Abstract

   Yellow to red colored betalains are a chemotaxonomic feature of
   Caryophyllales, while in most other plant taxa, anthocyanins are
   responsible for these colors. The carnivorous plant family Nepenthaceae
   belongs to Caryophyllales; here, red‐pigmented tissues seem to attract
   insect prey. Strikingly, the chemical nature of red color in Nepenthes
   has never been elucidated. Although belonging to Caryophyllales, in
   Nepenthes, some molecular evidence supports the presence of
   anthocyanins rather than betalains. However, there was previously no
   direct chemical proof of this. Using ultra‐high‐performance liquid
   chromatography‐electrospray ionization‐high‐resolution mass
   spectrometry, we identified cyanidin glycosides in Nepenthes species
   and tissues. Further, we reveal the existence of a complete set of
   constitutively expressed anthocyanin biosynthetic genes in Nepenthes.
   Thus, here we finally conclude the long‐term open question regarding
   red pigmentation in Nepenthaceae.

   Keywords: anthocyanins, betalain, Caryophyllales, Nepenthes, plant
   carnivory
     __________________________________________________________________

   Red colors are widely distributed in tissues of carnivorous Nepenthes
   plants. However, whether these colors are due to the presence of
   anthocyanins or betalains remained unclear. Here, we employed
   ultra‐high‐performance liquid chromatography‐electrospray
   ionization‐high‐resolution mass spectrometry and identified different
   cyanidin glycosides in Nepenthes species and tissues. In addition, the
   existence of a complete set of constitutively expressed anthocyanin
   biosynthetic genes in Nepenthes is shown.

   graphic file with name FEB4-11-2576-g003.jpg
     __________________________________________________________________

Abbreviations

   ADH
          arogenate dehydrogenase

   ANS
          anthocyanidin synthase

   C3′H
          5-O-(4-coumaroyl)-D-quinate 3′-monooxygenase

   CHI
          chalcone isomerase

   CHS
          chalcone synthase

   DFR
          dihydroflavonol 4-reductase/flavonone 4-reductase

   DODA
          DOPA 4,5-dioxygenase

   DOPA
          3,4-dihydroxyphenylalanine

   ESI-MS/MS
          electrospray ionization with tandem mass spectrometry

   F3′5′H
          flavonoid 3′,5′-hydroxylase

   F3′H
          flavonoid 3′-monooxygenase

   F3H
          flavanone 3-dioxygenase

   HCT
          shikimate O-hydroxycinnamoyl transferase

   UFGT
          anthocyanidin 3-O-glucosyltransferase

   UHPLC–ESI–HRMS
          ultra-high-performance liquid chromatography-electrospray
          ionization-high-resolution mass spectrometry

   The presence of betalains is a typical phytochemical feature of the
   plant order Caryophyllales [[32]1]. Betalains are violet to red
   (betacyanins) and orange to yellow (betaxanthins) pigments that are
   derived from the amino acid tyrosine. According to the Angiosperm
   Phylogeny Group classification [[33]2], the occurrence of betalains
   holds true for the so‐called core Caryophyllales, a defined clade of
   eudicots comprising c. 29 families. Only two Caryophyllales families,
   Caryophyllaceae and Molluginaceae, do not contain betalains but
   anthocyanins, also red and yellow pigments much more widely distributed
   in the plant kingdom [[34]3, [35]4]. Within the noncore Caryophyllales,
   betalains have never been documented. Strikingly, the presence of
   betalains and anthocyanins exclude each other [[36]1, [37]3, [38]5,
   [39]6]. Very likely, this is due to two events in the core
   Caryophyllales, the de‐regulation of the tyrosine biosynthesis and gene
   duplication events [[40]3, [41]6, [42]7]. While betalains are made from
   tyrosine, anthocyanins are made from phenylalanine. In both cases, the
   shikimate pathway provides the biosynthetic precursors; that is, the
   syntheses of tyrosine and phenylalanine compete for prephenate or
   arogenate as substrate. Whereas during tyrosine synthesis in bacteria
   and fungi prephenate is converted to 4‐hydroxyphenyl pyruvate, in
   plants prephenate is mainly converted to arogenate. Arogenate is
   further converted by arogenate dehydrogenases (ADH) into tyrosine.
   Typically, the ADH is negatively feedback‐regulated by tyrosine;
   however, in betalain accumulating species this regulation is partly
   lost. Here, a tyrosine‐insensitive ADH arose during evolution of the
   betalain synthetic pathway in the core Caryophyllales that accumulate
   high amounts of tyrosine and, as a consequence thereof, the substrate
   for betalain synthesis [[43]7]. In addition, two enzymes downstream of
   tyrosine synthesis, CYP76AD1 and a 3,4‐dihydroxyphenylalanine (DOPA)
   4,5‐dioxygenase (DODA), underwent gene duplication and concomitant
   neo‐functionalization. These duplications gave rise to DODA‐α and
   CYP76AD1‐α isoforms, which seem necessary for betalain synthesis; both
   new genes arose shortly before the origin of betalain pigmentation
   [[44]3].

