Non-strigolactone natural products interfering with parasitic weed development

Contents

• 1 Introduction

• 2 Terpenes and terpenoids
  • 2.1 Sesquiterpene lactones (1–38, Fig. 2–5)
  • 2.2 Iridoids (39–43; Fig. 6)
  • 2.3 Apocarotenoids, sesquiterpenoids, and diterpenoids (44–55; Fig. 7)
  • 2.4 Higher terpenoids (56–65; Fig. 8)

• 3 Aromatic metabolites
  • 3.1 Monocyclic aromatic metabolites (66–91; Fig. 9)
  • 3.2 Fused bicyclic aromatic metabolites (92–99; Fig. 10)
  • 3.3 Flavonoids and isoflavonoids (100–114; Fig. 11)
  • 3.4 Other aromatic metabolites (115–119; Fig. 12)

• 4 N-containing metabolites
  • 4.1 Amino acids and peptide-derived metabolites (120–143; Fig. 13)
  • 4.2 Alkaloids and N-heterocyclic metabolites (144–157; Fig. 14)
  • 4.3 Nitrile- and isothiocyanate-containing metabolites (162–173; Fig. 15)
  • 4.4 Other N-containing metabolites (174–177; Fig. 16)

• 5 Miscellaneous bioactive metabolites
  • 5.1 Volatile and signalling molecules (178–180; Fig. 17)
  • 5.2 Butenolides and related polyacetylenes (181–189; Fig. 18)
  • 5.3 Quinones and other oxygenated cyclic ketones (190–196; Fig. 19)
  • 5.4 Carbohydrates and polyols (197–199; Fig. 20)

• 6 Lipophilicity trends of the bioactive metabolites

• 7 Conclusions and prospects

• 8 Conflicts of interest

• 9 Data availability

• Acknowledgements

• References

---

Abstract

Covering: up to 2026

Strigolactones dominate the research on parasitic plant germination. Nevertheless, other types of natural products have roles in stimulating or inhibiting the germination and subsequent growth of parasitic weeds, including synergistic interactions with other compounds released by host roots that mediate host recognition and chemotropism. This review focuses on the bioactive non-strigolactone compounds, classified as terpenes and terpenoids, aromatic metabolites, N-containing metabolites and miscellaneous structures. Terpenes and terpenoids are the most common germination stimulants (51.0% of the total, among which sesquiterpene lactones represent the most common structures), while aromatic (41.7%) and N-containing (36.5%) metabolites are the leading inhibitors of germination or seedling development. The clog P trends suggest that molecular lipophilicity alone does not allow distinguishing between stimulants or inhibitors, but the data showed that two main intervals are particularly enriched in bioactive structures: moderately lipophilic compounds (47%, with clog P values between 1.0 and 3.0) and hydrophilic molecules (24%, with clog P below 0). An evaluation of germination-induced specificity indicated that sesquiterpene lactones are the strongest elicitors for O. cumana, P. aegyptiaca and S. asiatica, karrikin₁ and different terpenoids for O. minor and S. hermonthica, and isothiocyanates for P. ramosa. Notably, several sesquiterpene lactones and isothiocyanates induced germination at concentrations comparable to those of natural strigolactones. Beyond germination, recent findings implicate non-strigolactone cues in haustorium initiation and host chemotropism, underscoring the complexity of chemical signaling in parasitic plant development. Overall, the evidence gathered herein shows that parasitic development is influenced by a wider chemical space than strigolactones alone, opening perspectives for eco-rational bioherbicide development and for understanding host–parasite communication.

---

Jesús G. Zorrilla

Jesús García Zorrilla is Postdoctoral Researcher at the University of Cádiz (Spain) and the Institute of Biomolecules (INBIO), following extended international research stays. He completed his PhD on the synthesis of novel strigolactone analogues and eudesmanolide-based mimics for the sustainable management of parasitic weeds. His subsequent research has focused on the design of improved analogues and mimics, bioprospecting of non-strigolactone natural products that modulate parasitic weed germination and development, and, more broadly, applications in crop protection and health-related fields. His contributions have been recognized with awards from the Phytochemical Society of Europe and the Royal Spanish Society of Chemistry.

Antonio Cala Peralta

Antonio Cala Peralta received his PhD in Science from the University of Cádiz, where his doctoral research focused on the development, encapsulation, and biological evaluation of compounds for parasitic weed control. His PhD work was recognised with awards for the Best Doctoral Thesis from the University of Cádiz and the Royal Spanish Society of Chemistry. He subsequently held postdoctoral positions at the University of Cádiz and the University of Angers and established collaborations with research groups in Italy and Austria. Since 2023, he has been an Assistant Professor of Organic Chemistry at the University of Cádiz. His research focuses on bioactive natural products and related compounds for sustainable crop protection and parasitic weed management.

Marco Masi

Marco Masi received his PhD from the University of Naples Federico II (UNINA), performing a portion of his doctoral research at the New Mexico Institute of Mining and Technology (Socorro, NM, USA). He subsequently held one-year Postdoctoral positions at Brigham Young University (Provo, UT, USA) and UNINA. Following this, he spent three years as a researcher at the Biotechnology and Biological Control Agency (BBCA) in Rome and five years as a fixed-term researcher at UNINA. Since 2024, he has served as an Associate Professor of Organic Chemistry at UNINA. His research focuses on the isolation and chemical characterization of bioactive natural products derived from plants and microorganisms.

Francisco A. Macías

Professor Francisco A. Macías has been a Professor of Organic Chemistry at the University of Cádiz since 2000. Internationally recognised for his contributions to allelopathy and natural product chemistry, he has received major awards from the Phytochemical Society of Europe and the International Allelopathy Society and was listed among Stanford University's Top 2% scientists. He leads the Cádiz Allelopathy Group, a pioneering European team that has studied over 3500 allelochemicals and derivatives with applications in agriculture and medicine.

Alessio Cimmino

Along with being an Associate Professor of Organic Chemistry at the University of Naples Federico II, Dr Cimmino holds the National Scientific Qualification as a Full Professor. With over 20 years of expertise, his research focuses on the isolation and stereostructural characterization of bioactive metabolites produced by microorganisms and plants. He obtained his PhD in 2009, specializing in natural compounds for weed management. His career is marked by extensive international experience, with several years spent at prestigious institutions in Germany (Research Center Borstel) and Spain (CSIC). This international background underpins his research in advanced spectroscopy and hemi-synthesis, targeting structure–activity relationships for applications ranging from sustainable agriculture to medicinal chemistry.

---

1 Introduction

Parasitic plants are among the most destructive and difficult to control weedy angiosperms affecting crops.¹ The species most frequently responsible for global crop damage include the root-parasite including broomrapes (Orobanche spp. and Phelipanche spp.), witchweeds (Striga spp.), Rhamphicarpa spp. and Alectra spp. Other species, such as dodders (Cuscuta spp), parasitise the stems of host plants.

