Skip to main content
NCBI home page
Wiley Open Access Collection logoLink to Wiley Open Access Collection
. 2026 Jan 8;23(1):e01810. doi: 10.1002/cbdv.202501810

Sesquiterpene‐rich Essential Oil From Polygonum pubescens: Chemical Composition and Phytotoxicity Study

Sajjad Hussain 1,2, Tarani Biswas 2,3, Parthiv Kar 2,3, Siddhartha Proteem Saikia 1,2, Saikat Haldar 2,3,✉
PMCID: PMC13419962  PMID: 41504083

ABSTRACT

Polygonum pubescens, known as drooping or lax knotweed is a tropical and subtropical annual herb belonging to Polygonaceae. The weedy and herbaceous nature, wide distribution and availability make this herb an attractive target for useful phytochemicals. This study aimed to investigate the volatile metabolites of this herb which remained unexplored. First, a detailed morphological study established its identity and distinguishing features with closely related Polygonum species. The leaves of this herb produced highest yield of essential oil (0.20%–0.35%) among three aerial parts (leaf, stem, and inflorescence) hydrodistilled. The oil composition was grossly similar across the tissues and exclusively dominated by three eudesmane and cadalane‐type sesquiterpene hydrocarbons, namely, β‐selinene (36.3%–43.7%, 221.8–254.5 mg/g), α‐selinene (11.0%–14.4%, 74.2–86.5 mg/g), and δ‐cadinene (29.0%–41.6%, 140.6–197.6 mg/g). This oil was practically composed of sesquiterpenoids which collectively contributed 97.9%–100.0% relative abundance. When phytotoxicity was evaluated, the oil dose‐dependently (1.0–200.0 µg/mL) inhibited the growth of wheatgrass coleoptile. However, the inhibitory potency (59%–73%, 200.0 µg/mL) was significantly low for its consideration as an herbicidal candidate. The current study identified P. pubescens as a rich natural source of β‐selinene and δ‐cadinene, bearing significance in plant defense, chemical synthon and bioactive phytochemicals.

Keywords: Aromatic plant, Weed, Polygonaceae, Persicaria, Sesquiterpenoids, Phytotoxicity

graphic file with name CBDV-23-e01810-g006.jpg

1. Introduction

Polygonum pubescens Blume (Syn. Polygonum flaccidum Meisn., Persicaria pubescens Blume), commonly known as drooping knotweed, lax knotweed or nonspicy smartweed is an annual herb belonging to Polygonaceae [1, 2, 3]. It is predominantly distributed in the tropical and subtropical regions of Indian subcontinent, China, Japan, and southeast Asian countries. This weed species thrives in wet habitats such as swamps, agricultural fields and riverside. The plant is normally characterized by its purplish‐red flowers and drooping inflorescence, justifying its name. It is known as analgesic, anti‐inflammatory, diuretic, purgative, insecticidal and an antidote to snake bites in Bangladesh [3, 4]. The indigenous people of Arunachal Pradesh in northeast India use this herbal paste to stupefy and trap the fishes in the water streams [5]. In a study, α‐santalone, isolated from its aerial part, was reported to reduce acetic acid‐induced writhing (analgesic) and increase urinary excretion (diuretic) in mice [4]. The methanolic extract of this herb improved cognitive function (nootropic) and oxidative stress in rats by elevating the antioxidant markers in brain and reducing acetylcholinesterase activity [6]. Thus, modern scientific studies have validated its ethnomedicinal uses on several occasions. P. pubescens is also known as a manganese hyperaccumulator and demonstrated its effectiveness in phytoremediation of manganese‐contaminated soil [7, 8].

As a part of ongoing research in our laboratory on the bioprospection of under‐utilized medicinal and aromatic plants of northeast India [9, 10, 11, 12], the metabolites of P. pubescens were targeted. The genus Polygonum is a rich reserve of secondary metabolites in nature, and these species often bear volatile oil [13, 14, 15, 16, 17, 18]. Contextually, the authors hypothesized that P. pubescens may be a rich source of volatile and semi‐volatile metabolites. Although a number of sesquiterpenoids, phenylpropanoids, sterols, and flavonoids have been isolated and characterized from this herb; surprisingly, the essential oil content and its composition remained unexplored [3, 19, 20, 21]. Second, being an invasive and ubiquitous weed, its large‐scale availability for a sustainable future application is not a concern [5]. Therefore, the aims of the current study were (i) chemical profiling of essential oil from different aerial tissues of P. pubescens, (ii) identifying its importance and value from a chemical perspective, and (iii) assessing its phytotoxicity in vitro. There are various biological mechanisms acquired by the weed species which facilitate their growth, invasion and adaptability in different conditions [22]. Secondary metabolites released by these species sometimes act as allelochemicals to reduce interspecific competition in the ecosystem. The weedy nature of P. pubescens tempted the authors to evaluate the phytotoxicity of its essential oil.

2. Materials and Methods

2.1. Chemicals

The HPLC grade solvents, β‐caryophyllene, allyl alcohol and deuterated chloroform were procured from Sigma‐Aldrich (St. Louis, MO, USA). Tween 80, buffer components and sucrose were obtained from Avra Synthesis (Hyderabad, India). Water for the assays was collected from Milli‐Q water purification system. The organic seeds of wheatgrass (Triticum aestivum) for the phytotoxicity assay were purchased locally.

2.2. Collection of the Plant Material

The samples of P. pubescens were collected from the vicinities of CSIR‐North East Institute of Science and Technology (NEIST), Jorhat, Assam, India (Lat 26.736547° Long 94.153393°, altitude 94 m) during October–November 2024 (flowering and fruiting season). Three batches of samples were collected on different days. The plant material was identified by Sajjad Hussain, a botanist and first author of this article. The methods of identification and taxonomic details have been described in the following sections. The specimen of P. pubescens was deposited at the herbarium of CSIR‐NEIST, Jorhat, India with a voucher specimen no. 0833. The morphological traits of the collected sample were recorded; aerial tissues (leaves, stem, and inflorescence) were segregated and stored at −20°C prior subjecting to the hydrodistillation.

