Keeping it complicated: Mitochondrial genome plasticity across diplonemids

Valach, Matus; Moreira, Sandrine; Hoffmann, Steve; Stadler, Peter F.; Burger, Gertraud

doi:10.1038/s41598-017-14286-z

Download PDF

Article
Open access
Published: 26 October 2017

Keeping it complicated: Mitochondrial genome plasticity across diplonemids

Scientific Reports volume 7, Article number: 14166 (2017) Cite this article

1644 Accesses
18 Citations
3 Altmetric
Metrics details

Subjects

Abstract

Chromosome rearrangements are important drivers in genome and gene evolution, with implications ranging from speciation to development to disease. In the flagellate Diplonema papillatum (Euglenozoa), mitochondrial genome rearrangements have resulted in nearly hundred chromosomes and a systematic dispersal of gene fragments across the multipartite genome. Maturation into functional RNAs involves separate transcription of gene pieces, joining of precursor RNAs via trans-splicing, and RNA editing by substitution and uridine additions both reconstituting crucial coding sequence. How widespread these unusual features are across diplonemids is unclear. We have analyzed the mitochondrial genomes and transcriptomes of four species from the Diplonema/Rhynchopus clade, revealing a considerable genomic plasticity. Although gene breakpoints, and thus the total number of gene pieces (~80), are essentially conserved across this group, the number of distinct chromosomes varies by a factor of two, with certain chromosomes combining up to eight unrelated gene fragments. Several internal protein-coding gene pieces overlap substantially, resulting, for example, in a stretch of 22 identical amino acids in cytochrome c oxidase subunit 1 and NADH dehydrogenase subunit 5. Finally, the variation of post-transcriptional editing patterns across diplonemids indicates compensation of two adverse trends: rapid sequence evolution and loss of genetic information through unequal chromosome segregation.

The metazoan landscape of mitochondrial DNA gene order and content is shaped by selection and affects mitochondrial transcription

Article Open access 23 January 2023

Intragenomic rDNA variation - the product of concerted evolution, mutation, or something in between?

Article Open access 04 July 2023

Reconstruction of hundreds of reference ancestral genomes across the eukaryotic kingdom

Article Open access 16 January 2023

Introduction

Genomes balance between stability and plasticity, i.e., between conservation and alteration of the genetic information. Genomic plasticity includes reshuffling (inversions, translocations, fissions, and fusions), and elimination and acquisition of DNA, collectively referred to as genome rearrangements. In bacteria, genome plasticity can change the expression level of certain genes, cause antigenic variation, and protect against invasion by phages and mobile elements¹. Chromosome rearrangements in the nuclear genome contribute to speciation^2,3. In certain taxa, rearrangements take place in a programmed manner during development^4,5,6, while haphazard rearrangements drive progression of several human cancers (reviewed in⁷).

The large body of data on genome rearrangements in mitochondria documents not only sequence deletions, insertions, and reshuffling^8,9, but also change of topology by linearization of the originally circular chromosome, as well as disintegration into multiple chromosomes^10,11,12. A most spectacular kind of mitochondrial genome architecture is found in a group of unicellular marine flagellates: the diplonemids.

Traditionally, diplonemids (members of the Euglenozoa) have included two small genera, Diplonema and Rhynchopus. A recent addition is the genus Hemistasia, previously associated with Kinetoplastids, and containing a single recognized species, H. phaeocysticola ¹³. However, an environmental survey has uncovered two additional and diverse clades, named Deep-Sea Pelagic Diplonemids (DSPD) I and II¹⁴. Even more surprising are the recent data from the comprehensive Tara expedition, which classified diplonemids among the most cosmopolitan, most abundant, and most diverse eukaryotes in the oceans^15,16. Yet, as of now, not a single species of the DSPD-clades has been isolated.

The first diplonemid whose mitochondrial DNA (mtDNA) has been characterized is the model species Diplonema papillatum. We discovered that all genes except one are fragmented in up to 11 pieces (modules) of about 40–550 nt length, and almost each gene piece resides on its own chromosome^17,18. The fragmented structure of these mitochondrial genes is associated with a most unconventional architecture of the mitochondrial genome. This mtDNA consists of as many as 81 distinct circular chromosomes (each present in multiple copies) that fall into two classes based on molecule size (6 and 7 kbp) and sequence¹⁹. About 95% of a given chromosome’s length shares its sequence with that of the other members of its class. The remaining 5% of a chromosome is a distinctive, unique region, referred to as ‘cassette’, which includes a single gene module mentioned above²⁰. Gene-content wise, however, the D. papillatum mtDNA is quite conventional, specifying proteins of the respiratory chain and oxidative phosphorylation, and the ever-present large and small-subunit ribosomal RNAs (mt-LSU rRNA, mt-SSU rRNA).

Gene fragmentation in D. papillatum mtDNA is compensated at the post-transcriptional level. Separately transcribed gene pieces are first trimmed to coding-only sequences (modules), and then modules belonging to the same gene are covalently joined (trans-spliced) to form a full-length mRNA or rRNA^21,22. A number of mRNA and rRNA precursors is further decrypted by two different processes: (i) unique U-appendage RNA editing and (ii) substitution RNA editing at densely clustered sites, converting certain cytosines and adenosines into uridines and inosines, respectively (C-to-U; A-to-I)¹⁸. RNA editing by U additions also occurs in kinetoplastid mitochondria, but in that case, pre-mRNA is cut and resealed (reviewed in²³), whereas in diplonemids, Us are attached to module ends. Further, organellar C-to-U RNA editing has been reported before, mostly in plants, but also in certain amoeba and fungi²⁴. However, A-to-I substitutions in pre-mRNA and pre-rRNA are a first in diplonemid mitochondria.

Here, we investigate mitochondrial genome rearrangements across diplonemids, examining whether the eccentric, yet orderly mitochondrial genome structure of D. papillatum mitochondria is a landmark of diplonemids at large, or rather just one instance among a broad range of resourceful mtDNA architectures. We are also inquiring about the distribution and types of mitochondrial RNA editing across diplonemids. The present study is focusing on the Diplonema/Rhynchopus group, but also revisits data published by others on Hemistasia phaeocysticola.

Results

We have investigated mtDNAs from three diplonemid species available in culture collections, notably D. ambulator, D. sp. 2, and R. euleeides (Fig. 1). These species, together with D. papillatum examined earlier, represent the diversity of the Diplonema/Rhynchopus (D/R) clade according to nuclear-gene phylogenies^14,25.

The mitochondrial gene repertoire is conserved across diplonemids

Identification of gene modules encoded in diplonemid mtDNA is far from straightforward due to high sequence divergence and short module length. We used the following criteria: (i) sequence similarity with known mitochondrial genes and in particular with those from D. papillatum, (ii) evidence of directional transcription of the genomic region, (iii) inclusion in a mature transcript together with the more conserved module(s), and (iv) being flanked in the genome by the same repetitive motifs as those surrounding well conserved modules of a given mtDNA.

Using these criteria, we found in the mitochondrial genomes of the diplonemids investigated here the same 12 assigned genes present in D. papillatum, i.e., atp6 (ATP synthase subunit 6), cob (apocytochrome b), cox1–3 (cytochrome oxidase subunits 1, 2, and 3), nad1, 4, 5, 7, and 8 (NADH dehydrogenase subunits 1, 4, 5, 7, and 8), rnl, and rns (large and small-subunit ribosomal RNA, mt-LSU and mt-SSU rRNA). Among the six unassigned mitochondrial genes (y1-y6) of D. papillatum, all but y4 have been detected in the other diplonemids (Supplementary Fig. S1). Comparison of the inferred protein sequences of assigned mitochondrial genes shows rapid sequence evolution across diplonemids (Fig. 1). The cross-species identities range from moderate 46.6% for the most conservative protein Cox1 to merely 17.9% for the most variable, Nad5. With only ~9.5% overall identity, sequence conservation of the inferred Y proteins is extremely low.

