your protein of interest. Where there is a dark band, there is the secondary antibody/enzyme; where there is secondary antibody, there is primary antibody; and where there is primary antibody, there is your protein of interest.
The image to the right shows what this end result looks like. You can also run a protein ladder along side your sample, which will show the size of known proteins, so you can determine the size of your protein.The whole process of western blotting can take a few hours to complete (or all day if you're bad at it like me). There is, of course, alot more to western blotting than this; what I've presented is a very generalized idea of how western blotting works. Nevertheless, it is a reliable way to detect the presence of any protein you want in a sample, and has become a staple technique of molecular biology.
In general, western blots are a technique that is used to determine the presence of a particular protein from a biological sample. For example, let's say that you've introduced a plasmid carrying the gene for alcohol dehydrogenase into E.coli and you want to check to see if the bacteria are actively transcribing and translating the gene. An easy way to check this is with a western blot. Westerns are also a useful diagnostic tecnhique in medical labs: westerns can tell you whether a protein is the wrong size, or if a particular protein from a virus or bacterium is present in a sample from a patient.
The theory behind Western Blotting is rather simple. First, you need a sample of proteins. These can be obtained in a variety of ways but they way that I'm most familiar with is through the use of sonication. A culture of cells is grown which (supposedly) express the protein you're interested in. By subjecting these cells to very high frequency sound, the cells rupture and spill all of their contents. One can then put the sonicated sample into a centrifuge, so all of the cellular debris clumps together at the bottom of the test tube, and the proteins in the cell are left floating around in the supernatant.
Now that all the proteins have been isolated, you need to seperate all of them. This is done by using electrophoresis on a polyacrylamide gel. Polyacrylamide forms a mesh through which the proteins travel through when an electric current is applied. Bigger proteins travel though the gel slower than smaller proteins; this, then, allows us to seperate all of the proteins based on their size. At this stage, it is possible to use a protein stain to stain the gel. This will allow us to easily visualize the proteins on the gel. Unfortunately, unless you were working with a pure sample of your protein of interest, you'd see a large smear since the gell contains ALL the proteins from the sonicated cells. To determine the presence of your protein of interest, you'd need a way to specifically visualize your protein and not the rest. Western blotting allows us to do this with ease.
We can speficically visualize whichever protein we want, but polyacrylamide gels do not allow us to do this. Thus, the next step is to transfer the protins to a medium which we can use to probe for a specific protein. One commonly used material for this is nitrocellulose membrane. Nitrocellulose has a very useful property - pretty much any protein sticks to it like glue (this also makes it a little tricky to handle, because you dont want proteins from your hands sticking to it and messing up your results. By making a nitrocellulose-polyacrylamide gel sandwich, and applying an electric current, you can force the proteins in the gel to transfer to the nitrocellulose. The membrane should then contain the proteins seperated as they were on the gel.
Now you're ready to detect specific proteins. This can be accomplished by using antibodies against whichever protein you're interested in. Alot of companies sell antibodies for commonly used proteins. Alternatively, you can design your protein so that it contains a "tag" - a short sequence of amino acids - at one end of the protein which is absent in proteins naturally produced in cells. You can buy antibodies which recognize different tags, and thereby bind only to your protein of interest. These antibodies will bind directly to the proteins on the membrane. Next, secondary antibodies are applied. These will bind specifically to the first antibodies.
The secondary antibodies are the key to detection of your protein. Attached to the secondary antibodies is an enzyme. Which enzyme is attached is a matter of the method of detection used. There are multiple ways of detection, but the most commonly used methods are chemiluminescent detection and pigment production. Both methods work in a similar way; attached to the antibody is an enzyme which will convert a substrate into a product when applied to the membrane. In the case of pigment production, the product is a pigment which can be direcly visualized on the membrane. In the case of chemiluminescent detection, the conversion from substrate to product produces light, which can be detected on a piece of X-ray film. In both cases, the result is a dark band representing your protein of interest. Where there is a dark band, there is the secondary antibody/enzyme; where there is secondary antibody, there is primary antibody; and where there is primary antibody, there is your protein of interest.
The image to the right shows what this end result looks like. You can also run a protein ladder along side your sample, which will show the size of known proteins, so you can determine the size of your protein.The whole process of western blotting can take a few hours to complete (or all day if you're bad at it like me). There is, of course, alot more to western blotting than this; what I've presented is a very generalized idea of how western blotting works. Nevertheless, it is a reliable way to detect the presence of any protein you want in a sample, and has become a staple technique of molecular biology.
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