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Do DNA repressors exist?

Do DNA repressors exist?


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I know about enhancers and the mechanism that lead them to increase the gene expression of their targets but I was wondering if similarly DNA repressors exist. I know about protein repressors but I am looking for some kind of anti(or reversed)-enhancer equivalent in the genome which would act like an enhancer but reduce gene expression.

I am aware of repressed/poised enhancers which would kind of be that process, even if not really a pure repression of the target gene but rather a "non-overexpression". I also know about insulators but again this would be a different mechanism.

Some intergenic region binding protein repressors which by a folding mechanism similar to enhancers would repress a gene perhaps?


Yes, these sequences exist and they are called "silencers" (surprising, right?). There are different mechanisms by which this silencing of genes can happen.

In the "classical" way the silencer is bound by a transcription factor which either passively suppress the gene by hindering the binding of specific transcription factors or by actively preventing the assembly of the general transcription factors. See the figure from paper 1:

Additionally there are non-classical negative regulatory element (NRE), which are usually elements upstream of the promoter which inhibit the binding regulatory proteins. NRE can also be enhancers depending on the proteins bound on them. Some NRE can induce a bend of the DNA, inhibiting the access to enhancer or promoter elements.

References:

  1. Transcriptional control and the role of silencers in transcriptional regulation in eukaryotes.
  2. Transcriptional Regulatory Elements in the Human Genome

Difference Between Repressor and Corepressor

The key difference between repressor and corepressor is that repressor protein directly binds to the operator sequence of the gene and inhibits gene expression while corepressor protein binds to the repressor protein and indirectly regulates the gene expression.

Genes are the units of heredity. They have genetic information to make proteins. In order to make proteins, genes should be expressed via transcription and translation. Transcription factors should bind to promoters and enhancers and recruit RNA polymerase enzyme to initiate transcription. Gene expression can be regulated especially at the transcription level. The repressor is a protein that inhibits gene expression. Corepressor is a protein that indirectly regulates gene expression by binding to transcription factors. Repressors recruit corepressor complexes. In eukaryotes, both repressors and corepressors are proteins.

CONTENTS


Bacteriophages, Part A

Graham F. Hatfull , in Advances in Virus Research , 2012

2 Cluster G immunity systems

Putative repressor genes have also been identified in Cluster G phages, such as BPs and its closely related relatives Halo, Angel, and Hope ( Sampson et al., 2009 ). These phages are temperate, and stable lysogens can be recovered from infected cells. The BP repressor (gp33) is not closely related to Cluster A-encoded or any other phage repressors but does contain a putative helix-turn-helix DNA-binding motif, and expression of gp33 confers immunity to superinfection by all of the Cluster G phages (see Section III.G ). The repressor gene is located immediately upstream of the integrase gene (32) and the two genes are predicted to overlap. A notably unusual feature of genome organization is that the crossover site for integrative recombination at attP is located within the repressor gene itself, such that the gene product expressed from the prophage is 33 residues shorter than the virally encoded form. This suggests the possibility that integration plays a central regulatory role in the lytic–lysogenic decision.


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PcG proteins in the DNA damage response

PcG genes directly and indirectly regulate various aspects of the DDR. Bmi1 is required for maintenance of the redox balance. Bmi1 deficiency in mice leads to increased levels of reactive oxygen species, which results in DNA damage and DDR activation (Liu et al., 2009). With regard to the role of PRC2, cell-based assays support an involvement of EZH2 in the regulation of the G1/S and G2/M checkpoints upon treatment with clastogens (Wu et al., 2011b). In addition, in breast cancer cells, EZH2 might affect DSB repair indirectly through repression of the homologous recombination enzyme Rad51 (Chang et al., 2011) and of its paralogues (Zeidler et al., 2005), as well as through regulation of BRCA1 (Gonzalez et al., 2011 Gonzalez et al., 2009).

Recent work has begun to elucidate the involvement of specific PcG proteins in the local DDR at DSBs. A summary of PRC1 and PRC2 components that have been found to localize to these DNA damage sites is presented in Table 1.