Marginal variation in module number and gene breakpoint positions

In the four diplonemid species, the total number of modules that constitute the 17 shared mitochondrial genes is essentially identical (Table 1; Supplementary Table S1; Fig. 2b). The only exception is y3 of D. sp. 2, which is composed of four instead of five modules, the first module corresponding to modules m1 and m2 in the other species (Fig. 2d). Further, across these taxa, homologous modules with high sequence conservation are identical in length. Size differences up to 31 bp occur in modules with divergent sequences (e.g., modules of nad5 and y1-y5; Supplementary Table S1) and are almost all multiples of three in internal modules from protein-coding genes. Gene breakpoints are also highly conserved (Fig. 2c). Among the junctions that could be confidently aligned (59 out of 63 shared junctions), we found just seven cases with minor shifts (1 and 3 nt), each time in one of the four species (Fig. 2e, Supplementary Fig. S2).

Table 1 Mitochondrial chromosomes in D/R diplonemids.

Full size table

Number of mitochondrial chromosome classes differs among diplonemids

In the present study, we have obtained about 200 kbp of random mtDNA sequence for each, D. ambulator, D. sp. 2, and R. euleeides. Sequence assembly yielded four complete chromosomes from D. sp. 2, together with partial chromosomes sequences from all species (Supplementary Table S2). Despite the high coverage (>100×), sequence assembly of complete chromosomes is often hampered because of numerous intra- and inter-chromosomal repeats. Nevertheless, the new data confirm earlier reports that mitochondrial chromosomes in diplonemids are circular molecules of ~4–10 kbp in length^19,26,27.

For each of the three species, we aligned the sequences that surround gene modules. This allowed us to precisely locate module-adjacent unique regions constituting the distal portions of cassettes and, in addition, to identify recurring sequence motifs, which form the ‘constant regions’ that are shared with other chromosomes (see Fig. 3a). Then, chromosomes with similar constant regions were grouped together in a class (see Methods), revealing one or two classes more than in D. papillatum (Table 1; classes are referred to as Da_A, Da_B, etc.) There is no significant sequence similarity among the constant regions of the various classes from different species.

MtDNA of D. ambulator and D. sp. 2 also includes chromosomes of unique structure, which constitute the ‘unclassified’ (U) category (Table 1; Fig. 3a). Since these chromosomes do not share module-adjacent motifs with any other contig, cassette start and end points remain undefined. The four completely assembled chromosomes in D. sp. 2 belong to this category.

Diverse module/cassette arrangements in mitochondrial chromosomes across diplonemids

Unexpectedly, the three diplonemids investigated here do not adhere to the orderly mitochondrial chromosome architecture seen in D. papillatum—an architecture characterized by one module per cassette and one cassette per chromosome. In mtDNA of D. ambulator, D. sp. 2, and R. euleeides, we encountered, in addition to the ‘orthodox’ configuration (e.g., Fig. 3b, chromosome Da_A01), multiple modules congregated in a cassette, as well as, though less abundant, chromosomes harboring multiple cassettes (Table 1; Fig. 3a).

The two newly encountered architectures are most abundant and pronounced in D. ambulator. An example of a multi-module cassette resides in chromosome Da_A02, enclosing the third module of y2 (y2-m3) and nad1-m5 in opposite orientations (Fig. 3b). Multiple modules are also present in certain chromosomes of the ‘unclassified’ category and are arranged in cassette-like arrays. The largest array occurs in the chromosome Da_U01, containing as many as six modules spaced by merely ~100 bp of non-coding sequence (Fig. 3b). Throughout the diplonemids, modules grouped together in multi-module cassettes or arrays belong to different genes and are randomly combined. Only in one case are two such modules also neighbors in the mature transcript (chromosome Da_C14; Fig. 3c), and only one module combination occurs in two different species, namely nad7-m6 and y2-m3 in D. papillatum and D. sp. 2.

Da_A02 is a multi-cassette chromosome, whose configuration was readily resolvable by sequence assembly, because the two cassettes are only ~200 bp distant from one another (Fig. 3b). However, when cassettes on a chromosome are separated by long repeat stretches, assembly will likely result in several disconnected contigs. We confirmed four such instances by PCR, showing that contigs carrying a cassette c1 are part of a chromosome that also holds a cassette c2 in opposite position on the circular molecule (e.g., chromosome Da_C04; Fig. 3c). We infer that D. ambulator mtDNA contains 14 bi-cassette C-class chromosomes (Table 1). The unclassified chromosome Da_U02 (GenBank n. JF698652) has a similar cassette-like arrangement, but differs from C-class chromosomes in half of its non-coding sequence (Fig. 3c).

Finally, we found cassettes with tentative orphan modules in chromosomes of D. ambulator and D. sp. 2. Akin to X12 and X18 of D. papillatum, these hypothetical modules are transcribed bi-directionally at low level. Transcripts have variable termini and apparently are not post-transcriptionally edited, polyadenylated, or joined to other modules. In D. ambulator, all six orphan cassettes are short (~94 bp), share high nucleotide sequence identity to each other (86%), and reside on C-class chromosomes (Fig. 3c; Supplementary Table S2). Whether they have a biological function is currently unknown.

Certain modules code for pieces of two different genes

In the four diplonemids, about a third of the modules in multi-module cassettes are overlapping, either on the same strand or on opposite strands (Supplementary Figs S1, S3). Overlapping modules are exclusively pieces of protein-coding genes. In each of the three newly examined diplonemids, we encountered modules that are entirely nested one within the other (e.g., nad5-m3 embedded in nad5-m10 of the chromosome Da_C04 in Fig. 3c; Supplementary Fig. S3). All nested modules that are encoded on the same strand have the same reading frame. This results in two differently processed module transcripts that are incorporated into distinct mRNAs, yielding up to 22 amino acid-long identical sequences in two different proteins. In all cases, the embedded, shorter module is more divergent in protein sequence than the enclosing, larger counterpart (Table 2, Supplementary Fig. S3). As we discuss in more detail below, this may indicate that the enclosing ‘host’ module has secondarily shared its sequence with another gene.

Table 2 Embedded modules code for more divergent protein regions than their enclosing counterparts.

Full size table

U-appendage RNA editing: shared and species-specific sites

When comparing mitochondrial transcript and genome sequences of the three diplonemids studied here, we observed that RNA-DNA-indel differences consist exclusively of T insertions (in cDNA). As in D. papillatum, these insertions represent up to ~50-nt long U-additions at module-transcript 3′ ends, and occur in protein-coding and rRNA genes (Fig. 2a,b; Supplementary Table S3). About half of the 21 U-appendage sites are shared among all four diplonemids.

Only two sites (in cox1 and nad4) have identical U-tract length across the four D/R taxa. Remarkably, cross-species length variation of the U-tract at all other sites coincides with size deviation of the module upstream of the Us. For example, in D. ambulator and R. euleeides, the 3′-terminal nad5 module (nad5-m11) is about 25 nt shorter than those of D. sp. 2 and D. papillatum. In the former two species, RNA editing appends about 26 Us, but in the latter two taxa, only one or none (Fig. 2g). At internal modules of protein-coding genes, cross-species variations of U-tract length respect the corresponding reading frame (e.g., Fig. 2f; Supplementary Table S6). Apparently, U-appendage compensates for sequence loss in gene modules, as proposed earlier²⁴.

Substitution RNA editing: same cluster positions but different sites

The above-mentioned screening for mitochondrial RNA-DNA differences in the three diplonemids examined here also revealed nucleotide substitutions, notably A-to-I and C-to-U RNA editing as in D. papillatum ¹⁸. We experimentally confirmed the presence of inosines for mt-SSU rRNA of D. ambulator (Supplementary Fig. S4). The total number of substitution sites ranges from 78 (D. sp. 2) to 114 (D. papillatum) (Supplementary Table S3).

