An cornucopia of histone variant genes and RNA metabolism genes dominate the transcriptomic orchestra during sexual reproduction
A unifying mechanistic model for developmental sequence protection through sncRNAs integrating the many known factors and mechanisms, which contribute to programmed chromatin reorganization is lacking to date for spirotrichs like Stylonychia and Oxytricha. For the purpose of identifying important differentially expressed genes and to connect important gene networks, we performed deep sequencing of mRNA sampled at 8 stages in the sexual cycle of Stylonychia (i.e. vegetative cells, cells at the onset of conjugation (0 h post-conjugation [PC]) and after consecutive intervals 10–60 h PC). With emphasis on the probable important roles of histone variants in the course of macronuclear development, Fig. 1a summarizes the current knowledge on the spatiotemporal localization of several histone H3 variants. Our mRNA-seq results now underlined the superior roles of both histone variants and sncRNA metabolism factors in macronuclear differentiation, since many genes involved were ranked under the top differentially expressed candidates (Fig. 1b). Remarkably, during the same corresponding stages of macronuclear differentiation in Oxytricha trifallax, disproportionate numbers of mRNAs putatively encoding proteins involved in RNA (and DNA) functions were described recently [31]. Somewhat unexpectedly, we noted a paucity of chromodomain proteins and putative histone KMTs with only few candidates being developmentally expressed (Fig. 1c). This seemed to be a major difference to Tetrahymena, where such proteins are centrally for developmental DNA elimination [32, 33]. Notably, in contrast to the many Stylonychia histone variants, we could identify a uniform set of histone chaperones only, i.e. at a time one homolog of mammalian ASF1, CAF-1 and HIRA, respectively.
27nt-RNAs occur during development, whereas 21–22nt-RNAs become constitutively synthesized
Deep sequencing of small RNAs purified from the same samples as described above revealed two different size fractions becoming developmentally enriched, which were 21–22nt and 27nt in size, respectively, reminiscent of Oxytricha [3, 4]. We identified 21–22nt-RNAs at very low levels over threshold in isolates from vegetative cells and in the course of development (Fig. 2a). Although these small RNAs were almost undistinguishable from similarly sized fragments, their 5′- and 3′-bias as well as the internal base composition was reminiscent of 27nt-RNAs (Additional file 1: Figure S1A). This conservation appeared to rule out that the 21–22nt-RNAs were simple decay products.
In contrast, while an occurrence of 27nt-RNAs over baseline was not detected during vegetative growth, they emerged at the onset of conjugation and were clearly visible above threshold 10 h PC—at a time point, when the chromatin topology of prospective macronuclei (anlage[n]) exhibited a less condensed state when compared with micronuclei. Thereafter, we observed a massive 27nt-RNA gain (approx. 20 h PC, at the onset of polytene chromosome formation). Subsequently, 27nt-RNAs were still enriched until 60 h PC. At this time point, macronuclear anlagen entered a DNA-poor stage, which followed bulk DNA elimination. After rapidly reaching their maximum at ~ 20 h PC, the concentration of the 27nt-RNAs decreased continuously. Notably, the occurrence of other small-sized RNA (< 21nt) fragments overlapped with the synthesis of 27nt-RNAs, possibly representing debris of the sncRNA biogenesis.
Both 21–22nt-RNAs and 27nt-RNAs map exclusively to transcribed macronuclear sequences
We mapped both sncRNA fractions to the macronuclear genome and to all known micronuclear sequences (complete model genes and thousands of preliminary micronuclear contigs). Clearly and reminiscent of Oxytricha [3, 4], the vast majority of 21–22nt-RNAs (86.46%) and almost all 27nt-RNAs (98.04%) covered the transcribed regions of nanochromosomes at 20 h PC. Due to their greater abundance 27nt-RNAs had a much higher coverage than 21–22nt RNAs. Figure 2b shows exemplarily how both sncRNA fractions and mRNA reads mapped to four nanochromosomes—two containing single genes (TUBB and ASF1) and two dual gene nanochromosomes (Orf4995|PDI and GLRX|MSMO). Notably, the non-transcribed subtelomeric regions of the nanochromosomes were omitted from sncRNA read coverage, indicating that not the entire body of MDSs in the developing macronucleus became targeted by 27nt-RNAs—an interesting point we will take up below in the discussion.
