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. 2015 Mar 23;200(1):29–33. doi: 10.1534/genetics.114.170514

A Novel Approach for Studying Histone H1 Function in Vivo

Giorgia Siriaco 1, Renate Deuring 1, Gina D Mawla 1,1, John W Tamkun 1,2
PMCID: PMC4423370  PMID: 25805849

Abstract

In this report, we investigate the mechanisms that regulate Drosophila histone H1 expression and its association with chromatin in vivo. We show that histone H1 is subject to negative autoregulation and exploit this result to examine the effects of mutations of the main phosphorylation site of histone H1.

Keywords: Drosophila, histone H1, chromatin, gene expression

HISTONE H1 is a key determinant of higher-order chromatin structure. This conserved linker histone contains a winged helix domain that binds the nucleosome near the site of DNA entry and exit; the regions flanking this domain interact with core histones and linker DNA to package nucleosome arrays into 30-nm fibers in vitro (Harshman et al. 2013). Consistent with its biochemical properties, the loss of histone H1 function leads to chromosome decondensation, defects in gene expression, genomic instability, and lethality (Godde and Ura 2008; Happel and Doenecke 2009; Lu et al. 2009; Siriaco et al. 2009; Vujatovic et al. 2012; Lu et al. 2013). These findings have stimulated great interest in the mechanisms that regulate the expression, assembly, and function of histone H1.

Several factors make Drosophila an excellent model organism for studying histone H1 function in vivo. Unlike most higher eukaryotes, which contain multiple, functionally redundant histone H1 variants (Godde and Ura 2008; Happel and Doenecke 2009), Drosophila has only one somatic histone H1 variant, the product of the His1 gene. In addition, a subset of Drosophila tissues contains gigantic polytene chromosomes that arise from repeated rounds of DNA replication in the absence of cell division, making it much easier to directly score chromosome defects resulting from the loss of histone H1 function.

In this study, we used a Drosophila strain expressing histone H1 tagged with green fluorescent protein (H1-GFP) to investigate the regulation of histone H1 expression and its association with chromatin. A GAL4-inducible transgene encoding histone H1 with GFP fused to its C terminus (UAS–H1-GFP) (Fasulo et al. 2012) was expressed in the salivary glands of larvae bearing insertions of UAS–H1-GFP and an eyGAL4 driver (Hazelett et al. 1998). Live imaging revealed that H1-GFP is associated with polytene chromosomes of eyGAL4 UAS–H1-GFP larvae; no unbound H1-GFP was detected in the nucleoplasm (Figure 1A). The staining of polytene chromosome squashes with DAPI revealed that the banding patterns of H1-GFP and DNA are highly coincident (Figure 1, B and C), as previously observed for the endogenous histone H1 protein (Corona et al. 2007).

Figure 1.

Figure 1

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Histone H1 tagged with GFP is functionally equivalent to the endogenous histone H1 protein. (A) Confocal analysis reveals that H1-GFP is primarily associated with chromatin and localizes in a normal banding pattern in live eyGAL4 UAS–H1-GFP salivary gland polytene chromosomes. Chromosomes were imaged as previously described (Fasulo et al. 2012). Scale bar, 10 μm. (B) H1-GFP is normally distributed (as evidenced by GFP fluorescence) in fixed eyGAL4 UAS–H1-GFP salivary gland polytene chromosomes and colocalizes with DAPI. Arrowhead marks chromocenter. Chromosomes were prepared as previously described (Corona et al. 2007). Scale bars, 10 μm. (C) Comparison of the pixel intensity and pattern of the boxed chromosome arm in B reveals that H1-GFP and DAPI are highly overlapping. Chromosome boundaries were identified and the sum pixel intensity within chromosomes was calculated using Volocity (Improvision) software.

