SUB1 is required for efficient light induction of genes

We analyzed expression of SUB1 in the dark and light to confirm that sub1 is a direct target of WCC as shown previously [19,30]. When Neurospora was grown in constant darkness SUB1 was rhythmically expressed (S1A Fig.). Basal expression levels of SUB1 were essentially independent of WCC but upon light exposure expression of SUB1 was rapidly induced in a WCC-dependent manner (S1B Fig.). The data demonstrate that sub1 is a light-induced and clock-controlled gene.

To investigate a possible crosstalk of the GATA-family TFs SUB1 and WCC we analyzed by RNA-seq the transcriptomes of wt and Δsub1 strains grown in the dark and after light-exposure. We identified 519 light-inducible genes in wt (S1C Fig., S1 Table). 319 of these genes were also identified recently in a similar analysis [31]. More than 80% of the light-inducible genes (417) identified in our study responded rapidly to the light cue (>2x induction within 30 min), strongly suggesting that these are immediate early genes directly controlled by WCC. About 50% of the transcripts accumulated to maximal levels after 30 min and the RNA levels decreased subsequently (early genes). The remainder of the transcripts reached peak expression levels somewhat later (late genes). Light-induction of a substantial subset of genes (189) was severely impaired in Δsub1 (Fig. 1A). Both early and late accumulating light-induced transcripts were affected in Δsub1. The temporal transcription dynamics of SUB1-affected genes corresponds to the transient activity profile of light-activated WCC suggesting that SUB1 may cooperate with WCC rather than acting independently and downstream of WCC. In addition, deletion of sub1 affected expression of 593 genes that were not induced by light, indicating a major role of SUB1 beyond regulation of light-induced transcription (S1D Fig.).

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Fig 1. SUB1 is required for efficient recruitment of WCC and light-induction of a subset of genes.

A. Heat-maps showing the 189 light-inducible genes with significantly lower light-induced RNA levels in Δsub1 compared to wt. B. Quantification of light-induced RNA levels of rds1 and hyr1 by RT-PCR (± SEM, n = 4). 28s rRNA was used for normalization. wt RNA levels in dark were normalized to 1. C. ChIP-PCR analysis of SUB1 showing binding of SUB1 to the rds1 (upper panel) and hyr1 promoters (lower panel) determined by two-step FLAG-HIS ChIP. wt₉₇₁₈ was used as a control. 28s rDNA was used for normalization. SUB1 binding at t = 60 min was set to 1 (± SEM, n = 3). D. ChIP-PCR analysis of WC2 showing light-induced binding of the WCC to the rds1 (upper panel) and hyr1 promoters (lower panel) in wt and Δsub1 strains. 28s rDNA was used for normalization. WC2 ChIP at t = 60 min in wt was set to 1 (± SEM, n = 4).

https://doi.org/10.1371/journal.pgen.1005105.g001

We then analyzed by qPCR the SUB1-dependent regulation of the light-inducible genes rds1 and hyr1, which harbor a WCC binding site in their promoters [19]. Expression of both genes was rapidly induced by light and the light-induction was attenuated in a Δsub1 strain (Fig. 1B). In contrast, light-induction of the WCC-dependent vvd gene was not significantly affected by SUB1 (S1E Fig.). To assess the SUB1-dependence of light-induced transcription initiation at the rds1 and hyr1 promoters we analyzed the recruitment kinetics of RNA polymerase II (RNAPII) in wt and Δsub1 strains. SUB1-dependent recruitment of Ser5 phosphorylated RNAPII, which is indicative of transcription initiation, was already detected 5 min after light-induction (S1F Fig.), i.e. prior to the light-induced accumulation of newly synthesized SUB1 (S1G Fig.). The data indicate that the previously synthesized, old SUB1 cooperates with light-activated WCC to activate transcription at the rds1 and hyr1 promoters. Moreover, we analyzed recently published ChIP-seq data of recruitment kinetics of RNAPII in response to a single 1 min light-pulse [32]. The data revealed that 40% of the SUB1-dependent light-inducible genes and 43% of the SUB1-independent genes showed a rapid (5–10 min) and transient increase in RNAPII occupancy (S1H Fig), suggesting that these light-inducible target genes of SUB1 are directly activated by WCC. The rest of SUB1-dependent (60%) and independent (57%) light-inducible genes did not show significant RNAPII recruitment under these conditions (1 min light-pulse), suggesting that longer periods of light exposure are required for their maximal activation.

SUB1 binding to promoters of hyr1 and rds1 enhances WCC recruitment

Since SUB1 and WCC contain GATA-family DNA-binding domains we asked if these TFs bind to the same sites in the hyr1 and rds1 promoters. We therefore constructed by gene replacement a strain expressing a FLAG-HIS tagged version of SUB1 and performed tandem chromatin immunoprecipitation (ChIP) [33]. In dark-grown mycelia SUB1_FLAG-HIS was detected at a low level at the WCC binding sites in the rds1 and hyr1 promoters (Fig. 1C). Recruitment of SUB1_FLAG-HIS was enhanced after light exposure of mycelia, suggesting that light-activated WCC facilitates binding of SUB1 directly and/or indirectly by supporting light-induced expression of new SUB1.

To assess if SUB1 binding also affects recruitment of WCC we analyzed binding of WCC in wt and Δsub1 strains. Light-induced binding of WCC to the rds1 and hyr1 promoters was attenuated in a Δsub1 strain (Fig. 1D), indicating that SUB1 supports recruitment of light-activated WCC to these promoters. Hence, SUB1 and WCC mutually facilitated their recruitment to overlapping or nearby binding sites in the rds1 and hyr1 promoters.