   Nepenthales were classified as a noncore group of Caryophyllales
   [[45]8], which are characterized among other features by lacking
   betalains [[46]9]. Nepenthales cover noncarnivorous as well as
   carnivorous lineages. The latter lineage consists of five plant
   families, including Droseraceae with the genera Drosera (sundew) and
   Dionaea, represented by the only species Dionaea muscipula (Venus
   flytrap), and Nepenthaceae with the genus Nepenthes (pitcher plants).
   Almost all of these carnivorous plants have parts with intensive red
   colors, which are potentially involved in prey attraction. Maybe due to
   the classification into core and noncore Caryophyllales, there is still
   confusion about the nature of the red color in carnivorous plants
   belonging to Caryophyllales. While in Drosera spp. anthocyanins,
   cyanidin and pelargonidin glycosides, have been strongly suggested
   [[47]5, [48]10], for D. muscipula the presence of cyanidin‐3‐glucoside
   was demonstrated already in 1966 by chromatographic and spectroscopic
   (UV, IR) methods in comparison with an authentic standard [[49]11].
   Recently, a combination of spectrophotometry, HPLC co‐elution and
   electrospray ionization with tandem mass spectrometry (ESI‐MS/MS)
   proved the presence of delphinidin‐3‐O‐glucoside (myrtillin),
   cyanidin‐3‐O‐glucoside (kuromanin), and the cyanidin aglycone in
   D. muscipula [[50]12].

   For the genus Nepenthes, the situation is not as clear as for Drosera
   and Dionaea. There is still no evidence for anthocyanin presence in
   Nepenthes. Nevertheless, there are speculations about the nature of the
   red coloration in this genus, but no robust data yet. The optical
   properties of anthocyanins and betalains are very similar, and simple
   UV/Vis absorption measurements at, for example, 532 nm [[51]13] cannot
   discriminate both compounds and does not really justify a statement on
   the pigment's nature. Moreover, some citations are inaccurate and,
   thus, mere suggestions seem to become true the more often a reference
   is cited. For example, Moran and Moran [[52]14] are repeatedly cited
   for the presence of anthocyanins [[53]15, [54]16]. However, with the
   foliar reflectance analysis used in that study it was not possible to
   really prove the presence of anthocyanins as the spectral data of
   anthocyanin and betacyanin are very similar [[55]17]. Also, the study
   of [[56]18] was listed as a reference for anthocyanins [[57]19]
   although they only analyzed phenolic compounds and flavonols.

   Nevertheless, it has been common sense for many years that also
   Nepenthes species contain anthocyanins, not at least due to molecular
   studies [[58]1, [59]3, [60]6, [61]7, [62]20]. Unfortunately, the final
   proof based on reliable chemical analytics as in the case of Dionaea
   [[63]12] is still missing for Nepenthes. Here, we aim to evaluate the
   presence of anthocyanins in Nepenthes tissues by employing sensitive
   analytical techniques, that is, ultra‐high‐performance liquid
   chromatography–ESI–high‐resolution mass spectrometry (UHPLC–ESI–HRMS).
   We detected three different cyanidin derivatives and, moreover, found
   all genes that are necessary for the anthocyanin biosynthetic pathway
   constitutively expressed.

Materials and methods

Plant material

   Nepenthes x ventrata (the natural hybrid of Nepenthes ventricosa ×
   Nepenthes alata), Nepenthes thorelii, N. ventricosa plants were grown
   in the MPI greenhouse at 21–23 °C, 50–60% relative humidity and a 16 h
   light/8 h dark photoperiod. To keep the plants moistened, they were
   sprayed with distilled water for 25 s four times per day. Pitchers from
   Nepenthes robcantleyi, Nepenthes maxima, Nepenthes fusca, and
   Nepenthes mirabilis were provided from the Botanical Garden, Jena,
   Germany.

