Histone Gene Complement, Variant Expression, and mRNA Processing in a Urochordate Oikopleura dioica that Undergoes Extensive Polyploidization

Mariacristina Chioda, Ragnhild Eskeland and Eric M. Thompson2

Sars International Centre for Marine Molecular Biology, Bergen High Technology Centre, Bergen, Norway


    Abstract
 TOP
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Acknowledgements
 References
 
Considerable data exist on coding sequences of histones in a wide variety of organisms. Much more restricted information is available on total histone gene complement, gene organization, transcriptional regulation, and histone mRNA processing. In particular, there is a significant phylogenetic gap in information for the urochordates, a subphylum near the invertebrate-vertebrate transition. In this study, we show that the appendicularian Oikopleura dioica has a histone gene complement that is similar to that of humans, though its genome size is 40- to 50-fold smaller. At a total length of 3.5 kb, the H3, H4, H1, H2A, and H2B quintet cluster is the most compact described thus far, but despite very rapid early developmental cleavage cycles, no extensive tandem repeats of the cluster were present. The high degree of variation within each of the complements of O. dioica H2A and H2B subtypes resembled that found in plants as opposed to more closely related vertebrate and invertebrate species, and developmental stage–specific expression of different subtypes was observed. The linker histone H1 was present in relatively few copies per haploid genome and contained short N- and C-terminal tails, a feature similar to that of copepods but different from many standard model organisms. The 3'UTRs of the histone genes contained both the consensus stem-loop sequence and the polyadenylation signals but lacked the consensus histone downstream element that is involved in the processing of histone mRNAs in echinoderms and vertebrates. Two types of transcripts were found, i.e., those containing both the stem-loop and a polyA tail as well as those cleaved at the normal site just 3' of the stem-loop. The O. dioica data are an important addition to the limited number of eukaryotes for which sufficiently extensive information on histone gene complements is available. Increasingly, it appears that understanding the evolution of histone gene organization, transcriptional regulation, and mRNA processing will depend at least as much on comparative analysis of constraints imposed by certain life history features and cell biological characteristics as on projections based on simple phylogenetic relationships.


    Introduction
 TOP
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Acknowledgements
 References
 
The histones are a fundamental adaptation to the molecular processes occurring in the nucleus of a eukaryotic cell. The nucleosomal histone octamer, consisting of a central tetramer of two molecules of histone H3 and two molecules of histone H4, flanked by two histone H2A-H2B dimers, serves a structural role of efficiently packaging DNA in the nucleus. Although this structural role of the histones is arguably the best characterized aspect of their function, it is far from a static one, and the core and linker histones also collaborate intimately in transcriptional regulation, DNA repair, DNA replication, heterochromatin formation, chromosome condensation, and cell cycle progression. These regulatory activities are mediated by modifications of the N- and C-terminal tails of the histone molecules and through the use of a variety of histone variants.

Specific amino acid residues in the histone tails can be covalently modified through acetylation, methylation, phosphorylation, poly(ADP-ribosylation), and ubiquitination. These alterations may change interactions of the histone tails with DNA or other regulatory proteins (or both). It has been proposed that different combinations of these modifications on individual nucleosomes, or over regions of chromatin, may be an epigenetic code that is interpreted by downstream protein complexes in carrying out nuclear activities (Jeppesen 1997Citation ; Strahl and Allis 2000Citation ; Turner 2000Citation ). The code also may be important in mechanisms of cell memory in maintaining patterns of gene expression after mitotic cell division and in defining restricted outcomes in development and cellular differentiation. Current understanding of the histone code is extremely rudimentary, but already certain residues appear to have crucial importance. For example, acetylation of lysine 9 in the N-terminal tail of histone H3 favors the location of the associated locus in euchromatin, whereas methylation of this same residue is more characteristic of an address in heterochromatic regions (Nakayama et al. 2001Citation ; Rice and Allis 2001Citation ).

The regulatory repertoire of chromatin is not restricted to covalent modifications of histone tails and also includes the use of distinct histone variants. The use of variants is regulated both spatially and temporally. Homologues of the mammalian histone H3–like variant CENP-A occur throughout eukaryotes and are specifically found in centromeric regions of chromatin (Sullivan 2001Citation ). The variant H2A-Bbd is markedly deficient on the inactive X chromosome of mammalian cells (Chadwick and Willard 2001Citation ), a chromosome that also is enriched in hypoacetylated histone H4 (Jeppesen and Turner 1993Citation ). On a more local scale, the H2A.Z variant is preferentially linked to intergenic DNA in yeast at the PHO5 and GAL1 loci, and the extent of linkage varies with the state of transcriptional activation (Santisteban, Kalashnikova, and Smith 2000Citation ). In a temporal sense, one of the earliest observations on histone variants was the use of specific subtypes during cleavage stages at the onset of embryonic development. In the sea urchin, developmental variants of histones H2A, H2B, and linker histone H1 have been described (Mandl et al. 1997Citation ). Embryonic variants of histone H1 also have been found in Xenopus (Dworkin-Rastl, Kandolf, and Smith 1994Citation ), Drosophila (Ner and Travers 1994Citation ), and most recently, the mouse (Tanaka et al. 2001Citation ) and may be a general feature of early development. Later in development, the use of H1° linker variants in terminally differentiated cells also appears to be a common feature in a number of organisms.