WCC binds preferentially to tandem GATC motifs

We next determined by ChIP-seq the binding sites of WCC on a genome-wide scale. We, in collaboration with others, had previously identified binding sites of light-activated WCC on the basis of a ChIP-seq analysis with rather low sequence coverage [19]. Here, we determined WCC binding sites using two further independent ChIP-seq approaches. In the first approach we fragmented chromatin by sonification and identified 466 light-inducible putative WCC binding sites (S2A Fig., S2 Table) by tandem affinity ChIP of a TAP-tagged WC2 [34]. In the second approach, chromatin was gently fragmented by MNase digestion and we identified 218 putative binding sites of WCC by ChIP with WC2 antibodies (S2B Fig., S2 Table). The combined analyses revealed 582 light-dependent putative WCC binding sites with 92 highly confident binding sites that were identified by both approaches (Fig. 2A, S2 Table).

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Fig 2. Cistrome analysis of WCC and SUB1.

A. Heat-map showing the light-induced WCC occupancy at 92 binding sites identified by both, MNase-WC2 ChIP-seq and TAP-WC2 ChIP-seq. 5 kb region covering the binding sites are shown. Left panel: WCC binding in the dark. Right panel: WCC binding 30 min after light-exposure. B. Occurrence of tandem GATC motifs with the indicated spacing at WCC binding sites. 300 bp DNA regions covering the peaks of 92 highly confident WCC binding sites were analyzed. The dashed line corresponds to the occurrence of tandem GATC motifs in a set of randomly chosen 300 bp regions. C. Potential light response elements (LREs) at WCC binding sites contain multiple GATC motifs. GATC motifs in WCC binding sites of the indicated genes are shown. frq_as: frq antisense [19]. vvd_prox and vvd_dis: proximal and distal WCC binding sites in vvd promoter (S2 Table). D. Distribution of tandem GATC motifs with < 30 bp spacing at WCC binding sites. The grey area represents the sequence coverage of the WCC ChIP (MNase-WC2 ChIP, 30 min) at the highly confident 92 WCC binding sites. The red line shows the occurrence of tandem GATC motifs. E. Heat-map showing the SUB1 occupancy at binding sites. Left panel: SUB1 binding in the dark. Right panel: SUB1 binding 30 min after light-exposure. F. SUB1 binding motifs identified by MEME are shown. The major sequence motif shown in the upper panel is found in 171 sites. The GTA-rich motifs shown in the lower left and right panels are present in 82 and 63 sites, respectively. G. Distribution of the major SUB1 binding motif (a/cGAT-x6-a/cTGc/t) at SUB1 binding sites. The grey area represents the sequence coverage of the SUB1 ChIP (SUB1 30 min) at 617 SUB1 binding sites. The red line shows the occurrence of the SUB1 binding motif.

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Analysis of binding motifs by MEME [35] revealed a GATC-containing consensus motif (S2C Fig.), similar to previously identified WCC binding motifs [10,15,19]. Further analysis revealed that WCC sites were enriched in tandem GATC motifs with a preferential pairwise spacing of 10–30 bp (Fig. 2B, C and D). This arrangement of GATC motifs may reflect that light-activated WCC is a dimer of two WC1/WC2 protomers [21] and thus can potentially bind up to four GATC motifs.

The 92 highly confident WCC binding sites were associated with 91 genes and 51 of these genes were light-inducible. This enrichment of light-inducible genes suggests that these binding sites are functionally relevant. 41 WCC binding sites contain a consensus tandem GATC motif. Binding of WCC to 30 of these sites correlated with light-inducible expression of the associated genes. However, 11 sites with a consensus tandem GATC motif were not associated with light-inducible expression of neighboring genes under the conditions analyzed here (S5 Table).

Although the vast majority of light-inducible genes responded rapidly to light-cues (see above) we have not detected highly confident WCC binding sites (n = 92) in all light-inducible genes (n = 519), raising the question of whether they are directly controlled by the WCC. A visual inspection of the WC2 ChIP-seq coverage at promoters of light-inducible genes without significant binding sites revealed putative light-dependent WCC sites that were not detected by our peak-calling algorithm (S2D Fig.). A subsequent ChIP-PCR analysis revealed a light-dependent enrichment of WCC at such regions (S2D Fig.) indicating that they are true WCC binding sites and suggesting that the associated light-inducible genes are directly controlled by the WCC. Hence, not all of the WCC binding sites were detected by our ChIP-seq analysis.

SUB1 binding to the majority of sites is light-independent

In order to assess the possible effect of light on SUB1 binding on a genome-wide scale we performed tandem ChIP-seq [33] of SUB1_FLAG-HIS using dark grown and light-exposed mycelial cultures. We identified 617 binding sites that were associated with 562 genes (Fig. 2E, S2 Table). 63 genes were light-inducible suggesting a moderate but significant enrichment of SUB1 at light-inducible genes (p < e-07). However, binding of SUB1 to the majority of sites (527) was independent of light. Hence, the elevated expression of SUB1 in light-exposed mycelia does not support increased SUB1 binding on a genome-wide scale. Recruitment of SUB1 to 50 sites (associated with 50 genes) was, however, enhanced by light (S2E Fig.) and 27 of the associated genes were light-inducible. Interestingly, 23 of these 27 genes harbor overlapping (S2F Fig.) or close by WCC binding sites. The data indicate that light-induced SUB1 binding occurs mainly at light-inducible WCC target genes.

Analysis of SUB1 binding sites by MEME revealed the bipartite motif a/cGATc/g-x₆-a/cTGc/t (Fig. 2F, upper panel). This motif was highly enriched (223 / 617) and located in the center of the SUB1 binding sites (Fig. 2G). The MEME analysis revealed in addition two GTA-rich motifs, which were present in 82 and 63 sites, respectively (Fig. 2F, lower panels).

