Of moss and me

I recently had a look through some new papers published on moss (Physcomitrella patens) and found one published in Cell titled ‘Transcriptional control of gene expression by microRNAs’ [1]. This didn’t seem to fit the same pattern of a lot of moss papers I have read (ie in less high impact journals). This paper shows how useful moss is for studying gene expression using reverse genetics (so hopefully my PhD wont be a failure).

I think it is important to study important biological problems, such as RNA silencing and epigenetics, in a diverse range of organisms. I take this title from a short review published in Nature Medicine recently (as well as the Steinberg book) by Sir David Baulcombe (who holds a prestigious chair at Cambridge). He wrote about what we have learnt by studying things in plants (eg discovery of transposons) and their importance as a model, using his own research into RNA silencing and epigenetics as the main example [2]. I think this new moss paper goes some way to support this, but also to show the importance of this particular organism. My choice to work with Physcomitrella was because I wanted to stay in the plant world but wanted to work with something where I could knock out specific genes by homologous recombination and was simple to grow. Physcomitrella can be grown on simple agar media plates and joins the few species where genes can be knocked out.

Khraiwesh et al (2010) looked at what happened when you knocked out two Dicer-like (DCL) proteins in moss [1]. Dicer proteins in animals and plants cut long dsRNA to make small RNAs loaded onto Argonaute proteins (producing the effector complex). This effector complex is what causes the target mRNA to be cut or repress its translation. When PpDCL1a was knocked out you see a lack of microRNAs (miRNAs) being produced and the plant is restricted to its first developmental stage only. Not surprising. Strangely, when you knocked out PpDCL1b get production of miRNAs but not cleavage of the targets. It has been shown in animals, but not other plants, Dicer proteins can sometimes be used to help a small RNA in the effector complex to cut the target, as well as in miRNA or siRNA biogenesis. Very interestingly they found these targets had a lower level of expression in the mutant plants. If cleavage is not reducing the mRNA levels then what? The answer to this as many other odd questions in biology is epigenetics. Well DNA methylation at least. siRNAs have been shown to cause gene silencing but not much evidence for miRNAs doing this before. The authors went onto show the ratio of miRNA to mRNA target is critical. If you only have a little miRNA compared to mRNA target you get cleavage. If you have lots of miRNA to target mRNA it causes DNA methylation and silencing. But the methylation was only seen in the knockout so why should we care?

Finally they end the paper by showing ratio of miRNA to target mRNA is important in an endogenous gene involved with a hormone response. The plant hormone ABA mediates drought response and abiotic stress. In moss, when ABA treated you get a decrease in a transcription factor response gene. This is a target of a natural miRNA. This down-regulation is due to an increase in miRNA expression, changing the effect from cleaving the transcription factor’s mRNA to targeting its promoter for methylation and silencing expression. This study shows that miRNAs have the potential to cause DNA methylation, which had not yet been show, as well as suggesting a mechanism for it. Future studies can now go on to show the relevance of this in other systems from yeast to flies to mammals.

I end by simply saying MOSS ROCKS.

Khraiwesh, B., Arif, M., Seumel, G., Ossowski, S., Weigel, D., Reski, R., & Frank, W. (2010). Transcriptional Control of Gene Expression by MicroRNAs Cell, 140 (1), 111-122 DOI: 10.1016/j.cell.2009.12.023

Baulcombe, D. (2008). Of maize and men, or peas and people: case histories to justify plants and other model systems Nature Medicine, 14 (10), 1046-1049 DOI: 10.1038/nm1008-1046

Non-B DNA structures and disease

People seem to have started contributing actively to the blog again so I thought it was my turn to try again. And this is an achievement, as facing any science, in particular anything written by me about it, would have been impossible a month ago.

I decided to keep things simple for this one and just give a short introduction on the theme of non-B DNA structures and how they influence disease, the topic of my open essay.

