# What's, hows and whys of programmed DNA elimination in Tetrahymena

Tomoko Noto, Kazufumi Mochizuki

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Tomoko Noto, Kazufumi Mochizuki. What's, hows and whys of programmed DNA elimination in Tetrahymena. Open Biology, 2017, 7 (10), pp.170172. 10.1098/rsob.170172. hal-02387399

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**Cite this article:** Noto T, Mochizuki K. 2017
Whats, hows and whys of programmed DNA
elimination in *Tetrahymena*. *Open Biol.* **7**: 170172.

http://dx.doi.org/10.1098/rsob.170172

Received: 13 July 2017

Accepted: 12 September 2017

**Subject Area:**

genetics/genomics/molecular biology/
biochemistry/cellular biology

**Keywords:**

epigenetics, RNAi, DNA elimination,
transposon, *Tetrahymena*

**Author for correspondence:**

Kazufumi Mochizuki

e-mail: kazufumi.mochizuki@igh.cnrs.fr

Whats, hows and whys of programmed
DNA elimination in *Tetrahymena*

Tomoko Noto and Kazufumi Mochizuki

Institute of Human Genetics, UMR 9002, CNRS and University of Montpellier, Montpellier, France

ID KM, 0000-0001-7987-9852

Programmed genome rearrangements in ciliates provide fascinating examples of flexible epigenetic genome regulations and important insights into the interaction between transposable elements (TEs) and host genomes. DNA elimination in *Tetrahymena thermophila* removes approximately 12 000 internal eliminated sequences (IESs), which correspond to one-third of the genome, when the somatic macronucleus (MAC) differentiates from the germline micro-nucleus (MIC). More than half of the IESs, many of which show high similarity to TEs, are targeted for elimination in *cis* by the small RNA-mediated genome comparison of the MIC to the MAC. Other IESs are targeted for elimination in *trans* by the same small RNAs through repetitive sequences. Furthermore, the small RNA–heterochromatin feedback loop ensures robust DNA elimination. Here, we review an updated picture of the DNA elimination mechanism, discuss the physiological and evolutionary roles of DNA elimination, and outline the key questions that remain unanswered.

**1. Introduction**

Ciliated protozoa undergo extensive programmed genome rearrangements when the germline micronucleus (MIC) produces the new macronucleus (MAC) during sexual reproduction. In this process, many transposon-related sequences are removed from the MAC [1,2]. RNAi-related mechanisms play important roles in programmed genome rearrangements in at least three different ciliates, including *Oxytricha trifallax* [3,4], *Paramecium tetraurelia* [5–7] and *Tetrahymena thermophila* [8–11]. There are substantial (and very interesting) differences in the mechanisms that regulate programmed genome rearrangement in ciliates. This review aims to overview the mechanisms and the roles of programmed genome rearrangement in *Tetrahymena thermophila* (hereafter referred to as *Tetrahymena*). We refer the reader to previous reviews [12–16] for programmed genome rearrangements in other ciliates.

Although it has long been known that programmed genome rearrangements occur genome-wide and eliminate many sequences related to transposable elements (TEs), a global landscape concerning ‘what’ is eliminated—their numbers, chromosomal distributions and relationship with TEs—became available only after the recent establishment of the assemblies of nearly full-length MIC chromosomes [1]. In §2, we overview the programmed genome rearrangements of *Tetrahymena* in the context of the MIC genome.

Current knowledge of the eliminated sequences in combination with the accumulated genetic and biochemical data on the molecular machineries that regulate small RNA-mediated DNA elimination now provide a clear picture of ‘how’ eliminated sequences are recognized. In §3, we discuss how small RNAs target particular sequences for DNA elimination by a complex genome-wide network.

Whenever we present at conferences, the most frequently asked question is, ‘Why do ciliates perform DNA elimination?’ Dr Martin Gorovsky, our former supervisor, taught us not to ask *why* things happen but *how* they occur. We have faithfully followed this advice for over a decade. However, with a recently
<figure><img src="image_3.png" /></figure>

**Figure 1.** Life cycle and programmed genome rearrangements of *Tetrahymena*. (a) Life cycle of *Tetrahymena*. A single *Tetrahymena* cell contains a macronucleus (MAC) and a micronucleus (MIC). When sufficient nutrients are available, *Tetrahymena* grow by binary fission, and the MAC and the MIC divide independently (vegetative growth). After prolonged starvation, cells undergo conjugation, the sexual reproduction process: (i) the MIC undergoes meiosis to generate four haploid nuclei, three of which are discarded, and one selected haploid nucleus proceeds with post-meiotic division to form two haploid pronuclei; (ii) one of the two pronuclei is exchanged between the mating partner, and the stationary and the exchanged pronuclei fuse to form a diploid zygotic nucleus; and (iii) the zygotic nucleus divides twice to form two new MACs and two new MICs, while the parental MAC is degraded. Readers interested in the details of the *Tetrahymena* life cycle should refer to [22]. (b) Programmed genome rearrangements. In the newly formed MAC, chromosome breaks occur at CBSs and telomeres are formed *de novo*. In addition, internal eliminated sequences (IESs) are removed by DNA elimination. Each MAC chromosome is endoreplicated to approximately 45–70 copies (see text for discrepancy among the previous estimates of the MAC chromosome copy number).

improved understanding of the DNA elimination pathway, we think that it is time to consider the 'why' question because we can now form testable hypotheses (sorry, Marty). In §4, we discuss why *Tetrahymena* perform DNA elimination from the point of view of genome integrity and dynamics.

## 2. The genome of *Tetrahymena*: what is eliminated from the MAC?

### 2.1. The life of *Tetrahymena*

*Tetrahymena* is a free-living freshwater ciliate that is one of the most commonly studied protozoa in laboratories. Like most other ciliates, *Tetrahymena* display nuclear dimorphism [17] through the presence of a diploid germline MIC and a poly-ploid somatic MAC. Gene expression occurs in the MAC, whereas the MIC is transcriptionally inert (although there is an exception during early sexual reproduction; see below). Because the MAC in G1 phase was reported to contain approximately 23 times more DNA than the G1 MIC [18] and the MIC genome is approximately 1.5 times larger than the MAC genome (see below), the copy number of the MAC chromosomes in G1 phase is estimated to be approximately 70 [19]. However, this number does not agree well with the estimates (approx. 45 C) from the kinetics of phenotypic assortment [20,21], the process in which chromosome copies distribute randomly during MAC vegetative division. Recent sequencing and/or microscopic technologies would aid to reconcile this discrepancy.

Figure 1a summarizes the life cycle of *Tetrahymena*. During vegetative growth, both the MAC and the MIC divide and segregate to the daughter cells. During conjugation, the sexual reproduction process of *Tetrahymena*, the MIC undergoes meiosis, which is followed by fertilization to form the zygotic nucleus. The zygotic nucleus further divides to make the new MAC and the new MIC. The parental MAC is destroyed by autophagic degradation at the end of sexual reproduction [23]. The new

MAC genome is endoreplicated to 4 C immediately after its development, and then programmed genome rearrangements occur concomitant with the second round of endoreplication (4 C to 8 C) [24]. However, endoreplication is not necessary for genome rearrangement [25]. Further endoreplication of the MAC genome occurs later during vegetative growth.