In order to assess the difference between SUB1-dependent and independent light-activated genes we next analyzed recruitment of SUB1 and WCC to light-inducible genes. Sequence coverage of the SUB1 ChIP was higher at promoters of SUB1-dependent light-inducible genes while WCC occupancy was higher at promoters of SUB1-independent light-inducible genes (S2G and S2I Fig.). The data suggest that SUB1 may preferentially support light-induced transcription at promoters with lower affinity for WCC.

SUB1 supports WCC-dependent light-induced nucleosome eviction

To assess whether WCC and/or SUB1 support light-induced chromatin remodeling we performed nucleosome mapping of dark-grown and light-exposed wt, Δsub1 and Δwc2 strains by MNase digestion of DNA followed by paired-end sequencing. Two independent replicates were analyzed. The average length of protected fragments was about 140–150 bp and fragments longer than 100 bp were considered nucleosomal DNA, while protected smaller fragments were considered footprints of TFs or other DNA-binding proteins.

The nucleosome occupancy profiles at WCC binding sites were bimodal. A rather broad region (about ± 2kb) with moderately reduced nucleosome occupancy likely reflects that WCC binding sites are enriched in promoters, which are generally rather nucleosome free, while the confined nucleosome-free region in the center may reflect the actual binding site of WCC (Fig. 3A, dashed lines). In the dark nucleosome occupancy at WCC sites was similar in all the stains (S3A Fig., left panel). Upon light-exposure, the nucleosome occupancy decreased in wt and Δsub1 but not in Δwc2, demonstrating that the activated WCC reduces nucleosome occupancy at its binding sites (Fig. 3A, solid lines). Maximal depletion of nucleosomes was observed in wt, i.e. when SUB1 and light-activated WCC were present (Figs. 3A and S3A, right panel). Analysis of an independent nucleosome analysis supported these results (S3B Fig.). The light-induced nucleosome loss at 20 highly confident WCC binding sites was impaired in Δsub1 while nucleosome eviction was independent of SUB1 at the remaining 72 WCC sites (Fig. 3B).

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Fig 3. Light promotes WCC dependent nucleosome eviction at WCC binding sites.

A. Nucleosome occupancy at binding sites of WCC (n = 92) and in wt (red), Δsub1 (green) and Δwc2 (blue) strains in dark (dotted lines) and 20 min after light-exposure (solid lines). B. Nucleosome occupancy at binding sites of WCC for SUB1-dependent (upper panel) and SUB1-independent (lower panel) light induced nucleosome loss. wt (red) and Δsub1 (green) strains in dark (dotted lines) and 20 min after light-exposure (solid lines) are shown. C. Nucleosome occupancy at binding sites of SUB1 (n = 617) in wt (red), Δsub1 (green) and Δwc2 (blue) strains in dark (dotted lines) and 20 min after light-exposure (solid lines).

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Nucleosome occupancy at SUB1 binding sites was rather low in wt, Δwc2 and even in Δsub1 and essentially independent of light (Figs. 3C, S3A). The data suggest that SUB1 binding sites are either intrinsically free of nucleosomes or that other chromatin remodelers keep these sites open.

To obtain potential footprints of the WCC and SUB1, we analyzed MNase-resistant DNA fragments that were shorter than typical fragments protected by nucleosomes. Protected DNA fragments < 100 bp accumulated in light-dependent fashion at WCC binding sites (S3C Fig., left panel), suggesting that they correspond to a footprint of the light-activated WCC. In contrast, a potential footprint of SUB1 was not affected by light (S3C Fig., right panel).

Together these observations indicate that binding of the light-activated WCC triggers depletion of nucleosomes from its binding sites. SUB1 contributes to the light-induced nucleosome removal at WCC binding sites. Binding sites of SUB1 are also rather devoid of nucleosomes, even in the absence of SUB1, and the nucleosome occupancy of SUB1 sites was independent of light and WCC.

To identify on a genome-wide level light-induced nucleosome remodeling in promoters and genes, we aligned the +1 nucleosomes of all annotated transcription start sites (TSSs). In transcribed regions the nucleosomes were regularly spaced by 176 bp and nucleosome occupancy was rather high (Fig. 4A). In contrast, nucleosome spacing was irregular and occupancy was lower in promoters, similar to corresponding observations in other species [36,37,38]. Light, WCC and SUB1 did not affect nucleosome occupancy of genes and promoters on a genome-wide scale (Fig. 4A). However, light triggered nucleosome remodeling at the promoter of the SUB1-dependent rds1 gene. A light-induced loss of nucleosomes was detected at the overlapping WCC and SUB1 binding sites (Fig. 4B and C). The light-induced nucleosome loss was attenuated in Δsub1 and absent in a Δwc2 strain, indicating that removal was strictly dependent on the activated WCC and supported by SUB1. A light-induced loss a nucleosome was also observed at the WCC binding site of the hyr1 promoter (S4A Fig., left panel), which was, however, not dependent of SUB1.

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Fig 4. Light- and WCC-dependent nucleosome eviction is transcription independent.

A. Line graphs showing the averaged nucleosome occupancy in transcribed genes and promoters of all annotated Neurospora genes (n = 9733) in wt, Δsub1 and Δwc2 strains in dark (red) and 20 min (blue) after light-exposure of cultures. The center of the +1 nucleosome (nucleosome overlapping the annotated transcription start site) was used for alignment of sequence coverage of MNase-resistant fragments >100bp. B. Wig file showing the nucleosome position and occupancy at the rds1 promoter in wt, Δsub1 and Δwc2 strains in the dark and after light-exposure. MNase-WC2 ChIP-seq (blue) is shown below the nucleosome signals. Numbers on the ChIP-seq panels show the maximum read coverage shown in the wig file. C. ChIP-PCR analysis showing H2A occupancy in the dark and 20 min after light exposure at the binding sites of WCC and SUB1 in the rds1 promoter. ChIP was performed by immunoprecipitation with H2A antibody (± SEM, n = 4). actin gene was used for normalization. wt dark level was set to 1. D. Transcription-independent light-induced nucleosome eviction at WCC binding sites (BS). Four examples (wig files) of nucleosome position and occupancy at WCC BS in wt, Δsub1 and Δwc2 strains are shown. WCC binding (TAP-WC2 ChIP-seq) is shown above the nucleosome signals. The positions of GATC motifs are shown in the lower panels. Numbers on the ChIP-seq panels indicate the maximum nucleosome coverage shown in the Wig file. Regions used for ChIP-PCR analysis are indicated by black lines. E. ChIP-PCR analysis showing H2A occupancy in the regions shown in Fig. 4D. Occupancy of H2A was determined by immunoprecipitation with H2A antibody (± SEM, n = 4). actin gene was used for normalization. wt dark level was set to 1.