   Matured and well‐developed pitchers were sampled from different plants
   representing independent biological replicates. Digestive fluid from
   pitcher was discarded, and pitcher was rinsed three times with sterile
   ddH[2]O. Afterward, tissues of interest (peristome, digestive zone,
   leaf, branches) were sampled and directly frozen in liquid N[2].

Extraction and quantification of anthocyanins by HPLC‐UV

   Frozen tissue samples were ground and 100 mg fresh weight powder
   extracted with 1.0 mL ddH[2]O:MeOH (50 : 50 v/v). After mixing, samples
   were sonicated for 15 min on ice‐cold water bath. Therefore, shook for
   30 min at 4 °C using Rotator Mixer RM‐Multi‐1 (STARLAB GmbH, Hamburg,
   Germany) with the following settings: orbital at 100 r.p.m. for 15 s,
   reciprocal at 75° for 16 s, and vibro at 3° for 5 s. Samples were
   centrifuged afterward at 16 000 g at 4 °C for 30 min, and clear
   supernatants were collected and used for further analysis.

   Anthocyanins were analyzed by reversed‐phase HPLC with UV detection
   using an Agilent 1100 system (Agilent Technologies, Waldbronn,
   Germany): column used: Nucleodur Sphinx RP (250 × 4.6 mm, 5 µm;
   Macherey‐Nagel, Düren, Germany); injection volume was 50 μL; flow rate,
   1.0 mL·min^−1; solvent A, 0.5% (v/v) trifluoroacetic acid; solvent B,
   acetonitrile. The photodiode array detector was used in the range of
   250–650 nm. Samples were analyzed with the following chromatographic
   gradient: start 5% B, linear gradient from 5% B to 25% B in 20 min
   followed by a washing cycle. Peaks at 18.1 min and at 18.5 min in the
   HPLC‐UV/Vis chromatograms were identified by match of retention time
   with commercial standards as cyanidin‐3‐O‐galactoside (Extrasynthese,
   Genay, France) and as cyanidin‐3‐O‐glucoside (TransMIT GmbH, Gießen,
   Germany), respectively. Further identification is based on LC‐ESI‐HRMS
   (see below). Quantification was achieved by detection at 520 nm using a
   calibration curve generated from authentic cyanidin‐3‐O‐glucoside.

Identification of anthocyanins by LC‐ESI‐HRMS

   Chemical structures of anthocyanins were determined by UHPLC–ESI–HRMS
   performed with a Dionex Ultimate 3000 series UHPLC (Thermo Scientific,
   Schwerte, Germany) and a Bruker timsToF mass spectrometer (Bruker
   Daltonics, Bremen, Germany). UHPLC was used applying a Zorbax Eclipse
   XDB‐C18 column (100 mm × 2.1 mm, 1.8 µm; Agilent Technologies) with a
   solvent system of 0.1% (v/v) formic acid (A) and acetonitrile (B) at a
   flow rate of 0.3 mL·min^−1. The elution profile was the following:
   0–0.5 min, 5% B; 0.5–11.0 min, 5–60% B; 11.0–11.1 min, 60–100% B,
   11.1–12.0 min, 100% B and 12.1–15.0 min 5% B. ESI in positive
   ionization mode was used for the coupling of LC to MS. The mass
   spectrometer parameters were set as follows: capillary voltage 4.5 KV,
   end plate offset of 500 V, nebulizer pressure 2.8 bar, nitrogen at
   280 °C at a flow rate of 8 L·min^−1 as drying gas. Acquisition was
   achieved at 12 Hz with a mass range from m/z 50 to 1500 with
   data‐dependent MS^2. Fragmentation was triggered at the two most
   intense peaks applying a target intensity of 20 000 counts, with MS^2
   spectra acquisition at 2 Hz, and a limited total cycle time of 2 s.
   Collision energy was alternated between 20 and 50 to achieve mixed MS^2
   spectra.