2.3. Taxonomic Identification

The comprehensive online reference databases (eflora of China, eflora of India, eflora of Pakistan) were consulted for the comparison of morphological traits. Taxonomic revisionary reports, monographic literatures and scientific articles specifically focused on the taxonomy of family Polygonaceae and genus Polygonum were investigated thoroughly. Herbarium specimens from online herbarium databases and repositories (Naturalis Biodiversity Center, Netherlands; Royal Botanic Gardens (RBG), London, UK) were also studied to confirm the identity of P. pubescens.

2.4. Essential Oil Extraction

Three different fresh aerial tissues of P. pubescens, namely, leaves, stem, and inflorescence were individually chopped and subjected to hydrodistillation in Clevenger‐type apparatus for 6 h (3 tissues × 3 batches of collection) [10, 11, 12]. The light yellow‐colored oil with a dominant herbal note which was lighter than water was separated from the collection arm. A complete isolation of the oil samples was ensured through diethyl ether wash. The separated oil samples were passed through anhydrous sodium sulfate, solvent was removed and the weights were recorded. The oil yield was represented in % w/w and calculated as (oil extracted in g/fresh weight of the plant sample in g) × 100%.

2.5. Gas Chromatography–Mass Spectrometry

The gas chromatography–mass spectrometry (GC–MS) analyses were carried out in an Agilent 8890 gas chromatograph equipped with Agilent 7010B triple quadrupole mass spectrometer [10]. The oil samples were prepared in HPLC grade ethyl acetate (∼300 ppm) for the profiling. The split/splitless injector was maintained at 250°C throughout the run with a split ratio 10:1. An HP‐5MS GC column (30 m × 0.32 mm × 0.25 µm) and helium (carrier gas, 1.5 mL/min) were used for the separation of analytes. The temperature gradient program (28.75 min) was as follows: (i) 50°C for 1.0 min, (ii) then increased to 200°C with a ramp of 8°C/min and maintained for 2.0 min, (iii) then to 260°C at a rate of 15°C/min and held for 3.0 min. PAL3 autosampler was used for the automatic injection (0.5 µL) of the samples. The data were processed in Agilent Mass Hunter Qualitative Analysis 10.0 software integrated with NIST2017 mass spectral library. The retention indices (RI) of the identified peaks were calculated in reference to a series of linear C8–C28 n‐alkanes. The peak area sum percentage was considered as the relative abundance (%) of individual metabolites in the oil samples. It was represented as the range of percentages obtained through the analysis of three individual batches of samples.

The semi‐quantitative analysis was performed with respect to a known concentration of β‐caryophyllene as the internal standard (IS). The assay mixture contained 500 ppm oil sample and 100 ppm IS which was injected manually (1.0 µL) in triplicate under the same GC–MS conditions. The concentration of individual constituents was calculated using the equation: (PAA × CIS)/PAIS; where PAA = peak area of the analyte, PAIS = peak area of the IS, and CIS = concentration of the IS. The concentration was represented in mg/g of the oil.

2.6. Nuclear Magnetic Resonance Spectroscopy

The nuclear magnetic resonance (NMR) spectroscopy data were recorded in a Jeol 400YH FT NMR spectrometer (1H: 400 MHz, 13C: 100 MHz). The leaf essential oil sample was dissolved in deuterated chloroform for data recording. The solvent signals at δ H 7.26 ppm and δ C 77.0 ppm were used as the reference for 1H and 13C NMR data analysis, respectively. The data were visualized and analyzed in MestReNova software.

2.7. Wheatgrass Coleoptile Bioassay

Wheatgrass coleoptile bioassays were conducted separately for the oil samples of P. pubescens (leaves, stem, and inflorescence) to evaluate their phytotoxic potential [10]. The assays, set up in 5 mL glass vials contained eight different concentrations of the oil samples (1.0, 2.0, 5.0, 10.0, 25.0, 50.0, 100.0, 200.0 µg/mL), 0.5% v/v HPLC grade methanol and 0.1% v/v Tween 80 in phosphate‐citrate buffer of pH 5.0 added with 2.0% w/v sucrose in 2.0 mL assay volume. First, the oil samples prepared in HPLC grade methanol (10 µL) were taken in the vials followed by Tween 80 and phosphate‐citrate buffer. This mixture was vortexed vigorously for homogeneous mixing and then, added with eight instantly cut coleoptiles (4.0 mm) obtained from germinated (2.5 days) wheatgrass seeds. The control vials contained no oil samples. The assays were set up in duplicate. The vials were incubated at 23 ± 2°C for 2 days under 12/12 h light/dark photoperiod with occasional swirling. Then, the length of the coleoptiles in the assays was measured and % inhibition was calculated as [(C − T)/C] × 100%, where C is the average increase of coleoptile length in the control and T represents average increase in the treated sample.

2.8. Statistical Analysis

The analytical and bioassay results were tabulated in Microsoft Office Excel 2007 (Microsoft Corporation, Redmond, WA, USA) and the graphs were prepared in GraphPad Prism 8.0.2 software (GraphPad Software Inc., San Diego, CA, USA). The semi‐quantification (GC–MS) and % inhibition (coleoptile bioassay) data were represented as mean ± standard deviation (SD). A one‐way analysis of variance (ANOVA) followed by Tukey's post hoc test was employed in GraphPad Prism for the analysis of statistically significant difference in each treated samples compared to the control in coleoptile bioassay.

3. Results and Discussion

3.1. Taxonomic Authentication of P. pubescens

The genus Polygonum exhibits considerable variability in both vegetative and reproductive traits. The redundancy in vegetative and reproductive traits and wide distribution of these species increase the likelihood of conflicting or erroneous recognition. An accurate identification of Polygonum species requires careful and thorough taxonomic analysis so as to avoid misidentification and ensure accuracy. Therefore, in the current study, the identity of P. pubescens was validated based on following morphological and behavioral studies (Figure 1).

FIGURE 1.

FIGURE 1

Open in a new tab

Taxonomic plate of Polygonum pubescens: (A) whole plant (scale bar = 4.5 cm); (B, C) habitat: damp, water‐logged marshy areas; (D) stem (scale bar = 4.5 cm); (E) ocrea (scale bar = 4.0 mm); (F, G) leaf adaxial and abaxial surfaces (scale bar = 2.5 cm); (H) root (scale bar = 6.6 cm); (I) inflorescence (scale bar = 2.0 cm); (J) bract (scale bar = 1.1 mm); (K) flower (scale bar = 1.0 mm); (L) dissected flower showing eight stamens (scale bar = 1.2 mm, stamens are indicated with yellow arrows); (M) solitary carpel (scale bar = 1.2 mm); (N) an achene (scale bar = 1.8 mm).

