Review Article

The evolutionary origin of plant and animal microRNAs

  • Nature Ecology & Evolution 1, Article number: 0027 (2017)
  • doi:10.1038/s41559-016-0027
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MicroRNAs (miRNAs) are a unique class of short endogenous RNAs, which have become known in the past few decades as major players in gene regulation at the post-transcriptional level. Their regulatory roles make miRNAs crucial for normal development and physiology in several distinct groups of eukaryotes including plants and animals. The common notion is that miRNAs have evolved independently in those distinct lineages, but recent evidence from non-bilaterian metazoans, plants, and various algae raise the possibility that the last common ancestor of these lineages might already have employed an miRNA pathway for post-transcriptional regulation. In this Review we present the commonalities and differences of the miRNA pathways in various eukaryotes and discuss the contrasting scenarios of their possible evolutionary origin and their proposed link to organismal complexity and multicellularity.

MicroRNAs (miRNAs) are short (21–24 nucleotides (nt) long), endogenous, single-strand RNAs derived from hairpin transcripts that regulate gene expression in both animals and plants1,​2,​3,​4,​5. miRNAs repress gene expression by binding to a complementary mRNA target thereby mediating translational inhibition, degradation or cleavage3. They belong to a group of functional small RNAs, which also includes short interfering RNAs (siRNAs) responsible for RNA interference (RNAi) and PIWI-interacting RNAs (piRNAs) (for a review on piRNAs see ref. 6). Although siRNAs and miRNAs share important features and are both produced by the ribonuclease Dicer7, they are responsible for different cellular tasks and can be discerned from one another by unique characteristics (Fig. 1). Since their discovery in Caenorhabditis elegans more than two decades ago8,9 it has become clear that miRNAs play a role in a broad variety of biological processes and are essential for normal development of animals and plants2,3,10,11. The apparent rise in the number of miRNAs over the course of animal evolution and their involvement in regulating development raises the possibility that they might have contributed to the evolution of animal complexity12,13.

Figure 1: Differences between miRNAs and siRNAs.
Figure 1

a, A scheme of miRNA and siRNA precursors and duplexes. While miRNAs are usually produced from short hairpins carrying mismatches in their stem region, siRNAs are produced from long hairpins with stems of perfect complementarity. miRNA precursors usually give rise to a single duplex whereas siRNA precursors are a source for multiple duplexes. b, Small RNA profiles along a pre-miRNA sequence, here exemplified by miR-2024a of Nematostella vectensis. Note the homogenous product with the dominant guide strand (mature miRNA) and the neglectable passenger strand (miRNA*). c, Small RNA profiles along an siRNA precursor sequence, here exemplified by miR-2024c of N. vectensis. This siRNA locus was originally annotated as miRNA, but later determined to be an siRNA due to the fact it gives rise to multiple small RNAs45. The x-axis in b and c indicates the position along the hairpin sequence with paired (brackets) and unpaired nucleotides (dots). ppm = parts per million. b and c adapted from ref. 45, CSH Press.

The lack of sequence homology between miRNA families in plants and animals as well as differences in miRNA biogenesis and mode of action have led to the notion that miRNAs have evolved independently in both kingdoms from an ancient siRNA mechanism that already existed in the last common ancestor of all eukaryotes. However, recent studies relating plant and animal miRNAs unexpectedly raised the old questions: Did the common ancestor of all animals have miRNAs? Do miRNAs of plants and animals share a common origin? How many times did miRNAs evolve? This Review will attempt to re-address these questions.

miRNA sequence homology between distinct lineages

Is the lack of miRNA sequence homology between distinct lineages sufficient to determine convergence of the miRNA pathway? An early phylogenetic comparison of miRNAs revealed over 35 conserved families of miRNAs among bilaterian animals, and the pattern of conservation largely appeared to correlate with the accepted phylogeny. This led to the proposal that miRNAs are rarely lost once they have evolved14,15. Consequently, since no sequence similarity was detected between animal and plant miRNAs, they were considered to have evolved independently16. This line of thought extends also to several other lineages (Fig. 2). For instance, none of the eight miRNAs of the demosponge Amphimedon queenslandica is shared with other non-poriferan animals. Moreover, no conserved miRNAs could be detected between demosponge, calcareous and homoscleromorph sponges. This led to the extreme possibility that the miRNA pathway evolved convergently multiple times even within sponges13,17. Further, the traditional view that sponges are the sister group to all other animal lineages is under debate as several studies point to comb jellies (Ctenophora), which lack miRNAs, being the first animal lineage to diverge18,​19,​20. If Ctenophora was indeed the first animal lineage to diverge, an independent origin of animal miRNAs can be supported.

Figure 2: Phylogenetic tree of the major eukaryotic groups showing presence of miRNA systems.
Figure 2

Groups known to possess miRNAs are depicted in bold. Numbers in red count from top to bottom the maximum number of times miRNA systems evolved convergently. Figure adapted from ref. 85, The Royal Society.

Similarly, until recently, no shared miRNAs were observed between plants and green algae, suggesting a convergent origin of the miRNA pathway in these two main groups of Viridiplantae (Fig. 2)21. However, a new study annotating small RNAs of the liverwort Pellia endiviifolia revealed three miRNAs with high similarity to the green alga Chlamydomonas reinhardtii miRNAs22,23. This emphasizes the great importance of sampling many species from each phylum in order to fully perceive the extent of miRNA sequence conservation but also highlights the inherent difficulty of annotating miRNAs with high confidence24. Further, shallow sequencing depth as well as neglecting certain developmental stages and environmental conditions may mask the full miRNA repertoire of an organism and hence possible homology. Regardless of these caveats, the prevailing view is that the miRNA pathways of animals and plants evolved convergently14,16,21,25,​26,​27,​28. In fact, this would mean that the miRNA pathway evolved independently at least nine times (in bilaterians, in cnidarians; twice in sponges; in land plants; in green algae; in brown algae, in slime moulds and in excavates; Fig. 2)5,8,9,13,17,22,29,​30,​31,​32.