Search for betalains by HPLC‐UV and LC‐ESI‐HRMS

   The HPLC‐UV chromatograms at 520 nm from quantification of anthocyanins
   (see above) where searched for additional peaks that might correspond
   to betalains. However, in the HPLC‐UV chromatograms at 520 nm no other
   peaks apart from the three described anthocyanins were found.
   Additionally, betalains were searched for in the raw data from the
   LC‐ESI‐HRMS runs in positive ionization mode described above for
   structure elucidation of anthocyanins. Extracted ion chromatograms for
   the molecular ion peak [M + H]^+ of known betalains [[64]21, [65]22]
   with an isolation width of m/z 0.002 (Table [66]1) were inspected for
   possible peaks. For none of the tested known betalains an [M + H]^+
   peak could be detected, this means the compound(s) are not there or
   below the detection limit of the LC‐ESI‐HRMS system.

Table 1.

   Betalains searched in Nepenthes spp. peristomes by LC‐ESI‐HRMS.
   Compound Molecular sum formula Theoretical m/z for [M + H]^+ molecular
   ion Detected (D)/not detected (ND)
   Amaranthine C[30]H[34]N[2]O[19] 727.182853 ND
   Isoamaranthine C[30]H[34]N[2]O[19] 727.182853 ND
   Iresinin I C[36]H[42]N[2]O[23] 871.225112 ND
   Isoorientin I C[36]H[42]N[2]O[23] 871.225112 ND
   Celosianin I C[39]H[40]N[2]O[21] 873.219633 ND
   Isocelosianin I C[39]H[40]N[2]O[21] 873.219633 ND
   Celosianin II C[40]H[42]N[2]O[22] 903.230197 ND
   Isocelosianin II C[40]H[42]N[2]O[22] 903.230197 ND
   Gomphrenin I C[24]H[26]N[2]O[13] 551.150765 ND
   Isogomphrenin I C[24]H[26]N[2]O[13] 551.150765 ND
   Gomphrenin II C[33]H[32]N[2]O[15] 697.187545 ND
   Isogomphrenin II C[33]H[32]N[2]O[15] 697.187545 ND
   Gomphrenin III C[34]H[34]N[2]O[16] 727.198110 ND
   Isogomphrenin III C[34]H[34]N[2]O[16] 727.198110 ND
   Betanin C[24]H[26]N[2]O[13] 551.150765 ND
   Isobetanin C[24]H[26]N[2]O[13] 551.150765 ND
   Betanidin C[18]H[16]N[2]O[8] 389.097942 ND
   2‐Descarboxy‐betanidin C[17]H[16]N[2]O[6] 345.108113 ND
   Lampranthin II C[34]H[34]N[2]O[16] 727.198110 ND
   3‐Methoxytyramine‐betaxanthin C[18]H[20]N[2]O[6] 361.139413 ND
   (S)‐Tryptophan‐betaxanthin C[20]H[19]N[3]O[6] 398.134662 ND
   Indicaxanthin C[14]H[16]N[2]O[6] 309.108113 ND
   Miraxanthin‐V C[17]H[18]N[2]O[6] 347.123763 ND
   Betalamic Acid C[9]H[9]NO[5] 212.055349 ND
   [67]Open in a new tab

Transcriptome analysis: sampling, total RNA extraction, cDNA library
preparation, and sequencing

   For transcriptome analysis, N. x ventrata pitchers were collected first
   at the opening day of the lid as a reference time point and at the next
   two consecutive days. To avoid contamination, still‐closed pitchers
   were covered with a mesh as described in [[68]23]. Pitchers were rinsed
   three times with sterile ddH[2]O, digestive zone tissue was dissected
   sampled in 50 mL polypropylene tubes and directly frozen in liquid
   N[2]. Individual pitchers represent independent biological replicates
   from different plants. A total of 12 biological replicates were used
   for RNAseq analysis, with four replicates for each time point.

   Dissected digestive zones material was finely ground in liquid N[2]
   using mortar and pestle. Samples were stored at −80 °C until RNA
   extraction was performed. A 50 mg weighed powdery tissue material was
   used for total RNA isolation. The extraction was done at room
   temperature using RP InviTrap® Spin Plant RNA Mini kit (STRATEC
   Molecular, Berlin, Germany) according to the manufacturer's protocol
   with some modifications. Total RNA was dissolved in ddH[2]O. Each
   biological sample (digestive zone) was extracted from seven technical
   replicates and pooled in the final step of RNA elution.

