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Review

Carnivorous Plants from Nepenthaceae and Droseraceae as a Source of Secondary Metabolites

by
Magdalena Wójciak
1,*,
Marcin Feldo
2,
Piotr Stolarczyk
3 and
Bartosz J. Płachno
4,*
1
Department of Analytical Chemistry, Medical University of Lublin, Chodzki 4a, 20-093 Lublin, Poland
2
Department of Vascular Surgery and Angiology, Medical University of Lublin, 11 Staszica St., 20-081 Lublin, Poland
3
Department of Botany, Physiology and Plant Protection, Faculty of Biotechnology and Horticulture, University of Agriculture in Kraków, 29 Listopada 54 Ave., 31-425 Cracow, Poland
4
Department of Plant Cytology and Embryology, Institute of Botany, Faculty of Biology, Jagiellonian University in Kraków, 9 Gronostajowa St., 30-387 Cracow, Poland
*
Authors to whom correspondence should be addressed.
Molecules 2023, 28(5), 2155; https://doi.org/10.3390/molecules28052155
Submission received: 30 January 2023 / Revised: 13 February 2023 / Accepted: 20 February 2023 / Published: 24 February 2023
(This article belongs to the Special Issue Plant Metabolites: Accumulation, Profiling and Bioactivity)
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Abstract

:
Carnivorous plants are able to attract small animals or protozoa and retain them in their specialized traps. Later, the captured organisms are killed and digested. The nutrients contained in the prey bodies are absorbed by the plants to use for growth and reproduction. These plants produce many secondary metabolites involved in the carnivorous syndrome. The main purpose of this review was to provide an overview of the secondary metabolites in the family Nepenthaceae and Droseraceae, which were studied using modern identification techniques, i.e., high-performance liquid chromatography or ultra-high-performance liquid chromatography with mass spectrometry and nuclear magnetic resonance spectroscopy. After literature screening, there is no doubt that tissues of species from the genera Nepenthes, Drosera, and Dionaea are rich sources of secondary metabolites that can be used in pharmacy and for medical purposes. The main types of the identified compounds include phenolic acids and their derivatives (gallic, protocatechuic, chlorogenic, ferulic, p-coumaric acids, gallic, hydroxybenzoic, vanillic, syringic caffeic acids, and vanillin), flavonoids (myricetin, quercetin, and kaempferol derivatives), including anthocyanins (delphinidin-3-O-glucoside, cyanidin-3-O-glucoside, and cyanidin), naphthoquinones (e.g., plumbagin, droserone, and 5-O-methyl droserone), and volatile organic compounds. Due to the biological activity of most of these substances, the importance of the carnivorous plant as a pharmaceutical crop will increase.

1. Introduction

Carnivorous plants are mainly angiosperms, represented by approximately 850 carnivorous plant species [1,2]. Exceptions are some carnivorous liverworts of the genera Colura and Pleurozia [3,4]. Carnivorous plants are polyphyletic, and carnivorous syndromes have evolved independently in about 10 lineages of flowering plants [5,6,7,8,9]. They are capable of attracting small animals or protozoa and keep them in specialized traps, which are generally of foliar origin [10,11]. However, inflorescences can sometimes be trap organs as well [9]. Captured organisms are killed and digested, and nutrients from their bodies are absorbed by plants to be used for growth and reproduction [1,10,12]. Some researchers propose that carnivorous plant traps are analogous to the animal digestive tract [13].
Carnivorous species are also sources of various secondary metabolites (about 170 compounds), which are used by them to facilitate prey attraction, capture, digestion, and nutrient assimilation as well as defense against pathogens and herbivores or pollinator attraction [14,15]. As reported by Hatcher et al. [15], these metabolites provide a potentially powerful model system for exploring the role of metabolites in plant evolution and in adaptation to extreme nutrient-poor habitats. Moreover, they exhibit diverse biological activity and therefore, potential for use in medicine [14,16,17]. Recently, Miclea provided an overview of secondary metabolites found in carnivorous plants of the Sarraceniaceae family [14]. In our study, we focused on the phytochemistry of Nepenthaceae and Droseraceae species (Figure 1).
The genus Nepenthes has a center of diversity in tropical Asia. Dionaea muscipula is restricted to a small area of the Atlantic and Gulf Coastal Plain in North America, but the genus Drosera is almost cosmopolitan with a large number of species in Australia and South Africa [18]. Aldrovanda vesiculosa used to be widespread in the Old World and across various climatic zones; however, now it is rare and endangered [19]. Plants belonging to the Nepenthaceae and Droseraceae families are of great interest to researchers and, in recent years, they have been intensively studied in terms of the biological activity of both crude extracts and isolated components. Furthermore, some species, e.g., Di. muscipula and plants from the genus Drosera are common in cultivation and easy to propagate and introduce into in vitro cultures. Therefore, their biomass is relatively easily available, which makes these plants a valuable model for biological and phytochemical investigations [20,21,22].
In the review, investigations based on modern identification techniques, such as high-performance liquid chromatography (HPLC) or ultra-high-performance liquid chromatography (UPLC) with mass spectrometry (MS) and nuclear magnetic resonance spectroscopy (MNR) of isolated compounds, have been summarized. The LC-MS-based nontargeted approach was excluded from the study when no compounds were identified. Although it is a powerful tool for the investigation of chemical diversity and useful for the assessment of changes in metabolites in plants for which available data are scarce, it needs further analyses such as nuclear magnetic resonance to elucidate the real structures of the compounds. This approach was applied, for example, to study the ionizable metabolites of pitcher traps and leaf blades in Nepenthes × ventrata [23], in pitchers of N. ampullaria, N. rafflesiana and its hybrid (N. × hookeriana) [24], and in leaves of Drosera capensis [25].
The review is based mostly on papers published after 2010, as there are some review papers summarizing older investigations [16,26,27]. However, some oldest articles were also mentioned to give a relevant background or when they provide unique well-documented information. Our paper shows that phenolic acids and their derivatives (gallic, protocatechuic, chlorogenic, ferulic, p-coumaric, gallic, hydroxybenzoic, vanillic, syringic, and caffeic acids, as well as vanillin), flavonoids (myricetin, quercetin, and kaempferol derivatives), including anthocyanins (delphinidin-3-O-glucoside, cyanidin-3-O-glucoside, cyanidin), naphthoquinones (e.g., plumbagin, droserone, 5-O-methyl droserone), and volatile organic compounds, are the main types of compounds investigated in Nepenthaceae and Droseraceae. The role of these compounds in plant physiology, including growth, development, and prey attraction, has been previously described in detail by Hatcher et al. [15].

2. Secondary Metabolites in Nepenthaceae and Droseraceae

2.1. Phenolic Compounds

Plant phenolics are the most common secondary metabolites in the plant kingdom. This group includes both simple molecules such as phenolic acids and complex compounds with high molecular mass. Phenolics are divided into several classes, i.e., phenols, phenolic acids, coumarins, flavonoids, stilbenes, hydrolyzed and condensed tannins, lignans, and lignins [28,29]. They are synthesized from carbohydrates via the shikimate pathway or via the ‘polyketide’ acetate/malonate pathway during normal plant development and in defense against stress conditions [30].

2.1.1. Phenolic Acids

General Information

A characteristic structural feature of phenolic acids is the aromatic ring with one or more hydroxyl substituents and a carboxyl moiety. Two groups of phenolic acids are distinguished: derivatives of benzoic acid and derivatives of cinnamic acid based on the skeleton of C6–C1 and C6–C3, respectively. Phenolic acids can be esterified with alcohols, amino acids, and carbohydrates or may be bonded with malic, tartaric, shikimic, lactic, and quinic acids and may form conjugates with other natural compounds [28,29].
The role of phenolics in plants is varied. They may function as phytoalexins, antifeedants, attractants for pollinators, antioxidants, and protective agents against UV light [28,29,31]. They also have a regulatory function in plant growth and reproduction. Some phenolics, e.g., ferulic and p-coumaric acids, are found in cell walls and contribute to the mechanical strength.

Phenolic Acids and Their Derivatives in Nepenthaceae and Droseraceae

As mentioned above, phenolic acids may occur as esters or complex conjugates; thus, acidic or/and alkaline hydrolysis is often performed in the investigation of plant material to release the free form. This approach has been applied by Kováčik et al. [32]. They studied free and bonded phenolic acids in N. anamensis, Drosera capensis, and Dionaea muscipula. HPLC-MS analysis revealed the presence of gallic, protocatechuic, hydroxybenzoic, vanillic, chlorogenic, syringic, caffeic, ferulic, sinapic, and p-coumaric acids and their derivatives in both the leaves and traps of the species. In addition, free salicylic acid was found in Di. muscipula and its glycoside- and ester-bound derivatives were present in Di. muscipula and Dr. capensis. A phenolic aldehyde–vanillin was also found in all species. In turn, a protocatechuic acid derivative (1-O-protochatechoyl glucoside) was found in Drosera magna and the structure was established using NMR [33].
The detailed phytochemical profile of polyphenolic constituents, including flavonoids, phenolic acids, and their derivatives, in both field-grown and in vitro propagated leaves of Dr. rotundifolia was described by Tienaho et al. The components were tentatively identified based on their retention time, elution order, exact masses, and/or characteristic MS/MS patterns. Ellagic and dimethylellagic acids and their glycosides, as well as quinic acid, monogalloyl glucose, and coumaric acid glycosides were identified in both types of plant material. In turn, methylellagic acid and digalloyl glycose were found only in the field-grown plants [20]. The derivative of ellagic acid-3,3′-di-O-methylellagic acid 4′-glucoside was also identified based on 1H and 13C NMR in aboveground parts of Dr. tokaiensis [34]. Furthermore, HPLC-DAD-ESI/MS analysis revealed the occurrence of ellagic acid, the isomer of dimethylellagic acid, 3-O-methylellagic acid, and 3,3′-di-O-methylellagic acid in Di. muscipula [35] and 3,3-di-O-methylellagic acid in Dr. binata [36].
The presence of gallic, chlorogenic, protocatechuic, ferulic, and p-coumaric acids [22], as well as caffeic, salicylic, and ellagic acids [37] were observed in Di. muscipula, and elagic acid was detected in Dr. binata, Dr. indica, Dr. spatulata, and Di. muscipula [38]; however, the identification was only based on HPLC separation and comparison with the standards.
Structures of the most common phenolic compounds identified in Nepenthaceae and Droseraceae species are shown in Figure 2.