Parasitic plants depend on their hosts to varying degrees, functioning either as obligate root- or stem-holoparasites, which include broomrapes and many species of dodder, or as hemiparasites, which may be facultative (e.g., Rhamphicarpa spp.) or obligate (e.g., witchweeds and Alectra spp). All parasitic species use a specialised organ known as the haustorium to feed from either the root or stem of the host, extracting water and nutrients. In root holoparasites and obligate root hemiparasites, parasitism must occur shortly after germination; therefore, their seeds germinate only in response to chemical cues exuded by host plants into the soil. In contrast, facultative root hemiparasites and stem parasites are not subject to this requirement for germination.

Host-induced germination occurs in a species-specific manner along two dimensions: first, the qualitative and quantitative composition of exuded molecules is specific for each plant species, and second, the germination of each parasitic species is induced by specific compounds and their combinations. This coordination represents the first level of host range specialization, which is subsequently followed by compatible physical, chemical and hormonal interactions during haustorium initiation and vascular connections in susceptible hosts.²

The most common germination stimulants are the phytohormones strigolactones, which are derived from the carotenoid pathway via carlactone, consisting of three 6-, 5-, and 5-membered fused rings (ABC) linked to a butenolide (D) ring through an enol ether bridge (Fig. 1). Natural canonical strigolactones are divided into two groups, strigol and orobanchol types, depending on the orientations of the B/C junction (3aR,8aS and 3aS,8aR, respectively), while the D-ring is always R-configured.³ These compounds are phytohormones that control shoot branching, act as signals for hyphal branching in arbuscular mycorrhizal fungi, and influence parasitic weed development.⁴

Fig. 1 General structure of the canonical strigolactones.

Although the role of strigolactones as germination elicitors has been deeply studied,⁴ additional roles played these phytohormones have recently been discovered in facultative parasites.^5,6 Authors have described how strigolactones can act as chemoattractants guiding Orobanchaceae root development in a tropism process that involves the auxin efflux transport PIN2 in Phtheirospermum japonicum. However, they only stimulated germination in the absence of nitrate ions.⁶ Another facultative parasite, Castilleja foliolosa, exhibited parasitic behavior post-germination after treatment with strigolactones, triggering the formation of haustorium-like root structures.⁵

Parasitic weeds are widespread around the globe, and they cause a huge negative impact in the production of crops.⁷ For example, in Africa, 33 out of the 40 Striga species reported worldwide have been found infecting 11 major crops and, in sub-Saharan Africa alone, the crop losses were estimated at approximately US $1 billion, affecting 300 million people.^8,9

Management of parasitic weeds is not simple, since conventional methods do not work or have a limited impact, especially in low-input cropping systems. The most common strategies rely on crop resistance and chemical control. However, the use of host resistance is constrained by the scarcity of resistance sources in most crops affected by parasitic weeds and the heterogeneity of the parasitic seed bank, which in some cases, enables the parasitic weed population to overcome the available resistance.¹ Commercially, chemical control mainly involves the application of herbicides targeting amino acid biosynthesis. However, due to the ability of parasitic weeds to develop resistance to these herbicides, there is a continuous need to identify new herbicidal modes of action that inhibit parasitic weed development.¹

An alternative chemical approach to controlling these weeds is the use of germination agents, which induce parasitic seed germination in the absence of a host, leading to seedling death at an early developmental stage. This strategy is more sustainable in terms of management of emergence of parasitic populations with herbicide resistance since obligated parasitic weeds are unable to adapt to host absence and die once germinated without a host. A particularly important advantage of this strategy is also its preventive nature, allowing the weed to be managed before crops are planted in the field. Most studied germination inducers belong to the strigolactone class. However, less attention has been paid to non-strigolactone natural products that interfere with parasitic weed development either by affecting germination or by acting on later stages of the parasitic life cycle.

A literature search using the keywords of Orobanche OR Phelipanche OR Striga OR Cuscuta OR parasitic plant OR parasitic weed AND germination OR radicle OR haustorium AND stimulant OR inhibitor shows that approximately 42% of published articles since 1949 are dedicated to the strigolactone class of compounds. Therefore, this review aims to expand the current understanding by analyzing and summarizing the data reported in the literature regarding the compounds other than strigolactones that interfere with parasitic weed development and their associated bioactivities on these weeds. Due to the wide diversity of chemical structures under review, the compounds are classified in different sections and subsections according to structural and biosynthetic features. Given the substantial variability among studies in bioassay conditions and seed responsiveness, this review adopted an inclusive definition of activity, considering as bioactive any metabolite reported to produce a significant effect relative to the corresponding negative control.

2 Terpenes and terpenoids

Terpenes and terpenoids, also known as isoprenoids, comprise a large and structurally diverse family of natural products derived from isoprene units, including both hydrocarbon terpenes and oxygenated or rearranged derivatives. As secondary metabolites, they commonly function as defensive, signalling, and allelopathic agents that mediate ecological interactions and adaptation to environmental stress, which, together with their abundance and bioactivity, has made them a major focus of phytochemical research.¹⁰ In the field of parasitic weeds, sesquiterpene lactones are the most extensively studied subgroup due to the number and diversity of structures reported to stimulate broomrape and witchweed germination. As reviewed in this section, additional bioactive terpenoid subclasses of increasing structural complexity derived from both plant and fungal sources have also been identified, further highlighting the chemical diversity of allelochemicals and signalling molecules that modulate the biology of parasitic weeds.

2.1 Sesquiterpene lactones (1–38, Fig. 2–5)

Terpenoids are biosynthetically derived from three isoprene units and characterized by a lactone ring fused with a sesquiterpene scaffold, most often an α,β-unsaturated γ-lactone, which is an electrophilic moiety strongly related to their biological activity. This moiety makes them relevant not only for their bioactivity in parasitic weed germination, but also for the design of bioactive canonical strigolactone analogues like eudesmanestrigolactones and guaianestrigolactones.^11,12

Sesquiterpene lactones are typically bicyclic or tricyclic structures (although seco, elemane and other atypical skeletal variants also occur),^13,14 and their subclassification is based on their non-lactone sesquiterpene ring framework, among which the most common are germacranolides (10-membered ring), eudesmanolides (fused 6–6 rings), and guaianolides (fused 5–7 rings). Seco-lactones, in contrast, are characterized by cleavage of one of the rings, leading to open-chain or rearranged structures, whereas elemanolides are defined by an elemane-type rearranged sesquiterpene skeleton.