   For assessing a rough indicator of quality and yield, A260/A280 and
   A260/A230 ratios for RNA preparation samples were determined with
   NanoDrop UV/Vis Spectrophotometer (Thermo Fisher, Schwerte, Germany).
   To remove any DNA contamination, the isolated RNA was treated with
   Turbo DNA‐free Kit™ (Invitrogen™, Darmstadt, Germany).

   Finally, sample quality control was performed using the yield and the
   assessment of RNA integrity number. This was done based on comparative
   evaluation of 28S/18S rRNA on an Agilent 2100 Bioanalyzer system
   following manufacturer's protocol and performed on an Agilent RNA 6000
   Nano LabChip® Kit (Agilent Technologies). Transcriptome sequencing was
   carried out by the Max Planck Genome Center (Cologne, Germany)
   ([69]https://mpgc.mpipz.de/home/) using poly(A)+ enriched RNA
   fragmented to an average of 180 nucleotides. Sequencing was done on an
   Illumina HiSeq3000 Genome Analyzer platform, using standard TruSeq
   procedures and paired‐end (2 × 150 bp) read technology, yielding
   approximately 15 million reads for each of the 28 N. x ventrata
   samples.

Transcriptome assembly, mapping, and annotation

   Quality control measures and de novo transcriptome assembly, using the
   combined RNAseq sequence data was carried out using CLC Genomics
   Workbench v11.1 ([70]http://www.clcbio.com) To assess transcriptome
   completeness, we performed a BUSCO (Benchmarking Universal Single‐Copy
   Orthologs; [71]http://busco.ezlab.org) analysis by comparing our
   assembled transcript sets against a set of highly conserved single‐copy
   orthologs. This was accomplished using the BUSCO v3 pipeline [[72]24],
   comparing the predicted proteins of the N. x ventrata transcriptome to
   the predefined set of 1614 Embryophyta single‐copy orthologs from the
   OrthoDB v9.1 database. This resulted in 78.7% complete/partial and
   21.3% missing BUSCO genes for the pitcher transcriptome assembly.
   Digital gene expression analysis was carried out using CLC Genomics
   Workbench v9.1 to generate BAM (mapping) files, and qseq Software
   (DNAStar Inc., Madison, WI, USA) was then used to estimate gene
   expression levels. Sequence similarity searches of the transcriptome
   were performed using the NCBI BLAST suite on a Galaxy server against
   the NCBI nr database. Further sequence annotations were done using Gene
   Ontology (GO) and InterPro terms (InterProScan, EBI), enzyme
   classification (EC) codes, and metabolic pathways (Kyoto Encyclopedia
   of Genes and Genomes, KEGG) implemented in BLAST2GO v5.2
   ([73]https://www.biobam.com).

KEGG pathway enrichment analysis

   Based on the KEGG orthology (KO), 10 enzymes involved in the
   anthocyanin biosynthetic pathway were identified: chalcone synthase
   ([74]K00660); chalcone isomerase ([75]K01859); flavanone 3‐dioxygenase
   ([76]K00475); flavonoid 3′‐monooxygenase (K05280); flavonoid
   3′,5′‐hydroxylase (K13083); bifunctional dihydroflavonol
   4‐reductase/flavonone 4‐reductase (K13082); anthocyanidin synthase,
   (K05277); anthocyanidin 3‐O‐glucosyltransferase (K12930); shikimate
   O‐hydroxycinnamoyl transferase (K13065); 5‐O‐(4‐coumaroyl)‐d‐quinate
   3′‐monooxygenase (K09754). To verify the putative proteins, all
   sequences were searched via blastx (e‐value 1e‐3) against NCBI
   nonredundant database (available from:
   [77]https://www.ncbi.nlm.nih.gov/; April 23, 2021). Focusing on the
   essential enzymes, dominantly expressed transcripts with log[2]RPKM > 2
   (RPKM: reads per kilo base per million mapped reads) at least in one
   out of three independently on three consecutive days harvested
   N. x ventrata pitcher samples were selected in this study
   (Table [78]S1).