The requirement to package newly synthesized DNA into chromatin makes it logical that histone synthesis is regulated in concert with the cell cycle, and it has been observed that the abundance of histone mRNA increases 25- to 30-fold at the G1-S transition (Osley 1991Citation ; Ewen 2000Citation ). These replication-dependent histone mRNAs are intronless and are not polyadenylated but instead contain a conserved 16-nt stem-loop structure at the 3' end (Dominski and Marzluff 1999Citation ). The promoters of the replication-dependent histone genes usually contain core RNA polymerase II elements, a distal activation domain, and subtype-specific consensus elements that link promoter activity to the cell cycle. A second class of histone mRNAs is expressed at constitutive basal levels throughout the cell cycle, and these are both polyadenylated and can contain introns. These mRNAs encode the so called "replacement variant" histones implicated in chromatin repair and remodeling. The stoichiometric relationship of histone mass to DNA content might lead to expectation of some correlation between histone gene content and genome size; but thus far, no such relationship is obvious. High copy number tandem repeats of histone clusters do occur in some species with rapid cycles of DNA replication during early development, suggesting that histone gene organization may be related more to certain life history characteristics than to phylogenetic position, but the data remain limited. Overall, despite the large number of histone sequences available for a variety of organisms, information on histone gene complements and organization remains fragmentary in all but a few species, and evolutionary trends are unclear.

One significant phylogenetic gap in information on histone gene structure and organization is in the subphylum Urochordata. Of the five sister Urochordate classes, the Appendicularia (fig. 1 ) has interesting life history features with respect to histone gene organization, histone gene regulation, and spatial and temporal use of histone variants. The animal begins its short life cycle with very rapid cell cycles, but after metamorphosis, growth occurs almost entirely by increasing cell size through endoreduplication rather than mitotic cell division. Polyploidization, a process that is much more common in both plants and animals than is generally recognized, exceeds 1,000n in some tissues of the appendicularian genus Oikopleura (Fenaux 1971Citation ; Ganot and Thompson 2002Citation ) and raises questions concerning the decoupling of major histone synthesis from regular cell cycle progression. The accessible and transparent epithelium responsible for repeated synthesis of the elaborate houses in which the animal lives and filter feeds contains fields of cells with characteristic, distinctive variations in nuclear morphologies (Spada et al. 2001Citation ; Thompson, Kallesoe, and Spada 2001Citation ). This organ is ideally suited to exploring the spatial nuclear compartmentalization of histone variants, relative to defined patterns of gene expression.



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Fig. 1.—Schematic representation of the life cycle of O. dioica at 15°C. The transparent, 100-µm-diameter oocyte is fertilized and undergoes rapid mitotic cleavage cycles. At 5–6 h, the tadpole hatches from the chorion. During the next several hours, organogenesis occurs in the trunk and the length of the tail increases relative to the size of the trunk. At 12–14 h after fertilization, the animal undergoes metamorphosis that involves the tail shifting from a linear posteriorly directed orientation relative to the trunk to a more orthogonal position with the end of the tail lying in the same direction as the mouth. Immediately after tailshift, the animal inflates its first complex house structure (not shown in schema), in which it is entirely enclosed, and begins filter feeding. Over the subsequent 5–6 days, the animal renews the house structure every 3–4 h and grows principally by increasing the size of somatic cells as opposed to increasing cell number. From metamorphosis on cells in the majority of tissues of O. dioica exit mitotic cycles and enter endoreduplication cycles. Toward the end of the very short life cycle, the gonad increases dramatically in size, surpassing that of the trunk. Females produce 200–300 oocytes each, and upon spawning, both males and females die. Timing of events in the figure and the relative sizes of the embryos, tadpoles, and adults are not to scale.

 
In the present study, we show that O. dioica has a total histone gene complement that is very similar to that of humans, despite having a genome size that is approximately 50-fold smaller (Seo et al. 2001Citation ). Although O. dioica undergoes rapid early cleavage stages, there was no evidence of high copy number tandem repeats of histone clusters. The quintet histone cluster was the most compact of any described thus far, measuring only 3.5 kb in length. The promoters of the divergently transcribed H2A-H2B and H3-H4 histone pairs also were very compact, and some of these possessed only a single TATA box. Oikopleura dioica had an impressive array of diversity in H2A and H2B variants, and expression of the H2A variants was regulated developmentally. Many of the Oikopleura histone genes coded for both a stem-loop and polyadenylation signal in the 3'UTR, and transcripts containing both these elements as well as a polyA tail were produced.


    Materials and Methods
 TOP
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Acknowledgements
 References
 
Appendicularia Culture
Animals were collected from fjords around Bergen, Norway, and cultured in 6-liter beakers with constant stirring at 14–15°C. The duration of the life cycle was 5–6 days. Mature males and females were placed together in 4-liter volumes of seawater and allowed to spawn. Cultures were diluted 1:6 on day 1 and 1:1 on day 2; then animals were transferred to clean seawater by pipetting on each of the following days. The Appendicularia were maintained on cultured algal strains of Isochrysis galbana, Chaetoceros calcitrans, and Synecococcus sp., supplemented with Rhodomonas sp. from day 3 onward.