All these RNA editing sites are arranged in compact clusters (Fig. 2a,h; Supplementary Fig. S1). Out of the seven clusters in D. papillatum, five (in nad4, nad7-m3, rns, y2, and y5) are conserved across the four D/R group diplonemids, whereas those of y1 and y3 are only present in D. sp. 2 and R. euleeides, respectively. The latter two species share an additional small editing cluster in nad7-m5. Cluster sizes vary considerably, and individual editing sites within clusters only rarely coincide throughout the four species (Fig. 2h; Supplementary Fig. S1). As demonstrated earlier for nad4, substitution RNA editing makes protein sequences more similar within diplonemids and to homologs of other eukaryotes¹⁸.

We hypothesized that substitution editing equalizes the genomic A + T-content differences across diplonemids. Yet, this is only the case for mt-SSU rRNA, with a 16% across-diplonemid A + T-content difference among rns gene sequences shrinking to 7% among the edited transcripts. Interestingly, the A + T-content difference of diplonemid mt-LSU rRNA, which does not undergo substitution editing, is also 7%. The leveling of nucleotide composition in mt-SSU rRNA is due to predominant A-to-I substitutions in A + T-rich mtDNAs (R. euleeides) and predominant C-to-U edits in A + T-poor mtDNAs (D. papillatum; Fig. 2a). Apparently, secondary structure stability of the mature molecule favors the window of 51–58% A + T for diplonemid mt-rRNAs.

Hemistasia revisited: much more of the same compared to D/R diplonemids

A recent investigation of four mitochondrial genes (cob, cox1, cox2, and nad7) from Hemistasia phaeocysticola has shown that this apparently fast-evolving diplonemid¹³ has all particularities observed before in D. papillatum: multi-partite mtDNA, fragmented mitochondrial genes, and U-appendage, as well as C-to-U and A-to-I substitution RNA editing²⁸. Most gene breakpoints present in these four genes in the D/R diplonemids are found at same positions in H. phaeocysticola. Even in cases where junctions are ambiguous and remain to be experimentally confirmed in Hemistasia, potential shifts are only minor (e.g., Fig. 2h).

However, these four Hemistasia genes are much more fragmented than those from the D/R clade. For example, the Hp_cox1 gene includes according to the authors, in addition to 17 average-sized modules (29–132 bp), seven miniature modules (down to 7 bp), which are only seen in the transcript, but could not be identified in the available genome sequence²⁸. However, our data reanalysis leads to another conclusion. Most of these postulated mini-modules can be explained by large A-to-I and C-to-U editing clusters that impede proper alignment of mature transcript and genomic sequences, and thus hamper pinpointing gene-module starts and ends. For instance, at the gene breakpoint corresponding to the junction cox1-m6/m7 in the D/R group species, the postulated mini-module cox1-m18 of Hemistasia is in fact the 5′-moiety of the downstream module, camouflaged by heavy substitution RNA editing (Fig. 4; Supplementary Fig. S5). Larger modules reported as missing in the Hemistasia mtDNA are most likely encoded by chromosome classes that have not been sampled by the targeted PCR approach. To summarize, there are almost twice as many gene fragments in H. phaeocysticola mtDNA of about half the length compared to the D/R diplonemids (Supplementary Fig. S6). In fact, all modules larger than ~130 bp in the D/R taxa are split into two or more pieces in Hemistasia.

Our reexamination of the published Hemistasia data also allowed us to examine the cross-species distribution of substitution editing clusters. All four genes analyzed in Hemistasia appear to undergo substantial deamination RNA editing in that species, but not cob, cox1, and cox2 in the four D/R diplonemids (Supplementary Fig. S7). The two editing clusters present in nad7 of the D/R clade (Fig. 2b) are also found in Hemistasia, but the latter species contains 10 additional clusters with a total of 34 sites (Fig. 2h; Supplementary Fig. S7). Intriguingly, most additional substitution RNA editing clusters in Hemistasia occur near Hemistasia-specific module junctions, as does half of the U-appendage sites (Fig. 4; Supplementary Fig. S7).

Finally, the allelic state of the mitochondrial genomes appears to differ between that of D/R-diplonemids and Hemistasia. In contrast to an almost exclusively bi-allelic mode of single nucleotide polymorphic (SNP) sites in D. papillatum ¹⁸ and the three species investigated here (frequency of ~0.05–0.25% SNP per bp of coding sequence), Hemistasia’s cox1 gene modules were reported to contain ~7.4% SNPs with up to four different alleles²⁸. Unless the culture used in the study is not monoclonal, mitochondrial sequence heterogeneity of this diplonemid appears to be considerably higher than that of the D/R clade. Interestingly, certain SNP sites in Hemistasia undergo substitution RNA editing with the result that the inferred protein isoforms are more similar to one another (Supplementary Fig. S5C). This is consistent with our observation that substitution RNA editing renders diplonemid proteins more similar to homologs in other eukaryotes.

Discussion

Plasticity of the multipartite genome architecture in diplonemid mitochondria

We show here that in the D/R diplonemids, the configuration and module content of mitochondrial chromosomes vary considerably. First, the number of multi-member chromosome classes ranges from two to four. Second, some chromosomes (the ‘U’ category) are ‘outsiders’, as they do not cohere to any class. Third, in contrast to the regular architecture in the type species D. papillatum, mitochondrial chromosomes of the other three diplonemids may contain one or two cassettes (or module arrays) instead of only one, with ~15% of the cassettes including multiple modules (Table 1). Finally, when multiple gene pieces reside on the same chromosome, they seem grouped together at random, reflecting a high rate of genome rearrangement.

Multipartite organellar genomes are generally characterized by long repeat sequences that are shared by all chromosomes (e.g., in mitochondria of many metazoans²⁹, green algae³⁰, and protists³¹), or in plastids of dinoflagellates^32,33). A notable exception are the multiple unique chromosomes, as well as low-number member classes discovered in D. sp. 2 and R. euleeides, possibly indicating ongoing birth and death of chromosome classes. For instance, the chromosomes Ds_U02 and Ds_U03 might be in the process of founding a new common class, as they share two short regions (~100 and ~300 bp) of almost identical sequence. In contrast, the chromosomes Da_U01 and Da_U02 could be on their way to become members of the large classes, as they share some sequence similarity (~1 kbp with A and B classes, and ~2 kbp with the C class, respectively; Fig. 3). Alternatively, these chromosomes may be drifting away from their earlier affiliation with the larger classes.

Selection pressure imposed by multipartite genomes

When a genome is fragmented, there are essentially two ways to avoid gene loss during cell division: a well-organized partitioning system or, alternatively, stochastic repartition of massively and evenly amplified DNA. Cell microscopy studies of diplonemids show no indication of a kinetoplast body^19,26, a structure that ensures equal subdivision of the multipartite trypanosomatid mtDNA between daughter cells (reviewed in³⁴). The up to 130-fold variation in copy numbers of diplonemid mitochondrial chromosomes (correlating neither with a chromosome class, nor a particular gene; Supplementary Fig. S8; Supplementary Table S2), and the unusually high amount of mtDNA per cell (1:1 weight ratio of nuclear DNA versus mtDNA in D. papillatum) indicate a stochastic mode of mtDNA segregation and/or replication in diplonemids. This situation makes it quite plausible that certain chromosomes, and thus modules, are lost along generations. Therefore, a reduction in chromosome number via including multiple modules per circle instead of just one could provide an adaptive advantage.

