Overview of epigenetics and Friedreich's ataxia
In a eukaryotic nucleus, DNA is hierarchically packaged into chromosomes. This starts with the wrapping of DNA around histone proteins forming the so-called ‘nucleosome’ which is the basic subunit of chromatin. Chromatin could be less dense and easily accessible to RNAPII (i.e. euchromatin) or tightly packed and inaccessible (i.e. heterochromatin). Accessibility of chromatin is mainly mediated by the interaction of DNA with histone proteins which are subject to a plethora of biochemical modifications that determine the degree of such interactions and hence the compactness of chromatin (Kouzarides 2007; Bannister and Kouzarides 2011). In addition, direct methylation of DNA and RNA-interference based mechanisms have been shown to be involved in the dynamic regulation of chromatin. All of these key events are thought to be inherited independently of the genetic code; a phenomenon known as ‘epigenetics’ (Margueron and Reinberg 2010). Understanding of epigenetic mechanisms in the last decade elucidated novel layers in carcinogenesis as well as in the pathology of neurological diseases such as Alzheimer's or Parkinson's disease (Urdinguio et al. 2009).
Although epigenetics is defined to be independent of DNA sequence, the content of the genomic landscape seems to play an important role in determining the epigenetic chromatin status of a given gene. Importantly, 55% of the human genome is comprised of repetitive DNA sequences (Lander et al. 2001) and heterochromatin is actually thought to have evolved as a protective response against the disruptive effects of aberrant transcriptional noise which is likely to have been triggered by such elements (Yoder et al. 1997; Bashkirov 2002). Indeed, over-expression of repetitive DNA was linked to genomic instability that is associated with cancer and ageing (Schulz et al. 2006; Belancio et al. 2008; Ting et al. 2011; Zhu et al. 2011). Heterochromatinization of repetitive DNA is also important for the maintenance of nuclear architecture needed for the spatial organization of centromeres and sister chromatid segregation during cell division (Fisher and Merkenschlager 2002; Maison et al. 2002; Guenatri et al. 2004; Peters et al. 2008). Being first described by Heitz (1928), heterochromatin is visualised, after DNA staining, through the microscope as densely stained dark patches (Fig. 2a). Some of its important characteristics are that it is more prevalent in non-dividing differentiated cells, forms clusters close to the nuclear periphery, replicates late in S-phase, exhibits reduced recombination rates and is associated with transcriptional repression, particularly at centromeric and telomeric repeats (Dillon 2004).
Figure 2. Heterochromatin and its link with (GAA)n repeats. (a) A mammalian cell nucleus during interphase. When looked at through the microscope after DNA staining, a mammalian cell nucleus (presented as grey) appears to have light and dark regions. Lightly stained parts represent accessible and transcriptionally active ‘euchromatin’ whereas densely stained regions are condensed and transcriptionally silent ‘heterochromatin’. (b) An archetypal example of position effect variegation (PEV) on Drosophila eye. The ‘white’ gene residing on a long chromosome arm is responsible for red eye-colour (left). A coding sequence mutation in this gene typically causes white eye-colour (middle). Radiation induced translocation of the ‘white’ gene into a region close to pericentromeric heterochromatin causes a patchy white/red eye-colour as a result of heterochromatin's invasive nature (right). (c) PEV-like heterochromatic effects induced by (GAA)n in mouse T-cells. The hCD2 transgene with a truncated locus control region allows one to study PEV in mammals by using flow cytometry. The expression of this transgene does not variegate when it is located in a long chromosomal arm (left). When the transgene is placed into close proximity of pericentromeric heterochromatin, the expression of hCD2 variegates in a similar fashion to that observed in the Drosophila eye (middle). Strikingly, insertion of a long (GAA)n tract next to the hCD2 transgene results in a position independent PEV-like variegation (right) implying that (GAA)n repeats could nucleate heterochromatin.