In vitro fertilizations were performed by collecting oocytes from mature females in watch glasses and adding a diluted suspension of sperm obtained from 1 to 2 mature male ejaculates in sterile filtered seawater. Fertilized oocytes were rinsed two to three times in sterile filtered seawater when 90% of them had emitted polar bodies. Embryos were then left to develop at room temperature.

Genomic Library Screening
A partial O. dioica shotgun genomic database was searched for core histone sequences, and one partial H2A sequence was retrieved. Primers OdH2Ap1, 5'-GATGTTGGGTAGAACACCACC-3', and OdH2Ap2, 5'-GGACTCCAGTTCCCCGTTGG-3', were designed from this sequence to amplify a 250-bp fragment spanning the histone fold. This fragment was random prime labeled with 32P-{alpha}-dCTP and used to screen a genomic library that had been prepared by partial digestion of sperm DNA with Sau 3A and cloned into the lambda EMBL3 vector (Stratagene). Among positive clones obtained from this screening, a clone containing a five-histone cluster was used for producing probes for H1 and the other core histones (H2B, H3, and H4). A 241-bp HindIII or HindII fragment spanning the histone H1 globular domain was purified and used for further screening of the genomic sperm DNA library.

Quantitative Southern Blotting
Haploid genomic DNA was isolated from sperm, and 4-µg aliquots were digested with EcoRV, SpeI-SphI, or NcoI-BamHI. A genomic clone containing a five-histone cluster was loaded on the same gel as a copy number standard. After electrophoresis and transfer, the DNA was UV–cross-linked to the wet membrane (Nylon Hybond N+, Amersham) with 150 mJ/cm2 at 254 nm, using a Hoefer UVC 500 Ultraviolet Crosslinker (APBiotech). Probes spanning the histone fold of OdH2A, OdH2B, OdH3, OdH4, and the globular domain of OdH1 were labeled as above. Hybridizations were carried out overnight at 55°C in 6x SSC, 1% nonfat dried milk, 1% SDS, and 2.5 mM EDTA. Final washing was in 0.5x SSC, 0.5% SDS at 65°C. Hybridized blots were imaged with a FLA2000 phosphoimager (Fuji), and signal quantification was done using Image Gauge v2.01 software (Fuji).

Bacterial Artificial Chromosome and cDNA Library Screening
The bacterial artificial chromosome (BAC) library was prepared by partial HindIII digestion of sperm DNA. The BAC clones were spotted in ordered grids onto nylon filters (Amplicon Express, Washington). The cDNA library was prepared from polyA+ RNA isolated from early tadpoles, as described previously (Spada et al. 2001Citation ). The libraries were screened using probes spanning the globular domain of the linker histone H1 and the histone fold domains of H2A and H4. Hybridizations and washing steps were carried out as described above. Signals from hybridized filters were recorded on autoradiographic films (Kodak Biomax MS). For reprobing, BAC library filters were stripped by submerging them in boiling water–0.5% SDS for 15 min. Single-colony excision was performed on isolated plaques from the cDNA library to obtain DNA fragments cloned in pBK-CMV.

Reverse Transcriptase–Polymerase Chain Reaction
Total RNA from indicated developmental stages was isolated by the guanidium thiocyanate–acid phenol method. First strand cDNA synthesis was performed by incubating 2 µg of DNaseI-treated (PCR grade, GIBCO-BRL, Life Technologies) total RNA with 100 pmol of random hexamers, 10 mM DTT, 1 U/µl RNasin (Promega), 0.5 mM dNTPs, in 50 mM Tris-HCl–75 mM KCl–3 mM MgCl2, pH 8.3, for 1 h at 37°C in the presence or absence of 400 U of MMLV reverse transcriptase (GIBCO-BRL, Life Technologies). Each PCR reaction contained cDNA synthesized from an equivalent of 50 ng of total RNA, 0.4 U Vent polymerase (New England Biolabs), 0.2 mM of each dNTP, 0.2 µM primers, in 25 µl of 10 mM KCl, 20 mM Tris-HCl pH 8.8, 10 mM (NH4)2SO4, 2 mM MgSO4, and 0.1% Triton X-100. After initial denaturation for 2 min at 95°C, 35 amplification cycles (95°C, 30 s; 50–63°C [primer dependent, see table 1 ], 30 s; 72°C, 30 s) were carried out with a final extension for 5 min at 72°C. All RT-negative controls were run to 40 cycles of amplification. Primers used for the specific RT-PCR reactions are described in table 1 .


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Table 1 Primers Specific for Oikopleura dioica H1, H2A, and H4 Variants

 
RNase Protection Assay
Total RNA from early tadpoles (6–7 h after fertilization) and adults (d4/5) was isolated as described above. RNAse A/T1 mappings were performed according to Goodall, Wiebauer, and Filipowicz (1990)Citation , using 2 µg of total RNA per protection. The RNA antisense probes were prepared from clones isolated from the cDNA library. Plasmids were linearized with XhoI and used as templates for in vitro transcription using T3 RNA polymerase (Promega). Protections were run on 8 M urea, 6% polyacrylamide (19:1) gels. Autoradiographic images were processed with a FLA2000 phosphoimager (Fuji).