2.1.2. Flavonoids

General Information

Flavonoids are widespread plant secondary metabolites with a structure based on the C6–C3–C6 backbone. They can be divided into several subclasses: anthocyanins, chalcones, flavanones, flavones, flavonols, isoflavonoids, and flavanols on the basis of their characteristic structural features. They have multiple functions in plant organisms, e.g., protection against the harmful effects of ultraviolet–visible (UV-B) radiation, antimicrobial activity, and an inhibitory effect against herbivores such as insects and nematodes. They are also responsible for yellow-orange coloration and participate in copigmentation with anthocyanins, which increases color intensity and stability [39]. Flavonoids found in Nepenthaceae and Droseraceae plants mostly belong to derivatives of quercetin kaempferol and myricetin (Figure 3).

Flavonoids in Nepenthaceae and Droseraceae

The presence of flavonoids in the genera Nepenthes, Drosera, and Dionaea was previously reported in papers from the 1960s to the 1990s. For example, quercetin and its 3-O-galactoside, 3-O-glucoside, and 3-O-digalactoside, gossypetin and its 7-O-glucoside, and 3-O-glucoside kaempferol were detected in Dr. rotundifolia [40,41]. Iwashina et al. reported the presence of kaempferol and quercetin 3-O-glycosides in Di. muscipula and myricetin and quercetin 3-O-galactosides in Dr. spathulata [42]. Moreover, the presence of kaempferol and quercetin glycoside was determined based on the TLC screening of three Nepenthes hybrids: N. rajach, N. tentaculata, and N. muluensis [43]. In some studies, identification was based on a comparison of retention times with standards; for example, hyperoside, isoquercitrin, myricetin, quercetin, and ellagic acid were found in Dr. rotundifolia and Dr. madagascariensis [44], and in Dr. spatulata and Dr. tokaiensis [45]. However, retention parameters are often misleading, as coelution may occur in the case of compounds with similar polarity, and only further studies based on mass spectrometry and NMR identification can confirm the occurrence of specific derivatives of quercetin and kaempferol. In another study, epicatechin 3-gallate and quercetin gallate esters: quercetin 3-O-(2″-galloylarabinofuranoside), quercetin 3-O-(6″-galloylglucoside), quercetin 3-O-(3″-galloylrhamnoside), quercetin 3-O-(2″-galloylxylopyranoside), and quercetin 3-O-(3″-galloylxylopyranoside), were isolated from N. gracilis leaves using preparative chromatography and characterized using C NMR and ESI–MS [46]. The appearance of quercetin and kaempferol after acidic hydrolysis in the trap and leaves of N. anamensis, Dr. capensis, and Di. muscipula was confirmed in HPLC–MS by Kováčik et al. [32], proving the presence of many derivatives of these aglycones in the species. In turn, free flavonoid aglycones: methylated myricetin, quercetin, isorhamnetin, or rhamnetin, kaempferol or fisetin, and their glycosidic conjugates: 3-O-glycosides and 3-O-rhamnosylglucoside kaempferol or fisetin were observed in Dr. binata cultured in vitro. As can be seen, some flavonoids were not distinguished based on the collected MS data, because the formula and, therefore, the molecular mass of kaempferol and fisetin, as well as iorhamnetin and rhamnetin are the same (C15H10O6 and C16H12O7, respectively) [47], and more studies are needed to clarify this issue.
Furthermore, NMR and mass spectroscopic data allowed establishing the structure of the main flavonoid, quercetin 3-O-(6′-n-butyl-D-glucuronide isolated from N. thorellii × (ventricosa × maxima) [48]. In turn, Wong et al. conducted a comparative UHPLCQ/TOF-MS-based metabolomic analysis of four Nepenthes species, including N. minima, N. ampullaria, N. rafflesiana, and N. northiana. They tentatively identified 89 metabolites from different groups and, among others, flavonoid compounds, mostly derivatives of quercetin and kaempferol [49]. In another study, Fourier transform ion cyclotron resonance mass spectra (FT-ICR-MS) and 1H and 13C NMR data allowed establishing the structure of four flavonoids (quercetin, quercetin 3-O-(6′-n-butyl ß-D-glucuronide), quercitrin, kaempferol-3-O-a-L-rhamnoside) isolated from N. mirabilis leaves [50], and quercetin, hyperoside, and 2′-O-galloylhyperoside were identified in leaves of Dr. rotundifolia based on UHPLC-TOF-MS [51]. In turn, NMR analysis of Dr. magna components isolated using column fractionation revealed the presence of tamarixetin-3-rhamnoside, naringenin-6-C-β-D-glucopyranoside, hirsutrin, and four new flavonoid compounds (three flavonol diglycosides and flavan-3-ol glycoside) [33].
Flavonoids identified by MS-MS analysis in Dr. rotundifolia grown in the field and propagated in vitro included myricetin glycoside, hyperoside, galloylhyperoside, and hydroxybenzoylhyperin. Additionally, dihydromyricetin, hexahydroxyflavonegalloyl glycoside, tetrahydroxyflavone, kaempferol-galloylglycoside, quercetin, and its derivatives: glycoside and glycoside gallate, were found in plants grown in field conditions. In turn, syringetin glycoside and spinatoside were identified only in in vitro cultivated species [20]. Myricitrine and quercimelin were isolated from the aboveground parts of Dr. tokaiensis [34].

Anthocyanins

Anthocyanins are plant pigments from the flavonoid class. They are water-soluble compounds that display different colors (red, blue, and purple), and the color depends on pH, light, and temperature [52].
Anthocyanins are found both in reproductive organs (flowers and fruits) and vegetative organs (stems, roots, or leaves). They mostly occur as glycosides and aglycone forms (anthocyanidins) are rarely found in nature. Cyanidin derivatives followed by delphinidin derivatives are the two most common anthocyanins found in plants [52,53]. It should be mentioned that the coloration of plant organs may be a result of the presence of betalains—pigments derived from amino acid tyrosine; however, the presence of betalains and anthocyanins exclude each other [54].
Plant pigments play an important role in the interaction between plants and animals because they attract pollinators and seed dispersers. In carnivorous plants, these pigments are responsible for the distinctive red coloration of the trap, which probably attracts prey; however, the issue is still under debate because many invertebrate species cannot distinguish red wavelengths of light. The other suggested functions of trap pigments include protection against stress related to excess light exposure, nutrient deficiency, ultraviolet radiation, environmental conditions, pathogens, and predators [55].
Although speculation on the presence of anthocyanins in Nepenthes species was found in the literature previously, the detection methods used were not selective. A study conducted by Dávila-Lara et al. [56] gave clear evidence. Based on chromatographic parameters, UV–Vis spectra, and mass data obtained with the use of ultra-high performance liquid chromatography-electrospray ionization-high-resolution mass spectrometry (UHPLC–ESI–HRMS), three different cyanidin derivatives, i.e., 3-O-glucoside, 3-O-galactoside, and 3-O-glucuronide, were identified in mature and well-developed pitchers in seven Nepenthes species: N. × ventrata (natural hybrid of N. ventricosa and N. alata), N. thorelii, N. ventricosa, N. robcantleyi, N. maxima, N. fusca, and N. mirabilis. An analysis of the distribution of anthocyanins in Nepenthes × ventrata tissue showed that the pigments were most abundant in the peristome, compared to the digestive zone, and 3-O-glucoside was predominant (0.68 vs. ca 0.12 μmol/g of fresh weight, respectively). Low amounts of pigments were also identified in the branches and leaf blades of the plants. Interestingly, the ratio between the anthocyanins was constant in the different tissues [56].
Some studies report the presence of anthocyanins in Droseraceae, including cyanidin and delphinidin 3-O-glucosides in Di. muscipula [57], cyanidin and pelargonidin glycosides in Dr. anglica and Dr. intermedia [58], and cyanidin 3,5-di-O-glucoside, and 3-O-galactoside, as well as pelargonidin 3-O-galactoside and 3-O-glucoside in Dr. spatulata [59]. However, it should be noted that the reports were published in 1966–1999 and the investigation was carried out using poorly selective techniques such as column chromatography with spectral analysis or thin layer chromatography (or the methodology was not described); thus, the results need to be verified and the molecular structure should be established with the use of more selective modern techniques.
Only one recent study gives reliable information on anthocyanins in Droseraceae. Henarejos-Escudero et al. investigated pigments in digestive glands, epidermis of the base of the mature stems, and flower petals from Di. muscipula using HPLC with mass spectrometry (ESI-MS/MS), and three compounds were found, i.e., delphinidin-3-O-glucoside (myrtillin), cyanidin-3-O-glucoside (kuromanin), and cyanidin. Kuromanin was predominant in the snap trap, whereas myrtillin was the most abundant in mature stems. Aglycone (cyanidin) was found only in the snap trap [60].
Structures of the most common anthocyanins identified in Nepenthaceae and Droseraceae species are shown in Figure 4.

2.2. Naphthoquinones

2.2.1. General Information

Naphthoquinones derived from the naphthalene class are bicyclic structures with two carbonyl groups at positions 1, 4 or 1, 2. They are known toxins and insect ecdysis inhibitors. They also provide antimicrobial protection against visiting preys and preserve the prey during digestion. They show allelopathic effects that might protect plants from pathogen infection and can increase the ability of plants to compete with surrounding organisms for limited resources or to deter herbivores [15,61]. Different substituents may be attached to the ring structure; however, the presence of a hydroxyl and/or methyl group in the quinone structure is typically found in nature.