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Heterochromatin is invasive and tends to affect the expression of nearby genes. First lines of evidence regarding this phenomenon were reported in Drosophila, where the euchromatic ‘white’ gene responsible for the red eye-colour was translocated close to pericentromeric heterochromatin as a result of exposure to radiation (Fig. 2b) (Muller 1930). The resulting phenotype was a mosaic eye colour because of the stochastic expression of white in some cells but not in others, a phenomenon known as ‘position effect variegation (PEV)’. PEV takes place when competing factors involved in the formation of euchromatin or heterochromatin are in equilibrium (Fig. 3). Such factors include epigenetic marks including DNA methylation, histone modifications and antisense transcripts as well as sequence elements such as enhancers, silencers, insulators or locus control regions and repetitive DNA (Zuckerkandl 1974; Tartof et al. 1984, 1989; Locke et al. 1988; Festenstein et al. 1999; Dillon and Festenstein 2002). The presence or over-expression of chromatin modifiers could therefore shift the chromatin status in the opposite direction. Notably, many key modifiers of chromatin were discovered using PEV as a model tool (Reuter et al. 1987; Reuter and Spierer 1992; Tschiersch et al. 1994; Elgin 1996; Rea et al. 2000; Dillon and Festenstein 2002).
Figure 3. Competition between euchromatin and heterochromatin. Easily accessible euchromatin is associated with DNA demethylation, histone acetylation and H3K4 methylation. On the other hand, inaccessible and tightly packed heterochromatin is linked to DNA methylation, histone deacetylation, H3K4 demethylation, H3K9 and H3K27 methylation. ‘Glue proteins’ such as HP1 or PRC1 components are known to create a higher order chromatin structure in heterochromatin via their interactions with the nucleosome and each other. The concentration of modifiers responsible for these effects as well as the presence of the binding sites at a given locus determines the final status of transcription. It is thought that if heterochromatic and euchromatic factors are in balance, stochastic expression of genes (PEV) takes place.
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Importantly, microsatellite (GAA)n repeats are shown to behave like pericentromeric heterochromatin. Ohno et al. (2002) reported that (GAA)n tracts show similar spatial nuclear organizations as centromeric repeats. Moreover, expanded (GAA)n repeats are able to silence a nearby gene as shown using the human hCD2 transgene, which encodes a surface marker in thymocytes and T-lymphocytes when introduced into mouse genome. The expression of an hCD2 transgene with a truncated locus control region, when placed close to pericentromeric heterochromatin, exhibits a variegating pattern evident on flow cytometry analyses; providing a tool to study PEV in a mammalian system (Fig. 2c) (Festenstein et al. 1996). Similar to pericentromeric repeats, long microsatellite (GAA)n as well as (CTG)n tracts also induced heterochromatic silencing when fused with the hCD2 gene even when the construct was integrated into a euchromatic location (Saveliev et al. 2003). This discovery pointed to the possibility of abnormal heterochromatinization and thereby silencing of the FXN gene in FRDA, which is associated with expanded intronic (GAA)n elements. Notably, such silencing was also found to be highly sensitive to dosage of the classical PEV modifier heterochromatin protein 1 (HP1), raising the possibility that PEV modifiers were potentially disease modifiers in FRDA. Since then, numerous studies searched for the proof of such heterochromatic effects on the endogenous FXN locus and investigated the chromatin related mechanisms involved in its pathological silencing. The following sections summarise the major findings within the context of epigenetic gene silencing in FRDA.