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Light triggered a substantial loss and repositioning of nucleosomes at the vvd promoter and gene (S4A Fig., right panel). The light-induced nucleosome dynamics were similar in wt and Δsub1 but absent in Δwc2, indicating that chromatin remodeling of the vvd gene by the light-activated WCC was independent of SUB1. The pronounced depletion of nucleosomes in the transcribed region of vvd is likely due to the synchronous activation of the rather strong vvd promoter in the entire ensemble of nuclei. Similar losses of nucleosomes were observed in the transcribed region of other highly expressed light-inducible genes such as al-1, cry and con-10 but was less pronounced or not detectable in less active genes such as hyr1, ncu00309 and frq (S1 Table).

With the exception of highly transcribed genes the light-induced eviction of nucleosomes was generally confined to one or two nucleosomes overlapping the WCC binding sites. We observed eviction of individual nucleosomes at several high affinity binding sites of the WCC that were not associated with transcription initiation of a neighboring or close-by gene (Figs. 4D and S4B). We confirmed the light- and WCC- dependent nucleosome eviction at these sites by independent histone H2A ChIP-PCR (Fig. 4E). These observations suggest that the light-activated WCC supports eviction of nucleosomes at its binding sites independent of transcription.

SUB1 requires light-activated WCC to stimulate expression of light-inducible genes

To address whether SUB1 can activate transcription of light-inducible genes without the WCC we expressed in wc1-deficient (wc1^mut) [39] and wc1-proficient (wc1⁺) strains a FLAG-tagged SUB1 under control of the inducible quinic acid 2 (qa2) promoter. SUB1_FLAG was expressed at low level in the absence of QA and expression levels were elevated in the presence of QA (Figs. 5A, S5A). Expression levels of hyr1, which is a SUB1-affected light-inducible gene, were generally higher in a wc1⁺ background than in the corresponding wc1^mut strains (Fig. 5B). In light, QA-induced SUB1_FLAG supported expression of hyr1 at a high level in the presence of WCC (wc1⁺) but not in the absence of WCC (wc1^mut) (Figs. 5B and S5B). Together the data indicate that SUB1 cannot activate expression of hyr1 independently of the WCC. Maximal expression of hyr1 requires the presence of SUB1 and light-activated WCC.

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Fig 5. SUB1 requires light-activated WCC to induce hyr1 gene expression.

A. Western blot showing SUB1 expression levels of qa promoter driven FLAG-tagged sub1 in wc1⁺ and wc1^mut strains before and 4 h after QA-induction of cultures grown for 24 h in the dark or in constant light. B. Quantification of hyr1 RNA levels by RT-PCR (± SEM, n = 4) under the conditions described in (A). tubulin RNA was used for normalization. The hyr1 RNA level in wc1⁺qa2::sub1_FLAG in the dark (-QA) was set to 1. C. ChIP-PCR analysis of FLAG-SUB1 showing binding of SUB1 to the hyr1 promoter in wc1⁺qa2::sub1_FLAG, wc1^mut_{_}qa2::sub1_FLAG and Δsub1 strains grown 24 h in dark or in constant light. QA-induction, when indicated, was carried out for 4 h. Two-step ChIP was performed using FLAG and SUB1 antibodies. 28s rDNA was used for normalization. The ChIP-PCR signal of light grown wc1⁺qa2::sub1_FLAG was set to 1 (± SEM, n = 3). D. Western blot showing SUB1 levels after light-exposure of wt, tub::sub1 and ccg1::sub1 strains. E. Quantification by RT-PCR of light-induced accumulation of hyr1 RNA levels in wt, Δsub1, tub::sub1 and ccg1::sub1 strains. 28s rRNA was used for normalization. Dark RNA levels of wt were normalized to 1 (± SEM, n = 4).

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To analyze binding of SUB1_FLAG to the hyr1 promoter we performed ChIP-PCR. In a wc1⁺ background binding of QA-induced SUB1_FLAG was more efficient in light than in dark while binding of SUB1 was independent of light in a wc1-deficient strain (Fig. 5C). The data suggest that the light-activated WCC supports recruitment of SUB1 to the hyr1 promoter. Corresponding results were obtained for the rds1 promoter (S5C Fig.).