When we think about DNA, we think about its B-DNA structure, the one that Watson and Crick described in that famous Nature paper more than 50 years ago, i.e. a right-handed double helix, 20Å diameter, in which the base pairs are almost perpendicular to the axis of the helix, 10.5bps per turn. Although most DNA will be in this structure, it can also form other structures, the so called non-B DNA structures. Some of these are shown in the figure. These alternative structures have a few common characteristics: firstly, whether or not a structure is formed, and which specific structure, seems to be sequence dependent, as it often involves formation of new pairings of bases. Secondly, most of these structures are in higher energetic state than normal B-DNA, for example, because they require the separation and reformation of hydrogen bonds. Overall, therefore, DNA with a favourable sequence tends to remain in its B-DNA form, requiring events such as DNA replication, transcription or protein binding to be transiently converted to these unusual structures. Sequences with propensity to form unusual structures are quite common in the human genome, and it seems that in some cases they are necessary for the normal functioning of the cell. However, they have also been implicated in disease, and this is what I’ll be focusing till the end of this post.

There is a considerable list of diseases thought to involve, as part of their pathology, the formation of non-B DNA structures, but the mechanisms by which this is though to happen can vary. For example, certain sequences/structures have been thought to cause genomic instability, e.g. by causing chromosomal rearrangements due to the propensity of these structures to promote double strand breaks. Another interesting form of genomic instability associated with these unusual structures is repeat expansion, implicated in motor diseases such as Huntington’s disease. Non-B DNA structures, namely Z-DNA, have also been associated with viral infections.

Now, there isn’t really enough space here to talk about everything, so I’m going to give one examples of a situation in which a non-B DNA structure is though to be involved in disease. Friedreich ataxia (FRDA) is a disease caused by the expansion of GAA•TTC tracts in intron 1 of a gene encoding the protein frataxin, essential for mitochondrial function. In this disease, repeat expansion is associated with loss of protein expression. One of the current models by which frataxin production is thought to be reduced in expanded GAA•TTC repeats is based on their ability to form triplexes (an alternative model suggests that epigenetic changes may also be important). Duplex opening within the repeated region, due to the passage of RNA polymerase, is thought to allow one of the separated single strands to form Hoogsteen hydrogen bonds with the purine strand of a B-DNA duplex within the same repeated sequence. This leads to the formation of a 3-stranded helix. Its formation on the non-template GAA strand in frataxin probably interferes with RNA polymerase progression. The free template strand is then thought to base pair with the newly synthesised RNA transcript, forming stable RNA/DNA dimers and preventing further transcription.

Now, this is only a model, for a specific type of structure within the context of a specific disease. And although different sequences have been shown to form these unusual structures, and these structures to be somehow associated with specific diseases, to neatly demonstrate how exactly one influences the other seems to be quite hard. This is particularly difficult because different groups often use different model organisms or protocols, leading to sometimes contradictory results. Overall, although I liked writing about this topic, since I had never considered how the structure of DNA could have an impact on disease, I got the feeling that a lot of work still needs to be done before convincing evidence is given for how exactly these structures impact on our wellbeing, and what sort of therapeutics can be developed from this knowledge.

Red, GAA repeat strand; blue, GTT complementary strand; orange, RNA transcript; yellow, RNA polymerase. 1, Intramolecular triplex; 2, Stalled RNA polymerase; 3, DNA/RNA hybrid

Hebert, M. (2008). Targeting the gene in Friedreich ataxia Biochimie, 90 (8), 1131-1139 DOI: 10.1016/j.biochi.2007.12.005

Bacolla, A. (2004). Non-B DNA Conformations, Genomic Rearrangements, and Human Disease Journal of Biological Chemistry, 279 (46), 47411-47414 DOI: 10.1074/jbc.R400028200

WELLS, R. (2007). Non-B DNA conformations, mutagenesis and disease Trends in Biochemical Sciences, 32 (6), 271-278 DOI: 10.1016/j.tibs.2007.04.003

Wang, G., & Vasquez, K. (2009). Models for chromosomal replication-independent non-B DNA structure-induced genetic instability Molecular Carcinogenesis, 48 (4), 286-298 DOI: 10.1002/mc.20508

When a bit more fat is not that bad

Nothing is as simple as it looks in Biology. We can’t even rely on gene or protein sequences to tell us what is going to happen. At the chromosomal level, the new field of epigenetics is showing how the nucleotide sequence does not define everything. At protein level we have to consider post-translational modifications. My final year project, now at the final write up stages, concentrated on an aspect of Trypanosoma brucei molecular biology which I very pompously described as a potential drug target and the cure for African sleeping disease. This bit was probably ‘paper talk’ though. In any case, the project focused on protein palmitoylation.