The expression of histone H1 must be tightly regulated, as even modest changes in the level of histone H1 can have dramatic effects on nucleosome repeat length, global nucleosome density, and chromatin compaction (Blank and Becker 1995; Woodcock et al. 2006; Routh et al. 2008; Siriaco et al. 2009). In budding yeast, core histones are subject to negative autoregulation (Gunjan et al. 2006; Eriksson et al. 2012). Autoregulation also maintains constant levels of core histones in Drosophila (McKay et al. 2015). These observations prompted us to examine whether similar mechanisms are used to regulate histone H1 levels in Drosophila. As expected, the activation of the UAS–H1-GFP transgene in the larval salivary gland using the strong eyGAL4 driver caused a large (∼10-fold) increase in the total level of histone H1 RNA (i.e., endogenous plus GFP-tagged H1; Figure 2A). This increase was accompanied by a compensatory 10-fold decrease in the expression of the endogenous histone H1 protein, as determined by Western blotting using an antibody that recognizes both the endogenous and GFP-tagged H1 proteins (Figure 2, B and C). As a result, the total level of histone H1 protein did not increase significantly and H1-GFP became the predominant form of histone H1 in this tissue (Figure 2, B and C). The degree of endogenous H1 repression was sensitive to the dosage of the UAS–H1-GFP transgene, with the strongest repression observed in eyGAL4 UAS–H1-GFP homozygotes (data not shown). Similar results were obtained using the ubiquitously expressed daGAL4 driver (Gerber et al. 2004; see below). Flies ubiquitously expressing the UAS–H1-GFP transgene were viable and showed no developmental delay (data not shown). These findings indicate that negative autoregulation maintains relatively constant levels of histone H1 in vivo.

Figure 2.

Figure 2

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Histone H1 is negatively autoregulated in vivo. (A) Total levels of H1 RNA in eyGAL4 UAS–H1-GFP salivary gland nuclei were monitored by RT–PCR using primers specific to the RNAs encoded by RP49 and endogenous His1, described in Table 2. Activation of the UAS–H1-GFP transgene increased levels of total H1 RNA. (B) Endogenous H1 protein levels are greatly reduced in eyGAL4 UAS–H1-GFP salivary gland nuclei, as shown by Western blotting. Salivary gland protein extracts were prepared from third-instar larvae and analyzed by protein blotting as described previously (Corona et al. 2007) using antibodies against H1 (Active Motif, 39575) and H3 (Abcam, ab1791). (C) Quantitation of H1-GFP and endogenous H1 protein in eyGAL4 UAS–H1-GFP salivary gland nuclei shows a twofold increase in H1-GFP. For quantification, Western blots were scanned and the sum pixel intensity for each band was calculated using Volocity (Improvision) software.

The suppression of endogenous histone H1 levels via the expression of H1-GFP did not cause obvious changes in either the structure or banding pattern of salivary gland polytene chromosomes (Figure 1, A and B). By contrast, even the partial knockdown of histone H1 levels by RNAi blocks development and causes dramatic defects in polytene chromosome structure (Lu et al. 2009; Siriaco et al. 2009). These findings indicate that histone H1-GFP is functionally equivalent to the endogenous histone H1 protein and can be used to study histone H1 function in vivo.

The association of histone H1 with chromatin is highly dynamic, with the bulk of H1 in the genome undergoing exchange in less than a minute (Lever et al. 2000; Contreras et al. 2003; Siriaco et al. 2009). Recent studies implicating histone H1 exchange in the regulation of cellular pluripotency and differentiation have heightened interest in the molecular mechanisms underlying this process (Meshorer et al. 2006; Alvarez-Saavedra et al. 2014; Christophorou et al. 2014). In both Tetrahymena and mammals, the phosphorylation of histone H1 weakens its affinity for chromatin (Roth and Allis 1992) but its effect on histone H1 assembly and chromosome compaction remains poorly understood.

Drosophila is a good model organism for studying the role of this modification since it has a single zygotic H1 isoform that contains only one major phosphorylation site: histone H1 serine 10 (H1S10) (Villar-Garea and Imhof 2008; Bonet-Costa et al. 2012). Standard genetic approaches cannot be used to study histone H1 modification in Drosophila, however, due to the presence of ∼100 identical, functionally redundant copies of the gene encoding histone H1 (His1) interspersed with genes encoding the core histones H2A, H2B, H3, and H4 in the histone gene cluster (HIS-C) (McKay et al. 2015). Previous studies of histone H1 function in most higher eukaryotes, including Drosophila, have therefore been limited to the analysis of phenotypes resulting from the knockdown of histone H1 levels by RNAi (Lu et al. 2009; Siriaco et al. 2009). Our discovery that H1-GFP represses the expression of the endogenous H1 protein allowed us to circumvent this obstacle and characterize phenotypes associated with mutations affecting specific histone H1 residues.