Direct methylation of DNA
A key layer in regulating gene expression is based on the fact that DNA could be covalently modified at 5′CpG3′ residues by the addition of methyl groups (*-CH3) at the cytosines (C) (Holliday and Pugh 1975; Riggs 1975; Bestor and Ingram 1983; Li et al. 1992). This is performed via the activity of DNA methyltransferases (DNMTs) DNMT-1, -3a and -3b. Whereas, DNMT1 is mainly responsible for the inheritance of this epigenetic mark during DNA replication (Pradhan et al. 1999), the others were shown to be crucial in regulating gene expression at various stages of embryonic development and X chromosome inactivation (Beard et al. 1995; Okano et al. 1999; Geiman and Muegge 2010). The methylation event seems to be reversible (Ramchandani et al. 1999); however, no DNA demethylating enzymes have been identified so far. It is generally accepted that CpG methylation decreases the probability of transcription factor binding or creates additional binding sites for specific proteins (e.g. MeCP2) that recognise methylated DNA and recruit chromatin modifying enzymes (e.g. HDACs) (Dhasarathy and Wade 2008). Whether histone modifications take place before or after the DNA methylation is still a chicken-egg problem. Nevertheless, DNMTs were reported to act in synergy with chromatin modifiers. One important example is the interaction of DNMTs with the heterochromatin protein, HP1 (Fuks et al. 2003; Lehnertz et al. 2003). Importantly, DNA methylation is thought to regulate the majority of the human genome (Shen et al. 2007) and dysregulation of this key modification was associated with cancer and neurodegenerative disorders (Robertson 2005).
The tight link between DNA methylation and gene silencing prompted FRDA scientists to investigate whether this epigenetic mark is affected in the case of expanded (GAA)n repeats. According to the University of Southern California's CpG island search algorithm (Takai and Jones 2002), the FXN gene contains a CpG island covering the promoter, first exon and the beginning of first intron (Fig. 1). Most research, so far, focused on this region and some other CpG residues flanking the (GAA)n repeats. These are summarised in Table 1. Greene et al. (2007) reported a low level of DNA methylation flanking the unexpanded (GAA)n repeats in healthy-derived lymphoblastoid cell lines using bisulphite sequencing. They thought that this methylation pattern in healthy might be because of the Alu element, in which the (GAA)n repeats reside, triggering DNA methylation and causing its bidirectional spread (Liu and Schmid 1993). Strikingly, the number of methylated residues was significantly higher in patient cells. Greene et al. (2007) also reported that some of the cytosines were fully protected from methylation in healthy samples whereas they were extensively methylated in FRDA. One such key CpG residue overlaps with the E-box mentioned earlier.
Table 1. A summary of major studies investigating chromatin changes on the FXN locus
Al-Mahdawi et al. (2008) performed a similar type of analysis using FRDA autopsy samples including relevant tissues such as heart and brain, which exhibited elevated CpG methylation levels upstream of (GAA)n. The same study also reported similar methylation patterns in FRDA-affected tissues of two strains of FRDA disease model mice YG8 and YG22. In line with these findings, two large-scale studies on peripheral blood mononuclear cells also reported significantly higher levels of DNA methylation on the pathologically silenced FXN locus, particularly upstream of the expanded repeats (Castaldo et al. 2008; Evans-Galea et al. 2012). Both found a negative correlation with the age of disease onset and the level of DNA methylation on FXN. Castaldo et al. (2008) also reported that the degree of methylation is proportional to the extent of (GAA)n expansion and Evans-Galea et al. (2012) presented a positive correlation between FXN gene silencing, disease severity and the level of DNA methylation. Interestingly, 5′ untranslated region (UTR) region of the FXN gene did not show any difference in terms of DNA methylation between healthy and FRDA-patient fibroblasts (De Biase et al. 2009).
The fundamental subunit of chromatin - the nucleosome - consists of DNA wrapped around the highly conserved histone H3/H4 tetramer and two pairs of H2A/H2B dimers (Luger et al. 1997). In addition to these core histones, the linker DNA between the nucleosomes is occupied by the linker histone H1 (Allan et al. 1980). Histones are highly basic and positively charged and this explains their strong affinity towards the negatively charged DNA (Luger et al. 1997). As mentioned earlier, biochemical modifications of histones (mainly protruding N-terminal tails of H3 and H4), modulate the degree of this affinity towards the DNA; either by changing the electrostatic charge of the histone or creating conformational alterations (Luger et al. 1997; Kouzarides 2007; Bannister and Kouzarides 2011). The strength of intermolecular interactions with DNA as well as interactions between other histone proteins from the neighbouring nucleosomes are thought to play an important role in regulating the extent of chromatin compaction. In addition, histones could also interact with other proteins such as HP1 to form a higher order compactness of DNA (Bannister et al. 2001; Lachner et al. 2001).