However, we suggest an alternative explanation: it might be possible that sequence turnover rates are high in plants and non-bilaterian animals, leaving no trace of shared miRNA sequences between contemporary lineages. In support of this scenario, plant miRNA genes are born and lost at high rates33,34, hence only a handful of expressed miRNAs are conserved among distant plant lineages35. Comparison of Arabidopsis thaliana and Arabidopsis lyrata small RNAs has shown that 33% of the miRNA families are not conserved between the two species, hence were gained or lost during the 10 million years (Myr) since they diverged33. Analysis based on genome data and small RNA sequencing from Capsella rubella, a very close relative of Arabidopsis, estimated that the net flux rate (birth – death) for miRNA genes in Arabidopsis is 1.2–3.3 genes per Myr (ref. 34). A high turnover rate of miRNAs is also suggested in green algae as only one miRNA is conserved between the two green algae Chlamydomonas and Volvox that separated about 200 million years ago (Ma)36,37. Within Bilateria, a recent study demonstrates that despite impressive examples of conservation, miRNA loss is much more common than previously appreciated38. Further, major losses of miRNAs have been reported within flatworms, a rapidly evolving lineage, perhaps as part of a morphological simplification trend39. Regardless, while the turnover of most bilaterian miRNAs might be higher than initially estimated, it is still lower than that of plants and estimated at 0.8–1.6 (or possibly even as low as 0.3) genes per Myr in Drosophila40,41.

Evolution of miRNAs in understudied eukaryotic groups

Because bilaterians have preserved many conserved miRNA families in comparison to plants, it is useful to consider outgroups to bilaterians. Among the non-bilaterians, placozoans and ctenophores do not possess miRNAs18, while in the sponge Amphimedon only eight miRNAs have been identified so far13. The phylum Cnidaria (corals, sea anemones, jellyfish and hydroids) represents the sister group of Bilateria, which branched off about 600 Ma (refs 19,20). A pioneering study of small RNAs in the sea anemone Nematostella vectensis indicated that this species possesses at least 40 miRNAs, yet only miR-100, is shared with Bilateria13. Remarkably, miR-100 is conserved over almost the full length of the miRNA, yet shifted by one nucleotide in the seed region, which was predicted to have significant consequences on the target specificity13. miR-100 function and its targets are not well conserved in Bilateria42 making its extreme sequence conservation in the sea anemone puzzling. However, lack of target conservation is true for most of the well-conserved bilaterian miRNAs, as they are frequently rewired into different genetic networks43. Interestingly, sequencing of small RNAs of the cnidarian Hydra magnipapillata revealed 126 miRNAs with no sequence similarity to those of Bilateria (hence no evidence of miR-100) and only two shared miRNAs with Nematostella44. These results suggest a rather fast turnover of miRNAs within Cnidaria and highlight the risk of misinterpreting lack of miRNA sequence homology as a lack of homology of the miRNA system.

In a recent study of Nematostella small RNAs obtained from several developmental stages, the miRNA complement was expanded to 87 miRNAs45. Interestingly, miR-9422 of Nematostella shares 16 of its positions, including the seed sequence with Arabidopsis miR-156a. This sequence similarity is unlikely to have occurred randomly as the chance of observing such a sequence identity between Nematostella and shuffled Arabidopsis miRNAs is very small (random sampling P-value < 0.01)45. As miR-156 is conserved from mosses to higher plants5,46,47, its similarity to miR-9422 could be an evidence for an miRNA conserved between animals and plants, thus supporting the hypothesis that miRNAs were inherited from the last common ancestor of the two groups. Taken together, in light of the potential high turnover rate, it is questionable whether lack of sequence homology can serve as a proof for multiple cases of convergent evolution of this pathway. Hence, additional features must be taken into account.

Another indication for evolutionary homology of animal and plant miRNA pathways comes from sequencing the genome and the small RNA repertoire of the symbiotic unicellular dinoflagellate Symbiodinium kawagutii, an outgroup to both plants and animals (Fig. 2), which revealed many potential shared miRNAs with plants and animals48. miRNAs have also been described in excavates, a group of unicellular eukaryotes that includes the parasites Giardia lamblia and Trichomonas vaginalis32,49. Several putative miRNA families from excavates have sequence homology to conserved animal and plant miRNAs. However, unlike most plant and animal miRNAs those peculiar excavate miRNAs are produced from small nucleolar RNAs (snoRNAs) and tRNAs and exhibit some additional irregular features. Furthermore, the possibility of horizontal gene transfer from a host to its symbiont or parasite could be an alternative explanation for the potential homology in these cases. Thus, further confirmation of these data from dinoflagellate and excavates is required. Yet, if those small RNAs are verified as bona fide miRNAs, this will be of great importance because it will strongly support the placement of the pathway's origin before the split of plants and animals.

Can miRNA biogenesis clarify the pathway's origin?

The discussion of whether plant and animal miRNA pathways evolved in a common ancestor or independently from an ancient RNAi mechanism must include the differences and similarities in miRNA biogenesis (Fig. 3). miRNAs are synthesized as primary hairpins (pri-miRNAs) and are processed to pre-miRNAs by cutting the hairpin stem, followed by cleaving the hairpin loop3,7. Next, they are loaded on to an Argonaute (AGO) protein and one RNA strand is selected for complementary target mRNA inhibition or cleavage3,7. In animals, the first step is conducted by a specialized microprocessor complex comprised of the RNAse III Drosha with the aid of the RNA binding protein Pasha (DGCR8 in vertebrates), while the second step of cleavage is performed by the RNAse III Dicer (Fig. 3)7. In plants, the Dicer homologue, DICER-LIKE 1 (DCL1), is responsible for both processing events required for miRNA maturation, conducting the two exact same steps in the same order2,28. In both plants and animals Dicer is crucial for processing the precursor miRNA (pre-miRNA) into mature miR/miR* dsRNA duplexes2,7,25,50. However, Dicers are common in many eukaryotes and also take part in non-miRNA activities such as RNAi via endogenous siRNA biogenesis and viral defence25,51 and therefore cannot be used as an argument for the common origin of miRNAs. Phylogenetic and structural analyses indicate that animal Drosha proteins are related to plant and animal Dicers, suggesting that in animals Drosha may have evolved following a duplication of the common ancestor of Dicer and Drosha and further specialised in the first processing step of miRNAs25,52. A very recent study found that miRNAs in Chlamydomonas are processed by DICER-LIKE3 (DCL3), a Dicer homologue that exhibits a domain organization similar to that of Drosha53. This finding hints at a possible evolutionary link between miRNA biogenesis in green algae and animals.

Figure 3: Scheme describing the plant and animal canonical miRNA biogenesis pathways.
Figure 3

Homologues carrying similar functions such as Ars2 of animals and Serrate of plants are represented in the same colour. Figure adapted from ref. 54, Oxford Journals.