2.2.2. Naphthoquinones in Nepenthaceae and Droseraceae

Naphthoquinones are characteristic chemotaxonomic markers within Nepenthaceae and Droseraceae families. Several 1,4-naphthoquinone derivatives were found, and plumbagin was the main compound identified. The review paper by Devi et al. listed 114 species [26] in which plumbagin and sometimes other naphthoquinones were detected. Among them, 91 Nepenthes species were investigated by Schlauer et al. using TLC and GC-MS analysis of steam distillates and ether extracts from fresh leaves [62]. Furthermore, the studies conducted using GC-MS followed by isolation and elucidation of the structure using 1H-NMR, 13C-NMR have detected plumbagin in N. khasiana roots and in the waxy layers at the top prey capture region of the pitchers. In turn, its derivatives, i.e., droserone (3,5-dihydroxy-2-methyl-1,4-naphthoquinone, the oxygenated form of plumbagin) and 5-O-methyl droserone (2-methyl-3-hydroxy-5-methoxy -1,4-naphtho-quinone), were found in pitcher fluid after induction with chitin or prey capture [63,64].
Plumbagin was also detected in the extract from aerial parts of N. × thorelii (ventricosa × maxima) [65] and N. cv. ‘Miranda’ [66] and in the leaf and pitcher tissue of N. × ventrata, a natural hybrid of N. alata and N. ventricosa [67,68]. A detailed study of naphthoquinones in the extract of branches and leaves of N. mirabilis conducted by Thanh et al. [50] revealed the presence of droserone, plumbagin, 3-methoxy-7-methyljuglone, 2-methoxy-7-methyljuglone, nepenthone C, and nepenthones F and G.
Although there are some reports on naphthoquinones in Nepenthes genus, less is known about this group in Droseraceae family. Only plumbagin was found in Dr. intermedia [69] and D. peltata var. lunata [70], and its structure was established after isolation and NMR analysis. Norman et al. have recently provided more details on the phytochemical profiling of crude extracts from bulbs and leaves of Dr. magna followed by isolation and NMR characterization. The study showed the presence of 1,4-naphthoquinones including hydroxydroserone, hydroxydroserone -5-O-β-D-glucoside, droserone, and plumbagin, and two 2,3-dihydronapthalene-1,4-diones [33]. Plumbagin was also determined in Dr. burmanii by competitive ELISA using a monoclonal antibody against plumbagin [71], as well as in Aldrovanda vesiculosa L. [72] and in Dr. binata using HPLC and MS data [47]. In a recent study, 7-methyljuglone (ramentaceone), 7-methyljuglone diglycoside, and 7-methyljuglone glycoside have been found in Dr. rotundifolia [20]. Furthermore, plumbagin and 3-chloroplumbagin were identified in Dr. binata using HPLC-DAD-ESI/MS [36], plumbagin along with its derivatives (dihydroplumbagin, 3-chloroplumbagin, 8,8′-biplumbagin) were detected in Di. muscipula [35], and MS and NMR analysis proved that ramentaceone is a main naphthoquinone in Dr. aliciae from in vitro cultures [73,74]. HPLC and a comparison of retention parameters with standards showed the presence of droserone and plumbagin in Di. muscipula and Dr. binata, droserone and ramentaceone in Dr. spatulata and droserone, plumbagin and ramentaceone in Dr. indica [38].
Among recent reports, only one paper describes the distribution of naphthoquinones within particular plant organs [64]. Based on the analysis of the N. khasiana species, Raj et al. revealed that plumbagin was the most abundant in the roots (approx. 1.4%), followed by the leaf (0.4%) and the stem (0.2%), and only approx. 0.04% was found in the pitcher.
The structures of naphthoquinones identified in Nepenthaceae and Droseraceae are presented in Figure 5.

2.2.3. Acetogenic Tetralones

Acetogenic tetralones are regarded as derivatives of naphthoquinone differing in the presence of the hydroxyl group in position 4 instead of the carbonyl group. A few members of this group, namely isoshinanolone, shinanolone, and epishinanolone (Figure 6), were described by Aung et al. as a component of extract from leaves of N. gracilis. After isolation, identification was based on mass and NMR spectra; however, only shinanolone data were shown [75]. Cis-isoshinanolone was found in the extract of N. mirabilis branches and leaves [50] and in the aerial parts of N. thorellii × (ventricosa × maxima) [48]. After isolation, the structure was confirmed with the NMR method.

2.3. Other Metabolites

Volatile organic compounds (VOCs) involved in the attraction of prey are widely investigated secondary metabolites in carnivorous plants, and many different compounds have been identified. They are lipophilic components with low molecular weight and high vapor pressure at ambient temperature. The composition of VOC is very complex and includes a wide range of compounds from different classes, e.g., volatile terpenes (monoterpenes, diterpenes, and sesquiterpenes), phenylpropanoids, benzenoids, and volatile fatty acids [76]. Gas chromatography with mass spectrometry is the most common method for identification of VOCs. VOCs were identified, for example, in N. rafflesiana—fifty-four compounds [77], in N. rajah—44 components [78], and in Di. muscipula—over 60 VOCs [79]. More details on VOCs in carnivorous plants were provided by Hatcher et al. [25].
Furthermore, a detailed metabolomics analysis based on UHPLCQ/TOF-MS of the aerial parts of N. minima, N. ampullaria, N. rafflesiana, and N. northiana revealed the presence of non-phenolic compounds, for example alkaloids [49].
It should also be mentioned that, in addition to secondary metabolites, proteins in pitcher fluid have been intensively studied [80,81,82]. They belong to pathogenesis-related proteins and are responsible for the digestion of prey and the antibacterial effect. For example, 29 proteins belonging to serine carboxypeptidases, α and ß galactosidases, lipid transfer proteins, and esterases/lipases were found to be excreted in N. mirabilis, N. alata, N. sanguinea, N. bicalcarata, and N. albomarginata pitchers [81].
The secondary metabolites detected in Nepenthes and Droseraceae, as well as the methods applied for their determination, are summarized in Table 1, Table 2 and Table 3.

3. Biological Potential of Nepentheceae and Droseraceae Species

Carnivorous plants from Nepentheceae and Droseraceae have been used in folk medicine for a long time in the treatment of various disorders [17,27], and recent investigations have confirmed a wide range of activities of both crude extracts and isolated compounds [20,37,51,86].For example, it has been evidenced that Dr. rotundifolia and Dr. tokaiensis extracts have anti-inflammatory potential [45] and N. bicalcarata shows antimicrobial and antidiabetic activity [87]. Antibacterial activity was also evidenced for N. cultivar ‘Miranda’ against Klebsiella pneumonia [88], for N. gracilis against Bacillus subtilis and Escherichia coli [89], and for Di. muscipula against Staphylococcus aureus, Enterococcus faecalis, E. coli, and Pseudomonas aeruginosa [21,37]. A biofilm inhibitory effect against E. coli strains was also proved for Dr. rotundifolia and Dr. intermedia [90]. Additionally, Dr. rotundifolia extract exhibited antiviral activity and was effective against enteroviruses [20].
Moreover, it has been shown that some species are effective against different types of cancer. N. adrianii × clipeata showed cytotoxicity toward oral cancer cells [84], N. thorelii × (ventricosa × maxima) exerted anti-leukemic properties [65] and an anticancer effect toward breast cancer [48], and N. cultivar ‘Miranda’ was active in the case of lung cancer [88]. It should also be mentioned that the extracts had lower cytotoxicity to normal cells compared to cancer cells.
It is assumed that the majority of the biological properties of Nepenthaceae and Droseraceae species are related to naphthoquinones. It was found that ramentaceone isolated from Dr. aliciae [73,74], chloroplumbagin from Di. muscipula [91], and plumbagin derived from N. alata [92] showed significant anticancer activity. Furthermore, plumbagin (from N. gracilis) [93] and its isomer ramentaceone (from Dr. aliciae) [86] were shown to be have antibacterial and antifungal properties. Phytochemical screening of anti-inflammatory constituents of N. mirabilis also showed that naphthoquinone derivatives were the most potent inhibitors of production of proinflammatory cytokines in cells with lipopolysaccharide-induced inflammation; however, some other constituents, e.g., phenolic compounds, also exhibit an anti-inflammatory effect [94]. Both phenolic compounds and nepenthosides were responsible for the antiosteoporotic activity of N. mirabilis extract [50,95] and flavonoids: quercetin and 2″-O-galloylhyperoside were the main components of Dr. rotundifolia extract involved in antispasmodic action [51].

4. Conclusions and Future Directions

The review showed that species from the genera Nepenthes, Drosera, and Dionaea are rich sources of secondary compounds, which can be grouped into four major types: phenolic acids and derivatives, flavonoids (including anthocyanins), naphthoquinones (including acetogenic tetralones), and volatile organic compounds. Among them, the greatest attention is paid to flavonoids and naphthoquinones, and these metabolites have been intensively studied recently in terms of their biological activity. It has been evidenced that naphthoquinones are cytotoxic against several types of cancer, and they have significant antibacterial, antifungal, antiviral, insecticidal, anti-inflammatory, and antipyretic properties [35,36,94,95,96]. In turn, flavonoids have been shown to exert antiosteoporotic, anti-inflammatory, and antispasmodic effects [50,51,94,95]. This clearly indicates that Nephentaceae and Droseraceae plants have a great potential as a source of components for the development of new drugs and therapies. Furthermore, isolated components may serve as valuable additives to enhance biological effects, for example, antibacterial activity of nanosilver particles [35,36,38].
Obviously, Nephentaceae and Droseraceae are not the only sources of these compounds. For instance, plumbagin is commonly found in members of families Plumbaginaceae and Ebenceae [97]. However, it should be highlighted that many species from the genera Nepenthes, Drosera, and Dionaea can be relatively easily cultivated in vitro and, therefore, they can be a readily available alternative source of metabolites.
The review has shown that there is still scarce information on the chemistry of carnivorous plants including Nepenthaceae and Droseraceae families; in particular, the genus Aldrovanda is poorly explored. Therefore, this issue should be intensively studied in the future, especially in the context of reports on the biological activity of extracts or components isolated from some species. The bioactivity-guided approach, which combines the application of modern isolation and identification techniques with assays of the biological activity of particular fractions, seems to be a promising tool for a comprehensive exploration of the biological potential of these species, which may lead to discovering new therapeutic agents [93].
Moreover, future research should be focused on verification of older reports on the phytochemical composition with the use of modern analytical techniques. The investigations should be also extended to other groups of compounds, including primary metabolites, as it has recently been shown that Dionaea and Aldrovanda traps contain arabinogalactan proteins with potential in industry [98,99], and proteins from Nepenthes digestive fluid could help in the therapy of celiac disease [100].
Furthermore, detailed studies on the distribution of particular groups of metabolites in plant tissues should be conducted [101], given the scarcity of such papers shown in this review. Further phytochemical studies should be also focused on the investigation of the numerous cultivars and mutants of Dionaea and hybrids of Nepenthes because literature data on this topic is scarce so far. It should be noted that the investigation of such plants can lead to a breakthrough in science as has recently been demonstrated by Anda-Larisa et al. [102].
It is also worth increasing the efforts to propagate carnivorous species in vitro and test different elicitors and cultivation conditions, which may significantly increase biomass production and the content of metabolites [103,104].
Our paper summarizes the current knowledge on the occurrence of secondary metabolites in Nepenthes, Drosera, and Dionaea, which can be a good starting point for further investigations and can help researchers dealing with the phytochemistry of these genera.

