GTSF1 is required for transposon silencing in the unicellular eukaryote Paramecium tetraurelia

Abstract

The PIWI-interacting RNA (piRNA) pathway is crucial for transposon repression and the maintenance of genomic integrity. Gametocyte-specific factor 1 (GTSF1), a PIWI-associated protein indispensable for transposon repression, has been recently shown to potentiate the catalytic activity of PIWI in many metazoans. Whether the requirement of GTSF1 extends to PIWI proteins beyond metazoans is unknown. In this study, we identified a homolog of GTSF1 in the unicellular eukaryote Paramecium tetraurelia (PtGtsf1) and found that its role as a PIWI-cofactor is conserved. PtGtsf1 interacts with PIWI (Ptiwi09) and Polycomb Repressive Complex 2 and is essential for PIWI-dependent DNA elimination of transposons during sexual development. PtGtsf1 is crucial for the degradation of PIWI-bound small RNAs that recognize the organism's own genomic sequences. Without PtGtsf1, self-matching small RNAs are not degraded and results in an accumulation of H3K9me3 and H3K27me3, which may disturb transposon recognition. Our results demonstrate that the PIWI–GTSF1 interaction also exists in unicellular eukaryotes with a role in transposon silencing.

Graphical Abstract

Introduction

Transposable elements (TEs) are found in the genomes of nearly all organisms and can make up a significant portion of the genome (1). They contribute to genetic diversity and evolution by causing mutations, driving gene duplication and creating new genes (2). However, their uncontrolled activity can lead to genomic instability. Therefore, most organisms have evolved mechanisms to silence or limit the activity of TEs. The PIWI-interacting RNA (piRNA) pathway is essential for the repression of TEs in animal germlines as well as some somatic tissues of various non-mammalian species (3–5). piRNAs are transcribed from genomic loci known as piRNA clusters, which are further processed into primary piRNAs with the ability to guide its PIWI-partner to TE transcripts (6,7). Cleavage by a PIWI protein initiates the production of secondary piRNAs, which can further participate in TE repression (4,8). Transcriptional TE silencing is achieved by nuclear PIWI proteins (e.g. Piwi in flies and MIWI2 in mice), which are guided by piRNAs to nascent TE transcripts and generate heterochromatin via DNA or histone methylation (e.g. trimethylation of lysine 9 on histone H3, H3K9me3) (as reviewed in 9). Post-transcriptional TE silencing is mediated by cytoplasmic PIWI proteins (e.g. Aub and Ago3 in flies, MILI and MIWI in mice), which can directly slice target transcripts complementary to their bound piRNA.

Ciliates, one of the most diverse groups of unicellular eukaryotes that emerged approximately one billion years ago, employ a more extreme approach of controlling TEs than most organisms and do not rely on constant transcriptional or post-transcriptional silencing of such elements. They separate germline and somatic functions into distinct nuclei in the same cytoplasm and only allow TEs and TE-derived sequences known as internally eliminated sequences (IESs) to exist in the transcriptionally inactive germline micronuclear (MIC) genome (10–13). During sexual processes, they are eliminated from the active somatic macronuclear (MAC) genome through a piRNA-like DNA elimination pathway (as reviewed in 14,15). In the ciliate Paramecium tetraurelia (P. tetraurelia), about one third of germline-limited sequences are eliminated from the MAC during sexual processes, including TEs and ∼45 000 IESs (11).

The piRNA-like pathway in P. tetraurelia involves a small RNA-mediated comparison of the MIC and MAC genomes, known as the ‘scanning model’, a model originally proposed for a homologous pathway in Tetrahymena, another ciliate model organism (16–18). During this process, small RNAs generated from the germline MIC genome ‘scan’ the somatic MAC genome to remove MAC-matching sequences. This enriches for small RNAs containing MIC-specific sequences (including TEs and IESs), which are used to guide their elimination in the developing new MACs.

In P. tetraurelia, sexual development begins with the bidirectional transcription of the entire MIC genome by RNA polymerase II to produce double-stranded long non-coding RNAs (dsRNAs), which are cleaved into 25 bp small non-coding RNAs (scnRNAs) by Dicer-like proteins (19,20). Double-stranded scnRNAs are transported to the cytoplasm where they are loaded onto PIWI proteins (Ptiwi01/Ptiwi09), and the ‘passenger’ strand is removed (21). The PIWI–scnRNA complexes are transported to the old MAC where they ‘scan’ nascent transcripts by homologous pairing, and those that find complementary sequences are degraded by an unknown mechanism. The remaining scnRNAs containing MIC-specific sequences are transported to the developing new MACs, where they pair with nascent transcripts and recruit Polycomb repressive complex 2 (PRC2) to mediate the deposition of H3K9me3 and H3K27me3 (22–25). These modifications help recruit the domesticated PiggyBac transposase Pgm that excises the MIC-specific DNA (26,27). After excision, the excised DNA initiates the production of a secondary class of small RNAs known as iesRNAs, which serve as a positive feedback loop to ensure complete elimination of all MIC-specific DNA (20,28–29). Finally, the remaining DNA segments (called macronuclear destined sequences, MDSs) are either re-ligated by non-homologous end joining or de novo telomerized to form the new somatic MAC genome (30,31).

Although an increasing number of factors involved in this process have been identified in the past 20 years, many questions remain unanswered. One of these questions is how MAC-matching MDS-scnRNAs are selected and degraded to prevent their entry into the new MACs where they could guide PRC2 and lead to errant DNA elimination. In this study, we approached this question by identifying and characterizing a PIWI- and PRC2-interacting protein essential for this process. This protein is a homolog of Gametocyte-specific factor 1 (GTSF1), an evolutionarily conserved small zinc finger protein, which has been reported in many metazoans (32–36). GTSF1 was first identified in unfertilized eggs, ovaries and testes in mouse and later revealed to be an essential component of the piRNA pathway in various organisms, including mouse, Drosophila and silkworm (33–39). GTSF1 directly interacts with both cytoplasmic and nuclear PIWI proteins and is essential for transcriptional and post-transcriptional regulation of TEs. A recent study revealed that GTSF1 enhances the intrinsically weak RNA cleavage activities of catalytically active PIWI proteins (40).

In this study, we identified a homolog of GTSF1 in the unicellular eukaryote P. tetraurelia (PtGtsf1) and found the relationship between GTSF1 and PIWI to be conserved. PtGtsf1 plays a crucial role in the repression of TEs by facilitating the degradation of scnRNAs recognizing the organism's own genomic sequences, a mechanism that distinguishes MDS- from IES- and TE-DNA. Our results demonstrate that PtGtsf1 interacts with both PIWI and the PRC2 complex and regulates the scnRNA pool that guides the PRC2 complex to mark sequences for elimination. The identification of GTSF1 in protists indicates that GTSF1 was present in the last common ancestor of all animals with a role in RNA-mediated TE silencing.

Materials and methods

Paramecium tetraurelia cultivation and autogamy induction

Paramecium tetraurelia strain 51, mating type seven, was used to perform the experiments. Cells were cultured in wheat grass power (WGP) medium (Pine international, Lawrence, KS) bacterized with Klebsiella pneumoniae and supplied with 0.8 mg/L β-sitosterol (Shanghai Yuanye Bio-technology) (41). Autogamy was induced by starvation.

Gene silencing

Gene silencing was carried out as described in Beisson et al. (42). A fragment of the coding region from 1 to 366 bp of PtGTSF1 (Gene ID: PTET.51.1.G0490019) was cloned into the L4440 plasmid and then transformed into the HT115 (DE3) strain of Escherichia coli. Silencing constructs of EZL1, PtCAF1, DCL2, DCL3 and PGM were made according to published papers (19,23,26).

Briefly, silencing was performed as follows. Bacteria with silencing constructs were cultured overnight in LB medium, and then 1:100 diluted in 1× WGP medium and incubated overnight. The culture was further diluted with 1× WGP to an OD₆₀₀ of 0.04 and allowed to grow until the OD₆₀₀ reached a range between 0.07 and 0.1. Isopropyl-β-d-thiogalactopyranoside was added to a final concentration of 0.4 mM, and the culture was incubated for at least 4 h to induce the production of dsRNA. Prior to seeding, the silencing medium was supplemented with 0.8 mg/L β-sitosterol, and the P. tetraurelia cells were washed in Tris-HCl (10 mM, pH 7.5) and seeded at a density of 200 cells/ml.