Both Drosha and Pasha homologues were found in Cnidaria13,54 and in two sponge lineages13,17, for which independent evolution of the miRNA pathway has been proposed. Thus, Drosha and Pasha were already present in the common ancestor of all sponges, Cnidaria and Bilateria and probably served miRNA-related functions, although their involvement in the miRNA pathway in the first two lineages remains to be shown. If Drosha and Pasha have an ancestral function in miRNA processing in these lineages, the only explanation for the lack of homology of miRNA sequences in distinct sponge lineages would be high turnover rates of miRNA genes (see above). Nevertheless, it should be noted that Drosha and Pasha are known to be involved also in non-miRNA related functions in mammalian cells that range from ribosomal RNA maturation to cleavage of specific mRNAs (reviewed in ref. 55). The tendency to associate the presence of the microprocessor components with the presence of miRNAs in basally branching lineages may thus be misleading.

The site of miRNA biogenesis seems to differ between plants and animals. In plants, both processing steps by DCL1 take place in the nucleus, while in animals the first step performed by Drosha occurs in the nucleus and the second cleavage by Dicer occurs in the cytoplasm3,16. However, several studies in animals reported the presence of Dicer in the nucleus (reviewed in ref. 55). Whether the nuclear localization of Dicer in animals is a relic of an ancient miRNA-processing pathway or a secondary adaptation is an open question.

In plants and animals Dicer requires protein partners in order to accurately cleave pre-miRNAs (Fig. 3)2,7. In plants, DCL1 is assisted by the RNA binding proteins SERRATE (SE) and HYPONASTIC LEAVES1 (HYL1), both crucial for miRNA biogenesis and development2,56,57. Although an SE homologue, Ars2, is known in animals as a partner of the microprocessor and Dicer58, no HYL1 homologues were found in bilaterian animals. These differences in the biogenesis were taken as additional evidence for an independent evolution of plant and animal miRNAs. However, homologues for both SE and HYL1 were recently reported in the sponge Amphimedon and in several cnidarians including Nematostella, all possessing dsRNA binding motifs54. This suggests that a HYL1-like protein was present in the last common ancestor of plants and animals and was lost in multiple lineages, including Bilateria54. The lack of any known animal Dicer partners such as Loquacious (Loqs), TRBP or PACT7,59,​60,​61 in Nematostella54 suggests that HYL1 may constitute a Dicer partner in Cnidaria.

The miR/miR* duplexes in both plants and animals are similar: they are 22 nt long with imperfect complementarity between the two strands and a 2-nt 3′ overhang2,3,7. However, unlike in bilaterian animals, the stem-loop precursor in plants is long and variable16. Interestingly, sponges and slime moulds present longer pre-miRNAs than their bilaterian counterparts13,30. Those longer and more variable miRNA precursors are reminiscent of siRNA precursors. Interestingly, the miR-2024 family in Nematostella has members that are bona fide miRNAs whereas other members of the family show processing variability that is typical for siRNAs (Fig. 1b,c)45. This suggests that transformation of an miRNA into an siRNA or vice versa can happen in animals like in plants62 and that the siRNA-like features of some miRNAs are not sufficient in order to rule out a common origin for plant and animal miRNAs.

Differences can also be found in the genetic structure of miRNA genes in plants and animals. In animals, roughly 50% of miRNA genes are located in clusters, often comprised of different mature miRNAs16,63. In plants, fewer cases of miRNA clusters are found, mostly encoding miRNAs of the same family with noticeable homology16,64. Nevertheless, a few clusters of non-homologous miRNAs were discovered in plants, predicted to target related proteins64. Interestingly, the sea anemone Nematostella has only two repetitive clusters, both of miR-2024, similarly to plants45. Hence, it seems that despite having different frequencies, the same genomic topologies can be found in both kingdoms.

Another genomic pattern separating animal miRNAs from those of plants is the location of miRNA genes. Approximately 30% of animal miRNA genes are located in introns16,65. In contrast, only three tested examples of intronic miRNAs are known so far in plants66,​67,​68. The hypothesis that this distinction supports an independent origin of miRNA is undermined by a recent study on the green alga Chlamydomonas. This study found that unlike in plants, 50% of the miRNAs in Chlamydomonas are embedded within introns of protein-coding genes, similar to animals 53. These results can support a common origin of the animal and plant miRNA pathways, assuming higher plants lost this genomic feature.

In both plants and animals, binding of small RNAs to extensively complementary target RNAs induces degradation of the small RNA by adenosine or uracil addition (‘tailing’) and 3′-to-5′ exonucleolytic decay (‘trimming’)1,69. siRNAs in flies and piRNAs in germ cells of all animals are protected from these processes by 2′-O-methylation of their 3′ ends performed by the methyltransferase Hua enhancer 1 (HEN1)1,27. In plants, HEN1 is responsible for the methylation of both siRNAs and miRNAs1,2,70, while bilaterian miRNAs do not undergo this modification27. Interestingly, HEN1 was found to be expressed throughout the body in Nematostella54 and not only in germ cells, suggesting a non-restricted function. Moreover, periodate treatment proves that a major fraction of the Nematostella miRNAs is methylated, similar to plants13,45. One can speculate that the common ancestor of animals and plants possessed methylated miRNAs, a feature later lost in bilaterians possibly due to the loss of high complementarity between miRNAs and their targets (see next section).

miRNA mode of action varies between plants and animals

In both plants and animals siRNAs and miRNAs require a class of AGO proteins to execute gene regulatory functions71,​72,​73. The mature miRNA duplex is loaded onto AGO, a core component of the RNA-induced silencing complex (RISC) (Fig. 3)3,74,​75,​7675,76. The guide RNA strand is selected by AGO and directs RISC to the complementary RNA transcript3. AGO proteins are conserved throughout all classes of life, from Archaea and Bacteria to Eukarya, where they participate mostly in small non-coding RNA related mechanisms25,71,77,​78,​79,​80. Furthermore, the four structural domains of AGO proteins and their endonuclease and RNA-binding activity are highly conserved71,74. Target slicing by siRNA-guided AGO is used in a wide variety of organisms to protect cells from viruses and transposons81. In plants, in addition to those activities AGO proteins can cleave mRNA targets, thus controlling their expression. However, this gene regulation mechanism requires an miRNA guide loaded into the AGO protein3. Similar to siRNAs, plant miRNA target binding requires a nearly-full complementarity between the miRNA and its target, leading to the endonucleolytic cleavage of the target by AGO between position 10 and 11 of the miRNA, often resulting in a strong effect on a small number of targets1,82,​83,​84. The AGO proteins of prokaryotes that usually target foreign DNA also cleave their targets at the same position as plant AGOs, suggesting this is an ancient mode of action77,85,86. In contrast, in bilaterians, every miRNA potentially regulates a large number of targets because miRNA–mRNA recognition requires only a seven-nucleotide seed sequence, residing in positions 2–8 of the miRNA (Fig. 4)4. Unlike in plants, the vast majority of animal AGOs do not induce miRNA target cleavage. Instead, animal RISC induces translational repression of targets by blocking translation initiation or elongation or by deadenylation1,74. This mode of action results in relatively weak modulation of less than twofold both at the RNA and protein levels87,88. An advantage of translational repression is its reversibility, which allows rapid expression of existing blocked mRNAs in a given condition, time or location89,90. The destabilization and translational inhibition of targets is not performed directly by AGO, but by a partner protein known as GW182 in Drosophila or TNRC6 in vertebrates91,​92,​93,​94. Interestingly, GW182 proteins evolve very rapidly, making detection of homology even within bilaterians quite challenging95.

