So having finals upon us should be no excuse to stop the advancement of scientific and philosophical thought on here. So when I wandered into town today to do a favour for my dad (30 min walk for me, 3 hour drive for him) I saw a stall with a sign talking about truth. I still have no idea what a truth is so I thought I would have a look at what truths they were talking about. It was a stall full of Jehovah’s witnesses and I was very nice and polite before you say anything! I always am to there face. Being nasty and telling them, ‘you are probably was wrong as you can be’ doesn’t help win them over to your cause! Richard Dawkins QED. But over our chat I mentioned I supported evolution and was an atheist. But they suggested trusting some of the gaps in evolution had an explanation needed faith. This is an interesting one. It doesn’t but it requires something. I know very little physics and geology. If someone was to question the age of the earth in front of me I would have to turn around I say ‘I don’t know how they worked it out and don’t understand the sciences behind it’ so is it faith in these people I have in other scientists? Probably. But is it bad or wrong for me. I think it is a different sort of faith. First of all it is not blind absolute trust. I know I can easily be wrong about many things (it doesn’t happen often enough though) and I know other can be.
We all need to have this kind of faith to get through life. We cannot know everything so we need to trust others or ‘the system’. Imagine a friend offers you a life in there car but you have never seen them drive before in your life. You don’t know if they are good or bad or will get you killed or not. But you get in anyway. Perhaps because you are so lazy you don’t care if you could die as long as it gets you there faster like me. Or because you have faith that having passed a driving test and having belief in there abilities means you can trust them. It is like knowing peer reviewing produces mostly good knowledge and the author is confident in the results even if you don’t understand the experiments or the maths used to analyse it. This evidence based faith is not a bad thing but needs use to take the results with a large pinch of results. It is not perfect. Sometimes someone who you think would be a good driver turns out not to be while taking a ‘fact’ at face value because you trust the scientist you get it from could mean you are wrong but sometimes we need to take these ‘leaps of faith’. Just know that is what you are doing when you do.
Agree or not?
Tuesday, 31 March 2009
Monday, 16 February 2009
Drosophila: Virgins, balancers and jump-outs
A wise man once said that the sooner you find your model organism, the better. Ignoring the fact that the wise man I am quoting is actually me, I think I generally agree with him. This year I am working on an exciting project which extensively uses the model organism Drosophila melanogaster, geneticists’ favourite little invertebrate. In the 15 short weeks I have been working with flies, I have learned a great deal from these wonderful creatures and experienced emotions from adoration to amusement to frustrated rage in my dealings with them.
Briefly, this is your average everyday “fruit fly” (even though a more appropriate name would be “vinegar fly”) with the complete package of a head, a thorax with 2 wings and 6 legs, an abdomen, and everything else that makes flies flies. I will spare you the tediously detailed anatomy and physiology lecture and will instead jump straight to the cool stuff that interests us geneticists and you readers of genetics blogs.
One of the most interesting things about Drosophila is in fact the people that use them in research. These people are commonly known as “fly pushers”, since they do in fact spend a lot of their time pushing flies around under a microscope using an old half-mangled paintbrush. One thing that you will inevitably hear about once you start hanging out with fly pushers is how interested they are in virgin females. Flies, that is. You see, since experiments in fly genetics involve crossing male flies from one line to female flies of another in order to obtain progeny of the required genotype, one must ensure that the females have not already been fertilised by males before picked out and crossed to a different line. Collecting virgins involves looking for flies that have a dark spot in their abdomen called the meconium, which is visible for 2-3 hours from eclosion (i.e. emergence of the adult from the pupa). Females will not be receptive to males for up to 8 hours from eclosion so any fly that has a meconium can be safely considered a virgin. The most efficient way to collect virgins is to empty a vial of eclosing flies in the morning, selecting females with visible meconium, and returning in the evening. Any females in that vial will be virgins even if the meconium isn’t visible, since less than 8 hours will have passed since your last collection. Much like the world we live in these days, virgins can be rather hard to find, so if you ever hear a scientist worrying about the fact that they need as many virgins as possible, you now have an idea of what they are talking about.