Gene fragmentation allows multi-purpose use

Sequence overlaps of adjacent genes have been documented in numerous mitochondrial genomes across the eukaryotic tree (e.g., see^35,36). Diplonemids, however, take gene overlaps to a next level by reusing gene segments. One of the most spectacular cases involves nad5 and cox1 from D. sp. 2, where module nad5-m3 is completely embedded in cox1-m9 (Supplementary Fig. S3). This results in identical subsequences in the Nad5 and Cox1 proteins. Structural alignments of the deduced diplonemid proteins to homologs with known three-dimensional structure^37,38,39 indicate that all dual-purpose segments correspond to peripheral loops and short α-helices that connect trans-membrane helices, i.e., to generalizable domains of membrane proteins (Supplementary Fig. S9). We posit that a diplonemid mitochondrial genome with even smaller mitochondrial gene fragments would contain a larger number of dual-purpose modules than seen in this study, because the shorter the fragment, the more likely can the sequence it encodes be used for functionally different proteins—a hypothesis that would be worthwhile testing in Hemistasia.

The use of a given module transcript for distinct mRNAs observed in diplonemid mitochondria is equivalent to alternative trans-splicing in the metazoan nucleus (reviewed in^40,41). The difference is in the outcome: alternative splicing increases proteome complexity, whereas gene-module reuse results in the net reduction of coding sequence, indicating different evolutionary pressures.

U-appendage RNA editing restores missing coding sequence

Most gene breakpoints are precisely conserved across the four D/R-clade diplonemids, but the length of homologous modules can vary considerably when gene regions are poorly conserved. For example, the 3′-terminal gene module nad5-m11, which specifies a functionally important trans-membrane helix of the Nad5 protein³⁹, is about 25 nt shorter in D. ambulator and R. euleeides compared to the other two species. The missing coding sequences are added post-transcriptionally in the form of a U-tract of the corresponding length, which gives rise to a phenylalanine tract in the protein (Fig. 2g).

A similar restoration of incomplete genes occurs during maturation of tRNAs in the eukaryotic nucleus and many archaea and bacteria and involves addition of a CCA sequence at the transcript’s 3′ end (reviewed in⁴²). The completion of 3′ termini of several animal mitochondrial tRNAs by polyadenylation represents another well-known example of post-transcriptional ‘repair’^43,44. Lastly, in certain dinoflagellate mtDNAs, the gene cox3 is split into two fragments, which are brought together at the RNA level by an unknown mechanism⁴⁵. This trans-splicing is accompanied by insertion of an oligo-A tract⁴⁶, thus patching the missing information similar to U-appendage editing in diplonemids.

Module fusion or fission?

The breakpoints of D/R-clade mitochondrial genes are all identical, with one exception. The y3 gene of D. sp. 2 is lacking one split (Fig. 2d), i.e., the first ~300 nt of the corresponding mRNA are specified by a single gene module, but by two modules in the other D/R species. A comparison across all diplonemids shows that half of the Hemistasia gene modules are fused in the D/R species. Hemistasia might have diverged more rapidly than the D/R clade by accelerating gene fragmentation²⁸. In this case, the gene structure in the D/R group would represent a more ancestral state. Alternatively, the extreme gene fragmentation seen in Hemistasia may have already reigned in the common ancestor of all diplonemids, implying that the larger modules sizes in the D/R clade emerged by fusion of smaller gene pieces.

Evolutionary trends and strategies in diplonemid mitochondria

Under the premise of adaptive evolution, module nesting and U-appendage RNA editing in diplonemid mitochondria would be strategies to patch losses of coding sequence. Reduction of the genetic information would result in a more compact genome, thus countering the effect of unequal chromosome segregation. However, from the perspective of constructive neutral evolution^47,48, stochastic chromosome loss during segregation and ongoing gene decay were tolerated and allowed to accumulate because of the pre-existing, sophisticated post-transcriptional machinery.

According to nuclear rRNA phylogenies^13,14,16,49, the two DSPD clades evolve at a slower pace than the D/R clade and the Hemistasia group in particular. DSPD species might therefore provide a unique window on earlier stages of mitochondrial innovations in diplonemids. Work is underway to isolate and culture representatives of these environmentally and evolutionarily intriguing taxa.

Materials and Methods

Strains, culture, and DNA and RNA extraction

Diplonema papillatum (ATCC 50162), D. ambulator (ATCC 50223), Diplonema sp. 2 (ATCC 50224), and Rhynchopus euleeides (ATCC 50226) were obtained from the American Type Culture Collection and cultivated axenically as described earlier^22,27. Total cellular nucleic acids were extracted with a home-made Trizol substitute⁵⁰, allowing enrichment in small circular molecules, including diplonemid mitochondrial chromosomes⁵¹. DNA and RNA for library construction and other manipulations were obtained from the same sample. Residual RNA in DNA preparations was removed by RNase I treatment and samples were cleaned up using Genomic-tip 100/G (Qiagen). Residual DNA in RNA preparations was removed by either RNeasy (Qiagen) column purification, or digestion with TURBO DNase (Invitrogen) followed by extraction with the Trizol substitute.

Reverse transcription, PCR, 5′ RACE, RNase cleavage of glyoxalated RNA, and primer extension

Reverse transcription was performed with the AMV reverse transcriptase (Roche) or SuperScript IV Reverse Transcriptase (Thermo). DNA was amplified with Q5 High-Fidelity DNA Polymerase (New England BioLabs) or Platinum SuperFi DNA polymerase (Thermo). PCR products were purified using the Wizard SV Gel and PCR Clean-Up system (Promega) or Monarch DNA Gel Extraction Kit (New England BioLabs). To map the 5′ end of transcripts (5′ RACE), the RNA adaptor-oligonucleotide dp124 was ligated to total RNA with RNA ligase 1 (New England BioLabs) in the presence of 1 mM ATP and 15% PEG8000 for 4 h (1 h at 25 °C, 1 h at 16 °C, 2 h at 4 °C); RNA was then extracted and reverse-transcribed as described above. Inosines were mapped by primer extension on glyoxal-treated and RNase T1-digested RNA as described previously¹⁸. Detailed experimental procedures are available at https://www.protocols.io/u/matus-valach. Oligonucleotides (purchased from BioCorp and IDT) that were used as primers and adaptors are listed in Supplementary Table S4.

Genome and transcriptome sequencing and assembly

Illumina Tru-Seq genomic paired-end libraries were constructed from total DNA, multiplexed, and sequenced in a single MiSeq lane. Adaptor removal and quality clipping of reads were carried out as described previously¹⁸. Mitochondrial and nuclear genomic contigs were assembled from total DNA reads using Spades v3.8.0⁵². Total RNA-Seq libraries were constructed using an Illumina TruSeq kit, multiplexed, and sequenced in one MiSeq and one HiSeq lane. For de novo assembly, we used Trinity 2.2.0 with default parameters⁵³. Both library construction and sequencing of DNA and RNA were outsourced to the Genome Quebec Innovation Centre. Details on technology, read counts, and length are compiled in Supplementary Table S5. All data were deposited at NCBI: assembled mitochondrial genomic contigs and transcripts in GenBank (MF436742-MF436981; for details, see Supplementary Tables 2 and 7), and raw sequence data under BioProject PRJNA392339 (for accession numbers, see Supplementary Table S5).