Figure 4: Schematic comparison of miRNA network topology in land plants and bilaterian animals.
Figure 4

Solid lines represent inhibition of targets by miRNA either by seed match (bilaterian animals) or by nearly full complementarity (land plants). Dotted lines represent reciprocal effect of targets on miRNAs.

The degree of complementarity between the miRNA and its mRNA target has been considered a major factor in determining the mode of target repression. High complementarity, as seen in plants, promotes target cleavage by AGO, while seed-matching leads to translational inhibition and is the common mode of action of bilaterian miRNAs74,92,96. The boundaries between plant and animal miRNA mode of action are becoming more blurry as there is more translation inhibition in plants than previously appreciated83,84,97,98. However, even this type of inhibition requires nearly full complementarity as short matches limited to the seed region were shown to confer no target inhibition in plants83,84. Surprisingly, miRNA-mediated translational inhibition in plants also depends on a GW-repeat protein called SUO99. GW182 of animals and SUO of plants do not share detectable homology. However, it is tempting to consider the possibility of a common origin of those two proteins that cannot be detected anymore due to the extremely fast evolution of the GW182 family.

In Bilateria there are only a handful of examples for miRNA-mediated target cleavage due to full complementarity of the miRNA to the target100,101. For instance, in a genome-wide screen, only a single transcript was shown to be cleaved by a perfectly matching miRNA in C. elegans6. Further, the majority of AGO proteins in Bilateria have lost their endonucleolytic activity74,102,103. However, endonucleolytic activity of the vertebrate AGO2 protein was shown to be evolutionarily conserved due to its role in processing a single miRNA, miR-451, which cannot be detected by Dicer43,104,105. Thus, it is plausible that the conservation of this AGO activity in vertebrates is unrelated to target regulation. These are strong indications that target cleavage is far from being a major mechanism of action of bilaterian miRNAs. Recently, Nematostella miRNAs were reported to induce target cleavage by nearly perfect complementarity, more resembling siRNAs and plant miRNAs than bilaterian miRNAs45. Unlike in Bilateria, the target cleavage mechanism seems to be common in Cnidaria as it is also present in Hydra, a cnidarian that diverged from Nematostella at least 550 Ma (ref. 45). The fact that both Cnidaria and plants use target cleavage mechanisms combined with the existence of siRNAs in fungi, plants and animals suggest that cleavage was the ancestral form of small RNA action. The presence of a cleavage mechanism in sponges, the only other non-bilaterian lineage with miRNAs, would strongly support this view. Although experimental evidence is still missing, at least several of the eight miRNAs identified in Amphimedon queenslandica show high complementarity to putative target mRNAs. Additionally, the origin of the miRNA systems in the siRNA mechanism, which is based on slicing, by itself suggests that high complementarity is the ancestral state of the miRNA mode of action. This is also supported by the fact that seed-matching does not promote target inhibition in plants83,84. Those mechanistic differences indicate that AGO holds the guide strand very differently in plants and bilaterian animals leading to different binding characteristics106. It seems that seed-matching is a derived state that appeared in the last common ancestor of all bilaterians. It is likely that this mode of action allowed animals to expand their regulatory networks as one miRNA can potentially inhibit hundreds of targets (Fig. 4)4,32. Those complex networks involving miRNAs might be a reason for the lower turnover rates of miRNAs in bilaterians since the loss of a single miRNA might affect the expression level of numerous targets.

miRNAs and the evolution of complex organisms

In both animals and plants gene regulation by miRNAs allowed organisms to develop more complex gene regulatory networks4,107. Further, miRNAs were theorized by various authors to drive evolution to a multicellular state, making it possible for complex organisms to evolve3,12,​13,​14,​15. To assess this theory, one needs to look into the evolution of multicellularity and the correlation between the miRNA repertoire and the complexity of an organism. Multicellular life forms had evolved from a unicellular ancestor independently in multiple lineages leading to fungi, animals, plants and various algae108,109. In all cases, it was speculated that this transition required the evolution of cell adhesion molecules, cell communication, co-operation between cells and cellular differentiation (division of labour)108,110,111. miRNAs in both plants and animals are known to be involved in pathways related to multicellularity such as development timing112,​113,​114, differentiation36,115,​116,​117,​118 and morphogenesis10. Additionally, in multicellular organisms ‘fine tuning’ of gene expression by miRNAs can lower phenotypic variation (‘canalization’) among individuals in a population and even among cells, thus reducing conflicts between cells of different genetic background12,110,119,120. Nevertheless, their connection to the evolution of multicellularity is at best a mere correlation. Indeed, many multicellular organisms including plants and animals possess miRNA regulatory mechanisms. However, miRNAs were found in unicellular organisms such as the green alga C. reinhardtii22,29 and the reef-building coral dinoflagellate endosymbionts Symbiodinium microadriaticum121 and Symbiodinium kawagutii48. This proves that the presence of miRNAs does not necessarily predict multicellularity. It can be hypothesized that miRNAs may be part of the genetic toolkit allowing the transition to multicellularity under the right environmental conditions. For example, it was demonstrated that under strong artificial settlement selection Chlamydomonas can develop a multicellular life stage122, suggesting that ecological conditions are as important as genetic potential on the way to multicellularity.

miRNAs are not mandatory for multicellularity as there are multicellular organisms that do not possess miRNAs such as fungi 123, placozoans13 and ctenophores18. In the case of ctenophores, it is possible that their extraordinary large collection of RNA-binding proteins substitutes the functions that miRNAs cover in other animals, allowing this phylum to produce complex cell types such as muscles and neurons20. Is the loss of complex, allegedly integral features feasible? For example, it was suggested that sponges and Placozoa might have lost their nervous system for an adaptive advantage72. A similar logic can explain the possible loss of miRNAs in organisms such as fungi, Placozoa or choanoflagellates. An example for advantageous loss of the RNAi components in fungi can be found in a study on the yeast Saccharomyces cerevisiae. The study demonstrated that several strains that had lost RNAi became more susceptible to killer virus infection when RNAi components were restored73. If losing the entire RNAi machinery is possible, we propose that losing only miRNAs must be a possibility as well. Despite of all this, the alternative scenario arguing that miRNAs evolved independently in plants, animals and other clades cannot be ruled out.