Author Contributions

Conceptualization: M.W. and B.J.P.; methodology: M.W. and B.J.P.; investigation: B.J.P., M.W., M.F. and P.S.; resources: M.W. and B.J.P.; data curation: M.W. and B.J.P.; writing— preparing the original draft: M.W., B.J.P., M.F. and P.S.; writing—review and editing: M.W. and B.J.P.; visualization: B.J.P. and M.W.; supervision: M.W. and B.J.P.; project administration: M.W., B.J.P.; funding acquisition: M.W., B.J.P. and M.F. All authors have read and agreed to the published version of the manuscript.

Funding

In the case of B.J.P., this research was partially financially supported by the Ministry of Science and Higher Education of Poland as part of the statutory activities of the Department of Plant Cytology and Embryology, Institute of Botany, Faculty of Biology, Jagiellonian University in Kraków (N18/DBS/000002).

Institutional Review Board Statement

Not applicable.

Informed Consent Statement

Not applicable.

Data Availability Statement

Data are contained within the article.

Acknowledgments

We thank the Botanical Garden of the Jagiellonian University for the opportunity to use plants from the garden collection. We would like also to thank the Center for Technology Transfer CITTRU, Jagiellonian University in Krakow for their assistance and funding for the research.

Conflicts of Interest

The authors declare no conflict of interest.

