As in all metazoans, the replication-dependent histone genes of Caenorhabditis elegans lack introns and contain a short hairpin structure in the 3′ untranslated region. This hairpin structure is a key element for post-transcriptional regulation of histone gene expression and determines mRNA 3′ end formation, nuclear export, translation and mRNA decay. All these steps contribute to the S-phase-specific expression of the replication-dependent histone genes. The hairpin structure is the binding site for histone hairpin-binding protein that is required for hairpin-dependent regulation. Here, we demonstrate that the C. elegans histone hairpin-binding protein gene is transcribed in dividing cells during embryogenesis and postembryonic development. Depletion of histone hairpin-binding protein (HBP) function in early embryos using RNA-mediated interference leads to an embryonic-lethal phenotype brought about by defects in chromosome condensation. A similar phenotype was obtained by depleting histones H3 and H4 in early embryos, indicating that the defects in hairpin-binding protein-depleted embryos are caused by reduced histone biosynthesis. We have confirmed this by showing that HBP depletion reduces histone gene expression. Depletion of HBP during postembryonic development also results in defects in cell division during late larval development. In addition, we have observed defects in the specification of vulval cell fate in animals depleted for histone H3 and H4, which indicates that histone proteins are required for cell fate regulation during vulval development.

Introduction

Histones are the protein components of nucleosomes and are required for the packaging of DNA into chromatin. In animals, histone synthesis is determined by the expression of two classes of histone genes: replication-dependent and replication-independent, or replacement variant, histone genes (for reviews,see Maxson et al., 1983;Osley, 1991;Brown, 2001). Caenorhabditis elegans has clusters of replication-dependent histone genes as well as a small number of replacement histone genes(Table 1). Replacement histone genes are expressed either constitutively or in a tissue-specific manner (for reviews, see Maxson et al.,1983; Brown, 2001). In contrast, replication-dependent histone genes are responsible for the doubling of the histone content during S phase in dividing cells in order to maintain a regular chromatin structure upon replication of the nuclear DNA(for reviews, see Osley, 1991;Marzluff, 1992;Müller and Schümperli, 1997;Dominski and Marzluff, 1999). Replication-dependent histone genes are unusual in that they lack introns and contain a conserved RNA hairpin structure in the 3′ untranslated region of the RNA. This RNA hairpin structure is required for histone RNA cleavage, a nuclear processing event that occurs during S phase, and produces mRNA ending immediately after the hairpin structure and lacking a poly(A) tail. Furthermore, the RNA hairpin is required for nuclear export and translation of histone mRNA and the regulation of histone mRNA stability. The hairpin is the binding site for histone hairpin-binding proteins (HBPs), also called stem-loop binding proteins (SLBPs), which are responsible for the post-transcriptional regulation of histone genes(Müller and Schümperli, 1997;Dominski and Marzluff, 1999). HBP, in combination with the U7 small nuclear ribonucleoprotein particle, is required for histone RNA cleavage (Wang et al., 1996; Martin et al.,1997; Dominski et al.,1999;Müller et al., 2000). This activation of histone message is presumably the role of most of the characterised vertebrate HBPs(Fig. 1), with the exception of Xenopus SLBP2, which is responsible for translational silencing of histone mRNA in Xenopus oocytes(Wang et al., 1999). Given the crucial role of histones in the cell nucleus, it is likely that, as is the case in Drosophila (Sullivan et al., 2001), HBPs are required for development in all metazoans.

Table 1.

The C. elegans core histone genes

Chromosome/clusterH2AH2BH3H4
Replication-dependent gene clusters II/HIS3 F08G2.2 F08G2.1 F08G2.3 ZK131.1 
  ZK131.6 (his-12ZK131.5 (his-11ZK131.2 ZK131.4 (his-10
  ZK131.10 (his-16ZK131.9 (his-15ZK131.3 (his-9ZK131.8 (his-14
    ZK131.7 (his-13 
 IV/HIS5 F17E9.13 (his-33F17E9.9 (his-34F17E9.10 (his-32F17E9.12 (his-31
  F35H10.1 (his-30F35H10.11   
 IVa B0035.7 B0035.8 B0035.10 B0035.9 
 IVa F54E12.5 F54E12.4 F54E12.1 F54E12.3 
 IVa   F22B3.2 F22B3.1 
 IVa H02I12.7 H02I12.6   
 IV F55G1.10 F55G1.3 F55G1.2 F55G1.11 
 V/HIS1 T10C6.12 (his-3T10C6.11 (his-4T10C6.13 (his-2T10C6.14 (his-1
 V/HIS2 F45F2.4 (his-7F45F2.2(his-39)1 F45F2.13 (his-6F45F2.3 (his-5
   F45F2.12 (his-8  
 Vb/HIS4 K06C4.3 (his-21K06C4.4 (his-20K06C4.5 (his-17K06C4.2 (his-28
  K06C4.11 (his-19K06C4.12 (his-22) K06C4.13 (his-27K06C4.10 (his-18
 Vb F07B7.3 F07B7.4 F07B7.5 F07B7.9 
  F07B7.10 F07B7.11   
Other replication-dependent genes T23D8.6   T23D8.5 
 II   W09H1.22  
 III   E03A3.33  
 III   E03A3.44  
   F20D6.94  
   K03A1.15 K03A1.6 (his-38
Replacement variant gene cluster6 V/HIS6 C50F4.13 (his-35C50F4.5  C50F4.7 (his-37
Other replacement variant genes III   Y49E10.6  
    F58A4.3(hcp-3)2  
    F54C8.22  
 IV R08C7.37    
  ZK1251.1    
   W05B10.1  
   F45E1.6  
Histone genes were identified by sequence similarity searches of the C. elegans genome for histone sequences(Altschul et al., 1990). Histone genes that lack introns and contain the sequence encompassing the conserved hairpin structure (Fig. 1B) were categorised as replication-dependent genes, histone genes containing introns and lacking the hairpin element were categorised as replacement-variant histone genes. Gene names and clusters as originally described are indicated (Roberts et al.,1987; Roberts et al.,1989). Immediately adjacent histone genes are included into the clusters. Histone H1 genes are described elsewhere(Jedrusik and Schulze, 2001).      
Chromosome/clusterH2AH2BH3H4
Replication-dependent gene clusters II/HIS3 F08G2.2 F08G2.1 F08G2.3 ZK131.1 
  ZK131.6 (his-12ZK131.5 (his-11ZK131.2 ZK131.4 (his-10
  ZK131.10 (his-16ZK131.9 (his-15ZK131.3 (his-9ZK131.8 (his-14
    ZK131.7 (his-13 
 IV/HIS5 F17E9.13 (his-33F17E9.9 (his-34F17E9.10 (his-32F17E9.12 (his-31
  F35H10.1 (his-30F35H10.11   
 IVa B0035.7 B0035.8 B0035.10 B0035.9 
 IVa F54E12.5 F54E12.4 F54E12.1 F54E12.3 
 IVa   F22B3.2 F22B3.1 
 IVa H02I12.7 H02I12.6   
 IV F55G1.10 F55G1.3 F55G1.2 F55G1.11 
 V/HIS1 T10C6.12 (his-3T10C6.11 (his-4T10C6.13 (his-2T10C6.14 (his-1
 V/HIS2 F45F2.4 (his-7F45F2.2(his-39)1 F45F2.13 (his-6F45F2.3 (his-5
   F45F2.12 (his-8  
 Vb/HIS4 K06C4.3 (his-21K06C4.4 (his-20K06C4.5 (his-17K06C4.2 (his-28
  K06C4.11 (his-19K06C4.12 (his-22) K06C4.13 (his-27K06C4.10 (his-18
 Vb F07B7.3 F07B7.4 F07B7.5 F07B7.9 
  F07B7.10 F07B7.11   
Other replication-dependent genes T23D8.6   T23D8.5 
 II   W09H1.22  
 III   E03A3.33  
 III   E03A3.44  
   F20D6.94  
   K03A1.15 K03A1.6 (his-38
Replacement variant gene cluster6 V/HIS6 C50F4.13 (his-35C50F4.5  C50F4.7 (his-37
Other replacement variant genes III   Y49E10.6  
    F58A4.3(hcp-3)2  
    F54C8.22  
 IV R08C7.37    
  ZK1251.1    
   W05B10.1  
   F45E1.6  
Histone genes were identified by sequence similarity searches of the C. elegans genome for histone sequences(Altschul et al., 1990). Histone genes that lack introns and contain the sequence encompassing the conserved hairpin structure (Fig. 1B) were categorised as replication-dependent genes, histone genes containing introns and lacking the hairpin element were categorised as replacement-variant histone genes. Gene names and clusters as originally described are indicated (Roberts et al.,1987; Roberts et al.,1989). Immediately adjacent histone genes are included into the clusters. Histone H1 genes are described elsewhere(Jedrusik and Schulze, 2001).      
a