Since the WCC facilitates recruitment of SUB1, expression of SUB1-dependent light-inducible genes could be limited by SUB1 abundance. To test this hypothesis, we generated WCC-proficient strains expressing SUB1 under control of the tubulin (tub) and the ccg1 promoter, respectively. Dark-grown mycelial cultures of wt, Δsub1, tub::sub1 and ccg1::sub1 were exposed to light and SUB1 levels and the kinetics of hyr1 expression were measured (Fig. 5D and E). In wt, SUB1 was expressed at low level in the dark and accumulated, as expected, to high levels after light exposure. In contrast, SUB1 levels were constitutively low in tub::sub1 and constitutively high in ccg1::sub1 (Fig. 5D). In the dark hyr1 expression correlated well with the SUB1 levels in the respective strain. In response to light, hyr1 levels increased with similar kinetics (5–7 fold) in all strains, reaching the highest expression in ccg1::sub1 and the lowest level in Δsub1. Interestingly, hyr1 levels were similar in wt and tub::sub1 despite light-induced accumulation of substantial amounts of SUB1 in wt (Fig. 5D upper panel). Hence, the SUB1 that was newly synthesized under control of the WCC did not independently activate hyr1 on a second hierarchical tier. Corresponding results were obtained when light-induced expression of the SUB1-dependent genes rds1 and ncu00309 was analyzed (S5D, S5E Fig.). Over-expression of SUB1 in a WCC-deficient background (S5F Fig.) did neither support elevated expression of the SUB1 target genes (rds1 and ncu00309) in the dark nor in light (S5G and S5H Fig.). Together the data indicate that SUB1 functionally cooperates with the dark-form and the light-activated WCC but cannot activate transcription of light-inducible genes in the absence of WCC. Thus, SUB1 supports the activity of WCC in synergistic manner but is not a bona-fide transcription activator of light-inducible genes.

We noted that light-induction of rds1 was strongly attenuated by deletion of sub1 but was not affected by SUB1 overexpression. In contrast, light-induction of hyr1 was only moderately affected by deletion of sub1 but was strongly enhanced by SUB1 overexpression, suggesting that the SUB1 level in wt was limiting for hyr1 expression under the conditions analyzed. To detect hyr1-type genes that respond only to high levels of SUB1 we analyzed the light-inducible transcriptome of ccg1::sub1 in comparison to a wt strain. In wt (replicate 2) we identified 657 light inducible genes (S5I Fig., S3 Table). Most of these genes were also identified by the independent analysis (replicate 1) described above (S5J Fig.). Light-induction of 121 genes was significantly enhanced in ccg1::sub1 (S5K and S5L Fig.). Together with the group of genes down-regulated in Δsub1 (see Fig. 1) the data indicates that about 40% (264) of the light-inducible genes are co-regulated by SUB1.

SUB1 interacts and cooperates with FF7 to activate light-inducible and non light-inducible genes

To identify interaction partners of SUB1 we performed tandem affinity purification of SUB1_FLAG-HIS. By subsequent mass-spectrometry we identified Female Fertility-7 (FF7) as a potential interaction partner of SUB1. FF7 has a Gal4-type Zn(2)-Cys(6) binuclear cluster domain and a putative acetyl transferase domain with similarity to maltose acetyl transferase. To confirm the interaction we constructed a strain expressing FLAG-HIS tagged FF7 and performed reciprocal anti-FLAG and anti-SUB1 immunoprecipitations. SUB1 co-immunoprecipitated with FF7_FLAG-HIS (Fig. 6A) and vice versa, FF7_FLAG-HIS was pulled down with SUB1 antibodies (Fig. 6B). The pull-down efficiency was quite low, suggesting that the interaction is rather unstable. To identify the binding sites of FF7 and to investigate whether SUB1 and FF7 co-localize on the genome, we performed tandem ChIP-seq of FF7_FLAG-HIS from light-exposed mycelial cultures. We identified 2756 putative FF7 binding sites that were associated with 2315 genes (S4 Table), suggesting a rather ubiquitous role of FF7. Analysis of FF7 binding sites by MEME revealed a AACCGC motif (Fig. 6C, upper panel) that was highly enriched in the center of the FF7 binding sites (Fig. 6D). A “GTA” rich motif, similar to the one found in the SUB1 ChIP-seq, was also found in ChIPed FF7 sites (Fig. 6C, lower panel). This motif might be a more general element associated with promoters since it was not enriched at the center of the binding sites. To assess the potential relationship of FF7, SUB1 and WCC we analyzed the occupancies of the transcription factors at their own and the respective binding sites of the other TFs (S6A Fig.). FF7 binding was enriched at WCC and at SUB1 sites. Similarly, SUB1 binding was enriched at WCC and at FF7 sites. WCC sites were, possibly due to their low number (n = 92), neither enriched at SUB1 sites (n = 617) nor at FF7 sites (n = 2756). On a genome wide scale, about 70% (422 / 617) of the SUB1 binding sites were also occupied by FF7 indicating a highly significant (p < e-10) co-occurrence of these factors on the DNA (Fig. 6E). Similarly, 72% (68 / 92) of the WCC binding sites overlap with FF7. Interestingly, essentially all SUB1 and WCC overlapping sites (28 / 29) harbor also FF7 binding sites (Fig. 6E). Examples are shown in Figs. 6F and S6B. The 28 overlapping WCC, SUB1 and FF7 binding sites were associated with 25 expressed genes. 10 of these genes were light-inducible and dependent on SUB1 and/or FF7, 5 genes were light-inducible but independent of SUB1/FF7 and 10 genes were not light-inducible (S1 and S6 Tables). Thus, the majority of functional WCC sites with overlapping SUB1 and FF7 sites are also co-regulated by SUB1 and FF7.

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Fig 6. FF7 interacts weakly with SUB1 and co-regulates light-inducible and non light-inducible genes.