Sesquiterpene lactones are mainly found in plants from the Asteraceae family, playing ecological roles and contributing to plant allelopathy. Their relevance in parasitic weed management increased after the discovery that the germacranolide costunolide (1) and the guaianolide dehydrocostuslactone (14), exuded by the roots of sunflower (Helianthus annuus), worked as germination stimulants for O. cumana,¹⁵ a highly specific parasitic weed species to sunflower. Moreover, compound 1 has been recently reported to influence germ tube orientation in sunflower broomrape O. cumana suggesting its additional role as a chemotropic signal.¹⁶ Costunolide (1) and dehydrocostuslactone (14) were also found in other plants such as Dolomiaea costus (formerly known as Saussurea costus or Saussurea lappa),¹³ where they occur in the roots and are isolated in large quantities, or in different species from the Magnolia or Laurus genera, like L. azorica, L. nobilis or L. novocanariensis.^17–22

A total of 38 bioactive natural sesquiterpene lactones (Fig. 2–5), including germacranolides, guaianolides, eudesmanolides, seco-lactones, and elemanolides, are reported in the literature. The available data indicate that an exocyclic methylene group in the α,β-unsaturated lactone ring is not strictly required for activity against parasitic weeds, as some reduced analogues such as dihydrocostunolide (5), dihydroparthenolide (7), dihydroperuvin (22), and dihydrosantamarine (28) are active in some species.²³ However, comparisons between closely related compounds suggest that this feature can notably modulate potency, as illustrated by the weaker effect of inuloxin B (13) relative to inuloxin A (12). Likewise, the retention of bioactivity in seco-lactones, elemanolides, and even the correlated sesquiterpenoid α-costic acid (45) indicates that neither strict ring closure nor the canonical sesquiterpene lactone framework is indispensable for germination-modulating effects.

Fig. 2 Bioactive germacranolide-type sesquiterpene lactones.

Fig. 3 Bioactive guaianolide- and pseudoguaianolide-type sesquiterpene lactones.

Fig. 4 Bioactive eudesmanolide-type sesquiterpene lactones.

Fig. 5 Bioactive seco-lactones and elemanolides.

2.1.1 Germacranolides (1–13; Fig. 2). After discussing above the key case of costunolide (1) in O. cumana germination, several other germacranolides warrant consideration.

Melampomagnolide A (2) and parthenolide (6), isolated from Magnolia grandiflora, have also shown stimulating activity in O. cumana germination, with compound 2 (reaching 25% seed germination at 10 µM) notably less active than compound 6 (75% at 1 µM).^24–27

Different germacranolides have also shown stimulatory activity in Striga asiatica germination, i.e. costunolide (1), dihydrocostunolide (5, also mainly reported from S. lappa), parthenolide (6), dihydroparthenolide (7, isolated from Ambrosia artemisiifolia, but also present in other species such as Magnolia x alba), 1,10-epoxyparthenolide (8, mainly found in M. grandiflora and Ambrosia confertiflora) and its hydrogenated analog 1,10-epoxydihydroparthenolide (9, a rare sesquiterpene lactone reported only from Achillea micrantha aerial parts), 4,5-epoxy-8-epi-inunolide (10), and 11,13-dehydroeriolin (11, reported for the first time in Schkuhria schkuhrioides and later found in other species like Stevia ovata).^28–36

In some cases, a natural product can either induce germination or inhibit growth depending on the concentration and the species, as in the case of inuloxins isolated from Dittrichia viscosa. Among this structurally diverse group of sesquiterpene lactones, inuloxin A (12) stimulated the germination of O. cumana at 100 µM (close to 60% germination) and weakly stimulated that of P. ramosa (<10% germination).³⁷ In inhibition bioassays, compound 12 completely inhibited the germination of O. crenata and C. campestris. In the same study, inuloxin B (13) exhibited weaker inhibitory effects against C. campestris, which can be attributed to the absence of the exocyclic double bond in the lactone ring.³⁸ Inuloxin A (12) was complexed with β-cyclodextrins to increase its solubility in water and its bioavailability, and the complexes almost completely inhibited P. ramosa seed germination.³⁹ Formulations of the organic extract of D. viscosa, containing compound 12, in poly(butylene succinate)- and polycaprolactone-based films (PBS and PCL, respectively) were also proposed as an alternative herbicide for the management of this weed.⁴⁰

Salonitenolide (3) and cnicin (4), with germacranolide structures similar to costunolide (1), containing an extra hydroxyl group, and, in the case of cnicin (4), bearing an additional side chain, showed high inhibitory activity in the radicle growth of O. crenata, O. cumana O. minor and P. ramosa, reaching 80% inhibition at 0.5 mM in the first three species.⁴¹ These germacranolides were isolated from the aerial parts of Centaurea cineraria L. subsp. cineraria, with cnicin (4) being the major isolated metabolite. Cnicin (4) was first isolated as the bitter component from Cnicus benedictus and described by F. Scribe in 1826,⁴² but its correct structure was determined later, while salonitenolide (3) was subsequently isolated from Centaurea salonitana and chemically characterized by the same authors.^43,44

2.1.2 Guaianolides (14–25; Fig. 3). Apart from O. cumana, dehydrocostuslactone (14) also stimulated about 70% germination of P. ramosa at 0.1 µM, a concentration that also stimulated the germination of P. aegyptiaca, though to a lesser degree than in O. cumana (dropping from approximately 50% to 10% seed germination).^15,45,46 The 3-hydroxy derivative of dehydrocostuslactone, known as isozaluzanin C (15) and produced by S. costus (but also found in L. nobilis), stimulated the germination of O. cumana seeds, while lappalone (16) and pertyolide C (17) stimulated that of P. ramosa.^26,47–50

Guaianolide (and pseudoguaianolide) stimulants for S. asiatica germination include parthenin (18, isolated from Parthenium hysterophorus), desacetylconfertiflorin (19) and confertiflorin (20, from A. confertiflora), peruvin (21), dihydroperuvin (22, reported only from the inflorescences of A. artemisiifolia), burrodin (23, unique to the aerial parts of Inula hupehensis), dihydroburrodin (24, described exclusively in Stevia isomeca), and to a lesser extent, rudmollin (25, thus far restricted to the genus Rudbeckia, including R. mollis and R. subtomentosa).^{31,32,51–54}

2.1.3 Eudesmanolides (26–33; Fig. 4). Most of the bioactive eudesmanolides are members of the 12,6-subfamily, such as reynosin (26), which stimulated the germination of P. ramosa seeds by more than 60% at 0.1 µM and O. cumana seeds by more than 25% at 1 µM.^25,55 The double-bond isomers of reynosin (26), i.e. santamarine (27) and magnolialide (29), also stimulated P. ramosa and O. crenata germination, but to a lesser extent, suggesting that the position of the double bond within the eudesmane scaffold is also related to the bioactivity.⁵⁵ Studies using mixtures of eudesmanolides 26 and 27 reported more than 50% germination of S. asiatica up to 1 nM.³¹ The occurrence of eudesmanolides 26–29 has been documented in species such as S. costus, A. confertiflora, Artemisia scoparia, L. azorica, Inula racemosa, Michelia compressa, and M. grandiflora.^{18,29,56–60}

Dihydrosantamarine (28), found in species like Michelia alba or Leontodon hispidus subsp. hispidus,^55,61,62 lacking the double bond in the lactone ring, stimulated P. ramosa but not O. cumana and O. crenata. On the other hand, anhydrojudaicin (30), found only in Artemisia canariensis, with an additional unsaturated carbonyl group, exhibited an increase in germinatory activity compared with 28 in O. cumana, P. ramosa and O. crenata.^55,63 Inuloxin C (31), isolated from D. viscosa, completely inhibited the germination of C. campestris and O. crenata at 1.6 mM, while at 100 µM, it slightly stimulated O. cumana seed germination (approximately 10%) and reduced radicle growth in O. crenata and O. minor by approximately 80%, 70% in O. cumana, and 35% in P. ramosa.^37,38 Isoalantolactone (32) was found to stimulate the germination of P. ramosa seeds by approximately 60% at 100 µM.⁴⁸