We did not observe mapping of 21–22nt/27nt-RNA to micronucleus-specific genomic sequences. Almost all reads not covering MDSs could be assigned to other sources, e.g. sequences of ribosomal or mitochondrial origin, mRNA or repetitive sequences from ingested Chlorogonium algae (food source for Stylonychia), plastids or bacteria (Additional file 1: Figure S1B). Notably, these results challenge earlier observations in Stylonychia, where a small RNA probe preferentially appeared to hybridize to micronucleus DNA instead of macronucleus DNA in Southern analyses [13]. We assume that these earlier results were false positive. This could possibly be caused by probe purity and/or specificity issues, e.g. very short probe molecules (therefore less specific) were hybridized to micronuclear DNA, which is rich in repetitive sequences, at intermediate hybridization temperatures of 40 °C. Moreover, a relatively early sampling time point was selected in that previous study (10 h PC). Thus, enrichment of 27nt-RNAs was presumably not at its peak and the probe presumably contained other small nucleic acid contamination not originating from MDSs (compare Fig. 2a, c). We thus expect that the ratio between MDS-specific 27nt-RNAs and unspecified RNA fragments, which accumulated 10 h PC (Fig. 2a), led to an adverse probe composition.
Interestingly, the 27nt-RNAs coverage seemed to correlate directly with the mRNA copy number (R2 = 0.22586; p = 1.22 × 10−15) (Fig. 2b, d). This direct correlation left open the possibility that the substrate for the biogenesis of 27nt-RNAs could be pre-mRNA or mRNA. Supportive to this speculation was also that the 27nt-RNA reads covered introns to some extent when mapped to nanochromosomes, but not the non-transcribed subtelomeric regions. Thus, the 27nt-RNA read coverage overlapped well with the transcribed regions of the nanochromosomes.
Further, we tested whether the relative nanochromosomes copy numbers correlated with the 27nt-RNA coverage, since it is known that the mRNA copy numbers are influenced by the individual nanochromosome copy numbers [2, 17], whereby the different expression levels of genes residing on dual gene nanochromosomes show that the nanochromosome copy number is not the only determinant for mRNA quantity (Fig. 2b; Orf4995|PDI, GLRX|MSMO). However, we found a high positive correlation of both mRNA and nanochromosome copy numbers (R2 = 0.70728; p < 1.00 × 10−37) (Fig. 2d).
In addition, we observed that the mapped read count of 27nt-RNAs increased massively, transiently and selectively about 20 h PC (Fig. 2c), while the sequence composition of the 27nt-RNA pool remained stable from the onset of conjugation and in the course of successive development (Additional file 1: Figure S1C, D). This was reminiscent of a selective 27nt-RNA amplification by RNA-dependent RNA polymerase activity, possibly by one of the identified RDRP candidates (Fig. 1c).
In sum, 27nt-RNAs obviously occupied an elevated functional position also for Stylonychia. Sequence homology suggested that 27nt-RNAs target MDSs in developing macronuclei, whereby the omission of non-transcribed nanochromosomal sequences that are contained in MDSs suggested that this targeting might be incomplete.
In the subsequent part we focussed on 27nt-RNA and decided to neglect 21–22nt-RNA for further analyses.
27nt-RNAs bound by PIWI1 hybridize to complementary MDSs in developing macronuclei
To gain deeper insight into the putative 27nt-RNA-mediated protection mechanism for MDS retention, we studied the interactions of 27nt-RNAs with PIWI1 and with their target sequences, respectively. For this purpose, we purified macronuclear anlagen from Stylonychia, most of them being between 20 h and 40 h PC (beginning polytenization to beginning DNA elimination). These samples were allocated to different experiments.