    Results
 TOP
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Acknowledgements
 References
 
Oikopleura dioica Histone Gene Complement
Several Oikopleura histone genes were isolated from the genomic library and sequenced. After assessing restriction site frequencies, haploid genomic Southern blots were prepared from sperm DNA. Probes were made covering the conserved histone fold regions of the core histones and the globular region of the linker histone H1 and were hybridized to the Southern blots (fig. 2 ). Densitometric analyses of the blots yielded estimates of 9–11 H4 genes, 11–14 H3 genes, 15–19 H2A genes, 18–20 H2B genes, and 4–7 H1 genes. The very similar hybridization patterns on the H3 and H4 blots indicated significant coupling of these genes. This also was observed for H2A and H2B, though the patterns among these genes were more variable. Despite the fact that Oikopleura undergoes rapid cleavage cycles during early development, there was no evidence of any large tandem repeats of histone clusters as has been observed in a number of other organisms that also have rapid, early cleavage cycles.



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Fig. 2.—Histone gene complement of O. dioica. Haploid sperm DNA was digested with (1) BamH1-NcoI, (2) EcoRV, or (3) SpeI-SphI. The Southern blots were hybridized with probes corresponding to the globular domain of histone H1 and the conserved histone fold regions of histones H2A, H2B, H3, and H4, respectively. Copy number standards are shown at the bottom of each blot, and the positions of molecular mass standards of 1, 2, 3, and 10 kb are indicated by horizontal lines at the left of each blot

 
Further analysis of the histone complement was undertaken by screening an O. dioica BAC library using the same probes for H1, H2A, and H4 that were hybridized to the Southern blots. The BAC library was prepared using a partial HindIII digest of sperm DNA. The average insert size was 135 kb, and based on an estimated genome size of 72 Mb (Seo et al. 2001Citation ), the entire grided filter set of the library represents 15- to 17-fold coverage of the genome. This coverage has been confirmed by screening for a number of single-copy genes (data not shown) that consistently yielded between 13 and 20 positive signals. Results of the histone screening are summarized in table 2 . When the number of positive signals was divided by 15 to normalize to genome coverage, conclusions could be drawn that agreed well with the genomic Southern blots. At the resolution of 135 kb, it appeared that H1 occurred in two locations, was linked to other histone genes, and did not occur anywhere as an isolated locus. The results for H4 suggest that it occurred in equal proportions in quintet clusters, quartet clusters, and as isolated H4 loci or H3-H4 pairs. There were numerous clones positive for H2A but at least 30% were linked in quintet or quartet clusters. The ratio of total positive clones of 1:3:2 for H1-H2A-H4 agreed with the ratios of gene numbers obtained by the densitometric analysis of the Southern blots.


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Table 2 Representation of Histone Genes in the Oikopleura dioica BAC Library

 
The variants of each of the core histone subtypes are presented in figure 3 . The H4 and H3 histone sequences were highly conserved relative to H4 and H3 sequences from a variety of organisms. This was evident in both the histone fold domains and in the N-terminal tails implicated in regulating modifications of the chromatin structure. Within the H4 histone fold domain, the O. dioica and ascidian sequences share a change from methionine to leucine at position 85, but there are insufficient data from other Urochordate classes to determine whether this is characteristic of the subphylum. Oikopleura dioica showed extensive differences between both H2A and H2B variants. This was particularly noteworthy in the N-terminal tail regions. For example, the seven human H2A variants can be divided into two groups with respect to the N-terminal sequence, and both these groups have a common GKA(K/R)AKA core. In comparison, the N-terminal regions of the four O. dioica H2A variants identified thus far are all distinctly different, and only one contains a core KAKAKA sequence. The O. dioica H2A variants also show considerable variability in the last 10–15 residues of the C-terminal tail.



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Fig. 3.—Sequence comparisons of O. dioica core histones. Histone H4 and H3 sequences from O. dioica (Od) are compared with representative sequences from the related urochordate, ascidian Styela plicata (asc), the arthropod D. melanogaster (dme), the echinoderm sea urchin Paracentrotus lividus (pli), the nematode C. elegans (cel), the human (hum), and a consensus vertebrate H4 sequence (vert). For H2A and H2B, the O. dioica variants are compared amongst each other, and the level of variation in the N-termini of O. dioica H2A sequences is compared with that observed in human H2A sequences. We have not identified any O. dioica sequence related to the distinctly different macroH2A variants. Amino acid residues comprising the histone fold domains of the respective core histones are shaded in gray. Dots indicate amino acid residues identical to the upper consensus sequence, and dashes represent gaps

 
The linker histone H1 (fig. 4 ) was present in relatively few copies in the haploid genome of O. dioica. The most striking feature was the short length of the N- and C-terminal tails, a characteristic shared by copepod H1, and resembling terminal differentiation H1.0 variants in higher vertebrates. The two O. dioica H1's differed very little, the essential difference being an additional repeat unit (TKKAAS) in the C-terminal tail of H1.2.