Histone gene-rich region between nucleotides 11,335,000 and 11,420,000.

b

Histone gene-rich region between nucleotides 8,848,000 and 8,900,000.

1

Histone H2B gene described as H4 gene containing introns. It has a near complete H2B sequence in one intron, with the conserved sequence element with the hairpin structure downstream. It is tentatively classified as a replication-dependent gene.

2

Codes for a H3-like protein.

3

Codes for a H3 protein with a truncated N-terminus.

4

Codes for a H3-like protein, lacks hairpin element. Possible pseudogene.

5

Histone H3 gene, described as containing introns. Histone H3 open reading frame with one nucleotide deletion in exon three, with the hairpin structure immediately downstream. It is close to the K03A1.6 and is tentatively classified as a replication-dependent gene

6

C50F4.6 is annotated as his-36 but codes for an unrelated protein.

7

Similar to human H2A.Z.

Fig. 1.

C. elegans histone hairpin-binding protein and the histone hairpin RNA sequence element. (A) C. elegans HBP sequence aligned with the RNA-binding domains (RBD) of other HBPs. Both the full-length protein and the RBD (underlined) bind to the C. elegans histone hairpin structure(Michel et al., 2000). (B) The conserved sequence element in the 3′ untranslated region of histone genes encompassing the hairpin structure. Conserved residues in metazoan histone RNA hairpins are circled. C11 is unique to C. elegans histone RNAs. Nucleotide position numbering is arbitrary.

Fig. 1.

C. elegans histone hairpin-binding protein and the histone hairpin RNA sequence element. (A) C. elegans HBP sequence aligned with the RNA-binding domains (RBD) of other HBPs. Both the full-length protein and the RBD (underlined) bind to the C. elegans histone hairpin structure(Michel et al., 2000). (B) The conserved sequence element in the 3′ untranslated region of histone genes encompassing the hairpin structure. Conserved residues in metazoan histone RNA hairpins are circled. C11 is unique to C. elegans histone RNAs. Nucleotide position numbering is arbitrary.

The presence of the histone hairpin structure in the C. elegansreplication-dependent histone genes indicates that the post-transcriptional regulation of these histone genes is also dependent upon HBP activity. We previously identified the C. elegans HBP homologue on the basis of its sequence similarity to vertebrate HBP(Martin et al., 1997). Sequence conservation between HBPs is highest in the RNA binding domain, and an alignment of HBP sequences in this region, highlighting the conserved residues, is shown in Fig. 1A. Hairpin sequences in the 3′ UTRs of the C. elegans core histone genes deviate at position 11 from the vertebrate hairpin consensus sequence(Fig. 1B)(Marzluff, 1992). At this position, C. elegans hairpin sequences contain a C, whereas the vertebrate sequences contain a U. Uniquely, C11 is absolutely essential for C. elegans HBP RNA binding(Michel et al., 2000), which is indicative of a highly specific interaction between HBP and hairpin RNA. The binding specificity is intrinsic to the C. elegans RNA-binding domain (Michel et al.,2000).

Here, we analyse HBP expression during C. elegans development,using a HBP promoter-green fluorescent protein (GFP) fusion construct, and investigate the role of HBP during C. elegans development, using RNA-mediated interference (RNAi) (Fire et al., 1998) to deplete the endogenous levels of HBP during both embryonic and postembryonic development. Reducing HBP function in this way results in a decrease in histone protein levels and defects in mitosis associated with improperly condensed chromosomes, confirming its proposed role in regulating histone biosynthesis. An identical defect was observed in embryos depleted of histones H3 and H4 using RNAi. These data demonstrate that HBP is an essential component required for the correct regulation of histone biosynthesis during C. elegans development.