DNA preparation and microinjection

The entire PtGTSF1 gene, including flanking sequences (206-bp upstream and 179-bp downstream), was cloned into the pCE2 vector using the 5 min^TM TA/Blunt-Zero Cloning Kit (Vazyme Biotechnology). A codon-optimized GFP or FlagHA was then inserted after the start codon ATG of PtGTSF1. Plasmids containing GFP tagged PTIWI09, EZL1 and PGM were constructed as in the published papers (21,23,26). After confirmation of the plasmids by sequencing, the constructs were extracted from bacteria and linearized by enzyme digestion in the backbone, followed by purification using phenol chloroform (pH 8.0) and Ultrafree-MC Centrifugal Filters (Millipore).

To perform microinjections, post-autogamous P. tetraurelia cells were recovered to the vegetative stage and grown for 4–8 divisions. DNA was injected into the macronucleus as described in previous study (43). Successful injections were confirmed by polymerase chain reaction (PCR) and the cells were cultured for follow-up experiments.

Immunofluorescence

Around 150 000 cells were centrifuged and washed twice with 10 mM Tris-HCl (pH 7.5). The cells were permeabilized with 1% Triton X-100 in 1× PHEM buffer (10 mM EGTA, 25 mM HEPES, 2 mM MgCl₂, 60 mM PIPES (pH 6.9)) for 10 min and fixed with 2% Paraformaldehyde in 1× PHEM buffer for 10 min. Next, the cells were blocked in a 3% bovine serum albumin (BSA) in Tris buffered saline (0.15 M NaCl, 10 mM Tris-HCl) with 1% Tween 20, 10 mM ethylene glycol-bis (β-aminoethyl ether)-N,N,N′,N′-tetraacetic acid (EGTA), and 2 mM MgCl₂ (TBSTEM) for 30 min. After blocking, the cells were incubated with a 1:200 dilution of the following antibodies overnight at 4°C: anti-trimethyl-Histone H3 (Lys27) (07–449, Millipore), anti-trimethyl-Histone H3 (Lys9) (07–442, Millipore) or anti-gamma H2A.X (phosphor S139) antibody (ab11174, abcam). Afterward, the cells were washed twice with a 3% BSA in TBSTEM and incubated with a goat anti-rabbit Alexa Fluor 546 Secondary antibody (A-11071, Invitrogen) at a dilution of 1:4000 for an hour. Following two washes with 1× phosphate-buffered saline (PBS), the cells were spread onto glass slides and mounted with ProLong Glass Antifade Mountant (P36980, Invitrogen), containing the blue DNA stain NucBlue. Imaging was performed with a Axio Imager D2 (Zeiss) and the Zen 2 software.

Imaging and measurement of the mean fluorescence intensity

For cells injected with GFP-tagged constructs, approximately 150 000 cells at the desired timepoint were centrifuged and fixed in 75% ethanol. Before imaging, the cells were washed three times with 1× PBS and stained with 4,6-diamidino-2-phenylindole (DAPI). Then, the cells were spread onto glass slides for imaging. All imaging was performed with a Axio Imager D2 (Zeiss) and the Zen 2 software.

Images used to compare signal intensities were taken under the same conditions. ImageJ (44) was used to measure the fluorescence intensity with background subtraction and regions of interested selected by polygon selections.

Immunoprecipitation

Around 1.5 million cells were collected and washed twice with 10 mM Tris-HCl (pH 7.5), followed by two washes with cold 1× PBS. After centrifugation and removal of the supernatant, the cell pellet was resuspended in 2 ml of lysis buffer (50 mM Tris-HCl (pH 8.0), 150 mM NaCl, 5 mM MgCl₂, 1 mM DTT, 1× complete EDTA-free protease inhibitor cocktail tablet (Roche), 1% Triton X-100 and 10% glycerol). The samples were subjected to sonication using a Branson digital sonifier SFX250, with an amplitude of 55% for 15 s. Following sonication, the soluble fraction was collected after clearing by centrifugation at 15 060 rpm for 30 min. Next, 50 μl of Anti-HA Affinity Matrix (11815016001, Roche) was washed three times with IP buffer (10 mM Tris-HCl (pH 8.0), 150 mM NaCl, 1 mM MgCl₂, 0.01% NP40 and 5% glycerol), and the soluble fraction (1 ml) was added to the beads, followed by an overnight incubation at 4°C. After the incubation, the beads were washed six times with IP buffer, the supernatant was discarded, and the beads fraction were used for further experiments.

Western blot

Immunoprecipitated proteins were separated by sodium dodecyl-sulfate polyacrylamide gel electrophoresis (SDS-PAGE) and transferred to a nitrocellulose membrane (Amersham Protran, GE Healthcare Life Sciences) using a wet transfer method. The membrane was blocked with 10% skim milk in phosphate-buffered saline with 0.1% Tween 20 (PBST) for 1 h at room temperature. After blocking, the membrane was incubated overnight at 4°C with a Rabbit anti-HA primary antibody (3724S, Cell Signaling Technology) at a dilution of 1:1000. Next, the membrane was washed three times with PBST and incubated with a 1:8000 diluted secondary antibody (ProteinFind^® Goat Anti-Rabbit IgG (H + L), HRP Conjugate, HS101-01, Transgen Biotechnology) at room temperature for 1 h. The membrane was washed three times with PBST, followed by one wash with PBS. Pierce^TM ECL Western Blotting Substrate (32106, Thermo Scientific) was used to develop the signal.

Mass spectrometry

Immunoprecipitated proteins were separated by SDS-PAGE electrophoresis, stained with Coomassie brilliant blue R-250 (Solarbio) and cut into gel slices. After destaining and dehydration, the gel bands were subjected to reduction by incubation with 10 mM DTT at 56°C for 1 h, followed by alkylation with 55 mM IAA for 45 min. The proteins were digested with trypsin overnight at 37°C. The resulting peptides were separated by the UltiMate3000 RSLCnano ultra-high-performance liquid system using a gradient of 65 min at the rate of 400 nl/min. Solvent A was water with 0.1% FA, and solvent B was 98% ACN with 0.1% FA. The gradient was 5–8% B for 6 min, 8–30% B for 34 min, 30–60% B for 5 min, 60–80% B for 3 min, 80% B for 8 min, 80–5% B for 2 min and 5% B for 7 min. The peptides were then analyzed by the Thermo Scientific^TM Q Exactive^TM mass spectrometer in Data Dependent Acquisition mode. The ion source voltage was 1.8 kV, the full scan range of MS was 350–2000 m/z, and the scanning resolution was set to 70 000. The top 20 peptides were selected for MS2, and the resolution was 17 500. The proteomic data were queried using MaxQuant (2.6.2.0) (45) against the protein sequences of Paramecium tetraurelia downloaded from ParameciumDB (ptetraurelia_mac_51_annotation_v2.0.protein.fa). The search parameters were set as follows: the enzyme was specified as trypsin; the missed cleavage was set to 2; the minimum peptide length was 7; the fixed modification was cysteine carbamidomethyl (C); the variable modifications included Oxidation (M) and Acetyl (Protein N-term); LFQ of label-free quantification was selected. The output of MaxQuant was further analyzed by LFQ-Analyst (Dev.) (https://analyst-suites.org/apps/lfq-analyst-dev/) to compare the enrichment with imputation type of zero and t-statistics-based false discovery rate (FDR) correction (46).