Transgenic strains bearing GAL4-inducible transgenes encoding GFP-tagged histone H1 proteins with amino acid substitutions that mimic (H1S10E) or block (H1S10A) S10 phosphorylation were generated by P-element-mediated transformation. The UAS–H1S10A-GFP and UAS–H1S10E-GFP transgenes were then expressed in the larval salivary gland using the daGAL4 driver to study the effect of the S10E and S10A mutations on histone H1 function in vivo. As observed for H1-GFP (Figure 2B and Figure 3A), the expression of either H1S10A-GFP or H1S10E-GFP dramatically reduced the level of endogenous histone H1 such that the mutant form of histone H1 was the predominant linker histone in the larval salivary gland (Figure 3A).

Figure 3.

Figure 3

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Mutations in the major phosphorylation site do not alter histone H1 function in vivo (A) Western analysis of salivary gland protein extracts from UAS–H1-GFP; daGAL4, UAS–H1S10A-GFP; daGAL4 and daGAL4 UAS–H1S10E-GFP third-instar larvae. Endogenous H1 protein levels are reduced compared to wild-type control, suggesting that H1-GFP, H1S10A-GFP, and H1S10E-GFP are the main forms of histone H1 in salivary gland nuclei. Both endogenous H1 and H1-GFP were detected using antibodies against H1 (Active Motif, 39575). (B) Fixed salivary gland polytene chromosomes from UAS–H1-GFP; daGAL4, UAS–H1S10A-GFP; daGAL4 and daGAL4 UAS–H1S10E-GFP third instar larvae. H1-GFP, H1S10A-GFP, and H1S10E-GFP are normally distributed on polytene chromosomes and colocalize with DAPI. Scale bars, 10 μm. (C) Confocal images of UAS–H1-GFP; daGAL4, UAS–H1S10A-GFP; daGAL4 and daGAL4 UAS–H1S10E-GFP third-instar salivary gland nuclei. Confocal images were obtained as described in Fasulo et al. (2012). H1-GFP, H1S10A-GFP, and H1S10E-GFP localize normally to chromosomes. Bottom: magnification of a single section through the salivary gland nuclei shown in the top. Images were taken at comparable gains. Scale bars, 10 μm. (D) Exchange rate of H1S10A-GFP is reduced compared to H1S10E-GFP and H1-GFP.

Neither the S10A or S10E mutation caused obvious defects in the binding of histone H1 to chromatin in vivo. As observed for the wild-type histone H1 protein, the distribution of both H1S10A-GFP and H1S10E-GFP on fixed polytene chromosome squashes was coincident with DAPI staining (Figure 3B). Live analyses also revealed that the H1S10A-GFP and H1S10E-GFP proteins associated with chromatin in a normal banded pattern; the majority of both mutant proteins were bound to chromatin and neither H1S10A-GFP nor H1S10E-GFP was observed in the nucleoplasm (Figure 3C). The expression of either H1S10A-GFP or H1S10E-GFP caused a slight (∼25%) reduction in both the size and DNA content of salivary gland polytene chromosomes, suggesting they may have a subtle effect on endoreplication (Figure 3B and data not shown). However, the banding pattern, morphology and overall structure of polytene chromosomes of larvae expressing H1S10A-GFP or H1S10E-GFP in place of the endogenous H1 protein were surprisingly normal (Figure 3, B and C). We therefore conclude that H1S10 mutations have little or no effect on chromatin compaction in vivo.

We also investigated the potential effect of H1S10 phosphorylation on histone H1 exchange in salivary gland nuclei using FRAP assays. The H1S10A mutation increased the residence half-time (t1/2 = 31 sec) but not the mobile fraction of histone H1 (Figure 3D and Table 1). By contrast, the H1S10E mutation did not alter histone H1 exchange (t1/2 = 21 sec vs. 20 sec for H1-GFP; Figure 3D and Table 1). These data suggest that even large changes in the rate of histone H1 exchange do not cause obvious changes in chromosome structure in vivo. Thus, mutations that block or mimic the phosphorylation of H1S10—the main site of histone H1 phosphorylation—have minimal effect on histone H1 function in this tissue. Consistent with these findings, flies expressing high levels of either mutant protein under the control of the strong, ubiquitously expressed daGAL4 driver were healthy, fertile, and phenotypically normal (data not shown). It remains possible that the phosphorylation of other sites of the histone H1 protein modulate its activity in vivo.

Table 1. Quantitation of histone H1 exchange using FRAP assays.