Mammalian Polycomb group proteins that localize to DNA damage sites

Protein Protein motifs Biochemical activity Method of DNA damage induction Function in DDR References
PRC2EZH1 SET domain Histone methyltransferase Laser micro-irradiation (GFP-tagged protein) Prevent sensitivity to DSB-inducing agents (Chou et al., 2010)
EZH2 SET domain Histone methyltransferase UV Laser micro-irradiation endonuclease-induced DSBs (ChIP) H2O2 induced foci Prevent sensitivity to DSB-inducing agents (Chou et al., 2010 O'Hagan et al., 2008 O'Hagan et al., 2011)
SUZ12 Zinc finger Stimulates histone methyltransferase activity UV Prevent sensitivity to DSB-inducing agents (Chou et al., 2010)
PHF1 (interactor) Two PHD fingers and a Tudor domain Stimulates histone methyltransferase activity Laser micro-irradiation (GFP-tagged protein) Prevent sensitivity to DSB-inducing agents (Hong et al., 2008)
PRC1BMI1/PCGF4 RING domain E3 ubiquitin ligase Laser micro-irradiation hydroxyurea-, camptothecin- and IR-induced foci aphidicolin-induced fragile sites (ChIP) ZNF-induced DSBs (ChIP) H2A/H2AX ubiquitylation Prevent genomic instability Prevent sensitivity to DSB-inducing agents (Chagraoui et al., 2011 Chou et al., 2010 Ginjala et al., 2011 Ismail et al., 2010 Nacerddine et al., 2012 Pan et al., 2011)
MEL18/PCGF2 RING domain E3 ubiquitin ligase Laser micro-irradiation n.d. (Chou et al., 2010 Ginjala et al., 2011)
RING1A/RNF1 RING domain E3 ubiquitin ligase Laser micro-irradiation (GFP-tagged protein) n.d. (Chou et al., 2010)
RING1B/RNF2 RING domain E3 ubiquitin ligase Laser micro-irradiation IR-induced foci H2A/H2AX ubiquitylation Prevent sensitivity to DSB-inducing agents (Chou et al., 2010 Ginjala et al., 2011 Ismail et al., 2010 Wu et al., 2011a)
CBX2 Chromodomain Methyl-lysine binding Laser micro-irradiation ZNF-induced DSBs (ChIP) n.d. (Ginjala et al., 2011)
CBX4 Chromodomain Methyl-lysine binding SUMO E3 ligase Laser micro-irradiation IR-induced foci SUMOylation of BMI1 Prevents sensitivity to DSB-inducing agents (Chou et al., 2010 Ismail et al., 2012)
CBX6, 7, 8 Chromodomain Methyl-lysine binding Laser micro-irradiation (GFP-tagged protein) n.d. (Chou et al., 2010)
HPH1, 2 SAM and Zinc finger domain Laser micro-irradiation (GFP-tagged protein) n.d. (Chou et al., 2010)
Protein Protein motifs Biochemical activity Method of DNA damage induction Function in DDR References
PRC2EZH1 SET domain Histone methyltransferase Laser micro-irradiation (GFP-tagged protein) Prevent sensitivity to DSB-inducing agents (Chou et al., 2010)
EZH2 SET domain Histone methyltransferase UV Laser micro-irradiation endonuclease-induced DSBs (ChIP) H2O2 induced foci Prevent sensitivity to DSB-inducing agents (Chou et al., 2010 O'Hagan et al., 2008 O'Hagan et al., 2011)
SUZ12 Zinc finger Stimulates histone methyltransferase activity UV Prevent sensitivity to DSB-inducing agents (Chou et al., 2010)
PHF1 (interactor) Two PHD fingers and a Tudor domain Stimulates histone methyltransferase activity Laser micro-irradiation (GFP-tagged protein) Prevent sensitivity to DSB-inducing agents (Hong et al., 2008)
PRC1BMI1/PCGF4 RING domain E3 ubiquitin ligase Laser micro-irradiation hydroxyurea-, camptothecin- and IR-induced foci aphidicolin-induced fragile sites (ChIP) ZNF-induced DSBs (ChIP) H2A/H2AX ubiquitylation Prevent genomic instability Prevent sensitivity to DSB-inducing agents (Chagraoui et al., 2011 Chou et al., 2010 Ginjala et al., 2011 Ismail et al., 2010 Nacerddine et al., 2012 Pan et al., 2011)
MEL18/PCGF2 RING domain E3 ubiquitin ligase Laser micro-irradiation n.d. (Chou et al., 2010 Ginjala et al., 2011)
RING1A/RNF1 RING domain E3 ubiquitin ligase Laser micro-irradiation (GFP-tagged protein) n.d. (Chou et al., 2010)
RING1B/RNF2 RING domain E3 ubiquitin ligase Laser micro-irradiation IR-induced foci H2A/H2AX ubiquitylation Prevent sensitivity to DSB-inducing agents (Chou et al., 2010 Ginjala et al., 2011 Ismail et al., 2010 Wu et al., 2011a)
CBX2 Chromodomain Methyl-lysine binding Laser micro-irradiation ZNF-induced DSBs (ChIP) n.d. (Ginjala et al., 2011)
CBX4 Chromodomain Methyl-lysine binding SUMO E3 ligase Laser micro-irradiation IR-induced foci SUMOylation of BMI1 Prevents sensitivity to DSB-inducing agents (Chou et al., 2010 Ismail et al., 2012)
CBX6, 7, 8 Chromodomain Methyl-lysine binding Laser micro-irradiation (GFP-tagged protein) n.d. (Chou et al., 2010)
HPH1, 2 SAM and Zinc finger domain Laser micro-irradiation (GFP-tagged protein) n.d. (Chou et al., 2010)

BMI1, B lymphoma Mo-MLV insertion region 1 homolog CBX, chromobox ChIP, chromatin immunoprecipitation DSB, DNA double strand break GFP, green fluorescent protein EZH, enhancer of zeste homologue HPH, human polyhomeotic homologue PCGF, Polycomb group RING finger protein PHD finger, plant homeodomain finger PHF1, PHD finger protein 1 PRC Polycomb repressive complex RING, really interesting new gene RNF, RING Finger protein SAM, sterile alpha motif SET domain, Su(var)3-9, Enhancer-of-zeste, Trithorax domain SUMO, small ubiquitin-related modifier Suz12, Suppressor of Zeste homologue 12 ZNF, zinc finger protein. n.d., not determined.