Now let’s delve into some actual genetics. One of the multitude of reasons Drosophila are an excellent test tube for genetic experimentation is something called a “balancer chromosome”. A line of flies carrying a recessive lethal mutation can be maintained by tracking the mutation using dominant markers, or by “balancing” it over a different lethal mutation on its homologous chromosome. This way each chromosome “rescues” the mutation the other is carrying. However, if recombination takes place between these chromosomes, it is likely that you will then have one chromosome carrying both mutations over a wild type chromosome, which will quickly take over the stock and the mutations will be lost. You could always outcross males every generation (there is no recombination in male Drosophila) but that would be very laborious. In order to get around this problem, fly geneticists initially discovered and subsequently intentionally generated balancer chromosomes. These are full chromosomes which contain multiple inversions (essentially all the genes are still there and intact, but their order is jumbled up) in order to suppress recombination with its
homologous chromosome and make any gametes with recombined chromosomes non-viable due to aneuploidy. They also carry some dominant visible markers such as Cy (curly wings), Tb (tubby), Sb (stubble) and others, and some recessive lethal markers, meaning that they can never go homozygous.
So, to illustrate this, if you have a recessive lethal mutation on chromosome 2 (let’s now call that chromosome “Yorf”), in order to establish a line that will self-perpetuate and not need constant attention, all you need to do is cross it to a line that carries a balancer second chromosome, for example the chromosome CyO. You will now have a line whose flies will always have the same genotype, i.e. Yorf/CyO. As the Yorf/CyO flies interbreed, any Yorf/Yorf progeny will not survive because the mutation will be lethal, and any CyO/CyO flies will also not survive because of the recessive lethal marker that CyO carries. There will also never be any recombination between the Yorf and CyO chromosomes because of the multiple inversions in CyO that prevent homologous pairing. Wonderful, isn’t it?
Another great thing about Drosophila is insertional mutagenesis. P-elements are transposons, short stretches of DNA which, in the presence of transposase and absence of inhibitor, are excised from the genome and reinserted randomly into another site. Some P-elements are rendered inactive by having the transposase gene deleted. This temporarily immobilises the transposon, until that fly is crossed to a fly carrying an active transposase gene. In the progeny, the P-element will jump out, and if it inserts into the coding sequence of a gene, it can disrupt its function. A mutagenesis screen can then be carried out and the insertion locus can be mapped using standard complementation or cytological mapping. There is however an application of P-element mutagenesis that has accelerated Drosophila genetics in recent years.
The targeted gene knock-out system developed in mice is an excellent way to study the role of single genes, but the long generation time and ethical concerns make the mouse a rather cumbersome model organism. An analogous targeted gene disruption tool has been developed in Drosophila, taking advantage of a property of P-element transposition. When a P-element is transposed, its excision from the genome can sometimes be imprecise, taking some of its flanking DNA along with it, thus causing a deletion in that gene. Since an extensive library of mapped P-element inserts already exists (in over half the fly genome), a line carrying a P-element insert in a particular gene can be obtained and the P-element “jumped out” by crossing to a transposase line. In the cases where the jump-out has been imprecise, the result will effectively be a targeted gene knock-out.
This is, of course, only scratching the surface of the range of genetic tools that Drosophila offers. The best way to understand fly genetics is to try some crosses on paper yourself. Some quite amazing things can be achieved within a few weeks’ time simply by performing a series of crosses and studying the progeny. The waiting time is still a little too long for impatient people like myself, but careful time planning can ensure that you always have some flies to play with and are never stuck staring at a pupa, waiting for the damn fly to emerge.