### 2.2. The eliminated sequences

The latest studies on genome sequencing indicate that the sizes of the MIC and MAC genomes are 157 Mb and 103 Mb, respectively [1,26–28]. This difference in size is caused by the two types of programmed genome rearrangements that occur during the development of the MAC.

The first type of programmed genome rearrangement is chromosome breakage at the chromosome breakage sequences (CBSs), accompanied by short DNA trimming and *de novo* telomere formation (figure 1b). Because CBSs share a conserved 15 bp element [29], a yet unidentified endonuclease that specifically recognizes CBS probably catalyses the chromosome breakage. The telomere end binding protein homologue Pot2p localizes to CBSs exclusively at the time of chromosome breakage and is suggested to be involved in *de novo* telomere formation [30]. Chromosome breakages split 5 MIC chromosomes into approximately 230 MAC chromosomes, while approximately 50 of them, called the non-maintained chromosomes (NMCs, also called the eliminated mini-chromosomes), are either immediately degraded before telomere addition or lost within approximately 20 fissions during vegetative growth after telomere addition. The mechanism for the disappearance of NMCs is not known. The identified NMCs are generally short, ranging from 30 to 80 kbp, compared with the other MAC chromosomes (40 kb to 1.4 Mb) [1,26,31].

The second type of programmed genome rearrangement is the elimination of internal DNA segments, called internal eliminated sequence (IES), followed by the ligation of flanking MAC-destined sequences (figure 1b). Hereafter, we refer to this genome rearrangement event as DNA elimination. In
the articles published before 2012, the number of IESs in the *Tetrahymena* MIC was estimated to be approximately 6000. This was based on estimations from a limited sampling of the MIC genome [32] or low coverage sequencing of the MIC genome [33]. With the near-complete MIC genome sequence, the predicted number of IESs has been substantially increased and the latest estimation is approximately 12 000 [1], which corresponds to approximately 46 Mb. The predicted IESs range from 136 bp to 43.4 kb with the mean and the median length of 3.8 and 2.8 kb, respectively.

The excisions of nearly all IESs are believed to be catalysed by the domesticated piggyBac transposase Tpb2p. However, due to technical difficulties, it has not been possible to completely remove Tpb2p activity from the cell [34,35], and thus the absolute requirement of Tpb2p for individual DNA elimination remains elusive. Tpb2p-dependent IESs are imprecisely excised from the genome. End-point heterogeneities of boundaries, mostly within a few base pairs, were observed between siblings [1]. Accordingly, Tpb2p-dependent IESs are found within intergenic regions and introns. In contrast to the majority of IESs, the excision of 12 (and possibly slightly more) IESs is dependent on the other domesticated piggyBac transposases Tpb1p and Tpb6p [36,37]. All known Tpb1/6p-dependent IESs is within exons of protein-encoding genes and precisely excised [1,36,37]. These exceptional 12 IESs share a conserved 12 bp terminal repeat and produce few scnRNAs, if any [36,37]. Therefore, in contrast to the Tpb2p-dependent IESs that are targeted for DNA elimination by an RNAi-related mechanism (see below), these 12 IESs are probably recognized and targeted for DNA elimination directly by Tpb1/6p, and are excluded from IESs in the following discussion.

The majority of the genome that is eliminated from the developing MAC (157 Mb [MIC] – 103 Mb [MAC] = 54 Mb) is occupied by the predicted IESs (46 Mb). We speculate that the most of the remaining 8 Mb also consists of IESs, which were not predicted due to the insufficient quality of the current MIC genome assembly. The contribution of NMCs to genome down-sizing is limited because the sum of identified NMCs totals 0.56 Mb [31], although unidentified large NMCs may exist.

## 2.3. Kinship between transposable elements and IESs

The currently assembled MIC genome contains at least approximately 19 Mb of transposable element (TE)-related sequences [1]. Among the classifiable TE sequences, the majority (91%) correspond to DNA transposons, including Tc1/Mariner, Helitron and Maverick/Tlr families. A small portion (9%) of TEs consists of retrotransposons, all of which are related to non-LTR elements [1]. Approximately 18 Mb of these TE-related sequences are removed from the MAC genome. There is a retention bias toward the terminal regions of TEs, suggesting that occasionally IES elimination incompletely removes TEs, leaving the terminal regions of the TEs in the MAC [1]. Therefore, some (or most) of the approximately 1 Mb TE-related sequences that remain in the MAC are likely to be non-functional fragments of TEs.

Some IESs have intact open-reading frames and terminal repeats of TEs and are found in multiple copies in the MIC genome [1]. Therefore, some IESs are likely to be the result of recent TE activity. Consistently, IES positions are highly variable between closely related *Tetrahymena* species [38]. Many IESs consist of modules of sequences homologous to different TE families, which is probably a consequence of frequent TE

insertions into an IES and its following degeneration [1,39]. Some or many regions of IESs that do not show detectable homology with TEs might also be remnants of TEs.

# 3. The molecular mechanism of DNA elimination: how do small RNAs regulate DNA elimination?

It has been known that DNA elimination is regulated by the small RNA-mediated comparison of the MIC to the MAC genomes [8]. Although this view is still valid, recent studies have indicated that additional layers of mechanisms act in addition to the genome comparison. In §3.1, we discuss how a small RNA-based system harnesses nuclear dimorphism and the modular organization of IESs to robustly but flexibly regulate DNA elimination. Next, the molecular players behind this small RNA-mediated regulation are discussed in §3.2. Finally, in §3.3 we outline the unanswered questions regarding the recently discovered small RNA–heterochromatin positive feedback loop in DNA elimination.

## 3.1. Systems point of view of small RNA-directed DNA elimination

Here we present a small RNA-centric model of DNA elimination (figure 2a) that is based on genetic and biochemical studies of RNAi machineries and high-throughput sequencing of small RNAs [8–11,40–43].

In the early stages of conjugation in the MIC, approximately 3 h post-mixing (hpm), small RNAs of approximately 29 nt, called Early-scnRNAs, are produced from certain (approx. 60%) IESs (called Type-A IESs) and their surrounding regions, whereas the remaining (approx. 40%) IESs (called Type-B IESs) do not produce Early-scnRNAs [43]. Type-A IESs are primarily located at the middle (peri-centromeric) and the end (telomeric) regions of the MIC chromosomes, which are called A-regions, whereas Type-B IESs are primarily located at the chromosomal arms, called B-regions [1] (figure 2b). Therefore, contrary to previous 'scnRNA models' [8,44], Early-scnRNAs are produced heterogeneously from the MIC genome. There are two biases in the pattern of Early-scnRNA production: the global bias in which Early-scnRNAs are preferentially produced from the A-regions, and the local bias in which, within the A-regions, Early-scnRNAs are preferentially produced from IESs [43].

Early-scnRNAs then localize to the parental MAC at the mid stages of conjugation (approx. 3–7 hpm), where any Early-scnRNAs that are complementary to the MAC genome (i.e. non-IES sequences) are degraded by the process called scnRNA selection [40,42,43], as previously proposed [8,44]. As a result, only Early-scnRNAs that are complementary to the MIC-limited sequences are retained.