The role of PRC1 and PRC2 at DSBs

Ubiquitylated H2A fulfils a dual function it is a key epigenetic mark for transcriptionally silent chromatin (Cao et al., 2005 Wang et al., 2004), and also acts as a mark of DNA damage (Bergink et al., 2006 Doil et al., 2009 Huen et al., 2007 Ikura et al., 2007 Mailand et al., 2007 Marteijn et al., 2009 Nicassio et al., 2007 Stewart et al., 2009 Zhao et al., 2007). This raises the possibility that the PRC1 E3 ligase is also involved in DDR. Several laboratories confirmed this hypothesis by showing that both BMI1 and RING1B efficiently accumulate at ionizing radiation (IR)- and laser-induced DSBs (Chagraoui et al., 2011 Chou et al., 2010 Facchino et al., 2010 Gieni et al., 2011 Ginjala et al., 2011 Ismail et al., 2010 Nacerddine et al., 2012 Pan et al., 2011 Wu et al., 2011a) (Box 1 and Fig. 3). The kinetics of BMI1 and RING1B recruitment is similar to that of the early DDR factors, such as RNF8, meiotic recombination 11 (MRE11) and Nijmegen breakage syndrome 1 (NBS1) (Ismail et al., 2010), and their presence at DSBs is sustained for several hours after damage (Chou et al., 2010 Ginjala et al., 2011 Ismail et al., 2010). In contrast to BMI1, its homologue MEL18 is only transiently recruited to DSBs, suggesting that recruitment and maintenance of different PRC1 complexes occurs with distinct kinetics (Chou et al., 2010 Ginjala et al., 2011). The other PRC1 subunits, including human polyhomeotic homologue 1 and 2 (HPH1 and HPH2, respectively), as well as different chromodomain-containing proteins, such as CBX2, CBX4, CBX6, CBX7 and CBX8, are also found to localize to laser-induced DSBs (Chou et al., 2010 Ginjala et al., 2011 Ismail et al., 2012). Given the variety in the composition of PRC1 complexes (Gao et al., 2012 Gearhart et al., 2006 Ogawa et al., 2002 Sanchez et al., 2007 Tavares et al., 2012 Trojer et al., 2011 Vandamme et al., 2011 Yu et al., 2012), there is a large number of possible complexes that can be assembled at DSBs, which raises the question whether they perform distinct functions. Interestingly, depending on the experimental system used, in some cases the recruitment of BMI1 and RING1B was found to be independent from each other (Ginjala et al., 2011 Ismail et al., 2010), which suggests that in the absence of RING1B BMI1 can be recruited by RING1A and that in the absence of BMI1 other PCGF factors can recruit RING1B (Chou et al., 2010).

Regulation of PRC1 recruitment and activity at DSBs. ATM kinase activity and H2AX are required for BMI1 recruitment to and its maintenance at DSBs, possibly through RNF8. Independently from ATM, poly(ADP ribosyl)ation by PARP enzymes supports the recruitment of the PRC1 subunits MEL18 and CBX4 to DSBs. The other components of PRC1 have also been detected at sites of DSBs, suggesting that the entire complex is recruited. The precise molecular mechanisms by which the ATM–H2AX–RNF8 pathway and PARP promote PRC1 recruitment are not fully understood. In particular, it is unknown whether components of PRC1 can bind directly to phosphorylated or ubiquitylated substrates. However, it has been suggested that CBX4 directly binds to poly(ADP-ribose), which would provide a mechanism by which PARP enzymes promote PRC1 recruitment. Recruitment of BMI1 to DSBs has been suggested to depend on SUMOylation (Su) by CBX4. Once recruited, BMI1 is involved in H2A and H2AX monoubiquitylation at DSBs. The ubiquitylation activity of BMI1 at DSBs is stimulated by phosphorylation, and one function of BMI1 at DSBs is to promote homologous recombination. Moreover, since H2A ubiquitylation has been implicated in transcriptional silencing of genes that are in direct proximity to the break, we speculate that PRC1 is also involved in this process (as shown by the question mark).

Regulation of PRC1 recruitment and activity at DSBs. ATM kinase activity and H2AX are required for BMI1 recruitment to and its maintenance at DSBs, possibly through RNF8. Independently from ATM, poly(ADP ribosyl)ation by PARP enzymes supports the recruitment of the PRC1 subunits MEL18 and CBX4 to DSBs. The other components of PRC1 have also been detected at sites of DSBs, suggesting that the entire complex is recruited. The precise molecular mechanisms by which the ATM–H2AX–RNF8 pathway and PARP promote PRC1 recruitment are not fully understood. In particular, it is unknown whether components of PRC1 can bind directly to phosphorylated or ubiquitylated substrates. However, it has been suggested that CBX4 directly binds to poly(ADP-ribose), which would provide a mechanism by which PARP enzymes promote PRC1 recruitment. Recruitment of BMI1 to DSBs has been suggested to depend on SUMOylation (Su) by CBX4. Once recruited, BMI1 is involved in H2A and H2AX monoubiquitylation at DSBs. The ubiquitylation activity of BMI1 at DSBs is stimulated by phosphorylation, and one function of BMI1 at DSBs is to promote homologous recombination. Moreover, since H2A ubiquitylation has been implicated in transcriptional silencing of genes that are in direct proximity to the break, we speculate that PRC1 is also involved in this process (as shown by the question mark).

Besides PRC1, the recruitment of PRC2 core components, the PRC2-associated factor PHD finger protein 1 (PHF1), and the H3K27 methyl mark have also been detected at sites of DNA damage (Chou et al., 2010 Hong et al., 2008 O'Hagan et al., 2008 Seiler et al., 2011). This suggests that the entire PRC2 complex is present at DSBs.

The role of PRC1 in histone ubiquitylation and signaling at DSBs

As discussed above, the PRC1 E3 ligase is recruited to DSBs, which are main sites of ubiquitylation. These findings prompt the question whether PRC1 contributes to ubiquitylation at DSBs. Determining the contribution of PRC1 to overall ubiquitylation by immunofluorescence with an antibody that recognizes ubiquitin conjugates (FK2) has proven controversial (Ginjala et al., 2011 Ismail et al., 2010), most probably because of the abundance of ubiquitin conjugates at DSBs and the limitations of this approach in measuring quantitative differences. The role of RING1B and BMI1 in DSB-induced histone ubiquitylation has been addressed specifically, both biochemically and by using immunofluorescence. Immunoblot analysis of H2A and H2AX suggests that loss of RING1B or BMI1, or mutation of the canonical target lysine of H2A or H2AX interfere with both basal and IR-induced monoubiquitylation (Ginjala et al., 2011 Ikura et al., 2007 Pan et al., 2011 Wu et al., 2011a). In addition, ubiquitylation of H2A – as detected using the antibody E6C5 – at laser-induced damage is found to depend on BMI1 (Ginjala et al., 2011). Although the specificity of this antibody is not undisputed, collectively these findings support a role for BMI1 in H2A monoubiquitylation at the chromatin that surrounds the DNA lesion (Fig. 3).