Description: Annual erect herb, monoecious, 40‐ to 95‐cm tall, found in damp, water‐logged marshy areas. Stem: Slender, reddish, 30–80 cm, nodes swollen, hispidulous. Ocrea: pale red, membranous, hairy, tubular, stoutly ciliate, 1 to 2 cm long. Leaves: Simple, alternate, petiolate, ovate‐lanceolate in shape, 5–10 × 1.5–2.0 cm, both surfaces densely hispidulous; veins parallel, base cuneate, apex pointed, margin ciliate or fringed. Inflorescence: Terminal or sub‐terminal, spicate, drooping or pendulous, 7 to 15 cm long, 0.5 cm broad, interrupted below. Bracts: Green, funnel‐shaped, membranous, ciliate apex, 4–5 mm. Flowers: Bisexual, five‐parted, pedicellate with longer pedicels than bracts. Perianth: Green below, red above, tepals elliptic and punctate. Stamens: 8 and included. Styles: 3, fused at the middle, solitary carpel with hyaline style and pale green ovary, 1.5–2.5 mm. Seed: Trigonous, deep brown, densely pitted, 2.0–2.5 mm. Phenology: Vegetative growth period: August–October, flowering and fruiting period: October–November.

Several species of Polygonum including P. hydropiper, P. posumbu, P. longiseta, and P. maculosa share certain morphological characteristics with P. pubescens, notably terminal or axillary spicate inflorescences and pink‐colored, persistent perianth. Despite these similarities, certain unique features make P. pubescens distinguished. For instance, in spite of a similar lax and slender arrangement of the inflorescence, P. pubescens was differentiated from P. hydropiper on the basis of its reddish hue of the stem and lack of pungency in the leaves. Despite a similar leaf shape, P. posumbu can be distinguished by the lack of characteristic punctate pattern found on the perianth of P. pubescens. Furthermore, P. longiseta can be identified by the bristled margins on the flower bracts and at the apex of ocrea, a characteristic that is absent in P. pubescens. P. maculosa can be differentiated by the distinct dark triangular or lunate blotch in its leaves, despite other morphological resemblances like reddish‐brown stem and pink‐red inflorescence. P. glabrum is another species bearing morphological similarities to P. pubescens such as spicate inflorescence or pinkish perianth. However, lax and interrupted inflorescence of P. pubescens was in contrast to the dense and aggregated inflorescence in P. glabrum. These morphological and structural differences were critical in distinguishing P. pubescens from its closely related species.

3.2. Essential oil Profile of P. pubescens Aerial Tissues

3.2.1. Essential Oil Productivity

Essential oil productivity and composition vary with the plant species or subspecies, their tissues, ecological conditions, collection batch, postharvest processing, extraction techniques and associated parameters. Three individual aerial tissues of P. pubescens, namely, leaves, stem, and inflorescence obtained from three batches of collection (3 tissues × 3 batches) were extracted in fresh condition and a variation was clearly visible in their oil productivity. The investigation was conducted with three independent batches of samples to test their batch‐dependent variation and consistency in oil productivity and composition. Leaves produced highest yield of essential oil ranging 0.20%–0.35% w/w across three batches of samples, followed by inflorescence (0.12%–0.14%). The oil productivity of stem was relatively very low, furnishing in the range 0.012%–0.019% only. According to the literature reports, the essential oil yield of Polygonum species (e.g., P. odorata, P. minus, P. hydropiper, P. bistorta, P. orientale) varies in the range 0.01%–0.30% with respect to fresh weights of the aerial tissues [15, 23, 24, 25, 26]. Considering that, the oil productivity of P. pubescens was on the higher side of the usual yield range within the genus, indicating its potential for practical applications.

3.2.2. Essential Oil Composition

The GC–MS profile of the P. pubescens oil showed the presence of three major sesquiterpene hydrocarbons which were identified as β‐selinene (β‐eudesmene, 1), α‐selinene (2), and δ‐cadinene (3) (Figure 2). Selinene belongs to eudesmane‐type sesquiterpenes, whereas cadinene bears a cadalane‐type skeleton. The oil samples from all three tissues across different batches of samples showed a grossly similar compositional profile. The oil constituents were structurally characterized through the mass spectral library hits and their retention index (RI) values (Table 1). The identity of these three major constituents (1–3) was further confirmed through 1H and 13C NMR spectroscopy data of the leaf oil sample. The 1H NMR spectrum showed the characteristic peaks of vinylic, allylic and methyl protons of 1–3 [δ H (1): 4.69–4.74 (m, 12‐H and 14‐H), 4.44 (m, 14‐H), 2.28–2.34 (m, 3‐H), 1.75 (s, 13‐Me), 0.73 (s, 15‐Me); δ H (2): 5.32 (m, 3‐H), 4.69–4.71 (m, overlapped with 1, 12‐H), 1.75 (s, overlapped with 1, 13‐Me), 1.61 (s, 14‐Me), 0.80 (s, 15‐Me); δ H (3): 5.45 (s, 4‐H), 2.71 (ddd, J = 12.8, 4.8, 2.4 Hz, 1‐H), 2.49–2.55 (m, 5‐H), 1.67 and 1.65 (2×s, 14 and 15‐Me), 0.96 (d, J = 6.9 Hz, 12‐Me), 0.78 (d, J = 6.9 Hz, 13‐Me)] (Figure 3). The peak integration was checked for these selected peaks and they maintained the original expected ratio of a particular metabolite. The 13C NMR spectrum also displayed the characteristic peaks of 1 (δ C: 150.98, 150.81, 108.13, 105.35, 49.85, 45.82, 41.92, 41.14, 36.87, 35.95, 29.46, 26.76, 23.45, 21.01, 16.32), 2 (δ C: 150.90, 135.10, 120.90, 108.25, 46.78, 46.69, 40.22, 37.93, 32.32, 28.88, 26.80, 22.95, 21.18, 20.87, 15.60), and 3 (δ C: 134.17, 129.91, 124.67, 124.45, 45.33, 39.42, 32.32, 31.96, 26.76, 26.69, 23.57, 21.75, 21.15, 18.51, 15.65) (spectrum not shown), further confirming their identities. These results matched well with the previous literature, reporting the NMR data of these individual molecules [27, 28, 29, 30]. β‐Selinene (1, 36.3%–43.7%) and δ‐cadinene (3, 29.0%–41.6%) were found with higher abundance compared to α‐selinene (11.0%–14.4%) in all three tissues (Table 1). These three sesquiterpene hydrocarbons (1–3) collectively contributed 79.6%–98.3% of the total relative peak area. They were followed by α‐humulene (0.4%–10.2%), α‐calacorene (0.3%–1.2%), and neointermedeol (0.5%–2.9%). A few sesquiterpene hydrocarbons as well as alcohols such as β‐elemene, epi‐β‐caryophyllene, γ‐gurjunene, α‐muurolene, viridiflorol, globulol were detected with minor abundance (< 1%). Even, one sulfur‐containing sesquiterpene, namely, α‐mintsulfide was detected in very low percentage. Thus, P. pubescens oil was exclusively composed of sesquiterpenoids, sharing 97.9%–100.0% relative abundance.