Module and cassette annotation

Initial annotation. In general, de novo assembly of RNA-Seq reads produced partial mitochondrial transcripts because of the substantial amount of precursors in RNA preparations hampering full assembly. Transcripts were identified in two ways: by aligning them against known mitochondrial proteins, and in particular those of D. papillatum, using exonerate (–model protein2genome)⁵⁴, and by HMMER searches (HMMER 3.0)⁵⁵ against profile Hidden Markov Models (HMMs) constructed from excavate proteins. An in-house Python script (mitoTranscriptomeReconstruction.py; available upon request), based on the Overlap-Layout-Consensus (OLC) algorithm, compares the identified transcript fragments to each other with BLAT v. 35 × 1⁵⁶ and aligns identical regions. The script specifically tolerates poorly aligning transcript ends that correspond to the 5′ and 3′ flanking regions of unprocessed modules. To identify mitochondrial contigs and to delimit modules, full-length transcripts were reconstructed semi-manually and then compared with genomic contigs using BLAT. Identification of additional cassettes. We used the non-coding (or non-module) regions of initially identified mitochondrial contigs to search similar sequences in other contigs with NCBI-BLAST v2.3.0 (default options)^57,58. Among the contigs with significant hits, those with a length >200 bp and coverage >100× were considered mitochondrial candidate contigs. Occasionally, we detected chimeric contigs (part nuclear, part mitochondrial), which were produced by a single chimeric read pair. Such chimeric contigs were split to keep the mitochondrial and discard the nuclear moiety. Split point definition was based on their uneven coverage, since the coverage of validated mitochondrial contigs was >200× (Supplementary Table S2), whereas that for validated nuclear contigs was by far lower (~5× for unique and <100× for repetitive sequences). Mean coverage of a cassette or module array (modules + 50 bp up- and downstream) was calculated after mapping DNA-Seq reads onto mitochondrial contigs with Bowtie2 in the local mode. Prior to mapping, reads were merged with BBMerge (rem k = 62 extend2 = 50 ecct) and deduplicated with Dedupe (ac = f) from the BBMap 36.99 suite (http://jgi.doe.gov/data-and-tools/bbtools/). Box plots and data-point distributions were generated using BoxPlotR (http://shiny.chemgrid.org/boxplotr/)⁵⁹. Search for initially unrecognized modules and genes. Missing terminal modules were uncovered by iterative mapping of RNA-Seq reads on a partially reconstructed transcript reference and reads extending beyond the reference but not containing flanking regions were used for extending the partial transcript. In these cases, we confirmed the structure of the mature transcript by RT-PCR with an oligo-dT primer (for details, see above) and by sequencing the PCR amplicon. To pinpoint the highly divergent Y genes in the three diplonemids studied here, we first performed BLAST searches with DNA and inferred protein sequences of the D. papillatum Y genes against the de novo assembled transcripts and candidate mitochondrial genomic contigs. Once a Y gene’s most conserved module was spotted, the full transcript was reconstructed using the script mitoTranscriptomeReconstruction.py or by the aforementioned iterative mapping. Alternatively, we mapped the RNA-Seq data with Bowtie2⁶⁰ in local mode on the candidate mitochondrial genomic contigs, extracted soft-clipped sequences, and searched for those reads and read pairs that span two (or more) contigs. Finally, homologous Y gene modules were used to build HMM profiles to screen species in which certain modules were not identified with the above procedures. In unresolved cases, we used 5′ RACE (see above) and amplicon sequencing to precisely map the 5′ end of the mature transcript (e.g., y3 in D. sp. 2, R. euleeides, and D. papillatum). Pairwise nucleic acid and protein identities and similarities were calculated from a multiple sequence alignment, using the IUB and BLOSUM62 weight matrix, respectively, and visualized in Geneious 9.1⁶¹. Inference of module junctions. Out of the 251 module junctions across the four diplonemids investigated here, the exact position of 115 could not be inferred from the transcript sequence alone because of identical nucleotides at the 3′ end of the upstream module and the 5′ end of the downstream module (up to 6 nt identity). However, the inspection of mapped reads allowed us to precisely position the junction in most cases. The remaining 11 uncertain junctions are listed in Supplementary Table S6; all are adjacent to long U-appendages (two in D. papillatum, D. ambulator, and D. sp.2, and five in R. euleeides). In these cases, we chose the position downstream of a genome-encoded T residue. Similarly, when the last module ends with a T or an A, the residue could be genome-encoded or originate from U-appendage or polyadenylation, respectively. In cases where disambiguation was not feasible, we chose the genome-encoded base to represent the terminal nucleotide. Errors at module junctions. In contrast to earlier small-scale analyses²¹, the high coverage of our RNA-Seq data allowed us to detect a small number of imprecise trans-splicing events at certain junctions (usually <1%, but, for example, up to ~15% at the junction Da_y2-m3/m4). Four types of errors were noted: (i) over- or under-trimming of nucleotides at either the 5′, or 3′ end participating in the module-joining (usually <3 nt); (ii) U-addition at a 3′ end normally not undergoing U-appendage RNA editing (usually <3 nt); (iii) incompletely trimmed 3′ end instead of a U-tract; and (iv) mis-joining of non-cognate modules (usually <1%). These errors apparently represent the inherent noise of biological processes. An in-depth, comprehensive investigation of these phenomena is in preparation.

Chromosome classification

In D. papillatum, a cassette is defined as a unique sequence flanked by recurring motifs of the class-specific constant region (Fig. 3a). To classify mitochondrial chromosomes of the other diplonemids, module-containing contigs were compared among each other by BLAST to identify common sequences upstream and downstream of modules. Cassette-flanking motifs were defined as sequences 5′ or 3′ adjacent to modules, having >90% identity over >100 bp, and being shared by the highest number of contigs. Contigs with the same pair of cassette-flanking motifs are considered (partial) chromosomes of the same class. Cassette starts and ends were precisely determined by aligning these contigs using MAFFT v7.222 (–auto–adjustdirection)⁶², and placing boundaries at positions where sequence identities dropped below 90% over 100 bp.

Detection of RNA editing

Mapping of DNA-Seq reads onto mitochondrial contigs was performed with Bowtie2 in local mode, while RNA-Seq reads were mapped with segemehl⁶³ (available at http://www.bioinf.uni-leipzig.de/Software/segemehl-diplonema/). The latter tool allows correct mapping of intensely edited regions using a reduced alphabet during the extension step (-F 6), with an accuracy threshold of 95% (-A 95). Reads aligning with indels were discarded. In order to detect substitution editing sites, DNA and RNA variants were called with the GATK tool (-T UnifiedGenotyper -ploidy 100 -glm BOTH -stand_emit_conf 30 -stand_call_conf 30 -read_filter MappingQuality -drf DuplicateRead). DNA variants were then subtracted from RNA variants with the vcf-isec utility from the vcftool suite v0.1.12b. Variants were further filtered with the GATK tool (-T SelectVariants) according to the criteria “StrandOddsRatio” (SOR < 3), which detected the strand bias (only for DNA), and “QualByDepth” (QD > 2.0), which selected regions with sufficient mapping quality and coverage. Finally, variants were visually inspected. Incorrectly called variants were removed (<5%, occurring at module junctions due to read mapping artifacts), whereas variants with independent support were added (<1%, RNA editing sites that coincide, for example, with a genomic indel). Editing by U-appendage was detected by performing a local mapping of DNA-seq and RNA-seq reads onto reconstructed RNA transcripts with Bowtie2 and Segemehl, respectively. Regions not covered by DNA-Seq reads but overlapped by poly-T sequences on RNA-Seq reads specify loci of U-appendage. No other type of insertion editing was identified. The GenBank accession numbers for sequences of edited RNAs are listed in Supplementary Table S7.

The long conserved U-homopolymer downstream of y5-m1 could not be fully resolved by Illumina sequencing reads alone, as observed previously¹⁸. To more precisely measure its size, we performed RT-PCR with a high-fidelity DNA polymerase, and sequenced the amplicons from both sides of the U-tract in all four species. Although determining the precise sequence downstream of a homopolymer longer than 20 nt is virtually impossible by Sanger sequencing due to polymerase slippage, the size of the homopolymer can be determined more reliably based on the peak heterogeneity downstream of the homopolymer⁶⁴. Using this approach, the inferred length of the y5-m1 U-appendages was 48–50 nt.