Chromatin immunoprecipitation

Around 6 million cells were centrifuged and washed twice with 10 mM Tris-HCl (pH 7.5). The cells were crosslinked with 1% methanol-free formaldehyde (28908, Thermo Fisher Scientific) at room temperature for 10 min. After quenching with 0.125 M glycine (room temperature, 5 min), the cells were washed twice with 1× PBS. Then, the cells were incubated with 2.5 volume lysis buffer (0.25 M sucrose, 10 mM Tris pH 6.8, 10 mM MgCl₂, 0.2% Nonidet P-40, 1.5 mM PMSF, 1.5 × Complete EDTA-free Protease Inhibitor Cocktail tablets (Roche)) on ice for 5 min and lysed with a Potter–Elvehjem homogenizer, followed by two washes with wash buffer (0.25 M sucrose, 10 mM MgCl₂, 10 mM Tris pH 7.5, 1.5 mM PMSF, 1.5 × Complete EDTA-free Protease Inhibitor Cocktail tablets (Roche)). The pellet was resuspended with 3 ml RIPA buffer (50 mM Tris pH 8.0, 150 mM NaCl, 10 mM EDTA, 1% Triton X-100, 1% Sodium deoxycholate, 1.5 mM PMSF, 1.5 × Complete EDTA-free Protease Inhibitor Cocktail tablets (Roche), 1% Sodium Dodecyl Sulfate) and aliquoted to 200 μl per tube. The samples were sonicated with the Bioruptor (Diagenode) for 30 min (30 s on, 30 s off) to shear the chromatin to sizes between 200 and 1000 bp. After centrifugation at 20 000 rcf at 4°C for 20 min, the supernatant containing 16 μg of DNA was diluted 5-fold with RIPA buffer without sodium dodecyl sulfate and incubated with 3 μl H3K27me3 antibody (C15410195, Diagenode) overnight at 4°C. Meanwhile, 10% of the supernatant was taken as input. Antibody bound chromatin was captured with Dynabeads^TM M-280 Sheep anti-Rabbit IgG (11203D, Invitrogen) after five washes with IP buffer (10 mM Tris-HCl (pH 8.0), 150 mM NaCl, 1 mM MgCl₂, 0.01% NP40, 5% glycerol). The ChIP samples and input were reverse cross-linked at 65°C overnight with proteinase K and NaCl added, and the DNA was extracted with phenol chloroform.

Enrichment of the ChIP-DNA and input DNA was analyzed by quantitative real-time PCR (qPCR), which was performed with the ChamQ SYBR Color qPCR Master Mix (Vazyme) and the Applied Biosystems 7500 Fast Real-Time PCR system. Primers used for qPCR are listed in Supplementary Table S1.

Amino acid sequence alignment and phylogenetic tree construction

The amino acid sequences of GTSF1 from different organisms were downloaded from the UniProt and NCBI databases with the accession numbers shown in Supplementary Table S2. The alignment in Supplementary Figure S1 were performed in Geneious with MUSCLE alignment. The sequences for the phylogenetic tree was aligned by MEGA v.11 with Clustal using default parameters and the Neighbor-joining tree was constructed with 1000 bootstrap replications with Jones–Taylor–Thornton model (47).

Survival test

After undergoing autogamy, the cells were transferred into individual wells containing bacterized 0.2 × WGP (0.8 mg/L of β-sitosterol added) to recover to a vegetative stage. The growth of the cells was monitored for three consecutive days. If the cells grew similarly as the control, they were categorized as healthy, otherwise as sick (fewer cells than the control) or dead (only one or no alive cell).

Macronuclear isolation and DNA extraction

The macronuclei of post-autogamous cells were isolated following the protocol in Arnaiz et al., 2012 (11). Approximately 1.5 million cells were centrifuged and washed twice with 10 mM Tris-HCl (pH 7.5). The cell pellet was resuspended in 2.5 volumes of lysis buffer 1 (0.25 M sucrose, 10 mM MgCl₂, 10 mM Tris-HCl (pH 6.8) and 0.2% NP-40). After incubating on ice for 5 min, a Potter–Elvehjem homogenizer was used to disrupt the cell membranes while keeping the nuclei intact. Subsequently, the lysate was washed twice with wash buffer (0.25 M sucrose, 10 mM MgCl₂ and 10 mM Tris-HCl (pH 7.5)). The pellet was resuspended in three volumes of sucrose buffer (2.1 M sucrose, 10 mM MgCl₂ and 10 mM Tris-HCl (pH 7.5)) and carefully layered on top of 3 ml of sucrose buffer in a centrifuge tube (344060, Beckman Coulter). The tube was filled with wash buffer to create a sucrose gradient. The macronuclei were isolated by ultracentrifugation at 35 000 rpm, 4°C for an hour using a Beckman Optima L-90K Ultracentrifuge. Following ultracentrifugation, MAC DNA was extracted by phenol chloroform. Next-generation sequencing was performed by the Novogene company (Tianjin, China).

RNA extraction, mRNA-seq and sRNA-seq

Half a million cells at the desired timepoint were collected and washed twice with 10 mM Tris-HCl (pH 7.5). After removing the supernatant, the cells were frozen in liquid nitrogen until RNA extraction. Total RNA was extracted with TRIzol (Sigma-Aldrich), following the TRIzol reagent BD protocol.

The extracted total RNA was sent to Novogene company to do library preparation and sequencing. For the mRNA-seq, the mRNA was enriched using magnetic beads with Oligo(dT) and then broken into short fragments, followed by cDNA synthesis and AMPure XP beads purification. The purified cDNA then underwent end repair, A-tailing, adapter ligation, size selection, amplification, and purification with AMPure XP beads to obtain the final library. After quality assessment, the library was loaded on a Novaseq 6000 to obtain paired-end 2× 150 bp reads.

The small RNA libraries were prepared with NEBNext^® Multiplex Small RNA Library Prep Set for Illumina^® (NEB, USA). The purified RNA was ligated with the NEB 3′ SR Adapter and reverse-transcribed into double-stranded DNA molecules with M-MuLV Reverse Transcriptase (RNase H⁻). Then, the cDNA was amplified and purified in an 8% polyacrylamide gel in which fragments ranging from 140 to 160 bp were recovered and dissolved in elution buffer. Subsequently, the purified cDNA was loaded on a Novaseq 6000 to obtain single-end 1× 50 bp reads.

Reference genomes

The following reference genomes were used in the IES and TE analyses and for read mapping:

MAC: http://paramecium.cgm.cnrs-gif.fr/download/fasta/ptetraurelia_mac_51.fa

MAC + IES: http://paramecium.cgm.cnrs-gif.fr/download/fasta/ptetraurelia_mac_51_with_ies.fa

TE: https://paramecium.i2bc.paris-saclay.fr/files/Paramecium/tetraurelia/51/annotations/ptetraurelia_mic2/ptetraurelia_TE_consensus_v1.0.fa

sRNA-seq analysis

Single-end sRNA reads ranged from 20 to 45 nt were processed using fastp (-q 20 -l 20; v0.23.1; (48)) before mapping using Bowtie (v1.3.1; (49)) with default parameters. The size-selected reads were mapped to the following reference datasets sequentially: the genome of Klebsiella (CP002824.1); IESs from the MAC + IES genome (ptetraurelia_mac_51_with_ies.fa); the MAC genome (referred to as MDSs) (ptetraurelia_mac_51.fa). The 23 nt siRNA reads mapped to the RNAi targets were removed, and the scnRNA read counts were normalized using the total number of reads and siRNAs.

mRNA-seq analysis

Paired-end reads were filtered using fastp with -q 20, -l 75, -c, –detect_adapter_for_pe and then mapped against the MAC genome of P. tetraurelia (ptetraurelia_mac_51.fa) using hisat2 (v2.2.1; (50)) with default parameters. Reads were counted using featureCounts (v2.0.1, (51)) with the parameter of -p and the output used as the input for DESeq2 (52). Differentially expressed genes (DEGs) in the PtGTSF1 KD were identified as those with an adjusted P < 0.1 and with at least a 4-fold change relative to the corresponding control timepoint. Genes were classified as upregulated (fold change ≥ 4) or downregulated (fold change ≤¼). Gene ontology annotation of DEGs were obtained from the ParameciumDB, and the Gene ontology functional enrichment was performed using the clusterProfiler package (53).

IES retention analysis

Paired-end reads were filtered using fastp with -w 16, -I 75 and then mapped to the MAC and MAC + IES genomes sequentially with the Map module in ParTIES with default parameters (54). IES retention scores (IRSs) were calculated with the MIRET component of ParTIES using the whole score option. IGV (2.17.4) was used to visualize the IES deletion levels between different timepoints (55).

Results

PtGtsf1 interacts with Ptiwi09 and PRC2

In a previous study, we performed immunoprecipitation and mass spectrometry to identify the PRC2 complex and its interaction partners in P. tetraurelia (24). In addition to identifying the subunits of the complex, we discovered a homolog of PtGtsf1 (Supplementary Figure S2A–D). We also re-analyzed mass spectrometry data from a related study that reported the interaction partners of the scnRNA-binding PIWI protein Ptiwi09 and found that PtGtsf1 also co-precipitates with Ptiwi09 (Supplementary Figure S2E) (22). Since GTSF1 is an essential component of the piRNA pathway in animals and is well known to interact with PIWI proteins, we decided to further characterize this protein (32,34–36,38,40).