Next, the selected Early-scnRNAs translocate to the newly formed MAC (approx. 7 hpm), where they interact with Type-A IESs to induce heterochromatin formation [8,45,46]. Type-B IESs contain repetitive sequences, named A-repeats, which are complementary to Type-A IESs [43]. Thus, Early-scnRNAs also interact with Type-B IESs in *trans* via A-repeats to induce heterochromatin formation. Moreover, Early-scnRNAs induce the production of another group of approximately 29-nt small RNAs, called Late-scnRNAs, in *cis* at both Type-A and Type-B
<figure><img src="image_4.png" /><figcaption>**Figure 2.** Regulation of DNA elimination by small RNAs. (a) A model for small RNA-directed DNA elimination. Early-scRNAs are expressed from Type-A IESs and their surrounding regions (A-region) in the MIC and then move into the parental MAC, where Early-scRNAs derived from MAC-destined sequences are degraded (left). When the new MAC is formed in the progeny (right), Early-scRNAs move from the parental to the new MAC. There, Early-scRNAs recognize not only Type-A IESs but also Type-B IESs in *trans* through common repetitive sequences. This recognition triggers Late-scRNA production in *cis*. Early- and Late-scRNAs cooperatively induce DNA elimination. (b) Localizations of Type-A and Type-B IESs, transposons (TEs), and coding sequences and productions of Early- and Late-scRNAs in the MIC chromosome 5 are shown in histograms with 50 kb bins. Tlr1- and REP-elements were used as TEs. Twi1p-bound 26–32-nt RNAs at 3 hpm and Twi11p-bound 26–32-nt RNAs at 10.5 hpm were analysed as Early- and Late-scRNAs, respectively.</figcaption></figure>

IESs [43]. Late-scRNAs further trigger heterochromatin formation in *trans*, forming an RNAi-heterochromatin positive feedback loop [43]. The heterochromatinized IESs are eventually excised by Tpb2p [34,35]. When the accumulation of

Late-scRNAs was inhibited by genetically removing zygotically expressed Twi1p and Twi11p (see below), DNA eliminations of Type-A IESs were only weakly inhibited and those of Type-B IESs were more strongly, but not completely,
inhibited [43]. These indicate that Early-scnRNAs can directly
induce DNA elimination but Late-scnRNAs are necessary to
ensure complete removal of all IES copies from the MAC.

The RNAi-heterochromatin positive feedback loop, which is composed of *trans*-recognition and *cis*-spreading mechanisms, probably provides robustness to the system. Our simulations indicated that the feedback loop mechanism can secure the elimination of most IESs, even when 99% of Type-A IESs fails to express Early-scnRNAs [43]. Most of the potentially active TEs are found in Type-A IESs [43]. Active TE-containing (i.e. young) IESs may be constrained by natural selection to localize at the gene-poor A-regions (figure 2b), where the production of Early-scnRNAs ensures that their DNA is eliminated from the new MAC. As TE sequences degenerate (i.e. become older), IESs may be tolerated as residents of the gene-rich chromosomal arm regions and then transition into Type-B IESs.

## 3.2. Mechanistic point of view of small RNA-directed DNA elimination

The molecular machineries of small RNA-directed DNA elimination are illustrated in figure 3. The MIC is transcriptionally inert during all life stages, except in meiotic prophase [47,48], when the MIC genome is transcribed bi-directionally (figure 3a) [40,49]. The resulting double-stranded RNAs are processed to Early-scnRNAs by the Dicer protein Dcl1p in the MIC (figure 3b) [9,10]. The global and local biases in the pattern of Early-scnRNA production described above are determined at the level of transcription [42]. Because Rpb3p, a RNA polymerase II subunit, localizes to the MIC during meiotic prophase, while Rcp5p, a shared subunit of RNA polymerase I and III, does not [50], the MIC is probably transcribed by RNA polymerase II.

Early-scnRNAs are then exported by a yet unidentified mechanism to the cytoplasm (figure 3c) and loaded into the Argonaute protein Twi1p (figure 3d) [11,41]. This loading process requires Hsp90 and its co-chaperone [51]. Twi1p has endonuclease (Slicer) activity that cuts and removes one of the two strands of the loaded Early-scnRNAs (figure 3e) [41]. Then, the single-stranded Early-scnRNA-Twi1p complex interacts with Giw1p and is imported into the parental MAC (figure 3f) [41]. The MAC and the MIC have distinct nuclear pore proteins and use different importins/exportins [52–54]. Exactly how Giw1p links the Twi1p-Early-scnRNA complex to the MAC-specific nuclear import pathway is unclear.

In the parental MAC, Early-scnRNAs that are complementary to the MAC genome are degraded by snRNA selection (figure 3h) [11,40,42]. Although Early-scnRNAs are stabilized by Hen1p-catalysed 2'-O-methylation at their 3'-ends (figure 3g) [55], this modification does not appear to be involved in snRNA selection because Early-scnRNAs are complementary to the MAC genome were degraded faster than those to IESs even in the absence of Hen1p (T.N. & K.M. 2017, unpublished data). The putative RNA helicase Ema1p triggers Early-scnRNA degradation, probably by promoting the interaction between Early-scnRNA and chromatin through nascent transcripts [40]. The requirement of complementary transcripts for snRNA selection was directly demonstrated in *Paramecium* by RNAi-knockdown [7]. CnjBp and Wag1p also have a redundant role in snRNA selection [56]. They both contain GW repeats, also called the AGO hook, which are characteristics of several Argonaute-interacting proteins [57]. Although snRNA

selection is one of the key processes in the regulation of DNA elimination, its molecular mechanism remains elusive. Ema1p, CnjBp and/or Wag1p possibly link Early-scnRNAs to RNA degradation machinery. Alternatively, there may not be specific RNA degradation machinery for snRNA selection, but it is possible that the Twi1p-binding proteins promote dissociation of Early-scnRNAs from Twi1p, which would allow nonspecific RNase(s) access to Early-scnRNAs.

In parallel to snRNA selection, the MIC undergoes meiosis, zygote formation, and formation of the new MAC and MIC. When the new MAC develops, those Twi1p-Early-scnRNA complexes that are not degraded by snRNA selection move from the parental MAC to the new MAC (figure 3i) [8,41]. In the new MAC, Early-scnRNA-Twi1p complexes interact with chromatin through nascent transcripts in a Ema1p-dependent manner (figure 3j), and, by a yet unknown mechanism, recruit the histone methyltransferase Ez11p (figure 3k). As a result, methylated histones H3 at lysine 9 and lysine 27 (H3K9/K27me3) as well as the HP1-like protein Pdd1p, which binds to these methylated histones, accumulate on Type-A IESs and A-repeats on Type-B IESs (figure 3l) [40,43,45,46].