A-B. Western blots showing co-immunoprecipitation (co-IP) of (A) SUB1 with FF7_FLAG-HIS and (B) FF7_FLAG-HIS with SUB1. FLAG antibody was used for FF7_FLAG-HIS IP and α-SUB1 antibody was used for SUB1 IP. The asterisks (*) indicate cross-reactions of the FLAG antibody. C. FF7 binding motifs identified by MEME. The top 200 binding sites identified by FF7 ChIP-seq were used for the motif analysis. The upper motif is found in 117 / 200 binding sites whereas the lower motif is found in 36 / 200 binding sites. D. Occurrence of the major FF7 motif at FF7 binding sites. The grey area shows the occupancy of FF7 binding sites determined by ChIP-seq. The red line shows the occurrence of the FF7 binding motif “t/c AAGCG c/a”. E. Wig file showing MNase-WC2, SUB1 and FF7 ChIP-seq signals at the rds1 promoter. Numbers on the ChIP-seq panels correspond the maximum coverage shown in the wig file. F. Venn-diagram showing the overlap between SUB1, WC2 and FF7 ChIP-seq signals. G. Heat-map showing light-inducible genes with significantly lower RNA levels in Δsub1 and in Δff7 strains in comparison to wt. H. Wig file (left panel) showing the nucleosome position and occupancy at the rds1 promoter in wt and Δff7 strains in the dark and after light-exposure. The MNase-WC2 ChIP-seq (blue) is shown below the nucleosome signals. Numbers on the ChIP-seq panels show the maximum coverage shown in the wig file. ChIP-PCR analysis (right panel) of H2A occupancy at the binding sites of WCC and SUB1 at rds1 promoter in the dark and 20 min after light-exposure (± SEM, n = 4). actin DNA was used for normalization. wt dark level was set to 1. I. Nucleosome occupancy at binding sites of WCC (n = 92) and in Δff7 in dark (dotted lines) and 20 min after light-exposure (solid lines).

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To analyze the functional cooperation of FF7 with SUB1 and WCC, we determined by RNA-seq the transcriptome of a Δff7 strain grown in the dark and after light-exposure. RNA levels of 192 of 519 light-inducible genes were reduced in Δff7 (S6C Fig., S1 Table). 109 of these genes show impaired light-induction in Δff7 and in Δsub1 (Fig. 6G). We confirmed the impaired light-induction of the rds1 gene by independent RNA measurements (S6D Fig.). Furthermore, expression of 440 genes was reduced in Δff7 in a light-independent manner (S1 Table) and expression of 278 of these genes was also reduced in Δsub1 (S6E Fig.).

Finally, we performed nucleosome mapping of Δff7 after light-induction to assess whether FF7 contributes to light-induced nucleosome eviction at WCC binding sites. The light-induced nucleosome loss at the rds1 promoter was independent of FF7 despite impaired light-induction of the rds1 gene in Δff7 (Fig. 6H). Similarly, we did not detect impaired light-induced nucleosome removal at other WCC sites in Δff7 (Fig. 6I) suggesting that FF7 is required for transactivation rather than eviction of nucleosomes. Together the data suggest that WCC, SUB1 and FF7 have distinct functions and cooperate to regulate subsets of genes in a combinatorial fashion.

Neurospora strains

Neurospora strains; wt (FGSC #2489), Δsub1 (FGSC #11127), Δwc2 (FGSC #11124), Δff7 (FGSC #11073), wc1^mut (FGSC #4398), bd (FGSC #1859), his-3 (FGSC #6103) used in this study were acquired from FGSC. wc1⁺ (FGSC #9718) was used to generate ff7_FlagHis, sub1_FlagHis and wc1⁺qa2::sub1_Flag strains. wc1^mut was used to generate wc1^mut qa2::sub1_Flag. bd Δwcc, his-3 [52] was used for integration of ccg1::sub1 into the his-3 locus to obtain bd ΔWCC ccg1::sub1. his-3 strain was crossed to Δsub1 to create Δsub1,his-3 strain that was used to generate tub::sub1 and ccg1::sub1strains. Δsub1,his-3 strain transformed with empty vector was used as a sub1 KO strain in RNA measurements together with ccg1::sub1 and tub::sub1 strains.

Culture conditions

Standard growth medium contained 2% glucose, 0.5% L-arginine, 1× Vogel's medium, and 10 ng / mL biotin. For light-induction experiments, indicated strains were grown in petri plates until mycelial mats formed. Mycelial pads (1 cm) were cut out and grown for 1 day in light at 25°C and transferred to darkness for 24 h before cultures were exposed to light (100 μE) and harvested after the indicated time periods. For replicate 1 of the light-induction experiment wt, Δsub1 and Δff7 strains were analyzed whereas wt and ccg1:sub1 strains were analyzed for the replicate 2. For the QA induction experiments 0.3% QA (final) was added to 24 h dark grown and constant light grown cultures. Samples were harvested before and 4 hours after the addition of QA. For nucleosome mapping conidia of wt, Δsub1, Δwc2 and Δff7 (strains were inoculated in 200 ml media, grown in light for 2 days and transferred to darkness for 24 h. Dark grown and 20 min light exposed cultures were crosslinked with 0.5% paraformaldehyde (FA) for 10 min. FA was quenched with 125 mM glycine for 5 min. Δff7 was not included into the replicate 2 nucleosome mapping.

Generation of knock-in cassettes and Neurospora transformations

The yeast in vivo recombination system [53] was used to generate sub1_FlagHis, ff7_FlagHis and qa2::sub1_Flag strains. Transformation of Neurospora was performed as described [53]. Primers are listed in S6 Table.

Protein analysis

Extraction of proteins and subcellular fractionation were performed as described [54]. SUB1 rabbit antibody was generated against the peptide “RKRQLEQRSIRPKPTDDRN”. H2A antibody was generated against the peptide “CHQNLLPKKTGKTGKNASQEL” Western blotting was performed as described [55]. Protein concentration was estimated by measuring absorption at 280 nm (NanoDrop, PeqLab). Enhanced chemiluminescence signals were detected with X-ray films (Fuji Film Tokyo, Japan).

RNA analysis

RNA was prepared with peqGOLD TriFAST (peqLab, Erlangen, Germany) and reverse transcribed with the QuantiTect Reverse Transcription Kit (QIAGEN, Hilden, Germany). Transcript levels were analyzed by quantitative real-time PCR in 96-well plates with the StepOnePlus Real-Time PCR System (Applied Biosystems). TaqMan Gene Expression Master Mix (Applied Biosystems) and UPL probes (Roche) were used. Primers and probes are listed in S6 Table.

