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The Chloroplast Genome of Utricularia reniformis Sheds Light on the Evolution of the ndh Gene Complex of Terrestrial Carnivorous Plants from the Lentibulariaceae Family

  • Saura R. Silva,

    Affiliation Instituto de Biociências, UNESP - Univ Estadual Paulista, Câmpus Botucatu, São Paulo, Brazil

  • Yani C. A. Diaz,

    Affiliation Departamento de Biologia Aplicada à Agropecuária, Faculdade de Ciências Agrárias e Veterinárias, UNESP - Univ Estadual Paulista, Câmpus Jaboticabal, São Paulo, Brazil

  • Helen Alves Penha,

    Affiliation Departamento de Tecnologia, Faculdade de Ciências Agrárias e Veterinárias, UNESP - Univ Estadual Paulista, Câmpus Jaboticabal, São Paulo, Brazil

  • Daniel G. Pinheiro,

    Affiliation Departamento de Tecnologia, Faculdade de Ciências Agrárias e Veterinárias, UNESP - Univ Estadual Paulista, Câmpus Jaboticabal, São Paulo, Brazil

  • Camila C. Fernandes,

    Affiliation Departamento de Tecnologia, Faculdade de Ciências Agrárias e Veterinárias, UNESP - Univ Estadual Paulista, Câmpus Jaboticabal, São Paulo, Brazil

  • Vitor F. O. Miranda , (AMV); (VFOM)

    Affiliation Departamento de Biologia Aplicada à Agropecuária, Faculdade de Ciências Agrárias e Veterinárias, UNESP - Univ Estadual Paulista, Câmpus Jaboticabal, São Paulo, Brazil

  • Todd P. Michael,

    Affiliation Ibis Bioscience, Computational Genomics, Carlsbad, California, United States of America

  • Alessandro M. Varani (AMV); (VFOM)

    Affiliation Departamento de Tecnologia, Faculdade de Ciências Agrárias e Veterinárias, UNESP - Univ Estadual Paulista, Câmpus Jaboticabal, São Paulo, Brazil


The Chloroplast Genome of Utricularia reniformis Sheds Light on the Evolution of the ndh Gene Complex of Terrestrial Carnivorous Plants from the Lentibulariaceae Family

  • Saura R. Silva, 
  • Yani C. A. Diaz, 
  • Helen Alves Penha, 
  • Daniel G. Pinheiro, 
  • Camila C. Fernandes, 
  • Vitor F. O. Miranda, 
  • Todd P. Michael, 
  • Alessandro M. Varani


Lentibulariaceae is the richest family of carnivorous plants spanning three genera including Pinguicula, Genlisea, and Utricularia. Utricularia is globally distributed, and, unlike Pinguicula and Genlisea, has both aquatic and terrestrial forms. In this study we present the analysis of the chloroplast (cp) genome of the terrestrial Utricularia reniformis. U. reniformis has a standard cp genome of 139,725bp, encoding a gene repertoire similar to essentially all photosynthetic organisms. However, an exclusive combination of losses and pseudogenization of the plastid NAD(P)H-dehydrogenase (ndh) gene complex were observed. Comparisons among aquatic and terrestrial forms of Pinguicula, Genlisea, and Utricularia indicate that, whereas the aquatic forms retained functional copies of the eleven ndh genes, these have been lost or truncated in terrestrial forms, suggesting that the ndh function may be dispensable in terrestrial Lentibulariaceae. Phylogenetic scenarios of the ndh gene loss and recovery among Pinguicula, Genlisea, and Utricularia to the ancestral Lentibulariaceae cladeare proposed. Interestingly, RNAseq analysis evidenced that U. reniformis cp genes are transcribed, including the truncated ndh genes, suggesting that these are not completely inactivated. In addition, potential novel RNA-editing sites were identified in at least six U. reniformis cp genes, while none were identified in the truncated ndh genes. Moreover, phylogenomic analyses support that Lentibulariaceae is monophyletic, belonging to the higher core Lamiales clade, corroborating the hypothesis that the first Utricularia lineage emerged in terrestrial habitats and then evolved to epiphytic and aquatic forms. Furthermore, several truncated cp genes were found interspersed with U. reniformis mitochondrial and nuclear genome scaffolds, indicating that as observed in other smaller plant genomes, such as Arabidopsis thaliana, and the related and carnivorous Genlisea nigrocaulis and G. hispidula, the endosymbiotic gene transfer may also shape the U. reniformis genome in a similar fashion. Overall the comparative analysis of the U. reniformis cp genome provides new insight into the ndh genes and cp genome evolution of carnivorous plants from Lentibulariaceae family.


Carnivorous plants from the genus Utricularia (Lentibulariaceae) are distributed worldwide and are comprised of approximately 235 species occurring across every continent except the poles, some arid regions and oceanic islands [1]. They are highly specialized plants with modified leaves (traps) for capturing prey [2,3], and there are diverse life forms, such as aquatic, terrestrial, epiphytic, and reophytic forms [1]. In addition, the genus has some of the smallest nuclear genomes across the angiosperms, ranging from 77 to 561 megabases (Mb; 1C), surpassed only by the genus Genlisea with species possessing the smallest genomes [4]. Interestingly, angiosperms present extremes in genome sizes, from around 61 Mb of Genlisea to 150,000 Mb of one of the largest plant genomes known, the monocot Paris japonica [5,6]. From this perspective, Lentibulariaceae also provides outstanding candidates for model plants, with flexible genomes from around 61 to 1,500 Mb [4,7]; it is thus an important group to address evolutionary, genomic and phylogenetic as well as functional questions. For instance, it is well known that polyploidy, the amount of repetitive DNA such as transposable elements and other repeats, and whole genome duplications (WGD), along with other mechanisms such as small-scale genome duplications or fractionation/gene death rates are the key drivers of genome size differences [6,8,9].

Utricularia reniformis A.St.-Hil. is a terrestrial bladderwort endemic to the Brazilian Atlantic Forest and restricted to the mountaintops of the southeastern coast of Brazil [1,10]. All known populations are limited to elevations above 700 m, and live in wet grasslands, wet rocks, and in Bromeliaceae leaf axes [11]. Little is known about the U. reniformis genome structure. However, previous studies based on microsatellite markers have shown that different populations of U. reniformis may have tetraploid genomes with high levels of heterozygosity [10]. Recently, the aquatic U. gibba was sequenced, revealing that its 82 Mb genome encodes a typical number of genes for a plant at 28,500 but that the genome has gone through several whole genome duplications and reductions. In addition, it was reported that the 152kb chloroplast (cp) genome is similar to most other angiosperms [9]. Furthermore, additional studies have focused on genome contraction [12,13] and comparative transcriptomics comparing the different organs and tissues in the aquatic U. gibba genome [14]. In order to better understand the evolutionary dynamics of a terrestrial form, the Utricularia reniformis A.St.-Hil. genome and transcriptome of different tissues are being sequenced and analyzed by our research group.

Four complete cp genomes of the Lentibulariaceae have been published [9,15]. These are U. gibba and U. macrorhiza, which are found in aquatic environments as submersed plants, and Genlisea margaretae and Pinguicula ehlersiae, which are terrestrial species. These four cp genomes are composed of large and small single copy (LSC and SSC) regions and two inverted repeats (IRs), which is the typical circular cp genome quadripartite structure (LSC-IR-SSC-IR). In addition, independent deletions and pseudogenization of subunits of the NAD(P)H dehydrogenase complex (ndh), altered proportions of repeats, increase of substitution rates on coding regions and microstructural changes were observed as important landmarks of these cp genomes [15].

The ndh complex is composed of eleven genes (ndhA, ndhB, ndhC, ndhD, ndhE, ndhF, ndhG, ndhH, ndhI, ndhK, and ndhJ), and together with the nuclear genes nhdL, ndhM, ndhN, and ndhO encode the thylakoid NAD(P)H dehydrogenase complex. The complex is involved in the electron transfer from NAD(P)H to plastoquinone, protecting the plant cell against photo-oxidative stress, and maintaining optimal rates of cyclic photophosphorylation [16,17]. Interestingly, the ndh genes are related to land adaptation and photosynthesis, whereas small changes in any of the ndh genes significantly decrease the photosynthesis rate [18]. The ndh loss is mainly associated with heterotrophic (parasitic) plants and those endemic to an underwater environment [16]. Indeed, the plants found in submersed aquatic environments receive low levels of light, ultraviolet radiation and have specific limitations that require a number of adaptations, and therefore are under different selective pressures than terrestrial plants [19]. However, the ndh genes may be dispensable to the plant. Functional studies suggest that depending on the environmental condition and stress, alternative metabolic pathways might surpass the absence of the ndh genes, indicating that these genes may not be central for photosynthesis [19].

However, other questions about the evolution of the Lentibulariaceae cp genomes still remain. For instance, is lateral gene transfer of cp DNA to the nuclear and/or mitochondrial genomes occurring very frequently as it does in other angiosperms [20]. This process is termed endosymbiotic gene transfer [21], and has not been fully addressed in the Lentibulariaceae cp genomes. Furthermore, previous studies suggested that there is a close relationship between carnivorous and parasitic plants at the level of the cp genome [15,22]. Therefore, sequenced cp genomes from this order are still necessary to further understand this relationship, and also to shed light on the systematic relationships in the order Lamiales.

To provide additional insights into the plastid genomes and the driving forces related with plastid NAD(P)H dehydrogenase complex genes loss and pseudogenization in terrestrial carnivorous plants, we have sequenced the cp genome of the U. reniformis, and compared it to the other cp genomes among other Lamiales, with a specific focus on the carnivorous plants from the family Lentibulariaceae. We have also analyzed the gene content and expression by RNAseq, RNA editing, repeats and structure of the U. reniformis plastid genome.