Formulations with cyclodextrins significantly increased the bioactivity of (−)-α-santonin (33) in the germination of O. cumana seeds. At 100 µM, bioactivity increased from about 20% for the free lactone dissolved in 1% acetone (v/v) to approximately 80% for the product encapsulated in β- or γ-cyclodextrin dissolved in water.⁶⁴

2.1.4 Seco-lactones and elemanolides (34–38; Fig. 5). Other structurally diverse bioactive sesquiterpene lactones, including bicyclic seco-lactones and elemanolides, such as tomentosin (34) and 8-epixanthatin (35) (isolated from the host species H. annuus), have also been identified as germination stimulants of O. cumana.¹⁵ Inuloxin D (36) and E (37), isolated from D. viscosa, stimulated O. cumana germination, while compound 36 completely inhibited O. crenata and C. campestris seed germination at 1.6 mM.^38,65 In the case of elemanolides, it was discovered that isocnicin (38, isolated from C. cineraria L. subsp. cineraria) inhibited the radicle growth of P. ramosa, O. minor, O. cumana and O. crenata.⁴¹

2.2 Iridoids (39–43; Fig. 6)

Iridoids are monoterpenoids characterized by a cyclopentane ring fused to a six-membered oxygen-containing heterocycle, commonly occurring as glycosides in plants. Some iridoid glycosides (39–43, Fig. 6) isolated from Bellardia trixago, a facultative hemiparasitic plant (family Orobanchaceae), including bartsioside (39), melampyroside (40), and mussaenoside (42), inhibited O. cumana radicle growth by 60–80% at 100 µg mL⁻¹, while aucubin (41) and gardoside methyl ester (43) showed notably lower activity (below 20%).⁶⁶ Ecotoxicological studies revealed that compound 41 exhibited little or no toxicity toward the investigated organisms and could be considered as a potential antiparasitic weed agent with an optimal toxicity-selectivity ratio.⁶⁶

Fig. 6 Bioactive iridoids.

2.3 Apocarotenoids, sesquiterpenoids, and diterpenoids (44–55; Fig. 7)

These are carotenoid-derived oxidative cleavage products, C₁₅ terpenoids biosynthesized from three isoprene units, and C₂₀ terpenoids derived from four isoprene units, respectively.

In relation to apocarotenoids, the main bioactive metabolite is the phytohormone abscisic acid (44, Fig. 7). Compound 44 inhibited the germination of O. crenata and S. hermonthica, and it has been reported to play an essential role during seed dormancy and germination in S. hermonthica; it is suggested that the parasite disrupts host stomatal abscisic acid (44) signalling by impairing the tryptophan lock mechanism.^67,68

Fig. 7 Bioactive apocarotenoids (44), sesquiterpenoids (45–50), and diterpenoids (51–55).

Bioactive sesquiterpenoids (45–50, Fig. 7) include α-costic acid (45), which is commonly known as a metabolite produced by D. viscosa but has also been reported in Tessaria absinthioides and Zea mays.^38,69,70 Compound 45 stimulated the germination of O. cumana and P. ramosa seeds approximately by 40% when applied at 100 µM and 57% O. crenata at 400 µM.^37,38 Compound 45 also inhibited C. campestris germination by 76% at 1.7 mM but did not significantly affect the germination or radicle growth of O. crenata, O. cumana, O. minor and P. ramosa.^37,38

Other bioactive sesquiterpenoids include trichothecenes, which are produced as mycotoxins by several fungi including the genera Fusarium or Trichoderma, among others.⁷¹ Bioactive Fusarium trichothecenes include 8-acetylneosolaniol (46), which completely inhibited S. hermonthica germination at 24 µM, as well as other metabolites that differ in the number and position of acetyl groups, i.e. neosolaniol (47), acuminatin (48, which shares the name with a flavonoid),⁷² and tetraacetoxy T-2 tetraol (49), all of which also reduced S. hermonthica germination.⁷³ A study of fungal isolates from the rhizosphere of Vicia faba grown in soil infested with Orobanche spp. led to the discovery of the macrocyclic trichothecene verrucarin A (50), which is structurally related to the same scaffold of compounds 46–49; it is produced by Myrothecium verrucaria and reported as an inhibitor for the germination of O. crenata (50% at 2 µM).⁷⁴

In relation to diterpenoids (51–55, Fig. 7), the phytohormone gibberellic acid (GA₃, 51) has been shown to stimulate the germination of O. crenata and S. hermonthica.^75,76 Interestingly, when compound 51 is used in combination with other bioactive compounds it can act synergistically, increasing the germination of O. minor seeds induced by brassinolide (58).⁷⁷ Notably, alternative bioactive diterpenoids to compound 51 are fungal metabolites. Sphaeropsidin A (52), produced by the Diplodia genus, inhibited the germination of O. minor seeds by almost 100% at 100 µM, while the germination of O. crenata, O. cumana and P. ramosa was inhibited by 20–40%, and radicle growth was inhibited by almost 100% in all four species.³⁷ Moreover, fusicoccane-type diterpenoids with a characteristic 5-8-5 tricyclic ring system, such as cotylenol (53) and its glycosylated derivative cotylenin A (54), produced by the genus Cladosporium, as well as the structurally related fusicoccin A (55), produced by Phomopsis amygdali (formerly Fusicoccum amygdali), have been found to induce 50–80% germination of O. minor and S. hermonthica at 1 µM.⁷⁸

2.4 Higher terpenoids (56–65; Fig. 8)

The subgroup of higher terpenoids (56–65, Fig. 8) encompasses more structurally complex terpenoids, including sesterterpenoids (56), steroids (58–60), limonoids (61–65), and other triterpenoids (57).

Fig. 8 Bioactive higher terpenoids: sesterterpenoids (56), steroids (58–60), limonoids (61–65), and other higher triterpenoids (57).

The fungal toxin ophiobolin A (56) is a sesterterpenoid produced by the Bipolaris genus that induced approximately 50% germination of the broomrape species O. minor and P. ramosa at 0.1 µM, whereas the germination rates were about 20% for O. cumana and 10% for P. aegyptiaca.⁷⁹ Germination decreased at higher concentrations except for P. aegyptiaca, which reached 20% at 1 µM.⁷⁹ Soyasapogenol B (57), an oleanane-type triterpenoid isolated from Vicia sativa root exudates, stimulated the germination of O. minor (76% at 1 mM), but none of the other tested broomrapes.⁸⁰

Steroids, a specific subclass of triterpenoids that plays essential roles in plant growth and development, have also been shown to be relevant metabolites in the inhibition of parasitic weed germination. Brassinolide (58), known for stimulating germination in various plants such as rice and barley, and castasterone (59) have been reported to shorten the conditioning period needed for O. minor germination, but they do not induce the germination of O. minor seeds by themselves. On the other hand, compounds 58 and 52 increased germination rates when applied in combination with strigolactone-type stimulants.^77,81 A steroid isolated from Vicia sativa root exudates, trans-22-dehydrocampesterol (60), stimulated the germination of P. aegyptiaca, O. crenata, O. foetida and O. minor seeds, with germination rates ranging from 47% to 86% at 63 mM.⁸⁰

Lastly, diverse limonoid-type tetranortriterpenoids isolated from Nigerian Harrisonia abyssinica, including deoxyobacunone (61), obacunone (62), 12β-acetoxyharrisonin (64), and pedonin (65), as well as, to a lesser extent, harrisonin (63), were described as stimulators of S. hermonthica germination.⁸² These compounds were tested at 1, 0.1 and 0.01 mM, showing germination rates in the range of 98–12%. It is necessary to highlight that the structures of 63 and 64 reported by Rugutt et al. in 2001 (ref. 82) differ from those reported by Rajab et al. in 1997,⁸³ who determined them by modern 1D and 2D NMR spectroscopic methods and X-ray diffraction. Both structures are included in Fig. 8 for completeness.