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Fig. 4.—Oikopleura dioica linker histone H1. (A) The amino acid sequences of O. dioica histone H1 with the predicted globular domains shaded in gray. (B) Comparison of domain lengths in representative linker histone subtypes from O. dioica (Od), the echinoderm sea urchin Stongylocentrotus purpuratus (spu), the copepod Tigropius clifornicus (cop), the nematode C. elegans (cel), Drosophila (dme), the frog X. laevis (xla), and the human (hum). The N-terminal regions are represented by gray rectangles, the globular domains by striped rectangles, and the C-terminal tails by stippled rectangles. The length of each sequence in amino acids is given at the right

 
Organization of Histone Genes and Their Regulatory Elements
Sequencing of a number of genomic clones (fig. 5 ) revealed the presence of both full histone quintet clusters and isolated variants. The quintet cluster consisted of a central H1 gene flanked by divergently transcribed H3-H4 and H2A-H2B pairs, an organization found in other organisms such as humans. The quintet cluster is extremely compact, covering only 3.5 kb, and to our knowledge, is the most compact histone quintet to be described. H2A-H2B pairs were found with both divergent and same sense transcriptional directions. In view of the cell cycle regulation of transcription of many histone genes, there was an intriguing association of protein kinase A and pololike kinase genes with some of the histone gene variants. Protein kinase A is involved in regulating the G1-S transition of the cell cycle in response to intracellular levels of cAMP (Haddox, Magun, and Russell 1980Citation ) and is able to phosphorylate cyclin D1 (Sewing and Muller 1994Citation ). Pololike kinases are implicated in multiple processes during cell division, including activation of Cdk1-cyclinB, destruction of mitotic cyclins, and regulation of cytokinesis (Nigg 1998Citation ).



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Fig. 5.—Organization of O. dioica histone clusters. Four genomic clones (A–D) were sequenced. Differentially shaded arrows indicate the position and reading orientation of the histone genes. Open arrows indicate the position and reading orientation of genes with similarities to protein kinase A (PKA), pololike kinase (Plx-1), and chitinase. Parallel slashes indicate nonlinear breaks in the representations of the clones, and shaded vertical rectangles indicate junctions with the cloning vector

 
The promoter regions (fig. 6 ) of O. dioica histone genes were very small. The shared promoters for divergently transcribed histone pairs ranged from 220 to 240 bp in length. For human shared H2A-H2B promoters, two types have been described, those that contain the cell cycle–dependent elements CREB and E2F and those that do not. These two types of promoters also were observed in the O. dioica sequences. In addition to the CREB and E2F motifs, Trappe, Doenecke, and Albig (1999)Citation have described recently a functional RT-1 motif in type II human promoters. We now show that this RT-1 motif is present in O. dioica H2A-H2B promoters that also contain the CREB and E2F elements. The shared promoters of the appendicularians differ from those of the human in that some contain only one TATA box for the two divergently transcribed genes. In general, the frequency of occurrence of CAAT boxes was reduced greatly in O. dioica promoters when compared with their human counterparts, and in many cases they were absent completely.



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Fig. 6.—Elements in the promoter regions of O. dioica histone genes. (A) Monodirectional promoters. (B) Bidirectional promoters. The organization of the two types of bidirectional human H2A-H2B promoters (Albig et al. 1999Citation ) are shown for comparison. Elements that are generally considered to be cell cycle dependent are shaded in gray, core promoter elements are represented by open symbols, and the RT-1 box, an element that had thus far only been identified in the human H2A-H2B promoters, is represented by a striped vertical arrow. The Oct-1 element that has been implicated in the S-phase elevation of human H2B gene expression but also is found in the promoters of genes expressed independently of DNA replication is indicated by a stippled cylinder

 
All the O. dioica histones sequenced thus far code for a well-conserved stem-loop structure in the 3'UTR that is characteristic of replication-dependent histone mRNAs (table 3 ). Both visual inspection and PRATT analysis (Jonassen 1997Citation ) of the sequences 3' to the conserved stem-loop revealed no consensus 3' purine-rich histone downstream element (HDE) that interacts with U7 snRNA in the processing of histone mRNAs. In fact, no consensus sequence or pattern motif of any kind was discovered 3' of the stem-loop in O. dioica histone gene sequences. The interaction of U7 snRNA with the HDE is postulated to serve as a molecular guide defining the exact position of the cleavage site (Zhao, Hyman, and Moore 1999Citation ). However, despite the absence of an HDE in the O. dioica 3' sequences, cleavage at the normal site 3' of the stem-loop structure was observed by RNase protection analysis of histone H4 transcripts (fig. 7 ). Most of the O. dioica histone sequences also contained an AATAAA polyadenylation signal sequence downstream of the stem-loop structure (table 3 ). Both RT-PCR assays (see below) and direct sequencing (fig. 8 ) revealed that histone H1, H2A, and H4 transcripts can indeed be polyadenylated. The polyadenylated transcripts contained the conserved stem-loop structure in the 3'UTR.