FIGURE 2.

FIGURE 2

Open in a new tab

GC–MS chromatogram (R t 10.0–19.0 min) of Polygonum pubescens leaf essential oil, labelled with the major constituents and their chemical structures.

TABLE 1.

The list of constituents in Polygonum pubescens essential oil, their retention time, retention index and relative abundance (GC–MS %) in aerial tissues (RICalc: calculated retention index, RILit: retention index from the literature, nd: not detected). The ranges cover the relative percentage obtained through the analysis of three different batches of samples (n = 3).

R t (min) Identity RICalc RILit Relative % (GC–MS)
Leaves Stem Inflorescence
5.56 β‐Myrcene 997 991 nd nd 0.50–1.01
12.61 β‐Elemene 1395 1391 trace–0.44 trace–0.38 0.38–0.67
13.60 α‐Humulene 1459 1454 0.39–1.20 5.16–10.20 1.78–2.67
13.72 epi‐β‐Caryophyllene 1467 1466 0.22–0.52 0.31–0.49 0.36–0.52
13.93 γ‐Gurjunene 1481 1473 trace–0.48 trace–0.39 0.31–0.41
14.11 β‐Selinene (β‐Eudesmene) 1492 1486 40.40–43.72 37.36–43.72 36.30–38.98
14.25 α‐Selinene 1502 1494 12.98–14.42 12.85–13.20 10.99–11.79
14.31 α‐Muurolene 1506 1499 trace–0.43 trace–0.46 0.42–0.48
14.66 δ‐Cadinene 1529 1524 37.61–41.60 29.01–34.75 39.59–40.61
14.96 α‐Calacorene 1548 1542 0.29–1.23 0.43–1.19 0.39–0.40
15.58 Globulol 1589 1580 trace trace trace
15.69 Viridiflorol 1596 1591 trace–0.64 0.26–0.63 0.81–0.94
16.00 Humulane‐1,6‐dien‐3‐ol 1618 1619 trace 0.24–0.46 0.34–0.40
16.20 Muurola‐4,10(14)‐dien‐1β‐ol 1633 1635 0.26–0.83 0.60–1.76 0.56–0.60
16.58 Neointermedeol 1661 1660 0.54–1.81 1.42–2.86 2.39–2.41
17.74 α‐Mintsulfide 1745 1744 trace trace–0.38 0.52–0.76
Open in a new tab
FIGURE 3.

FIGURE 3

Open in a new tab

1H NMR spectrum (400 MHz, CDCl3, δ H 8.0–0.0 ppm) of Polygonum pubescens leaf essential oil labelled with the designated protons of major constituents 1–3. Label: (compound number) carbon number—group.

The major constituents were semi‐quantified in the oil samples using IS (β‐caryophyllene) in GC–MS and the concentration was represented in mg/g of oil (Figure 4). As indicated through relative composition, the quantity of β‐selinene and δ‐cadinene was highest in all three oil samples, ranging 221.8–254.5 and 140.6–197.6 mg/g, respectively. They were followed by α‐selinene and α‐humulene which were quantified as 74.2–86.5 and 26.8–95.8 mg/g, respectively. Neointermedeol showed a level in the range 25.9–31.6 mg/g. Three major constituents 1–3, together contributed 451.8–546.5 mg/g of the oil samples. These results revealed P. pubescens essential oil to be a rich source of sesquiterpenoids as a whole, and three eudesmane and cadalane‐type sesquiterpene hydrocarbons (1–3) in particular.

FIGURE 4.

FIGURE 4

Open in a new tab

The concentration (mg/g of oil, mean ± SD, n = 3) of major constituents in Polygonum pubescens oil from three different tissues (semi‐quantitative data obtained through GC–MS).

3.2.3. Significance of the Major Oil Constituents

Selinenes (α and β) have been demonstrated previously to play a critical role in maize defense as the precursor to its oxidized metabolites costals, costic acids promoting fungal pathogen resistance [31]. Following biotic stresses such as fungal infection or root herbivory, a bunch of selinene and analogous metabolites were produced in maize root, among which β‐selinene and β‐costic acid were predominant. Furthermore, in vitro assays showed the efficacy of β‐costic acid inhibiting maize fungal pathogens even at 25 µg/mL dose. This study revealed β‐costic acid as a potent phytoalexin in maize root, produced from β‐selinene under the biotic stress. Recently, a gram‐scale synthetic method has been reported to prepare β‐costic acid from β‐selinene [32]. Therefore, selinenes are ecologically and agrochemically important molecules. Other hand, δ‐cadinene (3) is a widely bioactive phytochemical possessing anticancer, antimicrobial, larvicidal and acaricidal activities. It has been shown to inhibit the proliferation of human ovarian cancer cells dose‐dependently (10–100 µM) through the induction of apoptosis, cell cycle arrest, nuclear membrane damage and caspase activation [33]. Bioassay‐guided fractionation of Schinus molle fruit essential oil identified δ‐cadinene as the principal active constituent against antibiotic resistant Streptococcus pneumonia with a minimum inhibitory concentration (MIC) 31.25 µg/mL [34]. This molecule also inhibited the growth of Listeria monocytogenes, a food‐borne pathogen with MIC 5.0 µL/mL through the suppression of biofilm formation and cell membrane damage. It was effective (10.0 µL/mL) in controlling the growth of this pathogen in milk [35]. Its larvicidal activity was reported against mosquito larvae with LC50 ranging 8.23–9.86 µg/mL [36]. The essential oil from dry shoots and leaves of Rhododendron nivale and its major component δ‐cadinene showed acaricidal activity (1.56–6.25 µL/mL) against Psoroptes cuniculi, a mite affecting livestock [37]. Thus, the major constituents of P. pubescens oil are important candidates from the perspective of plant defense and bioactive potency.