Among the many covalent histone modifications, the most prevalent and well-studied ones are acetylation and methylation of lysine (K) residues. All types of core histones can be acetylated and this correlates well with the transcriptionally active euchromatic state; most possibly because of the fact that the addition of acetyl groups (CH3CO-) reduces the positive charge and thereby the affinity towards the DNA (Allfrey et al. 1964; Sealy and Chalkley 1978; Halleck and Gurley 1981; Hebbes et al. 1988; Hong et al. 1993; Shahbazian and Grunstein 2007). In addition, acetylated histones are known to recruit chromatin-remodelling complexes such as SWI/SNF (Hassan et al. 2002). The occurrence of the acetylation mark on many different lysine residues is thought to create a cumulative effect in loosening up the chromatin structure. Among these many residues, H3K9 and H4K16 acetylation are known to be robust marks for transcriptionally active euchromatin (Mottus et al. 2000; Dion et al. 2005; Shogren-Knaak et al. 2006). While histone acetyltransferases catalyse the acetylation reaction, histone deacetylases (HDACs) remove this euchromatic mark. Phylogenetically classified, there are four major groups of histone acetyltransferases (GNAT, MYST, P300/CREB binding protein and nuclear receptor families) and also four groups of HDACs (HDAC I, II, III, IV – HDACIII also known as sirtuins). Various enzymes belong to each family and it is generally difficult to tell their specificity for a particular lysine residue. Still, some are known to dominate particular nuclear processes (e.g. heat shock or hormone response). These were reviewed in previous publications (Sterner and Berger 2000; Martin and Zhang 2005; Shahbazian and Grunstein 2007) and also presented in this issue by Pandolfo et al.
Deacetylated lysine (K) residues of histones could be subject to another biochemical modification; methylation, which can also take place on arginine (R) residues. Unlike acetylation, the addition of single or multiple methyl groups (*-CH3) does not change the electrostatic charge of histones but it introduces conformational changes because of hydrophobic alterations. Therefore, the outcome of methylation on transcription could be either activation or repression, depending on the place of the methylated residue (Lee et al. 2005; Kouzarides 2007). As reviewed by Kouzarides (2007) and Bannister and Kouzarides (2011), histone methylation is carried out by various histone methyltransferases (HMTs) and reversed by histone demethylases whose mechanisms of action were clearly defined for specific residues. Among the many different types of histone methylations, at least two of them are known to be hallmarks of heterochromatin. H3K9 trimethylation (H3K9m3) is associated with a highly compact chromatin structure and particularly enriched in constitutive heterochromatin such as satellite or telomeric repeats (Jenuwein and Allis 2001; Peters et al. 2001). On the other hand, mono- and di-methylated forms (H3K9me1 and H3K9me2) are mostly enriched on the silenced promoters of formerly euchromatic genes (Barski et al. 2007; Mikkelsen et al. 2007). In constitutive heterochromatin, the histone methyltransferase Drosophila suppressor of variegation 3-9 (human homologue) (SUV39H) catalyses H3K9 trimethylation which is recognised by heterochromatin protein 1 (HP1) (Rea et al. 2000; Bannister et al. 2001; Lachner et al. 2001). HP1 self-dimerises and is therefore thought to create a ‘glue effect’ on chromatin by holding adjacent nucleosomes together. Importantly, SUV39H methyltransferase was shown to interact with HP1 (Aagaard et al. 1999; Bannister et al. 2001; Schotta et al. 2002). This is thought to be the basis of the spreading behaviour of heterochromatin (Fig. 4). In addition to SUV39H, other HMTs such as G9a, SETDB1 and RIZ1 are known to act mostly on inactivated gene promoters (Kouzarides 2007).