Material and Methods

Plant Sampling

The Utricularia reniformis A.St.-Hil. plant samples were collected in the fall of 2015 near the Serra do Mar Atlantic Forest, located in Salesópolis Municipality, São Paulo State, Brazil (Geographic Location: -23.5047, -45.9018, 961 m a.s.l.) and deposited in the JABU Herbarium of São Paulo State University (voucher V.F.O de Miranda et al., 1670 –JABU). The sample was not collected in protected areas and U. reniformis is not a threatened species according to the global IUCN (The IUCN Red List of Threatened Species– and the Brazilian List of Threatened Plant Species [23].

Chloroplast Sequencing and Assembly

The DNA was extracted following the QIAGEN DNeasy Plant Maxi Kit extraction protocol (QIAGEN). The whole-genome shotgun sequencing was performed using the Illumina MiSeq technology with a paired-end library of 2x300bp and insert size of ~600 bp. The library construction followed the Illumina Nextera XT Preparation Guide (Illumina, USA). The DNA was tagmented (tagged and fragmented) by the Nextera XT transposome. The Nextera XT transposome simultaneously fragments the input DNA and adds adapter sequences to the ends. The tagmented DNA is amplified via a limited-cycle PCR program. The PCR step also adds index 1 (i7) and index 2 (i5) to sequences required for cluster formation. After that, a PCR clean-up was performed by AMPure XP beads to purify the library DNA, and provides a size selection step that removes very short library fragments from the population. A total of 40M paired-end reads were generated and used for the cp genome assembly. Furthermore, in our ongoing studies of the Utricularia reniformis genome, we also sequenced 160M mate-pair reads (2x100 bp) with an average of 3,500 bp insert size using the Illumina HiScanSQ technology. The library construction followed the Nextera mate pair gel free protocol (Illumina, USA). The mate-pair set of reads was used in this study to confirm the cp assembly. Poor quality sequences (phred < 24), contaminants, adapters, and sequences with less than 50bp were removed using seqyclean software (, leaving 36M (2x300 bp, paired-end) and 150M (2x100 bp, mate-pair) high quality reads.

The cp genome assembly was conducted in three steps. First, the trimmed reads were mapped back to Utricularia gibba, U. macrorhiza, Genlisea margaretae, and Pinguicula ehlersiae cp genomes (Accession numbers on S1 Table) with bowtie2 ( [24] using default parameters. In the second step, the resulting potential cp reads were then assembled separately with SPAdes v3.7.1 ( [25], and by iterative (mapping) assembly with MITObim ( [26], using U. gibba, U. macrorhiza, G. margaretae and P. ehlersiae cp genomes as references. In the third step, the cp genome was manually reconstructed using the SPAdes and MITObim results. The mate-pair reads were mapped back to the cp genome with bowtie2, with the—very-sensitive parameter, to confirm the assembly.

Annotation and Comparative Analysis of the Chloroplast Genomes

The cp genome was annotated using DOGMA (Dual Organellar GenoMe Annotator -[26] coupled with Prodigal v2.6.2 ( [27] and Blast ( [28] for additional gene location and refinements. The Aragorn software package ( [29] was used for tRNAs validation and intron identification. Corrections of start and stop codons, and annotation curation was made with Artemis genome browser ( [30]. In this study the potential pseudogenes were defined by Blast comparative analysis with the use of at least one of the following criteria: (a) presence of at least one stop codon in-frame with the predicted coding region; (b) absence of start and/or stop-codon; (c) frameshift; (d) lacking of at least 20% of the coding region when compared to the respective coding region of related species. For gene assignments the Uniprot ( [31] and InterProScan ( [32] databases were used. The codon and amino acid usage were calculated using CodonW v1.4.4 ( The circular gene maps were made with OGDRAW (OrganellarGenome DRAW - [33]. Comparative analysis was carried out with Interactivenn ( [34], and Blast.

The annotated sequence and the raw reads for the U. reniformis chloroplast genome have been deposited in the GenBank database under accession number [GenBank: KT336489 and SRR3277235, respectively] (BioProject PRJNA290588).

Microsatellite and other repeats analysis

Microsatellite analyses were carried out with the MISA software package ( [35], with thresholds of seven repeats for mononucleotide SSRs, four repeats for di- and trinucleotide SSRs, and three repeats for tetra-, penta- and hexanucleotide SSRs. Maximal number of bases interrupting 2 SSRs in a compound microsatellite was set to 100 bp. Direct and palindromic repeats were determined with REPuter ( [36], with a minimal size of ≥ 30 bp, sequence identity ≥ 90% (hamming distance of 3).

Phylogenomic and Phylogenetic Analyses: Maximum likelihood and Bayesian Inference

A total of 47 coding sequences from different chloroplasts were considered (S1A and S1B Table). The alignment of sequences was performed using MAFFT ( [37] with default parameters.

The phylogenetic analysis of the plastomes utilized Oleaceae as outgroup. For the probabilistic analysis, the best evolutionary models (best-of-fit) were chosen using ModelTest 3.7 ( [38]. Thus, the best-of-fit DNA model was evaluated for each data matrix with the corrected Akaike information criterion [39,40] to estimate the parameters. Maximum likelihood (ML) and Bayesian analyses were performed to estimate the phylogenetic hypothesis for each data matrix. The ML analyses were run with RAxML ( [41]. Prior to the probabilistic analyses, the Akaike information criterion was used to compare the fit to the data of different models as implemented in ModelTest, resulting in the selection of GTR+GAMMA+I as the best-of-fit model. Thus, the GTR+GAMMA+I model was also employed and bootstrapping was applied with 10,000 pseudoreplicates. Bayesian analyses were performed with MrBayes software version 3.2.5 ( [42] for each data set with 9x106 generations sampled for each 100 generations, using the default parameters. For each analysis, two runs (nruns = 2) with four chains (nchains = 4) were performed beginning from random trees. Initial samples were discarded after reaching stationary (estimated at 25% of the trees). Cladograms (except the one with optimizations of ancestral states) were drawn with the program TreeGraph2 beta version 2.0.52–347 ( [43].

The phylogenetic analyses of the evolution of ndh genes was carried out using a matrix of presence/absence (pseudogenes and frame-shifts were regarded as absences), and the plastome phylogenomic tree (described above) was considered for tracing the presence/absence of ndh genes with the Mesquite software version 3.04 ( The cloudgram was produced by DensiTree version 2.1 ( [44] based on 18,000 trees sampled with Bayesian inference (using the same parameters as described above for phylogenomic analyses of plastomes) but with the matK gene. The matK data set was produced by fifty-five sequences from NCBI and also by sequences produced by this study (S1C and S1D Table). The sequences produced in this study were made with 1R-KIM and 3F-KIM primers, following the PCR protocol and procedures recommended by the CBOL Plant Working Group [45].

RNA-Seq and RNA-edit analyses

Three different plant tissues were used for RNA-seq analysis; fresh leaves, stolons and utricules. The tissues were pooled in three replicates and the total RNA (including the rRNA) was extracted using the PureLink RNA Mini Kit (Thermo Fisher Scientific), according to the manufacturer’s protocol. DNase I (Thermo Fisher Scientific) was used to remove any genomic DNA contamination. The extracted RNA was evaluated using an Agilent 2100 Bioanalyzer (Agilent Technologies) and a Qubit 2.0 Fluorometer (Invitrogen). Only samples having an RNA integrity number (RIN) ≥ 7.0 were used for the sequencing. The cDNA libraries were sequenced on the Ion Proton System generating a yield of 180M of reads with an average read length of 200bp, representing the nuclear and organellar cDNAs. Poor quality sequences (phred < 20), adapters, bacterial contaminants such as photoautotrophic bacteria, and sequences with less than 20bp were removed using prinseq lite v0.20.4 [46]. Two different approaches were used to distinguish potential nuclear/mitochondrial transcripts from authentic plastid transcripts. First, filtered reads were mapped back to the assembled U. reniformis plastome with bowtie2, with the—very-sensitive and—end-to-end parameters. The libraries of all three tissues were pooled and this generated a total of 1,632,156 cp related reads with an average length of 170bp for downstream analysis. Second, the RNAseq reads were mapped with CLC Genomics Workbench v9 (QIAGEN Aarhus, Denmark - using the following parameters: mismatch cost of 3, insertion cost of 3, deletion cost of 3, minimal alignment coverage of 90% (Length fraction) and similarity fraction of >98%. The cp genes were considered for the RNAseq read mapping and transcription abundance by RPKM (Reads Per Kilobase Million) normalization, whereas only unique read mapping were considered. In addition, the intronic regions of intron-containing genes were also considered for the identification of spliced exons.

The RNA-editing analyses were conducted with TopHat2 ( [47] against the pooled reads using the following parameters: no coverage search (—no-coverage-search), filtering multiple mapped reads such as low complexity or repetitive reads (—prefiltered-multihits), reads spanned by junctions with a minimum of 10 bases (—min-anchor-length 10) or with a maximum of 1 base mismatch in the anchor region (—splice-mismatches 1), to align trans-spliced genes, fusion search was activated (—fusion-search) with the minimum distance of 10,000 (—fusion-min-dist 10,000). The final alignment was inspected for C to U or U to C nucleotide substitutions by a custom perl script, with the use of the following parameters: editing site with a minimum coverage of 10x and phred quality score of ≥25. In addition, the PREP-Cp tool ( [48] was used with default parameters to predict additional RNA editing sites.

The cp RNA-seq reads used in this study have been deposited in the GenBank database under accession number [GenBank: SRP072162] (BioProject PRJNA290588).