MNase digestion for nucleosome mapping

The previously published MNase digestion protocols [56,57] were optimized for Neurospora. 400 mg ground mycelial powder from each culture was resuspended in 3.75 ml MNase digestion buffer (250 mM sucrose, 60 mM KCl, 15 mM NaCl, 15 mM Tris-HCl pH 7.4, 3 mM MgCl_2, 1 mM CaCl_2, 0.2% NP-40) with freshly added 0.5 mM DTT, 0.5 mM Spermidine and protease inhibitors (EDTA-free, Roche). The suspension was mixed by vortexing and incubated on ice for 5 minutes. Next 750 μl aliquots were distributed to five 1.5 ml Eppendorf tubes. MNase powder (Sigma-N3755-500) was resuspended in 850 μl MNase resuspension solution (10 mM HEPES-KOH pH 7.6, 50 mM KCl, 1.5 mM MgCl₂, 0.5 mM EGTA, 10% glycerol). Aliquots were digested with different amounts (0 [control], 0.75, 1.5, 3, 6U) of MNase to cover both, sensitive and resistant sites. All samples were incubated at 25°C for 1 hour with shaking at 400 rpm in a Themo-mixer. The reaction was stopped by adding stop buffer (final 0.2% SDS, 10 mM EDTA pH 8.0). Samples are centrifuged at 20000 g for 20 min to pellet the cell debris. Supernatants were transferred to new tubes and 2 μl RNAse Cocktail Enzyme mix (Life Technologies, AM2286) was added and incubated at 37°C for 45 minutes to degrade RNA. To degrade proteins, 15 μl proteinase K solution (Life Techonolgies, AM2548) was added and samples were incubated at 65°C for 2 hours. DNA was precipitated with EtOH in the presence of 40 μg glycogen (Thermo) and further cleaned by using PCR clean-up kit (Promega). Aliquots were analyzed by electrophoresis in a 1.7% Agarose gel (80 volts for 1 hour) to visualize nucleosomal DNA ladders. Paired-end libraries for sequencing were prepared as described below.

MNase-WC2-ChIP

Neurospora cultures were crosslinked with 1% FA for 10 min. FA was quenched 5 min with 125 mM glycine. ∼ 400 μl ground mycelial powder was resuspended and digested in 600 μl MNase digestion buffer for 1 h at 25°C with 15 U of MNase (Sigma-N3755-500) (In addition an independent sample from a 1 min light-induced culture was digested with 5 U MNase). The reactions were stopped by adding final 5 mM EDTA pH 8.0. Then 400 μl ChIP lysis buffer (50 mM HEPES, 150 mM NaCl, 1 mM EDTA, 1% Triton-X, 0.1% SDS and 0.1% NaDOC) was added and samples were centrifuged at 4°C, 15000 g. The rest of the ChIP protocol was performed as described [19]. DNA for ChIP-seq was pooled from two-independent experiments.

Tandem chromatin immunoprecipitation with FLAG-His and protein A/Calmodulin

Light-induction was performed for the indicated time periods and cultures were cross-linked in constant light with 1% FA for 15 min. FA was quenched with 125 mM glycine for 5 min. 6 aliquots of 600 μl ground mycelial powder were used for each time point for TAP-WC2-ChIP-seq. Mycelia were dissolved in 1 ml ChIP lysis buffer (50 mM HEPES pH 7.4, 150 mM NaCl, 1 mM EDTA, 1% Triton-X, 0.1% SDS and 0.1% NaDOC) with freshly added 0.5 mM DTT and protease inhibitors (Roche cOmplete Protease Inhibitor Cocktail-EDTA free). Sonification was performed with SonoLab^TM Covaris Version 7.0.20.0 (average incident power 36 watt, peak incident power 180 watt, duty factor 20 percent, cycles / burst 200 count and duration 160 seconds). Samples were centrifuged at 15000 g for 20 min at 4°C. Supernatants of the six aliquots per time points were combined and incubated with 100 μl IgG Sepherose/time point (GE Healthcare) for 3 h at 4°C. Beads were washed 2x with ChIP lysis buffer, 1x Lindet (250 mM LiCl, 1% NP-40, 1% NaDOC, 1 mM EDTA, 10 mM Tris / HCl; pH 8) and 2x with TAP buffer (50 mM Tris–HCl, pH 7.5, 150 mM NaCl, 1.5 mM MgCl2, 0.1% NP-40, 0.5 mM DTT). Elution was performed with two consecutive digestions with 50 U TEV protease (Invitrogen) at 16 C° 2 h and ON at 4 C°. Final 3 mM CaCl₂ was added to the combined elution and incubated with 130 μl Calmodulin Sepharose 4B (GE Healthcare) for 3.5 h. Beads were washed 5 times with TAP buffer with 3 mM CaCl₂. The elution and DNA extraction was performed as described [19]. DNA for ChIP-seq was collected from three-independent experiments.

Tandem ChIP with Ni-NTA enrichment, followed by anti-Flag immunoprecipitation was performed as described [33]. Primers and probes used for the ChIP-PCR are listed in S6 Table.

RNA sequencing and data analysis

NEBNext Ultra RNA Prep kit with NEBNext Multiplex oligos was used for cDNA preparation at Bioquant Deep Sequencing Core Facility. PolyA selection was performed at the beginning of the protocol. The size and the quality of the libraries were checked with a 2100 Bioanalyzer. Un-paired sequencing with 50 bp read length was performed with a HiSeq 2000 at GeneCore EMBL Heidelberg. Raw reads can be accessed at SRA database under the accession numbers listed in S7 Table.

50 bp long raw reads were mapped to Neurospora crassa genome NC10 by using Bowtie [58]. Maximum 3 mismatches were allowed for the mapping. Reads mapping to more than one site were discarded. Gene expression was quantified by the number of reads falling into annotated exons. Normalization between samples was performed by using the size factor formula as described [59]. Genes with low RNA levels (lower 20% of all annotated genes) were excluded from further analysis.