Palmitoylation is part of a group of several lipid modifications that can occur in proteins. Examples include N-myristoylation, prenylation and GPI-anchors. N-myristoylation is the reason why I thought it was not pushing it to much to say that we can kill parasites by targeting these modifications. N-myristoylation corresponds to the co-translational addition of myristate to an N-terminus glycine, and is catalyzed by N-myristoyltransferases (NMT). It so happens that NMTs were shown to be essential for parasite viability, and several antifungal NMT inhibitors apparently work on reducing NMT activity in T.brucei as well(1).

But back to palmitoylation, the post-translational addition of palmitic acid to a cysteine residue (reviewed in 2 and 3). It can occur in any place in the protein, and so far no consensus palmitoylation motif has been identified. Often it occurs in an N-terminus cysteine, just next to a myristoylation site. These proteins are said to be dually acylated. Palmitoylation can also happen in close proximity to prenylation sites. Palmitoylation can perform many functions in the cell, the most obvious being protein tethering to membranes and cellular localization. It has also been shown to interfere with protein-protein interactions and even with protein degradation. For example, the yeast SNARE Tgl1, when is not palmitoylated, can interact with the ubiquitin ligase Tul1(4).

Those paying attention (or still reading) will think that it is a bit strange that palmitoylation can regulate protein degradation like that. In fact, most of these lipid modifications, N-myristoylation for example, last for the life time of the protein. Well, palmitoylation is characterized by a unique feature: it is reversible. This means that it can determine processes that other lipidations cannot. Dynamic trafficking of proteins of proteins, for example. This is the case of Ras protein, which through an deacylation/reacylation cycle is able to exchange between the plasma membrane and the Golgi apparatus (5). Reversibility can also be important for signalling. Regulator of G-protein signalling 2 (RGS2) was shown to increase or decrease its GTPase-activating ability depending on which of its cysteine residues are palmitoylated(6).

The process of palmitoylation in itself is catalyzed by the enzymes palmitoyl-acyltransferases (PATs). These were pretty hard to identify, apparently. Not only they were difficult to purify, but the reaction itself can occur spontaneously with biological significance(7), which made many sceptic on whether these enzymes existed at all. However, they were identified, first in yeast, now in mammals and in my dear T.brucei.

And what was supposed to be a short post if already long enough. I finish by adding that my project was about finding proteins that were palmitoylated and that would change their localization in the absence of this modification, in an attempt to identify substrates for RNAi PAT studies in T.brucei. I must say that the project was not particularly successful, but I developed the important skill of writing long discussions based on few results, which I bet will come handy in the future.

I now challenge Mel to continue in this post-translational modification topic and tell us about his project on glycosylation.


References:

1. Price, H. (2002). Myristoyl-CoA:Protein N-Myristoyltransferase, an Essential Enzyme and Potential Drug Target in Kinetoplastid Parasites Journal of Biological Chemistry, 278 (9), 7206-7214 DOI: 10.1074/jbc.M211391200

2. Nadolski MJ, Linder ME. Protein lipidation. FEBS J 2007;274:5202-10.

3. Greaves J, Chamberlain LH. Palmitoylation-dependent protein sorting. J Cell Biol 2007;176:249-54.

4. Valdez-Taubas J, Pelham H. Swf1-dependent palmitoylation of the SNARE Tlg1 prevents its ubiquitination and degradation. EMBO J 2005;24:2524-32.

5. Rocks O, Peyker A, Kahms M, et al. An acylation cycle regulates localization and activity of palmitoylated Ras isoforms. Science 2005;307:1746-52.

6. Ni J, Qu L, Yang H, et al. Palmitoylation and its effect on the GTPase-activating activity and conformation of RGS2. Int J Biochem Cell Biol 2006;38:2209-18.

7. Kummel D, Heinemann U, Veit M. Unique self-palmitoylation activity of the transport protein particle component Bet3: a mechanism required for protein stability. Proc Natl Acad Sci U S A 2006;103:12701-6.