It has been suggested that the relative number of miRNAs is in correlation with the relative level of organism morphological complexity. Although it is generally difficult to define complexity, miRNA complements might have contributed to the complexity of gene regulation13,​14,​15,124,​125,​126. For example, sea anemones and humans possess 80 and 500–1,000 miRNAs, respectively (but see also a recent higher count for humans in ref. 127)13,24,45. The abundance of miRNAs in human in relation to the sea anemone may help explain how despite high similarity in their genome and protein-coding genes humans have evolved a much more complex body form and cell type diversity than the relatively simple sea anemone13,128. However, on closer inspection it seems that no strong correlation actually exists between the number of miRNAs and organismal complexity. For example, sea anemones, annelids and fruit flies have a similar number of miRNA families (80, 105 and 110, respectively) despite differences in morphological complexity14,45,126,129. Moreover, the thalliform multicellular brown alga Ectocarpus possesses over 60 miRNA families despite its low complexity14. Yet, instead of the number of miRNAs, one should consider the number of targets per miRNA. We hypothesize that the seed target recognition approach in bilaterian animals, where each miRNA recognizes and regulates many targets, allowed the evolution of highly complex and interwoven regulatory networks (Fig. 4), which may have contributed to cell type diversification during evolution.

Conclusion and future prospects

The rapidly expanding field of non-coding RNA research allows us to constantly re-evaluate the evolutionary origin of miRNAs. This review presents claims for and against a common origin of miRNA in plants and animals. Given the fast turnover in miRNA sequences and significant loss of miRNAs in both plants and animals33,34,38, we consider the lack of miRNA sequence homology known today as not sufficient to conclude on convergent evolution. Instead, the available information provides support for the possibility that the last common ancestor of plants and animals did possess an miRNA system (Fig. 5).

Figure 5: A possible scenario of miRNA evolution in plants and animals where their last common ancestor possessed an miRNA system.
Figure 5

Appearances and losses of proteins and traits are depicted on the relevant branches.

Of course, we cannot rule out convergent evolution. If indeed miRNAs have evolved independently at least nine times among eukaryotes (Fig. 2), it is intriguing to consider what is so unique about them that will support the evolutionary pressures required to allow convergence of such a magnitude. Ctenophores certainly demonstrate that an animal with a wide variety of cell types, including neurons and muscles can exist in the absence of this class of small RNAs18 and single-celled green algae demonstrate that miRNAs are not exclusive for ‘complex’ multicellular organisms29. Further, the lack of classic RNAi components in some fungi and the loss of Pasha (DGCR8) in Placozoa13,105 suggests that losing the protein machinery of the miRNA system is possible (Fig. 5)13.

Taken together, we can suggest two alternative hypotheses regarding the evolution of miRNAs: (1) the miRNA pathways and their mode of action via slicing originated convergently from siRNA mechanisms in the plant and several animal lineages and a system based on seed-matching of miRNA to its targets evolved later, after bilaterians diverged from the rest of animals; (2) the common ancestor of plants and animals already had an miRNA system acting via slicing that was recruited from the siRNA system. Still, in this scenario bilaterian animals also evolved a derived system based on seed-matching. In both scenarios the siRNA system evolved to protect cells from viruses and transposons and was adapted to post-transcriptional regulation of gene expression by targeting mRNAs.

Given the recent new findings, we favour the hypothesis of a common origin of miRNAs in plants and animals (Fig. 5). To further test this idea, biochemical and genetic methods are required to map the mode of action and the biogenesis pathway of miRNAs in groups beyond bilaterian animals and higher plants. We believe that some of the key points to study in the near future should be:

  • Test whether miRNAs in basally branching lineages such as sponges or algae cleave their targets.

  • Test whether the cnidarian HYL1 proteins, known to be involved in miRNA biogenesis in plants, are also involved in biogenesis of miRNAs in Cnidaria.

  • Assess the impact of the slicing versus the seed-matching mechanism on the gene regulatory networks in animals and plants.

  • Test whether homologous miRNA-related proteins in algae are indeed functional in the miRNA pathway.

  • Sequence small RNAs at an adequate depth from more eukaryotic lineages and annotate them as miRNAs according to widely accepted guidelines to reveal pattern of evolutionary turnover.

When those five points are met, we might get a better understanding of the evolution of the miRNA pathway in eukaryotes and answer the question of how ancient this system is.

Additional information

How to cite this article: Moran, Y., Agron, M., Praher, D. & Technau, U. The evolutionary origin of plant and animal microRNAs. Nat. Ecol. Evol. 1, 0027 (2017).


  1. 1.

    & Diversifying microRNA sequence and function. Nat. Rev. Mol. Cell Biol. 14, 475–488 (2013).

  2. 2.

    Origin, biogenesis, and activity of plant microRNAs. Cell 136, 669–687 (2009).

  3. 3.

    MicroRNAs: genomics, biogenesis, mechanism, and function. Cell 116, 281–297 (2004).

  4. 4.

    MicroRNAs: target recognition and regulatory functions. Cell 136, 215–233 (2009).

  5. 5.

    , , , & MicroRNAs in plants. Genes Dev. 16, 1616–1626 (2002).

  6. 6.

    , & PIWI-interacting RNA: its biogenesis and functions. Annu. Rev. Biochem. 84, 405–433 (2015).

  7. 7.

    , & Biogenesis of small RNAs in animals. Nat. Rev. Mol. Cell Biol. 10, 126–139 (2009).

  8. 8.

    , & The C. elegans heterochronic gene lin-4 encodes small RNAs with antisense complementarity to lin-14. Cell 75, 843–854 (1993).

  9. 9.

    , & Posttranscriptional regulation of the heterochronic gene lin-14 by lin-4 mediates temporal pattern formation in C. elegans. Cell 75, 855–862 (1993).

  10. 10.

    et al. MicroRNAs regulate brain morphogenesis in zebrafish. Science 308, 833–838 (2005).

  11. 11.