Figure 4. Spreading of heterochromatin via HP1 and SUV39H. Heterochromatin is invasive. In a classical example, the spreading of chromatin takes place via the dimerisation of HP1 proteins as well as intermolecular interactions between HP1 and the H3K9 methyltransferase SUV39H. Here, SUV39H catalyses the H3K9 methylation, which is subsequently recognised by HP1. While HP1 molecules self-dimerise and create a more compact chromatin, they also recruit more SUV39H to the locus to exacerbate heterochromatin's spreading.
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Another vital methylated residue is H3K27. Di- and tri- methylated forms of this residue are associated with facultative heterochromatin formed at previously euchromatic genes and the inactivated X chromosome (Wutz and Jaenisch 2000; Barski et al. 2007; Mikkelsen et al. 2007). The activity of H3K27 methylation has been tightly linked to the polycomb system, where one polycomb complex (PRC2) catalyses the methylation and another one (PRC1) recognises methylated H3K27 residues. PRC1 monoubiquitinates the globular H2AK119, which prohibits chromatin remodelling and hence successive elongation of RNAPII (Fischle et al. 2003; Wang et al. 2004; Cao et al. 2005; Hublitz et al. 2009). Intramolecular interactions of PRC2 complexes are thought to be responsible for a low-level chromatin compaction and its spreading (Kim et al. 2002; Francis et al. 2004; Grau et al. 2011). Apart from H3K9 and H3K27 methylation, H4K20 methylation has also received particular attention recently for its association with heterochromatin, as reviewed by Balakrishnan and Milavetz (2010).
As opposed to the ones described above, some other methylation events are associated with the active state of chromatin. RNAPII is known to associate with specific SET methyltransfereases. Thus, the initiating form of RNAPII (phosphorylated on S5) recruits SET1 which catalyses the methylation of H3K4 residues (Ng et al. 2003). While the trimethylated residues of H3K4 are particularly enriched at active promoters, mono- and di- methylated forms are known to extend towards the coding regions (Santos-Rosa et al. 2002; Barski et al. 2007; Mikkelsen et al. 2007). Apart from SET1, there are other HMTs acting on H3K4 methylation (e.g. MLL and ASH1), which possibly act on the first nucleosome where the basal transcription machinery is initially recruited (Kouzarides 2007; Vermeulen et al. 2007). Furthermore, the elongating form of RNAPII (phosphorylated on S2) is known to interact with SET2 which methylates H3K36 residues (Xiao et al. 2003). This mark is therefore particularly enriched in the active coding regions towards the 3′ end (Saunders et al. 2006; Barski et al. 2007; Mikkelsen et al. 2007). Although not much is known about it, H3K79 methylation is another mark that is associated with active transcription (Nguyen and Zhang 2011).
FRDA scientists have collected elaborate ChIP data relating to the histone modification changes on the silenced FXN locus with expanded (GAA)n. In general, all of these studies reported high levels of heterochromatin marks in the first intron of the pathologically silenced FXN in a similar fashion with DNA methylation. An up-to-date summary of literature addressing histone modifications on the FXN locus is also given in Table 1. These studies establish that the expanded (GAA)n repeats are indeed linked to heterochromatinization, as had been suggested by Saveliev et al. (2003) and is consistent with the observations of Ohno et al. (2002). Because of the unstable nature of (GAA)n repeats and their non-specific distribution throughout the genome, ChIP DNA could not be analysed using primers specific to the (GAA)n. However, it was revealed that the classical heterochromatin marks H3K9me2, histone H3 lysine 9 tri-methylation (H3K9me3) and histone H3 lysine 27 tri-methylation (H3K27me3) are enriched particularly in the immediate flanking regions of expanded (GAA)n repeats whereas acetylation marks are reduced. This suggests that heterochromatin is bidirectionally spreading from the expanded (GAA)n tract within the FXN locus. The spreading of heterochromatin from (GAA)n repeats is particularly obvious at the ChIP results presented by Chan et al. (2013), a study that has been published while this review was in press.