To confirm that PIWI1 binds to sncRNAs, we performed PAR-CLIP. Therefore, we pulled down PIWI1 after 4TU-labelling of nascent RNA in exconjugants and cross-linking with UV light (365 nm). Enriched PIWI1-RNPs were end-labelled using polynucleotide kinase and 32P followed by denaturing SDS-PAGE. X-ray film detection of the radioactive tracer revealed a prominent signal being associated with the PIWI1 protein band (sncRNA1 in Fig. 2e) and another very faint smaller sized signal (sncRNA2 in Fig. 2e). Both signals were well in agreement with the expected molecular weight of PIWI1 loaded with 27nt-RNA (~ 98 kDa) or 21–22nt-RNA (~ 96 kDa), respectively, whereas higher molecular weight RNA was not present. The signal strength difference and the relative overrepresentation of the 27nt-RNAs over 21–22nt-RNAs suggested that PIWI1 binds predominantly 27nt-RNAs.
For further characterization RNA-seq and coverage studies on co-purified sncRNAs from pulled-down PIWI1 immunocomplexes were performed. For threshold subtraction and normalization, we used isotype control antibodies not specific for PIWI1. Taken together, we yielded substantial enrichment of 27nt-RNAs associated with pulled-down PIWI1. Deep sequencing revealed a coverage pattern on transcribed sequences of nanochromosomes, which was well in accordance with the data obtained for RNA-seq on purified small RNAs from serial stages of macronuclear development (Fig. 2f). Next, we intended to study the nature of MDS targeting by 27nt-RNAs, i.e. in particular, whether 27nt-RNAs bind directly to their target sequences via base-pairing: We purified nucleic acids from the same samples as described above and used the mouse monoclonal S9.6 antibody for immunoprecipitation of RNA/DNA hybrids [26].Again, negative isotype controls were performed. Subsequently, we made use of deep sequencing to analyse the coverage of 27nt-RNAs contained in these immunocomplexes. Strikingly, we observed a very high level of congruency between the coverages of 27nt-RNAs purified from RNA/DNA hybrids and previously purified sncRNAs at 20 h PC (Fig. 2b) as well as the coverage of 27nt-RNAs purified from pulled-down PIWI1 complexes (Fig. 2f). Here, R2 was as high as 0.99634 (Fig. 2f). In summary, these observations suggested that 27nt-RNAs are bound by PIWI1. The resulting PIWI1/27nt-RNA complexes could function as guides to target complementary sequences in MDSs by RNA/DNA base-pairing.
Intergenic bulk DNA elimination, but not IES removal depends on PIWI1
Although the above results revealed the deepest characterization of 27nt-RNAs bound to PIWI1 in Stylonychia so far, earlier studies already suggested that the sncRNA turnover is associated with PIWI1 [12, 14, 15]. By RNA-seq we now confirmed and complemented older results from microarrays [34], that PIWI1 was by far the most abundantly expressed developmental variant out of 13 PIWI-like proteins encoded in Stylonychia (Fig. 1). However, although we recently obtained evidence that PIWI1 influences the expression of some histone variants during macronuclear development [12], its mechanistic involvement is not yet understood. The quasi inverse sncRNA coverage pattern observed in Oxytricha as well as in Stylonychia—when compared with the ciliate Tetrahymena—could give support to the idea that alternative mechanisms could had evolved in the spirotrichous ciliate taxon, which fundamentally differ from Tetrahymena, where small non-coding ‘scan’ RNAs and the PIWI-homolog Twi1p specify micronuclear-limited sequences for heterochromatin formation [35]. Here, prior to chromatin elimination, heterochromatin formation involves H3K27me3 and H3K9me3 introduced by the histone lysine methyltransferase (KMT) EZL1 and binding of the chromodomain protein Pdd1p [32, 33]. This concept historically fed earlier speculations that the PIWI-domain proteins could directly interact with chromatin modifying factors also in spirotrichs. We now note that this view is challenged by the observed differences in sncRNA biology between these protozoa, and this impression is confirmed by the observed paucity of promising candidates for chromatin modifiers encoded in the Stylonychia genome, which would theoretically be required.