Briefly, this is your average everyday “fruit fly” (even though a more appropriate name would be “vinegar fly”) with the complete package of a head, a thorax with 2 wings and 6 legs, an abdomen, and everything else that makes flies flies. I will spare you the tediously detailed anatomy and physiology lecture and will instead jump straight to the cool stuff that interests us geneticists and you readers of genetics blogs.One of the most interesting things about Drosophila is in fact the people that use them in research. These people are commonly known as “fly pushers”, since they do in fact spend a lot of their time pushing flies around under a microscope using an old half-mangled paintbrush. One thing that you will inevitably hear about once you start hanging out with fly pushers is how interested they are in virgin females. Flies, that is. You see, since experiments in fly genetics involve crossing male flies from one line to female flies of another in order to obtain progeny of the required genotype, one must ensure that the females have not already been fertilised by males before picked out and crossed to a different line. Collecting virgins involves looking for flies that have a dark spot in their abdomen called the meconium, which is visible for 2-3 hours from eclosion (i.e. emergence of the adult from the pupa). Females will not be receptive to males for up to 8 hours from eclosion so any fly that has a meconium can be safely considered a virgin. The most efficient way to collect virgins is to empty a vial of eclosing flies in the morning, selecting females with visible meconium, and returning in the evening. Any females in that vial will be virgins even if the meconium isn’t visible, since less than 8 hours will have passed since your last collection. Much like the world we live in these days, virgins can be rather hard to find, so if you ever hear a scientist worrying about the fact that they need as many virgins as possible, you now have an idea of what they are talking about.
Now let’s delve into some actual genetics. One of the multitude of reasons Drosophila are an excellent test tube for genetic experimentation is something called a “balancer chromosome”. A line of flies carrying a recessive lethal mutation can be maintained by tracking the mutation using dominant markers, or by “balancing” it over a different lethal mutation on its homologous chromosome. This way each chromosome “rescues” the mutation the other is carrying. However, if recombination takes place between these chromosomes, it is likely that you will then have one chromosome carrying both mutations over a wild type chromosome, which will quickly take over the stock and the mutations will be lost. You could always outcross males every generation (there is no recombination in male Drosophila) but that would be very laborious. In order to get around this problem, fly geneticists initially discovered and subsequently intentionally generated balancer chromosomes. These are full chromosomes which contain multiple inversions (essentially all the genes are still there and intact, but their order is jumbled up) in order to suppress recombination with its
homologous chromosome and make any gametes with recombined chromosomes non-viable due to aneuploidy. They also carry some dominant visible markers such as Cy (curly wings), Tb (tubby), Sb (stubble) and others, and some recessive lethal markers, meaning that they can never go homozygous.So, to illustrate this, if you have a recessive lethal mutation on chromosome 2 (let’s now call that chromosome “Yorf”), in order to establish a line that will self-perpetuate and not need constant attention, all you need to do is cross it to a line that carries a balancer second chromosome, for example the chromosome CyO. You will now have a line whose flies will always have the same genotype, i.e. Yorf/CyO. As the Yorf/CyO flies interbreed, any Yorf/Yorf progeny will not survive because the mutation will be lethal, and any CyO/CyO flies will also not survive because of the recessive lethal marker that CyO carries. There will also never be any recombination between the Yorf and CyO chromosomes because of the multiple inversions in CyO that prevent homologous pairing. Wonderful, isn’t it?
Another great thing about Drosophila is insertional mutagenesis. P-elements are transposons, short stretches of DNA which, in the presence of transposase and absence of inhibitor, are excised from the genome and reinserted randomly into another site. Some P-elements are rendered inactive by having the transposase gene deleted. This temporarily immobilises the transposon, until that fly is crossed to a fly carrying an active transposase gene. In the progeny, the P-element will jump out, and if it inserts into the coding sequence of a gene, it can disrupt its function. A mutagenesis screen can then be carried out and the insertion locus can be mapped using standard complementation or cytological mapping. There is however an application of P-element mutagenesis that has accelerated Drosophila genetics in recent years.