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 Okazaki fragments are filled by DNA pol, and nick as sealed by DNA ligase. An important characteristic, however, is that the minicircles maintain at least one gap in their structure. This is thought to be a self-check feature of this system, as a way to guarantee that no minicircle is replicated twice.

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

The wonders of auxin

I am a complete sell out. I have worked for a plant biotech company and getting funding from Gatsby (a charity with a goal, alone with others, is to get scientists interesting in plants). These people have been very nice by helping me pay rent, buy DVDs and science magazines as well as sending me to Mexico for a conference. Therefore, I thought I should give something back by educating my friends in a bit of plant biology. Don’t worry faint hearted, I will keep it nice and molecular as that is the way I like it.

After returning from my Mexico trip I realised a few things, English seems to be spoken by everyone, don’t eat food from Mexico City airport and auxin is a fascinating plant hormone and virtually everyone in the plant community agrees. Most talks there were on auxin biosynthesis, receptor function and structure, its control of shoot branching and transport around the plant. However, I do not recall all of it. Or hardy any of it as I was jet lagged with little prior knowledge of plant hormones (plus it has been over a year since the conference).

Auxin was the most interesting of all the plant signalling molecules. It has been studied for over 100 years and only today we are starting to understand how it controls plant development and cell biology. Auxin is a small molecule made from an amino acid and its most biologically active for is called IAA. Auxin controls cell elongation and division, and can promote these or stop them depending on the tissue type auxin enters. Auxin causes lateral root growth and patterns the vascular system of plants. Auxin starts shaping the plant in the embryo. What is interesting is how it regulates many of these things.

It controls gene expression by activating Auxin Response Factors (ARFs). This is done by destabilizing proteins called Aux/IAAs. Genes activated by the addition of auxin contain Auxin Response Elements (ARE) which an ARF binds to. When no auxin is present, an ARF is bound to an ARE but it is diamerized with an Aux/IAA, which represses transcription of this gene. When auxin enters the cell, it gets ride of this Aux/IAA and the ARF diamerizes with another ARF. This causes transcription of the auxin regulated gene. How does auxin cause breakdown of Aux/IAA? (I thought I would sound more intelligent if I asked a lot of questions) Auxin doesn’t have a nice receptor at the cell membrane that activates a second message or a phosphorylation pathway, no that would be to simple. In 2005 Ottoline Leyser and Stefan Kepinski (then post-doc, now lecturer at Leeds) published in Nature that TIR1 was an auxin receptor. tir1 mutants had been known to be deficient in auxin signalling for a long time but not thought to be a mutation in the receptor. TIR1 is an F-box protein, which are not famous for being receptors (until now). Normally they simply act as an E3 ubiqutin (Ub) ligase, meaning they take Ub from one protein and add it to another creating a poly-ubiquitin tag that sends a protein for degradation by the 26S proteasome. Ubiqutination and targeting to the proteasome is found animals and fungi and plays an important role in signalling, including regulation of the cell cycle! TIR1 adds Ub to the Aux/IAA when auxin enters the cell. It was assumed that auxin was perceived by some other protein in the cell and caused some modification of the Aux/IAA or TIR1 to cause this to happen, probably by phosphorylation. Now we know that auxin binds toTIR1 and acts as a molecular glue, bridging the gap between TIR1 and Aux/IAA so it can be broken down. I think this is a brilliant method. I am glad to see plants are being original and creating cool new signalling pathways instead of being just like boring old animals and fungi that love there MAPK so much they should just go along and marry it. Please see my simplified diagram of the auxin signalling pathway, it is not perfect, but who is......except maybe Colombo. But then how does auxin have very different effects on different cell types. This appears to be because different tissues expresses different ARFs and Aux/IAAs and these turn on different genes but little is known right now.



One final thing, is TIR1 the only receptor? A good question indeed. Like I said earlier, I like asking questions to make me sound smart. Other F-box proteins appear to do the same job. The same only story of redundancy. However, this style of signalling cannot explain all the effects auxin has on plants. Some auxin responses occur very quickly after auxin addition. These happen so quickly, it is unlikely proteolysis followed by transcription and translation of effectors can account for them. Things like guard cell outward K+ current up, increased cytoplasm calcium, cell wall acidification starts (to help increase cell wall expansion) and elongation growth. How can we explain auxin’s influence over these physiological changes? There is debate and investigation over what causes these things. Some are caused by changes in membrane potential and perhaps Auxin Binding Protein 1 (ABP1) has some control over some changes but I am not convinced and other features are unrelated to this. After auxin addition MAP kinase activity increase but this is unrelated to both TIR1 and ABP1. There are still a lot of unanswered questions.