Results and Discussion

   Anthocyanins are a water‐soluble group of plant pigments derived from
   flavonoids. These pigments are produced in the central vacuole of plant
   cells and occur in reproductive (flowers and fruits) or vegetative
   (stems, roots, or leaves) plant organs [[79]25]. Still, in vegetative
   tissues the functional role of anthocyanins remains a controversial
   topic, ranging from stress response to drought and nutrient deficiency,
   photoprotection, free radical scavenging, and herbivory defense
   [[80]26]. In the taxon of core Caryophyllales, anthocyanins are
   replaced by betalains and the presence of one of these pigments
   excludes that of the other [[81]1, [82]3, [83]5, [84]6]. Betalains,
   therefore, have been seen and used as chemotaxonomic markers [[85]27].
   Due to their similar locations in plant tissues and cells and their
   comparable optical features, both pigments are considered to be
   functional homologues in plant environment interactions [[86]28].

   Carnivorous plants of the Droseraceae and Nepenthaceae have been
   usually assigned to the Caryophyllales sensu stricto, but not to the
   core group of this order. Thus, for a long time it was not clear which
   pigments occur in these plants. While for D. muscipula (Droseraceae) a
   recent investigation by Henarejos‐Escudero et al. [[87]12] eventually
   demonstrated the presence of the anthocyanins delphinidin‐3‐O‐glucoside
   and cyanidin‐3‐O‐glucoside, as well as the aglycone cyanidin, thereby
   confirming earlier but from a chemical point of view less conclusive
   results; for Nepenthaceae such study was still missing. As shown in
   Fig. [88]1, for N. x ventrata peristomes there are only three peaks
   detected in an HPLC‐UV analysis at 520 nm, all three showing the
   typical UV spectrum of anthocyanins (see inset Fig. [89]1A). By match
   of retention time and UV spectra with commercial standards, we proved
   the presence of cyanidin‐3‐O‐glucoside (Fig. [90]1A, peak #2, major
   peak; Fig. [91]1D) and cyanidin‐3‐O‐galactoside (Fig. [92]1A, peak #1;
   Fig. [93]1D). The identity was further supported by analysis by
   LC‐ESI‐HRMS (Fig. [94]1C), the molecular ion peak ([M + H]^+) of peak
   #2 at m/z 449.1085 fits the molecular formula of C[21]H[21]O[11](Δ
   −1.47 p.p.m.), the fragment by collision‐induced dissociation at m/z
   287.0554 suggests a cyanidin aglycone structure (C[15]H[11]O[6], Δ
   −1.34 p.p.m.). The analysis of the commercial cyanidin‐3‐O‐glucoside
   standard resulted in almost identical values (Fig. [95]1B). The full
   scan MS data and MS^2 fragmentation pattern data strongly suggest the
   presence of cyanidin‐3‐O‐glucuronide (m/z 463.0874, C[21]H[19]O[12], Δ
   −0.64 p.p.m.; Fig. [96]1D), fragment after CID: m/z 287.0553,
   C[15]H[11]O[6], Δ −0.99 p.p.m.), represented as peak #3 in the
   HPLC‐UV/Vis chromatogram at 19.2 min (Fig. [97]1A). These results
   represent the missing proof for the existence of anthocyanins in
   Nepenthaceae.

Fig. 1.

   Fig. 1
   [98]Open in a new tab

   Identification of cyanidin‐3‐O‐glycosides as the major anthocyanidin
   compounds in Nepenthes x ventrata peristome tissue. (A) HPLC‐UV/Vis
   chromatograms at 520 nm for a N. x ventrata peristome extract (upper
   chromatogram) and a cyanidin‐3‐O‐glucoside standard (lower
   chromatogram). Insert: UV spectrum of (peak #2). Peak #1 was identified
   as cyanidin‐3‐O‐galactoside by comparison to a commercial standard.
   Peak #3 was tentatively identified by HRMS as cyanidin‐3‐O‐glucuronide.
   (B) Full scan HR‐MS spectrum (positive mode) and MS^2 fragmentation
   spectrum for cyanidin‐3‐O‐glucoside standard and (C) peak #2 of
   N. x ventrata peristome extract. (D) Structures of anthocyanins
   identified from N. x ventrata peristome extract. 1.
   Cyanidin‐3‐O‐galactoside corresponding to peak #1 in (A); Chemical
   formula: C[21]H[21]O[11] ^+; 2. Cyanidin‐3‐O‐glucoside corresponding to
   peak #2 in (A); Chemical formula: C[21]H[21]O[11] ^+; 3.
   Cyanidin‐3‐O‐glucuronide corresponding to peak #3 in (A); Chemical
   formula: C[21]H[19]O[12] ^+.