    Small RNAs and their roles in plant development. Annu. Rev. Cell Dev. 25, 21–44 (2009).

  12. 12.

    , & MicroRNAs and metazoan macroevolution: insights into canalization, complexity, and the Cambrian explosion. Bioessays 31, 736–747 (2009).

  13. 13.

    et al. Early origins and evolution of microRNAs and Piwi-interacting RNAs in animals. Nature 455, 1193–1197 (2008).

  14. 14.

    et al. microRNAs and the evolution of complex multicellularity: identification of a large, diverse complement of microRNAs in the brown alga Ectocarpus. Nucleic Acids Res. 43, 6384–6398 (2015).

  15. 15.

    et al. The Cambrian conundrum: early divergence and later ecological success in the early history of animals. Science 334, 1091–1097 (2011).

  16. 16.

    , & Vive la différence: biogenesis and evolution of microRNAs in plants and animals. Genome Biol. 12, 221 (2011).

  17. 17.

    et al. The identification of microRNAs in calcisponges: independent evolution of microRNAs in basal metazoans. J. Exp. Zool. B 320, 84–93 (2013).

  18. 18.

    , , , & MicroRNAs and essential components of the microRNA processing machinery are not encoded in the genome of the ctenophore Mnemiopsis leidyi. BMC Genomics 13, 714 (2012).

  19. 19.

    et al. The genome of the ctenophore Mnemiopsis leidyi and its implications for cell type evolution. Science 342, 1242592 (2013).

  20. 20.

    et al. The ctenophore genome and the evolutionary origins of neural systems. Nature 510, 109–114 (2014).

  21. 21.

    , & Do miRNAs have a deep evolutionary history? Bioessays 34, 857–866 (2012).

  22. 22.

    , , , & miRNAs control gene expression in the single-cell alga Chlamydomonas reinhardtii. Nature 447, 1126–1129 (2007).

  23. 23.

    et al. The liverwort Pellia endiviifolia shares microtranscriptomic traits that are common to green algae and land plants. New Phytol. 206, 352–367 (2015).

  24. 24.

    et al. A uniform system for the annotation of human microRNA genes and the evolution of the human microRNAome. Annu. Rev. Genet. 49, 213–242 (2015).

  25. 25.

    & On the origin and functions of RNA-mediated silencing: from protists to man. Curr. Genet. 50, 81–99 (2006).

  26. 26.

    & Plant and animal microRNAs: similarities and differences. Funct. Integr. Genomics 5, 129–135 (2005).

  27. 27.

    & Small silencing RNAs: an expanding universe. Nat. Rev. Genet. 10, 94–108 (2009).

  28. 28.

    , & MicroRNAs and their regulatory roles in plants. Annu. Rev. Plant Biol. 57, 19–53 (2006).

  29. 29.

    et al. A complex system of small RNAs in the unicellular green alga Chlamydomonas reinhardtii. Genes Dev. 21, 1190–1203 (2007).

  30. 30.

    , , & MicroRNAs in Amoebozoa: deep sequencing of the small RNA population in the social amoeba Dictyostelium discoideum reveals developmentally regulated microRNAs. RNA 18, 1771–1782 (2012).

  31. 31.

    et al. The small RNA repertoire of Dictyostelium discoideum and its regulation by components of the RNAi pathway. Nucleic Acids Res. 35, 6714–6726 (2007).

  32. 32.

    et al. Identification of putative miRNAs from the deep-branching unicellular flagellates. Genomics 99, 101–107 (2012).

  33. 33.

    , & Evolution and functional diversification of MIRNA genes. Plant Cell 23, 431–442 (2011).

  34. 34.

    et al. MicroRNA gene evolution in Arabidopsis lyrata and Arabidopsis thaliana. Plant Cell 22, 1074–1089 (2010).

  35. 35.

    , & Common functions for diverse small RNAs of land plants. Plant Cell 19, 1750–1769 (2007).

  36. 36.

    , & MicroRNAs in a multicellular green alga Volvox carteri. Sci. China Life Sci. 57, 36–45 (2014).

  37. 37.

    , , & Triassic origin and early radiation of multicellular volvocine algae. Proc. Natl Acad. Sci. USA 106, 3254–3258 (2009).

  38. 38.

    , , & A critical appraisal of the use of microRNA data in phylogenetics. Proc. Natl Acad. Sci. USA 111, E3659–E3668 (2014).

  39. 39.

    , , , & Substantial loss of conserved and gain of novel microRNA families in flatworms. Mol. Biol. Evol. 30, 2619–2628 (2013).

  40. 40.

    et al. The birth and death of microRNA genes in Drosophila. Nat. Genet. 40, 351–355 (2008).

  41. 41.

    et al. Evolutionary flux of canonical microRNAs and mirtrons in Drosophila. Nature Genet. 42, 6–9 (2010).

  42. 42.

    , , & The conserved miR-51 microRNA family is redundantly required for embryonic development and pharynx attachment in Caenorhabditis elegans. Genetics 185, 897–905 (2010).

  43. 43.

    & The evolution of gene regulation by transcription factors and microRNAs. Nat. Rev. Genet. 8, 93–103 (2007).

  44. 44.

    et al. Deep sequencing reveals unique small RNA repertoire that is regulated during head regeneration in Hydra magnipapillata. Nucleic Acids Res. 41, 599–616 (2013).

  45. 45.

    et al. Cnidarian microRNAs frequently regulate targets by cleavage. Genome Res. 24, 651–663 (2014).

  46. 46.

    et al. Cloning and characterization of micro-RNAs from moss. Plant J. 43, 837–848 (2005).

  47. 47.

    et al. Specific effects of microRNAs on the plant transcriptome. Dev. Cell 8, 517–527 (2005).

  48. 48.

    et al. The Symbiodinium kawagutii genome illuminates dinoflagellate gene expression and coral symbiosis. Science 350, 691–694 (2015).

  49. 49.

    & snoRNA, a novel precursor of microRNA in Giardia lamblia. PLoS Pathog. 4, e1000224 (2008).

  50. 50.

    et al. Multiple dicer genes in the early-diverging metazoa. Mol. Biol. Evol. 26, 1333–1340 (2009).

  51. 51.

    RNA-based antiviral immunity. Nat. Rev. Immunol. 10, 632–644 (2010).

  52. 52.

    et al. Structure of Human DROSHA. Cell 164, 81–90 (2016).

  53. 53.

    et al. Most microRNAs in the single-cell alga Chlamydomonas reinhardtii are produced by Dicer-like 3-mediated cleavage of introns and untranslated regions of coding RNAs. Genome Res. 26, 519–529 (2016).