As a progressive step towards elucidating the mechanistic involvement of PIWI-domain proteins in spirotrichs, we continued to dissect the function of PIWI1. We used RNAi to study the effects of PIWI1 knock-down on the elimination of two different micronucleus-specific sequence classes—IESs and bulk DNA, since it was unknown to date whether the elimination of both depends on PIWI1 or not. Using PCR primer pairs amplifying only IES-containing micronucleus-specific sequences of selected genes, we observed that IES elimination from the macronucleus development protein 2 (Mdp2)-gene in PIWI1-minus samples was almost indistinguishable from controls—most IESs were removed between 24 and 30 h PC (Fig. 3a, Additional file 1: Figure S2). Contrary, a reporter sequence from the repetitive bulk DNA element MaA81 that is present in ~ 5000–7000 copies per haploid micronucleus genome [36] was almost completely eliminated 48 h PC in controls, whereas no DNA diminution was seen during prolonged observation periods up to 72 h PC in PIWI1-minus samples (Fig. 3b, Additional file 1: Figure S2). These results indicate that bulk DNA elimination, but not IES removal depended on PIWI1.
PIWI1 and H3K27me3 exhibit mutually exclusive localization patterns in polytene chromosomes
To complement the hypothesis that 27nt-RNAs in association with PIWI1 supposedly protect sequences that must be retained during macronuclear development [3], we reinvestigated the nuclear localization of PIWI1 and H3K27me3 with improved microscopic resolution (see visualization of an intact macronuclear anlagen nucleus with polytene chromosomes in Additional file 1: Figure S3A and Fig. 3c1-8, d, e). A first time observed feature was that PIWI1 appeared to travel physically within extranuclear spheres from parental macronuclei to the young anlage, which at this time point morphologically could not yet be distinguished from micronuclei (m) (Fig. 3c2). In particular, extranuclear PIWI1 overlapped with faint To-Pro-3-positive spheres, indicating that they might have contained significant RNA concentrations. Improved resolution elucidated that PIWI1 subsequently swamped the young anlage, and only in later anlagen stages subtle PIWI1 foci shaped out (Fig. 3c3-7). Subsequently, these foci became gradually fewer in number. These patterns suggested that PIWI1/27nt-RNA complexes were specifically targeted to discrete sites. Strikingly, this suggestion became obvious when higher magnifications of polytene chromosomes were examined (Fig. 3d). Here, condensed chromatin bands were devoid of PIWI1, whereas PIWI1 signals traced the longitudinal shape of the polytene chromosomes in areas not associated with dense chromatin. This suggested that PIWI1 colocalized with uncondensed chromatin.
Complementary, H3K27me3 was selected as a marker for condensed chromatin (Fig. 3c–e). Inversely, when compared with PIWI1 patterns, solely the condensed chromatin bands were strongly stained when antibodies against H3K27me3 were used for immunofluorescence microscopy on polytene chromosomes (Fig. 3e).
Notably, with respect to a potential H3K27me3/K9me3 ambiguity (compare epitopes in Additional file 1: Figure S3B), we observed non-identical H3K27me3/H3K9me3 staining patterns demonstrating the discriminability of both PTMs by the antibodies in use (Additional file 1: Figure S3C): In contrast to H3K27me3, H3K9me3 did not occur in condensed chromatin bands of polytene chromosomes. During DNA elimination H3K9me3 occurred only within chromatin bodies that were pinched off the polytene chromosomes. Within those amorphous bodies also H3K27me3 and H3K9ac occurred. These bodies were reminiscent of ‘swollen compartments’ observed earlier by transmission electron microscopy in anlagen after polytene chromosome breakdown, which were interpreted to contain material from former individual bands of a polytene chromosome [37]. All these observations supported the idea that PIWI1/27ntRNA complexes covered MDSs, possibly to protect them against the formation of condensed chromatin by means of H3K27me3.