Materials and Methods

Database searches

The analysis of the C. elegans histone genes was carried out using the Wormbase database (http://www.wormbase.org/) and the C. elegansgenome sequence(http://www.sanger.ac.uk/Projects/C_elegans/blast_server.shtml). Information about Drosophila histones was obtained from the flybase database(http://flybase.bio.indiana.edu/), and information about human histone genes was obtained from the National Centre for Biotechnology Information human genome databases (http://www.ncbi.nlm.nih.gov/genome/guide/human/).

Nematode strains

Standard C. elegans culturing techniques were used. N2 (Bristol)and MG152 were used as wild type. MG152 carries an integrated transgene expressing a GFP-tagged histone H2B(Kaitna et al., 2000). All experiments involving this strain were carried out at 25°C.

Construction of R06F6.1::GFP reporter constructs

The genomic region upstream of the R06F6.1 coding region was amplified with primers ceHBPA (5′-CAATCAGCTGTTCGCGCCGG-3′) and ceHBPB(5′-CTAGAGTCGACCTGCAGGCGTCGGCGAAATCCTCGAC-3′) from pFM#6 containing the 7,976 bp PstI fragment of T19E10 encompassing the R06F6.1 gene, and GFP with a nuclear localisation signal was amplified from plasmid pPD95.67 (a generous gift from Andy Fire and co-workers) with primers ceHBPC(5′-GTCGAGGATTTCGCCGACGCCTGCAGGTCGACTCTAG-3′) and GFPD(5′-GGGAGCTGCATGTGTCAGAG-3′) using the Expand High Fidelity PCR system (Roche). The two PCR products were then combined and the fusion product was amplified with primers ceHBPD (5′-CGGTGCGAACACACTCACGC-3′) and GFPE (5′-GGCCGACTAGTAGGAAACAG-3′) as described(Hobert et al., 1999). The fusion product contains 2923 bp upstream of the AUG (and spans the sequence between R06F6.1 and the preceding gene, T19E10.1), followed by the sequence coding for the first 41 amino acids of R06F6.1,spanning 1 intron, fused to the GFP coding region. Products were gel purified and, after diagnostic restriction digest, used to establish transformed C. elegans lines employing standard microinjection technique(Mello and Fire, 1995).

RNA interference

RNAi by injection was performed essentially as described previously(Fire et al., 1998). Sense and antisense transcripts were synthesised separately from linearised pFM#4(containing a full length HBP cDNA) using appropriate Megascript RNA synthesis kits (Ambion) and annealed at 70°C for 5 minutes, followed by 20 minutes at 37°C. Production of double-stranded RNA was verified by non-denaturing gel electrophoresis. dsRNA was injected at approximately 1 mg/ml into one gonad arm per animal (this results in RNAi effects in the progeny of both arms). Injected animals were cultured together for 6-12 hours post-injection. Progeny laid during this period typically displayed no, or weak, RNAi phenotypes and were discarded. Single injected animals were then cultured separately for 24 hours, before being transferred to a fresh plate every 24 hours until no further progeny were produced. The broods from these plates were then scored for phenotype analyses.

RNAi by feeding was carried out as described previously(Kamath et al., 2000;Timmons et al., 2001). A 580 bp BamHI XhoI R06F6.1 fragment containing HBP sequence was excised from pFM#4 and inserted into the feeding vector L4440(Timmons et al., 2001). The resultant plasmid, pPE#R7, was then transformed into HT115(DE3) bacteria. Overnight cultures of the transformed bacteria were used to seed fresh NGM plates containing 1 mg/ml IPTG and 25 μg/ml carbenicillin. L4 larvae, or eggs harvested from gravid hermaphrodites, were then added to the plates and grown at 20°C unless otherwise stated. In the case of L4 larvae, the animals were transferred to fresh plates 24 hours after the beginning of egglaying and then transferred again every 24 hours until no more fertilised oocytes were produced. The progeny laid during the first 24 hours were discarded since these sometimes showed weaker, incomplete phenotypes. Where eggs were used to initiate cultures, 10-20 eggs were co-cultured to adulthood on the same seeded plate, and the postembryonic phenotypes monitored throughout this period. A 981 bp fragment encompassing the full his-10 gene (a histone H4 gene) and the his-9 gene (a histone H3 gene) lacking the most 3′ 14 bp was amplified directly from C. elegans genomic DNA using the primers ceHis34F(5′-TTATCCTCCGAATCCGTACA-3′) and ceHis34R(5′-CTCGGATACGTCTTGCCAATT-3′) using the Expand High Fidelity PCR System (Roche). The fragment was inserted into pGEM-Teasy (Promega), analysed by DNA sequencing and excised with EcoRI, and finally inserted into the feeding vector L4440 to produce plasmid pPE#R11.

Western blotting

Protein extracts were prepared as described previously(http://info.med.yale.edu/mbb/koelle/protocol_list_page.html). To obtain R06F6.1(RNAi)-treated animals for preparation of protein extracts,newly hatched L1 larvae were grown on HT115 bacteria transformed with pPE#R7 until they reached adulthood. Extracts were analysed by 15% SDS-PAGE. Proteins were transferred onto Hybond-P membrane (Amersham Pharmacia Biotech) using a semi-dry electroblotting system. Rabbit polyclonal anti-histone H3 antibodies,goat polyclonal anti-histone H4 antibodies raised against a C-terminal epitope(Santa Cruz Biotechnology Inc) and mouse monoclonal anti-tubulin antibodies(Sigma) were used as primary antibodies. Secondary antibodies were anti-rabbit IgG HRP and anti-goat IgG HRP conjugate (Sigma) and anti-mouse IgG HRP conjugate (Promega). Antibodies were detected using ECL western blotting detection reagents (Amersham Pharmacia Biotech)

Immunofluorescent detection of histone H3

Embryos were dissected from gravid hermaphrodites in egg salts(Edgar, 1995), attached to poly-L-lysine slides and permeabilised by freeze-cracking(Miller and Shakes, 1995). The embryos were fixed for 5 minutes in methanol at -20°C, followed by 5 minutes in acetone at -20°C. They were then re-hydrated through an ethanol/PBS wash series, blocked in 30% donkey serum for an hour and incubated with rabbit anti-histone H3 antibody (Santa Cruz Biotechnology Inc), diluted 1:100 in PBS, overnight at -4°C. After three 10 minute PBS washes, the slides were incubated for 45 minutes with a rhodamine-conjugated goat anti-rabbit secondary antibody (TCS Biological), diluted 1:50 in PBS at room temperature. They were then washed three times in PBS (the first wash including 1 μg/ml 4′,6-diamidino-2-phenolindole dihydrochloride(DAPI) and mounted in antifade reagent. Fluorescent images of fixed embryos were obtained as Z-series captured using a Bio-Rad MRC1024 confocal laser scanning microscope and processed using Confocal Assistant and Adobe Photoshop.