Here is a short, concise review by Ottoline’s old post-doc (now at Leeds), which I found yesterday and does a far better job of introducing people to this that I have!
The anatomy of auxin perception
http://www.ncbi.nlm.nih.gov/pubmed/17876776?ordinalpos=57&itool=EntrezSystem2.PEntrez.Pubmed.Pubmed_ResultsPanel.Pubmed_DefaultReportPanel.Pubmed_RVDocSum

Another one talking a more detailed look at auxin receptors and the debate around them. I have not read it fully, sadly.
Receptors for auxin: will it all end in TIRs?
http://www.ncbi.nlm.nih.gov/sites/entrez?db=pubmed&cmd=search&term=Receptors+for+auxin%3A+will+it+all+end+in+TIRs%3F

I have not referenced here for two reasons. First, I can’t be bothered and secondly, a lot of this is from conferences or meetings I have been to. I hope you enjoyed this little rant from the plant person.

Yet another twist in the world of gene expression - transcription factories

When I went to a conference on post translational modification and chromatin, one talk really grabbed me, ironically it was the one that didn’t relate to the topic of the conference. I was going to facebook people about it and then remembered this existed.

First of all I will ask you, did you know that transcription only happened at a few sites within the nucleus? In mouse cells from the animal there are between 100-300 of these but in cultured cells such as HeLa cells there are many more. Transcription factories also known as RNAPII foci are where most, if not all mRNA is produced. This amazed me and raises the obvious questions of why and how. The why may be obvious. It is a good idea to keep the nucleus as I see it ‘tidy’ but in more technical terms it is a way of keeps gene expression organised and regulating it (see below). The how this work I don’t think has been addressed! All I can say is watch this space.

Another interesting feature of mammalian gene expression that has come out of these studies is that most ‘active genes’ are not on all the time like classically thought. Most genes appear to show a pulse fashion of expression. Travelling to transcription factories and being expressed then returning to a dormant stage until they travel back to transcription foci for another round of expression and so on. However, some genes show the classical continuous expression pattern and are seen at these sites all the time. One example is Beta-globin. Genes like these are relatively rare compared to those showing the pulse type expression but both are continuously on in there cell type. If elements of the beta-globin promoter is deleted then not only is overall expreesion decreased but it shows the pulse type expression.

One of the most interesting things is that some genes will co-localize to the same transcription factory more often than you expect by pure chance. What is more interesting is these genes can be separated by several megabases! or even be on different chromosome. Using the 3C and 4C assay you can join different pieces of DNA together from different parts of the genome that have transcribed within the same transcription foci. These have shown along with RNA FISH the physical movement of genes to the same place to be transcribed. One of the two Beta-globin alleles within a cell was found to enter the same factory with some other genes at a rate of ~25% with a background rate expected to be less than 2%. These gene’s promoters were searched and a simple motif was found: CCACC. This is actually a binding site for a regulator of Haem biosynthesis and haemoglobhin production called Eklf. When Eklf is knocked out the association between these genes stops! This transcription factor must recruit to these genes to the transcription factories it is located at. I can see two mutually exclusive models here (tell me what you think or extra ones to consider). Either 1) something, perhaps Eklf recruits genes to these sites and physically moves them there or 2) genes with this motif move to all transcription foci and only stay to be transcribed at ones that have Eklf present. I would guess the second is more likely but I see no evidence for either model at this time. Another question I would like answered is whether other or maybe all transcription factors work in this way. I think it is unlikely but perhaps that is the case!

One more interesting thing related to this is the story in B cells of mice. It has been shown that the proto-oncogene Myc is expressed in the same foci as immunoglobulin heavy chain (IgH) a very high proportion of the time. The interesting part is that these two genes have been show in many blood cancers to be translocated to be on the same chromosome so perhaps there is a link between transcription factories and cancer. Well that is what the people doing the research on it want to think. After all related to cancer equal more funding as I told a friend of mine earlier. A new lab has been set up by the post-doc who led most of the described research to explore the cancer link.