3 Aromatic metabolites

Aromatic compounds are secondary metabolites widely produced by plants, fungi, and microorganisms. Characterized by one or more benzene rings as their main scaffolds, they play essential ecological roles, including defense against herbivores and pathogens, UV protection, and signalling in plant–plant and plant–microbe interactions. Their structural diversity and bioactivity make them valuable sources for practical applications in different fields.⁸⁴ Regarding their bioactivity in parasitic weed germination, both stimulatory and inhibitory effects have been reported. Bioactive aromatic compounds (66–119, Fig. 9–12) are discussed in Sections 3.1–3.4 according to the number of rings and the connectivity pattern of their aromatic frameworks. N-containing aromatic metabolites are addressed separately in Section 4 for a clearer structural classification and discussion.

Fig. 9 Bioactive monocyclic aromatic metabolites.

Fig. 10 Bioactive bicyclic coumarins (92–94), benzodioxoles (95, 96), quinones (97, 98), and polyketides (99).

Fig. 11 Bioactive flavonoids and isoflavonoids.

Fig. 12 Bioactive aromatic metabolites produced by the pea plant (Pisum sativum).

3.1 Monocyclic aromatic metabolites (66–91; Fig. 9)

This subgroup covers benzoic acid derivatives, phenylpropanoids and related phenolics. Simple monocyclic aromatic metabolites are commonly associated with inhibitory activity, either by suppressing seed germination or inhibiting radicle growth. However, earlier studies reported an exception in parasitic weed research, identifying the first stimulant of S. asiatica germination in the host species Sorghum bicolor, the hydroquinone 5-methoxy-3-(8Z,11Z)-8,11,14-pentadecatrien-1-yl-1,2,4-benzenetriol (66).⁸⁵

Broomrape species are affected by benzoic acid (67) derivatives and their methyl esters. Methyl 4-hydroxybenzoate (68) inhibited radicle growth in O. cumana, O. minor, and P. ramosa by about a 60% at 1 mM, which was higher than that induced by methyl 4-hydroxy-3-methoxybenzoate (69), which caused about 20% inhibition of those species.⁸⁶ Benzoic acid (70) also inhibited radicle growth in O. cumana (about 30% inhibition at 820 µM) and O. crenata (40% inhibition at 131 µM).^66,87

Another monocyclic phenolic is 2,4-diacetylphloroglucinol (71), produced by Pseudomonas ogarae. The presence of this polyketide in culture supernatants was correlated with a reduction of Brassica napus infection by P. ramosa.⁸⁸

A series of studies on the effects of structurally diverse aromatic metabolites on C. campestris germination found that these metabolites can exert phytotoxic effects on this parasitic weed through different modes of action.^89–91 Benzenepropanal (74) and hydrocinnamic acid (75) strongly inhibited seedling growth at 1 mM (about 70–90%), with 75 also inducing strong overproduction of protuberances in trichomes at root apices.^89,90 Other plant metabolites structurally related to compound 75, i.e. melilotic acid (76), 3-(4-methoxyphenyl)propionic acid (77), 3-(3-hydroxy-4-methoxyphenyl)propionic acid (78) and ethyl 3-phenylpropanoate (79), also inhibited C. campestris seedling growth to a lower degree.⁹¹ Milder inhibition of C. campestris seedling growth was also observed for benzoic acid (67), vanillic acid (70), benzenepropanol (72), and p-coumaric acid (81).^70–72 In addition, dihydrocaffeic acid (80), caffeic acid (82), and, to a lesser extent, benzoic acid (67) and ferulic acid (83), caused necrosis of the tissue.^90,91 Compounds 81–83 were found to inhibit radicle growth of O. crenata with little or no effect on germination inhibition. Interestingly, inhibition of P. ramosa radicle growth was reported for the plant metabolite methyl p-coumarate (84).^87,92

Regarding hydrocinnamic acid (75), structurally related compounds with unsaturation in the side chain, such as cinnamaldehyde (85) and cinnamic acid (86), have also been explored. In a structure–activity relationship study of these compounds, cinnamaldehyde (85) and methyl cinnamate (87) strongly inhibited C. campestris seedling growth (by almost 100% at 1 mM), while lower inhibition was observed (10–40% at 1 mM) for cinnamyl alcohol (73), cinnamic acid (86), 4-methoxycinnamic acid (88), 4-methylcinnamic acid (89), o-coumaric acid (90), and m-coumaric acid (91).⁸⁹

3.2 Fused bicyclic aromatic metabolites (92–99; Fig. 10)

This subgroup covers fused-bicyclic coumarins (92–94), benzodioxoles (95, 96), quinones (97, 98), and polyketides (99).

The benzopyrone coumarin (92), a common plant secondary metabolite and known germination inhibitor, was reported to stimulate the germination of P. aegyptiaca seeds (up to 87% at 10 µM), while simultaneously inhibiting radicle elongation and inducing tip bifurcation.⁹³ In another study, coumarin (92) was reported as an inhibitor of the germination of O. crenata seeds.⁷⁶ The structurally related aromatic lactones umbelliferone (93) and scopoletin (94), which are also common in plants, inhibited C. campestris seedling growth (approximately 50% and 40% inhibition at 1 mM, respectively), and compound 94 also induced necrosis.⁹⁰ Scopoletin (94) was also found to inhibit the germination of seeds and radicle growth of O. crenata and cause necrosis, suggesting additional roles in allelopathic or defense-related responses.⁸⁷ Other bioactive plant metabolites include sesamol (95) and 4-(methylenedioxy)cinnamic acid (96), with mild inhibition of C. campestris seedling growth in comparison with some of the monocyclic aromatic structures discussed in Section 4.1, but with necrosis of the tissue observed in the case of compound 95.^89,90

Fusarium solani isolates contain bioactive compounds named fusaric acids (discussed in Section 4.2 for bearing a nitrogen atom). Further studies on these isolates have also reported the naphthoquinones javanicin (97) and solaniol (98), whose absolute configuration was later demonstrated as that of (+)-solaniol, inhibiting Striga (species not specified in the report) germination by approximately 80% at 0.1 µg mL⁻¹.^94,95

A number of bicyclic lactones with a substituted aromatic ring have also demonstrated activity against parasitic weeds. Cyclopaldic acid (99), a fungal metabolite produced by genera such as Seiridium, Diplodia, Botryosphaeria, and Neofusicoccum, was found to inhibit radicle growth at 100 µM by 40–60% relative to the control in O. crenata, O. cumana, O. minor and P. ramosa. Inhibition of seed germination was not significant for these species, while stimulation of germination was found for O. cumana seeds (approximately 20% at the same concentration).³⁷

3.3 Flavonoids and isoflavonoids (100–114; Fig. 11)

Flavonoids and isoflavonoids are polyphenolic natural products built on a C6–C3–C6 scaffold, differing in the position of the B ring, which is attached at C-2 in flavonoids and at C-3 in isoflavonoids. Diverse bioactive flavonoids, isoflavonoids, and related structures have also been reported to interfere with different stages of the parasitic weed life cycle, with all showing inhibition of germination or seedling growth of broomrape, witchweed or dodder weeds.