Phylogenetic inference

We used all 10 assigned mitochondrion-encoded protein sequences (Atp6, Cob, Cox1–3, and Nad1,4,5,7,8) from the four diplonemids and the corresponding homologs from other Discoba, notably Hemistasia phaeocysticola (Diplonemea); Trypanosoma brucei (Kinetoplastida); Euglena gracilis (Euglenida); Acrasis kona, Naegleria gruberi, and Stachyamoeba lipophora (Heterolobosea); Andalucia godoyi, Reclinomonas americana (ATCC 50394), and Seculamonas ecuadoriensis (Jakobida). Homologs from slowly evolving species were added as well, i.e., two from Malawimonada (Malawimonas californiana and M. jakobiformis) and four from Archaeplastida (Porphyra purpurea (Rhodophyta), Prototheca wickerhamii, Klebsormidium flaccidum, and Marchantia polymorpha (Chlorophyta)). Sequences were downloaded from the NCBI GenBank Protein database. Only one of the multiple allelic variants for each of the H. phaeocysticola genes was selected for the analysis. The sequences of Cox1, Cox2, and Nad1 from M. jakobiformis, including minor substitutions due to RNA editing, can be obtained upon request. Multiple sequence alignments (MSAs) of proteins were generated with Mams, a tool developed in-house (Lang B.F. and Rioux P., unpublished; available upon request), which produces, by employing Muscle v3.8.31⁶⁵, a preliminary MSA for each protein class. These MSAs served to build HMMs, to which the protein sequences were aligned using hmmalign3 from the HMMER v3.0 package⁵⁵. Alignment columns with posterior probabilities <1 were eliminated, and subsequently, protein sequences were concatenated for each species. The final MSA contained 19 taxa and 3,137 amino acid positions (the alignment is available from the authors upon request). Phylogenetic inferences were performed by a Bayesian approach using posterior probabilities as support values (PhyloBayes v3.2e⁶⁶, MrBayes v3.2.6⁶⁷) and by a Maximum Likelihood (ML) approach with bootstrapping (RAxML v8.2.11)⁶⁸. Bayesian methods were executed in four independent chains and the first 25% cycles were discarded as burn-in. For Phylobayes, we chose the substitution model CAT-GTR, six discrete categories of gamma rate variation, and the -dc parameter to eliminate constant sites, and ~8,700 cycles corresponding to ~270,000 generations. For MrBayes, we chose the GTR model with six discrete categories of gamma rate variation, and 400,000 MCMC generations. For the ML tree, we used the substitution matrix LG for amino acid frequencies, eight distinct rate categories, a gamma model of rate heterogeneity with estimated parameters, the algorithm “rapid bootstrap analysis”, an optimization precision of 0.01 log likelihood units, and 100 distinct alternative runs on distinct starting trees for bootstrap support values.

Analysis of structural domains of proteins

Sequences of Cox1, Nad4, and Nad5 proteins were aligned to their mitochondrial homologs with known three-dimensional structures from the Protein Data Bank (PDB)⁶⁹ as described above. Structural alignments and models were inferred with Phyre2⁷⁰ and visually examined in UCSF Chimera⁷¹.