Genotype t1/2 (sec) Mobile fraction (%)
UAS–H1-GFP; daGAL4 20.63 ± 6.9 79 ± 15
UAS–H1S10A-GFP; daGAL4 31.27 ± 13.5 72 ± 16
daGAL4 UAS–H1S10E-GFP 20.03 ± 10.3 72 ± 11
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The mobile fraction and residence half-times of wild-type and mutant histone H1 tagged with GFP were determined by FRAP. Each value represents the average of at least 10 experiments (average n = 15). For FRAP experiments, 10 single imaging scans were acquired followed by 60 bleach pulses of 267 ms within a square region of interest (ROI) measuring 2 × 2 μm. Every second, 160 images were then collected. For imaging, the laser power was attenuated to 8% of the bleach intensity. Two further ROIs, measuring 2 × 2 μm, were used to measure total fluorescence and background. Recovery curves were generated and analyzed using easyFRAP software (Rapsomaniki et al. 2012).

Table 2. List of oligonucleotides used for the construction of transgenes and the quantification of RNA levels by RT-PCR.

Primer Sequence
RP49-F 5′ CACCAGGAACTTCTTGAATCCCGG 3′
RP49-R 5′ AGATCGTGAAGAAGCGCACCAAG 3′
His1-F 5′ CAGCAAATGGTGGACGCTTC 3′
His1-R 5′ GCGAACATGTACCAAATACTGC 3′
H1-F 5′ TATCTCGAGTGCTTTTTGGCAGCCGTAGTCTCGC 3′
H1S10A-R 5′ACCGAATTCGCATGTCTGATTCTGCAGTTCGAACGTCCGCTGCCCCAGTG 3′
H1S10E-R 5′CCGAATTCGCATGTCTGATTCTGCAGTTGCAACGTCCGCTGAGCCAGTGG 3′
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The primer names and corresponding sequences are shown. To generate GAL4-regulated transgenes encoding C-terminal GFP-tagged histone H1 S10A and S10E substitutions, fragments were amplified from Drosophila genomic DNA using the H1-F, H1S10A-R, and H1S10E-R primers, subcloned into the EcoRI and XhoI sites of pENTR1A, and transferred to the P-element transformation vector pTWG (Brand and Perrimon 1993; Rørth et al. 1998). Transformants were generated by P-element-mediated transformation using the Df(1)w67c2 strain (Spradling 1986). Homozygous viable transformants used in this study include UAS–H1S10A-GFP (line 1A4B) and UAS–H1S10E-GFP (line 2G1), which are located on the second and third chromosomes, respectively.

Although the mechanism by which histone H1 regulates its own expression is unknown, we suspect that it occurs at the level of protein stability. In our studies, we rarely observed free H1-GFP in the nucleoplasm, even using the strongest available GAL4 drivers, suggesting that the unbound histone H1 pool is unstable and rapidly degraded. Consistent with this possibility, we have been unable to express mutant forms of the histone H1 protein lacking the winged-helix domain required for nucleosome binding (data not shown). Histone H1 stability could also be dependent on a limiting interaction partner, as has been observed for the centromere-associated histone H3 variant CENP-A, which is unstable unless it is bound to the CENP-A assembly factor Cal1 (Schittenhelm et al. 2010; Chen et al. 2014).

Our serendipitous finding that histone H1 is autoregulated allowed us to establish a system for investigating mutations in the main phosphorylation site of histone H1. Until recently, it was impossible to conduct studies of Drosophila histone genes due to the presence of multiple, functionally redundant copies of the genes encoding histone H1 and all four core histones in the HIS-C gene cluster. Although a strategy for overcoming this obstacle has recently been developed and used to analyze phenotypes associated with Drosophila core histone mutations (Gunesdogan et al. 2010; Hodl and Basler 2012; Pengelly et al. 2013; McKay et al. 2015), it is relatively labor intensive and its utility for studying histone H1 function has not been established. By expressing transgenes encoding mutant H1 proteins under the control of a strong, GAL4-inducible promoter, we were able to test their ability to functionally replace the endogenous histone H1 protein. In theory, this approach could be used to study mutations in other residues of histone H1 or other proteins subject to negative autoregulation.

Acknowledgments

We thank the members of our laboratory for helpful discussions during the course of this work and the University of California—Santa Cruz Life Sciences Microscopy Center for technical support. Stocks obtained from the Bloomington Drosophila Stock Center (National Institutes of Health P40OD018537) were used in this study. This work was supported by National Institutes of Health grant GM49883 to J.W.T.

Footnotes

Communicating editor: P. K. Geyer

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