In addition to the key observation of alternating PIWI1/H3K27me3 patterns in polytene chromosomes, we noticed that prior to its enrichment in macronuclear anlagen cytoplasmic H3K27me3 accumulations occurred, which did not overlap with any type of nuclei (Additional file 1: Figure S3D-L). This discovery suggested that trimethylation of H3K27 took place before it was assembled into polytene chromosomes. Consequently, since a pool of cytoplasmic H3K27me3 appeared to exist prior to its enrichment in anlagen chromatin, a putative nuclear H3K27-specific histone KMT activity that selectively targets micronucleus-specific sequence-associated nucleosomes would not be required.
Considering all these data we developed the model that sequence protection could mechanistically be achieved, if 27nt-RNAs together with PIWI1 could cause an ‘RNA-induced DNA replication interference’—i.e. if 27nt-RNAs would impair DNA replication at covered MDS loci during polytenization, thus locally and temporarily reducing the de novo deposition of nucleosomes. If in the meantime one or more specific histone variants and/or specific H3K27me3 activity would be transiently enriched, heterochromatin formation should be limited to those regions not protected by 27nt-RNAs. Our earlier work suggested that the late introduction of H3K27me3 during polytenization might correlate with the best time window for nucleosome deposition into chromatin, which opens during DNA replication—here during polytenization [14]. We hypothesize in particular that meanwhile MDSs are protected through stalled replication, the transient availability of H3K27me3 would lead to its preferential enrichment in bulk DNA regions.
DNA replication does not take place simultaneously at all sites during polytene chromosome formation in vivo
Accordingly, during developmental chromosome polytenization DNA replication should not occur simultaneously at all sites. In fact, we recognized spatiotemporally regulated replication patterns in the micro- and macronuclei of vegetative Stylonychia previously [20]. To test whether differentially replicated regions exist also in polytene chromosomes, we performed pulse (5 h)–chase (1 h)–pulse (5 h) labelling of newly replicated DNA using the nucleotide analogues iododeoxyuridine (IdU; early pulse) and chlorodeoxyuridine (CldU; late pulse). Subsequently, labelled nascent DNA in anlagen or spread polytene chromosomes was analysed by immunofluorescence microscopy. We observed that IdU (green, 1st pulse) and CldU (red, 2nd pulse) occurred mostly separated from each other, with IdU being tentatively associated with condensed bands and CldU with less condensed chromatin areas (Fig. 4a). These observations suggested that non-synchronous replication occurred during polytene chromosome formation. To confirm, we used qPCR to examine a time course during polytenization using reporter amplicons from bulk DNA (MaA81 and Stad5, respectively) and two micronuclear model genes, whose MDS/IES architecture and flanking sequences are well characterized (telomere binding protein alpha [TEBPalpha] and actin I [ACT1]). We found that MaA81 and Stad5 started replication earlier than MDSs (Fig. 4b). Sequences, which directly flanked micronuclear genes (and eventually became eliminated) became replicated later than bulk DNA, but earlier than MDSs.
The transient expression of histone variant H3.4 and H3K27me3 overlaps with the occurrence of polytene chromosomes in macronuclear anlagen
Interestingly, three histone variants of interest—H3.7, H3.5 and H3.4—exhibited differential developmental expression patterns (Fig. 4c), which at least for H3.4 fitted well with the hypothesized differential availability during heterochromatin formation. In contrast, H3.7 is an hyperacetylated replacement variant being only transiently incorporated very early in developing macronuclei [12]. Its function might be related to the replacement of micronucleus-specific H3.8 and thus to the loosening of the highly compacted zygote nucleus chromatin formed after haploid micronuclei fusion. Variant H3.5, though becoming gradually enriched in developing macronuclei, occurs in macronuclei throughout the life cycle [12]. Importantly, although both variants, H3.4 and H3.5, possess a conserved lysine 27 and were expressed in almost equal amounts 30 h PC and thereafter during further development, our data suggested that H3.4 is the preferential substrate for histone KTM activity directed to lysine 27. In particular, two H3 variants in Stylonychia—H3.4 and H3.8—harboured an identical motif (H3.4: K27TAPA; H3.8: K32TAPA) reminiscent of canonical H3 that was recognized by anti-H3K27me3 antibodies, if lysine 27 in H3.4 or lysine 32 in H3.8 were trimethylated (compare Additional file 1: Figure S3B and [12, 14]).