References

  1. Król, E.; Płachno, B.J.; Adamec, L.; Stolarz, M.; Dziubińska, H.; Trębacz, K. Quite a Few Reasons for Calling Carnivores ‘the Most Wonderful Plants in the World’. Ann. Bot. 2012, 109, 47–64. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  2. Cross, A.T.; Krueger, T.A.; Gonella, P.M.; Robinson, A.S.; Fleischmann, A.S. Conservation of Carnivorous Plants in the Age of Extinction. Glob. Ecol. Conserv. 2020, 24, e01272. [Google Scholar] [CrossRef]
  3. Barthlott, W.; Fischer, E.; Frahm, J.-P.; Seine, R. First Experimental Evidence for Zoophagy in the Hepatic Colura. Plant Biol. 2000, 2, 93–97. [Google Scholar] [CrossRef]
  4. Hess, S.; Frahm, J.P.; Theisen, I. Evidence of Zoophagy in a Second Liverwort Species, Pleurozia purpurea. Bryologist 2005, 108, 212–218. [Google Scholar] [CrossRef]
  5. Albert, V.A.; Williams, S.E.; Chase, M.W. Carnivorous Plants: Phylogeny and Structural Evolution. Science 1992, 257, 1491–1495. [Google Scholar] [CrossRef]
  6. Cameron, K.M.; Wurdack, K.J.; Jobson, R.W. Molecular Evidence for the Common Origin of Snap-Traps among Carnivorous Plants. Am. J. Bot. 2002, 89, 1503–1509. [Google Scholar] [CrossRef]
  7. Fleischmann, A.; Schlauer, J.; Smith, S.A.; Givnish, T.J. Evolution of Carnivory in Angiosperms; Oxford University Press: Oxford, UK, 2018; Volume 1. [Google Scholar] [CrossRef]
  8. Silva, S.R.; Gibson, R.; Adamec, L.; Domínguez, Y.; Miranda, V.F.O. Molecular Phylogeny of Bladderworts: A Wide Approach of Utricularia (Lentibulariaceae) Species Relationships Based on Six Plastidial and Nuclear DNA Sequences. Mol. Phylogenet. Evol. 2018, 118, 244–264. [Google Scholar] [CrossRef] [Green Version]
  9. Lin, Q.; Ané, C.; Givnish, T.J.; Graham, S.W. A New Carnivorous Plant Lineage (Triantha) with a Unique Sticky-Inflorescence Trap. Proc. Natl. Acad. Sci. USA 2021, 118, e2022724118. [Google Scholar] [CrossRef]
  10. Ellison, A.M.; Adamec, L. Introduction: What Is a Carnivorous Plant? Oxford University Press: Oxford, UK, 2018; Volume 1. [Google Scholar] [CrossRef]
  11. Płachno, B.J.; Muravnik, L.E. Functional Anatomy of Carnivorous Traps; Oxford University Press: Oxford, UK, 2018; Volume 1. [Google Scholar]
  12. Givnish, T.J.; Sparks, K.W.; Hunter, S.J.; Pavlovič, A. Why Are Plants Carnivorous? Cost/Benefit Analysis, Whole-Plant Growth, and the Context-Specific Advantages of Botanical Carnivory; Oxford University Press: Oxford, UK, 2018; Volume 1. [Google Scholar] [CrossRef]
  13. Freund, M.; Graus, D.; Fleischmann, A.; Gilbert, K.J.; Lin, Q.; Renner, T.; Stigloher, C.; Albert, V.A.; Hedrich, R.; Fukushima, K. The Digestive Systems of Carnivorous Plants. Plant Physiol. 2022, 190, 44–59. [Google Scholar] [CrossRef]
  14. Miclea, I. Secondary Metabolites with Biomedical Applications from Plants of the Sarraceniaceae Family. Int. J. Mol. Sci. 2022, 23, 9877. [Google Scholar] [CrossRef]
  15. Hatcher, C.R.; Ryves, D.B.; Millett, J. The Function of Secondary Metabolites in Plant Carnivory. Ann. Bot. 2020, 125, 399–411. [Google Scholar] [CrossRef] [Green Version]
  16. Mithöfer, A. Carnivorous Pitcher Plants: Insights in an Old Topic. Phytochemistry 2011, 72, 1678–1682. [Google Scholar] [CrossRef]
  17. Sanusi, S.B.; Abu Bakar, M.F.; Mohamed, M.; Sabran, S.F.; Mainasara, M.M. Ethnobotanical, Phytochemical, and Pharmacological Properties of Nepenthes Species: A Review. Asian J. Pharm. Clin. Res. 2017, 10, 16–19. [Google Scholar] [CrossRef] [Green Version]
  18. Brewer, J.S.; Schlauer, J. Biogeography and Habitats of Carnivorous Plants; Oxford University Press: Oxford, UK, 2018; Volume 1. [Google Scholar]
  19. Adamec, L. Biological Flora of Central Europe: Aldrovanda vesiculosa L. Perspect. Plant Ecol. Evol. Syst. 2018, 35, 8–21. [Google Scholar] [CrossRef]
  20. Tienaho, J.; Reshamwala, D.; Karonen, M.; Silvan, N.; Korpela, L.; Marjomäki, V.; Sarjala, T. Field-Grown and In Vitro Propagated Round-Leaved Sundew (Drosera rotundifolia L.) Show Differences in Metabolic Profiles and Biological Activities. Molecules 2021, 26, 3581. [Google Scholar] [CrossRef]
  21. Makowski, W.; Królicka, A.; Nowicka, A.; Zwyrtková, J.; Tokarz, B.; Pecinka, A.; Banasiuk, R.; Tokarz, K.M. Transformed Tissue of Dionaea muscipula J. Ellis as a Source of Biologically Active Phenolic Compounds with Bactericidal Properties. Appl. Microbiol. Biotechnol. 2021, 105, 1215–1226. [Google Scholar] [CrossRef]
  22. Makowski, W.; Królicka, A.; Tokarz, B.; Miernicka, K.; Kołton, A.; Pięta, Ł.; Malek, K.; Ekiert, H.; Szopa, A.; Tokarz, K.M. Response of Physiological Parameters in Dionaea muscipula J. Ellis Teratomas Transformed with RolB Oncogene. BMC Plant Biol. 2021, 21, 564. [Google Scholar] [CrossRef]
  23. Dávila-Lara, A.; Rodríguez-López, C.E.; O’Connor, S.E.; Mithöfer, A. Metabolomics Analysis Reveals Tissue-Specific Metabolite Compositions in Leaf Blade and Traps of Carnivorous Nepenthes Plants. Int. J. Mol. Sci. 2020, 21, 4376. [Google Scholar] [CrossRef]
  24. Rosli, M.A.F.; Mediani, A.; Azizan, K.A.; Baharum, S.N.; Goh, H.-H. UPLC-TOF-MS/MS-Based Metabolomics Analysis Reveals Species-Specific Metabolite Compositions in Pitchers of Nepenthes ampullaria, Nepenthes rafflesiana, and Their Hybrid Nepenthes × hookeriana. Front. Plant Sci. 2021, 12, 655004. [Google Scholar] [CrossRef]
  25. Hatcher, C.R.; Sommer, U.; Heaney, L.M.; Millett, J. Metabolomic Analysis Reveals Reliance on Secondary Plant Metabolites to Facilitate Carnivory in the Cape Sundew, Drosera capensis. Ann. Bot. 2021, 128, 301–314. [Google Scholar] [CrossRef]
  26. Devi, S.P.; Kumaria, S.; Rao, S.R.; Tandon, P. Carnivorous Plants as a Source of Potent Bioactive Compound: Naphthoquinones. Trop. Plant Biol. 2016, 9, 267–279. [Google Scholar] [CrossRef]
  27. Egan, P.A.; van der Kooy, F. Phytochemistry of the Carnivorous Sundew Genus Drosera (Droseraceae)—Future Perspectives and Ethnopharmacological Relevance. Chem. Biodivers. 2013, 10, 1774–1790. [Google Scholar] [CrossRef] [PubMed]
  28. Quideau, S.; Deffieux, D.; Douat-Casassus, C.; Pouységu, L. Plant Polyphenols: Chemical Properties, Biological Activities, and Synthesis. Angew. Chem. Int. Ed. 2011, 50, 586–621. [Google Scholar] [CrossRef] [PubMed]
  29. Cheynier, V.; Comte, G.; Davies, K.M.; Lattanzio, V.; Martens, S. Plant Phenolics: Recent Advances on Their Biosynthesis, Genetics, and Ecophysiology. Plant Physiol. Biochem. 2013, 72, 1–20. [Google Scholar] [CrossRef] [PubMed]
  30. Hano, C.; Tungmunnithum, D. Plant Polyphenols, More than Just Simple Natural Antioxidants: Oxidative Stress, Aging and Age-Related Diseases. Medicines 2020, 7, 26. [Google Scholar] [CrossRef]
  31. Li, Z.-H.; Wang, Q.; Ruan, X.; Pan, C.-D.; Jiang, D.-A. Phenolics and Plant Allelopathy. Molecules 2010, 15, 8933–8952. [Google Scholar] [CrossRef] [Green Version]
  32. Kováčik, J.; Klejdus, B.; Repčáková, K. Phenolic Metabolites in Carnivorous Plants: Inter-Specific Comparison and Physiological Studies. Plant Physiol. Biochem. 2012, 52, 21–27. [Google Scholar] [CrossRef]
  33. Norman, E.O.; Tuohey, H.; Pizzi, D.; Saidah, M.; Bell, R.; Brkljača, R.; White, J.M.; Gasser, R.B.; Taki, A.C.; Urban, S. Phytochemical Profiling and Biological Activity of the Australian Carnivorous Plant, Drosera magna. J. Nat. Prod. 2021, 84, 964–971. [Google Scholar] [CrossRef]
  34. Tominaga, Y.; Sugawa, H.; Hirabayashi, K.; Ikeda, T.; Hoshi, Y.; Nagai, R. Drosera Tokaiensis Extract Containing Multiple Phenolic Compounds Inhibits the Formation of Advanced Glycation End-Products. Arch. Biochem. Biophys. 2020, 693, 108586. [Google Scholar] [CrossRef]
  35. Krychowiak-Maśnicka, M.; Krauze-Baranowska, M.; Godlewska, S.; Kaczyński, Z.; Bielicka-Giełdoń, A.; Grzegorczyk, N.; Narajczyk, M.; Frackowiak, J.E.; Krolicka, A. Potential of Silver Nanoparticles in Overcoming the Intrinsic Resistance of Pseudomonas aeruginosa to Secondary Metabolites from Carnivorous Plants. Int. J. Mol. Sci. 2021, 22, 4849. [Google Scholar] [CrossRef]
  36. Krychowiak, M.; Grinholc, M.; Banasiuk, R.; Krauze-Baranowska, M.; Głód, D.; Kawiak, A.; Królicka, A. Combination of Silver Nanoparticles and Drosera binata Extract as a Possible Alternative for Antibiotic Treatment of Burn Wound Infections Caused by Resistant Staphylococcus aureus. PLoS ONE 2014, 9, e115727. [Google Scholar] [CrossRef] [Green Version]
  37. Makowski, W.; Tokarz, K.M.; Tokarz, B.; Banasiuk, R.; Witek, K.; Królicka, A. Elicitation-Based Method for Increasing the Production of Antioxidant and Bactericidal Phenolic Compounds in Dionaea muscipula J. Ellis Tissue. Molecules 2020, 25, 1794. [Google Scholar] [CrossRef]
  38. Banasiuk, R.; Krychowiak, M.; Swigon, D.; Tomaszewicz, W.; Michalak, A.; Chylewska, A.; Ziabka, M.; Lapinski, M.; Koscielska, B.; Narajczyk, M.; et al. Carnivorous Plants Used for Green Synthesis of Silver Nanoparticles with Broad-Spectrum Antimicrobial Activity. Arab. J. Chem. 2020, 13, 1415–1428. [Google Scholar] [CrossRef]
  39. Jiang, N.; Doseff, A.; Grotewold, E. Flavones: From Biosynthesis to Health Benefits. Plants 2016, 5, 27. [Google Scholar] [CrossRef] [Green Version]
  40. Bienenfeld, W.; Katzlmeier, H. Flavonoide aus Drosera rotundifolia L. Arch. Pharm. 1966, 299, 598–602. [Google Scholar] [CrossRef]