PtGtsf1 (PTET.51.1.P0490019 in ParameciumDB) is predicted to have two CHHC-type zinc fingers, which is a hallmark of GTSF1 proteins (Figure 1B; Supplementary Figure S1) (56). Mammals have three GTSF1 paralogs (GTSF1, GTSF1-like and GTSF2) and Drosophila has four (Asterix/DmGTSF1/CG3893, CG14036, CG32625 and CG34283) (39,57). In a phylogenetic tree of GTSF1 and GTSF1-like proteins from various organisms, PtGtsf1 clusters with the GTSF1 of Drosophila melanogaster and Caenorhabditis elegans (Figure 1C). Although a previous study identified three homologs of GTSF1 in P. tetraurelia by an iterative PSI-BLAST search (PTET.51.1.G0170076, PTET.51.1.G0860105 and PTET.51.1.G0160267), PtGtsf1 was not one of them, and none of these three homologs were detected in the immunoprecipitated proteins of Ptiwi09 (58). To further confirm the interactions identified in the previously published mass spectrometry data, we expressed a HA-tagged PtGtsf1 in P. tetraurelia and performed immunoprecipitation using an anti-HA antibody to identify interaction partners of PtGtsf1 (Supplementary Figure S2F, G). Both Ptiwi09 and PRC2 subunits were co-immunoprecipitated with PtGtsf1, confirming the interactions (Figure 1D). Considering the previously established interaction between Ptiwi09 and PRC2 (22,24), we conclude that PtGtsf1, PRC2 and Ptiwi09 interact in P. tetraurelia (Figure 1E).

Figure 1.

PtGtsf1 interacts with Ptiwi09 and PRC2 and is exclusively expressed during sexual development in Paramecium tetraurelia. (A) Sexual reproduction (autogamy) and photographs of P. tetraurelia. When cells are starved, two micronuclei (MICs) undergo meiosis to produce eight haploid nuclei. One of the eight haploid nuclei divides mitotically and the rest degrade. Two haploid nuclei fuse to form the zygotic nucleus, which divides twice before two products become the new MICs, and the other two develop into the new macronuclei (MACs). Finally, the cell divides once, distributing the nuclei into two cells. When the post-autogamous cells are refed, they recover to the vegetative stage of the next generation. The timepoints collected in this study are labeled in red (Early, Late, Late++ and F1 (Filial generation 1)). Arrowheads point to micronuclei. Scale bar: 30 μm. (B) Protein domains of GTSF1 in Paramecium tetraurelia (Pt), Drosophila melanogaster (Dm) and Caenorhabditis elegans (Ce). (C) Neighbor-Joining tree of GTSF1 and its paralogs in P. tetraurelia and metazoans. The support values higher than 50 were shown in the tree. Created with BioRender.com. (D) Mass spectrometry of proteins immunoprecipitated by PtGtsf1 with wild type as the control (CTRL). (E) Diagram to show the interaction between PtGtsf1, PRC2 and Ptiwi09. Created with BioRender.com. (F) RNA-seq expression profiles of PtGTSF1, PGM, EZL1, DCL2 and DCL3. Veg: vegetative growth before starvation; Mei: meiosis, corresponding to Early in (A); Frg: fragmented old MAC; Dev1-4: stages of new MAC development with Dev2/3 aligns to Late and Dev4 aligns to Late++ in (A). (G) Localization of PtGtsf1 during vegetative and sexual development (autogamy). From left to right, progression of autogamy. Arrows indicate new MACs. Scale bar: 10 μm.

PtGtsf1 is essential for sexual development

The life cycle of Paramecium is separated into vegetative and sexual stages (Figure 1A). According to publicly available mRNA-seq data, PtGTSF1 is specifically expressed during sexual development (autogamy), with the highest levels detected at the meiotic stage (Figure 1A, F) (56). This expression pattern is reminiscent of genes encoding core components of PRC2 and scnRNA-related proteins, such as Ptiwi01 and Ptiwi09, and Dicer-like proteins Dcl2 and Dcl3 (20,21).

We next assessed the localization of PtGtsf1 by fusing it with a green fluorescent protein (GFP) (Figure 1G). To confirm that the fusion protein is functional, we silenced PtGTSF1 in wild type and PtGTSF1-GFP transformed cells. The wild type was unable to recover to a vegetative stage after autogamy in the absence of PtGtsf1, while the introduction of PtGTSF1-GFP could rescue this effect and successfully recover the cells to a vegetative stage, confirming that the protein of PtGtsf1 fused with GFP is functional. No PtGtsf1-GFP fluorescence was observed during vegetative growth (Figure 1G). Upon induction of autogamy by starvation, PtGtsf1 initially appeared in the intact old MAC and remained there when the MAC threaded and fragmented. In early new MAC development, PtGtsf1 exists both in the old and new MACs simultaneously. As the new MACs continue to develop, the signal gradually diminishes, leaving only faint signals in the new MACs that eventually disappear. Through the entire life cycle, PtGtsf1 was never detected in the MICs.

Both the expression profile and protein localization suggest that PtGtsf1 has a function specifically during sexual development in P. tetraurelia (Figure 1F, G). To further investigate its function, we performed a knockdown of PtGTSF1 (PtGTSF1 KD) and used an empty vector (EV) as control. During the vegetative stage, PtGTSF1 KD cells grow similarly to the EV control, confirming that it is dispensable for vegetative growth. To accurately assess its function during autogamy, we studied four representative stages (Figure 1A): Early, corresponding to the meiotic stage, in which 30–50% of cells have a fragmented old MAC; Late, approximately 14 h after Early with more than 70% of cells having visible new MACs. At this stage, TEs and IESs are being eliminated from the new MACs; Late++, 36 h after Late when the new MACs are almost fully developed; and F1 (Filial generation 1), 24 h after feeding Late++ cells, corresponding to the vegetative stage of next life cycle.

PtGTSF1 KD cells can enter autogamy and there are no noticeable differences compared to the control until the Late++ stage, at which point the cells have an increased number of old MAC fragments compared to the control (median: 11.0 versus 22.5), and this difference remains significant until the F1 stage (4.0 versus 11.5) (Figure 2A, B). Unlike the control, PtGTSF1 KD cells cannot recover to the vegetative stage after autogamy. After re-feeding, PtGTSF1 KD cells survive the first day; however, their morphology is aberrant (Figure 2C). The posterior end changed from broadly rounded to irregular, and the cells became shorter and wider with a decreased length/wide ratio (Figure 2C, D). From the second day, cell numbers in the PtGTSF1 KD culture show obvious differences to the control with nearly all of them morphologically or behaviorally abnormal (Figure 2E). Taken together, the completion of sexual development, but not vegetative growth, requires PtGtsf1 in P. tetraurelia.

Figure 2.

PtGtsf1 is essential for sexual development and the repression of TEs. (A) DAPI staining to show the old MAC fragments in Late++ and F1 stages of EV (empty vector, negative control) and PtGTSF1 KD. Arrows indicate new MACs. Scale bar: 10 μm. (B) Numbers of old MAC fragments in EV and PtGTSF1 KD. Numbers in brackets under the boxes are the numbers of individuals whose fragments were counted. Horizontal line and the number to the right of the line denotes the median. ****, P < 0.0001 (unpaired t-test). (C) Photographs of Paramecium tetraurelia on the first day after feeding. Scale bar: 30 μm. (D) The length, width and length/width ratio of P. tetraurelia cells on the first day after feeding. (E) Survival test of EV and PtGTSF1 KD cells. Cells were monitored for three consecutive days after feeding, denoted by D1, D2 and D3. Bars with left diagonal slashes, horizontal stripes, and dots represent healthy, sick, and dead, respectively. (F) Venn diagram depicting the overlap of IESs (IRSs ≥ 0.1) affected by PtGTSF1 KD, DCL2/3 KD, EZL1 KD and PGM KD. (G) Transposons (TEs) retained in EV, PtGTSF1 KD, DCL2/3 KD and EZL1 KD. The retention level of TEs is reflected by FPKM values. (H) Expression of TEs in Late, Late++ and F1 stages of EV and PtGTSF1 KD. (I) Distribution of IRSs in EV, PtGTSF1 KD, DCL2/3 KD and EZL1 KD. Higher scores correspond to a more severe IES retention. The break on the axis indicates that the axis is discontinuous.