The targeted gene knock-out system developed in mice is an excellent way to study the role of single genes, but the long generation time and ethical concerns make the mouse a rather cumbersome model organism. An analogous targeted gene disruption tool has been developed in Drosophila, taking advantage of a property of P-element transposition. When a P-element is transposed, its excision from the genome can sometimes be imprecise, taking some of its flanking DNA along with it, thus causing a deletion in that gene. Since an extensive library of mapped P-element inserts already exists (in over half the fly genome), a line carrying a P-element insert in a particular gene can be obtained and the P-element “jumped out” by crossing to a transposase line. In the cases where the jump-out has been imprecise, the result will effectively be a targeted gene knock-out.
This is, of course, only scratching the surface of the range of genetic tools that Drosophila offers. The best way to understand fly genetics is to try some crosses on paper yourself. Some quite amazing things can be achieved within a few weeks’ time simply by performing a series of crosses and studying the progeny. The waiting time is still a little too long for impatient people like myself, but careful time planning can ensure that you always have some flies to play with and are never stuck staring at a pupa, waiting for the damn fly to emerge.
Thursday, 22 January 2009
Meaning out of nonsense
Imagine you are a eukaryotic cell. You have many genes and make many mRNAs. One problems is mutated genes, pseudogenes and mistakes by RNAPII all make mRNAs which contain Premature Stop Codons (PTC). These then make truncated proteins which could be toxic to the cell. Eukaryotes have evolved a mechanism to recognise these and degrade them. I guess prokaryotes do not have an analogous system (that I know of) because the half-life of these mRNAs are minutes long not hours long like most eukaryotic mRNAs. Eg the average mRNA half-life in Arabidopsis is between 4-6 hours and is very variable between mRNAs. Personally I am not convinced this is an important enough reason to have a quality control check mechanism. The system I am referring to is called Nonsense-Mediated mRNA Decay (NMD). Mediators of it have been found in the early branching Giardia and is conserved between fungi/animals to plants. A PTC is recognised as wrong rather than a correct stop codon at the end of a ORF during the first round of translation by the ribosome. When the ribosome reaches a stop codon it has two choices, terminate correctly or stall and cell effectors of NMD to take it away for degradation by whatever nucleases that cell uses. It was first found in yeast and the mutants were called upf1-3 (for up-frame shift 1-3) and these yeast grow fairly normally but have increased levels of transcripts with stop codons. Raises the question why did it evolve if not very important when you KO it. In C. elegance SMG1-7 were identified having genital defects. Strangely, these were involved NMD with SMG2-4 corresponding to UPF1-3. Homologues of most of these proteins are found in many other eukaryotes. UPF1 is generally phosphorylated when ribosome stalling takes place and condemns it to degradation. Interestingly, it is a DEAD box RNA helicase (SF2 I believe).
The two questions I find interesting is 1) how do organisms tell the difference between a PTC and normal stop codon and 2) how this can be used to regulated gene expression in cell signalling. You may ask ‘James, you charismatic stallion, how can something degrading useless mRNAs be useful or regulated for controlling gene expression’ and I would reply ‘wait and read the rest of this post you $#£!&%’. The first question is rather interesting as different organisms do different things. Yeast and invertebrates such as C. elegance and the fly pushers Drosphilia use something called the 3’ faux model. Something in the 3’ end allows normal termination when the stop codon is here but NMD starts when the stop codon (PTC) is distant from the 3’ end. It has been found the distance from the poly-A tail and therefore the Poly-A Binding Protein (PABP) determines this. When a long 3’ UTR is present the mRNA is degraded. When PABP is tethered close to the stop codon termination occurs normally. This also works in plants and mammals. However, another mechanism also works in both of them. When an introns is spliced out an Exon-Junction Complex (EJC) is left behind and if a EJC is found near a stop codon by the ribosome NMD starts. Why this operates in both plants and mammals but not yeast and invertebrates is curious. Did both mechanisms operate in the last common ancestor of plants and animals and has been lost a few times in the fungi and animal kingdoms or is it convergent evolution? I hope to better understand that.