H3.4 is differentially expressed during chromosome polytenization. Although the K27me3 epitope is also present in H3.8, this 20 kDa H3 variant is restricted to micronuclei [12]. Other H3 variants, which were investigated in macronuclear anlagen, do not exhibit H3K27me3 (1. H3.7, a development-specific 20 kDa variant has no conserved motif homologous to H3K27; 2. H3.5 and H3.5 are abundant in developing and mature macronuclei at stages when H3K27me3 is not present.). Both variants H3.5 and H3.3 have deviant epitopes adjacent to K27 with additional amino acid substitutions/insertions (H3.5: K27TAQVAQ/H3.3: H3.5: K27STNVN), which most probably influence its availability as a KTM substrate or readout by putative effector proteins. The expression of other H3 variants seemed to be insignificant with respect to macronuclear development.
We observed that the expression of H3.4, which supposedly was target for K27 trimethylation, was marginal in vegetative Stylonychia and until 20 h PC. Then its expression increased massively during early polytenization from 20 to 30 h PC, before dropping again after 50 h PC (Fig. 4c). Thereby, the availability of both H3.4 seemed to outlast the process of heterochromatin formation, whereas other genes putatively relevant for condensed chromatin structure formation became only transiently expressed within a restricted critical time frame (~ 20 to 40 h PC), i.e. the putative histone KMT SET4 and the histone chaperones ASF1, CAF1 and HIR1, furthermore two histone H1 variants (Additional file 1: Figure S4). They all decreased from 40 h PC. We assume that a temporally restricted activity of either the H3K27me3 activity or the chromatin assembly machinery in form of histone chaperones could lead to the preferential enrichment of H3K27me3 nucleosomes at bulk DNA sequences, while MDS replication was stalled through PIWII1/27nt-RNA. Later, after replication at MDSs would have become completed and H3.4 or H3.5 became assembled into these sequences, a lack of H3.4K27me3 activity could have counteracted the formation of MDS-associated heterochromatin. Alternatively, but not mutually exclusive, a lack of histone chaperone activity could suppress the assembly of new nucleosomes into MDS chromatin. While this remains unknown, these proposed mechanisms would depend largely on replication timing and the coordinated availability of involved factors. We thus interrogated whether PIWI1/27nt-RNA complexes could have indeed the mechanistic potency to interfere with DNA replication and thus to be involved in the control of replication timing.
Small RNAs and Argonaute/oligonucleotide complexes impair DNA replication in vitro
To elucidate the influence of 27nt-RNAs and PIWI1 on the activity of several polymerases, we tested in vitro:
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1.
Whether 27nt-RNAs can modify DNA replication using Pfu and Taq polymerases for PCR (Fig. 5a);
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2.
Whether oligonucleotides (5′-phosphorylated 21nt-DNA or analogous 21nt-ssRNA) and a heat stable Argonaute protein—Thermus thermophilus Argonaute (TtAgo)—can modify DNA replication in a qPCR (Additional file 1: Figure S5);
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3.
Whether 27nt-RNAs in combination with Stylonychia PIWI1 can modify linear DNA amplification in a Klenow reaction (Fig. 5b).