PtGtsf1 is required for transposon silencing

GTSF1 is an essential component of the piRNA pathway in animals and plays a crucial role in the repression of TEs (34–36,38,40). In ciliates, the repression of TEs is achieved through DNA elimination during sexual development, and we therefore investigated whether DNA elimination is affected in the absence of PtGtsf1. To do this, we isolated new MACs from post-autogamous cells and extracted genomic DNA in EV control and PtGTSF1 KD cultures, as previously described (11), followed by next-generation sequencing. We found the elimination of TEs belonging to LINE, SINE, SOLO-ORF and TIR families to be impaired by PtGTSF1 KD, albeit to a weaker extent than DCL2/3 and EZL1 KD (Figure 2G). This was also accompanied with an increase in RNA expression of the corresponding TE families (Figure 2H).

Silencing of PtGTSF1 also affected the elimination of ancient TE remnants known as IESs. The retention level of IESs was estimated by calculating IRSs, in which a higher IRS indicates a greater extent of IES retention (54). Overall, 2 783 IESs (IRS ≥ 0.1) were retained in the new MAC genome in the absence of PtGtsf1 (Figure 2I). Similar to the effect on TEs, the influence on IESs appears subtle, only occupying 6.2% of the total IES pool (44 817). Nearly all the retained IESs (2 740, 98.5%) in PtGTSF1 KD are a subset of the IESs affected by EZL1 KD (31 250 IESs), and two thirds (1 727 IESs) are shared with DCL2/3 KD (4 897 IESs) (Figure 2F). However, as previously mentioned, PtGTSF1 KD cells had significantly higher numbers of old MAC fragments than the control, suggesting that the degradation of the old MAC is affected (Figure 2A, B). Since this increases the contamination of old MAC DNA in PtGTSF1 KD but not the control, the impact on TEs and IESs is likely underestimated.

PtGtsf1 is required for the degradation of PIWI-bound small RNAs

Our results thus far are consistent with a role of PtGtsf1 in TE repression, similar to the role of GTSF1 in metazoans. However, the piRNA-like pathway found in P. tetraurelia differs from the metazoan piRNA pathway in many ways, and the role of PtGtsf1 is still unclear. Additionally, the precise role of GTSF1 complexed with nuclear PIWI proteins has not yet been demonstrated in any organism. We therefore sought to determine which step of the process is affected by the absence of PtGtsf1 by interrogating the small RNA (sRNA) population. We extracted and deep sequenced sRNAs from control and PtGTSF1 KD cells. In the control, the percentage of 25 nt scnRNAs decreased from Early to Late stages, corresponding to the degradation of MAC-matching MDS-scnRNAs in the old MAC (Figure 3A, B); however, this decrease did not take place in PtGTSF1 KD (Figure 3E, F).

Figure 3.

PtGtsf1 is required for the degradation of PIWI-bound small RNAs. (A–D) The proportion of sRNAs mapping to various features in EV from Early to F1 stages. (E–H) The proportion of sRNAs mapping to various features in PtGTSF1 KD from Early to Veg stages. (I–L) Normalized counts of scnRNAs mapped to MDSs, IESs and TEs in EV and PtGTSF1 KD. IES: internal eliminated sequence; TE: transposon; Kleb: Klebsiella pneumoniae (food bacteria of Paramecium); MDS: macronuclear destined sequence.

To exclude the possibility that the lower percentage of MDS-scnRNAs in the Late timepoint of the control is due to the massive production of iesRNAs, a secondary class of sRNAs ranging from 26 to 31 nt, we normalized the scnRNA counts to 23 nt siRNAs. After normalization, we found the amount of MDS-scnRNAs in the Late timepoint of PtGTSF1 KD to be much higher than that in the control, and this was also the case in Late++ and F1 stages (Figure 3J–L). Already in the Early stage, the amount of scnRNAs in PtGTSF1 KD was higher than that in the control (Figure 3I). According to previous studies, the decrease of MDS-scnRNAs compared to IES-scnRNAs between Early and Late stages is due to their degradation after pairing with the MAC genome, a process known as scnRNA selection or scanning (21,59). The higher proportion of MDS-scnRNAs in PtGTSF1 KD may be due to an increased production of scnRNAs or because MDS-scnRNAs cannot be degraded after scanning. In P. tetraurelia, scnRNAs are produced from the entire MIC genome, which include MDS, IES and TE sequences; however, only the amount of MDS-scnRNAs increased in PtGTSF1 KD, while scnRNAs containing IES and TE sequences were comparable to the control (Figure 3I). Considering that PtGtsf1 does not enter MICs (Figure 1G), and a related study on bioRxiv found that the maternal RNA production (i.e. precursors of scnRNAs) is not affected by PtGTSF1 KD (60), this is likely a problem of scnRNA degradation rather than production.

Loss of PtGtsf1 also affected another class of sRNAs involved in DNA elimination, the iesRNAs (20). In the control, iesRNAs are massively generated in the Late stage of development and gradually degrade after the elimination of IESs and TEs (Figure 3B–D). In the absence of PtGtsf1, this process is significantly delayed, and iesRNAs cannot be produced until the Late++ stage, 36 h later than the Late stage (Figure 3F, G). While the production of iesRNAs is delayed in PtGTSF1 KD, their degradation does not appear to be affected as they were almost entirely degraded by the F1 stage (Figure 3H). Taken together, PtGtsf1 is involved in the genome scanning process in P. tetraurelia by regulating PIWI-bound sRNAs, but its effects on scnRNAs and iesRNAs differ. PtGtsf1 is required for the degradation but not the production of scnRNAs and in contrast, it is needed for the duly production rather than the degradation of iesRNAs.

Ezl1 can enter new MACs and deposit H3K9me3 and H3K27me3 in the absence of PtGtsf1

Since PtGtsf1 is required for scnRNA degradation and interacts with the scnRNA-binding PIWI protein Ptiwi09, we next sought to assess the location of these undegraded MDS-scnRNAs. In P. tetraurelia, scnRNAs bound to Ptiwi09 shuttle between different nuclei, and scnRNAs not complexed with PIWI were found to be unstable (21,28). Thus, we can visualize the location of scnRNAs using a GFP-fused Ptiwi09. Upon PtGTSF1 KD, Ptiwi09-GFP was still able to enter the new MACs by the Late stage and was not arrested in the old MAC or the cytoplasm, suggesting that the undegraded MDS-scnRNAs can be transferred into the new MACs (Figure 4A). Moreover, the signal of Ptiwi09-GFP in new MACs of PtGTSF1 KD was stronger than in the control, and remained detectable even in the F1 stage (Figure 4A, B). These findings align well with our sRNA-seq results, supporting the conclusion that MDS-scnRNAs cannot be degraded without PtGtsf1 (Figure 3). Surprisingly, however, these undegraded MDS-scnRNAs appears to have entered the new MACs.

Figure 4.

Ptiwi09 can enter new MACs and the intensity and distribution of PRC2 are altered in the absence of PtGtsf1. (A, C) Localization of Ptiwi09-GFP (A) and Ezl1-GFP (C) in EV and PtGTSF1 KD. Arrows indicate new MACs. Scale bar: 10 μm. (B, D) Mean intensity of Ptiwi09-GFP (B) and Ezl1-GFP (D) in new MACs measured by ImageJ. Numbers in brackets under the boxes are the numbers of individuals measured. The median is listed near its line. ****, P < 0.0001 (unpaired t-test).

In the new MACs, scnRNAs normally guide the PRC2 complex to catalyze H3K9me3 and H3K27me3 on sequences destined for elimination (22–25). Since the loss of PtGtsf1 affected the scnRNA population but Ptiwi09 was still able to enter the new MACs, we wondered whether these undegraded MDS-scnRNAs were ‘functional’ (i.e. able to guide PRC2). We therefore investigated the localization of PRC2 in the absence of PtGtsf1 using a GFP-tagged Ezl1 (the catalytic subunit of PRC2). In the control, the Ezl1-GFP entered the new MACs by the Late stage of development; however, when PtGtsf1 was absent, we observed many small PRC2 foci remaining in the fragmented old MAC in the late stage (Figure 4C). Moreover, while the complex was able to enter the new MACs, it did not form foci as the control.