High-quality assembly of the chloroplast genome sequence

A total of 1,259,272 (paired-end, 2x300bp), and 2,087,893 (mate-pair, 2x100bp) chloroplast (cp) related reads were filtered from the raw reads generated by the Illumina MiSeq and HiScanSQ platforms, respectively. These reads shows an average phred value above 30. A total of 1,029,442 (81.7%) paired-reads were assembled to the U. reniformis cp contigs. This resulted in the assembly of the entire LSC region as well as part of the IRs and the SSC, in three different contigs. The plastid supercontig was manually closed by iterative read mapping and Blast searches. This resulted in a circular sequence with 139,725 bp, with a GC content of 38.15% and average coverage of 3,300x (maximum coverage peak of 8,563x, and standard deviation of 1,725). No high-quality discrepancies were observed, thus indicating a high quality cp genome assembly. A total of 1,751,896 (84%) mate-pair reads were mapped in pairs to the assembled cp genome according to the average insert size of the library (~3,500bp), thus confirming the cp genome assembly. In addition, the mate-pairs read mapping confirmed the assembly of the LSC/IRa, LSC/IRb, and IRa/SSC/IRb boundaries, and therefore the circularity of the plastid supercontig.

Interestingly, the remaining paired-end reads (2x300bp; 229,830; 18.3%) were assembled into contigs containing fragments of incomplete or truncated cp genes (at least 28 contigs encoding 48 truncated cp genes, which span a total of 23,606kbp—S2 Table). Interestingly, these truncated genes are present as full-length gene copies in the cp genome. It is worth noting that the boundaries of some of these contigs do not have similarity to the assembled U. reniformis cp genome, nor the other Lentibulariaceae cp genomes analyzed. Indeed, during the ongoing genome sequence of the Utricularia reniformis A.St.-Hil., we have noted that these cp-derived sequences are interspersed within U. reniformis mitochondrial (mt) and nuclear genome scaffolds (data not shown). Therefore, this finding suggests that these contigs do not belong to the cp genome sequence itself, and most likely represents distinct endosymbiotic gene transfer events. In addition to these 28 contigs, we have noted an additional number of truncated cp genes interspersed among the U. reniformis mt genome. For instance, truncated copies of the ndhJ, ndhK and ndhC genes, which are absent in the cp genome (see below), are present in the mt genome. These findings strongly support that gene transfer events between the organelles and nuclear DNA occurred and may shaped the U. reniformis genome.

Organization and gene content of the U. reniformis plastid genome

Utricularia reniformis circular cp genome size is 139,725 bp in length, showing the typical quadripartite structure found in most land plants. The IRs spans 24,064bp each (34%), whereas the small single copy (SSC), and large-single-copy region (LSC) span 12,661bp (10%) and 78,936bp (56%), respectively (Fig 1). The ndhJ, ndhK and ndhC, which are commonly located on the LSC in angiosperms, are lost in the U. reniformis cp genome (Fig 1). However, with that exception, the U. reniformis cp LSC is collinear in gene content and arrangement to the respective region of the closely related species, U. gibba, U. macrorhiza, G. margaretae and P. ehlersiae and to other angiosperms, such as Arabidopsis thaliana and Nicotiana tabacum. The GC content of the LSC and SSC regions are 36% and 31.75%, respectively, whereas that of the IR regions is 43%. As observed in other land plants, such as from Cynara humilis, a species of family Asteraceae, the higher GC content in the IRs is due to the GC rich rRNA genes rrn16, rrn23, rrn4.5, and rrn5 [49].

Fig 1. Genomic map of the Utricularia reniformis cp genome.

Genes shown on the outside of the map are transcribed clockwise, whereas genes on the inside are transcribed counter-clockwise. Genes are color coded by their function in the legend.

There are a total of 136 predicted coding regions, 90 of which are single copy (68 CDS and 22 tRNAs), and 46 of which are duplicated in the IRs (22 CDS, 16 tRNAs and 8 rRNAs) (Table 1). In addition to the predicted coding regions, 14 pseudogenes, 6 of which are single copies, mostly located on the SSC and related with ndh complex genes, were identified (Fig 1 and Table 1). A total of 22,601 codons represent the coding repertoire of the protein coding regions (Table 2). The most prevalent codon encodes for leucine (2,326–5.99%), whereas the least is cysteine (243–1.07%). Only four coding regions have alternative start codons, these are the rpl16 (AUC), rps19 (GUG) and ycf15 (GUG). All 3 of the stop codons are present, with UAA being the most frequently used (UAA 51%, UAG 27% and UGA 21%). The predicted tRNA genes enable the U. reniformis cp genome to decode all amino acids, but not all 61 codons (29 out 64 codons; Table 2). A similar codon distribution is also observed with the related species U. gibba, U. macrorhiza, G. margaretae and P. ehlersiae.

Table 1. List of genes encoded by the Utricularia reniformis chloroplast (cp) genome.

Table 2. Codon usage table.

Codon usage and codon—anticodon recognition pattern for tRNA in Utricularia reniformis cp genome.

Other features are related with the genes atpF, rpl2, rps16, rpoC1, rps16, petB and petD which each contain one intron, and clpP and ycf3 which each contain two introns. The introns of all protein-coding genes share the same splicing mechanism as Group II introns. The rps12 gene is transpliced, the 5’ end exon is located in the LSC region and the 3’ exon and intron are duplicated and located in the IR regions. This is frequently observed in other angiosperms as well [50,51]. The splicing was also evidenced by mapping of the RNA-seq reads on spliced exons including those from transpliced genes rps12. Conversely, six tRNAs contain introns. In the IRs, the trnI-GAU intron includes the ycf68 pseudogene, and trnA-UGC intron includes the orf42 and orf56 pseudogenes. On the LSC, trnG-UCC, trnV-UAC and trnL-UAA each contain one intron, and the trnK-UUU intron includes the matK gene. The matK encodes for a maturase which is not able to promote intron mobility due to the absence of the reverse transcriptase domain, and the trnL-UAA contains a group I intron. This genomic organization is highly conserved in the U. gibba, U. macrorhiza, G. margaretae and P. ehlersiae.

In summary, 68,969 bp (49.4%) of the cp genome is made up of coding DNA, whereas 57,159 bp (40.9%) 2,766 bp (2%), and 9,044 bp (6.4%) corresponds to CDS, tRNA and rRNAs, respectively. Introns represent 16,811 bp (12%), pseudogenes 6,453 bp (4.6%), and the remaining 47,492 bp (34%) of the genome is made up of non-coding and intergenic spacers.

Transcription evidence of the plastidial genes

Due to its endosymbiotic origin, the chloroplast retains a prokaryotic biochemistry, with its own gene expression machinery. However, the expression of cp genes in many land plants requires different nuclear-encoded proteins, mostly to bind transcripts and regulate translation [52]. Conversely, transcription of some genes is regulated by light-dark cycles or nutrient availability, such as those from photosystem I and II, and rbcL, and the results presented here show that U. reniformis follows this trend. Moreover, the chloroplast RNAs shows a poly(A) tail, which may target the cp RNAs for rapid decay [53]. In order to explore the U. reniformis cp genome expression, cDNA libraries from three different tissues (leaves, stolon, and utricules) were created and sequenced (RNAseq).

The libraries were merged, and a total of 113,224 (7%, out from 1,632,156) reads were uniquely mapped to the annotated cp genes, and the expression profiles of the cp genes were investigated. Overall, >95% of the coding regions and pseudogenes have at least one read of coverage (average base coverage of 183x and median of 33x). The remaining reads (1,087,926; 65.5%) map to the cp rRNAs, such as the 23S rRNA, and also to the tRNAs, and 431,006 (27.5%) reads are non-specific matches. These results show that the cp genome is transcriptionally active, and that all predicted genes, including the pseudogenes are transcribed at some level (Fig 2 and S3 Table). The transcription of genes from photosystem I and II are highly expressed compared to the other genes. In particular the psaJ and psbA genes show the highest expression levels as measured in RPKM (323,421 and 437,421, respectively; S3 Table), and were removed from Fig 2 for brevity and clarity. Moreover, the genes psaC, psaI, psbB, psbC, petB, atpA, atpH, rpl2, clpP, and rbcL show expression near or above 20,000. The least expressed gene is the petG, which encodes the cytochrome b6/f complex subunit V,with expression value of 77, and only one RNAseq read mapped (covering >95% of petG with 99% of identity). Other genes with low RPKM expression values are: psbF (15 transcripts), psbI (11 transcripts), psbJ (12 transcripts), psbK (7 transcripts), psbT (6 transcripts), petL (2 transcripts), rpoB (409 transcripts), rpoC1 (246 transcripts), rpoC2 (566 transcripts), rpl23 (11 transcripts), rpl32 (5 transcripts), ycf15 (11 transcripts) and ycf2 (575 transcripts)(More details in S3 Table).

Fig 2. Expression profile of the Utricularia reniformis cp genes.

The expression values are normalized in Reads Per Kilobase per Million mapped reads (RPKM) on the Y-axis. The psaJ and psbA genes show the highest expression levels as measured in RPKM (323,750 and 437,421, respectively) and were suppressed from the figure for clarity.