References

Darmon, E. & Leach, D. R. F. Bacterial genome instability. Microbiol. Mol. Biol. Rev. 78, 1–39 (2014).
Article CAS PubMed PubMed Central Google Scholar
Brown, J. D. & O’Neill, R. J. Chromosomes, conflict, and epigenetics: chromosomal speciation revisited. Annu. Rev. Genomics Hum. Genet. 11, 291–316 (2010).
Article CAS PubMed Google Scholar
Hou, J., Friedrich, A., de Montigny, J. & Schacherer, J. Chromosomal rearrangements as a major mechanism in the onset of reproductive isolation in saccharomyces cerevisiae. Current Biology 24, 1153–1159 (2014).
Article CAS PubMed PubMed Central Google Scholar
Bracht, J. R. et al. Genomes on the edge: Programmed genome instability in ciliates. Cell 152, 406–416 (2013).
Article CAS PubMed PubMed Central Google Scholar
Wang, J. & Davis, R. E. Programmed DNA elimination in multicellular organisms. Curr. Opin. Genet. Dev. 27, 26–34 (2014).
Article CAS PubMed Google Scholar
Bryant, S. A., Herdy, J. R., Amemiya, C. T. & Smith, J. J. Characterization of somatically-eliminated genes during development of the sea lamprey (Petromyzon marinus). Mol. Biol. Evol. 33, 2337–2344 (2016).
Article CAS PubMed PubMed Central Google Scholar
Leibowitz, M. L., Zhang, C.-Z. & Pellman, D. Chromothripsis: A new mechanism for rapid karyotype evolution. Annu. Rev. Genet. 49, 183–211 (2015).
Article CAS PubMed Google Scholar
Sloan, D. B. et al. Rapid evolution of enormous, multichromosomal genomes in flowering plant mitochondria with exceptionally high mutation rates. PLoS Biol. 10, e1001241–17 (2012).
Article CAS PubMed PubMed Central Google Scholar
Bernt, M. et al. A comprehensive analysis of bilaterian mitochondrial genomes and phylogeny. Mol. Phylogenet. Evol. 69, 352–364 (2013).
Article CAS PubMed Google Scholar
Shao, R., Kirkness, E. F. & Barker, S. C. The single mitochondrial chromosome typical of animals has evolved into 18 minichromosomes in the human body louse, Pediculus humanus. Genome Res. 19, 904–912 (2009).
Article CAS PubMed PubMed Central Google Scholar
Valach, M. et al. Evolution of linear chromosomes and multipartite genomes in yeast mitochondria. Nucleic Acids Res. 39, 4202–4219 (2011).
Article CAS PubMed PubMed Central Google Scholar
Kayal, E. et al. Evolution of linear mitochondrial genomes in medusozoan cnidarians. Genome Biol. Evol. 4, 1–12 (2012).
Article CAS PubMed Google Scholar
Yabuki, A. & Tame, A. Phylogeny and reclassification of Hemistasia phaeocysticola (Scherffel) Elbrächter & Schnepf, 1996. J. Eukaryot. Microbiol. 62, 426–429 (2015).
Article PubMed Google Scholar
Lara, E., Moreira, D., Vereshchaka, A. & López-García, P. Pan-oceanic distribution of new highly diverse clades of deep-sea diplonemids. Environmental Microbiology 11, 47–55 (2009).
Article CAS PubMed Google Scholar
de Vargas, C. et al. Eukaryotic plankton diversity in the sunlit ocean. Science 348, 1261605–1261605 (2015).
Flegontova, O. et al. Extreme diversity of diplonemid eukaryotes in the ocean. Curr. Biol. 26, 3060–3065 (2016).
Article CAS PubMed Google Scholar
Vlcek, C., Marande, W., Teijeiro, S., Lukeš, J. & Burger, G. Systematically fragmented genes in a multipartite mitochondrial genome. Nucleic Acids Res. 39, 979–988 (2011).
Article CAS PubMed Google Scholar
Moreira, S., Valach, M., Aoulad-Aissa, M., Otto, C. & Burger, G. Novel modes of RNA editing in mitochondria. Nucleic Acids Res. 44, 4907–4919 (2016).
Article CAS PubMed PubMed Central Google Scholar
Marande, W., Lukeš, J. & Burger, G. Unique mitochondrial genome structure in diplonemids, the sister group of kinetoplastids. Eukaryot. Cell 4, 1137–1146 (2005).
Article CAS PubMed PubMed Central Google Scholar
Marande, W. & Burger, G. Mitochondrial DNA as a genomic jigsaw puzzle. Science 318, 415–415 (2007).
Kiethega, G. N., Yan, Y., Turcotte, M. & Burger, G. RNA-level unscrambling of fragmented genes in Diplonema mitochondria. RNA Biol. 10, 301–313 (2013).
Article CAS PubMed PubMed Central Google Scholar
Valach, M., Moreira, S., Kiethega, G. N. & Burger, G. Trans-splicing and RNA editing of LSU rRNA in Diplonema mitochondria. Nucleic Acids Res. 42, 2660–2672 (2014).
Article CAS PubMed Google Scholar
Read, L. K., Lukeš, J. & Hashimi, H. Trypanosome RNA editing: the complexity of getting U in and taking U out. Wiley Interdiscip. Rev. RNA 7, 33–51 (2016).
Article CAS PubMed Google Scholar
Burger, G., Moreira, S. & Valach, M. Genes in hiding. Trends Genet. 32, 553–565 (2016).
Article CAS PubMed Google Scholar
Heyden von der, S., Chao, E. E., Vickerman, K. & Cavalier-Smith, T. Ribosomal RNA phylogeny of bodonid and diplonemid flagellates and the evolution of euglenozoa. J. Eukaryot. Microbiol. 51, 402–416 (2004).
Article Google Scholar
Roy, J., Faktorová, D., Lukeš, J. & Burger, G. Unusual mitochondrial genome structures throughout the Euglenozoa. Protist 158, 385–396 (2007).
Article CAS PubMed Google Scholar
Kiethega, G. N., Turcotte, M. & Burger, G. Evolutionarily conserved cox1 trans-splicing without cis-motifs. Mol. Biol. Evol. 28, 2425–2428 (2011).
Article CAS PubMed Google Scholar
Yabuki, A., Tanifuji, G., Kusaka, C., Takishita, K. & Fujikura, K. Hyper-eccentric structural genes in the mitochondrial genome of the algal parasite Hemistasia phaeocysticola. Genome Biol. Evol. 8, 2870–2878 (2016).
PubMed Central Google Scholar
Lavrov, D. V. & Pett, W. Animal mitochondrial DNA as we do not know it: mt-genome organization and evolution in nonbilaterian lineages. Genome Biol. Evol. 8, 2896–2913 (2016).
Article CAS PubMed PubMed Central Google Scholar
Smith, D. R., Hua, J. & Lee, R. W. Evolution of linear mitochondrial DNA in three known lineages of Polytomella. Curr. Genet. 56, 427–438 (2010).
Article CAS PubMed Google Scholar
Burger, G., Forget, L., Zhu, Y., Gray, M. W. & Lang, B. F. Unique mitochondrial genome architecture in unicellular relatives of animals. Proc. Natl. Acad. Sci. USA 100, 892–897 (2003).
Article ADS CAS PubMed PubMed Central Google Scholar
Zhang, Z., Green, B. R. & Cavalier-Smith, T. Single gene circles in dinoflagellate chloroplast genomes. Nature 400, 155–159 (1999).
Article ADS CAS PubMed Google Scholar
Mungpakdee, S. et al. Massive gene transfer and extensive RNA editing of a symbiotic dinoflagellate plastid genome. Genome Biol. Evol. 6, 1408–1422 (2014).
Article PubMed PubMed Central CAS Google Scholar
Jensen, R. E. & Englund, P. T. Network news: the replication of kinetoplast DNA. Annu. Rev. Microbiol. 66, 473–491 (2012).
Article CAS PubMed Google Scholar
Boore, J. L. Animal mitochondrial genomes. Nucleic Acids Res. 27, 1767–1780 (1999).
Article CAS PubMed PubMed Central Google Scholar
Burger, G., Gray, M. W., Forget, L. & Lang, B. F. Strikingly bacteria-like and gene-rich mitochondrial genomes throughout jakobid protists. Genome Biol. Evol. 5, 418–438 (2013).
Article PubMed PubMed Central CAS Google Scholar
Tsukihara, T. et al. The low-spin heme of cytochrome c oxidase as the driving element of the proton-pumping process. Proc. Natl. Acad. Sci. USA 100, 15304–15309 (2003).
Article ADS CAS PubMed PubMed Central Google Scholar
Solmaz, S. R. N. & Hunte, C. Structure of complex III with bound cytochrome c in reduced state and definition of a minimal core interface for electron transfer. J. Biol. Chem. 283, 17542–17549 (2008).
Article CAS PubMed Google Scholar
Fiedorczuk, K. et al. Atomic structure of the entire mammalian mitochondrial complex I. Nature 538, 406–410 (2016).
Article ADS CAS PubMed PubMed Central Google Scholar
Lasda, E. L. & Blumenthal, T. Trans-splicing. Wiley Interdiscip. Rev. RNA 2, 417–434 (2011).
Article CAS PubMed Google Scholar
Lei, Q. et al. Evolutionary insights into RNA trans-splicing in vertebrates. Genome Biol. Evol. 8, 562–577 (2016).
Article CAS PubMed PubMed Central Google Scholar
Tomita, K. & Yamashita, S. Molecular mechanisms of template-independent RNA polymerization by tRNA nucleotidyltransferases. Front. Genet. 5 (2014).
Yokobori, S. & Pääbo, S. Transfer RNA editing in land snail mitochondria. Proc. Natl. Acad. Sci. USA 92, 10432–10435 (1995).
Article ADS CAS PubMed PubMed Central Google Scholar
Rammelt, C. & Rossmanith, W. Repairing tRNA termini: News from the 3′ end. RNA Biol. 13, 1182–1188 (2016).
Article PubMed PubMed Central Google Scholar
Jackson, C. J. et al. Broad genomic and transcriptional analysis reveals a highly derived genome in dinoflagellate mitochondria. BMC Biol 5, 41–17 (2007).
Article PubMed PubMed Central CAS Google Scholar
Jackson, C. J. & Waller, R. F. A widespread and unusual RNA trans-splicing type in dinoflagellate mitochondria. PLoS ONE 8, e56777 (2013).
Article ADS CAS PubMed PubMed Central Google Scholar
Stoltzfus, A. On the possibility of constructive neutral evolution. J. Mol. Evol. 49, 169–181 (1999).
Article ADS CAS PubMed Google Scholar
Gray, M. W., Lukeš, J., Archibald, J. M., Keeling, P. J. & Doolittle, W. F. Irremediable complexity? Science 330, 920–921 (2010).
Gawryluk, R. M. R. et al. Morphological identification and single-cell genomics of marine diplonemids. Curr. Biol. 26, 3053–3059 (2016).
Article CAS PubMed Google Scholar
Rodríguez-Ezpeleta, N., Teijeiro, S., Forget, L., Burger, G. & Lang, B. F. Construction of cDNA libraries: focus on protists and fungi. Methods Mol. Biol. 533, 33–47 (2009).
Article PubMed CAS Google Scholar
Valach, M., Moreira, S., Faktorová, D., Lukeš, J. & Burger, G. Post-transcriptional mending of gene sequences: Looking under the hood of mitochondrial gene expression in diplonemids. RNA Biol. 13, 1204–1211 (2016).
Article PubMed PubMed Central Google Scholar
Bankevich, A. et al. SPAdes: A new genome assembly algorithm and its applications to single-cell sequencing. J. Comput. Biol. 19, 455–477 (2012).
Article CAS PubMed PubMed Central MathSciNet Google Scholar
Haas, B. J. et al. De novo transcript sequence reconstruction from RNA-seq using the Trinity platform for reference generation and analysis. Nat. Protoc. 8, 1494–1512 (2013).
Article CAS PubMed Google Scholar
Slater, G. S. C. & Birney, E. Automated generation of heuristics for biological sequence comparison. BMC Bioinformatics 6, 31 (2005).
Article PubMed PubMed Central CAS Google Scholar
Eddy, S. R. Accelerated profile HMM searches. PLoS Comput. Biol. 7, e1002195 (2011).
Article ADS CAS PubMed PubMed Central MathSciNet Google Scholar
Kent, W. J. BLAT—the BLAST-like alignment tool. Genome Res. 12, 656–664 (2002).
Article CAS PubMed PubMed Central Google Scholar
Altschul, S. F. et al. Gapped BLAST and PSI-BLAST: a new generation of protein database search programs. Nucleic Acids Res. 25, 3389–3402 (1997).
Article CAS PubMed PubMed Central Google Scholar
Camacho, C. et al. BLAST+: architecture and applications. BMC Bioinformatics 10, 421 (2009).
Article PubMed PubMed Central CAS Google Scholar
Spitzer, M., Wildenhain, J., Rappsilber, J. & Tyers, M. BoxPlotR: a web tool for generation of box plots. Nat. Methods 11, 121–122 (2014).
Article CAS PubMed PubMed Central Google Scholar
Langmead, B. & Salzberg, S. L. Fast gapped-read alignment with Bowtie 2. Nat. Methods 9, 357–359 (2012).
Article CAS PubMed PubMed Central Google Scholar
Kearse, M. et al. Geneious Basic: an integrated and extendable desktop software platform for the organization and analysis of sequence data. Bioinformatics 28, 1647–1649 (2012).
Article PubMed PubMed Central Google Scholar
Katoh, K. & Standley, D. M. MAFFT multiple sequence alignment software version 7: Improvements in performance and usability. Mol. Biol. Evol. 30, 772–780 (2013).
Article CAS PubMed PubMed Central Google Scholar
Otto, C., Stadler, P. F. & Hoffmann, S. Fast and sensitive mapping of bisulfite-treated sequencing data. Bioinformatics 28, 1698–1704 (2012).
Article CAS PubMed Google Scholar
Kieleczawa, J. Fundamentals of sequencing of difficult templates—an overview. J. Biomol. Tech. 17, 207–217 (2006).
PubMed PubMed Central Google Scholar
Edgar, R. C. MUSCLE: multiple sequence alignment with high accuracy and high throughput. Nucleic Acids Res. 32, 1792–1797 (2004).
Article CAS PubMed PubMed Central Google Scholar
Lartillot, N., Lepage, T. & Blanquart, S. PhyloBayes 3: a Bayesian software package for phylogenetic reconstruction and molecular dating. Bioinformatics 25, 2286–2288 (2009).
Article CAS PubMed Google Scholar
Ronquist, F. et al. MrBayes 3.2: efficient Bayesian phylogenetic inference and model choice across a large model space. Systematic Biology 61, 539–542 (2012).
Stamatakis, A. RAxML version 8: a tool for phylogenetic analysis and post-analysis of large phylogenies. Bioinformatics 30, 1312–1313 (2014).
Article CAS PubMed PubMed Central Google Scholar
Berman, H. M. The Protein Data Bank. Nucleic Acids Res. 28, 235–242 (2000).
Article ADS CAS PubMed PubMed Central Google Scholar
Kelley, L. A., Mezulis, S., Yates, C. M., Wass, M. N. & Sternberg, M. J. E. The Phyre2 web portal for protein modeling, prediction and analysis. Nat. Protoc. 10, 845–858 (2015).
Article CAS PubMed PubMed Central Google Scholar
Pettersen, E. F. et al. UCSF Chimera—a visualization system for exploratory research and analysis. J. Comput. Chem. 25, 1605–1612 (2004).
Article CAS PubMed Google Scholar