  41. Schölly, T.; Kapetanidis, I. Flavonol and Naphthoquinone Glycosides of Drosera rotundifolia. Planta Med. 1989, 55, 611–612. [Google Scholar] [CrossRef]
  42. Iwashina, T. Flavonoid Properties of Five Families Newly Incorporated into the Order Caryophyllales (Review). Bull. Natl. Mus. Nat. Sci. Ser. 2013, 39, 25–51. [Google Scholar]
  43. Adam, J.H.; Omar, R.; Wilcock, C.C. Phytochemical Screening of Flavonoids in Three Hybrids of Nepenthes (Nepenthaceae) and Their Putative Parental Species from Sarawak and Sabah. J. Biol. Sci. 2002, 2, 623–625. [Google Scholar] [CrossRef] [Green Version]
  44. Paper, D.H.; Karall, E.; Kremser, M.; Krenn, L. Comparison of the Antiinflammatory Effects of Drosera rotundifolia and Drosera madagascariensis in the HET-CAM Assay. Phytother. Res. 2005, 19, 323–326. [Google Scholar] [CrossRef]
  45. Fukushima, K.; Nagai, K.; Hoshi, Y.; Masumoto, S.; Mikami, I.; Takahashi, Y.; Oike, H.; Kobori, M. Drosera rotundifolia and Drosera tokaiensis Suppress the Activation of HMC-1 Human Mast Cells. J. Ethnopharmacol. 2009, 125, 90–96. [Google Scholar] [CrossRef]
  46. Fan, D.-H.; Wang, H.; Zhi, D.; Shen, Y.-M. CE Analysis of Endogenous Flavonoid Gallate Esters from Nepenthes gracilis (Nepenthaceae). Chromatographia 2010, 72, 1013–1016. [Google Scholar] [CrossRef]
  47. Marczak, Ł.; Kawiak, A.; Łojkowska, E.; Stobiecki, M. Secondary Metabolites In in Vitro Cultured Plants of the Genus Drosera. Phytochem. Anal. 2005, 16, 143–149. [Google Scholar] [CrossRef] [PubMed]
  48. Ou-Yang, F.; Tsai, I.-H.; Tang, J.-Y.; Yen, C.-Y.; Cheng, Y.-B.; Farooqi, A.A.; Chen, S.-R.; Yu, S.-Y.; Kao, J.-K.; Chang, H.-W. Antiproliferation for Breast Cancer Cells by Ethyl Acetate Extract of Nepenthes thorellii x (ventricosa x maxima). Int. J. Mol. Sci. 2019, 20, 3238. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  49. Wong, C.; Ling, Y.S.; Wee, J.L.S.; Mujahid, A.; Müller, M. A Comparative UHPLC-Q/TOF–MS-Based Eco-Metabolomics Approach Reveals Temperature Adaptation of Four Nepenthes Species. Sci. Rep. 2020, 10, 21861. [Google Scholar] [CrossRef] [PubMed]
  50. Thanh, N.V.; Thao, N.P.; Huong, P.T.T.; Lee, S.H.; Jang, H.D.; Cuong, N.X.; Nam, N.H.; Kiem, P.V.; Kim, Y.H.; Minh, C.V. Naphthoquinone and Flavonoid Constituents from the Carnivorous Plant Nepenthes mirabilis and Their Anti-Osteoporotic and Antioxidant Activities. Phytochem. Lett. 2015, 11, 254–259. [Google Scholar] [CrossRef]
  51. Hake, A.; Begrow, F.; Spiegler, V.; Symma, N.; Hensel, A.; Düfer, M. Effects of Extracts and Flavonoids from Drosera rotundifolia L. on Ciliary Beat Frequency and Murine Airway Smooth Muscle. Molecules 2022, 27, 6622. [Google Scholar] [CrossRef]
  52. Mattioli, R.; Francioso, A.; Mosca, L.; Silva, P. Anthocyanins: A Comprehensive Review of Their Chemical Properties and Health Effects on Cardiovascular and Neurodegenerative Diseases. Molecules 2020, 25, 3809. [Google Scholar] [CrossRef]
  53. Bendokas, V.; Stanys, V.; Mažeikienė, I.; Trumbeckaite, S.; Baniene, R.; Liobikas, J. Anthocyanins: From the Field to the Antioxidants in the Body. Antioxidants 2020, 9, 819. [Google Scholar] [CrossRef]
  54. Timoneda, A.; Feng, T.; Sheehan, H.; Walker-Hale, N.; Pucker, B.; Lopez-Nieves, S.; Guo, R.; Brockington, S. The Evolution of Betalain Biosynthesis in Caryophyllales. New Phytol. 2019, 224, 71–85. [Google Scholar] [CrossRef] [Green Version]
  55. Foot, G.; Rice, S.P.; Millett, J. Red Trap Colour of the Carnivorous Plant Drosera rotundifolia Does Not Serve a Prey Attraction or Camouflage Function. Biol. Lett. 2014, 10, 20131024. [Google Scholar] [CrossRef] [Green Version]
  56. Dávila-Lara, A.; Reichelt, M.; Wang, D.; Vogel, H.; Mithöfer, A. Proof of Anthocyanins in the Carnivorous Plant Genus Nepenthes. FEBS Open Bio 2021, 11, 2576–2585. [Google Scholar] [CrossRef]
  57. Gregorio, G.J.D.; Dipalma, J.R. Anthocyanin in Dionaea muscipula Ellis (Venus Flytrap). Nature 1966, 212, 1264–1265. [Google Scholar] [CrossRef]
  58. Bendz, G.; Lindberg, G.; Buchardt, O.; Norin, T. Naphthoquinones and Anthocyanins from Two Drosera Species. Acta Chem. Scand. 1968, 22, 2722–2723. [Google Scholar] [CrossRef] [Green Version]
  59. Ichiishi, S.; Nagamitsu, T.; Kondo, Y.; Iwashina, T.; Kondo, K.; Tagashira, N. Effects of Macro-Components and Sucrose in the Medium on in Vitro Red-Color Pigmentation in Dionaea muscipula Ellis and Drosera spathulata Labill. Plant Biotechnol. 1999, 16, 235–238. [Google Scholar] [CrossRef]
  60. Henarejos-Escudero, P.; Guadarrama-Flores, B.; García-Carmona, F.; Gandía-Herrero, F. Digestive Glands Extraction and Precise Pigment Analysis Support the Exclusion of the Carnivorous Plant Dionaea muscipula Ellis from the Caryophyllales Order. Plant Sci. 2018, 274, 342–348. [Google Scholar] [CrossRef]
  61. Meyer, G.W.; Bahamon Naranjo, M.A.; Widhalm, J.R. Convergent Evolution of Plant Specialized 1,4-Naphthoquinones: Metabolism, Trafficking, and Resistance to Their Allelopathic Effects. J. Exp. Bot. 2021, 72, 167–176. [Google Scholar] [CrossRef]
  62. Schlauer, J.; Nerz, J.; Rischer, H. Carnivorous Plant Chemistry. Acta Bot. Gall. 2005, 152, 187–195. [Google Scholar] [CrossRef] [Green Version]
  63. Eilenberg, H.; Pnini-Cohen, S.; Rahamim, Y.; Sionov, E.; Segal, E.; Carmeli, S.; Zilberstein, A. Induced Production of Antifungal Naphthoquinones in the Pitchers of the Carnivorous Plant Nepenthes khasiana. J. Exp. Bot. 2010, 61, 911–922. [Google Scholar] [CrossRef] [Green Version]
  64. Raj, G.; Kurup, R.; Hussain, A.A.; Baby, S. Distribution of Naphthoquinones, Plumbagin, Droserone, and 5-O-Methyl Droserone in Chitin-Induced and Uninduced Nepenthes khasiana: Molecular Events in Prey Capture. J. Exp. Bot. 2011, 62, 5429–5436. [Google Scholar] [CrossRef] [Green Version]
  65. Liu, W.; Lin, L.-C.; Wang, P.-J.; Chen, Y.-N.; Wang, S.-C.; Chuang, Y.-T.; Tsai, I.-H.; Yu, S.-Y.; Chang, F.-R.; Cheng, Y.-B.; et al. Nepenthes Ethyl Acetate Extract Provides Oxidative Stress-Dependent Anti-Leukemia Effects. Antioxidants 2021, 10, 1410. [Google Scholar] [CrossRef]
  66. Guan, H.-H.; Huang, Y.-H.; Lin, E.-S.; Chen, C.-J.; Huang, C.-Y. Plumbagin, a Natural Product with Potent Anticancer Activities, Binds to and Inhibits Dihydroorotase, a Key Enzyme in Pyrimidine Biosynthesis. Int. J. Mol. Sci. 2021, 22, 6861. [Google Scholar] [CrossRef] [PubMed]
  67. Rahman-Soad, A.; Dávila-Lara, A.; Paetz, C.; Mithöfer, A. Plumbagin, a Potent Naphthoquinone from Nepenthes Plants with Growth Inhibiting and Larvicidal Activities. Molecules 2021, 26, 825. [Google Scholar] [CrossRef] [PubMed]
  68. Dávila-Lara, A.; Rahman-Soad, A.; Reichelt, M.; Mithöfer, A. Carnivorous Nepenthes x ventrata Plants Use a Naphthoquinone as Phytoanticipin against Herbivory. PLoS ONE 2021, 16, e0258235. [Google Scholar] [CrossRef] [PubMed]
  69. Grevenstuk, T.; Gonçalves, S.; Domingos, T.; Quintas, C.; van der Hooft, J.J.; Vervoort, J.; Romano, A. Inhibitory Activity of Plumbagin Produced by Drosera intermedia on Food Spoilage Fungi. J. Sci. Food Agric. 2012, 92, 1638–1642. [Google Scholar] [CrossRef]
  70. Tian, J.; Chen, Y.; Ma, B.; He, J.; Tong, J.; Wang, Y. Drosera peltata Smith var. lunata (Buch.-Ham.) C.B. Clarke as a Feasible Source of Plumbagin: Phytochemical Analysis and Antifungal Activity Assay. World J. Microbiol. Biotechnol. 2014, 30, 737–745. [Google Scholar] [CrossRef]
  71. Putalun, W.; Udomsin, O.; Yusakul, G.; Juengwatanatrakul, T.; Sakamoto, S.; Tanaka, H. Enhanced Plumbagin Production from in Vitro Cultures of Drosera burmanii Using Elicitation. Biotechnol. Lett. 2010, 32, 721–724. [Google Scholar] [CrossRef]
  72. Adamec, L.; Gastinel, L.; Schlauer, J. Plumbagin Content in Aldrovanda vesiculosa Shoots. Carniv. Plant Newslett. 2006, 35, 52–55. [Google Scholar]
  73. Kawiak, A.; Królicka, A.; Łojkowska, E. In Vitro Cultures of Drosera aliciae as a Source of a Cytotoxic Naphthoquinone: Ramentaceone. Biotechnol. Lett. 2011, 33, 2309–2316. [Google Scholar] [CrossRef]
  74. Kawiak, A.; Zawacka-Pankau, J.; Wasilewska, A.; Stasilojc, G.; Bigda, J.; Lojkowska, E. Induction of Apoptosis in HL-60 Cells through the ROS-Mediated Mitochondrial Pathway by Ramentaceone from Drosera aliciae. J. Nat. Prod. 2012, 75, 9–14. [Google Scholar] [CrossRef]
  75. Aung, H.H.; Chia, L.S.; Goh, N.K.; Chia, T.F.; Ahmed, A.A.; Pare, P.W.; Mabry, T.J. Phenolic Constituents from the Leaves of the Carnivorous Plant Nepenthes gracilis. Fitoterapia 2002, 73, 445–447. [Google Scholar] [CrossRef]
  76. Picazo-Aragonés, J.; Terrab, A.; Balao, F. Plant Volatile Organic Compounds Evolution: Transcriptional Regulation, Epigenetics and Polyploidy. Int. J. Mol. Sci. 2020, 21, 8956. [Google Scholar] [CrossRef]
  77. Di Giusto, B.; Bessière, J.-M.; Guéroult, M.; Lim, L.B.L.; Marshall, D.J.; Hossaert-McKey, M.; Gaume, L. Flower-Scent Mimicry Masks a Deadly Trap in the Carnivorous Plant Nepenthes rafflesiana: Flower-Scent Mimicry in Nepenthes rafflesiana. J. Ecol. 2010, 98, 845–856. [Google Scholar] [CrossRef]
  78. Wells, K. Pitchers of Nepenthes rajah Collect Faecal Droppings from Both Diurnal and Nocturnal Small Mammals and Emit Fruity Odour. J. Trop. Ecol. 2011, 27, 347–353. [Google Scholar] [CrossRef]