  54. 54.

    , , & The evolution of microRNA pathway protein components in Cnidaria. Mol. Biol. Evol. 30, 2541–2552 (2013).

  55. 55.

    & Swiss army knives: non-canonical functions of nuclear Drosha and Dicer. Nat. Rev. Mol. Cell Biol. (2015).

  56. 56.

    , , & The nuclear dsRNA binding protein HYL1 is required for microRNA accumulation and plant development, but not posttranscriptional transgene silencing. Curr. Biol. 14, 346–351 (2004).

  57. 57.

    , , & The Arabidopsis double-stranded RNA-binding protein HYL1 plays a role in microRNA-mediated gene regulation. Proc. Natl Acad. Sci. USA 101, 1093–1098 (2004).

  58. 58.

    et al. Ars2 regulates both miRNA-and siRNA-dependent silencing and suppresses RNA virus infection in Drosophila. Cell 138, 340–351 (2009).

  59. 59.

    et al. The role of PACT in the RNA silencing pathway. EMBO J. 25, 522–532 (2006).

  60. 60.

    et al. Normal microRNA maturation and germ-line stem cell maintenance requires Loquacious, a double-stranded RNA-binding domain protein. PLoS Biol. 3, 1187 (2005).

  61. 61.

    et al. TRBP recruits the Dicer complex to Ago2 for microRNA processing and gene silencing. Nature 436, 740–744 (2005).

  62. 62.

    et al. Evolution of microRNA genes by inverted duplication of target gene sequences in Arabidopsis thaliana. Nat. Genet. 36, 1282–1290 (2004).

  63. 63.

    & Genomics of microRNA. Trends Genet. 22, 165–173 (2006).

  64. 64.

    , , & Plant polycistronic precursors containing non-homologous microRNAs target transcripts encoding functionally related proteins. Genome Biol. 10, R136 (2009).

  65. 65.

    , , & Identification of mammalian microRNA host genes and transcription units. Genome Res. 14, 1902–1910 (2004).

  66. 66.

    , , & A diverse and evolutionarily fluid set of microRNAs in Arabidopsis thaliana. Genes Dev. 20, 3407–3425 (2006).

  67. 67.

    et al. A diverse set of microRNAs and microRNA-like small RNAs in developing rice grains. Genome Res. 18, 1456–1465 (2008).

  68. 68.

    et al. Identification of mirtrons in rice using MirtronPred: a tool for predicting plant mirtrons. Genomics 99, 370–375 (2012).

  69. 69.

    et al. Target RNA–directed trimming and tailing of small silencing RNAs. Science 328, 1534–1539 (2010).

  70. 70.

    et al. Methylation as a crucial step in plant microRNA biogenesis. Science 307, 932–935 (2005).

  71. 71.

    et al. The evolutionary journey of Argonaute proteins. Nat. Struct. Mol. Biol. 21, 743–753 (2014).

  72. 72.

    & Where is my mind? How sponges and placozoans may have lost neural cell types. Phil. Trans. R. Soc. B 370, 20150059 (2015).

  73. 73.

    , & Compatibility with killer explains the rise of RNAi-deficient fungi. Science 333, 1592–1592 (2011).

  74. 74.

    & Argonaute proteins: key players in RNA silencing. Nat. Rev. Mol. Cell Biol. 9, 22–32 (2008).

  75. 75.

    Argonaute proteins: functional insights and emerging roles. Nat Rev. Genet. 14, 447–459 (2013).

  76. 76.

    , , & Distinct roles for Argonaute proteins in small RNA-directed RNA cleavage pathways. Genes Dev. 18, 1655–1666 (2004).

  77. 77.

    et al. DNA-guided DNA interference by a prokaryotic Argonaute. Nature 507, 258–261 (2014).

  78. 78.

    , , & Crystal structure of Argonaute and its implications for RISC slicer activity. Science 305, 1434–1437 (2004).

  79. 79.

    et al. Structure of an argonaute silencing complex with a seed-containing guide DNA and target RNA duplex. Nature 456, 921–926 (2008).

  80. 80.

    , , , & Bacterial argonaute samples the transcriptome to identify foreign DNA. Mol. Cell 51, 594-605 (2013).

  81. 81.

    & Origins and mechanisms of miRNAs and siRNAs. Cell 136, 642–655 (2009).

  82. 82.

    et al. Prediction of plant microRNA targets. Cell 110, 513–520 (2002).

  83. 83.

    . & Molecular insights into microRNA-mediated translational repression in plants. Mol. Cell 52, 591–601 (2013).

  84. 84.

    , & Analysis of complementarity requirements for plant microRNA targeting using a Nicotiana benthamiana quantitative transient assay. Plant Cell 26, 741–753 (2014).

  85. 85.

    , , & The evolutionary history of haptophytes and cryptophytes: phylogenomic evidence for separate origins. Proc. R. Soc. Lon. B 279, 2255–2261 (2012).

  86. 86.

    et al. Resolving difficult phylogenetic questions: why more sequences are not enough. PLoS Biol. 9, e1000602 (2011).

  87. 87.

    et al. Widespread changes in protein synthesis induced by microRNAs. Nature 455, 58–63 (2008).

  88. 88.

    et al. The impact of microRNAs on protein output. Nature 455, 64–71 (2008).

  89. 89.

    et al. Reversible inhibition of PSD-95 mRNA translation by miR-125a, FMRP phosphorylation, and mGluR signaling. Mol. Cell 42, 673–688 (2011).

  90. 90.

    , , , & Relief of microRNA-mediated translational repression in human cells subjected to stress. Cell 125, 1111–1124 (2006).

  91. 91.

    & The mechanics of miRNA-mediated gene silencing: a look under the hood of miRISC. Nat. Struct. Mol. Biol. 19, 586–593 (2012).

  92. 92.

    & Gene silencing by microRNAs: contributions of translational repression and mRNA decay. Nat. Rev. Genet. 12, 99–110 (2011).

  93. 93.

    et al. miRNA repression involves GW182-mediated recruitment of CCR4–NOT through conserved W-containing motifs. Nat. Struct. Mol. Biol. 18, 1218–1226 (2011).

  94. 94.

    et al. Identification of novel argonaute-associated proteins. Curr. Biol. 15, 2149–2155 (2005).

  95. 95.

    , , & The Caenorhabditis elegans GW182 protein AIN-1 interacts with PAB-1 and subunits of the PAN2-PAN3 and CCR4-NOT deadenylase complexes. Nucleic Acids Res. 40, 5651–5665 (2012).