Download references

Acknowledgements

We acknowledge G. N. Kiethega and S. Teijeiro for culturing diplonemids and extracting DNA and RNA. We thank B.F. Lang for assistance in phylogenetic analyses, B.F. Lang and M. Sarrasin for critically reading the manuscript, and M. Aoulad Aissa for excellent technical assistance (all at the Université de Montréal, Montreal, Canada). We also thank A. Yabuki (JAMSTEC, Yokosuka, Japan) for sharing Hemistasia data prior to their release in GenBank. This work was supported by Canadian Institute of Health Research (CIHR, operating grant number MOP-79309); Gordon and Betty Moore Foundation (grant number GBMF-4983); and NSERC (grant number RGPIN-2014-05286).

Author information

Sandrine Moreira
Present address: Department of Biochemistry and Molecular Biophysics, Columbia University, Hammer Health Science Center, 701 W 168th St, New York, NY, 10031, USA
Matus Valach and Sandrine Moreira contributed equally to this work.

Authors and Affiliations

Department of biochemistry and Robert-Cedergren Centre for Bioinformatics and Genomics, Université de Montréal, 2900 Edouard-Montpetit, Montreal, H3T 1J4, QC, Canada
Matus Valach, Sandrine Moreira & Gertraud Burger
Leipzig University, LIFE - Leipzig Research Center for Civilization Diseases, Haertelstrasse 16-18, Leipzig, D-04107, Germany
Steve Hoffmann
Bioinformatics Group, Department Computer Science, and Interdisciplinary Center for Bioinformatics, University Leipzig, Härtelstrasse 16-18, D-04107, Leipzig, Germany
Peter F. Stadler
German Centre for Integrative Biodiversity Research (iDiv) Halle-Jena-Leipzig, Competence Center for Scalable Data Services and Solutions, and Leipzig Research Center for Civilization Diseases, University Leipzig, D-04107, Leipzig, Germany
Peter F. Stadler
Max Planck Institute for Mathematics in the Sciences, Inselstrasse 22, D-04103, Leipzig, Germany
Peter F. Stadler
Fraunhofer Institute for Cell Therapy and Immunology, Perlickstrasse 1, D-04103, Leipzig, Germany
Peter F. Stadler
Department of Theoretical Chemistry of the University of Vienna, Währingerstrasse 17, A-1090, Vienna, Austria
Peter F. Stadler
Center for RNA in Technology and Health, University of Copenhagen, Grønnegårdsvej 3, 1870, Frederiksberg C, Denmark
Peter F. Stadler
Santa Fe Institute, 1399 Hyde Park Road, Santa Fe, NM, 87501, USA
Peter F. Stadler

Authors

Matus Valach
View author publications
You can also search for this author in PubMed Google Scholar
Sandrine Moreira
View author publications
You can also search for this author in PubMed Google Scholar
Steve Hoffmann
View author publications
You can also search for this author in PubMed Google Scholar
Peter F. Stadler
View author publications
You can also search for this author in PubMed Google Scholar
Gertraud Burger
View author publications
You can also search for this author in PubMed Google Scholar

Contributions

G.B. conceived and orchestrated the study. S.M. and M.V. designed and conducted the bioinformatics and experimental approaches, respectively. M.V., S.M., and G.B. performed data analysis. S.H. and P.F.S. contributed bioinformatics tools and critical discussions. The manuscript was written by M.V., G.B., and S.M. All authors have approved the final manuscript.

Corresponding authors

Correspondence to Matus Valach or Gertraud Burger.

Ethics declarations

Competing Interests

The authors declare that they have no competing interests.

Additional information

Publisher's note: Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.

Electronic supplementary material

Supplementary Material

Rights and permissions

Open Access This article is licensed under a Creative Commons Attribution 4.0 International License, which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license, and indicate if changes were made. The images or other third party material in this article are included in the article’s Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the article’s Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder. To view a copy of this license, visit http://creativecommons.org/licenses/by/4.0/.

Reprints and permissions

About this article

Cite this article

Valach, M., Moreira, S., Hoffmann, S. et al. Keeping it complicated: Mitochondrial genome plasticity across diplonemids. Sci Rep 7, 14166 (2017). https://doi.org/10.1038/s41598-017-14286-z

Download citation

Received: 30 June 2017
Accepted: 06 October 2017
Published: 26 October 2017
DOI: https://doi.org/10.1038/s41598-017-14286-z

This article is cited by

Origin of minicircular mitochondrial genomes in red algae
- Yongsung Lee
- Chung Hyun Cho
- Hwan Su Yoon
Nature Communications (2023)
Recent expansion of metabolic versatility in Diplonema papillatum, the model species of a highly speciose group of marine eukaryotes
- Matus Valach
- Sandrine Moreira
- Gertraud Burger
BMC Biology (2023)
Single-cell genomics unveils a canonical origin of the diverse mitochondrial genomes of euglenozoans
- Kristína Záhonová
- Gordon Lax
- Jeremy G. Wideman
BMC Biology (2021)
Extensive molecular tinkering in the evolution of the membrane attachment mode of the Rheb GTPase
- Kristína Záhonová
- Romana Petrželková
- Marek Eliáš
Scientific Reports (2018)

Comments

By submitting a comment you agree to abide by our Terms and Community Guidelines. If you find something abusive or that does not comply with our terms or guidelines please flag it as inappropriate.