The plant‐based source of selinenes, especially β‐selinene is not rare in nature. A thorough literature survey revealed several plant sources with comparably high β‐selinene content (% GC–MS > 20%) such as Apium graveolens seeds (celery, 11.2%–22.2%) [38], Phlomis cancellata stem (34.7%) [39], Piper mollipilosum leaves (32.4%) [40], Eryngium kotschyi aerial parts (53.0%) [41], Litsea glauca leaves (30.8%) [42], various tissues of Callicarpa macrophylla (29.0%–57.0%) [43], Helichrysum italicum herb (21.7%) [44], Torilis japonica aerial parts (22.9%) [45], Senecio polyanthemoides leaves (32.7%) [46], Mangifera indica leaves (mango, 24.0%) [47], Peucedanum cervariifolium aerial parts (27.4%) [48], Cinnamomum chemungianum leaves (21.0%) [49], Phlomis aucheri aerial parts (27.0%) [50], Pluchea purpurascens stem and flowers (26.9%–27.6%) [51]. However, the occurrence scenario is different in case of δ‐cadinene. Though it is quite well‐distributed, there are only a handful of sources known with high δ‐cadinene content including Litsea calicaris leaves (38.2%) [52], Pterocarya stenoptera leaves (24.8%) [53], Teucrium yemense leaves (34.9%) [54], Alpinia blepharocalyx rhizome (28.4%) [55], and Cedar atlantica wood (36.3%) [56]. Again, among these sources, majority are trees which reduces the convenience of their cultivation, collection or processing. On the other hand, P. pubescens is a weedy herb and widely distributed across tropics and subtropics, making it a promising sustainable source of δ‐cadinene.

3.2.4. Phytotoxicity of P. pubescens Oil Samples

Many essential oils and their constituents are known to possess phytotoxicity which inhibits the seed germination and plant growth. Considering the weedy nature of this herb and established involvement of the major volatiles in plant defense (as discussed in the Section 3.2.3), its essential oil was evaluated for phytotoxicity against the growth of wheatgrass coleoptile. A dose‐dependent inhibition was exerted by the oil samples from all three tissues in the range 1.0–200.0 µg/mL (Figure 5). However, the observed phytotoxicity was not highly potent and only 52%–72% growth inhibition was found at 100.0 µg/mL. A plateau was reached at > 100.0 µg/mL, showing a comparable inhibition even at 200.0 µg/mL (59%–73%). In comparison, allyl alcohol, a standard chemical herbicide (positive control) achieved nearly 95% inhibition at 50.0 µg/mL in the same assay (data not shown). These results indicated that P. pubescens essential oils, rich in sesquiterpene hydrocarbons were relatively weak phytotoxin and could not exert a complete inhibition of coleoptile growth up to 200.0 µg/mL. To the best of our knowledge, there is no strong evidence available in the literature on phytotoxic potential of α‐/β‐selinene and δ‐cadinene‐rich oils or individual molecules. Though the phytotoxicity of P. pubescens oil may not be overruled, it is not a competent candidate to be considered for the herbicidal application.

FIGURE 5.

FIGURE 5

Open in a new tab

Wheatgrass coleoptile bioassay: dose‐dependent (1.0–200.0 µg/mL) growth inhibition (%, mean ± SD, n = 8 coleoptiles per assay × 2 sets) of wheatgrass coleoptiles by Polygonum pubescens oil samples [ns, not significant, p > 0.05; #, significant, 0.05 > p > 0.001; *, highly significant, p < 0.001 vs. control].

4. Conclusion

In conclusion, P. pubescens, a weedy herb of tropics and subtropics was studied morphologically and identified as a potential source of volatile oil. Among three tissues tested, the leaves of this herb produced relatively high yield (0.20%–0.35%) of essential oil. The oil was exclusively dominated by three eudesmane and cadalane‐type sesquiterpene hydrocarbons, namely, β‐selinene, α‐selinene, and δ‐cadinene, contributing 79.6%–98.3% relative abundance, collectively. Though the oil showed a dose‐dependent phytotoxicity on wheatgrass coleoptiles, the efficacy was much lower than a chemical herbicide. However, major sesquiterpene hydrocarbons in the oil bear importance from plant defense, chemical synthesis, and bioactivity points of view; indicating this herb as a sustainable reserve of these phytochemicals.

Author Contributions

Sajjad Hussain: investigation, methodology, writing – original draft. Tarani Biswas: methodology, formal analysis. Parthiv Kar: methodology. Siddhartha Proteem Saikia: writing – review and editing. Saikat Haldar: conceptualization, supervision, writing – original draft.

Conflicts of Interest

The authors declare no conflicts of interest.

Acknowledgments

The current work was supported by the CSIR‐NEIST, Jorhat In‐house Project (OLP 2503C) and the Director, CSIR‐NEIST, Jorhat (Manuscript No. CSIR‐NEIST/PUB/2025/093). S.H., T.B., and P.K. acknowledge UGC, New Delhi for the fellowship.

Data Availability Statement

The data that support the findings of this study are available from the corresponding author upon reasonable request.