Another and perhaps more important question is what is NMD role in plants and animals. Does it provide another function other than keeping toxic protein levels low? Short answers, yes. It affect development in C. elegance and embryo lethality in mice. Clearly it plays a role in animal development. In yeast the cells senesce sooner in upf1-3 mutants because telomeres shorten. In mammals, amino acid deletion causes problems for the ribosome and NMD is down regulated and genes for amino acids biosynthesis are up-regulated so more amino acids will be made. In Arabidopsis, when upf-1 is completely KO we see embryo lethality but Knock-down alleles see some developmental phenotypes (such as in the flowers) and altered stress responses.
I will finished up now but feel free to ask questions. I have just finished writing a 4-page grant proposal on this. Just to piss Mel off, I will say this. I didn’t do any reading for this. This was all from memory when I got home. No PDF files were opened – not one!
The two questions I find interesting is 1) how do organisms tell the difference between a PTC and normal stop codon and 2) how this can be used to regulated gene expression in cell signalling. You may ask ‘James, you charismatic stallion, how can something degrading useless mRNAs be useful or regulated for controlling gene expression’ and I would reply ‘wait and read the rest of this post you $#£!&%’. The first question is rather interesting as different organisms do different things. Yeast and invertebrates such as C. elegance and the fly pushers Drosphilia use something called the 3’ faux model. Something in the 3’ end allows normal termination when the stop codon is here but NMD starts when the stop codon (PTC) is distant from the 3’ end. It has been found the distance from the poly-A tail and therefore the Poly-A Binding Protein (PABP) determines this. When a long 3’ UTR is present the mRNA is degraded. When PABP is tethered close to the stop codon termination occurs normally. This also works in plants and mammals. However, another mechanism also works in both of them. When an introns is spliced out an Exon-Junction Complex (EJC) is left behind and if a EJC is found near a stop codon by the ribosome NMD starts. Why this operates in both plants and mammals but not yeast and invertebrates is curious. Did both mechanisms operate in the last common ancestor of plants and animals and has been lost a few times in the fungi and animal kingdoms or is it convergent evolution? I hope to better understand that.
Another and perhaps more important question is what is NMD role in plants and animals. Does it provide another function other than keeping toxic protein levels low? Short answers, yes. It affect development in C. elegance and embryo lethality in mice. Clearly it plays a role in animal development. In yeast the cells senesce sooner in upf1-3 mutants because telomeres shorten. In mammals, amino acid deletion causes problems for the ribosome and NMD is down regulated and genes for amino acids biosynthesis are up-regulated so more amino acids will be made. In Arabidopsis, when upf-1 is completely KO we see embryo lethality but Knock-down alleles see some developmental phenotypes (such as in the flowers) and altered stress responses.
I will finished up now but feel free to ask questions. I have just finished writing a 4-page grant proposal on this. Just to piss Mel off, I will say this. I didn’t do any reading for this. This was all from memory when I got home. No PDF files were opened – not one!
Thursday, 1 January 2009
Arch nemesis
Evidence is the cornerstone of science. It is what allows scientists to reject or accept hypotheses, and a robust piece of evidence can force the staunchest defender of a theory to completely reject it. People tend to like evidence when it supports their "beliefs", and dislike it when it refutes them, but scientists have no choice, they need to be apathetic and disinterested towards any results, so as to avoid introducing bias. However, scientists are also (rumoured to be) people, and the tendency to like or dislike a piece of evidence can be quite hard to resist.