Since the PRC2 complex can enter new MACs but not form foci, we next assessed whether it was catalytically active by immunofluorescence of H3K9me3 and H3K27me3. In P. tetraurelia, H3K9me3 and H3K27me3 are found in the old MAC in the early stage and in the new MACs in the late stage of development (23,24,61). In both EV and PtGTSF1 KD, H3K27me3 became detectable in the old MAC at the earliest stages of autogamy (prior to fragmentation), and H3K9me3 became detectable when the old MAC fragmented (Figure 5A, B). Surprisingly, the signal in PtGTSF1 KD was stronger than in the control for both modifications. To verify that this phenomenon was not a coincidence, we repeated the experiment and observed similar results. We also measured the mean signal intensities of both modifications in the fragmented old MAC by ImageJ, revealing that the signal intensities in PtGTSF1 KD are significantly higher than those in the EV control (Figure 5C, D).

Figure 5.

Loss of PtGtsf1 affects the intensity and distribution of H3K9me3 and H3K27me3. (A, B) Immunofluorescence of H3K9me3 (A) and H3K27me3 (B) during autogamy. Arrows indicate new MACs. The inset shows one of the new MACs. Scale bar: 10 μm. (C, D) Mean intensity of H3K9me3 (C) and H3K27me3 (D) in the fragmented old MAC in the Early stage. Numbers in brackets under the boxes are the numbers of individuals measured. Broken lines within the boxes are the 25% percentile, median and 75% percentile, respectively. The median is listed near its line. ****, P < 0.0001 (unpaired t-test). (E, G) Immunofluorescence of H3K9me3 (E) and H3K27me3 (G) in EV/PGM KD and PtGTSF1/PGM KD. (F, H) Mean intensity of H3K9me3 (F) and H3K27me3 (H) in new MACs of EV/PGM and PtGTSF1/PGM KD cells. (I, J) Enrichment of H3K27me3 on house-keeping genes (Actin, GAPDH, ST PPase, ST Kinase and Helicase) and TEs in EV/PGM KD (I) and PtGTSF1/PGM KD (J), measured by ChIP-qPCR. Quantitative data are expressed as percentage of ChIP over input.

Moreover, H3K9me3 and H3K27me3 persisted in the new MACs of PtGTSF1 KD cells much beyond that of the control (Figure 5A, B). By the Late++ stage, both modifications have disappeared from the new MACs in the control, whereas in PtGTSF1 KD, they are still detectable even at the F1 stage (Figure 5A, B). This phenotype could be the result of increased methylations by the PRC2 complex due to undegraded scnRNAs. Moreover, the duration of both scnRNAs/Ptiwi09 and PRC2 in the new MACs were extended by PtGTSF1 KD (Figure 4A-D). We further tested this hypothesis by comparing H3K9me3 and H3K27me3 levels in new MACs of control and PtGTSF1 KD cells. However, the amount of H3K9me3 and H3K27me3 gradually decreases throughout new MAC development, making a direct comparison between control and PtGTSF1 KD difficult. To tackle this problem, we simultaneously silenced the domesticated PiggyBac transposase PiggyMac (Pgm) to retain all DNA containing H3K9me3 and H3K27me3 in both conditions. Even when DNA elimination is blocked, the intensity of H3K9me3 and H3K27me3 in PtGTSF1/PGM KD is higher than in EV/PGM KD (Figure 5E–H), indicating that PRC2 deposits more H3K9me3 and H3K27me3 modifications in the new MACs.

As previously described, silencing of PtGTSF1 did not affect scnRNA biogenesis and the amount of MDS-scnRNAs in the control and PtGTSF1 KD should therefore be similar. Moreover, loss of scnRNAs by DCL2/3 KD (Dicer-like proteins that cleave double-strand RNA to produce scnRNAs) leads to an absence of H3K9me3 and H3K27me3 (20,23,61), suggesting that the increase of H3K9me3 and H3K27me3 may be due to undegraded MDS-scnRNAs guiding the PRC2 complex. Taken together, MDS-scnRNAs can enter new MACs and may guide the PRC2 complex to catalyze H3K9me3 and H3K27me3 in the absence of PtGtsf1.

Loss of PtGtsf1 affects the distribution of H3K9me3 and H3K27me3

The distribution of H3K9me3 and H3K27me3 in new MACs was also affected by PtGTSF1 KD (Figure 5A, B). Normally, H3K9me3 and H3K27me3 form foci in the new MACs, which become fewer and larger as development progresses, and finally disappear from the new MACs altogether (23). The formation of these foci is due to an enrichment of H3K9me3 and H3K27me3 on specific regions of the genome, such as TEs. In contrast, PtGTSF1 KD cells had a uniform distribution of H3K9me3 and H3K27me3 in the developing new MACs, showing a consistent distribution pattern with the localization of the PRC2 complex (Figure 4C). This suggests that they are no longer confined to specific regions and may instead be present throughout the entire genome. Since PRC2 can still catalyze the formation of H3K9me3 and H3K27me3, this dispersed distribution is not due to an inability to bind to chromatin.

One possibility is that the loss of PtGtsf1 abolishes the interaction between PRC2 and Ptiwi09, resulting in an unspecific activity of PRC2. To test this, we performed immunoprecipitation and mass spectrometry of PtCaf1 (one of the core components of PRC2) in EV control and PtGTSF1 silenced cells (Supplementary Figure S2H–J). After PtGTSF1 KD, no PtGtsf1 was detected, confirming the silencing efficiency (Supplementary Figure S2I). However, the interaction between PRC2 and Ptiwi09 was not affected by the knockdown of PtGTSF1, and the intensity of Ptiwi09 is comparable between EV and PtGTSF1 KD (Supplementary Figure S2I, J). Therefore, the uniform distribution of H3K9me3 and H3K27me3 in the absence of PtGtsf1 is not due to a loss of the interaction between Ptiwi09 and PRC2.

Since these methylation marks are normally present on TEs, we next assessed the enrichment of H3K27me3 on TEs by ChIP-qPCR. The domesticated PiggyBac transposase PiggyMac (Pgm) was co-silenced to block the elimination of TEs and prevent H3K27me3 from diminishing during new MAC development, as Pgm is responsible for the excision of TEs and IESs in P. tetraurelia (26,27). In the control (EV/PGM KD), H3K27me3 is enriched on TEs; however, when PtGTSF1 is silenced, there is no enrichment on TEs (Figure 5I, J). We also attempted H3K9me3 ChIP-qPCR with the same conditions, but our attempts were unsuccessful with the commercial antibody we used. However, a recent study on bioRxiv corroborates and extends on our findings by demonstrating that PtGtsf1 also affects the accumulation of H3K9me3 on TEs, using a home-made antibody (60). Thus, the removal of PtGtsf1 prevents the accumulation of H3K9me3 and H3K27me3 on TEs, which may explain their de-repression (Figure 2H).

Since the deficiency of PtGtsf1 increases the amount of H3K9me3 and H3K27me3 in both new and old MACs, we wonder whether the overexpression of PtGTSF1 gives the opposite result. Contrary to this hypothesis, the overexpression of PtGTSF1 did not affect the intensity of H3K9me3 or H3K27me3 (Supplementary Figure S3A–F). Their distribution was also not affected, and post-autogamous cells were able to return to vegetative growth, indicating that all processes of autogamy were successfully completed (Supplementary Figure S3G). Hence, the loss of PtGtsf1, but not its overexpression, affects the level and distribution of H3K9me3 and H3K27me3 in P. tetraurelia.

The localization of PtGtsf1 is independent of Ptiwi01/09 and PRC2

Although GTSF1 depletion did not impact the nuclear import of Ptiwi09, it led in an accumulation of Ptiwi09 in the new MACs by impairing the degradation of scnRNAs (Figure 4A). In other organisms, it has been reported that the localization of GTSF1 depends on PIWI (34,38). To determine if this is also the case for PtGtsf1 in P. tetraurelia, we assessed the localization of PtGtsf1 when the scnRNA-binding PIWI proteins Ptiwi01 and Ptiwi09 were mislocated by silencing the Dicer-like proteins Dcl2/3. In the absence of Dcl2/3, scnRNAs cannot be produced and Ptiwi09 cannot enter the old MAC; however, this did not affect the localization of PtGtsf1 (Supplementary Figure S4A, B). Since PtGtsf1 also interacts with PRC2 and affects its localization, we assessed whether loss of PRC2, achieved by EZL1 KD, affected the localization of PtGtsf1. Still, the localization of PtGtsf1 remained unchanged (Supplementary Figure S4A, C). We therefore conclude that the nuclear localization of PtGtsf1 does not depend on Ptiwi01/09 nor PRC2.