   We next analyzed the red colored peristomes and digestive zone of the
   pitcher of six additional Nepenthes species growing from Philippines
   (N. x ventrata; N. ventricosa; N. robcantleyi, endemic on the island
   Mindanao), Borneo (N. fusca), Sulawesi (N. maxima), Vietnam
   (N. thorelii), all together representing the huge area of Southeast
   Asia. In all species, the dominating anthocyanin was
   cyanidin‐3‐O‐glucoside, followed by cyanidin‐3‐O‐galactoside and
   cyanidin‐3‐O‐glucoronide. In all cases, the peristome contained more
   anthocyanins than the digestive zone, which matches well with the red
   color of the tissues (Fig. [99]2). Within the different species, the
   highest anthocyanin concentration was found in N. fusca (c.
   6.56 µmol·g^−1 fresh weight in peristome and 2.27 µmol·g^−1 fresh
   weight in digestive zone, respectively). The lowest peristome
   concentrations were determined in N. ventricosa and N. x ventrata both
   with 0.68 µmol·g^−1 fresh weight, the lowest digestive zone
   concentration in N. maxima, N. mirabilis, and N. thorelii with less
   than 0.1 µmol·g^−1 fresh weight. In N. robcantleyi, no anthocyanins at
   all could be detected in the digestive zone (Fig. [100]2). These
   results indicate that the presence of anthocyanins is widely
   distributed within Nepenthaceae but also that the different species
   contain different levels of these pigments.

Fig. 2.

   Fig. 2
   [101]Open in a new tab

   Proof of principle for the presence of anthocyanins in various species
   of the genus Nepenthes. (A) Distribution and relative amounts of
   different cyanidin anthocyanins in peristomes and digestive zones of
   seven Nepenthes species. (B) Pitcher tissue pigmentation in three
   Nepenthes species (Nepenthes fusca, Nepenthes maxima, and
   Nepenthes mirabilis). Black boxes show dissected pitcher tissues
   harvested for anthocyanin quantification (DZ, digestive zone; P,
   peristome).

   A more detailed, tissue‐specific anthocyanin analysis was performed
   with branches, leaf blades, pitcher digestive zone, and peristomes of
   N. x ventrata (Fig. [102]3A). While in the nonred branch and leaf
   tissues almost no anthocyanins could be found, both the digestive zone
   and the peristome tissues contained mainly cyanidin‐3‐O‐glucoside; the
   level of cyanidin‐3‐O‐galactoside and cyanidin‐3‐O‐glucoronide were
   similar (Fig. [103]3B). The ratio between the three anthocyanins were
   constant in the different tissues.

Fig. 3.

   Fig. 3
   [104]Open in a new tab

   Distribution and concentrations of anthocyanins in Nepenthes x ventrata
   tissues. (A) Indication of four tissues analyzed for anthocyanin
   presence: branch, leaf blade, digestive zone, peristome. (B) Determined
   concentrations of different anthocyanins in N. x ventrata tissues
   (n = 3; mean ± SEM).

   The biosynthetic pathways to anthocyanins are well known, starting with
   the shikimate pathway, followed by the phenylpropanoid pathway and
   different possible related routes within the flavonoids biosynthetic
   pathways leading to the anthocyanidins cyanidin, pelargonidin, and
   delphinidin and to their respective anthocyanin glycosides [[105]29,
   [106]30, [107]31] (Fig. [108]4A). Most of the biosynthetic enzymes are
   employed in the generation of the anthocyanidins. This was evident in
   the transcriptome analysis of the corresponding genes, all of which are
   constitutively expressed (Fig. [109]4B), suggesting that the
   biosynthetic pathways for anthocyanins are active, indicated also by
   the permanent red coloration. Thus, both molecular evidence (a complete
   set of constitutively expressed anthocyanin biosynthetic genes) and
   analytical chemistry‐based evidence now demonstrates the existence of
   anthocyanins in Nepenthes.

Fig. 4.