DSB-induced ubiquitin signaling entails a complex network of ubiquitin substrates, which are mostly regulated by RNF8–RNF168 (Al-Hakim et al., 2010 Lukas et al., 2011 Tang and Greenberg, 2010). The reported requirement of BMI1 for the monoubiquitylation of H2A and H2AX (Ginjala et al., 2011 Pan et al., 2011) and that of RNF8–RNF168 for the generation of K63-linked ubiquitin chains (Doil et al., 2009 Pinato et al., 2009 Stewart et al., 2009) suggest that these ubiquitin signals have distinct functions at DSBs and it will be of great interest to characterize these functions. Other important unanswered questions include: which lysine(s) on H2A or H2AX are modified by RNF8–RNF168 and do they overlap with the PRC1 target lysine? Do PRC1 and RNF8–RNF168 modify histone H2A molecules in the same nucleosome? Can they act on the same ubiquitin chains, and are there other ubiquitylation substrates that are targeted by these E3 ligases?

Another crucial question is whether PRC-complex-mediated histone modifications coordinate DDR signaling. As mentioned above, 53BP1 and also UIMC1 (also known as RAP80) are important DDR mediators, whose recruitment is dependent on the RNF8–RNF168–ubiquitin pathway (Al-Hakim et al., 2010 Lukas et al., 2011 Tang and Greenberg, 2010). Analysis of Bmi1-knockout cells shows that RAP80, as well as 53BP1, efficiently accumulate at DNA damage sites that were generated by using the UV-laser scissor approach (Box 1), suggesting that BMI1-mediated ubiquitylation of H2A does not affect the canonical ubiquitin pathway but, instead, impacts on a parallel pathway that involves the recruitment of yet unidentified factors (Ginjala et al., 2011). However, Ismail et al. reached the opposite conclusion using a different method to induce DSBs (the near-infrared laser-based approach see Box 1) and assessing recruitment at earlier time points, they found that Bmi1 loss affects accumulation of 53BP1, RAP80 and BRCA1 at DNA damage sites (Ismail et al., 2010). Detailed time course experiments following induction of specified DNA damage are needed to clarify the exact contribution of BMI1 to DSB signaling.

Several methods can be used to induce localized DNA lesions in the nucleus of mammaliancells (Nagy and Soutoglou, 2009 Polo and Jackson, 2011). Laser lines are used to locally irradiate specific regions in the cell nucleus, thereby generating focal DNA damage. PcG protein recruitment to DNA lesions has been studied by using micro-irradiation with an UV-A laser (337 nm) on cells that were sensitized with halogenated nucleotide analogues such as BrdU or IdU (referred to as the UV-A laser scissor approach) (Chou et al., 2010 Ginjala et al., 2011 Nacerddine et al., 2012), and with a near-infrared laser line (750 nm) on cells sensitized with the DNA-intercalating dye Hoechst 33258 (Ismail et al., 2010 Ismail et al., 2012). A limitation of these laser-based methods is that, besides DSBs, they also give rise to a wide spectrum of other DNA lesions, including cyclobutane pyrimidine dimers, 6,4 pyrimidine-pyrimidones and ssDNA breaks (Bekker-Jensen et al., 2006 Dinant et al., 2007 Kong et al., 2009). Furthermore, structural changes in the DNA that are caused by intercalation of Hoechst dye may lead to non-physiological DDR (Dinant et al., 2007 Kong et al., 2009 Williams et al., 2007a). UV-A treatment, if applied at high power, can also elicit aberrant cellular responses by inducing protein damage or protein–protein and protein–DNA crosslinks. Thus, the type and relative contribution of DNA lesions need to be considered when comparing different studies, as they may affect the dynamics of DDR factor recruitment.

Nuclease-based systems are used to induce DSBs at defined genomic loci and are based on the use of endonucleases that recognize and cut highly specific DNA sequences. An inducible DSB-inducing system that uses endonuclease I-SceI (Rouet et al., 1994), as well as a zinc-finger nuclease (ZFN) strategy (Miller et al., 2007), has been employed to investigate the recruitment of PcG factors to DSBs (Ginjala et al., 2011 O'Hagan et al., 2008).

Requirements of PRC1 and PRC2 for survival following clastogen treatment and for DSB repair

Work from several laboratories established that PcG proteins have functional relevance in DDR and in genome maintenance. Indeed, loss of components of PRC1 (Chagraoui et al., 2011 Ginjala et al., 2011 Ismail et al., 2010 Pan et al., 2011) and PRC2 (Chou et al., 2010 Hong et al., 2008) leads to increased sensitivity to IR and other clastogens (Table 1). Several authors examined a role for PcG proteins in DSB repair. In particular, experiments in which a GFP-based reporter assay was used to detect homologous recombination efficiency, support a contributing role of BMI1 in the homologous recombination repair pathway (Chagraoui et al., 2011 Ginjala et al., 2011 Nacerddine et al., 2012) (Fig. 3). Regardless of the exact underlying mechanisms, this provides an at least partial explanation for the requirement of BMI1 for cell survival upon treatment with DSB inducing agents (Chagraoui et al., 2011 Ginjala et al., 2011 Ismail et al., 2010 Pan et al., 2011). However, it should be noted that the effects on both homologous recombination and cell survival are relatively mild, which suggests that the PcG factors are not core repair factors, but rather have a modulatory role in the establishment of conditions that favour repair. The potential involvement of PcG proteins in NHEJ awaits further investigation.