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Table 3 Oikopleura dioica Histone 3'UTR Elements

 


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Fig. 7.—RNAse A/T1 protection using an antisense RNA probe complementary to a cDNA sequence encoding histone H4 mRNA. The RNA probe was hybridized against total RNA extracted from early tadpoles (ET, 6–7 h after fertilization) and a mixture of day 4 and 5 adults (d4/5). The portion of the probe sequence corresponding to the coding region of the histone H4 cDNA is in bold italics and of the stem-loop consensus sequence is in bold and underlined. The presence of a band at 188 bp (*) shows that although a consensus purine-rich HDE is absent, cleavage of the histone mRNA occurs at the usual site after the stem-loop consensus sequence. The other bands present on the gel result from polymorphisms in the 3'UTR sequences of the O. dioica histone H4 gene complement compared with the 3'UTR region of the specific histone H4 cDNA probe used. The intense double band at 129 and 133 nt corresponds to the coding portion of the probe where sequences among the H4 genes are much more strongly conserved

 


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Fig. 8.—Polyadenylation of O. dioica histone transcripts. The sequences of histone transcripts were obtained by screening a cDNA library made from polyA+ RNA at the early tadpole stage. Portions of sequences in italics highlighted in light gray are the ends of the coding regions, terminating with the TAA stop codon highlighted in black. The stem-loop structure is highlighted in gray with key conserved consensus nucleotides in bold and underlined. The polyadenylation signal AATAAA is in bold and double underlined, with the polyA tail indicated in bold. H2A.1-a and H2A.1-b are two different 3'UTR sequences associated with genes coding for the same variant H2A.1

 
Developmental Stage–Specific Expression of Histone Variants
Developmental expression profiles were obtained by RT-PCR using primer pairs for histones H1.1/2 and H4.1, and the specific variants H2A.1, H2A.2, H2A.3, and H2A.4 (fig. 9 ). Absence of cross-reactivity of primer pairs among the H2A variants was verified using phage or plasmid DNA containing each of the respective variants. The reverse transcription reaction was primed with either oligodT or random hexamers to distinguish polyadenylated histone transcripts from total histone transcripts. Primers specific to transcripts coding for the ribosomal protein L8 were included as internal controls.



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Fig. 9.—Developmental expression profiles of O. dioica histone H1, H2A, and H4 variants. Total RNA was prepared from oocytes (ooc), 2–4 cell embryos (4c), 2-h embryos (2hrs), early tadpoles (ET, 6–7 h after fertilization), and day 1 (d1), day 2 (d2), and maturing day 4–5 (d4/5) animals. First-strand synthesis cDNA was obtained by priming total RNA with either random hexamers (left panel) or oligo dT (centre panel). Controls for the variant specificity of the H2A primers were performed on phage or bacterial clones containing only the specific variant (right panel), with none of the primer pairs exhibiting cross-amplification of other variants. Primers for the ribosomal protein L8 (RbL8) were used as internal controls at each developmental stage

 
Stage-specific expression of the variants was observed. Histone H4.1 was expressed throughout development. Transcripts were present in mature oocytes, rose in abundance during gastrulation and organogenesis in tadpoles, and then declined in abundance after the shift from cell division and proliferation to subsequent growth and maturation principally due to increase in cell sizes through endoreduplicative cell cycles (days 1–5). Histone H1.1/2 transcripts did not appear to be stocked maternally because no amplification was detected in oocytes or four-cell embryos. H1.1/2 was first detected in gastrulating embryos 2 h after fertilization, a time when zygotic transcription is underway. Transcript levels appeared to attain a peak during organogenesis at the tadpole stage and were less abundant throughout the remainder of the life cycle. Histone H2A.1 showed an expression profile that was similar to that of H4.1, whereas variants H2A.2 and H2A.4 were expressed only in maturing animals at days 4 to 5. Using this assay, we were unable to detect the expression of H2A.3 and could not rule out the possibility that it may be a pseudogene. Polyadenylated transcripts were observed for H1.1/2, H2A.1, and H4.1 but were not found for H2A.2 or H2A.4, although both these genes did contain a downstream polyadenylation signal at a similar distance from the termination codon as H2A.1.

EMBL accession numbers for all sequences reported in this study are from AJ494848 to AJ494856.


    Discussion
 TOP
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Acknowledgements
 References
 
Sequencing of genes coding for core histones from a wide variety of species clearly shows that they are among the most phylogenetically conserved proteins. However, studies on the organization of the histone complement within organisms, the transcriptional regulation of histone expression, and the processing of histone mRNA have had a much narrower focus on relatively few model organisms or cell culture systems. Nonetheless, a number of generalizations have been made, and it is of interest to consider the data presented here on the urochordate appendicularians in the context of these previous trends and observations.

No clear correlation has been observed between the total histone gene complement of an organism and its genome size. The data from O. dioica agree that if any such correlative constraint does exist, it is very weak. Despite an estimated genome size of 72 Mb (Seo et al. 2001Citation ) compared with 3,200 Mb for humans (International Human Genome Sequencing Consortium 2001Citation ), O. dioica has a total histone gene complement that is very similar to that of humans. Thus, size of the histone gene complement does not appear to be a major factor that regulates the stoichiometry of histone to DNA content in the cell nucleus. One trend that has appeared previously is that organisms undergoing embryonic development with rapid, early cleavage cycles have extended tandem repeats of histone gene clusters, whereas those with longer cleavage stage cell cycles, such as mammals, do not. Tandem histone gene cluster repeats are found both in rapidly developing organisms with relatively small genome sizes such as Drosophila (Lifton et al. 1978Citation ; Matsuo and Yamazaki 1989Citation ) and in those with large genomes such as Xenopus (Perry, Thomsen, and Roeder 1985Citation ; Turner et al. 1988Citation ). Oikopleura dioica diverges from this trend in that it has very rapid cleavage cycles (5–10 min duration at 20°C) during early development but does not contain extended tandem histone gene repeats.