Another remarkable feature observed is related to the transcriptional activity of pseudogenes. Interestingly, all ndh complex genes have some sort of mutation, which lead to stop codons in-frame, frameshift and deletions (detailed analysis in the next sections). However, RNAseq reads were mapped to each of the ndh genes (≥98% identity considering 100% of the read length), thus suggesting that these pseudogenes are transcribed. Moreover the ycf68, orf42 and orf56 pseudogenes also show expression values in RPKM. The ycf68 gene has a frameshift and one stop codon in-frame, whereas orf42 has one stop codon in-frame, and orf56 has a frameshift. The ycf68 gene encodes a functional protein present in grasses, gymnosperm and Nymphaeales, whereas orf42 and orf56 are commonly found in the chloroplast genomes of the other species [54,55]. However, orf42 and orf56 are related to mitochondrial genes [55], showing a considerable sequence similarity, and therefore the expression results observed in both genes should not be fully considered due to non-specific and misleading alignments, and therefore they are not shown in Fig 2. Comparative analysis did not detect copies of the ndh, orf42, orf56 and ycf68 genes in nuclear or mitochondrial scaffolds, suggesting that these are really lost from U. reniformis, and thus indicating that these represent real transcripts from a pseudogene template. Interestingly, previous studies indicate that truncated, transcribed molecules may exist in the chloroplast [56], and may support these findings. However, whether these transcripts encode stable RNAs, or even if they are translated to functional proteins is yet to be established, and their roles remain unknown.

Overall, these results indicate that essential cp genes are expressed. However, whether these pseudogenes are not completely inactivated yetor if they encoding regulatory RNAs, orifthe photosystem is impaired or not, due to the loss and pseudogenization of ndh complex, and if some sort of ndh functional replacement could or is occurring remains unknown.

Identification and prediction of RNA editing sites

RNA editing is a post-transcriptional modification that normally changes a cytosine (C) to a uracil (U) or U to C nucleotides, producing transcripts that are different from their DNA template [57]. These modifications can alter the amino-acid sequence of protein, and can also introduce new start and/or stop codons [57]. At least 44 potential editing sites were identified using PREP-Cp and RNAseq analysis. These are all in the coding region, whereas six editing sites were exclusively detected by RNAseq data corresponding to potential novel editing sites (Tables 3 and 4). The editing level from RNAseq data was inferred from the C versus U ratio of the transcripts derived from the respective loci. Among the RNAseq detected sites, three editing sites alters the first and the second nucleotide of a codon, and one editing site alters the third nucleotide of a codon. Except for rpoA, all editing sites lead to non-synonymous substitutions. Four sites located in rpoB, rps14, petB and rps2 are the most frequent editing sites (> 81%), whereas the remaining sites located in atpA, psaB and rpoA were edited at low frequency (11 to 20%) (Table 3). Moreover, the PREP-Cp predicted 37 additional sites, whereas one site located on the rpoB gene was confirmed by RNAseq data (Table 3). All predicted sites by PREP-Cp lead to non-synonymous substitutions. Overall, except for the six editing site detected by RNAseq, the RNA editing sites are quite similar to those observed in other angiosperms [58]. Therefore, these results suggest that the prediction tools may fail to identify authentic RNA editing sites, and that RNAseq data can be used preferentially.

Table 3. RNA editing pattern in Utricularia reniformis cp genome identified by RNAseq reads alignment.

Table 4. RNA editing pattern in Utricularia reniformis cp genome predicted by PREP-Cp.

Several plastids transcripts require C to U editing, whereas ndh genes contain about 50% of the editing sites of angiosperm plastid transcripts [18]. However, the RNA edition was also observed in pseudogenes, such as the ndhB [58]. Interestingly none of the editing sites were associated with transcripts that align with ndh genes loci, supporting the idea that despite the fact that RNAseq reads align to these genes they may be indeed correspond to non-functional genes.

Microsatellite and other repeats

It was previously described that Lentibulariaceae plastomes carry a large number of chloroplast microsatellite (cpSSR) and small number of repeats longer than 60 bp [15]. The U. reniformis plastome follows this trend, having at least 331 cpSSR ranging from 7 to 179 bp (Fig 3). Among those, mono and di repeats were the most common, representing 86% (284 cpSSRs) and 4% (13 cpSSRs), respectively. No pentanucleotides or hexanucleotides repeats were found, and low frequencies of tri-, and tetra repeats were observed (Fig 3). Among the 284 mononucleotide repeats, only 16 C/G type repeats were found, with all other repeats belonging to the A/T type. Repeat number of mononucleotide motifs ranged from seven (48%) to 15. Furthermore, at least 55 cpSSRs mononucleotide repeats with a length of at least 10 bp were detected. It is noteworthy that the AT-rich di repeats are commonly found in others carnivorous plant plastomes [15]. In general, these results are also quite similar to those observed in Tanaecium tetragonolobum, from the family Bignoniaceae, where 347 cpSSR were identified [59]. The distribution of the mononucleotide, di-repeats, tri-repeat and compound/polymorphic cpSSRs are shown in Table 5, and indicate a similar distribution of cpSSRs to that present in the coding regions of U. gibba, U. macrorhiza, G. margaretae and P. ehlersiae. However, the majority of the cpSSR are located in non-coding/intergenic regions, accounting for up to 197 (60%) occurrences, whereas 38 were polymorphic, representing good regions for the development of cpSSRs molecular markers for population studies and to estimate the relationship between different Lentibulariaceae.

Fig 3. Frequency of SSR motifs found in Utricularia reniformis cp coding and intergenic regions, taking into account sequence complementarities.

Table 5. Distribution of the cpSSRs present in the U. reniformis cp coding regions.

A plethora of forward and palindromic repeats were also identified in the U. reniformis cp genome (Table 6). A total of 24 pairs of repeats (30 bp or longer, and up to 58 bp) were identified. These repeats are spread out over the LSC (42%), IRs (50%) and SSC (8%), and no introns were found to contain repeated elements. These repeats are found predominantly in coding regions (58%), which are not commonly found in other angiosperm lineages [51], but are frequent in other Lentibulariaceae [15]. This may indicate that the Lentibulariaceae cp coding regions are repeat hotspots acting as a source of recombination and rearrangements. For instance, two palindromic repeats were identified within the genes psaC, ndhDΨ and ccsA, suggesting a potential role during the pseudogenization process of ndhD. In addition, the region containing psaC, ndhD and ccsA in U. reniformis, U. gibba, U. macrorhiza, P. ehlersiae and G. margaretae is quite variable in terms of nucleotide and gene composition, and order (see below). Palindromic repeats located on the LSC are identified within the genes accD, rbcL, ycf3, psaA, psaJ, rpl2, rpl33, rps14, rps19 and psaB, and the majority of repeats located on the IRs are associated with the gene ycf2. In summary, the number and distribution of these sequences vary from one species to another. However, comparative analyses indicate that this repeat repertoire is quite similar to those previously observed in the terrestrial forms G. margaretae and P. ehlersieae.

Table 6. Sequence repeats in the cp genome of Utricularia reniformis.

Type, location and size (in bp) of repeated elements (IGS, Intergenic spacer).

Comparative analysis among other Lentibulariaceae

Terrestrial and aquatic forms show distinct SSC organization.

The plastomes of the sequenced Lentibulariaceae are highly conserved in terms of gene synteny. Fig 4 shows comparisons of the SSC region and the IR-LSC and IR-SSC boundaries. In spite of the high level of synteny noted on the LSC/IRb and IRa/LSC junctions, the IRb/SSC, SSC/IRa boundaries and the SSC region itself show distinct organization. Gene deletions, rearrangements, expansions and contractions in the SSC and IR/SSC boundaries of these plants are markedly noted.In the SSC, only the rpl32, trnL, cssA, psaC and rpl15 genes are conserved among the analyzed plants. Moreover, a distinct repertoire for the ndh gene complex is observed in each plastome. Indeed, previous studies have indicated that several mutational hotspots were found in the entire SSC as well as the region around the ndh genes of U. macrorhiza, G. margaretae and P. ehlersiae [15]. This may be one of the evolutionary mechanisms related to the ndh and ycf1 pseudogenization process.

Fig 4. Comparison of the boundaries of the LSC, SSC, and IR regions in the currently available cp genomes of Lentibulariaceae.

The ndhF gene is exclusively present in aquatic species, and only U. gibba carries two copies of the ndhF gene, delimiting the IR/SSC boundaries. It is worth noting that the ndhF genes, which are commonly located on the IR/SSC boundaries in other angiosperms plastomes, are lost in the other Lentibulariaceae terrestrial taxa. The loss of the ndhF is not an exclusive feature of the terrestrial forms, since this gene can be either present or absent in angiosperms [51,60]. However ndhF is often found in Coniferophyta, Filicophyta, Ginkgophyta, Gnetophyta, Lycophyta, Psilophyta and Sphenophyta plastomes [61,62]. Interestingly, the ndhF loss may be related to shifts in the position of the junction of the IR and SSC regions in Orchidaceae [62]. Indeed these shifts may lead to losses due to recombination, as observed in the Lentibulariaceae (Fig 4).

For all the other species, two copies of the ycf1 gene, one larger and other smaller, delimits the IR/SSC boundaries. Interestingly, assuming that there have been no major errors in genome assembly, the ycf1 gene shows different sizes among the analyzed plants, indicating that together with ndhF, these genes may be used as a potential hotspot for the study of the evolution of the IR/SSC junction in the Lentibulariaceae. For instance, in G. margaretae, P. ehlersiae and U. gibba, the larger copy of ycf1 is a pseudogene, whereas all smaller copies are intact genes. In addition, ycf1 is often associated with many rearrangements in other angiosperms [63], and this is indeed observed within the Lentibulariaceae plastomes. Therefore, as observed in other land plants, during the course of evolution, the Lentibulariaceae plastomes displayed rearrangements, deletions and gene loss. Overall, these finding also suggests a correlation of the plant life style with plastome genomic structure. For instance, whereas aquatic forms of Lentibulariaceae tend to maintain larger SSC regions, retaining the ndh complex genes intact, the terrestrial forms have smaller SSC regions and have lost ndh genes.

The Lentibulariaceae plastome gene repertoire varies mainly in the ndh genes.