Quercetin (100) and some of its methoxylated derivatives, i.e. 7,4′-dimethoxyquercetin (101), 3,7-dimethoxyquercetin (102) and 3,3′,4′,7-tetramethoxyquercetin (103), induced haustorium formation and inhibited radicle growth of P. ramosa by 60% at 100 µM.⁹² Notably, a recent study demonstrated that (±)-catechin (108 and (−)-108) suppresses prehaustorium formation in the parasitic weed P. ramosa and reduces its infestation in tomato, while showing no effect on GR24-induced germination or radicle elongation.⁹⁶ Radicle growth of P. ramosa was also inhibited by approximately 40% at 1 mM by 3-acetylpadmatin (109) and hispidulin (104), the latter of which also active against O. cumana and O. minor (approximately 20% growth inhibition).^86,92 The flavone ephedroidin (105), isolated from Retama raetam, strongly inhibited radicle growth of O. cumana (80% at 100 µM) and weakly inhibited that of O. crenata, O. minor and P. ramosa (around 20%). Other related flavonoids and isoflavonoids, licoflavone C (106), laburnetin (111), alpinumisoflavone (112) and hydroxyalpinumisoflavone (113), also produced by R. raetam, inhibited radicle growth of O. cumana in the range of 20–40%, while inhibition in other species did not reach 20% (only worth highlighting the case of O. crenata for compounds 106, 111–113, O. minor for compounds 111 and 112, and P. ramosa for compounds 106, 111 and 113).⁹⁷ To our knowledge, only one bioactive glycosylated flavonoid has been reported in the literature, isoschaftoside (107), isolated from Desmodium uncinatum root exudate, which showed radicle inhibition of S. hermonthica by 48% at 100 ppm.⁹⁸

Vestitol (114), a legume-specific isoflavonoid phytoalexin produced by Lotus japonicus, was reported as a strong inhibitor of S. hermonthica seedling growth, causing browning of root tips at 100 µM, germination was inhibited by 30%, suggesting its role in the host defense mechanism against parasitic intrusion.⁹⁹ Naringenin (110), mainly found in plants of the genus Citrus, inhibited C. campestris seedling growth by less than 20% at 1 mM but also induced necrosis in the apices.^90,100

3.4 Other aromatic metabolites (115–119; Fig. 12)

This final subgroup mainly focuses on different aromatic metabolites produced by pea plant (Pisum sativum). The polyphenols 1-(2,4-dihydroxyphenyl)-3-(4-methoxyphenyl)propenone (115), 1-(2,4-dihydroxyphenyl)-3-hydroxy-3-(4-hydroxyphenyl)-1-propanone (116) and peapolyphenol B (117) were found to selectively stimulate O. foetida seed germination compared with other broomrape species (up to 35% at 1 mM), highlighting a degree of host-specific chemical signalling in pea–parasite interactions.¹⁰¹

Roots of P. sativum exude the strigolactone-like aromatic metabolite peagol (118), which stimulated 42% germination of P. aegyptiaca seeds selectively at 500 µM. Compound 118 also stimulated the germination of O. foetida by 27% at the same concentration, but its activity on O. crenata and O. minor was negligible.^101,102 Pisatin (119) is also an interesting metabolite for its activity on C. campestris, since it strongly inhibited seedling growth at 1 mM (about 70–90%) and caused tissue necrosis.^89,90

4 N-containing metabolites

This section highlights the diversity of bioactive nitrogen-containing structures discovered through bioprospecting against parasitic weeds, originating not only from plants and fungi but also from less common sources such as bacteria. The incorporation of nitrogen atoms provides a particularly broad range of biological and physicochemical properties, including basicity, enhanced polarity, and increased capacity for hydrogen bonding or electrophilic interactions, which can strongly influence bioactivity. As reviewed in the following subsections 4.1–4.4, this chemical diversity is reflected in the wide range of bioactive N-containing metabolites identified in this field, which have been reported to interfere with different stages of the parasitic weed life cycle.

4.1 Amino acids and peptide-derived metabolites (120–143; Fig. 13)

They range from simple proteinogenic amino acids to oligopeptides and cyclic peptide-derived scaffolds, being widespread metabolites of primary and secondary metabolism in plants, fungi, and microorganisms (120–143, Fig. 13).

Fig. 13 Bioactive amino acids and peptide-derived metabolites.

A key discovery was the identification of several amino acids capable of inhibiting P. ramosa germination: glycine (120), alanine (121), methionine (122), lysine (123), arginine (127), proline (128), histidine (129), tryptophan (130), and phenylalanine (131). Inhibition patterns triggered by exogenous amino acids differ among parasitic species, as demonstrated by the fact that tryptophan (130) does not inhibit the development of S. hermonthica, whereas glycine (120) and proline (128) did not inhibit the germination of O. minor.^103,104 In contrast, the development of S. hermonthica was inhibited by lysine (123), tyrosine (132), and leucine (133), with compound 132 producing the lowest dry weight.¹⁰³

The germination of O. minor is inhibited by alanine (121), methionine (122), lysine (123), cysteine (124), histidine (129), tryptophan (130), phenylalanine (131), tyrosine (132), leucine (133), and isoleucine (134). Furthermore, radicle growth is inhibited by methionine (122), lysine (123), serine (125), homoserine (126), histidine (129), tryptophan (130), phenylalanine (131), tyrosine (132), leucine (133), isoleucine (134), and valine (135). Inhibition increased up to 100% at the highest concentration tested in some cases (5 mM, i.e., compounds 123, 130, and 131), reaching 90% inhibition at 1.25 mM in the case of leucine (133).^87,104 The inhibitory effect of tryptophan (130) was reported to be related to its metabolization to indole-3-acetic acid (140).^104,105 To our knowledge, the only amino acid that has been reported as an inductor of the germination of parasitic weed seeds is methionine (122, by about 30% in S. asiatica).^{75,87,104–107}