Loss of PtGtsf1 increases DNA damage in MICs

Depletion of PtGtsf1 increased the levels of H3K9me3 and H3K27me3, both of which are associated with gene expression regulation. We therefore assessed whether this increase also altered gene expression by mRNA-seq. In the Early and Late timepoints, only 10 and 3 genes, respectively, were differentially expressed (Figure 6A, B). The Late++ timepoint had 984 differentially expressed genes (DEGs) with 837 (85.06%) of them upregulated, while 708 DEGs (415, 58.62% upregulated) were found in the F1 timepoint (Figure 6C, D; Supplementary Figure S5A, B).

Figure 6.

Loss of PtGtsf1 increases DNA damage in MICs. (A–D) Differentially expressed genes (DEGs) between PtGTSF1 KD and EV in Early, Late, Late++ and F1 stages. (E–G) GO annotations of DEGs. Rich factor is the ratio of DEG numbers to the total number of genes in a certain GO term. (H) Localization of Pgm-GFP in EV and PtGTSF1 KD. Arrows point to new MACs. Scale bar: 10 μm. (I) Immunofluorescence of γH2A.X in EV, PtCAF1, PtGTSF1, PGM, PtCAF1/PGM and PtGTSF1/PGM KD. Arrowheads indicate MICs. Numbers at bottom represent the percentage of MICs with γH2A.X signal.

DEGs in PtGTSF1 KD include both genes that are differentially expressed during sexual development and genes that are not (Supplementary Figure S5C–F). Notably, all genes that are known to play a role in the genome rearrangement process and are differentially expressed upon PtGTSF1 KD are upregulated (Supplementary Figure S5G). Furthermore, neither PTIWI09 nor EZL1 are upregulated in the PtGTSF1 KD; hence, the stronger signals of Ptiwi09 and Ezl1 in Late++ and F1 stages of PtGTSF1 KD cannot be attributed to increased mRNA levels (Supplementary Figure S5G). We further predicted the functions of the DEGs in Late++ and F1 timepoints by GO analysis (Figure 6E–G). Many upregulated genes are related to DNA damage and repair, such as DNA repair, DNA recombination, mismatch repair, mismatched DNA binding and damaged DNA binding, suggesting that during these stages, there may be more DNA damage or slower DNA repair when PtGTSF1 is silenced.

Since DNA elimination by default massively induces DNA double-strand breaks (DSBs), we first investigated the localization of the excision complex using a GFP-fused Pgm. In the wild-type situation, Pgm disappears after the Late++ stage, when DNA elimination is completed (Figure 6H) (26). Interestingly, Pgm was still present in some cells even in the F1 stage in the PtGTSF1 KD. We next investigated whether blocking DNA elimination by PGM KD affects DNA damage levels during autogamy by immunofluorescence of phosphorylated γ-H2A.X (Ser139) (Figure 6I). Silencing PGM effectively blocks DNA elimination but did not lead to lower DNA damage levels compared to the EV control, suggesting that DSBs caused by DNA elimination are quickly repaired (Figure 6H). This is likely due to efficient coupling of DNA elimination and repair, as previously reported (31).

In contrast, PtGTSF1 KD cells have significantly higher levels of phospho-γ-H2A.X (Ser139) signal in the MICs than the EV and PGM KD (Figure 6I). This suggests that there are more DSBs in the absence of PtGtsf1, which may be due to an increase in the number of DSBs, changes to their positions in the genome or slower DSB repair. Interestingly, even higher DNA damage levels were detected in PtCAF1 KD (Figure 6I), which abolishes the PRC2 complex as well as H3K9me3 and H3K27me3 marks on TEs (24). Because both PtGtsf1 and PtCaf1 has a dual effect on DNA elimination, first ensuring transcriptional repression of TEs followed by DNA elimination, we next investigated the effect of double silencing with PGM KD to block DNA elimination. In both cases, the DNA damage levels in the MICs were much higher than in the EV or PGM KD alone, suggesting that the increase in DNA damage is not due to PGM-mediated DNA elimination (Figure 6I).

Loss of PtGtsf1 or PRC2 both resulted in higher levels of phospho-γ-H2A.X (Ser139) in the MICs as well as de-repression of TEs (Figure 2H) (23,25). Since the activity of TEs can induce DNA damage, one possible explanation of the higher DNA damage in MICs is the de-repression of TEs. However, PtGtsf1 is not observed in the MICs (Figure 1G), and this effect therefore seems to be indirect. In conclusion, loss PtGtsf1 or the PRC2 complex increases DNA damage in MICs during sexual development, but the underlying mechanism remains unknown and requires further investigation.

Discussion

Findings in several organisms have established GTSF1 as an essential component of the piRNA pathway in animals. Here, we expanded on these findings by revealing that the PIWI–GTSF1 interaction also exists in the unicellular eukaryote P. tetraurelia, with a role in TE silencing. In P. tetraurelia, PtGtsf1 is essential for sexual development and required for the degradation of small RNAs recognizing the organism's own genomic sequences, as a way to distinguish MDS- from IES- and TE-DNA.

PtGtsf1 is required for the elimination of Ptiwi01/09 after scnRNA-target RNA pairing

Previous studies have demonstrated that the PRC2 complex requires scnRNAs to set H3K9me3 and H3K27me3 on TEs, which aids in their recognition (23,25). In the wild-type situation, only scnRNAs corresponding to TE and IES sequences enter the new MACs to guide PRC2. In the absence of PtGtsf1, the degradation of MDS-scnRNAs is disrupted, likely blocked after pairing with their targets (Figure 3). We also noted that PRC2 accumulates in the old MACs and localizes to the new MACs with a diffused distribution pattern (Figure 4). Consequently, the level of H3K9me3 and H3K27me3 increase in both old and new MACs, and their distribution is dispersed mimicking the localization of PRC2 (Figure 5). However, ChIP-qPCR indicates that H3K27me3 is not enriched on TEs (Figure 5). Moreover, a recent paper on the same topic obtained results consistent with ours and found that the enrichment of H3K27me3 and H3K9me3 on TEs disappear in the absence of PtGtsf1 (60). Combining all of these results together, we speculate that the distribution changes of PRC2 and methylation marks in the absence of PtGtsf1 may be due to the undegraded MDS-scnRNAs. Since the PRC2 complex requires scnRNAs to deposit these marks, pairing between scnRNAs and the nascent transcripts of the old MAC appear to still take place in the absence of PtGtsf1, which places PtGtsf1 downstream of scnRNA-target RNA pairing.

Interestingly, this appears to be a PtGtsf1-specific phenotype, as this does not occur in known cases of scnRNA-degradation defects. In our previous work, we constructed a ‘Late-PRC2’ strain of P. tetraurelia whose PRC2 complex only locates and acts in the new MACs, unlike the wild type PRC2 complex that is active in both the old and new MACs (24). By removing the complex from the old MAC, we were able to assess the role of PRC2 in the old MAC specifically. Using this method, we found that loss of PRC2 in the old MAC impaired the degradation of MDS-scnRNAs, similar to what we found for PtGTSF1 KD in this study. This was also accompanied by a diffused distribution of PRC2 in the new MACs, presumably due to undegraded MDS-scnRNAs; however, and in contrast to the case in PtGTSF1 KD, H3K9me3 and H3K27me3 still formed foci as the control. Therefore, the scanning process appears to be blocked at different stages in the Late-PRC2 strain and the PtGTSF1 KD: in the Late-PRC2 strain, pairing still takes place and the MDS-scnRNAs are marked but they are not degraded, whereas in the PtGTSF1 KD, pairing still takes place but the MDS-scnRNAs can’t be marked or degraded.

These findings suggest that the characteristics of scnRNA–Ptiwi09 complexes before and after pairing differ. Thus, we speculate that scnRNA–Ptiwi09 complexes are marked after pairing with its targets, and that this process depends on PtGtsf1. This mark, on scnRNAs or Ptiwi09, would prevent MDS-scnRNAs from pairing with its targets in the new MACs and result in the degradation of the scnRNA–Ptiwi09 complexes.