Results

C. elegans histone genes

The conserved hairpin structure in the 3′ untranslated region of replication-dependent histone genes is the binding site for HBPs(Fig. 1) and is required for the regulation of gene expression. We decided to investigate the role of HBPs during development using C. elegans as animal model system. To identify the potential targets for regulation by HBP, we surveyed the C. elegans histone genes. Database searches revealed that the C. elegans genome contains 64 histone genes that have all the hallmarks of the replication-dependent histone genes: they lack introns and encode the RNA hairpin structure downstream of the stop codon(Table 1)(Roberts et al., 1989). These code for the core histones H2A, H2B, H3 and H4, and 58 of these genes are in clusters on chromosomes II, IV and V(Roberts et al., 1987). Normally in these clusters H2A and H2B genes, and H3 and H4 genes, are paired and divergently transcribed. In addition, there are two pairs of replication-dependent histone genes on chromosomes I and X. The search also detected two further genes coding for replication-dependent histone H3 gene variants (E03A3.3 and W09H1.2, on chromosomes III and II,respectively). In addition, near one of these genes (E03A3.3) there is a related sequence (E03A3.4) that is lacking a hairpin structure in the 3′ untranslated region and may be a pseudogene. A further H3-like gene lacking the hairpin structure (F20D6.9) is located on chromosome V.

The C. elegans replication-dependent histone gene clusters lack genes for the linker histone H1 family(Roberts et al., 1987), but there are eight genes coding for histone H1 proteins that have the structure of replacement variant histone genes, that is they contain introns and lack the conserved RNA hairpin in the 3′ untranslated region(Jedrusik and Schulze, 2001). C. elegans also has a small number of core histone genes with introns. Replacement variant genes for one histone H2A, H2B and H4 each are located on chromosome V, two more histone H2A genes are on chromosome IV; and three further histone H3 genes are on chromosomes III, V and X, respectively. In addition, two H3-like genes (F58A4.3, F54C8.2) are located on chromosome V.

The apparent lack of replication-dependent C. elegans histone H1 genes is unusual. Replication-dependent histone H1 genes are found from Drosophila to man. In Drosophila, the replication-dependent histone genes are organised as discontinuous arrays of ∼100 repeats containing histone H2A, H2B, H3, H4 and H1 genes on chromosome 2(Pardue et al., 1977;Matsuo and Yamazaki, 1989). In addition, single copies of replacement variant histone genes, H2A and H4, and two copies of replacement variant Histone H3 were described(Adams et al., 2000). In humans, genome sequence data indicate that at least 42 replication-dependent core histone and five replication-dependent H1 histone genes, as well as a testis-specific H1 gene are clustered on chromosome 6. A further group of at least three replication-dependent histone genes is located on chromosome 1. At least another nine histone genes, including replacement variant histone,testis-specific histone and macroH2A histone genes were found singly on different chromosomes (Albig and Doenecke,1997). Other experimental data suggest that the number of human histone genes may be higher (Albig et al.,1993; Albig et al.,1997; Albig and Doenecke,1997). In conclusion, the number and organisation of histone genes in humans and C. elegans is similar: replication-dependent genes are clustered, and a low number of other histone genes are distributed throughout the genome. One difference is that C. elegans is apparently lacking replication-dependent histone H1 genes but has a similar number of replacement variant H1 genes.

The majority of C. elegans histone genes are of the replication-dependent type. We have previously described the interaction between HBP and the RNA hairpin structure in the 3′ untranslated region of these genes (Michel et al.,2000). Here, we test directly whether the C. elegans HBP is involved in regulation of histone gene expression.

R06F6.1::GFP reporter construct is expressed in all somatic cells throughout development

The C. elegans HBP is the gene product of the gene R06F6.1. To determine where R06F6.1 is expressed, we generated transgenic lines that expressed a GFP reporter construct under the control of R06F6.1 upstream sequences. Several lines were generated and all displayed the same expression pattern. Nuclear GFP expression was detected in all cells during embryonic and postembryonic development, with expression appearing strongest in actively dividing cells(Fig. 2). This was particularly obvious during postembryonic development, where strong GFP expression was restricted to proliferating cells and cells undergoing endoreduplication(Fig. 2B,C). For example, the cells of the lateral hypodermis divide in a stem-cell-like fashion throughout postembryonic development and show very strong GFP expression compared to cells that have left the cell cycle (Fig. 2B,C). Thus, HBP promoter activity is highest in cells undergoing DNA replication, consistent with a role for HBP in regulating histone gene expression.

Fig. 2.

R06F6.1::gfp is expressed in all somatic cells throughout development. (A) Confocal Z-series projection of an embryo approximately 300 minutes after the first cell division. (B) An epifluorescent image of the expression of R06F6.1::GFP in an L2 larva. The actively dividing nuclei of the lateral hypodermal cells are indicated by arrowheads, and an intestinal nucleus (which undergoes endoreduplication during postembryonic development) is indicated by an arrow. These cells show stronger GFP expression than non-dividing cells. (C) Epifluorescence image showing that nuclei of recently divided cells strongly express R06F6.1::GFP. The cells that form the postdeirid (boxed) and two lateral hypodermal cells(arrows) show stronger expression than neighbouring cells. Scale bars represent 10 μm.

Fig. 2.