Next, Late-scnRNAs are produced from IESs in the new MAC in an Ez11p- and Pdd1p-dependent manner (figure 3m) and are loaded into the two Argonaute proteins Twi1p and Twi11p (figure 3n) [43]. Late-snRNA and heterochromatin form a positive feedback loop to facilitate Late-snRNA biogenesis and heterochromatin formation on IESs (figure 3o). Because heterochromatin eventually induces DNA excision [34], this activity must be precisely stopped at the borders of the IESs (figure 3p). Several items remain unclear, including how heterochromatin components induce Late-snRNA production, how the borders of IESs are determined, and the functional distinction, if any, between the two Argonaute proteins. These issues are discussed in §3.3.

Heterochromatinized IESs are assembled into large nuclear foci called heterochromatin bodies (HBs) before or during DNA elimination. Pdd1p is hyper-phosphorylated upon heterochromatin establishment [58]. The phosphorylation and subsequent dephosphorylation of Pdd1p are required for the assembly of HBs [59,60]. The dephosphorylation of Pdd1p also promotes RNA-Pdd1p interaction by reducing the net negative charge of Pdd1p [59]. RNAs may act as glue to assemble multiple heterochromatinized IESs into an HB. The RNA-binding aggregation-prone protein Jub6p is also required for HB formation, suggesting that RNA-mediated phase separation may play a role in heterochromatin aggregation [61].

The final excision step of IESs require Tpb2p (except for the 12 Tpb1/6p-dependent IESs) and non-homologous end-joining machinery for their repair steps [34,35,62]. Although Tpb2p can directly interact with H3K9me3, such interaction does not fully explain how Tpb2p executes DNA excision at the IES border, but not within the body of the IES. Tpb2p may become active only when it interacts with a specific chromatin environment at the IES border.

## 3.3. Three unknowns of the small RNA–heterochromatin positive feedback loop

### 3.3.1. What mechanism drives the small RNA–heterochromatin positive feedback loop?

How does heterochromatin induce Late-snRNA production?
Clearly, the production of Late-snRNAs is preceded by the
<figure><img src="image_5.png" /><figcaption>**Figure 3.** (Caption opposite.)</figcaption></figure>
<figure><img src="image_6.png" /><figcaption>**Figure 4.** Co-deletion (coDel). A DNA construct in which a non-IES sequence (target) is inserted into the middle of a Type-A IES (i) is introduced into the developing MAC (ii). The Type-A IES of the construct is recognized by Early-scnRNAs that are produced from the corresponding endogenous Type-A IES (iii) and trigger Late-scnRNA production from the adjacent target sequence in *cis* (iv). Late-scnRNAs then recognize the endogenous target locus in *trans* (v) and induce ectopic DNA elimination (vi). Yellow circles indicate the hypothetical IRHs.</figcaption></figure>

**Figure 3.** (Opposite.) Machinery of DNA elimination. Bi-directional transcription in the MIC by RNA polymerase II produces double-stranded RNAs (a), which are processed by the Dicer-like protein Dcl1p to Early-scnRNAs (b). Early-scnRNAs are exported to the cytoplasm (c) where they are loaded into the Argonaute protein Twi1p with the aid of Hsp90 and its co-chaperone Coi12p (d). The passenger strand of the Early-scnRNA duplex is cut and removed by the Slicer activity of Twi1p (e). Giw1p transports the Early-scnRNA–Twi1p complex to the parental MAC (f). The RNA methyltransferase Hen1p introduces 2'-*o*-methylation to the last nucleotides of Early-scnRNAs (g). The RNA helicase Ema1p facilitates the interaction between the Early-scnRNA–Twi1p complex and complementary nascent transcripts, which induce the degradation of Early-scnRNAs (scnRNA selection). Two GW-repeat proteins, Wag1p and CnjBp, are also necessary for SCN RNA selection (h). The remaining Early-scnRNA–Twi1p complexes are transported into the new MAC, possibly by Giw1p (i). Like in the parental MAC, Ema1p facilitates the interaction between the Early-scnRNA–Twi1p complex and nascent RNAs (j). This interaction recruits the histone methyltransferase Ezl1p, which catalyses H3K9me3 and H3K27me3 (k). The HP1-like protein Pdd1p binds to these methylated histones (l). Heterochromatin induces bi-directional transcription in *cis*, resulting in Dcl1p-dependent production of Late-scnRNAs (m). Late-scnRNAs are loaded into Twi1p and Twi11p and then are 2'-*o*-methylated by Hen1p (n). The Late-scnRNA–Twi1/11p complex recruits Ezl1p, forming an RNAi-heterochromatin positive feedback loop (o). The feedback loop is downregulated by ‘boundary-protecting factors’ Coi6p (HP1-like), Coi7p, Lia5p (domesticated piggyBac transposase) and Jmj1p (H3K27 demethylase) (p). The domesticated piggyBac transposase Tpb2p eventually excises heterochromatinized IESs (q).

transcription of their precursors. A heterochromatin component may directly promote bi-directional transcription. While transcriptional enhancement by heterochromatin may not be intuitive, heterochromatin-induced transcription has been observed in different eukaryotes [63,64]. Alternatively, promiscuous transcription may occur independently of heterochromatin and a heterochromatin component may direct the transcripts to the Late-scnRNA-producing pathway.

We hypothesize that there are at least two consecutive heterochromatin states on an IES: heterochromatin state I, which induces the production of Late-scnRNAs; and heterochromatin state II, which induces heterochromatin aggregation and subsequent DNA elimination. Over 20 heterochromatin components have been reported to be involved in DNA elimination [19,59]. Investigations of chromatin localization with high temporal resolution would provide a basis to understand how different heterochromatin states exert distinct biological effects.

The phosphorylation states of Pdd1p are correlated with the two heterochromatin states: Pdd1p is hyper-phosphorylated when Late-scnRNAs are produced and hypo-phosphorylated when DNA elimination begins [58,59]. Although the phosphorylation of Pdd1p is probably not necessary for the biogenesis of Late-scnRNAs [60], enzymes regulating Pdd1p phosphorylation might also regulate Late-scnRNA production through another heterochromatin component. Future studies should identify kinase(s) and phosphatase(s) of Pdd1p to determine if they play a role in Late-scnRNA production.

### 3.3.2. How is the positive feedback loop downregulated at the ends of IESs?

Exactly how the borders of IESs are determined is still unclear because no consensus sequence has been identified around the

IES boundaries. An implication can be obtained from the phenomenon of co-deletion (coDel), which involves targeted ectopic DNA elimination that is induced at any non-IES sequence by introducing an IES–target–IES chimeric construct into the new MAC [43,65]. The *cis*-spreading of the RNAi–heterochromatin signal and *trans*-recognition by SCN RNAs are the bases of coDel (figure 4). Although the boundaries of coDel-induced elimination sites are not as homogeneous as those of IESs, there appear to be preferred locations for their boundaries [65]. We hypothesize that there are insulators of RNAi–heterochromatin *cis*-spreading (IRHs) at the IES boundaries as well as at non-IES loci in the MIC genome (figure 4, yellow circles). When a non-IES region is targeted by coDel, two IRHs adjacent to the target may insulate the spreading of the RNAi–heterochromatin feedback loop, and thus DNA excision occurs at the two IRHs. Identification of potential IRHs by a large-scale coDel screening may allow us to extract sequence features of IRHs, which may provide insight into this long-standing mystery.