The elimination of successfully paired scnRNAs is of outmost importance as this allows the organism to distinguish MDS from IES- and TE-DNA. In the absence of such selection, essential DNA sequences might be deleted; alternatively, the retention of IESs would disrupt gene expression in the new MACs. Hence, the organism requires an efficient degradation system that can distinguish paired and unpaired scnRNAs. While the details of this process are still unknown, one can imagine that this may occur either at the sRNA level or at the protein level. Interestingly, previous studies found that Argonaute carrying an extensively paired miRNA triggers its degradation through ubiquitination in a process known as target-directed miRNA degradation (62,63). Considering that Ptiwi09-bound scnRNAs have a perfect match with its targets, we envision that a similar mechanism may be at play in P. tetraurelia, in which Ptiwi09 proteins carrying perfectly matched scnRNAs undergo a conformational change that results in their degradation, in a process dependent on PtGtsf1. In support of this hypothesis, a recent study on bioRxiv reported that the lack of PtGtsf1 decreased the level of ubiquitination during autogamy, which may be linked to the degradation of Ptiwi09 and MDS-scnRNAs (60).

Our study provided insight into the genome scanning process, summarized in the following model (Figure 7). At the beginning of autogamy, MICs are transcribed to generate scnRNAs corresponding to MDSs, IESs and TEs, all of which bind with Ptiwi01/09 and enter the old MAC to identify MAC-matching MDS-scnRNAs by pairing with transcripts originating from the old MAC genome. After pairing, MDS-scnRNAs guide PRC2 to form H3K9me3 and H3K27me3 in the old MAC, after which MDS-scnRNAs are degraded in a PtGtsf1-dependent manner. The unpaired IES- and TE-scnRNAs are transferred into the new MACs to guide the elimination of IESs and TEs. PtGtsf1 also enters the new MACs during early new MAC development; however, its function there remains unknown.

Figure 7.

Model depicting the role of PtGtsf1 in the genome rearrangement process of Paramecium tetraurelia. During autogamy, micronuclei (MICs) are bidirectionally transcribed to produce long ncRNAs that are processed into scnRNAs, which then bind with Ptiwi01/09 and enter the old macronucleus (MAC). In the old MAC, scnRNAs pair with nascent transcripts generated from the old MAC genome. Paired scnRNAs (MDS-scnRNAs) guide the PRC2 complex to set H3K9me3 and H3K27me3 and are then marked in a PtGtsf1-dependent manner, resulting in their degradation. Unpaired scnRNAs enter the new MACs to identify TEs and IESs. Targeting by scnRNAs guide the PRC2 complex to set H3K9me3 and H3K27me3 on TEs, resulting in heterochromatin formation and recruitment of PiggyMAC (Pgm) and related proteins. Pgm then cleaves and eliminates the TEs. When PtGtsf1 is absent, MDS-scnRNAs cannot be degraded and aberrantly enter the new MACs, resulting in an abnormal distribution of H3K9me3 and H3K27me3, thereby affecting the recognition of TEs. Additional mechanisms yet to be discovered likely help recognize TEs. Figure created with BioRender.com.

The possibility of protective marks in Paramecium tetraurelia

The enrichment of H3K9me3 and H3K27me3 on TE sequences is essential for their repression and elimination (25). These modifications facilitate the heterochromatinization of TEs regions, which are subsequently recognized by Pgm and related proteins. Pgm is then believed to cleave the boundaries of heterochromatin and euchromatin to eliminate the TEs. The loss of PtGtsf1 leads to the aberrant entry of MDS-scnRNAs into the new MACs, which may cause the increase of H3K9me3 and H3K27me3 by guiding the PRC2 complex. Despite the presence of these methylation marks, they are no longer accumulated on TEs, and we suspect that these modifications may cover a large region of the genome due to targeting by MDS-scnRNAs. It has been suggested that the excision complex may require open chromatin at the boundaries of sequences destined for excision (64). The involvement of both nucleosome depletion and heterochromatic marks for their removal may suggest a mechanism in which the excision complex uses the boundaries of open and closed chromatin to identify such sequences. If this is the case, a dispersed distribution may pose a challenge for the excision complex to correctly recognize and eliminate these sequences. Additionally, since most IESs and TEs are still eliminated from the new MACs, there may also be additional mechanisms to recognize MDSs, TEs and IESs. In the related ciliate Tetrahymena thermophila, the boundaries of MDSs are protected to avoid excessive elimination (65). Though this may differ from the case in P. tetraurelia, it is possible that the MDSs might also be protected in this organism. This, in turn, may aid in the identification of TEs and IESs. However, further work is required to fully understand the mechanism by which the excision complex recognizes sequences for elimination in Paramecium.

The role of GTSF1 in regulating the activity of nuclear PIWI proteins

It is important to note that the role of GTSF1 has only been elucidated for catalytically active PIWI proteins, in which it appears to enhance the intrinsically weak RNA cleavage activities of PIWI (40). In mouse, silk moth and sponges, GTSF1 interacts with PIWI and recognizes the pre-catalytic piRISC state, facilitating a conformational change in PIWI and stabilizing its catalytically active conformation (40). Although Ptiwi09 is also a catalytically active PIWI protein, its role in the nucleus suggests that its main function is not to directly cleave its targets, but rather that it acts similarly to the catalytically inactive Drosophila Piwi, which guides downstream factors to deposit histone methylation marks (66,67). Of note, Drosophila Piwi also requires GTSF1 for its activity (34,36,39). We can envision a model in which the binding of nuclear PIWI proteins to its targets induces a conformational change in PIWI, and that the interaction with GTSF1 can only happen after this change.

Our results are consistent with a similar role of PtGtsf1 as in metazoans, in which its canonical function is TE silencing through its interaction with, and regulation of, PIWI proteins. Although it is also involved in TE silencing, it is important to note that the phenotype of PtGTSF1 KD also has important differences to its metazoan counterparts. For instance, the depletion of PtGtsf1 did not affect the catalytic activities of Ptiwi09 or the biogenesis of sRNAs, but disturbed the degradation of scnRNAs after pairing with the genome. Moreover, PtGtsf1 regulates the levels of H3K9me3 and H3K27me3 in P. tetraurelia. Interestingly, while the intensity of H3K9me3 and H3K27me3 increased in the absence of PtGtsf1, these aberrant methylation marks do not concentrate on TEs; counterintuitively, the enrichment on TEs decreased, resulting in their de-repression. A similar phenotype was also reported in D. melanogaster, in which the loss of GTSF1 also decreases the levels of H3K9me3 on TEs (36). Our characterization of PtGtsf1 in P. tetraurelia demonstrates that transposon repression is a common function of GTSF1 proteins. Protists are the ancestors of multicellular organisms; hence, the existence of GTSF1 in protists suggests that this protein appeared early in eukaryotic evolution, with the function of TE repression.

Data availability

The mRNA-seq, sRNA-seq and genomic DNA sequencing data were submitted to NCBI database under the BioProject PRJNA1020915. The mass spectrometry data was submitted to PRIDE Proteomics Identifications Database with project IDs of PXD049029 and PXD054662.

Supplementary data

Supplementary Data are available at NAR Online.

Acknowledgements

We thank Prof. Weibo Song from Ocean University of China for his suggestion and help in drafting the manuscript. We acknowledge the computing resources provided on IEMB-1 computing cluster in the Institute of Evolution and Marine Biodiversity, Ocean University of China, and SDU-LMPBE computing cluster in the Laboratory of Marine Protozoan Biodiversity and Evolution, Shandong University, respectively.

Author contributions: C.W.: Conceptualization; data curation; formal analysis; investigation; visualization; writing-original draft; writing – review and editing; funding acquisition. L.L.: Formal analysis; visualization; writing – original draft. T.S.: Conceptualization; writing-review and editing. H.Z.: Investigation. Z.W.: Investigation; F.G.: Supervision; writing – original draft; writing – review and editing; funding acquisition.

Funding

Science & Technology Innovation Project of Laoshan Laboratory [No. LSKJ202203202]; National Natural Science Foundation of China [32100382, 32470512, 31961123002]; Natural Science Foundation of Shandong Province [ZR2021QC104]; Young Elite Scientists Sponsorship Program by CAST [2022QNRC001], Young Taishan Scholar Program of Shandong Province, Fundamental Research Funds for the Central Universities [202141004]; World Premier International Research Center Initiative (WPI), MEXT, Japan.

Conflict of interest statement. None declared.