The gene repertoires of the plastome of the sequenced carnivorous plants (U. gibba, U. macrorhiza, G. margaretae, and P. ehlersiae) are quite similar (Fig 5). The Lentibulariaceae cp core gene repertoire is composed of 69 genes, mostly involved in photosynthesis, energy metabolism, and other housekeeping functions (Fig 5, center). Indeed, this result is similar to the core cp genome of essentially all photosynthetic organisms [52]. The main differences are related to a combination of losses and pseudogenization of ndh genes among the five plastomes. It is worth noting that the ndh gene loss/pseudogenization observed in terrestrial forms are clearly derived from independent events (Table 7), which is corroborated by previous studies [15]. In Genlisea, the genes ndhC, D, F, G, H, J, and K were lost from the plastome, and the genes ndhA, B, E, and I are truncated, whereas the ndhA, D, E, G, H, I, J, and K retain insertion/deletions and frame shifts in Pinguicula[15]. However, U. reniformis shows a different pattern, in that the genes ndhC, F, J and K were lost from the plastome, and the genes ndhA, B, DE, G, H and I reside as truncated pseudogenes. Previous studies indicate that the ndh gene loss in Genlisea and Pinguicula occurred two times independently within Lentibulariaceae [15]. Interestingly, considering only carnivorous plants, the ndh gene complex spans to up to 16kb in aquatic forms, representing about of 10% of the plastome, whereas terrestrial forms vary from 5kb to 10kb (3–7%—Table 7).

Fig 5. Venn diagram showing the full complement of genes present in the sequenced Lentibulariaceae cp genomes (pan genome).

The tRNAs and rRNAs are not included. The numbers below each species represent the unique genes used in the comparison.

Table 7. Distribution and comparisons of the eleven genes in the ndh complex, encoded in aquatic and terrestrial cp genomes from carnivorous plants and the ancestral lineages Sesamum indicum, Tanaecium tetragonolobum and Andrographis paniculata.

Black box represents that the given ndh gene is present and intact. Gray box indicates truncated ndh gene according to the legend. White box indicate that the given ndh gene is absent in the cp genome.

Phylogenomic and phylogenetic analysis

The loss and recovery of the ndh gene complex among the Lentibulariaceae.

Moreover, adding the U. reniformis plastome to more complete phylogenetic analyses, and tracing the possible evolutionary hypotheses according to the presence (functional gene) or absence (truncated or gene in frame-shift) of ndh genes, a more comprehensible evolutionary scenario can be determined. The phylogenetic history resulting from the cp phylogenomic analyses (for further discussion see below), indicates that Lentibulariaceae is a monophyletic family with the Pinguicula clade as a sister group of the Genlisea-Utricularia clade; the topology is also corroborated by previous studies [64,65]. By tracing the evolution of ndh genes in the trees, it was determined that the presence of functional ndhA, ndhC, ndhE, ndhF, ndhG, ndhH, ndhI, ndhJ, and ndhK is plesiomorphic, but were lost by pseudogenization or frame-shift in the ancestors of the Lentibulariaceae clade (Fig 6A). However, those ndh genes were recovered in a reversion process for the aquatic group, formed by Utricularia gibba and U. macrorhiza. A similar situation may have occurred with the ndhB gene, but this gene is functional for Pinguicula (Fig 6B). Even assuming the presence of this gene in the ancestral lineage, two possibilities can be explored by optimizing the transformations in the tree, and having both accepted with the parsimony approach (ACCTRAN and DELTRAN—[66]) since both hypotheses assume the same number of transformations (two in this case). Therefore, it is possible to accept the alternative scenario of the loss of ndhB gene from the ancestors of the Genlisea-Utricularia clade, or independent (parallel) losses for Genlisea and Utricularia reniformis (Fig 6B). For the same reasons, it is necessary to assume two alternative histories for the ndhD gene as well. The ndhD is functional for the aquatic species, Utricularia gibba and U. macrorhiza. It exists as a pseudogene for Pinguicula and U. reniformis, but as a frame-shift for Genlisea (Table 7). After surveying all Lamiales taxa represented in the phylogenomic analyses (Fig 6), we found that most species present a functional version of ndhD (Tanaecium tetragonolobum as an exception), thus it is reasonable to suppose this state as plesiomorphic. Assuming the topology as presented in Fig 6C, the ndhD gene can be lost independently more than once. In the present scenario, ndhD was lost for the Tanaecium and Lentibulariaceae clades, and afterwards reverted in the ancestors of the aquatic species Utricularia gibba and U. macrorhiza (Fig 6C), similar to the other ndh genes (Fig 6A and 6B). There is also the possibility that ndhD was lost in the ancestors of the Tanaecium-Andrographis-Pinguicula-Genlisea-Utricularia clade, and reverted twice for the Andrographis and U. gibba—U. macrorhiza clades. Both of these hypotheses for the ndhD gene assume three events, thus both are plausible.

Fig 6. Phylogenetic history of ndh genes.

(A) represents the only scenario for the ndhA, ndhC, ndhE, ndhF, ndhG, ndhH, ndhI, ndhJ, and ndhK genes evolution. (B) represents the phylogenetic history and the two possible scenarios for the ndhB gene and (C) for the ndhD gene (blue bars indicate the arising of the functional genes and red bars their loss).

Relationships with the Lamiales order.

The relationships between the families in the order Lamiales are only partially resolved [67]. Despite the attempts based on several plastomes and mitochondrial genes [67,68] to identify the sister family of Lentibulariaceae, this issue still remains unclear. Therefore, phylogenomic analyses based on whole plastomes might contribute significantly to the elucidation of the systematic relationships inside of this order. With the aim to identify the phylogenetic position of U. reniformis and Lentibulariaceae among the other families of Lamiales, plastomes from 23 different taxa were compared. The phylogenomic analyses were constructed using 47 genes from the Lamiales plastomes. All Lamiales with less than 47 genes were excluded from our analysis, including parasitic plant cp genomes (truncated/pseudogenes and tRNAs were not considered). Both maximum likelihood (ML) and Bayesian analyses recovered the same tree topology with high support values (Fig 7), and agree with a previous study based on the rapidly evolving genes trnK/matK, trnL-F and rps16 [68]. Our results indicate that Lentibulariaceae is monophyletic (100 maximum likelihood bootstrap—BML; 100 Bayesian posterior probability—PP) and Pinguicula is a sister clade of the Genlisea-Utricularia clade (100 BML; 100 PP), corroborating previous studies [65,69]. In addition, that U. reniformis is a sister to the U. gibba-U. macrorhiza clade, is also well supported (100 BML; 100 PP). According to our results based on matK, and also corroborated by previous studies [64,65], the ancestral lineage of Lentibulariaceae was possibly terrestrial, with life forms adapted for this environment developed by most species of Pinguicula and Genlisea (Fig 8). The alternative life forms present in Utricularia species [1,70] are thus derived from the ancestral terrestrial state, representing the occupation of different environments and the consequent diversification of body forms in an adaptive response to several ecological niches. Very specialized alternative life forms were developed by Utricularia lineages, for instance the rare reophytes, which inhabit rapids, cascades and streams at flood level during torrential conditions. For Utricularia this life form is represented only by the four species U. neottioides, U. oliveriana, U. rigida, and U. tetraloba [1], which have at least two independent phylogenetic origins (Fig 8, black circles 4 and 4’). Most aquatic species are represented by Utricularia sect. Utricularia(Taylor 1989), despite this life form having arisen at least twice (Fig 8, blue circles 2 and 2’), with species found in both the Northern and Southern Hemispheres (including U. gibba and U. macrorhiza). Utricularia reniformis is nested within a very specialized clade (Fig 8, green circle 3), with species adapted for terrestrial (the case of U. reniformis) and also epiphytic life forms, represented by species from sections Iperua and Orchidioides. The sister species of U. reniformis is U. nelumbifolia ([69]; this study), a rare endemic Brazilian Utricularia, very similar morphologically to U. reniformis, but which lives among the leaves of the giant bromeliads found on inselbergs [71]. These results also support positioning the Lentibulariaceae close to the Acanthaceae, Bignoniaceae, Pedaliaceae, Orobanchaceae, and Lamiaceae (Fig 7), composing the higher core of the Lamiales group previously proposed [68].

Fig 7. Phylogenomic analysis based on 23 complete Lamiales cp genomes.

The phylogenetic position of Utricularia reniformis was inferred by maximum likelihood and Bayesian analyses. Numbers above and below the lines indicate the maximum likelihood bootstrap values and Bayesian posterior probabilities values > 50% for each clade, respectively. The table on the right indicates the genome features in base pairs (chloroplast genome length, LSC, SSC and IRs regions). The histograms located to the left of each feature (CP size, LSC, SSC and IRs), graphically illustrate the size distribution for each feature. Sub-titles: I Lentibulariaceae; II Acanthaceae; III Bignoniaceae; IV Pedaliaceae; V Orobanchaceae; VI Lamiaceae; VII Scrophulariaceae; VIII Gesneriaceae, and IX Oleaceae.

Fig 8. Cloudgram (Bayesian inference) of Lentibulariaceae from 18,000 Bayesian trees based on matK cp gene.

The red circle 1 indicates the terrestrial ancestral lineage for the Lentibulariaceae family. The blue circles, 2 and 2’, represent the independent radiations to the aquatic habitat of Utricularia lineages. The green circle 3 indicates the possible ancestor of the epiphytic species from the plesiomorphic terrestrial life form, and the black circles, 4 and 4’, represent the independent origins for the rare reophytic life form for Utricularia lineages. Numbers below clades represent the support (maximum parsimony bootstrap/ maximum likelihood bootstrap/ posterior probabilities based on Bayesian inference). Higher color densities represent higher levels of certainty represented by congruent trees (from the 18,000 trees).