It has been reported that some IESs have *cis*-acting elements that play an important role in determining the borders of IESs [66,67]. A subset of *cis*-acting elements potentially forms a G-quadruplex structure. Additionally, the G-quadruplex binding protein Lia3p is important for properly determining the boundaries of G-quadruplex-associated IESs [66]. It is important to investigate the role of Lia3p, the G-quadruplex, and the other known *cis*-acting elements in limiting the
<figure><img src="image_7.png" /><figcaption>**Figure 5.** TE invasion and outcomes. Scenario 0: a schematic representation of DNA elimination without naive TE invasion. Scenario 1: a new insertion of a naive TE into an Early-scRNA-producing region. This results in Early-scRNA production from the TE and elimination of the TE from the new MAC. Scenario 2: a new insertion of a naive TE into a Type-B IES. This results in production of Late-scRNAs from the TE by cis-spreading and thus the elimination of the TE with the adjacent IES from the new MAC in the next generation. Scenario 3: a new insertion of a naive TE into a non-IES and non-Early-scRNA-producing region of the MIC genome. This does not immediately provoke the small RNA pathway and the TE may freely jump. The TE also retains in the new MAC (there may be a TE-silencing mechanism in vegetative cells). However, once a copy of the TE is trapped in an IES or Early-scRNA-producing region (such probability is high as IESs occupy one-third of the MIC genome), Scenario 1 or 2 occurs. Additionally, once this copy produces Early- and/or Late-scRNAs, all other copies of the TE are recognized in trans and targeted for elimination.</figcaption></figure>

surveillance of TEs. Although the effect of DNA elimination on replication is limited in *Tetrahymena* (which eliminates only 30% of the genome from the MAC), elimination is more prevalent in other ciliates, some of which remove over 90% of their genome [84]. DNA elimination may have evolved from an ancestral ciliate that encountered a heavy TE load. Another interesting question is: why do human somatic cells not undergo DNA elimination when approximately 45% of their genome consists of TEs? We speculate that TEs are required for faithful chromosome segregation and genome integrity in diploid somatic cells to avoid aneuploidy and cancer in the context of multicellularity.

DNA elimination also acts as a device for transgenerational epigenetic inheritance, as previously discussed in other ciliates [13,85]. The approximately 12 000 IESs are mostly located in intergenic regions of the gene-dense genome (approx. 27 000 predicted genes, one gene/3.7 kb of the MAC genome) [26]. Therefore, spontaneous DNA elimination ‘errors’ (IES retention or ectopic DNA elimination) could influence the expression of nearby genes in the MAC. Because DNA elimination occurs during the second round of the MAC genome endoreplication

[24], DNA elimination potentially forms eight variants per locus. The phenotypic assortment [20,21], in which amitotic chromosome segregation of the polyploid MAC randomly assort a variant, may provide increased fitness to cells that are adapting to environmental change (figure 6). Moreover, the scnRNA selection mechanism allows an advantageous DNA elimination variant to be epigenetically inherited by sexual progeny without phenotypic assortment (figure 6). Future studies should be designed to investigate the frequency of DNA elimination errors, and test if such errors and the following transgenerational epigenetic inheritance indeed aid in environmental adaptation.

**5. Concluding remarks**

TEs are considered necessary evils in genome dynamics and evolution; although they must be tightly downregulated, their activities are the drivers of genome reorganization. As discussed in this review, this balance has probably shaped small RNA-directed DNA elimination in *Tetrahymena*. Although
<figure><img src="image_8.png" /><figcaption>**Figure 6.** Potential impact of variations in DNA elimination on environmental adaptation. (Top) Eliminations of IESs (boxes) make the MAC genome streamlined. (Left) Spontaneous retention of IES-Y, which promotes the expression of Gene-Z or an ectopic DNA elimination at Gene-X make phenotypic variations. (Middle) These variations have no selective advantage in Environment A. The expression of Gene-Z (yellow cells) provides adaptive advantage in Environment B. The absence of Gene-X (purple cells) is beneficial in Environment C. (Right) The altered pattern of DNA elimination can be trans-generationally transmitted through conjugation by scnRNA selection.</figcaption></figure>

the overall pathway of DNA elimination of *Tetrahymena* has emerged, key questions regarding its molecular mechanism, physiological function and evolutionary significance remain. For example, how are IES-specific Early-scnRNAs selected? How does heterochromatin induce Late-scnRNA production? How are the borders of IESs determined? What are the roles of IESs in the MIC? To what extent do spontaneous DNA elimination errors contribute to environmental adaptation? Further genetic, biochemical and comparative genomic analyses of the DNA elimination mechanism, in the context of TE regulation

and genome evolution, should provide an exciting new entry point to answer these long-standing questions.

**Data accessibility.** This article has no additional data.

**Competing interests.** We declare we have no competing interests.

**Funding.** K.M. is supported by funds from the French National Research Agency (an Advanced Grant from the Investissements d'avenir programme Labex EpiGenMed (ANR-10-LABX-12-01) and an Accueil de Chercheurs de Haut Niveau (ACHN) grant (ANR-16-ACHN-0017) and the Sumitomo Foundation (a Grant for Basic Science Research Projects (150112)).

## References

1. Hamilton EP et al. 2016 Structure of the germline genome of *Tetrahymena thermophila* and relationship to the massively rearranged somatic genome. *Elife* 5, e19090. (doi:10.7554/ eLife.19090)

2. Arnaiz O et al. 2012 The *Paramecium* germline genome provides a niche for intragenic parasitic DNA: evolutionary dynamics of internal eliminated sequences. *PLoS Genet.* 8, e1002984. (doi:10.1371/ journal.pgen.1002984)

3. Fang W, Wang X, Bracht JR, Nowacki M, Landweber LF. 2012 Piwi-interacting RNAs protect DNA against loss during *Oxytricha* genome rearrangement. *Cell* 151, 1243–1255. (doi:10.1016/j.cell.2012.10.045)

4. Zahler AM, Neeb ZT, Lin A, Katzman S. 2012 Mating of the stichotrichous ciliate *Oxytricha trifallax* induces production of a class of 27 nt small RNAs derived from the parental macronucleus. *PLoS ONE* 7, e42371. (doi:10.1371/journal.pone.0042371)

5. Bouhouche K, Gout JF, Kapusta A, Bétermier M, Meyer E. 2011 Functional specialization of Piwi proteins in *Paramecium tetraurelia* from post-transcriptional gene silencing to genome remodelling. *Nucleic Acids Res.* 39, 4249–4264. (doi:10.1093/nar/gkq1283)

6. Sandoval PY, Swart EC, Arambasic M, Nowacki M. 2014 Functional diversification of Dicer-like proteins and small RNAs required for genome sculpting. *Dev.*

Cell 28, 174–188. (doi:10.1016/j.devcel.2013. 12.010)

7. Lepère G, Bétermier M, Meyer E, Duharcourt S. 2008 Maternal noncoding transcripts antagonize the targeting of DNA elimination by scanRNAs in *Paramecium tetraurelia*. *Genes Dev.* 22, 1501–1512. (doi:10.1101/gad.473008)