In a study on the metabolites produced by Azospirillum brasilense,¹⁰⁸ the authors tested a series of peptides with inhibitory activity against the germination of P. aegyptiaca seeds. Among the tested compounds, the most active were of synthetic origin (up to 80% inhibition and causing browning at 0.2 mg mL⁻¹), while the ones with natural occurrence, i.e. Leu–Gly (136), Leu–Gly–Gly (137), N-formylmethionyl-leucyl-phenylalanine (fMLF, 138), and the disulfide-linked cyclic peptide Cys–Asn–Gly–Arg–Cys (139), inhibited germination by up to 30%. A later study reported more bioactive cyclic dipeptides (diketopiperazines) isolated from Bacillus velezensis, including cyclo(Pro–Phe) (140), cyclo(Pro–Tyr) (141), cyclo(4-hydroxyproline-Phe) (142), and cyclo(4-hydroxyproline-Leu) (143), which strongly inhibited the germination of P. aegyptiaca. The complete inhibition of germination was observed at 4 mM with compounds 140 and 141, while approximately 70% and 80% inhibition was induced by compounds 142 and 143, respectively.¹⁰⁹ Regarding proteins, H. annuus defensin protein 1 (Ha-DEF1), a sunflower-specific defensin, was found to induce cell death in the parasitic plants O. cumana and P. ramosa, but not in S. hermonthica.¹¹⁰ However, most efforts have been focused not on the use pure proteins but on the application of antagonistic microorganisms, which release hydrolytic enzymes to damage parasitic weeds, such as Fusarium oxysporum against O. cumana or Aspergillus alliaceus against O. cernua, in order to reduce the seed bank.^111,112 These biocontrol approaches are promising, but challenges or limitations, like those related to their selectivity, should be carefully evaluated since microorganisms active against parasitic weeds may also affect the host plant directly or indirectly or the environment.

4.2 Alkaloids and N-heterocyclic metabolites (144–157; Fig. 14)

This subsection comprises metabolites characterized by the presence of a nitrogen atom within a heterocyclic scaffold, apart from the cyclic peptides discussed in Section 4.1 (144–157, Fig. 14). Alkaloids are typically distinguished by their basic nitrogen atom, but this feature does not apply to all N-containing natural products, and therefore a clear distinction is required to avoid conflating true alkaloids with other N-containing metabolites of different structural and biosynthetic origins. Most of these structures showed inhibiting properties in parasitic weeds, with the only exceptions of indole-3-acetic acid (144) and α-tomatine (146). Compound 144 is an auxin phytohormone biosynthesized from tryptophan (130) that also inhibits radicle elongation in O. minor, induces haustorium formation in P. aegyptiaca, and stimulates germination in O. crenata.^87,105,113 On the other hand, gramine (145), a plant alkaloid structurally related to tryptophan (130) and indole-3-acetic acid (144), mainly found in barley (Hordeum vulgare) but also in other plants, inhibited the germination of O. crenata by 26% and radicle growth by approximately 20% at 16 µg mL⁻¹.⁸⁷

Fig. 14 Bioactive alkaloids and other N-heterocyclic metabolites.

Other N-heterocyclic structures with stimulating activity of parasitic germination was only found for the glycoalkaloid α-tomatine (146), isolated from tomato (Solanum esculentum) roots, which stimulated the germination of P. ramosa following a dose-dependent profile (with the highest stimulation close to 80% at 100 µM), and it did not induce germination in O. cumana and O. crenata.¹¹⁴ On the other hand, some iminosaccharides (sugar mimics in which the ring oxygen is replaced by nitrogen) showed inhibiting properties. This is the case for 1-deoxynojirimycin (147), an antibiotic iminosaccharide produced by plants, insects, and bacteria, including Streptomyces and Bacillus species, which inhibited radicle elongation of O. minor at 0.1 mM but did not affect seed germination.¹¹⁵ Moreover, nojirimycin (148) completely inhibits the seed germination of O. minor at 10 µM and slightly inhibits the germination of Striga gesnerioides, which is the only non-strigolactone metabolite found to be active against this species to date.¹¹⁶ Due to its low stability, compound 148 is frequently used as a bisulfite.¹¹⁷

Aromatic N-containing phytohormones are also classified in this subgroup. In particular, kinetin (149), was reported as an inhibitor of O. crenata germination.¹¹⁸ Moreover, compound 149 and the structurally related cytokinin zeatin (150) were found to promote ethylene biosynthesis (a known stimulant of S. asiatica germination), especially when applied in combination, consistent with previous reports on the germination activity described for both compounds 149 and 150 in S. asiatica.^75,119,120 A related bioactive structure was also the nucleotide adenosine 3′,5′-cyclic monophosphate (cAMP, 151), which appears to play a role in the germination of the root parasitic plant O. minor.¹²¹ Addition of compound 151 restored germination rates in seeds previously exposed to light or supraoptimal temperatures during the conditioning period. Endogenous levels of 151 increased following treatment with gibberellic acid (51).¹²¹

Fusarium species have been considered as potential control agents of Orobanche and Striga species.^122,123 Their effectiveness against the Striga species has been known since the end of the last century and has been attributed to the presence of bioactive metabolites like fusaric acid (152) and 9,10-dehydrofusaric acid (153), which inhibit S. hermonthica germination at low concentrations (1 µM).¹²⁴ Moreover, compound 152 is a fungal toxin that can elicit various plant defense responses at 0.1 µM without toxic effects on Arabidopsis thaliana, while it inhibits the germination of P. ramosa seeds at the same concentration.^123,125

More complex structures included different plant metabolites. The plant-derived benzoxazinoid 2-benzoxazolinone (154) strongly inhibited radicle growth of O. crenata by 97% at 74 µM, and germination by 60% at 30 µM.^87,90 Compound 154 inhibited C. campestris growth by 89% at 1 mM, whereas 6-hydroxy-benzoxazolinone (155), a metabolite only reported in Acanthus arboreus, caused mild inhibition (by approximately 20%).^90,126 Castanospermine (156), a plant indolizidine alkaloid originally isolated from the seeds of Castanospermum australe (which remains its best-established natural source), inhibited O. minor seed germination by 16% at 10 µM.^127,128 Trigoxazonane (157) was isolated from Trigonella foenum-graecum (fenugreek), a legume species that can reduce O. crenata infection in intercropped strategies, and inhibited O. crenata seed germination (by approximately 50% at 10.32 µg cm⁻²).¹²²

Cycloheximide (158), a glutarimide metabolite produced by Streptomyces, inhibited the seed germination of O. minor by 50% at 9.2 nM.¹²⁹ The most complex structures in this subgroup are fungal lactams known as cytochalasans, produced by Pyrenophora semeniperda, a species shown to possess a diverse metabolite profile.¹³⁰ Cytochalasins A (159) and B (160), along with deoxaphomin (161), inhibited the germination of P. ramosa (by 57% relative to the control at 100 µM). Compound 161 also inhibited the germination of O. crenata (38% inhibition) and strongly inhibited radicle growth in O. crenata, O. cumana, O. minor, and P. ramosa (80% inhibition relative to the control).³⁷

4.3 Nitrile- and isothiocyanate-containing metabolites (162–173; Fig. 15)

This subgroup comprises N-containing metabolites in which the nitrogen atom is part of a nitrile or isothiocyanate functional group, which have been identified through bioprospecting studies of rye (Secale cereale L.) and the genus Brassica (162–173, Fig. 15).