In general, the order Lamiales maintains the quadripartite structure, except for the parasitic family of Orobanchaceae that was not included in our analysis due to the small genome size (45kb to 120kb) and number of genes (21 to 42) [22]. In general the SSC and IR are 17 and 25kb long, respectively and only a few differences in genome size are observed (Fig 7). The biggest cp genome belongs to the family Oleaceae. The Jasminum nudiflorum cp genome is 165 kb with an IR and LSC expansion followed by a contraction in the SSC. This is also observed in Schwalbea americana from the Orobanchaceae family, which is 160kb long. In addition, SSC contraction and IR expansion were observed in the Lathraea squamaria from the Orobanchaceae family. However, major differences were noted in the Lentibulariaceae; SSC contraction was mainly observed in Utricularia-Genlisea and Pinguicula (Fig 7). This suggests that this family is under different selective pressures, resulting in dynamic plastome structures. It is noteworthy that only the Lentibulariaceae are carnivorous, suggesting their carnivorous syndrome may impact metabolism and photosynthesis.


The U. reniformis plastome contribution to the study of the evolution of terrestrial carnivorous plants from the Lentibulariaceae family

We sequenced the cp genome of Utricularia reniformis and compared it against other carnivorous plants from the Lentibulariaceae family. This study revealed that the 139kbp cp genome of U. reniformis is quite similar to the cp genome of U. gibba, U. macrorhiza, G. aurea, and P. ehlersiae in terms of gene synteny, repeats and cpSSRs content; whereas the main differences are located on the SSC region and the ndh genes repertoire (Figs 3 and 5 and Tables 1, 2, 5 and 6). In spite of the similarity of the gene repertoire of the U. reniformis cp genome to essentially all photosynthetic organisms, comparative genomics analysis corroborated previous studies [15], which show that whereas aquatic forms maintain the complete ndh gene complex composed of eleven genes, the terrestrial forms have shown a number of losses of the ndh genes, and these losses are exclusive for each species (Table 7). In addition, the proposed phylogenetic history of ndh genes shown in Fig 6, suggests that independent ndh losses occurred during the course of the evolution of the genera; whether other terrestrial Utricularia, Genlisea, and Pinguicula species have also lost the ndh gene set, and whether the ndh pseudogenes found in the terrestrial forms were lost recently, remains to be investigated. Indeed, this is an important question to be explored in future work. Moreover, phylogenomic analysis supports that the family Lentibulariaceae is monophyletic, belonging to the higher core of the Lamiales clade, and thus corroborating the hypothesis that the first Utricularia lineage emerged in terrestrial habitats and then evolved to epiphytic and aquatic forms, as shown by the Fig 8.

Furthermore, the transcriptome analysis by RNAseq approach indicate that mostly cp genes are transcribed (Fig 2 and S3 Table), whereas even the truncated ndh genes, orf42, orf56 and ycf68 show some level of transcription. In addition to the previous observation that truncated transcribed molecules may exist in the chloroplast [56], this finding supports that the pervasive transcription, which is commonly found in bacterial genomes, may also occur in cp genomes, thus suggesting that these transcripts have an important role in gene regulation and genome evolution, as previous discussed elsewhere [72]. However, further studies are necessary to uncover the potential role of these transcripts. In addition, this study also shed some light on the RNA editing in cp genomes, with novel editing site being uncovered (Table 3), suggesting that the methodology used in this study represent a powerful tool to identify novel RNA editing sites.

Endosymbiotic cp gene transfer to the nuclear and mitochondrial genome of U. reniformis

It was well known that during the course of evolution cp genes can be transferred to the nucleus, and their protein products can be reimported into the organelle lumen by the action of a transit peptide [20]. This indeed is a very widespread phenomenon in nature [73]. In addition, gene transfers from organelles often lead to functional replacement of host genes in a process called endosymbiotic gene replacement [21]. For instance, a large chunk of endosymbiotic cp genome was observed on chromosome 10 of rice, which contains a recent 33 kb insertion of cp DNA in addition to a 131 kb insertion representing nearly the entire plastid genome [74]. Furthermore, endosymbiotic gene transfer is also observed in smaller plant genomes, such as A. thaliana [75]. In our ongoing analysis of the U. reniformis genome we have noticed the presence of an endosymbiotic gene transfer of truncated cp genes to the nuclear and mitochondrial genomes. For instance, a total of 26kbp of cp-derived sequences were assembled in 28 contigs and mapped to mt and nuclear DNA assembled scaffolds (S2 Table). In addition to these contig-derived regions, during the ongoing assembly of the U. reniformis mt genome, we have found a truncated copy of thendhJ, ndhK and ndhC genes (data not shown). Interestingly the ndhJ, ndhK and ndhC genes are absent from the U. reniformis cp genome (shown in details in the Fig 1 and Table 1). Indeed, this suggests that an ancient lineage of U. reniformis had these genes in their cp genome, which were subsequently transferred to the mt genome by an endosymbiotic event. However, due to evolutionary pressures, yet to be established, the ndhJ, ndhK and ndhC genes were decayed from the cp copies, and remained as relics in the mt genome. These observations also suggest that during the course of the evolution of the ndh complex in U. reniformis, endosymbiotic gene replacement events from the mt ndhJ, ndhK and ndhC copies, may have occurred. Further investigation is needed based on the sequencing of new species, and the presence and absence of the of ndh genes of others carnivorous plant cp and mt genomes. Therefore, the endosymbiotic gene transfer events are shaping the U. reniformis nuclear and mitochondrial genomes.

A detailed analysis of the U. reniformis nuclear and mt genomes, including functional annotation and comparative genomics showing the endosymbiotic transfer eventsbetween the organelles and the nuclear DNA are in progress.

The impact of the carnivorous syndrome in the ndh complex genes evolution

It was previously observed that lineages that have lost photosynthetic function, trend toward reduced cp genome size [52]. The analysis of the cp genomes of the terrestrial forms, Utricularia reniformis, Genlisea margaretae and Pinguicula ehlersiae, may support this observation. However, U. reniformis, G. margaretae and P. ehlersiae are all photosynthetic organisms that lack the ndh genes, which apparently does not affect the fitness of these plants. Indeed, previous studies suggested that the ndh function might be dispensable under favorable growth conditions [19], suggesting that the carnivorous syndrome may act in favor of the functional ndh absence. Interestingly, the ndh gene loss or pseudogenization is relatively rare among the Viridiplantae clade [17,19]. However, it seems that the ndh genes were not essential during plant evolution, and their loss may also be related to early events leading to parasitic behavior [18]. In addition, the ndh genes are related to photosynthetic response to environmental stress, indicating its participation in the transition to terrestrial habitats [18,19]. However the first lineages of the Lentibulariaceae emerged in a terrestrial habitat, and then evolved to aquatic environments, suggesting that the evolutive history of the ndh genes among the Lentibulariaceae followed an opposite direction. For instance, we propose that the plastid ndh genes present in the aquatic forms U. gibba and U. macrorhiza were recovered in a reversion process, and that the ndh function may be dispensable in terrestrial forms (Fig 6).

In order to explain this genomic trend related to the loss of the ndh genes observed in the terrestrial Lentibulariaceae, a hypothesis posits that carnivorous plants are metabolically similar to parasitic plants in that they use organic carbon obtained through their prey, or their host for parasitic plants, to overcome environmental stress [15]. In addition, different and variable levels of nutritional stress to the plant may occur in aquatic, terrestrial, epiphytic and reophytic forms. This hypothesis is quite interesting, since it suggests that nutritional stress, which is a common feature of the carnivorous plants [2], can impact molecular and biochemistry characteristics shaping the cp genome. Moreover, over an evolutionary time scale these differences can lead to morphological changes, such as the adaptation to aquatic or land environments, and thus supporting the ndh gene repertoire differences observed among these species (Fig 8). However this hypothesis is yet to be established.

Conversely, it has been proposed that under relaxed selection, unequal efficiency of DNA repair, and high levels of mutagenic reactive oxygen species (ROS), the genome architecture of the Lentibulariaceae may also have been shaped in a fashion similar as those observed in the parasitic plants [9,14,15]. Indeed, a previous study has shown a genome-level convergence between carnivorous and parasitic plants [22]. Moreover, the ndh loci have accumulated several nucleotide substitutions and repeats [15], which may have resulted in the loss and pseudogenization process observed in U. reniformis, G. margaretae and P. ehlersiae. Indeed the sequence repeats are located mostly in the coding regions, and this is particularly noted with ndhDΨ gene in U. reniformis (Table 6). However the sequencing of additional terrestrial and aquatic forms is necessary to corroborate the role of the sequence repeats with the pseudogenization process.

Overall, we propose that sequencing of additional cp and nuclear genomes from other individuals and species from the Lentibulariaceae family will shed light on the relationships between the rearrangement and loss of ndh genes, life style (aquatic, terrestrial, epiphytic and reophytic) and endosymbiotic gene transfer of cpDNA. Indeed, a recent study has shown that the endosymbiotic gene transfer has also occurred in other carnivorous plants, such as the Genlisea nigrocaulis and G. hispidula genomes [76], thus suggesting that this may be an evolutionary trend. Whether the ndh genes loss in terrestrial forms and the endosymbiotic gene transfer is an evolutionary trend of this group, which is leading to biochemistry and plastome-scale convergence with the parasitic plants, remains as an important question to be answered in the near future.

Supporting Information

S1 Table. Phylogenomic and phylogenetic analysis.

(A) List of the chloroplast genomes used in the phylogenomics analysis with their respective GenBank accession number. (B) List of chloroplast genes used in the phylogenomics analysis. (C) The matK genes used in the phylogenetic analysis with their respective GenBank accession number. (D) The matK genes generated in this study and used in the phylogenetic analysis with their respective GenBank accession number.