The invasive nature of heterochromatin prompted scientists to pursue experiments to understand the boundaries of (GAA)n induced heterochromatin. Importantly, De Biase et al. (2009) discovered a CTCF binding site 154bp downstream of TSS1 within the 5′UTR of the FXN gene. CTCF is a chromatin insulator protein known to prevent the spreading of heterochromatin and generate long-range organization of genes via intramolecular interactions (Bushey et al. 2008). How CTCF exerts the chromatin insulation activity is not yet clearly defined; however, it is thought to recruit euchromatic enzymes and RNAPII or serve as a physical barrier against heterochromatin (Chernukhin et al. 2007; Bushey et al. 2008; Bartkuhn et al. 2009; Witcher and Emerson 2009). Importantly, De Biase et al. (2009) reported depletion in CTCF binding in FRDA patient fibroblasts and associated this with increased antisense transcription (see the next section).
Whether the CTCF binding prevents the spreading of heterochromatin from the expanded (GAA)n repeats towards the promoter, has not been fully answered. De Biase et al. (2009) showed increased H3K9me3 and H3K27me3 levels as well as HP1 on the CTCF binding site of the silenced FXN locus of FRDA derived fibroblasts. Nevertheless, there is still no substantial functional evidence yet to confirm that CTCF prohibits the spreading of (GAA)n centred heterochromatin towards the FXN promoter. Knocking-down CTCF with siRNAs further down-regulated FXN, but whether this is a specific or indirect effect has not yet been fully elucidated. To further check whether CTCF is acting as a barrier it would be interesting to examine the chromatin marks on the promoter region, upstream of the CTCF binding site in the presence and absence of CTCF.
Although there is consensus on the heterochromatinization of the intronic region close to (GAA)n, there is not complete agreement regarding to the promoter/5′ UTR region of the FXN. Al-Mahdawi et al. (2008) reported the enrichment of H3K9me2 and H3K9me3 on the promoter region based on ChIP using human brain tissue. The same study did not observe any reduction in the acetylation marks on the promoter of the silenced FXN. In contrast, Punga and Buhler (2010) and Kim et al. (2011) presented no significant enrichment in heterochromatic methylation marks on the FXN promoter region of FRDA patient derived lymphoblastoid cell lines compared to healthy. In line with this, FRDA lymphoblastoid cells also did not exhibit lower levels of acetylation on the FXN promoter region of FRDA patients compared to healthy (Herman et al. 2006; Kim et al. 2011). However, another study reported lower levels of the majority of acetylation marks in the case of expanded (GAA)n repeats in lymphoblastoid cell lines (Kumari et al. 2011). The discrepancy between these results could be because of technical differences between ChIP experiments and different control cell lines. Thus, whether the promoter of the pathologically silenced FXN is heterochromatinised, is still unclear.
As mentioned before, the effects of (GAA)n induced heterochromatin on the initiation and elongation of transcription is another matter of discussion. In line with other histone marks and RNAPII ChIP data, transcription related marks such as H3K4 methylation as well as H3K36 and H3K79 methylations also seem to be unaffected in the promoter region of silenced FXN alleles as suggested by Kim et al. (2011). The same study reports that all of these key methylated residues are reduced on the coding regions of FXN in FRDA lymphoblastoid cells. The results regarding to FXN promoter and the coding regions are in agreement with a previous study (Punga and Buhler 2010) and overall suggest a transcription elongation defect rather than a problem with initiation. It is noteworthy though, that Kumari et al. (2011) observed reduced H3K4me2 methylation levels at the promoter as well.