It is not surprising that with respect to a broad range of species, O. dioica histones H3 and H4 are highly conserved in both the histone fold and in the regulatory N-terminal tails. Thus far, most investigations of the histone code have concentrated on modifications of the N-terminal tails of H3 and H4. There is logic to this in that the high level of conservation implies that findings in one or a few species are likely to be generalized to many others. It also may imply, however, that these modifications will be primarily implicated in more basal processes in chromatin remodeling. The greater variability among diverse organisms in H2A and H2B sequences may reflect less functional constraint on these proteins or a greater variety of more specialized functions layered on to the regulatory roles of H3 and H4 (or both). Specific variants of H2A are known to be vital to the viability of some organisms, suggesting that at least some more specialized roles are critical for development and survival (Clarkson et al. 1999Citation ). There is a much greater degree of variability within the set of O. dioica H2A sequences, particularly in the N- and C-terminal tails, than among the H2A complements found in vertebrates or in invertebrates, such as Drosophila and Caenorhabditis elegans, where sufficient sequence information is available for comparison. Curiously, the extent of variation in O. dioica H2A sequences resembles that found in phylogenetically distant plants such as rice (Oryza sativa, TIGR rice genome database, http://www.tigr.org/tdb/e2k1/osa1) or Arabidopsis (TIGR A. thaliana genome database, http://www.tigr.org/tdb/e2k1/ath1). In plants, polyploidization is a frequent occurrence and is particularly prevalent in species with small genome sizes (De Rocher et al. 1990Citation ). Oikopleura dioica also makes extensive use of polyploidization in its development and rapid growth (Ganot and Thompson 2002Citation ), and a speculative correlation might link this aspect to the common greater diversity observed in H2A sequences in such distant organisms. The stage-specific expression of the H2A variants in O. dioica warrants further investigation in this regard.

The organization of histone genes in O. dioica shares features found in other organisms, including quintets with a central H1 gene flanked by pairs of divergently transcribed H3-H4 and H2A-H2B pairs as well as more isolated variants. At a total length of 3.5 kb, the quintet cluster is the most compact reported thus far, and high gene density appears to be a general feature of the O. dioica genome (Seo et al. 2001Citation ). The shared promoter regions of divergently transcribed histone genes were very small, ranging from 220 to 240 bp in length. In mitotically dividing cells, where most studies of histone synthesis have been carried out, primary regulation of transcription and translation occurs near the G1-S cell cycle transition. In O. dioica, after metamorphosis at 14 h of development, growth through the remainder of the life cycle occurs principally through endoreduplication of DNA and increased cell size rather than through mitotic cell division. Thus, this organism provides an appealing system for examining regulation of histone synthesis in an endoreduplicating as opposed to a mitotic environment. One notable feature in the regulatory regions of several O. dioica histone genes is the absence of an appropriately positioned TATA box upstream of the transcription start site. This is the case for genes that are indeed transcriptionally active (fig. 9 ). Elements found in the promoters of histone genes described in other species also are present in the O. dioica promoters. The shared promoter regions of divergently transcribed O. dioica H2A-H2B gene pairs can be divided into two classes as has been done for the shared H2A-H2B promoters in human and mouse (Albig et al. 1999Citation ). The human and murine type I promoters contain TATA boxes, Oct-1 elements, and CAAT boxes, whereas the type II promoters contain in addition E2F and CREB elements. The E2F element is characteristic of S-phase–regulated genes (La Thangue 1994Citation ), and the CREB element may adjust H2A and H2B expression to fluctuating growth conditions (Trappe, Doenecke, and Albig 1999Citation ). In human type II promoters, there is an additional octanucleotide RT-1 element adjacent to the E2F element (Albig et al. 1999Citation ). Database searches by these authors revealed this sequence to be unique to human H2A-H2B promoters. The O. dioica type I H2A-H2B promoters lack CAAT boxes when compared with the mammalian counterparts. The type II promoters contain the E2F and CREB elements as well as the RT-1 box described thus far only in human promoters. By running a PRATT analysis on promoter sequences in databases, we confirmed the absence of this element in promoters other than those of human and O. dioica. In O. dioica, the RT-1 box is not immediately adjacent to the E2F element, but interestingly, it is present in promoters that lack a TATA box 5' of the H2A gene and absent in promoters that contain the TATA box. Among genes coding for the O. dioica H2A.1 variant, examples of both type I and type II promoters were found. This variant was expressed throughout development. On the other hand, promoters of the H2A.2 and H2A.4 variants were of type I, lacking cell cycle–dependent elements, and were expressed exclusively in day 4 and 5 animals, a time when most growth of the animal occurs through endoreduplication rather than through mitotic division.