  96. 96.

    & A microRNA in a multiple-turnover RNAi enzyme complex. Science 297, 2056–2060 (2002).

  97. 97.

    et al. Widespread translational inhibition by plant miRNAs and siRNAs. Science 320, 1185–1190 (2008).

  98. 98.

    , , , & Gene regulation by translational inhibition is determined by Dicer partnering proteins. Nat. Plants 1, 14027 (2015).

  99. 99.

    , & Mutations in the GW-repeat protein SUO reveal a developmental function for microRNA-mediated translational repression in Arabidopsis. Proc. Natl Acad. Sci. USA 109, 315–320 (2012).

  100. 100.

    et al. Diverse endonucleolytic cleavage sites in the mammalian transcriptome depend upon microRNAs, Drosha, and additional nucleases. Mol. Cell 38, 781–788 (2010).

  101. 101.

    et al. Expanding the microRNA targeting code: functional sites with centered pairing. Mol. Cell 38, 789–802 (2010).

  102. 102.

    , , , & The making of a slicer: activation of human Argonaute-1. Cell Rep. 3, 1901–1909 (2013).

  103. 103.

    et al. Turning catalytically inactive human Argonaute proteins into active slicer enzymes. Nat. Struct. Mol. Biol. 20, 814–817 (2013).

  104. 104.

    , , & A dicer-independent miRNA biogenesis pathway that requires Ago catalysis. Nature 465, 584–589 (2010).

  105. 105.

    , & Evolution and diversification of RNA silencing proteins in fungi. J. Mol. Evol. 63, 127–135 (2006).

  106. 106.

    , , , & Single-molecule imaging reveals that Argonaute reshapes the binding properties of its nucleic acid guides. Cell 162, 84–95 (2015).

  107. 107.

    & Computational identification of plant microRNAs and their targets, including a stress-induced miRNA. Mol. Cell 14, 787–799 (2004).

  108. 108.

    & The origins of multicellular organisms. Evol. Dev. 15, 41–52 (2013).

  109. 109.

    & Evolutionary Transitions to Multicellular Life: Principles and mechanisms Vol. 2 (Springer, 2015).

  110. 110.

    & Cooperation and conflict in the evolution of multicellularity. Heredity 86, 1–7 (2001).

  111. 111.

    , , , & Life-history evolution and the origin of multicellularity. J. Theor. Biol. 239, 257–272 (2006).

  112. 112.

    & The lin-4 regulatory RNA controls developmental timing in Caenorhabditis elegans by blocking LIN-14 protein synthesis after the initiation of translation. Dev. Biol. 216, 671–680 (1999).

  113. 113.

    A hierarchy of regulatory genes controls a larva-to-adult developmental switch in C. elegans. Cell 57, 49–57 (1989).

  114. 114.

    & Regulation of flowering time and floral organ identity by a microRNA and its APETALA2-like target genes. Plant Cell 15, 2730–2741 (2003).

  115. 115.

    & Differential patterns of microRNA expression in neuroblastoma are correlated with prognosis, differentiation, and apoptosis. Cancer Res. 67, 976–983 (2007).

  116. 116.

    , & Opposing microRNA families regulate self-renewal in mouse embryonic stem cells. Nature 463, 621–626 (2010).

  117. 117.

    et al. The interplay between the master transcription factor PU. 1 and miR-424 regulates human monocyte/macrophage differentiation. Proc. Natl Acad. Sci. USA 104, 19849–19854 (2007).

  118. 118.

    et al. Functional profiling reveals critical role for miRNA in differentiation of human mesenchymal stem cells. PLoS ONE 4, e5605 (2009).

  119. 119.

    & Biological principles of microRNA-mediated regulation: shared themes amid diversity. Nat. Rev. Genet. 9, 831–842 (2008).

  120. 120.

    & Canalization of development by microRNAs. Nat. Genet. 38, S20–S24 (2006).

  121. 121.

    et al. Integrating microRNA and mRNA expression profiling in Symbiodinium microadriaticum, a dinoflagellate symbiont of reef-building corals. BMC Genom. 14, 1 (2013).

  122. 122.

    et al. Experimental evolution of an alternating uni-and multicellular life cycle in Chlamydomonas reinhardtii. Nat. Commun. 4, 2742 (2013).

  123. 123.

    et al. Diverse pathways generate microRNA-like RNAs and Dicer-independent small interfering RNAs in fungi. Mol. Cell 38, 803–814 (2010).

  124. 124.

    , , , & MicroRNAs and the advent of vertebrate morphological complexity. Proc. Natl Acad. Sci. USA 105, 2946–2950 (2008).

  125. 125.

    , , & The phylogenetic distribution of metazoan microRNAs: insights into evolutionary complexity and constraint. J. Exp. Zool. Part B 306, 575–588 (2006).

  126. 126.

    et al. The deep evolution of metazoan microRNAs. Evol. Dev. 11, 50–68 (2009).

  127. 127.

    et al. Analysis of 13 cell types reveals evidence for the expression of numerous novel primate-and tissue-specific microRNAs. Proc. Natl Acad. Sci. USA 112, E1106–E1115 (2015).

  128. 128.

    et al. Sea anemone genome reveals ancestral eumetazoan gene repertoire and genomic organization. Science 317, 86–94 (2007).

  129. 129.

    et al. MicroRNAs resolve an apparent conflict between annelid systematics and their fossil record. Proc. R. Soc. B 276, 4315–4322 (2009).

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Small RNA research in the Moran lab is supported by a European Research Council Starting Grant (CNIDARIAMICRORNA, 637456) and a Young Investigator Grant by the German–Israeli Foundation for Scientific Research and Development (I-1058-203.7-2013). Research in the Technau group is supported by grants of the Austrian Research Fund FWF (P24858 and P22618).

Author information


  1. Department of Ecology, Evolution and Behavior, Alexander Silberman Institute of Life Sciences, The Hebrew University Jerusalem, Jerusalem 91904, Israel.

    • Yehu Moran
    •  & Maayan Agron
  2. Department of Molecular Evolution and Development, Centre of Organismal Systems Biology, University of Vienna, Althanstr. 14, 1090 Vienna, Austria.

    • Daniela Praher
    •  & Ulrich Technau


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Y.M. and U.T. conceived the manuscript, Y.M., M.A., D.P. and U.T. wrote the paper.

Competing interests

The author declares no competing financial interests.

Corresponding authors

Correspondence to Yehu Moran or Ulrich Technau.