How could a scientist maintain an interest in their research while avoiding interest in the data generated from it? It seems almost self-contradictory. What goals do scientists usually have when carrying out research? For a scientist working at a pharmaceutical company (assuming they wanted to keep their job), this goal would be to get a drug on the market. For a graduate student, the goal could be to find something surprising or revolutionary in order to jump-start their career. For a hardened senior professor, it could be to prove their own hypotheses right and boost their reputation (and maybe get in line for a Nobel prize?). All these goals are fundamentally selfish, and you would be hard pressed to find pure altruism in even the "most moral" scientist (whatever that means...). Perhaps that's because scientists tend to be ambitious, competitive animals, and that's a selfish motivator in itself, but I would say it's because people's goals tend to be selfish in nature.
These selfish goals scientists have certainly pose the most danger in making them introduce bias into their data. However, the drive that these goals give to scientists is what has led to so many historical discoveries. I'm sure some discoveries had altruistic motives but let's just say Watson and Crick weren't really thinking about saving humanity from terrible doom. Not to paint a sad picture of scientists, but sometimes the only thing that scientists get up for in the morning is the tiny chance that today's experiment will get them even slightly closer to achieving their goal.
I think that scientists manage to minimize the danger of introducing bias by finding the one person on Earth they most disagree with and establishing an intimate, life-long professional relationship with them, manifested mostly through heckling at conferences and angry e-mail exchanges. This works much more efficiently in academia than in industry, because industry scientists are usually either unwilling or not allowed to talk to outsiders about anything they do. Academics are also secretive to a certain extent in order to avoid being scooped, but the spirit of collaboration and criticism is much stronger. This kind of mutually abusive relationship is what every scientist needs, and I personally can't wait to meet my arch nemesis!
P.S. Happy 2009!
How could a scientist maintain an interest in their research while avoiding interest in the data generated from it? It seems almost self-contradictory. What goals do scientists usually have when carrying out research? For a scientist working at a pharmaceutical company (assuming they wanted to keep their job), this goal would be to get a drug on the market. For a graduate student, the goal could be to find something surprising or revolutionary in order to jump-start their career. For a hardened senior professor, it could be to prove their own hypotheses right and boost their reputation (and maybe get in line for a Nobel prize?). All these goals are fundamentally selfish, and you would be hard pressed to find pure altruism in even the "most moral" scientist (whatever that means...). Perhaps that's because scientists tend to be ambitious, competitive animals, and that's a selfish motivator in itself, but I would say it's because people's goals tend to be selfish in nature.
These selfish goals scientists have certainly pose the most danger in making them introduce bias into their data. However, the drive that these goals give to scientists is what has led to so many historical discoveries. I'm sure some discoveries had altruistic motives but let's just say Watson and Crick weren't really thinking about saving humanity from terrible doom. Not to paint a sad picture of scientists, but sometimes the only thing that scientists get up for in the morning is the tiny chance that today's experiment will get them even slightly closer to achieving their goal.
I think that scientists manage to minimize the danger of introducing bias by finding the one person on Earth they most disagree with and establishing an intimate, life-long professional relationship with them, manifested mostly through heckling at conferences and angry e-mail exchanges. This works much more efficiently in academia than in industry, because industry scientists are usually either unwilling or not allowed to talk to outsiders about anything they do. Academics are also secretive to a certain extent in order to avoid being scooped, but the spirit of collaboration and criticism is much stronger. This kind of mutually abusive relationship is what every scientist needs, and I personally can't wait to meet my arch nemesis!
P.S. Happy 2009!
Friday, 17 October 2008
Kinetoplastid DNA
It seems to be that time of the year in which we post about something to do with our posts, as these are the topics we are supposed to be reading about now. Therefore, a post on parasites. That’s where the connection with my project finishes, but anyway.As you might know, tryponosomatids are organisms from the family Troponosomatidae, which include some pretty nasty unicellular parasites such as Trypanosoma brucei, causer for African sleeping disease, and Leishmania, causer of leishmaniasis. As other higher eukaryotes, these parasites are characterised by two genomes: a nuclear genome and a mitochondrion genome, 10-30% of all the cell DNA, situated in the kinetoplast (the name given to the mitochondrion in these organisms).