S2 Table. The remaining paired-end reads (2x300bp; 229,830; 18.3%) which were assembled into contigs containing fragments of incomplete or truncated cp genes.

The distribution, number of these contigs, truncated gene content and alignment to the Utricularia reniformis cp genome is shown.


S3 Table. RNAseq experiment table, showing the expression profile of all chloroplast related genes of Utricularia reniformis.



We would like to thank Dr. Cristine G. Menezes for the plant sample preparation, and all the members of the Plant Systematics Laboratory, UNESP-FCAV. We also would like to thank Dr. Lubomír Adamec who kindly provided the Utricularia samples used in the cloudogram analysis, and to the anonymous reviewers for their contributions to this manuscript.

Author Contributions

  1. Conceptualization: SRS VFOM TPM AMV.
  2. Data curation: SRS DGP AMV.
  3. Formal analysis: SRS VFOM AMV.
  4. Funding acquisition: AMV.
  5. Investigation: SRS DGP VFOM AMV.
  7. Project administration: VFOM AMV.
  8. Resources: SRS YCAD HAP DGP VFOM AMV.
  9. Software: SRS DGP AMV.
  10. Supervision: VFOM TPM AMV.
  11. Validation: SRS YCAD HAP DGP CCF AMV.
  12. Visualization: SRS VFOM AMV.
  13. Writing – original draft: SRS VFOM TPM AMV.
  14. Writing – review & editing: SRS VFOM TPM AMV.