R06F6.1::gfp is expressed in all somatic cells throughout development. (A) Confocal Z-series projection of an embryo approximately 300 minutes after the first cell division. (B) An epifluorescent image of the expression of R06F6.1::GFP in an L2 larva. The actively dividing nuclei of the lateral hypodermal cells are indicated by arrowheads, and an intestinal nucleus (which undergoes endoreduplication during postembryonic development) is indicated by an arrow. These cells show stronger GFP expression than non-dividing cells. (C) Epifluorescence image showing that nuclei of recently divided cells strongly express R06F6.1::GFP. The cells that form the postdeirid (boxed) and two lateral hypodermal cells(arrows) show stronger expression than neighbouring cells. Scale bars represent 10 μm.

Depleting HBP function in early embryos results in cell division arrest

We used RNAi to address the function of the C. elegans HBP. Depleting HBP in the early embryo, either by growing the parent hermaphrodites on R06F6.1 dsRNA expressing bacteria or by injecting R06F6.1dsRNA into the syncitial germline of the parent hermaphrodite, led to the same embryonic lethal phenotype (Table 2). The first few cell divisions occurred with approximately the same timing as was observed for untreated, wild-type embryos, but after the embryos reached approximately 30 cells in size, no further cell divisions were observed (data not shown).

Table 2.

RNAi of HBP and histone H3/H4 results in similar, but distinct,phenotypes

Phenotype*
RNAi targetEmbSteEvl/Pvl§MuvGro#
R06F6.1 100 (>500) 100 (>200) 91 (161) 0 (>200) 0 (200) 
his-9/10 100 (>200) 100 (>200) 0 (>200) 19 (36) 98% (>200) 
Phenotype*
RNAi targetEmbSteEvl/Pvl§MuvGro#
R06F6.1 100 (>500) 100 (>200) 91 (161) 0 (>200) 0 (200) 
his-9/10 100 (>200) 100 (>200) 0 (>200) 19 (36) 98% (>200) 
*

Percentage of total number of animals scored (in brackets) that displayed each phenotype, early embryonic arrest, sterile, no progeny produced,

§

everted/ protruding vulva. one or more ectopic pseudovulvae and #slow growth accompanied by reduced body size and impaired movement.

In order to better characterise the molecular basis of the cell-proliferation defects that we observed in R06F6.1(RNAi) embryos,we used a strain that expresses a histone H2B-GFP fusion protein, which allows the visualisation of the chromosomes in intact, living embryos(Kaitna et al., 2000). The 3′ untranslated region of this fusion protein is derived from the unc-54 gene and therefore is not dependent upon HBP for its expression. Using this strain we followed the chromosomal behaviour during development in R06F6.1(RNAi) embryos(Fig. 3). Abnormalities in chromosomal morphology became apparent during the first mitotic division of these embryos, with the chromosomes appearing less well condensed during metaphase and anaphase, than in wild-type embryos; however it was not until the second mitotic division that more severe defects became apparent. Chromatin bridges were evident between the two AB daughter cells upon reaching the end of anaphase, and cytokinesis occurred even though some chromatin remained at the midline (Fig. 3C,D). Although the nuclear morphology of these AB daughter cells appeared abnormal as visualised by differential interference contrast (DIC)optics, they initiated the next round of cell division appropriately. The P1 blastomere division, which occurs 2-3 minutes later, was less abnormal since,although the chromosomes appeared less well condensed than wild type during metaphase and anaphase, no chromatin bridges were evident between the P1 daughter cells. The nuclear morphology of the P1 daughter cells under DIC optics was normal (Fig. 3C,D). Chromosome condensation during all subsequent cell divisions became increasingly abnormal, with the chromosomes forming irregular condensations,rather than tight metaphase plates, accompanied by the presence of chromatin bridges between daughter cells at the end of anaphase(Fig. 3G,H).

Fig. 3.

Depleting HBP levels in the early embryo results in defects in chromatin structure. (A) A DIC photomicrograph of a wild-type embryo immediately after the division of the AB and P1 blastomeres, which generates ABa/ABp and EMS/P2,respectively. (B) An epifluorescent image of the same embryo in A showing the H2B::GFP expression associated with the newly divided nuclei. The arrowhead indicates one of the polar bodies that has been drawn between the two AB nuclei upon cytokinesis (the other polar body is visible at the anterior of the embryo). (C) A DIC photomicrograph of an embryo derived from an animal raised on R06F6.1 dsRNA expressing HT115 bacteria at an equivalent developmental stage to the embryo in (A,B). (D) An epifluorescence image of the same embryo as in (C), with H2B::GFP labelling the chromatin. Arrow indicates fluorescent material present between the dividing cells. Note the presence of the polar body (arrowhead), which indicates that this cell has undergone cytokinesis. (E) A DIC photomicrograph of a wild-type 4-cell embryo. Note that all four blastomeres have round nuclei. (F) A DIC photomicrograph of a his-9/10(RNAi) 4-cell embryo showing abnormal nuclear morphology in the two AB daughter cells (white arrowheads). (G,H). Epifluorescence images of wild-type (G) and R06F6.1(RNAi) (H) fixed embryos stained with DAPI. Arrows indicate cells in anaphase showing the poorly condensed chromosomes of R06F6.1(RNAi) embryos compared to wild- type. Anterior is to the left in all panels. Scale bar represents 10 μm.

Fig. 3.

Depleting HBP levels in the early embryo results in defects in chromatin structure. (A) A DIC photomicrograph of a wild-type embryo immediately after the division of the AB and P1 blastomeres, which generates ABa/ABp and EMS/P2,respectively. (B) An epifluorescent image of the same embryo in A showing the H2B::GFP expression associated with the newly divided nuclei. The arrowhead indicates one of the polar bodies that has been drawn between the two AB nuclei upon cytokinesis (the other polar body is visible at the anterior of the embryo). (C) A DIC photomicrograph of an embryo derived from an animal raised on R06F6.1 dsRNA expressing HT115 bacteria at an equivalent developmental stage to the embryo in (A,B). (D) An epifluorescence image of the same embryo as in (C), with H2B::GFP labelling the chromatin. Arrow indicates fluorescent material present between the dividing cells. Note the presence of the polar body (arrowhead), which indicates that this cell has undergone cytokinesis. (E) A DIC photomicrograph of a wild-type 4-cell embryo. Note that all four blastomeres have round nuclei. (F) A DIC photomicrograph of a his-9/10(RNAi) 4-cell embryo showing abnormal nuclear morphology in the two AB daughter cells (white arrowheads). (G,H). Epifluorescence images of wild-type (G) and R06F6.1(RNAi) (H) fixed embryos stained with DAPI. Arrows indicate cells in anaphase showing the poorly condensed chromosomes of R06F6.1(RNAi) embryos compared to wild- type. Anterior is to the left in all panels. Scale bar represents 10 μm.