References

  • 1. “Persicaria pubescens (Blume) H.Hara,” Royal Botanic Gardens, Kew | Plants of the World Online, accessed March 22, 2025 , https://powo.science.kew.org/taxon/urn:lsid:ipni.org:names:695122‐1.
  • 2. “Lax Knotweed,” Flowers of India, accessed March 22, 2025 , https://www.flowersofindia.net.
  • 3. Dash P. R., Sohrab M. H., and Rana M. S., “ Polygonum flaccidum (polygonaceae): A Review,” International Journal of Life Sciences and Review 2, no. 6 (2016): 109–113, 10.13040/IJPSR.0975-8232. [DOI] [Google Scholar]
  • 4. Mazid A., Datta B. K., and Nahar L., et al., “Analgesic and Diuretic Properties of α‐Santalone From Polygonum flaccidum ,” Phytotherapy Research 24 (2010): 1084–1087, 10.1002/ptr.3053. [DOI] [PubMed] [Google Scholar]
  • 5. Choudhary R. K., Oh S., and Lee J., “An Ethnomedicinal Inventory of Knotweeds of Indian Himalaya,” Journal of Medicinal Plants Research 5 (2010): 2095, 10.5897/JMPR.9000068. [DOI] [Google Scholar]
  • 6. Sahab Uddin M., Nasrullah M., and Hossain M. S., et al., “Evaluation of Nootropic Activity of Persicaria flaccida on Cognitive Performance, Brain Antioxidant Markers and Acetylcholinesterase Activity in Rats: Implication for the Management of Alzheimer's Disease,” American Journal of Psychiatry and Neuroscience 4 (2016): 26, 10.11648/j.ajpn.20160402.12. [DOI] [Google Scholar]
  • 7. Yu F., Li C., Dai C., Liu K., and Li Y., “Phosphate: Coupling the Functions of Fertilization and Passivation in Phytoremediation of Manganese‐Contaminated Soil by Polygonum pubescens Blume,” Chemosphere 260 (2020): 127651, 10.1016/j.chemosphere.2020.127651. [DOI] [PubMed] [Google Scholar]
  • 8. Yu F., Li Y., Li F., Li C., and Liu K., “The Effects of EDTA on Plant Growth and Manganese (Mn) Accumulation in Polygonum pubescens Blume Cultured in Unexplored Soil, Mining Soil and Tailing Soil From the Pingle Mn Mine, China,” Ecotoxicology and Environmental Safety 173 (2019): 235–242, 10.1016/j.ecoenv.2019.01.086. [DOI] [PubMed] [Google Scholar]
  • 9. Bora P. K., Saikia J., Kemprai P., Saikia S. P., Banik D., and Haldar S., “Evaluation of Postharvest Drying, Key Odorants, and Phytotoxins in Plai (Zingiber montanum) Essential Oil,” Journal of Agricultural and Food Chemistry 69 (2021): 5500–5509, 10.1021/acs.jafc.1c00256. [DOI] [PubMed] [Google Scholar]
  • 10. Borah G., Samia B. R., Hussain S., Kemprai P., Saikia S. P., and Haldar S., “Eryngial: An α,β‐Unsaturated Fatty Aldehyde as the Major Phytotoxin in Spiny Coriander (Eryngium foetidum L.) Essential Oil,” Chemistry and Biodiversity 21 (2024): e202400195, 10.1002/cbdv.202400195. [DOI] [PubMed] [Google Scholar]
  • 11. Kemprai P., Bora P. K., and Mahanta B. P., et al., “ Piper betleoides C. DC.: Edible Source of Betel‐Scented Sesquiterpene‐Rich Essential Oil,” Flavour and Fragrance Journal 35 (2020): 70–78, 10.1002/ffj.3537. [DOI] [Google Scholar]
  • 12. Mahanta B. P., Sut D., Lal M., and Haldar S., “Hydrodistillation Alters the Compositional Originality in Black Turmeric (Curcuma caesia Roxb.) Essential Oil,” Journal of Essential Oil Research 33 (2021): 240–246, 10.1080/10412905.2021.1873868. [DOI] [Google Scholar]
  • 13. Seimandi G., Álvarez N., and Stegmayer M. I., et al., “An Update on Phytochemicals and Pharmacological Activities of the Genus Persicaria and Polygonum ,” Molecules 26 (2021): 5956, 10.3390/molecules26195956. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 14. Baharum S. N., Bunawan H., Ghani M. A., Mustapha W. A. W., and Noor N. M., “Analysis of the Chemical Composition of the Essential Oil of Polygonum minus Huds. Using Two‐Dimensional Gas Chromatography‐Time‐of‐Flight Mass Spectrometry (GC‐TOF MS),” Molecules 15 (2010): 7006–7015, 10.3390/molecules15107006. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 15. Cai J., Wang S., Gao Y., and Wang Q., “Antibacterial Activity and Mechanism of Polygonum orientale L. Essential Oil Against Pectobacterium carotovorum subsp. carotovorum ,” Foods 11 (2022): 1585, 10.3390/foods11111585. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 16. Chen F., Jia J., Zhang Q., Yang L., and Gu H., “Isolation of Essential Oil From the Leaves of Polygonum viscosum Buch‐ham. Using Microwave‐Assisted Enzyme Pretreatment Followed by Microwave Hydrodistillation Concatenated With Liquid–Liquid Extraction,” Industrial Crops and Products 112 (2018): 327–341, 10.1016/j.indcrop.2017.12.024. [DOI] [Google Scholar]
  • 17. Demirpolat A., “Chemical Composition of Essential Oils of Seven Polygonum Species From Turkey: A Chemotaxonomic Approach,” Molecules 27 (2022): 9053, 10.3390/molecules27249053. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 18. Tong X., Li X., and Ayaz M., et al., “Neuroprotective Studies on Polygonum hydropiper L. Essential Oils Using Transgenic Animal Models,” Frontiers in Pharmacology 11 (2021): 580069, 10.3389/fphar.2020.580069. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 19. Ahmed M., Datta B. K., Rouf A. S. S., and Jakupovic J., “Flavone and α‐Santalene Derivatives From Polygonum flaccidum ,” Phytochemistry 30 (1991): 3155–3156, 10.1016/S0031-9422(00)98279-7. [DOI] [Google Scholar]