  1. 1. Taylor P. Genus Utricularia: a taxonomic monograph. 2nd edition. Royal Botanic Gardens, Kew; 1989.
  2. 2. Juniper B, Robins R, Joel D. The Carnivirorous Plants. Academic Press, London; 1989.
  3. 3. Reifenrath K, Theisen I, Schnitzler J, Porembski S, Barthlott W. Trap architecture in carnivorous Utricularia (Lentibulariaceae). Flora—Morphol Distrib Funct Ecol Plants. 2006;201: 597–605.
  4. 4. Fleischmann A, Michael TP, Rivadavia F, Sousa A, Wang W, Temsch EM, et al. Evolution of genome size and chromosome number in the carnivorous plant genus Genlisea (Lentibulariaceae), with a new estimate of the minimum genome size in angiosperms. Ann Bot. 2014;114: 1651–1663. pmid:25274549
  5. 5. Pellicer J, Fay MF, Leitch IJ. The largest eukaryotic genome of them all? Bot J Linn Soc. 2010;164: 10–15.
  6. 6. Michael TP. Plant genome size variation: bloating and purging DNA. Brief Funct Genomics. 2014;13: 308–317. pmid:24651721
  7. 7. Greilhuber J, Borsch T, Müller K, Worberg A, Porembski S, Barthlott W. Smallest angiosperm genomes found in lentibulariaceae, with chromosomes of bacterial size. Plant Biol Stuttg Ger. 2006;8: 770–777. pmid:17203433
  8. 8. Kelly LJ, Renny-Byfield S, Pellicer J, Macas J, Novák P, Neumann P, et al. Analysis of the giant genomes of Fritillaria (Liliaceae) indicates that a lack of DNA removal characterizes extreme expansions in genome size. New Phytol. 2015;208: 596–607. pmid:26061193
  9. 9. Ibarra-Laclette E, Lyons E, Hernández-Guzmán G, Pérez-Torres CA, Carretero-Paulet L, Chang T-H, et al. Architecture and evolution of a minute plant genome. Nature. 2013; pmid:23665961
  10. 10. Clivati D, Gitzendanner MA, Hilsdorf AWS, Araújo WL, Oliveira de Miranda VF. Microsatellite markers developed for Utricularia reniformis (Lentibulariaceae). Am J Bot. 2012;99: e375–378. pmid:22947485
  11. 11. Clivati D, Cordeiro GD, Płachno BJ, de Miranda VFO. Reproductive biology and pollination of Utricularia reniformis A.St.-Hil. (Lentibulariaceae). Plant Biol. 2014;16: 677–682. pmid:24834508
  12. 12. Carretero-Paulet L, Librado P, Chang T-H, Ibarra-Laclette E, Herrera-Estrella L, Rozas J, et al. High Gene Family Turnover Rates and Gene Space Adaptation in the Compact Genome of the Carnivorous Plant Utricularia gibba. Mol Biol Evol. 2015;32: 1284–1295. pmid:25637935
  13. 13. Carretero-Paulet L, Chang T-H, Librado P, Ibarra-Laclette E, Herrera-Estrella L, Rozas J, et al. Genome-wide analysis of adaptive molecular evolution in the carnivorous plant Utricularia gibba. Genome Biol Evol. 2015;7: 444–456. pmid:25577200
  14. 14. Ibarra-Laclette E, Albert VA, Pérez-Torres CA, Zamudio-Hernández F, Ortega-Estrada M de J, Herrera-Estrella A, et al. Transcriptomics and molecular evolutionary rate analysis of the bladderwort (Utricularia), a carnivorous plant with a minimal genome. BMC Plant Biol. 2011;11: 101. pmid:21639913
  15. 15. Wicke S, Schäferhoff B, dePamphilis CW, Müller KF. Disproportional Plastome-Wide Increase of Substitution Rates and Relaxed Purifying Selection in Genes of Carnivorous Lentibulariaceae. Mol Biol Evol. 2014;31: 529–545. pmid:24344209
  16. 16. Braukmann TWA, Kuzmina M, Stefanović S. Loss of all plastid ndh genes in Gnetales and conifers: extent and evolutionary significance for the seed plant phylogeny. Curr Genet. 2009;55: 323–337. pmid:19449185
  17. 17. Peredo EL, King UM, Les DH. The Plastid Genome of Najas flexilis: Adaptation to Submersed Environments Is Accompanied by the Complete Loss of the NDH Complex in an Aquatic Angiosperm. PLoS ONE. 2013;8. pmid:23861923
  18. 18. Martín M, Sabater B. Plastid ndh genes in plant evolution. Plant Physiol Biochem PPB Société Fr Physiol Végétale. 2010;48: 636–645. pmid:20493721
  19. 19. Ruhlman TA, Chang W-J, Chen JJ, Huang Y-T, Chan M-T, Zhang J, et al. NDH expression marks major transitions in plant evolution and reveals coordinate intracellular gene loss. BMC Plant Biol. 2015;15: 100. pmid:25886915
  20. 20. Martin W. Gene transfer from organelles to the nucleus: Frequent and in big chunks. Proc Natl Acad Sci. 2003;100: 8612–8614. pmid:12861078
  21. 21. Brown JR. Ancient horizontal gene transfer. Nat Rev Genet. 2003;4: 121–132. pmid:12560809
  22. 22. Wicke S, Müller KF, de Pamphilis CW, Quandt D, Wickett NJ, Zhang Y, et al. Mechanisms of functional and physical genome reduction in photosynthetic and nonphotosynthetic parasitic plants of the broomrape family. Plant Cell. 2013;25: 3711–3725. pmid:24143802
  23. 23. Miranda VFO, Borges RAX, Hering RLO, Monteiro NP, Santos-Filho LAF. Lentibulariaceae. Livro Vermelho da Flora do Brasil. 1st ed. Instituto de Pesquisas Jardim Botânico do Rio de Janeiro; 2013. pp. 614–615. Available:
  24. 24. Langmead B, Salzberg SL. Fast gapped-read alignment with Bowtie 2. Nat Methods. 2012;9: 357–359. pmid:22388286
  25. 25. Bankevich A, Nurk S, Antipov D, Gurevich AA, Dvorkin M, Kulikov AS, et al. SPAdes: a new genome assembly algorithm and its applications to single-cell sequencing. J Comput Biol J Comput Mol Cell Biol. 2012;19: 455–477. pmid:22506599
  26. 26. Hahn C, Bachmann L, Chevreux B. Reconstructing mitochondrial genomes directly from genomic next-generation sequencing reads—a baiting and iterative mapping approach. Nucleic Acids Res. 2013;41: e129. pmid:23661685
  27. 27. Wyman SK, Jansen RK, Boore JL. Automatic annotation of organellar genomes with DOGMA. Bioinforma Oxf Engl. 2004;20: 3252–3255. pmid:15180927
  28. 28. Camacho C, Coulouris G, Avagyan V, Ma N, Papadopoulos J, Bealer K, et al. BLAST+: architecture and applications. BMC Bioinformatics. 2009;10: 421. pmid:20003500
  29. 29. Laslett D, Canback B. ARAGORN, a program to detect tRNA genes and tmRNA genes in nucleotide sequences. Nucleic Acids Res. 2004;32: 11–16. pmid:14704338
  30. 30. Carver T, Harris SR, Berriman M, Parkhill J, McQuillan JA. Artemis: an integrated platform for visualization and analysis of high-throughput sequence-based experimental data. Bioinforma Oxf Engl. 2012;28: 464–469. pmid:22199388
  31. 31. Magrane M, Consortium U. UniProt Knowledgebase: a hub of integrated protein data. Database. 2011;2011: bar009–bar009. pmid:21447597
  32. 32. Mulder N, Apweiler R. InterPro and InterProScan: tools for protein sequence classification and comparison. Methods Mol Biol Clifton NJ. 2007;396: 59–70.
  33. 33. Lohse M, Drechsel O, Kahlau S, Bock R. OrganellarGenomeDRAW—a suite of tools for generating physical maps of plastid and mitochondrial genomes and visualizing expression data sets. Nucleic Acids Res. 2013;41: W575–581. pmid:23609545
  34. 34. Heberle H, Meirelles GV, da Silva FR, Telles GP, Minghim R. InteractiVenn: a web-based tool for the analysis of sets through Venn diagrams. BMC Bioinformatics. 2015;16: 169. pmid:25994840
  35. 35. Thiel T, Michalek W, Varshney RK, Graner A. Exploiting EST databases for the development and characterization of gene-derived SSR-markers in barley (Hordeum vulgare L.). TAG Theor Appl Genet Theor Angew Genet. 2003;106: 411–422. pmid:12589540
  36. 36. Kurtz S, Choudhuri JV, Ohlebusch E, Schleiermacher C, Stoye J, Giegerich R. REPuter: the manifold applications of repeat analysis on a genomic scale. Nucleic Acids Res. 2001;29: 4633–4642. pmid:11713313
  37. 37. Katoh K, Standley DM. MAFFT multiple sequence alignment software version 7: improvements in performance and usability. Mol Biol Evol. 2013;30: 772–780. pmid:23329690
  38. 38. Posada D, Crandall KA. MODELTEST: testing the model of DNA substitution. Bioinforma Oxf Engl. 1998;14: 817–818.
  39. 39. Akaike H. A new look at the statistical model identification. IEEE Trans Autom Control. 1974;19: 716–723.
  40. 40. Burnham KP, Anderson DR. Multimodel Inference Understanding AIC and BIC in Model Selection. Sociol Methods Res. 2004;33: 261–304.
  41. 41. Stamatakis A. RAxML version 8: a tool for phylogenetic analysis and post-analysis of large phylogenies. Bioinforma Oxf Engl. 2014;30: 1312–1313. pmid:24451623
  42. 42. Huelsenbeck JP, Ronquist F. MRBAYES: Bayesian inference of phylogenetic trees. Bioinforma Oxf Engl. 2001;17: 754–755.
  43. 43. Stöver BC, Müller KF. TreeGraph 2: Combining and visualizing evidence from different phylogenetic analyses. BMC Bioinformatics. 2010;11: 7. pmid:20051126
  44. 44. Bouckaert RR. DensiTree: making sense of sets of phylogenetic trees. Bioinforma Oxf Engl. 2010;26: 1372–1373. pmid:20228129
  45. 45. CBOL Plant Working Group. A DNA barcode for land plants. Proc Natl Acad Sci U S A. 2009;106: 12794–12797. pmid:19666622
  46. 46. Schmieder R, Edwards R. Quality control and preprocessing of metagenomic datasets. Bioinforma Oxf Engl. 2011;27: 863–864. pmid:21278185
  47. 47. Kim D, Pertea G, Trapnell C, Pimentel H, Kelley R, Salzberg SL. TopHat2: accurate alignment of transcriptomes in the presence of insertions, deletions and gene fusions. Genome Biol. 2013;14: R36. pmid:23618408
  48. 48. Mower JP. PREP-Mt: predictive RNA editor for plant mitochondrial genes. BMC Bioinformatics. 2005;6: 96. pmid:15826309
  49. 49. Curci PL, De Paola D, Danzi D, Vendramin GG, Sonnante G. Complete Chloroplast Genome of the Multifunctional Crop Globe Artichoke and Comparison with Other Asteraceae. PLoS ONE. 2015;10: e0120589. pmid:25774672
  50. 50. Yi D-K, Kim K-J. Complete Chloroplast Genome Sequences of Important Oilseed Crop Sesamum indicum L. PLoS ONE. 2012;7: e35872. pmid:22606240
  51. 51. Yao X, Tang P, Li Z, Li D, Liu Y, Huang H. The First Complete Chloroplast Genome Sequences in Actinidiaceae: Genome Structure and Comparative Analysis. PloS One. 2015;10: e0129347. pmid:26046631
  52. 52. Barbrook AC, Howe CJ, Kurniawan DP, Tarr SJ. Organization and expression of organellar genomes. Philos Trans R Soc B Biol Sci. 2010;365: 785–797. pmid:20124345
  53. 53. Lisitsky I, Klaff P, Schuster G. Addition of destabilizing poly(A)-rich sequences to endonuclease cleavage sites during the degradation of chloroplast mRNA. Proc Natl Acad Sci. 1996;93: 13398–13403. pmid:8917603
  54. 54. Raubeson LA, Peery R, Chumley TW, Dziubek C, Fourcade HM, Boore JL, et al. Comparative chloroplast genomics: analyses including new sequences from the angiosperms Nuphar advena and Ranunculus macranthus. BMC Genomics. 2007;8: 174. pmid:17573971
  55. 55. Do HDK, Kim JS, Kim J-H. Comparative genomics of four Liliales families inferred from the complete chloroplast genome sequence of Veratrum patulum O. Loes. (Melanthiaceae). Gene. 2013;530: 229–235. pmid:23973725
  56. 56. Schuster G, Lisitsky I, Klaff P. Polyadenylation and Degradation of mRNA in the Chloroplast. Plant Physiol. 1999;120: 937–944. pmid:10444076
  57. 57. Lin C-P, Ko C-Y, Kuo C-I, Liu M-S, Schafleitner R, Chen L-FO. Transcriptional Slippage and RNA Editing Increase the Diversity of Transcripts in Chloroplasts: Insight from Deep Sequencing of Vigna radiata Genome and Transcriptome. PloS One. 2015;10: e0129396. pmid:26076132
  58. 58. Zeng W-H, Liao S-C, Chang C-C. Identification of RNA Editing Sites in Chloroplast Transcripts of Phalaenopsis aphrodite and Comparative Analysis with Those of Other Seed Plants. Plant Cell Physiol. 2007;48: 362–368. pmid:17169923
  59. 59. Nazareno AG, Carlsen M, Lohmann LG. Complete Chloroplast Genome of Tanaecium tetragonolobum: The First Bignoniaceae Plastome. PloS One. 2015;10: e0129930. pmid:26103589
  60. 60. Zhang T, Fang Y, Wang X, Deng X, Zhang X, Hu S, et al. The Complete Chloroplast and Mitochondrial Genome Sequences of Boea hygrometrica: Insights into the Evolution of Plant Organellar Genomes. PLoS ONE. 2012;7: e30531. pmid:22291979
  61. 61. Neyland R, Urbatsch LE. The ndhF chloroplast gene detected in all vascular plant divisions. Planta. 1996;200: 273–277. pmid:8904810
  62. 62. Kim HT, Kim JS, Moore MJ, Neubig KM, Williams NH, Whitten WM, et al. Seven New Complete Plastome Sequences Reveal Rampant Independent Loss of the ndh Gene Family across Orchids and Associated Instability of the Inverted Repeat/Small Single-Copy Region Boundaries. PLoS ONE. 2015;10: e0142215. pmid:26558895
  63. 63. Li R, Ma P-F, Wen J, Yi T-S. Complete Sequencing of Five Araliaceae Chloroplast Genomes and the Phylogenetic Implications. PLoS ONE. 2013;8: e78568. pmid:24205264
  64. 64. Müller K, Borsch T, Legendre L, Porembski S, Theisen I, Barthlott W. Evolution of carnivory in Lentibulariaceae and the Lamiales. Plant Biol Stuttg Ger. 2004;6: 477–490. pmid:15248131
  65. 65. Müller KF, Borsch T, Legendre L, Porembski S, Barthlott W. Recent progress in understanding the evolution of carnivorous lentibulariaceae (lamiales). Plant Biol Stuttg Ger. 2006;8: 748–757. pmid:17203430
  66. 66. Agnarsson I, Miller JA. Is ACCTRAN better than DELTRAN? Cladistics. 2008;24: 1032–1038.
  67. 67. Refulio-Rodriguez NF, Olmstead RG. Phylogeny of Lamiidae. Am J Bot. 2014;101: 287–299. pmid:24509797
  68. 68. Schäferhoff B, Fleischmann A, Fischer E, Albach DC, Borsch T, Heubl G, et al. Towards resolving Lamiales relationships: insights from rapidly evolving chloroplast sequences. BMC Evol Biol. 2010;10: 352. pmid:21073690
  69. 69. Jobson RW, Playford J, Cameron KM, Albert VA. Molecular Phylogenetics of Lentibulariaceae Inferred from Plastid rps16 Intron and trnL-F DNA Sequences: Implications for Character Evolution and Biogeography. Syst Bot. 2003;28: 157–171.
  70. 70. Raynal-Roques A, Jérémie J. Biologie diversity in the genus Utricularia (Lentibulariaceae). Acta Bot Gallica. 2005;152: 177–186.
  71. 71. Płachno BJ, Stpiczyńska M, Davies KL, Świątek P, de Miranda VFO. Floral ultrastructure of two Brazilian aquatic-epiphytic bladderworts: Utricularia cornigera Studnička and U. nelumbifolia Gardner (Lentibulariaceae). Protoplasma. 2016; pmid:26945989
  72. 72. Wade JT, Grainger DC. Pervasive transcription: illuminating the dark matter of bacterial transcriptomes. Nat Rev Microbiol. 2014;12: 647–653. pmid:25069631
  73. 73. Bock R, Timmis JN. Reconstructing evolution: Gene transfer from plastids to the nucleus. BioEssays. 2008;30: 556–566. pmid:18478535
  74. 74. Rice Chromosome 10 Sequencing Consortium. In-depth view of structure, activity, and evolution of rice chromosome 10. Science. 2003;300: 1566–1569. pmid:12791992
  75. 75. Arabidopsis Genome Initiative. Analysis of the genome sequence of the flowering plant Arabidopsis thaliana. Nature. 2000;408: 796–815. pmid:11130711
  76. 76. Vu GTH, Schmutzer T, Bull F, Cao HX, Fuchs J, Tran TD, et al. Comparative Genome Analysis Reveals Divergent Genome Size Evolution in a Carnivorous Plant Genus. Plant Genome. 2015;8: 0.