8. Mochizuki K, Fine NA, Fujisawa T, Gorovsky MA. 2002 Analysis of a piwi-related gene implicates small RNAs in genome rearrangement in *Tetrahymena*. *Cell* 110, 689–699. (doi:10.1016/ S0092-8674(02)00909-1)

9. Mochizuki K, Gorovsky MA. 2005 A dicer-like protein in *Tetrahymena* has distinct functions in genome
rearrangement, chromosome segregation, and
meiotic prophase. *Genes Dev.* **19**, 77–89. (doi:10.
1101/gad.1265105)

10. Malone CD, Anderson AM, Motl JA, Rexer CH, Chalker DL. 2005 Germ line transcripts are processed by a Dicer-like protein that is essential for developmentally programmed genome rearrangements of *Tetrahymena thermophila*. *Mol. Cell. Biol.* **25**, 9151–9164. (doi:10.1128/MCB.25.20. 9151-9164.2005)

11. Mochizuki K, Gorovsky MA. 2004 Conjugation-specific small RNAs in *Tetrahymena* have predicted properties of scan (scn) RNAs involved in genome rearrangement. *Genes Dev.* **18**, 2068–2073. (doi:10.1101/gad.1219904)

12. Chalker DL, Meyer E, Mochizuki K. 2013 Epigenetics of ciliates. *Cold Spring Harb. Perspect. Biol.* **5**, a017764. (doi:10.1101/cshperspect.a017764)

13. Allen SE, Nowacki M. 2017 Necessity is the mother of invention: ciliates, transposons, and transgenerational inheritance. *Trends Genet.* **33**, 197–207. (doi:10.1016/j.tig.2017.01.005)

14. Duharcourt S, Betermier M. 2014 Programmed rearrangement in ciliates: paramecium. *Microbiol. Spectr.* **2**. (doi:10.1128/microbiolspec.MDNA3-0035-2014)

15. Yao MC, Duharcourt S, Chalker DL. 2002 Genome-wide rearrangements of DNA in ciliates. In *Mobile DNA II* (eds N Craig, R Craigie, M Gellert, A Lambowiz), pp. 730–758. New York, NY: Academic Press.

16. Yerlici VT, Landweber LF. 2014 Programmed genome rearrangements in the ciliate *Oxytricha*. *Microbiol. Spectr.* **2**. (doi:10.1128/microbiolspec. MDNA3-0025-2014)

17. Karrer KM. 2000 *Tetrahymena* genetics: two nuclei are better than one. *Methods Cell Biol.* **62**, 127–186. (doi:10.1016/S0091-679X(08)61529-0)

18. Woodard J, Kaneshiro E, Gorovsky MA. 1972 Cytochemical studies on the problem of macronuclear subnuclei in *Tetrahymena*. *Genetics* **70**, 251–260.

19. Cheng C-Y, Yao M-C, Chao J-L. 2014 Programmed genome rearrangements in *Tetrahymena*. *Microbiol. Spectr.* **2**. (doi:10.1128/microbiolspec.MDNA3-0012-2014)

20. Orias E, Flacks M. 1975 Macronuclear genetics of *Tetrahymena*. I. Random distribution of macronuclear *genecopies* in *T. pyriformis*, syngen 1. *Genetics* **79**, 187–206.

21. Allen SL, Nanney DL. 1958 An analysis of nuclear differentiation in the selfers of *Tetrahymena*. *Am. Nat.* **92**, 139–160. (doi:10.2307/2458858)

22. Cole E, Sugai T. 2012 Developmental progression of *Tetrahymena* through the cell cycle and conjugation. *Methods Cell Biol.* **109**, 177–236. (doi:10.1016/
B978-0-12-385967-9-00007-4)

23. Akematsu T, Pearlman RE, Endoh H. 2010 Gigantic macroautophagy in programmed nuclear death of *Tetrahymena thermophila*. *Autophagy* **6**, 901–911. (doi:10.4161/auto.6.7.13287)

24. Austerberry CF, Allis CD, Yao MC. 1984 Specific DNA rearrangements in synchronously developing nuclei

of *Tetrahymena*. *Proc. Natl Acad. Sci. USA* **81**,
7383–7387. (doi:10.1073/pnas.81.23.7383)

25. Nikiforov MA, Smothers JF, Gorovsky MA, Allis CD. 1999 Excision of micronuclear-specific DNA requires parental expression of Pdd2p and occurs independently from DNA replication in *Tetrahymena thermophila*. *Genes Dev.* **13**, 2852–2862. (doi:10.1101/gad.13.21.2852)

26. Eisen JA et al. 2006 Macronuclear genome sequence of the ciliate *Tetrahymena thermophila*, a model eukaryote. *PLoS Biol.* **4**, e286. (doi:10.1371/journal.pbio.0040286)

27. Coyne RS et al. 2008 Refined annotation and assembly of the *Tetrahymena thermophila* genome sequence through EST analysis, comparative genomic hybridization, and targeted gap closure. *BMC Genomics* **9**, 562. (doi:10.1186/1471-2164-9-562)

28. Coyne RS, Stover NA, Miao W. 2012 Whole genome studies of *Tetrahymena*. *Methods Cell Biol.* **109**, 53–81. (doi:10.1016/B978-0-12-385967-9.00004-9)

29. Hamilton EP, Williamson S, Dunn S, Merriam V, Lin C, Vong L, Russell-Colantonio J, Orias E. 2006 The highly conserved family of *Tetrahymena thermophila* chromosome breakage elements contains an invariant 10-base-pair core. *Eukaryot. Cell* **5**, 771–780. (doi:10.1128/EC.5.4.771-780.2006)

30. Cranert S, Heyse S, Linger BR, Lescasse R, Price C. 2014 *Tetrahymena* Pot2 is a developmentally regulated paralog of pot1 that localizes to chromosome breakage sites but not to telomeres. *Eukaryot. Cell* **13**, 1519–1529. (doi:10.1128/EC.00204-14)

31. Lin CYG, Lin IT, Yao MC. 2016 Programmed minichromosome elimination as a mechanism for somatic genome reduction in *Tetrahymena thermophila*. *PLoS Genet.* **12**, e1006403. (doi:10.1371/journal.pgen.1006403)

32. Yao MC, Choi J, Yokoyama S, Austerberry CF, Yao CH. 1984 DNA elimination in *Tetrahymena*: a developmental process involving extensive breakage and rejoining of DNA at defined sites. *Cell* **36**, 433–440. (doi:10.1016/0092-8674(84)90236-8)

33. Fass JN et al. 2011 Genome-scale analysis of programmed DNA elimination sites in *Tetrahymena thermophila*. *G3* **1**, 515–522. (doi:10.1534/g3.111.000927)

34. Vogt A, Mochizuki K. 2013 A domesticated *piggyBac* transposase interacts with heterochromatin and catalyzes reproducible DNA elimination in *Tetrahymena*. *PLoS Genet.* **9**, e1004032. (doi:10.1371/journal.pgen.1004032)

35. Cheng C-YYC-Y, Vogt A, Mochizuki K, Yao M-CM-CC. 2010 A domesticated *piggyBac* transposase plays key roles in heterochromatin dynamics and DNA cleavage during programmed DNA deletion in *Tetrahymena thermophila*. *Mol. Biol. Cell* **21**, 1753–1762. (doi:10.1091/mbc.E09-12-1079)