Regulation of PRC1 recruitment to sites of DSBs

The studies presented above support a broad involvement of PcG proteins in DDR but, nevertheless, raise several questions with regard to the underlying molecular mechanisms. For example, it remains to be determined which signaling events are required to induce the accumulation of PcG complexes at DSBs. With regard to PRC1, it has been shown that BMI1 recruitment at laser-induced DNA damage sites is dependent on ATM- and ATR-mediated signaling (Fig. 3) (Ginjala et al., 2011 Ismail et al., 2010). In response to ATM activation, a very early recruitment of BMI1 but not its persistence could be detected in mouse H2ax-knockout cells (Ginjala et al., 2011 Ismail et al., 2010). This suggests a biphasic mode of BMI1 accumulation, with an initial phase that is independent of H2AX, and a second phase that depends on ATM- or ATR-mediated phosphorylation and, possibly, ubiquitylation by RNF8 (Ginjala et al., 2011 Ismail et al., 2010) (Fig. 3). The DDR regulatory factors BRCA1 and 53BP1, which act further downstream in the ATM-RNF8–RNF168 pathway, were shown to be dispensable for BMI1 recruitment (Ginjala et al., 2011 Ismail et al., 2010).

In search for additional factors that influence the recruitment of PcG proteins to DNA damage sites, PARP-dependent signaling has also been investigated (Chou et al., 2010 Ginjala et al., 2011 Ismail et al., 2012). PARP inhibition or loss was reported to affect the recruitment of CBX4 (Ismail et al., 2012) and of the BMI1 homologue MEL18, but not of BMI1 itself (Chou et al., 2010 Ginjala et al., 2011) (Fig. 3). Taken together, the data discussed above suggest that multiple signaling pathways contribute to the recruitment of PcG proteins to DNA lesions (Fig. 3). In this context, it will be important to further elucidate the functional crosstalk between PcG proteins and the RNF8–RNF168–ubiquitin pathway, as well as the role of PARP-mediated signaling.

To dissect the molecular mechanism in detail it will be necessary to identify the molecular interactions that allow the recruitment of PcG complexes to damaged DNA, as PcG proteins do not have obvious binding domains for phosphorylated or ubiquitylated proteins. The observation that the H3K27Me3 mark has been detected at DSBs (O'Hagan et al., 2008) suggests that PRC2, to some extent, promotes PRC1 binding at these sites. However, against this hypothesis, recent experiments suggest that PRC1 recruitment to DSBs is not affected upon depletion of EZH2 or embryonic ectoderm development (EED) (Ismail et al., 2010), which are required for H3K27 trimethylation (Cao et al., 2002 Czermin et al., 2002 Kuzmichev et al., 2002 Müller et al., 2002). In addition, methyl-lysine binding by CBX4 appeared to be dispensable for recruitment to sites of microirradiation (Ismail et al., 2012).

Finally, recent data indicate that the function of BMI1 in DDR is also regulated by post-translational modification (Fig. 3). For example, modification of BMI1 with SUMO by CBX4 has been suggested to favor its recruitment to DSBs (Ismail et al., 2012). In addition, phosphorylation of BMI1 stimulates the ubiquitylation activity of PRC1 at DSBs (Nacerddine et al., 2012).

Taken together, is it clear that PcG proteins are required for cell survival upon the induction of DSBs and are physically recruited to DSBs, where BMI1 promotes histone H2A monoubiquitylation and efficient DDR. There are at least two ways by which PcG complexes might promote DSB repair. First, PcG proteins might contribute to DNA damage site recognition and, as we have discussed above, help to recruit DDR factors to the chromatin surrounding the break. Second, given that the PRC1-induced ubiquitylation of H2A acts as a repressive mark, an attractive means by which PcG proteins might facilitate DSB repair is by simply repressing transcription at the break, thereby coordinating the repair and transcriptional machineries (Chagraoui et al., 2011 Lukas et al., 2011). In the next section, we will discuss this possibility in detail.


IV. Plant hormones

Hormones affect almost every aspect of plant development and physiology, including anthocyanin biosynthesis. The signaling pathway from hormone perception to anthocyanin activation or inhibition is relatively clear for jasmonate, auxin, strigolactone, and gibberellic acid (Fig. 4). As will be elaborated on below, the basic logic of the signaling mechanisms for these four different hormones is remarkably similar. They all involve a Skp1/Cullin/F-box (SCF) E3 ubiquitin ligase complex that targets a protein substrate for poly-ubiquitination and degradation by the 26S proteasome. What distinguishes each signaling pathway are the specific hormone receptors and the targets of the F-box protein (Fig. 4). Other plant hormones, including ethylene, abscisic acid, and cytokinins, are also known to affect anthocyanin biosynthesis (Loreti et al., 2008 Das et al., 2012a ). However, whether they have a positive or negative effect on anthocyanin accumulation seems to vary from species to species (Loreti et al., 2008 ), and the signaling mechanisms that relay the hormone perception to anthocyanin biosynthesis are still poorly understood for these hormones.

1. The SCF COI1 -JAZ module and jasmonate response

Jasmonates (e.g. jasmonic acid (JA) and methyl jasmonate (MeJA)) are the sweet-smelling hormones released by plants undergoing environmental challenges, such as herbivory. The bioactive jasmonyl-L-isoleucine (JA-Ile) conjugate mediates the physical interaction between jasmonate zim-domain (JAZ) proteins and the F-box protein CORONATINE-INSENSITIVE PROTEINI (COI1), a component of the Skp1/Cullin/F-box (SCF COI1 ) complex that flags JAZ proteins for ubiquitination and subsequent degradation (reviewed in Pauwels & Goossens, 2011 ).