And I now realise one very important thing, I should have been writing my result section now I have a deadline as well as lab work, S#£!!%$! Hell!




Cameron S. Osborne, Lyubomira Chakalova, Jennifer A. Mitchell, Alice Horton, Andrew L. Wood, Daniel J. Bolland, Anne E. Corcoran, Peter Fraser (2007). Myc Dynamically and Preferentially Relocates to a Transcription Factory Occupied by Igh PLoS Biology, 5 (8) DOI: 10.1371/journal.pbio.0050192

Cameron S Osborne, Lyubomira Chakalova, Karen E Brown, David Carter, Alice Horton, Emmanuel Debrand, Beatriz Goyenechea, Jennifer A Mitchell, Susana Lopes, Wolf Reik, Peter Fraser (2004). Active genes dynamically colocalize to shared sites of ongoing transcription Nature Genetics, 36 (10), 1065-1071 DOI: 10.1038/ng1423

The Fanconi anaemia DNA repair pathway

Fanconi anaemia (FA) is an autosomal recessive chromosomal instability, characterised by congenital abnormalities, defective haemopoiesis, a high risk of leukaemia and development of solid tumours. FA patients have a 10% incidence of leukaemia and are 50 times more likely to develop solid tumours (particularly hepatic, oesophageal, oropharyngeal and vulval) by early adulthood. The disease consists of 13 complementation groups, each connected to mutations in 13 corresponding FANC genes (A, B, C, D1, D2, E, F, G, I, J, L, M, and N) which make up the FA pathway. Two thirds of FA cases are caused by inherited mutations in FANCA.

Interstrand crosslink damage

FA cells are characterised by hypersensitivity to damage by DNA Interstrand CrossLinking (ICL) agents such as mitomycin C and diepoxybutane. Cells fail to repair ICL damage, resulting in polysomies, radial chromosomal structures and unrepaired breaks (Figure 1). ICL agents tether two DNA strands together, creating a physical barrier to replication and transcription, which causes arrest of DNA replication. In normal cells, several mechanisms come in place to minimize damage and maintain the replication event. This necessitates removal of the physical barrier, repair of the gap and re-establishment of the replication fork. This complex repair mechanism involves the Fanconi anaemia proteins, translesion synthesis (TLS) polymerases, and the homologous recombination (HR) machinery. The FA pathway has also been implicated in spontaneous DNA damage repair, despite early speculation that the pathway was specific to ICL damage. In addition, there is some uncertainty as to whether the FA pathway also functions in intra-strand crosslink repair.

The FA pathway

The FA proteins are part of a complex DNA repair pathway, which is not yet fully understood. They are commonly arranged in three distinct groups according to their function. The first group is the FA core complex, which consists of FANC A, B, C, E, F, G, L and M. This complex stabilises stalled replication forks, processes the ICL site and activates a second group, which consists of FANCD2 and FANCI. Finally, group 3 consists of the remaining FANC proteins, BRCA2 (which is the same gene as FANCD1), FANCJ and FANCN, which have accessory or uncertain function in the pathway (Figure 2). The most recent model of this pathway is described in Wang (2007), Kennedy & D’Andrea (2005) and Niedernhofer (2007), and a detailed description of the model follows.

An ICL physically blocks replication by preventing separation of the strands by the advancing DNA helicase, which causes the enzyme to arrest. The replication fork arrest activates the ATR kinase, which subsequently phosphorylates FANCD2. A double strand break (DSB) upstream of the ICL is carried out by XPF-ERCC1, MUS81-EME1 or MUS81-EME2. Simultaneously, the monoubiquitin ligase FANCL, part of the FA core complex, monoubiquitinates FANCD2 and FANCI. This stage is regulated by the deubiquitinating enzyme USP1, and the monoubiquitin acts as a chromatin localisation signal. UPS1 is itself regulated by cell cycle controls, and self-cleaves in response to DNA damage. The integrity of the whole of the core complex is essential for this stage, as proven by the lack of chromatin localisation in FA core complex mutants. The downstream function of FANCI-Ub is uncertain, but FANCD2-Ub is arguably the most important component of this pathway, acting as a “fire captain” to recruit the proteins needed for the rest of the pathway (Figure 3).