36. Cheng C-Y, Young JM, Lin C-YG, Chao J-L, Malik HS, Yao M-C. 2016 The piggyBac transposon-derived genes TPB1 and TPB6 mediate essential transposon-like excision during the developmental rearrangement of key genes in *Tetrahymena*

thermophila. *Genes Dev.* **30**, 2724–2736. (doi:10.1101/gad.290460.116)

37. Feng L *et al*. 2017 A germline-limited piggyBac transposase gene is required for precise excision in *Tetrahymena* genome rearrangement. *Nucleic Acids Res.* **45**, 9481–9502. (doi:10.1093/nar/gkx652)

38. Huvos PE. 2007 Extensive changes in the locations and sequence content of developmentally deleted DNA between *Tetrahymena thermophila* and its closest relative, *T. malaccensis*. *J. Eukaryot. Microbiol.* **54**, 73–82. (doi:10.1111/j.1550-7408.2006.00148.x)

39. Huvos P. 2004 Modular structure in developmentally eliminated DNA in *Tetrahymena* may be a consequence of frequent insertions and deletions. *J. Mol. Biol.* **336**, 1075–1086. (doi:10.1016/j.jmb.2003.12.065)

40. Aronica L, Bednenko J, Noto T, DeSouza LVV, Siu KWMW, Loidl J, Pearlman REE, Gorovsky MAA, Mochizuki K. 2008 Study of an RNA helicase implicates small RNA-noncoding RNA interactions in programmed DNA elimination in *Tetrahymena*. *Genes Dev.* **22**, 2228–2241. (doi:10.1101/gad.481908)

41. Noto T, Kurth HM, Kataoka K, Aronica L, DeSouza LV, Siu KW, Pearlman RE, Gorovsky MA, Mochizuki K. 2010 The *Tetrahymena* Argonaute-binding protein Giw1p directs a mature argonaute-siRNA complex to the nucleus. *Cell* **140**, 692–703. (doi:10.1016/j.cell.2010.02.010)

42. Schoeberl EEE, Kurth HMM, Noto T, Mochizuki K. 2012 Biased transcription and selective degradation of small RNAs shape the pattern of DNA elimination in *Tetrahymena*. *Genes Dev.* **26**, 1729–1742. (doi:10.1101/gad.196493.112)

43. Noto T, Kataoka K, Suhren JH, Hayashi A, Woolcock KJ, Gorovsky MA, Mochizuki K. 2015 Small-RNA-mediated genome-wide trans- recognition network in *Tetrahymena* DNA elimination. *Mol. Cell* **59**, 229–242. (doi:10.1016/j. molcel.2015.05.024)

44. Mochizuki K, Gorovsky MA. 2004 Small RNAs in genome rearrangement in *Tetrahymena*. *Curr. Opin. Genet. Dev.* **14**, 181–187. (doi:10.1016/j.gde.2004.01.004)

45. Liu Y, Mochizuki K, Gorovsky MA. 2004 Histone H3 lysine 9 methylation is required for DNA elimination in developing macronuclei in *Tetrahymena*. *Proc. Natl Acad. Sci. USA* **101**, 1679–1684. (doi:10.1073/ pnas.0305421101)

46. Liu Y, Taverna SD, Muratore TL, Shabanowitz J, Hunt DF, Allis CD. 2007 RNAi-dependent H3K27 methylation is required for heterochromatin formation and DNA elimination in *Tetrahymena*. *Genes Dev.* **21**, 1530–1545. (doi:10.1101/gad.1544207)

47. Sugai T, Hiwatashi K. 1974 Cytologic and autoradiographic studies of the micronucleus at meiotic prophase in *Tetrahymena pyriformis*. *J. Protozool.* **21**, 542–548. (doi:10.1111/j.1550-7408.1974.tb03695.x)

48. Martindale DW, Allis CD, Bruns PJ. 1985 RNA and protein synthesis during meiotic prophase in
*Tetrahymena thermophila*. *J. Protozool.* **32**, 644–649. (doi:10.1111/j.1550-7408.1985.tb03094.x)

49. Chalker DL, Yao MC. 2001 Nongenic, bidirectional transcription precedes and may promote developmental DNA deletion in *Tetrahymena thermophila*. *Genes Dev.* **15**, 1287–1298. (doi:10.1101/gad.884601)

50. Mochizuki K, Gorovsky MA. 2004 RNA polymerase II localizes in *Tetrahymena thermophila* meiotic micronuclei when micronuclear transcription associated with genome rearrangement occurs. *Eukaryot. Cell* **3**, 1233–1240. (doi:10.1128/EC.3.5.1233-1240.2004)

51. Woehrer SLL, Aronica L, Suhren JHH, Busch CJ-LJ, Noto T, Mochizuki K. 2015 A *Tetrahymena* Hsp90 co-chaperone promotes siRNA loading by ATP-dependent and ATP-independent mechanisms. *EMBO J.* **34**, 559–577. (doi:10.15252/embj.201490062)

52. Iwamoto M, Mori C, Kojidani T, Bunai F, Hori T, Fukagawa T, Hiraoka Y, Haraguchi T. 2009 Two distinct repeat sequences of Nup98 nucleoporins characterize dual nuclei in the binucleated ciliate *Tetrahymena*. *Curr. Biol.* **19**, 843–847. (doi:10.1016/j.cub.2009.03.055)

53. Malone CD et al. 2008 Nucleus-specific importin alphas and nucleoporins regulate protein import and nuclear division in the bi-nucleate *Tetrahymena thermophila*. *Eukaryot. Cell* **7**, 1487–1499. (doi:10.1128/EC.00193-08)

54. Iwamoto M, Osakada H, Mori C, Fukuda Y, Nagao K, Obuse C, Hiraoka Y, Haraguchi T. 2017 Compositionally distinct nuclear pore complexes of functionally distinct dimorphic nuclei in ciliate *Tetrahymena*. *J. Cell Sci.* **130**, 1822–1834. (doi:10.1242/jcs.199398)

55. Kurth HMM, Mochizuki K. 2009 2'-O-methylation stabilizes Piwi-associated small RNAs and ensures DNA elimination in *Tetrahymena*. *RNA* **15**, 675–685. (doi:10.1261/rna.1455509)

56. Bednenko J, Noto T, DeSouza LVV, Siu KWMW, Pearlman REE, Mochizuki K, Gorovsky MAA. 2009 Two GW repeat proteins interact with *Tetrahymena thermophila* Argonaute and promote genome rearrangement. *Mol. Cell. Biol.* **29**, 5020–5030. (doi:10.1128/MCB.00076-09)

57. Trujillo JT, Mosher RA. 2017 Identification and evolutionary characterization of Argonaute-binding platforms. In *Methods Mol. Biol.* **1640**, 257–266. (doi:10.1007/978-1-4939-7165-7_19)

58. Madireddi MT, Coyne RS, Smothers JF, Mickey KM, Yao MC, Allis CD. 1996 Pdd1p, a novel chromodomain-containing protein, links heterochromatin assembly and DNA elimination in *Tetrahymena*. *Cell* **87**, 75–84. (doi:10.1016/S0092-8674(00)81324-0)