Among all plant hormones, the effect of jasmonates on anthocyanin biosynthesis is the least contentious: exogenous application of MeJA induces anthocyanin production in various plants (e.g. Tamari et al., 1995 Shan et al., 2009 ). In Arabidopsis, this is because the JAZ proteins interfere with the anthocyanin-activating MBW complex by directly binding the R2R3-MYB and the bHLH components. Methyl jasmonate application triggers the degradation of the JAZ proteins and thus the release of the R2R3-MYB and the bHLH proteins to reform the anthocyanin-activating MBW complex (Fig. 4) (Shan et al., 2009 Qi et al., 2011 ). The same mechanism has been shown to regulate anthocyanin biosynthesis in response to MeJA in apple (An et al., 2014 ) and presumably is conserved among other plants as well.

2. The SCF TIR1 -Aux/IAA-ARF module and auxin response

Early experiments in Brassica found that exogenous application of the auxin indole acetic acid (IAA) inhibits anthocyanin in a dose-dependent manner (Kang & Burg, 1973 ). Later studies in various species, mostly using callus culture, showed the same general trend – that high auxin concentration inhibits anthocyanin biosynthesis (Makunga et al., 1997 Jeong et al., 2004 Zhou et al., 2008 Liu et al., 2014 Ji et al., 2015 Y. Wang et al., 2018 but see Mori et al., 1994 ). A study of red-fleshed apple calli (Y. Wang et al., 2018 ) revealed that an Auxin Response Factor, MdARF13, represses anthocyanin biosynthesis both by directly binding the promoter of the ABP gene MdDFR and repressing its transcription, and by physically interacting with the subgroup 6 R2R3-MYB activator MdMYB10 and destabilizing the MBW complex. Under low auxin concentration, the auxin/indole-3-acetic acid (Aux/IAA) repressor MdIAA121 interacts with MdARF13 and prevents it from binding the MdDFR promoter or interacting with MdMYB10. Exogenous auxin application causes MdIAA121 degradation by the 26S proteasome (Y. Wang et al., 2018 ), presumably through mediating SCF TIR1 -Aux/IAA interactions, and subsequently MdARF13 is released from Aux/IAA to exert its repressive function on anthocyanin biosynthesis (Fig. 4). While callus culture provides a convenient platform for the study of hormone response, whether this paradigmatic model of auxin response also explains anthocyanin response to auxin in whole plants remains to be tested.

3. The D14-SCF D3 -D53 module and strigolactone response

Strigolactones (SLs) are carotenoid-derived plant hormones that have been shown to enhance anthocyanin accumulation (L. Wang et al., 2020 ). The SL signaling pathway was not characterized until very recently. With the presence of SL, the receptor protein DWARF14 (D14) undergoes a conformational change to facilitate its interaction with the F-box protein D3, a component of the SCF D3 complex, and the DWARF53 (D53) repressor, leading to ubiquitination and degradation of D53 (Jiang et al., 2013 Zhou et al., 2013 Yao et al., 2016 ). The homolog of D53 in Arabidopsis, AtSMXL6, was recently shown to repress transcription of AtPAP1 by directly binding to its promoter (L. Wang et al., 2020 ). Hence, SLs trigger degradation of D53/AtSMXL6, de-repressing the subgroup 6 R2R3-MYB genes and activating anthocyanin biosynthesis (Fig. 4).

4. The GID1-SCF GID2 -DELLA module and gibberellic acid response

Gibberellic acids (GA) have been reported to have both enhancing (Weiss et al., 1995 Hosokawa, 1999 ) and inhibiting effects (Kim et al., 2006 Jiang et al., 2007 Xie et al., 2016 ) on anthocyanin biosynthesis, depending on species or tissue types. In plant cells, GA is perceived by the receptor GID1 (GA-INSENSITIVE DWARF1). Gibberellic acid perception promotes the interaction between GID1 and the DELLA proteins, triggering a conformational change that enhances the binding affinity of the GID1-DELLA complex to the SCF SLY1/GID2 complex and leads to the proteasome-mediated degradation of DELLA (reviewed in Davière et al., 2008 ).

In Arabidopsis, the DELLA proteins are positive regulators of anthocyanin biosynthesis, as they directly interact with and sequester the AtMYBL2 and AtJAZ repressors, resulting in higher MBW complex activities (Xie et al., 2016 ). Gibberellic acid degrades DELLA and hence negatively regulates anthocyanin biosynthesis. Conversely, in another study it was shown that the DELLA proteins can also directly interact with the R2R3-MYB and bHLH components of the MBW complex, thereby destabilizing the activation complex (Qi et al., 2014 ). As such, DELLA proteins can theoretically be negative regulators of anthocyanin biosynthesis and GA can potentially enhance anthocyanin production. This offers a plausible explanation for the seemingly contradicting observations on the effect of GA in different species: perhaps it is the different DELLA-interacting proteins in different species or tissue types that determine whether GA has an enhancing or inhibiting effect on anthocyanin biosynthesis (Fig. 4).

Notably, both phosphate deficiency and sucrose induced anthocyanin accumulation in Arabidopsis seem to depend on the GA–DELLA pathway (Jiang et al., 2007 Li et al., 2014 ). Phosphate starvation was shown to reduce the concentration of bioactive GA, leading to the accumulation of DELLA proteins and enhanced anthocyanin production (Jiang et al., 2007 ). Sucrose was shown to specifically inhibit the GA-mediated degradation of DELLA proteins, also leading to the stabilization of DELLA proteins and enhanced anthocyanin accumulation (Li et al., 2014 ).


DNA Question: Why do introns exist?

Not sure if this is the right thread, but I was reading some biology literature and:

DNA -> transcription -> RNA -> Translation -> Aminos/Protiens

And somewhere in the process, (I believe transcription), in eukaryotes, the introns need to be removed before translation, or a different amino acid chain will be created.