  79. Kreuzwieser, J.; Scheerer, U.; Kruse, J.; Burzlaff, T.; Honsel, A.; Alfarraj, S.; Georgiev, P.; Schnitzler, J.-P.; Ghirardo, A.; Kreuzer, I.; et al. The Venus Flytrap Attracts Insects by the Release of Volatile Organic Compounds. J. Exp. Bot. 2014, 65, 755–766. [Google Scholar] [CrossRef]
  80. Hatano, N.; Hamada, T. Proteomic Analysis of Secreted Protein Induced by a Component of Prey in Pitcher Fluid of the Carnivorous Plant Nepenthes alata. J. Proteom. 2012, 75, 4844–4852. [Google Scholar] [CrossRef]
  81. Rottloff, S.; Miguel, S.; Biteau, F.; Nisse, E.; Hammann, P.; Kuhn, L.; Chicher, J.; Bazile, V.; Gaume, L.; Mignard, B.; et al. Proteome Analysis of Digestive Fluids in Nepenthes Pitchers. Ann. Bot. 2016, 117, 479–495. [Google Scholar] [CrossRef] [Green Version]
  82. Jakšová, J.; Adamec, L.; Petřík, I.; Novák, O.; Šebela, M.; Pavlovič, A. Contrasting Effect of Prey Capture on Jasmonate Accumulation in Two Genera of Aquatic Carnivorous Plants (Aldrovanda, Utricularia). Plant Physiol. Biochem. 2021, 166, 459–465. [Google Scholar] [CrossRef]
  83. Buch, F.; Rott, M.; Rottloff, S.; Paetz, C.; Hilke, I.; Raessler, M.; Mithöfer, A. Secreted Pitfall-Trap Fluid of Carnivorous Nepenthes Plants Is Unsuitable for Microbial Growth. Ann. Bot. 2013, 111, 375–383. [Google Scholar] [CrossRef] [Green Version]
  84. Tang, J.; Peng, S.; Cheng, Y.; Wang, C.; Farooqi, A.A.; Yu, T.; Hou, M.; Wang, S.; Yen, C.; Chan, L.; et al. Ethyl Acetate Extract of Nepenthes adrianii x clipeata Induces Antiproliferation, Apoptosis, and DNA Damage against Oral Cancer Cells through Oxidative Stress. Environ. Toxicol. 2019, 34, 891–901. [Google Scholar] [CrossRef]
  85. Zehl, M.; Braunberger, C.; Conrad, J.; Crnogorac, M.; Krasteva, S.; Vogler, B.; Beifuss, U.; Krenn, L. Identification and Quantification of Flavonoids and Ellagic Acid Derivatives in Therapeutically Important Drosera Species by LC–DAD, LC–NMR, NMR, and LC–MS. Anal. Bioanal. Chem. 2011, 400, 2565–2576. [Google Scholar] [CrossRef]
  86. Krolicka, A.; Szpitter, A.; Maciag, M.; Biskup, E.; Gilgenast, E.; Romanik, G.; Kaminski, M.; Wegrzyn, G.; Lojkowska, E. Antibacterial and Antioxidant Activity of the Secondary Metabolites from in Vitro Cultures of the Alice sundew (Drosera aliciae). Biotechnol. Appl. Biochem. 2008, 53, 175–184. [Google Scholar] [CrossRef]
  87. Ismail, N.A.; Kamariah, A.S.; Lim, L.B.L.; Ahmad, N. Phytochemical and Pharmacological Evaluation of Methanolic Extracts of the Leaves of Nepenthes bicalcarata Hook. F. Int. J. Pharmacogn. Phytochem. Res. 2015, 7, 1127–1138. [Google Scholar]
  88. Lin, E.-S.; Huang, C.-Y. Cytotoxic Activities and the Allantoinase Inhibitory Effect of the Leaf Extract of the Carnivorous Pitcher Plant Nepenthes Miranda. Plants 2022, 11, 2265. [Google Scholar] [CrossRef] [PubMed]
  89. Rodzali, N.N.; Mydin, M.M. Antibacterial Activity of Leaves and Pitchers Extract of Nepenthes gracilis against Bacillus subtilis and Escherichia coli. J. Fundam. Appl. Sci. 2018, 9, 81–88. [Google Scholar] [CrossRef]
  90. Gerschler, S.; Guenther, S.; Schulze, C. Antibiofilm Activity of Sundew Species against Multidrug-Resistant Escherichia coli Strains. Int. J. Mol. Sci. 2022, 23, 13720. [Google Scholar] [CrossRef] [PubMed]
  91. Kawiak, A.; Domachowska, A.; Krolicka, A.; Smolarska, M.; Lojkowska, E. 3-Chloroplumbagin Induces Cell Death in Breast Cancer Cells Through MAPK-Mediated Mcl-1 Inhibition. Front. Pharmacol. 2019, 10, 784. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  92. De, U.; Son, J.Y.; Jeon, Y.; Ha, S.-Y.; Park, Y.J.; Yoon, S.; Ha, K.-T.; Choi, W.S.; Lee, B.M.; Kim, I.S.; et al. Plumbagin from a Tropical Pitcher Plant (Nepenthes alata Blanco) Induces Apoptotic Cell Death via a P53-Dependent Pathway in MCF-7 Human Breast Cancer Cells. Food Chem. Toxicol. 2019, 123, 492–500. [Google Scholar] [CrossRef]
  93. Gwee, P.S.; Khoo, K.S.; Ong, H.C.; Sit, N.W. Bioactivity-Guided Isolation and Structural Characterization of the Antifungal Compound, Plumbagin, from Nepenthes gracilis. Pharm. Biol. 2014, 52, 1526–1531. [Google Scholar] [CrossRef]
  94. Thao, N.P.; Luyen, B.T.T.; Koo, J.E.; Kim, S.; Koh, Y.S.; Thanh, N.V.; Cuong, N.X.; Kiem, P.V.; Minh, C.V.; Kim, Y.H. In Vitro Anti-Inflammatory Components Isolated from the Carnivorous Plant Nepenthes mirabilis (Lour.) Rafarin. Pharm. Biol. 2016, 54, 588–594. [Google Scholar] [CrossRef] [Green Version]
  95. Thanh, N.V.; Thao, N.P.; Dat, L.D.; Huong, P.T.T.; Lee, S.H.; Jang, H.D.; Cuong, N.X.; Nam, N.H.; Kiem, P.V.; Minh, C.V.; et al. Two New Naphthalene Glucosides and Other Bioactive Compounds from the Carnivorous Plant Nepenthes mirabilis. Arch. Pharm. Res. 2015, 38, 1774–1782. [Google Scholar] [CrossRef]
  96. Babula, P.; Adam, V.; Havel, L.; Kizek, R. Naftochinony a Jejich Farmakologické Vlastnosti [Naphthoquinones and Their Pharmacological Properties]. Ceska Slov. Farm. 2007, 56, 114–120. [Google Scholar]
  97. Liu, Y.; Cai, Y.; He, C.; Chen, M.; Li, H. Anticancer Properties and Pharmaceutical Applications of Plumbagin: A Review. Am. J. Chin. Med. 2017, 45, 423–441. [Google Scholar] [CrossRef]
  98. Płachno, B.J.; Kapusta, M.; Stolarczyk, P.; Świątek, P. Arabinogalactan Proteins in the Digestive Glands of Dionaea muscipula J.Ellis Traps. Cells 2022, 11, 586. [Google Scholar] [CrossRef]
  99. Płachno, B.J.; Kapusta, M.; Stolarczyk, P.; Świątek, P.; Strzemski, M.; Miranda, V.F.O. Immunocytochemical Analysis of the Wall Ingrowths in the Digestive Gland Transfer Cells in Aldrovanda vesiculosa L. (Droseraceae). Cells 2022, 11, 2218. [Google Scholar] [CrossRef]
  100. Rey, M.; Yang, M.; Lee, L.; Zhang, Y.; Sheff, J.G.; Sensen, C.W.; Mrazek, H.; Halada, P.; Man, P.; McCarville, J.L.; et al. Addressing Proteolytic Efficiency in Enzymatic Degradation Therapy for Celiac Disease. Sci. Rep. 2016, 6, 30980. [Google Scholar] [CrossRef]
  101. Mithöfer, A. A Spotlight on Prey-Induced Metabolite Dynamics in Sundew. A Commentary on: ‘Metabolomic Analysis Reveals Reliance on Secondary Plant Metabolites to Facilitate Carnivory in the Cape Sundew, Drosera capensis. Ann. Bot. 2021, 128, v–vi. [Google Scholar] [CrossRef]
  102. Iosip, A.-L.; Scherzer, S.; Bauer, S.; Becker, D.; Krischke, M.; Al-Rasheid, K.A.S.; Schultz, J.; Kreuzer, I.; Hedrich, R. DYSCALCULIA, a Venus Flytrap Mutant without the Ability to Count Action Potentials. Curr. Biol. 2023, 33, 589–596.e5. [Google Scholar] [CrossRef]
  103. Wawrosch, C. An Improved 2-Step Liquid Culture System for Efficient In Vitro Shoot Proliferation of Sundew (Drosera rotundifolia L.). Sci. Pharm. 2009, 77, 827–836. [Google Scholar] [CrossRef] [Green Version]
  104. Wawrosch, C.; Vackar, E.; Grauwald, B. Variations of Naphthoquinone Levels in Micropropaaated Drosera Species In Vitro, under Qreenhouse and Outdoor Growth Conditions. Sci. Pharm. 2005, 73, 251–262. [Google Scholar] [CrossRef] [Green Version]
Molecules 28 02155 g001 550
Figure 1. Examples of plants from Nepenthaceae and Droseraceae. (A) Nepenthes cv. ‘Miranda’, (B) Drosera binata Labill. var. dichotoma, (C) Drosera capensis L., (D) Dionaea muscipula J.Ellis.
Figure 1. Examples of plants from Nepenthaceae and Droseraceae. (A) Nepenthes cv. ‘Miranda’, (B) Drosera binata Labill. var. dichotoma, (C) Drosera capensis L., (D) Dionaea muscipula J.Ellis.
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Figure 2. Structures of the most common derivatives of (a) benzoic acid, (b) cinnamic acid, and (c) ellagic acid found in Nepenthaceae and Droseraceae.
Figure 2. Structures of the most common derivatives of (a) benzoic acid, (b) cinnamic acid, and (c) ellagic acid found in Nepenthaceae and Droseraceae.
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Figure 3. General structure of flavonols (a) quercetin, (b) kaempferol, and (c) myricetin; R—glycoside moiety.
Figure 3. General structure of flavonols (a) quercetin, (b) kaempferol, and (c) myricetin; R—glycoside moiety.
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Figure 4. Structure of the most common anthocyanins: (a) cyanidin derivatives; (b) delphinidin derivatives found in Nepenthaceae and Droseraceae species.
Figure 4. Structure of the most common anthocyanins: (a) cyanidin derivatives; (b) delphinidin derivatives found in Nepenthaceae and Droseraceae species.
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Figure 5. Structure of the most common 1,4-naphthoquinone derivatives identified in Nepenthaceae and Droseraceae.
Figure 5. Structure of the most common 1,4-naphthoquinone derivatives identified in Nepenthaceae and Droseraceae.
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Figure 6. Structure of shinanolone (a) and isoshinanolone (b).
Figure 6. Structure of shinanolone (a) and isoshinanolone (b).
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Table
Table 1. Secondary metabolites identified in Nepenthes.
Table 1. Secondary metabolites identified in Nepenthes.
Investigated SpeciesCompoundMethodRef.
phenols/phenolic acids and derivatives
N. anamensis (leaf, trap)gallic: 58.9 (l), 53.7 (t) µg/g; protocatechuic: 4.48 (l), 15.1 (t) µg/g; hydroxybenzoic: 4.28 (l), 9.21 (t) µg/g; vanillic: 2.53 (l), 9.93 (f) µg/g; chlorogenic: 32.4 (l) 53.8 (f) µg/g; syringic: 10.7 (l), 14.7 (f) µg/g; caffeic: 2.6 (l), 3.57 (f) µg/g; ferulic: 1.07 (l), 7.97 (f) µg/g; sinapic: 7.09 (l) 15.7 (f) µg/g; p-coumaric 2.08 (l), 6.77 (t) µg/g; vanillin: 0.85 (l), 0.94 (f) µg/g dwHPLC-MS[32]