59. Kataoka K, Mochizuki K. 2015 Phosphorylation of an HP1-like protein regulates heterochromatin body assembly for DNA elimination. *Dev. Cell* **35**, 775–788. (doi:10.1016/j.devcel.2015.11.017)

60. Kataoka K, Noto T, Mochizuki K. 2016 Phosphorylation of an HP1-like protein is a prerequisite for heterochromatin body formation in *Tetrahymena* DNA elimination. *Proc. Natl Acad. Sci. USA* **113**, 9027–9032. (doi:10.1073/pnas.1606012113)

61. Kataoka K, Mochizuki K. 2017 Heterochromatin aggregation during DNA elimination in *Tetrahymena* is facilitated by a prion-like protein. *J. Cell Sci.* **130**, 480–489. (doi:10.1242/jcs.195503)

62. Lin IT, Chao JL, Yao MC. 2012 An essential role for the DNA breakage-repair protein Ku80 in programmed DNA rearrangements in *Tetrahymena thermophila*. *Mol. Biol. Cell* **23**, 2213–2225. (doi:10.1091/mbc.E11-11-0952)

63. Hediger F, Gasser SM. 2006 Heterochromatin protein 1: don't judge the book by its cover! *Curr. Opin. Genet.* **Dev.** **16**, 143–150. (doi:10.1016/j.gde.2006.02.013)

64. Grewal SI, Jia S. 2007 Heterochromatin revisited. *Nat. Rev. Genet.* **8**, 35–46. (doi:10.1038/nrg2008)

65. Hayashi A, Mochizuki K. 2015 Targeted gene disruption by ectopic induction of DNA elimination in *Tetrahymena*. *Genetics* **201**, 55–64. (doi:10.1534/genetics.115.178525)

66. Carle CM, Zaher HS, Chalker DL. 2016 A parallel g quadruplex-binding protein regulates the boundaries of DNA elimination events of *Tetrahymena thermophila*. *PLoS Genet.* **12**, e1005842. (doi:10.1371/journal.pgen.1005842)

67. Chalker DL, La Terza A, Wilson A, Kroenke CD, Yao MC. 1999 Flanking regulatory sequences of the *Tetrahymena* R deletion element determine the boundaries of DNA rearrangement. *Mol. Cell. Biol.* **19**, 5631–5641. (doi:10.1128/MCB.19.8.5631)

68. Wiley EA, Horrell S, Yoshino A, Schornak C, Bagnani C, Chalker D. In press. Diversification of HP1-like chromo domain proteins in *Tetrahymena thermophila*. *J. Eukaryot. Microbiol.* (doi:10.1111/jeu.12443)

69. Suhren JH, Noto T, Kataoka K, Gao S, Liu Y, Mochizuki K. 2017 Negative regulators of an RNAi-heterochromatin positive feedback loop safeguard somatic genome integrity in *Tetrahymena*. *Cell Rep.* **18**, 2494–2507. (doi:10.1016/j.celrep.2017.02.024)

70. Chung PH, Yao MC. 2012 *Tetrahymena thermophila* JMJD3 homolog regulates H3K27 methylation and nuclear differentiation. *Eukaryot. Cell* **11**, 601–614. (doi:10.1128/EC.05290-11)

71. Shieh AW, Chalker DL. 2013 LIA5 is required for nuclear reorganization and programmed DNA rearrangements occurring during *Tetrahymena macronuclear* differentiation. *PLoS ONE* **8**, e75337. (doi:10.1371/journal.pone.0075337)

72. Deragon JM, Capy P. 2000 Impact of transposable elements on the human genome. *Ann. Med.* **32**, 264–273. (doi:10.3109/07853890009011771)

73. Oliver KR, Greene WK. 2009 Transposable elements: powerful facilitators of evolution. *Bioessays* **31**, 703–714. (doi:10.1002/bies.200800219)

74. Vogt A, Goldman ADDAD, Mochizuki K, Landweber LFLF. 2013 Transposon domestication versus mutualism in ciliate genome rearrangements. *PLoS Genet.* **9**, e1003659. (doi:10.1371/journal.pgen.1003659)

75. Volff J-N. 2006 Turning junk into gold: domestication of transposable elements and the creation of new genes in eukaryotes. *Bioessays* **28**, 913–922. (doi:10.1002/bies.20452)

76. Chuong EB, Elde NC, Feschotte C. 2016 Regulatory activities of transposable elements: from conflicts to benefits. *Nat. Rev. Genet.* **18**, 71–86. (doi:10.1038/nrg.2016.139)

77. Cui B, Gorovsky MA. 2006 Centromeric histone H3 is essential for vegetative cell division and for DNA elimination during conjugation in *Tetrahymena thermophila*. *Mol. Cell. Biol.* **26**, 4499–4510. (doi:10.1128/MCB.00079-06)

78. Cervantes MD, Xi X, Vermaak D, Yao M-CC, Malik HS. 2006 The CNA1 histone of the ciliate *Tetrahymena thermophila* is essential for chromosome segregation in the germline micronucleus. *Mol. Biol. Cell* **17**, 485–497. (doi:10.1091/mbc.E05-07-0698)

79. Lhuillier-Akakpo M, Guérin F, Frapporti A, Duharcourt S. 2016 DNA deletion as a mechanism for developmentally programmed centromere loss. *Nucleic Acids Res.* **44**, 1553–1565. (doi:10.1093/nar/gkv1110)

80. Howard-Till RA, Lukaszewicz A, Novatchkova M, Loidl J. 2013 A single cohesin complex performs mitotic and meiotic functions in the protist *Tetrahymena*. *PLoS Genet.* **9**, e1003418. (doi:10.1371/journal.pgen.1003418)

81. Cervantes MD, Coyne RS, Xi X, Yao MC. 2006 The condensin complex is essential for amitotic segregation of bulk chromosomes, but not nucleoli, in the ciliate *Tetrahymena thermophila*. *Mol. Cell. Biol.* **26**, 4690–4700. (doi:10.1128/MCB.02315-05)

82. Liu Y, Song X, Gorovsky MA, Karrer KM. 2005 Elimination of foreign DNA during somatic differentiation in *Tetrahymena thermophila* shows position effect and is dosage dependent. *Eukaryot. Cell* **4**, 421–431. (doi:10.1128/EC.4.2.421-431.2005)

83. Howard-Till RA, Yao MC. 2007 Tudor nuclease genes and programmed DNA rearrangements in *Tetrahymena thermophila*. *Eukaryot. Cell* **6**, 1795–1804. (doi:10.1128/EC.00192-07)

84. Ammermann D. 1971 Morphology and development of the macronuclei of the ciliates *Stylonychia mytilus* and *Euplotes aediculatus*. *Chromosoma* **33**, 209–238. (doi:10.1007/BF00285634)

85. Nowacki M, Landweber LF. 2009 Epigenetic inheritance in ciliates. *Curr. Opin. Microbiol.* **12**, 638–643. (doi:10.1016/j.mib.2009.09.012)