Is there a reason for the existence of introns? Or is it just an observation of nature

I did my PhD on splicing and introns! This is such a great discussion :)

Quick brief definitions: intron (mostly non-coding RNA), exon (mostly coding RNA), isoform (the same gene, spliced in different ways)

The benefit to splicing (the removal of introns) is to create more genetic diversity. It was once thought that the number of genes correlated with organism complexity. But it turns out that some worms have more genes than we do!

However, when introns are removed, sometimes exons are also removed. Additionally, sometimes little pieces of exons are removed with the introns. Also, sometimes introns can be kept! The same gene can be spliced into several different ways. This is called alternative splicing. For example: Gene A may skip exon 3 sometimes, keep half of exon 2 other times, keep intron 7 sometimes. Then those three isoforms go to the cytoplasm and makes 3 similar but different proteins. Sometimes these proteins can have totally different purposes and functions.

As it turns out, complexity DOES correlate with the number of splicing variants (isoforms). Humans do A LOT of splicing.

Main purposes for making different variations of the same gene: Sometimes different cell types (kidney cells versus brain cells) will make two different isoforms. Developing embryos have an entirely different set of isoforms that are never used again once development is done. Cells can also change what isoforms are being produced in response to cellular signals like stress.


How Do Tigers Get Their Stripes? Science Not So Certain Now

A decades-old explanation for how tigers get their stripes has come into question as researchers challenge what’s called the morphogen theory. The research does not nix the theory, but science may now have a hypothetical tiger by the tail as they try to figure out this aspect of how Nature works.

The morphogen theory posits that proteins controlling traits are arranged as gradients, with different amounts of proteins activating genes to create specified physical features.

This theory was first put forth in the 1950s by mathematician and World War II code breaker Alan Turing and refined in the 1960s by Lewis Wolpert. It has been used to explain why a tiger has stripes, among other phenomena.

But some biologists have raised questions about the theory, which contends that physical features are necessarily tied to absolute concentrations of proteins within the morphogen gradient.

If a certain critical mass of protein is present, then a given physical feature&mdashfor example, cells that make the skin on your forehead&mdashwill appear. If less than that critical mass is present, a different structure&mdashsay, the skin that makes your eyebrows&mdashwill appear, and a boundary will be formed between the two structures.

Alternative views have suggested physical features are not necessarily the result of a specified number of proteins, but, rather, come from more complex interactions between multiple gradients that work against one another.

New York University biologists explored this process by studying the fruit fly Drosophila, a powerful model for studying genetic development as it is amenable to precise genetic manipulations. They focused on one protein, Bicoid (Bcd), which is expressed in a gradient with highest levels at the end of the embryo that will become the mature fly’s head.

The researchers, headed by Stephen Small, chair of NYU’s biology department, examined a large number of target genes that are directly activated by Bcd. Each target gene is expressed in a region of the embryo with a boundary that corresponds to a specific structure.

By examining DNA sequences associated with these target genes, the researchers discovered binding sites for three other proteins&mdashRunt, Capicua, and Kruppel&mdashwhich all act as repressors. All three proteins are expressed in gradients with highest levels in the middle part of the embryo, and thus are positioned in exactly the opposite orientation compared to the Bcd activation gradient.

By changing the spatial distribution of the repressors and by manipulating their binding sites, Small and his colleagues showed that these repressors antagonize Bcd-dependent activation and are absolutely critical for establishing the correct order of boundaries that are found in a normal embryo.

In other words, contrary to Turing’s theory, a single gradient of proteins does not have sufficient power to form the same body plan in each member of a species however, if there are multiple gradients that work against each other, then the system becomes robust enough for normal development.

While the results, reported in the journal Cell, raise questions about morphogen theory, the researchers explained that their findings did not &ldquofalsify&rdquo it, but, rather, suggested it needed some additional refinement.


The A, B, Z's of DNA

April 25th (4/25) is national DNA day. Digital World Biology™ celebrates by sharing some of our favorite DNA structures. We created these photos with Molecule World™ Molecule World is a tools for exploring molecular and chemical structures on an iPhone or iPad.

Update: These same structures can also be viewed on the iPhone with Molecule World for iPhone. Visit the iTunes app store to download either app and explore these DNA structures on your phone or iPad.

Update: If you have Molecule World on an iPad, you can download a collection of these structures in Molecule World and explore them while you read the post. Go to our new DNA Exploration Collection. Download the file named DNA_exploration.mwc and open it in Molecule World on an iPad. Alternatively, you can download the structures below and open them in Molecule World on an iPhone.

As we are taught in school, the double stranded DNA molecule is a right-handed helix as determined by Watson and Crick using Franklin's x-ray diffraction images [1]. This B-form of DNA has approximately 10 nucleotides per turn of the helix and is the most common form of DNA found in nature.

Classic structure with the elements colored .

Classic structure with the bases colored.

Classic structure with the strands colored.

However, we've heard it said that the first crystal structure of DNA was not right-handed (pers. communication S. Elgin, Washington University). Instead, the high salt and GC base-pairs, used to form the DNA crystals caused the helix to twist in a left-handed way, creating a structure called Z-DNA. Z-DNA occurs in nature, but is most frequently used by marketing departments to [incorrectly] create company logos.

Z-DNA with the elements colored.

Z-DNA with the bases colored.

Z-DNA with the stands colored.

In addition to B- and Z-DNA, DNA can exist in another form known as A-DNA. A-DNA occurs when DNA is dehydrated, but also in DNA/RNA hybrids and double stranded RNA.

A-DNA with the elements colored.

A-DNA with the bases colored.

A-DNA with the strands colored.

Here's another look at the same molecules, this time head on with the structures rendered in tubes and the bases colored.

B-DNA "on end," showing the double helix.

Z-DNA "on end," showing the double helix.

A-DNA "on end," showing the double helix.