R06F6.1 is required for postembryonic divisions

During postembryonic development, the 550 cells present in the newly hatched larva are added to by the proliferation of multiple cell lineages(Sulston and Horvitz, 1977). To determine whether HBP is required during postembryonic cell division we examined the effect of depleting HBP function by growing newly hatched larvae on R06F6.1 dsRNA-expressing bacteria. Such R06F6.1(RNAi)larvae grow up to become sterile adults, which almost invariably show an abnormally everted vulva (Ev1 phenotype)(Table 2;Fig. 4).

Fig. 4.

Late larval lineages are affected by depleting HBP expression during postembryonic development. DIC photomicrographs of postembryonic developmental stages are shown. (A) A vulva and uterus of a wild-type mid-L4 stage larva,showing the so-called `Christmas tree' morphology of the vulva at this developmental stage and obvious uterine cavity (ut). The arrow indicates the lamina that separates the uterine cavity from the vulva. (B) Abnormal vulva and uterus in a mid-L4 stage larvae fed R06F6.1 dsRNA from hatching is shown. Vulval morphology is grossly abnormal and the uterine cavity is completely absent, owing to multiple failures in the execution of uterine development. Note also the presence of germline nuclei proximal to the vulva.(C) Adult wild-type vulva. (D) Evl phenotype of adult fed R06F6.1dsRNA from hatching. (E) Wild-type male tail, showing the spicules (arrow). Sensory rays are not visible in this focal plane. (F) R06F6.1(RNAi)male tail showing crumpled spicules (arrow). Scale bars represent 10 μm in(A,B,E,F) and 20 μm in (C,D).

Fig. 4.

Late larval lineages are affected by depleting HBP expression during postembryonic development. DIC photomicrographs of postembryonic developmental stages are shown. (A) A vulva and uterus of a wild-type mid-L4 stage larva,showing the so-called `Christmas tree' morphology of the vulva at this developmental stage and obvious uterine cavity (ut). The arrow indicates the lamina that separates the uterine cavity from the vulva. (B) Abnormal vulva and uterus in a mid-L4 stage larvae fed R06F6.1 dsRNA from hatching is shown. Vulval morphology is grossly abnormal and the uterine cavity is completely absent, owing to multiple failures in the execution of uterine development. Note also the presence of germline nuclei proximal to the vulva.(C) Adult wild-type vulva. (D) Evl phenotype of adult fed R06F6.1dsRNA from hatching. (E) Wild-type male tail, showing the spicules (arrow). Sensory rays are not visible in this focal plane. (F) R06F6.1(RNAi)male tail showing crumpled spicules (arrow). Scale bars represent 10 μm in(A,B,E,F) and 20 μm in (C,D).

The sterile phenotype suggested possible defects in the proliferation of the germ cells. The germline in C. elegans originates from two germline precursor cells, which proliferate mitotically during the first three larval stages (Kimble and Hirsh,1979). This phase of germline development is unaffected in R06F6.1(RNAi) larvae, and most (82%; n=32) animals are able to produce the first set of mature germ cells, which develop as sperm. In wild-type animals, the later germ cells mature into oocytes; however, in R06F6.1(RNAi) animals very few mature oocytes were produced(n=32), and germline proliferation appeared to cease soon after reaching adulthood. Thus, it is likely that the sterility reflects a combination of failure of mitosis in the adult germline coupled with defects in the maturation of the germ cells into oocytes.

We also examined the development of the vulva and the uterus in R06F6.1(RNAi) animals, since the Ev1 phenotype arises from defects in these structures (Seydoux et al.,1993). The vulva is produced from cells that divide and undergo morphogenesis during the third and fourth larval stages (L3 and L4,respectively) (Sulston and Horvitz,1977; Sternberg and Horvitz,1986). At the same time, the uterus and spermathaecae are generated from the somatic gonad precursor cells(Newman et al., 1996). In R06F6.1(RNAi) larvae, the three vulval precursor cells, P5.p, P6.p and P7.p are induced to form vulval cells as in wild type, but fewer than wild type numbers of vulval cells are generated from these precursors, resulting in abnormal vulval morphogenesis (Fig. 4). Similarly, the later divisions of the somatic gonad cells,which generate the uterus and spermathecae, also fail to occur, resulting in the absence of these structures (Fig. 4).

Postembryonic development is also defective in R06F6.1(RNAi)males. Like the vulva and uterus, the male copulatory apparatus is also generated during mid-late postembryonic development, from a limited set of blast cells that proliferate at around the same time as the hermaphrodite uterus and vulva (Sulston et al.,1980). R06F6.1(RNAi) males display defects in the morphology of this organ (100%, n=24)(Fig. 4). Although we have not followed the proliferation of the cell lineages involved in the development of the copulatory structures in R06F6.1(RNAi) males, the defects we observed are consistent with failures in the generation of the cells responsible for producing them.

Thus, R06F6.1(RNAi) results in multiple defects in postembryonic somatic cell divisions that are consistent with the cell division defects that we observed in R06F6.1(RNAi) embryos. There are however many cell lineages that proliferate during early larval development, and it is striking that we did not observe defects in early cell proliferation events. Mutations that affect all postembryonic cell divisions result in a sterile, thin,uncoordinated phenotype reflecting failures in neuroblast and epidermal cell divisions that generate the mature nervous system and epidermis, respectively(Albertson et al., 1978;Sulston and Horvitz, 1981;O'Connell et al., 1998). That we did not see such defects in R06F6.1(RNAi) larvae and adults may be attributable to the fact that many of these cells are generated during early development, before significant depletion of HBP function has been induced by the ingested dsRNA. However, it has been observed that the nervous system is generally refractile to RNAi effects (Fire et al., 1998), and this may also account for the lack of defects associated with the generation of this tissue in R06F6.1(RNAi)animals.