  • 20. Dash P. R., Rana M. S., Hasan C. M., and Muhit M. A., “Friedelin Derivatives from Polygonum flaccidum Showed Cytotoxic Activity on Vero Cell Line,” Tropical Journal of Natural Product Research 5 (2021): 1456, 10.26538/tjnpr/v5i8.22. [DOI] [Google Scholar]
  • 21. Liu Q., Li B., and Zhao J., et al., “A New Sucrosephenylpropanoid Ester From Polygonum pubescens Blume,” Natural Product Research 31 (2017): 1725–1732, 10.1080/14786419.2017.1289208. [DOI] [PubMed] [Google Scholar]
  • 22. Bryson C. T. and Carter R., “Biology of Pathways for Invasive Weeds,” Weed Technology 18 (2004): 1216–1220, 10.1614/0890-037X(2004)018[1216:BOPFIW]2.0.CO;2. [DOI] [Google Scholar]
  • 23. Cecotti R., Carpana E., Falchero L., Paoletti R., and Tava A., “Determination of the Volatile Fraction of Polygonum bistorta L. at Different Growing Stages and Evaluation of Its Antimicrobial Activity Against Two Major Honeybee (Apis mellifera) Pathogens,” Chemistry and Biodiversity 9 (2012): 359–369, 10.1002/cbdv.201100326. [DOI] [PubMed] [Google Scholar]
  • 24. Dũng N. X., Van Hac L., and Leclercq P. A., “Volatile Constituents of the Aerial Parts of Vietnamese Polygonum odoratum L.,” Journal of Essential Oil Research 7 (1995): 339, 10.1080/10412905.1995.9698534. [DOI] [Google Scholar]
  • 25. Miyazawa M. and Tamura N., “Components of the Essential Oil From Sprouts of Polygonum hydropiper L. (‘Benitade’),” Flavour and Fragrance Journal 22 (2007): 188–190, 10.1002/ffj.1779. [DOI] [Google Scholar]
  • 26. Yaacob K. B., “Essential Oil of Polygonum minus Huds,” Journal of Essential Oil Research 2 (1990): 167–172, 10.1080/10412905.1990.9697855. [DOI] [Google Scholar]
  • 27. Chen Y., Xiong Z., Zhou G., Liu L., and Li Y., “Total Synthesis and Stereochemistry of 13‐hydroxy‐α‐eudesmol,” Tetrahedron: Asymmetry 9 (1998): 1923, 10.1016/S0957-4166(98)00184-0. [DOI] [Google Scholar]
  • 28. Chou T.‐S., Lee S.‐J., and Yao N.‐K., “Total Synthesis of Racemic Selina‐3,7(11)‐Diene, α‐Selinene, and α‐Eudesmol,” Tetrahedron 45 (1989): 4113–4124, 10.1016/S0040-4020(01)81307-7. [DOI] [Google Scholar]
  • 29. Chu S. S., Jiang G. H., and Liu Z. L., “Insecticidal Compounds From the Essential Oil of Chinese Medicinal Herb Atractylodes chinensis ,” Pest Management Science 67 (2011): 1253–1257, 10.1002/ps.2180. [DOI] [PubMed] [Google Scholar]
  • 30. Hu Y., Chou W. K. W., Hopson R., and Cane D. E., “Genome Mining in Streptomyces clavuligerus: Expression and Biochemical Characterization of Two New Cryptic Sesquiterpene Synthases,” Chemistry & Biology 18 (2011): 32–37, 10.1016/j.chembiol.2010.11.008. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 31. Ding Y., Huffaker A., and Köllner T. G., et al., “Selinene Volatiles Are Essential Precursors for Maize Defense Promoting Fungal Pathogen Resistance,” Plant Physiology 175 (2017): 1455–1468, 10.1104/pp.17.00879. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 32. Matsushima Y. and Iwata K., “Gram‐Scale Approach for β‐Costic Acid via Allylic Oxidation of β‐Selinene,” Bioscience, Biotechnology, and Biochemistry 88 (2024): 742–746, 10.1093/bbb/zbae052. [DOI] [PubMed] [Google Scholar]
  • 33. Hui L.‐M., Zhao G.‐D., and Zhao J.‐J., “δ‐Cadinene Inhibits the Growth of Ovarian Cancer Cells via Caspase‐Dependent Apoptosis and Cell Cycle Arrest,” International Journal of Clinical and Experimental Pathology 8 (2015): 6046. [PMC free article] [PubMed] [Google Scholar]
  • 34. Pérez‐López A., Cirio A. T., Rivas‐Galindo V. M., Aranda R. S., and de Torres N. W., “Activity Against Streptococcus pneumoniae of the Essential Oil and δ‐Cadinene Isolated from Schinus molle Fruit,” Journal of Essential Oil Research 23 (2011): 25, 10.1080/10412905.2011.9700477. [DOI] [Google Scholar]
  • 35. Qin R., Yang S., and Fu B., et al., “Antibacterial Activity and Mechanism of the Sesquiterpene δ‐Cadinene Against Listeria monocytogenes ,” LWT 203 (2024): 116388, 10.1016/j.lwt.2024.116388. [DOI] [Google Scholar]
  • 36. Govindarajan M., Rajeswary M., and Benelli G., “δ‐Cadinene, Calarene and δ‐4‐Carene From Kadsura Heteroclita Essential Oil as Novel Larvicides against Malaria, Dengue and Filariasis Mosquitoes,” Combinatorial Chemistry & High Throughput Screening 19 (2016): 565–571, 10.2174/1386207319666160506123520. [DOI] [PubMed] [Google Scholar]
  • 37. Guo X., Shang X., Li B., Zhou X. Z., Wen H., and Zhang J., “Acaricidal Activities of the Essential Oil From Rhododendron nivale Hook. F. and Its Main Compund, δ‐Cadinene Against Psoroptes cuniculi ,” Veterinary Parasitology 236 (2017): 51–54, 10.1016/j.vetpar.2017.01.028. [DOI] [PubMed] [Google Scholar]
  • 38. Philippe J., Suvarnalatha G., Sankar R., and Suresh S., “Kessane in the Indian Celery Seed Oils,” Journal of Essential Oil Research 14 (2002): 276–277, 10.1080/10412905.2002.9699852. [DOI] [Google Scholar]
  • 39. Akhlaghi H. and Rustaiyan A., “Composition of the Essential Oils from the Leaves, Stems and Ether‐Extracted Aerial Parts of Phlomis cancellata Bunge from Northeast of Iran,” Journal of Essential Oil Research 14 (2011): 278, 10.1080/0972060X.2011.10643934. [DOI] [Google Scholar]