As group 2 is activated, the FA core complex is also carrying out DNA processing in preparation for the next step in the pathway. FANCM acts as a DNA translocase. It anchors the rest of the FA core complex onto DNA, which stabilises the broken replication fork and facilitates removal of the ICL. A nucleotide excision repair (NER) endonuclease “unhooks” the ICL (which remains attached to the one strand), and translesion synthesis (TLS) polymerases recruited by the FA core complex fill in the gap. The TLS polymerases’ ability to pass through the damaged site has the trade-off of being error-prone, introducing random mutations in the repair site. The ICL adduct is then completely removed by a NER exonuclease, and the result is a replication fork broken by a DSB, with the ICL removed and the gap filled in (Figure 4). The next step is to repair the DSB, which is carried out by homologous recombination (HR) proteins.

HR-mediated replication fork re-establishment

FANCD2-Ub forms a complex with BRCA2 and chromatin. All the FANC proteins, along with BRCA1, NBS1, PCNA, RAD51 and other related proteins are recruited by the FANCD2-Ub/BRCA2-chromatin complex into nuclear foci at the damage site. BRCA1 forms a complex with BRCA2 and is essential for the formation of nuclear foci, even though its exact function is unknown. FANCJ, also identified as the helicase BRIP1 (BRCA1-interacting protein), depends on BRCA1 to translocate to the DNA damage site and unwind the replication fork in order to promote HR. The MRN complex, which consists of MRE11, RAD50 and NBS1 and has 3’-5’ exonuclease activity, processes either side of the DSB to prepare it for HR.

DSB repair can occur in two ways. Firstly by single-strand annealing, an error-prone non-homologous end joining (NHEJ) pathway, where the two strands are trimmed until homologous repetitive sequences are reached, and the strands are then annealed at that region of homology. This is mutagenic since the non-homologous intermediate region is deleted, but this is preferable to the arrest of replication. Secondly by strand invasion HR, where RAD51 is loaded onto chromatin by BRCA2 to form a nucleoprotein filament, within which Sister Chromatid Exchange (SCE) occurs and a Holliday junction forms between the strand being repaired and a homologous strand (Figure 5). Holliday junctions are intersections of four strands of DNA, which appear most commonly during meiosis when “crossing-over” occurs, and are resolved by endonucleases such as MUS81-EME1. FA cells show high levels of SCE in FANCC mutants, while other FANC mutant cells have normal SCE levels, suggesting a regulatory role for FANCC over SCE formation and a generally significant role in HR. HR thus re-establishes the replication fork and at the end of the S-phase, USP1 deubiquitinates FANCD2 to turn the pathway off.

In summary, ICL damage causes replication arrest, one of the strands is separated by a DSB, the FA core complex stabilises the broken fork and TLS repairs the ICL. At the same time, the structural integrity of the FA core complex is essential for the monoubiquitination of the “fire captain” FANCD2, which forms a complex with BRCA2 and chromatin and recruits HR and accessory proteins into nuclear foci. Finally, the MRN complex processes the ends of the DSB for HR to repair it and allow replication to restart.

There has been increased interest in the FA pathway in the past decade or so due to the discovery of its connection with cancer and complex relationship with the HR pathway. The pathway model is still in its early stages but already there are important questions raised about cell cycle regulation and its connection with DNA repair.

References

Kennedy, R.D. and D'Andrea, A.D. (2005) The Fanconi Anemia/BRCA pathway: new faces in the crowd. Genes Dev. 19: 2925-2940.

Niedernhofer,L.J., Lalai,A.S., and Hoeijmakers,J.H.J. (2005) Fanconi Anemia (Cross)linked to DNA Repair. Cell 123: 1191-1198

Patel,K.J. and Joenje,H. (2007) Fanconi anemia and DNA replication repair. DNA Repair 6: 885-890

Tischkowitz,M. and Dokal,I. (2004) Fanconi anaemia and leukaemia - clinical and molecular aspects. British Journal of Haematology 126: 176-191

Wang,W. (2007) Emergence of a DNA-damage response network consisting of Fanconi anaemia and BRCA proteins. Nat Rev Genet 8: 735-748