HBP is required for histone gene expression

Taken together, our observations indicate that C. elegans HBP is required for mitosis during both embryonic and postembryonic development. The most likely explanation for this is a reduction in histone expression, caused by decreased HBP-dependent post-transcriptional modification of histone RNA. In order to confirm this hypothesis, we examined the levels of endogenous H3 and H4 histones in both wild-type and R06F6.1(RNAi) animals. Using western blot analysis of protein extracts derived from adult worms fed R06F6.1 dsRNA from hatching, we observed a reduction in the levels of both H3 and H4 histones when compared with extracts prepared from wild-type adults, with H4 histone expression being reduced beyond the level of detection(Fig. 5A). It is not clear why there are apparent differences in the reduction of histone H3 and H4 levels in R06F6.1(RNAi) animals. It may be that the histone H3 signal is stronger because of cross-reactivity of this antibody with the replacement variants histone H3 proteins and other H3-like proteins(Table 1). Alternatively, this may reflect a real difference in histone H3 and H4 protein levels in response to reduced HBP function.

Fig. 5.

HBP is required for histone gene expression. (A) Extracts prepared from wild-type and R06F6.1(RNAi) C. elegans were analysed on a 15%SDS-PAGE. Proteins were detected by western blotting using antibodies against histone H3 (lanes 1, 2) or histone H4 (3, 4), as well as with an anti-tubulin antibody to control for loading. (B,C) Epifluorescence image showing immunostaining with anti-histone H3 antibody of a wild-type (B) and an R06F6.1(RNAi) embryo (C). Scale bars represent 10 μm.

Fig. 5.

HBP is required for histone gene expression. (A) Extracts prepared from wild-type and R06F6.1(RNAi) C. elegans were analysed on a 15%SDS-PAGE. Proteins were detected by western blotting using antibodies against histone H3 (lanes 1, 2) or histone H4 (3, 4), as well as with an anti-tubulin antibody to control for loading. (B,C) Epifluorescence image showing immunostaining with anti-histone H3 antibody of a wild-type (B) and an R06F6.1(RNAi) embryo (C). Scale bars represent 10 μm.

The reduction of histone H3 upon treatment with R06F6.1(RNAi) was confirmed by immunostainings in whole, fixed embryos with the anti-histone H3 antibody (Fig. 5B,C) (in our hands, the anti-histone H4 antibody did not recognise its epitope in fixed embryos, using any of the commonly available fixation protocols). Using the anti-H3 antibody, all nuclei were immunostained in 90% of wild-type embryos tested (n=30). In contrast, only 17% of R06F6.1(RNAi)embryos (n=35) showed equivalent levels of immunostaining.

Histone gene expression is required for embryonic and postembryonic development

As reducing HBP function led to reduced histone expression, it seemed likely that this was the cause of the RNAi phenotypes we observed. To further confirm this, we tested whether reducing histone expression directly using RNAi would reproduce the R06F6.1(RNAi) phenotypes. We therefore carried out RNAi against his-9 and his-10 (which encode histone H3 and H4 paralogues, respectively). The nucleotide sequences of both replication-dependent and replacement histone H4 genes are >85% identical,and nucleotide sequence identity between histone H3 genes exceeds 77%, with the replication-dependent H3 genes being >88% identical. Such levels of identity mean that dsRNA derived from his-9 and his-10 is likely to inhibit the expression of all H3 and H4 genes, leading to a general depletion of histone H3 and H4 proteins. his-9/10(RNAi) resulted in an embryonic-lethal phenotype (Table 2), similar to previous experiments with the his-10 gene(Maeda et al., 2001), as well as with a replacement histone H3 gene (Y49E10.6) and a replication-dependent H2A gene (T23D8.6)(Gonczy et al., 2000;Fraser et al., 2000). In contrast, a reduced lethality was observed when another replication-dependent histone H4 gene was depleted (T23D8.5)(Fraser et al., 2000). his-9/10(RNAi) embryos arrested at the same stage of development as R06F6.1(RNAi) embryos (Table 2) (data not shown). However, his-9/10(RNAi) additionally affected the localisation of the H2B::GFP fusion protein, which resulted in significant decreases in the amount of chromosomal fluorescence. This was accompanied by increased levels of diffuse cytoplasmic and nuclear fluorescence (data not shown), suggesting that the H2B fusion protein, while still expressed, failed to localise to the chromosomes correctly. Assembly of H2B::GFP into nucleosomes is almost certainly dependent upon the presence of endogenous histones. Therefore, the greatly reduced incorporation of H2B::GFP into chromatin is likely to reflect reduced levels of endogenous H3 and H4 core histones caused by his-9/10(RNAi). Since we did not observe the same reduction in chromosomal fluorescence in R06F6.1(RNAi) embryos,we take this to mean that his-9/10(RNAi) may be more effective at depleting H3 and H4 histone levels than R06F6.1(RNAi). This may be due to reduction in the expression of the replacement histone genes in addition to the replication-dependent genes. Alternatively, a fraction of the histone mRNAs may acquire a poly(A) tail in a HBP-depleted background, as is the case in Drosophila (Sullivan et al., 2001), which allows for translation of these mRNAs. Despite this difference, his-9/10(RNAi) embryos exhibited identical defects in chromosome behaviour to those we observed in R06F6.1(RNAi)embryos (Fig. 3F). This supports our conclusion that the chromosomal defects in embryos depleted for HBP function are due to reduced levels of histone biosynthesis.

We also examined the effect of depleting histone expression during postembryonic development, and in this case we observed differences between the R06F6.1(RNAi) and the his-9/10(RNAi) postembryonic phenotypes (Table 2). Whereas R06F6.1(RNAi) larvae displayed a normal growth rate, and appeared essentially wild type under the dissecting microscope until the late L4/adult stage, his-9/10(RNAi) larvae grew slowly, were thin and displayed a translucent appearance under the dissecting microscope. Those animals that reached adulthood were sterile, as we found for the R06F6.1(RNAi)animals. In some cases his-9/10(RNAi) adults also displayed a multivulval (Muv) phenotype (Table 2; Fig. 6),characteristic of the inappropriate activation of the vulval cell fate in the vulval precursor cells that normally adopt a non-vulval cell fate(Fay and Han, 2000). Thus,although displaying the sterility observed in R06F6.1(RNAi) animals, his-9/10(RNAi) results in additional postembryonic defects, including abnormal regulation of cell fate, not observed in R06F6.1(RNAi)animals.