Processing of histone mRNA has been most extensively studied using echinoderm sea urchin and mammalian sequences. The replication-dependent histones are up-regulated at G1-S, and this involves a conserved stem-loop structure in the 3'UTR. Replacement variants lack the stem-loop structure, are polyadenylated, and are expressed at more basal levels throughout the cell cycle. Important factors in processing replication-dependent histone RNAs are the stem-loop–binding protein and the U7 snRNP. Positioning of U7 is determined by RNA-RNA base pairing between U7 and a purine-rich element (HDE) that serves as a molecular guide in defining the point of cleavage (Scharl and Steitz 1994Citation ). The HDE is located 13–17 nucleotides downstream of the cleavage site that itself is immediately 3' of the stem-loop sequence in the histone 3'UTR. The HDE sequence is strictly conserved (CAAGAAAGA) in the sea urchin where it was first described (Georgiev and Birnstiel 1985Citation ) but is more variable around a consensus core AAAGAG sequence in mammals (Dominski and Marzluff 1999Citation ). Inspection of the flybase reveals that in Drosophila there is not a strictly conserved HDE consensus sequence but there are clearly purine-rich stretches in the 3'UTRs of the histone genes in the appropriate location downstream of the stem-loop structure. Analysis of C. elegans histone gene sequences in the wormbase reveals no consensus HDE sequence or purine-rich stretch downstream of the stem-loop. In O. dioica, there also was no evidence of an HDE consensus sequence or purine-rich element 3' of the stem-loop. In contrast, inspection of the sequence database for the related urochordate ascidian Ciona intestinalis (http://bahama.jgi-psf.org/prod/bin/blast.ciona.cgi) revealed a clear putative consensus core HDE element (AAAGAGA for H2A, H3, and H4; AAA[G/C]GAG for H1 and H2B). The U7 snRNA of echinoderms and vertebrates is small (55–70 nt), but thus far, no U7 candidates or obvious orthologues of the U7-specific Sm protein Lsm10 have been identified in the genomic sequences of Drosophila and C. elegans (Lanzotti et al. 2002Citation ). Despite lacking an HDE element, we did verify that cleavage at the normal site 3' of the stem-loop does occur in O. dioica histone H4 messages. It remains to be determined whether U7-like snRNA sequences are involved in the processing of Drosophila, C. elegans, and O. dioica histone mRNAs. Phylogenetically, it is intriguing that histone genes in the echinoderm sea urchin, the urochordate ascidian, and higher vertebrates all have HDE elements, whereas the appendicularian O. dioica does not.

In the O. dioica histone gene complement, there were frequently AATAAA polyadenylation signals downstream of the stem-loop sequence. This also is characteristic of the Drosophila and C. elegans histone genes, and it has been proposed that this might serve as a safeguard to prevent read-through into neighboring histone genes with the generation of antisense RNA (Lanzotti et al. 2002Citation ). We have shown in this study that O. dioica transcripts can contain both the stem-loop sequence and a polyA tail. This has been observed in other species, for example, Drosophila (Lanzotti et al. 2002Citation ) and bovine (Gendron et al. 1998Citation ). In Drosophila embryos, mutants for the stem-loop–binding protein accumulate polyadenylated histone mRNA. The turnover of polyadenylated RNA in this case appeared to be dependent on cell type, and the stability of the polyA transcripts was more pronounced in proliferating or endoreduplicating cells than in quiescent G0 cells. There was a clear decrease in O. dioica H1.1/2 gene expression that paralleled the transition from growth by cell proliferation to growth through endoreduplication, but there was no evident qualitative shift in the use of polyadenylation for any of the histone variants examined (fig. 9 ). The functional significance, if any, of the O. dioica histone transcripts containing both the stem-loop and a polyA tail is at present unclear.

The packaging of DNA into nucleosomes is a fundamentally conserved property of the eukaryotic nucleus, and this is evident in the conservation of histone sequences, particularly those of the central H3-H4 tetramer. In contrast, there appears to be much more plasticity in the organization of histone gene complements among organisms, the degree of variation within histone subclasses, the regulation of histone gene expression, and perhaps some of the fundamentals of histone mRNA processing. The results here show that differences can be quite marked even over relatively short phylogenetic distances (e.g., echinoderm-appendicularian-ascidian) and that similarities may be linked more to certain life history or cell biological characteristics of an organism. With the increasing number of eukaryotic genome sequences coming available, studies of these aspects would appear to be an important complement to the current focus on deciphering the histone code in a few selected organisms.


    Acknowledgements
 TOP
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Acknowledgements
 References
 
We thank Albert Poutska, Max-Planck Institute for Molecular Genetics, Berlin, for a partial O. dioica H2A sequence that was used to initiate screening and cloning of histone clusters and genes, Rita Tewari, Sars Centre, for providing the O. dioica genomic library, Hee-Chan Seo, Sars Centre, for providing the ligation mix from which plasmid clones C and D (fig. 5 ) were isolated, and Philippe Ganot, Sars Centre, for advice on the RNase protection assay. This work was supported by grants from the Norwegian Research Council and Ministry of Education.


    Footnotes
 
William Jeffery, Reviewing Editor

Keywords: U7 snRNP RT-1 box histone stem-loop Appendicularia histone polyadenylation histone promoter Back

Address for correspondence and reprints: Eric M. Thompson, Sars International Centre for Marine Molecular Biology, Bergen High Technology Centre, Thormøhlensgt. 55, N-5008 Bergen, Norway. E-mail: eric.thompson{at}sars.uib.no Back


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Accepted for publication August 22, 2002.