  • 40. Araujo C. A., da Camara C. A. G., de Moraes M. M., de Vasconcelos G. J. N., Pereira M. R., and Zartman C. E., “Chemical Composition of Essential Oils From Four Piper Species, Differentiation Using Multivariate Analysis and Antioxidant Activity,” Natural Product Research 36 (2021): 436–439, 10.1080/14786419.2020.1774761. [DOI] [PubMed] [Google Scholar]
  • 41. Pasayeva L., Karadeli H., Ulukus D., Fatullayev H., and Tugay O., “Chemical Composition and Biological Activity of the Essential Oil from Eryngium kotschyi Boiss.,” Journal of Essential Oil Research 26 (2023): 1055, 10.1080/0972060X.2023.2265959. [DOI] [Google Scholar]
  • 42. Salleh W. M. N. H. W., Salihu A. S., and Ab Ghani N., “Essential Oils Composition of Litsea glauca and Litsea fulva and Their Anticholinesterase Inhibitory Activity,” Natural Product Research 38 (2024): 629–633, 10.1080/14786419.2023.2180507. [DOI] [PubMed] [Google Scholar]
  • 43. Chandra M., Prakash O., and Kumar R., et al., “β‐Selinene‐Rich Essential Oils From the Parts of Callicarpa macrophylla and Their Antioxidant and Pharmacological Activities,” Medicines 4 (2017): 52, 10.3390/medicines4030052. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 44. Oliva A., Garzoli S., and Sabatino M., et al., “Chemical Composition and Antimicrobial Activity of Essential Oil of Helichrysum italicum (Roth) G. Don Fil. (Asteraceae) From Montenegro,” Natural Product Research 34 (2020): 445–448, 10.1080/14786419.2018.1538218. [DOI] [PubMed] [Google Scholar]
  • 45. Chen J., Xu X.‐J., Fang Y.‐H., Li S., and Zhang Y.‐L., “Chemical Composition and Antibacterial Activity of the essential oil from the aerial parts of Torilis japonica ,” Journal of Essential Oil Research 16 (2013): 499, 10.1080/0972060X.2013.831575. [DOI] [Google Scholar]
  • 46. Oladipupo L. A. and Adebola O. O., “Chemical Composition of the Essential Oils of the Flowers, Leaves and Stems of Two Senecio polyanthemoides Sch. Bip. Samples From South Africa,” Molecules 14 (2009): 2077–2086, 10.3390/molecules14062077. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 47. Simionatto E., Peres M., and Hess S. C., et al., “Chemical Composition and Cytotoxic Activity of Leaves Essential Oil From Mangifera indica var. coquinho (Anacardiaceae),” Journal of Essential Oil Research 22 (2010): 596–599, 10.1080/10412905.2010.9700408. [DOI] [Google Scholar]
  • 48. Bazgir A., Shaabani A., and Sefidkon F., “Composition of the Essential Oil of Peucedanum cervariifolium C.A. Mey. From Iran,” Journal of Essential Oil Research 17 (2005): 380–381, 10.1080/10412905.2005.9698936. [DOI] [Google Scholar]
  • 49. Rameshkumar K. B., George V., and Shiburaj S., “Chemical Constituents and Antibacterial Activity of the Leaf Oil of Cinnamomum chemungianum Mohan et Henry,” Journal of Essential Oil Research 19 (2007): 98–100, 10.1080/10412905.2007.9699238. [DOI] [Google Scholar]
  • 50. Javidnia K., Miri R., Soltani M., and Khosravi A. R., “Essential Oil Composition of Two Species of Phlomis L. (Phlomis aucheri Boiss. and Phlomis elliptica Benth.) (Lamiaceae) From Iran,” Journal of Essential Oil Research 22 (2010): 314–317, 10.1080/10412905.2010.9700334. [DOI] [Google Scholar]
  • 51. Pino J. A., Perera W. H., Sarduy R., Oviedo R., and Quijano C. E., “Essential Oil From the Stems, Leaves and Flowers of Pluchea rosea Godfrey and Pluchea purpurascens (Sw.) DC,” Journal of Essential Oil Research 20 (2008): 497–501, 10.1080/10412905.2008.9700069. [DOI] [Google Scholar]
  • 52. Scora R. W. and Scora P. E., “Essential Leaf Oil of Persea Subgenus eriodaphne and Closely Related Perseoid Genera,” Journal of Essential Oil Research 13 (2001): 37–42, 10.1080/10412905.2001.9699599. [DOI] [Google Scholar]
  • 53. Yin C., Sun F., Rao Q., and Zhang Y., “Chemical Compositions and Antimicrobial Activities of the Essential Oil From Pterocarya stenoptera C. DC,” Natural Product Research 34 (2020): 2828–2831, 10.1080/14786419.2019.1587426. [DOI] [PubMed] [Google Scholar]
  • 54. Ali N. A. A., Wurster M., Arnold N., Lindequist U., and Wessjohan L., “Chemical Composition of the Essential Oil of Teucrium yemense Deflers,” Records of Natural Products 2 (2008): 25. [Google Scholar]
  • 55. Hung N. D., Huong L. T., Dai D. N., Hoi T. M., and Ogunwande I. A., “Chemical Composition of Essential Oils of Alpinia strobiliformis TL Wu & SJ Chen and Alpinia blepharocalyx K. Schum. from Vietnam,” Journal of Essential Oil Research 21 (2018): 1585, 10.1080/0972060X.2019.1567396. [DOI] [Google Scholar]
  • 56. Kačániová M., Galovičová L., and Valková V., et al., “Chemical Composition, Antioxidant, In Vitro and In Situ Antimicrobial, Antibiofilm, and Anti‐Insect Activity of Cedar atlantica Essential Oil,” Plants 11 (2022): 358, 10.3390/plants11030358. [DOI] [PMC free article] [PubMed] [Google Scholar]

Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Data Availability Statement

The data that support the findings of this study are available from the corresponding author upon reasonable request.

Articles from Chemistry & Biodiversity are provided here courtesy of Wiley

RESOURCES