Fig. 6.

his-9/10(RNAi) causes defects in vulval precursor cell fate. (A) A DIC photomicrograph of the central region of a wild-type adult hermaphrodite.(B) A DIC photomicrograph of a his-9/10(RNAi) adult hermaphrodite displaying a Muv phenotype. Arrowheads indicate the vulva in both animals. The ectopic pseudovulvae are indicated by asterisks. Scale bar represents 50μm.

Fig. 6.

his-9/10(RNAi) causes defects in vulval precursor cell fate. (A) A DIC photomicrograph of the central region of a wild-type adult hermaphrodite.(B) A DIC photomicrograph of a his-9/10(RNAi) adult hermaphrodite displaying a Muv phenotype. Arrowheads indicate the vulva in both animals. The ectopic pseudovulvae are indicated by asterisks. Scale bar represents 50μm.

Discussion

The C. elegans HBP binds tightly and with great sequence specificity to the hairpin structure in the 3′ untranslated region of C. elegans histone mRNA that is required for histone gene expression(Michel et al., 2000). It is therefore reasonable to assume that the C. elegans HBP is involved in regulating histone gene expression at the post-transcriptional level. The work presented here provides evidence supporting this hypothesis. Firstly, we showed that the HBP gene promoter is most active in dividing cells, which require histone biosynthesis in order to package the newly replicated DNA into chromatin. Secondly, we showed that depleting the function of HBP results in failure of early embryonic cell divisions, apparently caused by abnormalities in chromosome condensation. Thirdly, we were able to demonstrate that HBP depletion decreases histone protein levels. Finally, we showed that depleting early embryos of histones H3 and H4 produces the same phenotype as HBP depletion. These findings indicate that HBP is an essential component of the histone biosynthesis machinery in C. elegans.

Our observation that HBP promoter activity, as measured by the expression of GFP fused to the HBP promoter region, is highest in dividing cells is in apparent contradiction to the observation that in synchronised cell culture,HBP levels are regulated post-transcriptionally, with mRNA levels being constant throughout the cell cycle(Whitfield et al., 2000). However the situation in C. elegans may be different as during postembryonic development cells do not continuously go through subsequent cell cycles, but cell division and DNA replication are programmed and can occur after long periods of quiescence. Thus, transcriptional regulation of C. elegans HBP may reflect this difference in the mode of cell division. However, from our current data we cannot address possible post-transcriptional regulation of HBP expression.

The phenotype of R06F6.1(RNAi) embryos is similar to the phenotype of Drosophila embryos lacking the maternal contribution of the Drosophila HBP homologue, dSLBP(Sullivan et al., 2001). As in C. elegans, dSLBP is required for mitotic cell divisions in the early embryo, and loss of maternal dSLBP function results in defects associated with chromosome condensation. Similarly, absence of maternal dSLBP, as we found for R06F6.1(RNAi), does not prevent the first few mitotic divisions, but results in the gradual accumulation of defects in chromosomal condensation. We take this to indicate that sufficient processed core histone mRNAs, or core histone proteins, are provided maternally to support these first few divisions and that in wild-type embryos these limited supplies of histones are supplemented by the HBP-dependent expression from maternally loaded,unprocessed histone mRNA or newly synthesised zygotic histone mRNAs. It is also possible that in HBP-depleted embryos, as was found for Drosophila (Sullivan et al.,2001), polyadenylation of histone mRNA may occur that would result in some histone synthesis, thereby allowing a limited number of cell divisions.

Reducing HBP/SLBP function in the early embryo would thus result in a chromosome condensation defect that grew worse as the limited source of histones became depleted, until insufficient histones were present to allow the packaging of newly synthesised DNA into chromosomes, at which point cell division would cease. This is what was observed in Drosophila embryos with a reduced dSLBP function and is also what we found in the R06F6.1(RNAi) embryos.

Although depletion of histones H3 and H4 caused a similar phenotype to depleting HBP in early embryos, his-9/10(RNAi) larvae exhibited a more severe postembryonic phenotype compared to R06F6.1(RNAi)animals. Some of the differences may be caused by a more severe reduction in histone synthesis in his9/10(RNAi) animals. This was suggested by the finding that the chromosomal localisation of the H2B::GFP fusion protein was significantly impaired by depleting H3 and H4 histones but was not appreciably affected by reducing HBP function. Higher histone synthesis in R06F6.1(RNAi) animals may be due to the expression of the replacement variant histone genes. In addition, it is possible that this is supplemented by replication-dependent histone gene expression brought about by some residual HBP synthesis or by some polyadenylation of histone RNA as discussed above.

More curious is the discrepancy between the effect of R06F6.1(RNAi) and his-9/10(RNAi) on the fates of the vulva precursor cells. The Muv phenotype in his-9/10(RNAi) animals may be caused by a general reduction in histone proteins. In yeast, depletion of histone H4 protein does not lead to a general change in transcription(Kim et al., 1988); however in at least one case this leads to gene activation, presumably due to a change of the nucleosome structure of the promoter sequence(Han et al., 1988). Thus it is possible that local changes in chromatin structure, brought about by the depletion of histones, may lead to the aberrant expression of genes involved in vulval cell fate determination. Alternatively, since the his-9 and his-10 genes also show high nucleotide identity with replacement variant histones (i.e. histones not regulated by the HBP-hairpin interaction),it is possible that the his-9/10(RNAi) inhibits the expression of replacement variant histone genes that may be essential for vulval cell fate. Reducing the function of the linker histone H1.1 results in the activation of otherwise silent transgenes in the germline(Jedrusik and Schulze, 2001). Thus, it is possible that the inhibition of one or more of the replacement variant histone genes results in the defects in vulval precursor cell fate that we observed. Further characterisation of the individual role of these proteins will shed light on their involvement in the regulation of vulval cell fate.

Acknowledgements

We thank Franck Martin for plasmids pFM#4 and pFM#6 containing HBP genomic and cDNA, and Andy Fire and co-workers for the GFP-based reporter construct and RNAi feeding vector. Some C. elegans strains were obtained from the C. elegans Genetic Stock Centre, which is funded by a grant from the NIH National Centre for Research Support. This work was supported by the Wellcome Trust (Grants to J.P. and B.M.) and The Royal Society (J.P.).

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