As template for all assays we used a cloned 856 bp PCR fragment from the PCNA1 gene on the nanochromosome Contig13765—a differentially expressed PCNA-form during macronuclear development. To minimize the possibility that PCR primers were bound by Argonautes, we used excess length oligos with 5′-non-matching nucleotides forming hairpin loops (Additional file 1: Figure S6A). We used Pfu or Taq polymeases in different reactions to consider the effects of their specific exonuclease activities (Pfu: 3′ → 5′; Taq: 5′ → 3′). When mixes of serially diluted 27nt-RNAs (25–12.8 pM) matching PCNA1 were added to the PCR reactions (Additional file 1: Figure S6A), we observed impaired amplification at higher 27nt-RNA concentrations using Pfu polymerase and end-point PCR, whereas a specific band appeared at 1 µM or below (Fig. 5a; top). Nonsense 27nt-RNAs had no effect on the PCR reaction (Additional file 1: Figure S6B). Using Taq polymerase under identical conditions, a band became visible in all reactions showing that 5′ → 3′ exonuclease activity counteracted replication impairment through 27nt-RNAs. Nonetheless, qPCR analyses under similar conditions demonstrated that 27nt-RNAs from 25 µM to 1 µM in fact impaired DNA amplification, even if Taq polymerase was used (Fig. 5a; bottom). Comparatively, the efficiency of PCR impairment measured by qPCR was far stronger, when Pfu polymerase was used at 25–40 nM. These experiments showed clearly that 27nt-RNAs in principle could impair DNA amplification using different polymerases in vitro. Remarkably, at least Taq polymerase can exhibit strong strand displacement activity, i.e. it displaces downstream encountered DNA. Although we observed a clear dependency of DNA replication interference on the 27nt-RNA concentration used, it does not seem that displacement activity can also lead to the degradation of RNA in RNA/DNA hybrids.
To discover whether PIWI1 and 27nt-RNAs can block DNA replication, we synthesized PIWI1 with ‘universalized’ genetic code (since ciliates use an alternative genetic code) and N-terminal His-tag. Subsequently, we cloned His-PIWI1 into pCMV-TNT (Promega) for in vitro expression. Prior to assaying the inhibitory potency on replication, we tested whether in vitro produced His-PIWI1 can bind 27nt-RNAs. We therefore incubated His-PIWI1 with 4TU-RNA mixture obtained after purification of RNA from 4TU-labelled exconjugants, followed by UV cross-linking (365 nm) and PIWI1-RNPs pull-down with PIWI1-specific antibodies [14]. Using the Small RNA Analysis Kit (Agilent) for microcapillary electrophoresis of RNA a faint 27nt band purified from PIWI1 immunocomplexes could be detected (Fig. 5b, top). To assay the efficiency of DNA replication in the presence of PIWI1 and 27nt-RNAs, we set up various Klenow reactions using pGEM-T-easy-PCNA1 as template. Here, we used the T7 primer-binding site in pGEM-T-easy for priming of linear DNA amplification and a modified T7 primer with non-matching, hairpin-forming additional 5′-nucleotides. Differences in the amount of amplified PCNA1 were then detected using qPCR and PCNA1 primers (Fig. 5b, bottom). The addition of PIWI1 and 27nt-RNAs in a 1:2 molar ratio revealed a significant reduction of amplified DNA between 25 and 1 µM, and this effect was apparently stronger as the effect of 27nt-RNAs alone. The addition of PIWI1 alone, in contrast, did not differ from controls without PIWI1 and 27nt-RNA.
Such ‘replication interference’ through oligonucleotides shown in vitro would provide a mechanism, which could underlie the control of replication timing during polytenization in vivo, and this could be a prerequisite for the temporally coordinated deposition of histone variant-containing nucleosomes within a given timeframe. Notably, according to the current doctrine on the RNAi-induced transcriptional silencing (RITS) for several model eukaryotes (e.g. S. pombe, C. elegans, A. thaliana), it is not assumed that Argonaute/sncRNA complexes bind directly to a DNA target. Rather, they apparently bind via base-pairing to tethered nascent transcripts, which origin from the target regions. Binding of such transcripts through Argonaute/scnRNAs leads to the localized recruitment of chromatin modifiers [38]. For a hypothetical ‘nascent transcript’ IES-targeting model in Stylonychia (Fig. 5c1), we assumed that a putative nascent transcript from a micronuclear MDS locus should contain sequences complementary to IESs. To test whether such a nascent transcript that origins from micronuclear genes could exist, we screened the developmental transcriptomes for reads matching to known IES sequences. Figure 5d shows that the reads mutually exclusively covered MDSs, whereas IESs were sharply omitted (here, 20 h PC). This observation did not support MDS targeting via a nascent transcript, thus pinpointing to a possible 27nt-RNA/dsDNA interaction (Fig. 5c2) or direct 27nt-RNA/ssDNA base-pairing in an open replication bubble (Fig. 5c3), with the latter being favoured through our finding that 27nt-RNAs became enriched, when RNA/DNA hybrids were pulled down with specific antibodies.