Fig.1 A- Schematic minicircle organization. B- in vivo network organization, seen sideways.
(Adapted from reference 1)
So far nothing extraordinary. Things get interesting, when one looks at the structure of the kinetoplast DNA (kDNA) in these species. The kDNA is constituted by thousands of small DNA circles, the minicircles, and a few dozen of large DNA circles, the maxicircles. These are interlocked, as in a chain mail of medieval armour. Each circle is interlocked with 3 other neighbouring circles. The minicircles and maxicircles are stretched into a disk-shaped structure, situated near to the flagellar basal body (Fig.1). The maxicircles are those more similar to normal mitochondrion genome. They include rRNAs and genes encoding proteins associated with the respiratory processes that takes place in this organelle. However, the maxicircles mRNA requires extensive editing, namely the introduction or deletion of uridylate. The minicircles encode the guide RNAs used as templates for this editing. Since there are many types of editing required, as well as maxicircles with different sequences, there is the need for the thousand minicircles with different sequences. Seems a bit of a wasteful system, but what do I know!
More interesting is to think of the replication process. The minicircles must stop being interlocked, replicated, interlocked again, and in the end originate two new networks of thousands of minicircles that separate into the two new daughter cells. Let us look at the different steps, following an individual minicircle (Fig.2):
Fig.2 kDNA replication model. (From reference 2)
1- A minicircle must first be released from the network into the so-called kinetoflagellar zone (KTZ), where several proteins involved in the process exist. Here the unidirectional replication of the circle occurs, although the details of this process are not well known.
2- The two daughter minicircles move to opposite antipodal sites of the circle, where there are two protein assemblies. Here a variety of reactions occur: the RNA primers are removed, the gaps between
3- Minicircles are attached by topoisomerase II to the network periphery adjacent to the antipodal sites.
The way the minicircles are attached to the disk varies with the species of trypanosomatid. In T.brucei, for example, the minicircles accumulate at the network poles. In T.cruzi, and C.fasciculata, the minicircles seem to be uniformly attached around the periphery of the disk, creating a ring of new minicircles, of increased thickness. How this uniform attachment is done is not certain, but it is though that either the antipodal protein complexes or the disk itself must rotate! (Fig.3)
Fig.3 Replication in C.fasciculata. New minicircles have been labelled with fluorescent nucleotides. The arrangement of the new minicircles around the periphery of the disk is obvious (from reference 2).
4- Regardless of the method, new minicircles are attached to the disk, and the valence must increase to 4 to 6 attachments per minicircle, in order to ensure twice as many minicircles in the same space (the mitochondrion membrane has not doubled at this point). When the space increases, topoisomerase II ensures that valence returns to 3.
5- Finally, the nicks are repaired and the network splits into 2, although how exactly this division occurs is not well known. Most importantly, it is not known for sure how to guarantee that the two daughter networks have the exactly same minicircles. Considering their importance in maxicircle mRNA editing, the loss of one minicircle could have dramatic consequences.
There is much to be explained in this process, namely identify exactly how the minicircles are moved to the different antipodal sites, or the details of individual minicircle replication, but such understanding will require the identification of the proteins involved. Considering the complexity of the process, many must be involved, but only a few have been identified so far.
This is probably not life-changing research, but it is surely interesting. It is important to remind ourselves once in a while that though we learn the textbook pathways and processes, Nature seems to like using hundreds of alternatives. Just to make scientists lives harder, I assume!
Liu, B., Liu, Y., Motyka, S., Agbo, E., & Englund, P. (2005). Fellowship of the rings: the replication of kinetoplast DNA Trends in Parasitology, 21 (8), 363-369 DOI: 10.1016/j.pt.2005.06.008
(2) Smith D. and Parsons M. (1996). Molecular Biology of Parasitic Protozoa, Kinetoplast DNA: Structure and Replication, Oxford University Press, USA
Subscribe to:
Posts (Atom)