In order to identify significant light-induced genes, differential gene expression analysis was performed. Read counts of the genes were assumed to follow negative binomial (NB) distribution, Gi≈NB(μi, σi), where μi is mean and σi is variance. Since no replicates information were available, σi was estimated based on the mean and variance correlation. A “locfit” R package was used to fit the relationship between the mean and variance. By adapting the Robinson and Smyth Exact Test [60], the two side p-value can be computed as follow, where , . is the read counts of i_th gene in treatment condition, is the read counts of i_th gene in control condition, f(a,b) is the product of f(a) and f(b), which can be computed using dnbinom of R package. After finding the genes that have a significant (p < 0.05) increase in the reads in either 30, 60 or 120 minutes compared to DD reads, we set another cut off (2 fold) to further filter the candidates. In order to identify genes that show impaired light-induced in Δsub1 light-induced time points were compared and p values were calculated as described above. The time points that have significantly lower reads (p < 0.05) in the knock-out strains were identified. Similar analysis and p values were used to identify up-regulated genes in ccg1:sub1 compared to wt (replicate 2).

We analyzed ChIP-seq data from Cesbron et al. to assess the kinetics of RNAPII recruitment to 519 light-inducible genes (RNA-seq replicate 1). Significantly increased levels (p < 0.05) of transcribing RNAP (RNAPII-S2P) were detected at 221 genes already 1, 5 and 10 min after a 1 min light-pulse.

Nucleosome mapping and data analysis

Libraries from purified MNase-digested DNA were prepared without size selection to detect nucleosomal DNA and putative footprints of transcription factors. NEBNext Ultra™ DNA Library Prep Kit for Illumina (E7370L) was used for library preparation at the Bioquant Deep Sequencing Core Facility by using manufacturer`s instructions. The size and the quality of the libraries were analyzed with a 2100 Bioanalyzer. Paired-end sequencing with 100 bp read length was performed with a HiSeq 2000 at BGI, Hong Kong. Raw reads can be accessed at SRA database under the accession numbers listed in S7 Table.

For nucleosome mapping the fragment length was set to be between 100 bp and 1000 bp with forward and reverse conformation. For mismatches and multiple alignments the same settings were used as in the single end mapping. The non-mapped reads were further processed to remove the adapters, mapped to the genome again and analyzed independently. The middle 50 bp of the pair end position was used to generate the wiggle file format genome nucleosome coverage. Wig files were visualized with IGV genome browser [61]. The normalization factor was computed by using 90^th percentile of each experiment. Smoothing was carried out by using Kernel Regression Smoother package of R. 1500 bp upstream and downstream of annotated TSS position were used for genome wide nucleosome coverage analysis. The center of the +1 nucleosome of genes was defined by the maximum read coverage in a window of 200 bp around the annotated TSS. Each nucleosome was estimated to cover 176 bp, and the nucleosome coverage was estimated by the area under the curve.

To analyze the nucleosome coverage at the transcription factor binding sites (BS) the nucleosome mean coverage at the summit of the BS was calculated. As the position of a BS relative to a TSS follows a NB, a random NB using the same parameter was generated. The background nucleosome mean coverage was then computed by using the random relative position of BS to TSS. The nucleosome mean coverage was determined by using the normalized nucleosome mean coverage and plotted as ratio to remove the possible bias related to the promoters.

ChIP-Sequencing and data analysis

ChIP DNA libraries were prepared with NEBNext ChIP-Seq Library Prep Reagent Set for Illumina with NEBNext Multiplex oligos at Bioquant Deep Sequencing Core Facility. A 2100 Bioanalyzer was used to check the quality and size of the libraries. Un-paired sequencing with 50 bp read length was performed with a HiSeq 2000 at BGI, Hong Kong.

Mapping was performed as in RNA-seq analysis. In order to identify peaks a sliding window of 150 bp with a step of 50 bp was used to scan the genome and quantify the read intensity. The read intensity was normalized by using the total mapped reads. Wig files were visualized with IGV genome browser [61]. The significantly enriched windows were computed by fitting the read intensity to a Poisson distribution. A ChIP binding site was called if 4 continuous windows were statistically higher (p value < 1e-05) compared to the control ChIP-seq. FGSC #9718 strain was used as a background to identify the significant windows for FLAG-HIS ChIP-seq. Dark ChIP of each experiment was used as a background for TAP-WC2 and MNase-WC2 ChIP. Another cut off was determined based on the coverage of the identified ChIP peak. We excluded the binding sites with low enrichment (< 1.5 fold, < 90^th percentile of the coverage) by comparing the peak coverage to general coverage of the ChIP-seq. Raw reads can be accessed at SRA database under the accession numbers listed in S7 Table.

Upstream and downstream genes were analyzed for the annotation of the peaks. The gene was annotated to a peak if there was a peak detected within 1000 bp upstream of the TSS and 500 bp downstream of TSS. If the peak was not close to a promoter of any gene, the binding site was annotated to the nearest genes with the expected orientation (i.e. binding sites are upstream of the annotated TSS) with no distance limitation. If there were no genes in the upstream/downstream of the peak with the correct orientation, the peak was not annotated to any gene.

The SUB1 binding sites of DD and 30 min light-induction samples were merged based on the coordinate location to find light-induced SUB1 binding sites. Using linear regression between the DD and 30 min light-induction read intensity of the peaks, light-regulated SUB1 binding sites were identified. The linear regression was fitted using R, and the clustering was based on the 80% confidence prediction interval.

Motif analysis was done by using MEME motif search software [35]. 300 bp DNA regions surrounding the summit of the ChIP-seq peaks were used. For the WCC ChIP-seq “GATC” motif the pair wise distribution was calculated within these 300 bp regions.