Double Helix- Watson, J. D., & Crick, F. H. (1953). Molecular structure of nucleic acids a structure for deoxyribose nucleic acid. Nature, 171(4356), 737-8.

B-DNA Structure (PDB:1BNA)- Drew, H., Wing, R., Takano, T., Broka, C., Tanaka, S., Itakura, K., & Dickerson, R. (1981). Structure of a B-DNA dodecamer: conformation and dynamics. Proceedings of the National Academy of Sciences, 78(4), 2179-2183 DOI:10.1073/pnas.78.4.2179

First Z-DNA Crystal- Wang, A., Quigley, G., Kolpak, F., Crawford, J., van Boom, J., van der Marel, G., & Rich, A. (1979). Molecular structure of a left-handed double helical DNA fragment at atomic resolution Nature, 282(5740), 680-686 DOI:10.1038/282680a0

Z-DNA Structure (PDB:1D48)- Egli, M., Williams, L., Gao, Q., & Rich, A. (1991). Structure of the pure-spermine form of Z-DNA (magnesium free) at 1-.ANG. resolution Biochemistry, 30(48), 11388-11402 DOI:10.1021/bi00112a005

A-DNA Structure (PDB:371D)- Fernandez, L., Subirana, J., Verdaguer, N., Pyshnyi, D., Campos, L., & Malinina, L. (1997). Structural Variability of A-DNA in Crystals of the Octamer d(pCpCpCpGpCpGpGpG) Journal of Biomolecular Structure and Dynamics, 15(1), 151-163 DOI:10.1080/07391102.1997.10508954

Molecule World™ was developed with funding from the National Science Foundation (SBIR IIP1315426). Any opinions, findings, conclusions, or recommendations expressed on this website are those of the authors and do not necessarily represent the official views, opinions, or policy of the National Science Foundation.


Repressors as modulators: mechanisms of action

As highlighted above, genome-wide analyses in yeast and in metazoans have demonstrated that repressors associate with actively transcribed loci and that their absence is associated with both increases and decreases of transcriptional output at different genes. These findings contribute to a body of compelling, yet circumstantial, evidence that repressors can play a role in activation of gene expression. The first convincing demonstration that repressor-mediated activation can regulate specific groups of genes came from work in S. cerevisiae. Indeed, the initial description of the Sin3 co-repressor complex identified it as a protein with dual functions: acting as both an activator and a repressor in vivo (Nawaz et al., 1994 Vidal and Gaber, 1991 Vidal et al., 1991). Other studies focused on the specific regulation of individual genes by a variety of repressor complexes, and these studies have perhaps been the most powerful in changing the orthodox view that repressor complexes merely inhibit transcription (summarised in Box 1). Such studies have identified clear roles for so-called repressors in activating gene expression, but how might these be mediating both activating and repressive functions? Below, we summarise existing evidence for how repressors function at actively transcribed genes.

Box 1. Repressors as activators?

The association of repressor complexes with sites of active transcription does not necessarily mean that they activate transcription, but a number of detailed studies confirm that, at least in some instances, repressors can and do activate transcription:

Lysine deacetylase activity promotes transcription of galactose- and inositol-responsive genes in S. cerevisiae (Wang et al., 2002).

Transcription induced by either osmotic or heat stress in S. cerevisiae requires the Sin3-Rpd3 KDAC complex for transcriptional activation to occur (De Nadal et al., 2004).

Mi-2 is required for activation of heat-shock genes in fruit flies (Murawska et al., 2011).

Hdac1 promotes expression of a subset of neural-specific genes in zebrafish (Harrison et al., 2011).

The NuRD complex mediates transcriptional activation during erythropoiesis in mice (Miccio et al., 2010).

The RNA polymerase connection

In many cases, mapping protein distribution by ChIP shows that repressor complex occupancy can extend to differing extents throughout the body of a gene (Johnsson et al., 2009 Joshi and Struhl, 2005 Kurdistani et al., 2002 Mathieu et al., 2012 Miccio et al., 2010 Morey et al., 2008 Murawska et al., 2011 Murawska et al., 2008 Reynolds et al., 2012a Reynolds et al., 2012b Wang et al., 2002 Wang et al., 2009). This is in contrast to the more familiar pattern seen for sequence-specific transcription factors, which generally localise tightly to the promoter region of regulated genes. Consistent with a broader association across gene loci, loss of KDAC activity through mutation or chemical inhibition can result in increased acetylation of promoters, which often spreads well into coding regions (Johnsson et al., 2009 Joshi and Struhl, 2005 Keogh et al., 2005 Li et al., 2007b Reid et al., 2004 Wang et al., 2002 Wirén et al., 2005). What factors might influence such a broad range of protein association? Often, the distribution of transcriptional repressors across genes or chromosomes closely resembles that of the transcribing RNA polymerase, suggestive of an interaction between them (Brookes et al., 2012 Mathieu et al., 2012 Murawska et al., 2008 Srinivasan et al., 2005).

Interactions between KDAC-containing repressor complexes and elongating RNA polymerase are a well-described phenomenon in yeast. At actively transcribed genes in S. cerevisiae, lysine deacetylase activity, in the form of the Rpd3C(S) protein complex, has been found to be recruited to gene bodies via association with dimethylated H3K36. This specific association is mediated by the Rpd3C(S) component proteins Eaf3 and Rco1. The consequence of this association is the deacetylation of histones H3 and H4 throughout the bodies of transcribed genes, which somewhat reduces elongation efficiency but, importantly, suppresses intragenic transcription initiation (Carrozza et al., 2005 Joshi and Struhl, 2005 Keogh et al., 2005 Li et al., 2007a Li et al., 2007b Li et al., 2009) (Fig. 1). More recent work has indicated that Rpd3C(S) can also be recruited via a direct interaction with the elongating form of RNA polymerase (Drouin et al., 2010 Govind et al., 2010).