flavonoids: flavonols
N. minima, N. ampullaria, N. rafflesiana, N. northianaquercetin (Q), Q 3-O-rhamnoside, Q 3-(6″-galloylglucoside),
Q 3-O-(6″-n-butyl ß-D-glucuronide), rutin, miquelianin, kaempferol (K) 3-glucoside, K 3-O-beta-D-xyloside, 6,8-dihydroxy-K 3-rutinoside, afzelin, luteolin, baicalin, butin, myrciacitrin		UHPLC-Q/TOF–MS[49]
N. gracilis
(leaves)		quercetin esters: 3-O-ß-(3″-O-galloylxylopyranoside),
3-O-α-(3″-O-galloylrhamnopyranoside), 3-O-ß-(2″-O-galloylxylopyranoside),
3-O-α-(2″-galloylarabinofuranoside), 3-O-ß-(6″-galloylglucopyranoside)		CC/NMR, MS[46]
N. thorellii × (ventricosa × maxima) (aerial parts)quercetin 3-O-(6″-n-butyl-D-glucuronide) HPLC-DAD, CC/NMR[48]
N. mirabilis
(branches and leaves)		quercitrin, quercetin (Q), Q-3-O-b-D-glucuronide,
K-3-O-a-L-rhamnoside 		CC/NMR, MS[50]
flavonoids: anthocyanins *
N. × ventrata, N. thorelii, N. ventricosa, N. robcantleyi, N. maxima, N. fusca, N. mirabilis (pitchers: digestive zone—dz, peristome—p)cyanidin-3-O-glucoside: ca 0.5–6 (p), up to ca 2 (dz) µM/g fw
cyanidin-3-O-galactoside: ca 0.1–0.6 (p), up to ca 0.3 (dz) µM/g fw
cyanidin-3-O-glucuronide: ca 0–0.1 (p, dz) µM/g fw 		UHPLC/HRMS[56]
other phenolic compounds
N. mirabili (branches and leaves)sepicatechin CC/NMR, MS[50]
N. gracilis (leaves)epicatechin 3-O-gallate CC/NMR, MS[46]
N. minima, N. ampullaria, N. rafflesiana, N. northiana (aerial parts)syringin, catechin 5-O-gallate, coniferin,
5-galloylshikimic acid 		UHPLC-Q/TOF–MS[49]
naphthoquinones
N. cv. ‘Miranda’ (leaves)plumbagin GC-MS[66]
Nepenthes × ventrata (N. alata × ventricosa) (pitcher, leaves)plumbagin NMR[67]
Nepenthes thorellii × (ventricosa × maxima) (aerial parts)plumbagin HPLC-DAD, CC/NMR[48]
N. × ventrata (N. alata × N. ventricosa) (leaves)plumbagin LC-MS/MS[68]
N. khasiana
(root, stem, leaves, pitchers),		plumbagin: ca 1.4% (r), 0.2% (s), 0.4% (l), 0.04% (pi) dw *
droserone, 5-O-methyl droserone (pi)		GC-MS/NMR[64]
N. khasiana (pitcher liquid)5-O-methyl droserone, droserone HPLC/MS, UV, NMR[63]
N. mirabilis (branches and leaves)3-methoxy-7-methyljuglone2-methoxy-7-methyljuglone, plumbagin, droserone,
nepenthones C, F and G		CC/NMR, MS[50]
N. alata, N. fusca, N. gracilis, N. mirabilis, N. superba, N. thorelii,
N. ventricosa (pitcher fluids)		plumbagin, 7-methyl-juglone LC/MS, NMR[83]
N. adrianii × clipeata
(twigs and leaves)		isoplumbagin Isolation/NMR[84]
acetogenic tetralone
N. gracilis (leaves),Isoshinanolone, shinanolone epishinanolone Isolation/MS, NMR[75]
N. thorellii × (ventricosa × maxima) (aerial parts)isoshinanolone HPLC-DAD, NMR[48]
N. mirabilis
(branches and leaves)		cis-isoshinanolone CC/NMR, MS[50]
alkaloids
N. minima, N. ampullaria, N. rafflesiana, N. northiana (aerial parts)trigonelline, anatoxin a, berberastine UHPLC-Q/TOF–MS[49]
* values were estimated from the graph; Q—quercetin; K—kaempferol; (dz)—digestive zone; (p)—peristome; (r)—root; (s)—stem; (l)—leaf; (t)—trap; (pi)—pitcher; fw—fresh weight; dw-dried weight.
Table
Table 2. Secondary metabolites identified in Drosera.
Table 2. Secondary metabolites identified in Drosera.
SpeciesCompoundMethodRefs.
phenols/phenolic acids and derivatives
Dr. capensis (leaf, trap)gallic: 167.8 (l), 195.5 (t) µg/g; protocatechuic: 1.26 (l), 12.1 (t) µg/g; chlorogenic: 28.5 (l), 29.6 (t) µg/g; ferulic: 1.15 (l), 2.21 (t) µg/g; p-coumaric: 1.52 (l), 2.06 (t) µg/g; hydroxybenzoic: 3.72 (l), 17.5 (t) µg/g; vanillic: 1.57 (l) 2.07 (t) µg/g; syringic: 7.86 (l) 10.1 (t) µg/g; caffeic: 1.91 (l) 3.01 (t) µg/g; sinapic: 0.18 (l), 0.69 (t) µg/g; vanillin: 1.93 (l), 1.62 (t) µg/g dwHPLC-MS[32]
Dr. rotundifolia, Dr. tokaiensis,
Dr. spatulata		ellagic acid (EA)HPLC[45]
Dr. tokaiensis (aboveground part)3,3-di-O-methylellagic acid 4′-glucoside HPLC/NMR[34]
Dr. magna (bulbs and leaves)1-O-protochatechoyl glucoside Isolation/NMR[33]
Dr. rotundifolia
field (f), in vitro propagated (p)		quinic acid (fp), monogalloyl glucose (fp), digalloyl glucose (f), coumaric acid glycoside (fp), EA (fp), EA glycoside (fp), dimethylEA glycoside ((fp), dimethylEA (fp), methylEA (f)UPLC/DAD/MS-MS[20]
Dr. anglica, Dr. intermedia,
Dr. madagascariensis, Dr. rotundifolia (aerial part)		ellagic acid (0.137–1.107%), 3,3′-di-O-methylEA (0.084–0.143%) dwLC-MS/NMR[85]
Dr. binata in vitro culture3,3-di-O-methylellagic acidHPLC-DAD-MS[36]
flavonoids
Dr. binatamethylated myricetin (M) methylated M 3-O-glucoside, quercetin (Q), Q 3-O-glucoside;
isorhamnetin (iRh) or rhamnetin (Rh); Rh/isoRh 3-O-glucoside; kaempferol (K)/fisetin (F)
K/F 3-O-glucoside; K/F-3-O-rhamnosylglucoside,		HPLC-UV/MS[47]
Dr. rotundifolia field (f), in vitro propagated (p)myricetin glycoside (fp), hyperoside (fp), galloylhyperoside (fp), hydroxybenzoylhyperin (fp),
dihydromyricetin (f), hexahydroxyflavonegalloyl glycoside (f), tetrahydroxyflavone (f), kaempferol-galloylglycoside (f), quercetin (Q)(f), Q-glycoside (f) Q-glycoside gallate (f), syringetin glycoside (p), spinatoside (p)		UPLC/DAD/MS[20]
Dr. rotundifoliaquercetin, hyperoside and 2″-O-galloylhyperosideUHPLC-TOF-MS[51]
Dr. tokaiensis (aboveground part)myricitrine, quercimelin HPLC/NMR[34]
Dr. magna
(bulbs and leaves)		tamarixetin-3-rhamnoside, naringenin-6-C-β-D-glucopyranoside, hirsutrin, three new flavonol diglycosides, flavan-3-ol glycoside Isolation/NMR[33]
Dr. rotundifolia Dr. anglica, Dr. intermedia,
Dr. madagascariensis,
(aerial part)		myricetin (M) (<0.003–0.097%), M-3-O-β-galactopyranoside (0.038–0.275%), M-3-O-β-glucopyranoside, hyperoside (H) (0.048–1.530%), 2″-O-galloylH (0–2.515%), isoquercitrin (0.032–0.421%), kaempferol (K), K-3-O-β-galactopyranoside, K-3-O-(2″-O-galloyl)-β-galactopyranoside, astragalin, quercetin (0.056–0.187%) dwLC-MS/NMR[85]
naphthoquinones
Dr. binata, Dr. adelae, Dr. aliciae, Dr. capensis, Dr. cuneifolia, Dr. ramentaceaplumbagin: 0.001–0.059% fwHPLC-UV/MS[47]
Dr. binata, Dr. giganteaplumbagin: 2.04 and 0.15 mg/g fwHPLC-DAD-MS[35]
Dr. binataplumbagin, 3-chloroplumbaginHPLC-DAD-MS[36]
Dr. intermedia,plumbaginHPLC-MS/NMR[69]
Dr. burmaniiplumbaginELISA[71]
Dr. peltata Smith var. lunataplumbagin: 11.05 mg/g dwTLC/NMR[70]
Dr. magna (bulbs and leaves)hydroxydroserone (H), H-5-O-β-D-glucoside, droserone, plumbagin, 2,3-dihydronapthalene-1,4-diones Isolation/NMR[33]
Dr. rotundifolia
field (f), in vitro propagated (p)		7-methyljuglone, 7-methyljuglone diglycoside, 7-methyljuglone glycoside (f,p)UPLC/DAD/MS-MS[20]
Dr. aliciae in vitro cultureramentaceoneIsolation/NMR, MS[73,74]
* Q—quercetin; K—kaempferol; M—myricetin; EA—ellagic acid; fw—fresh weight; dw—dried weight; (l)—leaf; (t)—trap.
Table
Table 3. Secondary metabolites identified in Dionaea.
Table 3. Secondary metabolites identified in Dionaea.
SpeciesCompoundMethodRef.
phenols/phenolic acids and derivatives
Di. muscipula
(leaf, trap)		gallic: 187.8 (l), 167.4 (t) µg/g; protocatechuic: 2.74 (l), 3.21 (t) µg/g; chlorogenic: 24.5 (l), 30.8 (t) µg/g; ferulic: 0.65 (l), 0.69 (t) µg/g; p-coumaric: 0.57 (l), 0.76 (t) µg/g; salicylic: 0.28 (l), 0.19 (t) µg/g; hydroxybenzoic: 4.26 (l), 4.53 (t) µg/g; vanillic: 4.84 (l), 6.87 (t) µg/g; syringic: 8.78 (l), 12.2 (t) µg/g; caffeic: 10.8 (l), 12.0 (t) µg/g; sinapic: 17.0 (l), 42.2 (t) µg/g; vanillin: 2.45 (l), 1.97 (t) µg/g dwHPLC-MS[32]
Di. muscipulaellagic, dimethylellagic acid isomer, 3-O-methylellagic acid, 3,3′-di-O-methylellagic acidHPLC-DAD-MS[35]
Di. muscipulacaffeic: ca 0.18 mg/g, salicylic: ca 300 mg/g, ellagic: ca 0.18 mg/g *HPLC-DAD[37]
Di. muscipulachlorogenic:0.26 mg/g, p-coumaric: 0.04 mg/g ferulic: 0.16 mg/g, gallic: 0.31 mg/g,
protocatechuic: 0.29 mg/g dw		HPLC-DAD[22]
flavonoids: flavonols
Di. muscipulamyricetin: ca 20 mg/g, hyperoside: ca 0.3 mg/g, quercetin: 15 mg/g dw *HPLC-DAD[37]
Di. muscipulakaempferol, quercetin after hydrolysis (leaf, trap)HPLC-MS[32]
Di. muscipulakaempferol: 0.59 mg/g dwHPLC-DAD[22]
Di. muscipulahyperoside, quercetin-3-(6″-O-galloyl)-glucoside/galactoside, kaempferol-3-(6″-O-galloyl)-glucosideHPLC-DAD-MS[35]
flavonoids: anthocyanins
Di. muscipula (digestive glands)delphinidin-3-O-glucoside, cyanidin-3-O-glucoside,
cyanidin 		HPLC/MS-MS[60]
naphthoquinones
Di. muscipulaplumbagin: 3.45 mg/g fw, dihydroplumbagin, 3-chloroplumbagin, 8,8′-biplumbaginHPLC/DAD-MS[35]
Di. muscipulaplumbagin: ca 50 mg/g dw *HPLC-DAD[37]
* values were estimated from the graph; dw—dried weight; (l)—leaf; (t)—trap.
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Wójciak, M.; Feldo, M.; Stolarczyk, P.; Płachno, B.J. Carnivorous Plants from Nepenthaceae and Droseraceae as a Source of Secondary Metabolites. Molecules 2023, 28, 2155. https://doi.org/10.3390/molecules28052155

AMA Style

Wójciak M, Feldo M, Stolarczyk P, Płachno BJ. Carnivorous Plants from Nepenthaceae and Droseraceae as a Source of Secondary Metabolites. Molecules. 2023; 28(5):2155. https://doi.org/10.3390/molecules28052155

Chicago/Turabian Style

Wójciak, Magdalena, Marcin Feldo, Piotr Stolarczyk, and Bartosz J. Płachno. 2023. "Carnivorous Plants from Nepenthaceae and Droseraceae as a Source of Secondary Metabolites" Molecules 28, no. 5: 2155. https://doi.org/10.3390/molecules28052155

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