   Fig. 4
   [110]Open in a new tab

   Molecular evidence for anthocyanin biosynthetic pathways in Nepenthes x
   ventrata pitchers. (A) Anthocyanin biosynthesis pathways. Red, blue,
   and green colors represent putative pathways for cyanidin 3‐O‐glucoside
   synthesis in N. x ventrata. In black, the metabolic pathway for
   pelargonidin and delphinidin synthesis; both compounds were not found
   in this study. Enzymes involved in the pathway are indicated in
   italics: ANS, anthocyanidin synthase; C3ˈH, 5‐O‐(4‐coumaroyl)‐D‐quinate
   3′‐monooxygenase; CHI, chalcone isomerase; CHS, chalcone synthase; DFR,
   bifunctional dihydroflavonol 4‐reductase/flavonone 4‐reductase; F3′5′H,
   flavonoid 3′,5′‐hydroxylase; F3′H, flavonoid 3′‐monooxygenase; F3H,
   naringenin 3‐dioxygenase; HCT, shikimate O‐hydroxycinnamoyl
   transferase; UFGT, anthocyanidin 3‐O‐glucosyltransferase. (B) KEGG
   pathway enrichment analysis. Gene expression profiles of 30 unique
   transcripts in anthocyanin biosynthetic pathway, based on RPKM values.
   Candidate genes with log[2]RPKM > 2 in at least one sample (columns 1,
   2, and 3; independent N. x ventrata pitcher samples harvested on three
   consecutive days) are represented.

   Although this result is not surprising and was expected, the final
   proof of anthocyanins in Nepenthaceae was still pending and furthermore
   supports the recent results for the Droseraceae [[111]12]. We also
   searched the LC‐ESI‐HRMS datasets for molecular ion peaks [M + H]^+ of
   known betalains, but did not detect any betalain‐corresponding peak
   (Table [112]1). Moreover, among the enzymes necessary for betalain
   biosynthesis we only found three transcripts for two basic enzymes
   related to DOPA metabolism, for example, an weakly expressed
   aromatic‐L‐amino‐acid/L‐tryptophan decarboxylase ([113]K01593; GenBank
   Acc [114]MZ322092) that may generate dopamine, and two 4,5‐DOPA
   dioxygenase transcripts (DODA) (K15777; GenBank Acc [115]MZ322091,
   [116]MZ322092); no other transcripts of betalain biosynthesis‐related
   genes (CYP76AD1, CYP76AD6, 5GT, 6GT) were detected. Although the
   expression of DODA sounds interesting as it may result in the formation
   of betalamic acid, homologs of DODA have been found in many anthocyanin
   generating taxa within angiosperms [[117]3]. Strikingly, a deep search
   for betalamic acid was not successful (Table [118]1) suggesting that
   the enzyme was not built or it remained inactive.

   Hence, it can be postulated that at least all carnivorous plants of the
   taxon Nepenthales contain anthocyanins rather than betalains. From an
   ecological and economical point of view, the absence of
   nitrogen‐containing pigments such as betalains makes sense. In
   particular carnivorous plants that catch insects in order to supplement
   nutrients with additional nitrogen derived from digested prey
   [[119]32], should not consume the limited nitrogen for betalain
   synthesis when anthocyanins might very likely perform similar
   functions. Nevertheless, not all functions of the red coloration in
   carnivorous plants are known. The coloration could have initially
   developed as an adaptive trait, because anthocyanin accumulation is
   often associated with stress responses [[120]16]. At the same time, it
   increased prey capture efficiency of the traps by providing attractive
   visual signals. As insect prey capture rates positively correlate with
   levels of red pigmentation, it might enhance the trap efficiency by the
   red color itself or by providing a special background for better
   recognition [[121]16]. Concerning herbivores, anthocyanins may protect
   the tissue from attack by herbivores, which are attracted by green
   color [[122]19, [123]33]. Moreover, anthocyanins have antioxidant
   activities which might protect the plant against reactive oxygen
   species [[124]33]. The strong red pigmentation of the peristome in all
   Nepenthes species supports these hypotheses. Here, more studies need to
   be done.

Conflict of interest

   The authors declare no conflict of interest.

Author contributions

   AD‐L and AM designed the study. AD‐L and MR performed experiments.
   AD‐L, MR, DW, HV, and AM analyzed and interpreted the results. AD‐L and
   AM wrote the manuscript. All authors revised and approved the final
   version and agree to be held accountable for the article content.

Supporting information

   Table S1. Anthocyanin biosynthesis gene index and GenBank accession
   numbers.
   [125]Click here for additional data file.^ (11KB, xlsx)

Acknowledgements