Transcription is not solely specific for protein-coding genes and the majority of the human transcriptome has been shown to consist of non-coding RNAs arising from intergenic regions, repetitive sequences and antisense strands of coding genes (Djebali et al. 2012; Dunham et al. 2012; Harrow et al. 2012). Various models have been suggested regarding how antisense transcription may regulate/dysregulate gene expression. As reviewed by Faghihi and Wahlestedt (2009), antisense transcripts could dominate transcription and stall the coding transcription by prohibiting the movement of the transcription bubble in the sense direction or by creating RNA•DNA hybrids. Antisense transcription is also likely to increase the chances of mutation or recombination, which may result in the recruitment of the affected gene to a different subnuclear environment (Bolland et al. 2004, 2007; Ronai et al. 2007). Furthermore, antisense transcripts are also important for genomic imprinting, alternative splicing and nuclear transport or retention (Faghihi and Wahlestedt 2009). Finally, one of the most intriguing discoveries regarding antisense transcription is the finding in the fission yeast, Schizosaccharomyces pombe. Here, double-stranded RNA targets heterochromatinization of the affected genomic region via the activity of the RNA-induced transcriptional gene silencing complex complex with an RNA-interference induced mechanism that involves the DICER enzyme (Fire et al. 1998; Caplen et al. 2001; Elbashir et al. 2001; Reinhart and Bartel 2002; Volpe et al. 2002; Djupedal and Ekwall 2009). Intense efforts are underway to discover whether a similar mechanism occurs in higher-eukaryotes; however, the results so far are inconclusive (Djupedal and Ekwall 2009).
Literature suggesting the presence of a link between heterochromatin and antisense transcription inspired De Biase et al. (2009) and they identified an FXN antisense transcript called ‘FXN antisense transcript 1′ (FAST1) which seems to be produced at higher levels in the fibroblasts obtained from FRDA patients compared to the levels obtained in healthy. Reverse transcription with strand specific primers revealed that FAST1 extends from the first exon and intron towards the promoter. However, it is as-yet unclear where this transcript initiates. Interestingly, siRNA knockdown of CTCF up-regulated FAST1 and reduced FXN sense transcription. As mentioned earlier, the question as to whether the depletion of CTCF actually provokes the spreading of heterochromatin arising from the (GAA)n repeats has not been addressed but was suggested as a potential mechanism. Accordingly, CTCF might prohibit the antisense transcription and therefore the spreading of heterochromatin. Importantly though, the tight link between antisense transcription and heterochromatin was predominantly shown in S. pombe, where heterochromatin is much simpler compared to mammals (Rando and Chang 2009; Beisel and Paro 2011). Although early reports suggest that the RNA interference pathway is indispensable for mammalian cellular lifespan and development (Bernstein et al. 2003; Fukagawa et al. 2004), DICER knock-out experiments did not result in significant changes in mammalian heterochromatin (Murchison et al. 2005; Wang et al. 2006; Ip et al. 2012). This implies that RNAi induced heterochromatinization mechanisms may not be fully conserved in mammals. Still, antisense transcription was linked to transcriptional silencing of oncogenes p15 (Yu et al. 2008) and p21 (Morris et al. 2008) as well as the progesterone receptor (Schwartz et al. 2008). Morris et al.(2008) and Schwartz et al. (2008) suggested a mechanism whereby Argonaute proteins recognise antisense transcripts in mammals.
Overall, it is tempting to think that (GAA)n induced heterochromatinization involves a mechanism where antisense transcription plays a key role; however, the elusive results so far obtained in mammals bring up the need for further characterisation of the link between FAST1 and the pathological heterochromatinization in FRDA. Moreover, the replication of data presented by De Biase et al. (2009) in different disease models (i.e. primary cells, induced pluripotent stem cells or mouse models) will strengthen this hypothesis further.