27nt-RNAs can impair DNA replication in the replication band of vegetative Stylonychia in vivo
To evaluate whether the ‘RNA-induced DNA replication interference’ could in principle exist in vivo, we exploited another feature characteristic for spirotrichous ciliates—the linear spatiotemporal progression of replication during the S-phase of vegetative macronuclei within a migrating morphological structure called ‘replication band’ (Fig. 6a). Replication bands in Stylonychia macronuclei are disc-shaped accumulations of hundreds of synchronously firing replication foci, which appear at the distal tips of the macronuclei at the onset of S-phase and migrate thereafter to the centre of the macronucleus. Hereby, the motility of macronuclear chromatin is strictly constrained, ensuring that each nanochromosome becomes evenly replicated and its copy number remains preserved (Fig. 6b) [20]. We described above that 27nt-RNAs were absent in vegetative Stylonychia, whereas a low level 21–22nt-RNAs seemed to exist over the entire life cycle (Fig. 2). Concomitantly, at least one PIWI-domain protein is constitutively expressed (PIWI2, see Fig. 1 and Additional file 1: Figure S7). A prediction of the ‘RNA-induced DNA replication interference’ concept would imply that the copy numbers of specific nanochromosomes became influenced, if 27nt-RNAs targeted them during replication. Therefore, we microinjected several 27nt-RNAs (Additional file 1: Table S1) specifically targeting genes (bait RNA), which become differentially expressed during macronuclear development (i.e. H3.7 [Contig610.g675]; MDP2 [Contig9714.g10391]; PCNA1 [Contig13765.g14680]). For control a mock oligo was injected not targeting any known Stylonychia macronuclear DNA sequence. We assumed that during vegetative growth a change in the copy numbers of these genes, including PCNA1—one out of three PCNA genes in Stylonychia, which is only expressed during macronuclear development, would not affect any cellular function during vegetative growth. Importantly, in non-synchronized vegetative Stylonychia cells the copy numbers of several nanochromosomes were studied extensively previously, and these experiments did not give any hint for a theoretically imaginable DNA degradation event under RNAi influence [17].
Then, using qPCR on single cells we compared the relative copy numbers of the targeted nanochromosomes with copy numbers in control cells, wherein nonsense-27nt-RNAs (mock) were injected, at three successive time points. No conspicuous differences or restrictions in growth between mock and bait RNA-treated cells were observed: Following microinjection, cells were grown individually in microtiter plates and inspected in daily routine. The observed growth rates were 2.31-fold/d for bait RNA-treated cells and 2.18-fold/d for mock RNA-treated cells. Intriguingly, we observed a significant reduction in the copy numbers of targeted nanochromosomes between the 1st and 2nd replication cycles after microinjection (mean fold change: 0.17314, p < 0.01) (Fig. 6c). When we assessed the copy numbers of targeted nanochromosomes in the course of continuous cycles (3rd to 4th/5th to 6th), we observed an attenuation of the copy number effects observed after the first two cycles, whereas a weak interference effect on the targeted nanochromosome copy numbers still appeared to be recognizable. Taken together, this observation substantiated the initial prediction that 27nt-RNAs targeting specific nanochromosomes during replication would influence their copy numbers. Consequently, these experiments provided first evidence that an ‘RNA-induced DNA replication interference’-mechanism is possible in principle in vivo and could therefore exist.