Fig. 15 Bioactive nitrile (162–165)- and isothiocyanate (166–173)-containing metabolites.

Specifically, bioactive metabolites containing a cyano group in their structure correspond to a series of ryecyanatines and ryecarbonitrilines (162–165) isolated from rye (Secale cereale L.) root exudates, encompassing monocyclic (ryecyanatine A, 162) and bicyclic (163–165) aromatic structures.¹³¹ Metabolites 162–165 showed significant activity in bioassays against parasitic weeds, affecting both germination and radicle growth inhibition, with key structure–activity relationships observed. Ryecarbonitriline A (164) stimulated the germination of O. cumana seeds by approximately 45% at 0.6 mM, while null germination was observed for O. crenata and O. minor. Ryecarbonitriline B (165) and ryecyanatines A (162) and B (163) did not stimulate the germination of the tested Orobanche spp. The germination of O. cumana was completely inhibited by compound 162 at 0.6 mM, and strongly by ryecarbonitriline B (165; 80% at 0.676 mM). Ryecyanatine A (162) also inhibited O. crenata by approximately 40% and O. minor by 30% at 0.6 mM, while compound 165 only inhibited O. crenata by 20% at 0.676 mM. Moreover, compound 162 completely inhibited radicle growth in three Orobanche spp. At 0.6 mM, while radicle growth in O. cumana was strongly inhibited by 165 and only mildly by compounds 163 and 164. On the other hand, radicle growth in O. crenata and O. minor was weakly inhibited by compounds 163–165.¹³¹

Isothiocyanates 166–173, derived from species of the genus Brassica, represent another important group of metabolites that induce broomrape germination and are notable as the only sulfur-containing metabolites reviewed. In particular, isothiocyanates from Brassica napus, including, sulforaphane (166), erucin (167), berteroin (168), 4-pentenyl isothiocyanate (171), benzyl isothiocyanate (172) and 2-phenylethyl isothiocyanate (173), stimulated P. ramosa germination (with EC₅₀ of 0.1–0.001 µM).¹³² This study showed that changes in the side chain only moderately affect potency within this series, with sulforaphane (166) as the least active and 4-pentenyl isothiocyanate (171) as the most potent, suggesting that a simple aliphatic unsaturated chain may be slightly more favorable than functionalized substituents.¹³² A later structure–activity relationship study was carried out, which also included heptyl isothiocyanate (170), with a stimulatory activity of 55% at 1 mM, and other derivatives of non-natural origin.¹³³ P. aegyptiaca germination was also significantly stimulated by 169, 170, 172, and 173 at 1 mM, reporting 20–30% germination rates, highlighting how structural requirements of isothiocyanates for germination stimulation were similar but not the same between P. ramosa and O. aegyptiaca.¹³³

4.4 Other N-containing metabolites (174–177; Fig. 16)

This final subgroup comprises structurally diverse metabolites that do not fit readily into the previous subsections 4.1–4.3 (174–177, Fig. 16). It is worth first highlighting the case of the simplest N-containing structure, namely, urea (174), whose application after conditioning of O. crenata seeds slightly reduced their germination by 18% at 8 mM. Its inhibitory effect is hypothesized to be indirect (potentially resulting from its conversion to ammonium), which has prompted its use as scaffold for the design of strigolactone antagonists against P. aegyptiaca and S. asiatica germination.^134,135

Fig. 16 Other bioactive N-containing metabolites.

Two amino acid derivatives, 1-aminocyclopropane-1-carboxylic acid (175) and aminoethoxyvinylglycine (176), were classified in this subgroup because they are specialized non-proteinogenic derivatives rather than canonical amino acids or peptide-based metabolites (reviewed in Section 4.1). 1-Aminocyclopropane-1-carboxylic acid (175) promoted ethylene biosynthesis (stimulant of S. asiatica germination), as aforementioned for kinetin (149) and zeatin (150), while aminoethoxyvinylglycine (176) was shown to inhibit germination in S. hermonthica and S. asiatica acting as an ethylene biosynthesis inhibitor.¹¹⁹

With a more complex structure, AAL-toxin (177), a sphinganine-analog mycotoxin produced by Alternaria alternata (one of the most relevant phytopathogenic fungi, also with recognized allelopathic roles), induced programmed cell death in the parasitic weeds O. cumana and P. ramosa, triggering responses rarely observed in common plants, which highlights the potential of A. alternata as biocontrol agent.^136,137

5 Miscellaneous bioactive metabolites

This final section comprises the bioactive metabolites that could not be assigned unambiguously to the structural classes discussed above, ranging from volatile and signalling molecules to quinones and other oxygenated cyclic ketones, as well as carbohydrates and polyols.

5.1 Volatile and signalling molecules (178–180; Fig. 17)

This subsection includes key low-molecular weight signalling molecules involved in plant communication and stress responses (178–180, Fig. 17). Ethylene (178) is a phytohormone that has been shown to stimulate the germination of S. asiatica and S. hermonthica seeds, but its application in agronomy is limited due to its gaseous nature, instability, and non-selective action.¹²¹ This compound was applied to reduce Striga emergence by 90% in one of the first practical applications of suicidal germination for management of parasitic weeds.¹³⁸ As previously discussed, metabolites 149, 150 and 175 could enhance its biosynthesis in planta.¹¹⁹

Fig. 17 Miscellaneous volatile and signalling bioactive molecules.

Jasmonic acid (179), a lipid-derived plant growth regulator and phytohormone, was identified as a germination elicitor for S. hermonthica, inducing 26% stimulation at 1 mM. In the same study, the related phytohormone methyl jasmonate (180) almost doubled the germination activity at the same concentration. 179 did not significantly stimulate O. minor germination, while 180 induced 67% stimulation at 100 µM.¹³⁹

5.2 Butenolides and related polyacetylenes (181–189; Fig. 18)

This subsection comprises diverse plant metabolites characterized by butenolide rings and related polyacetylene-derived scaffolds, many of which are associated with allelopathic interactions and other ecological functions (181–189, Fig. 18).

Fig. 18 Bioactive butenolides and related polyacetylenes.

Conyza bonariensis produces (4Z)-lachnophyllum lactone (181) and (4Z,8Z)-matricaria lactone (182), two strong inhibitors of radicle growth in C. campestris, O. crenata, O. cumana, O. minor, and P. ramosa, inducing 80–100% inhibition at 1 mM. In contrast, (4E,8Z)-matricaria lactone (183) and (4Z)-lachnophyllum methyl ester (184) isolated from this plant showed 0–20% inhibition at the same concentration when tested on the same broomrape species.^87,140 A comparative ecotoxicological study with conventional herbicides on aquatic and terrestrial organisms highlighted compounds 181 and 182 as lower-impact alternatives.¹⁴¹ Bioactive lactones isolated from Streptomyces albus bearing a side chain, 5-(6-methyloctyl)-2(5H)-furanone (185), 5-(6-hydroxy-6-methyloctyl)-2(5H)-furanone (186) and 5-(6-methyl-7-oxooctyl)-2(5H)-furanone (187), stimulated the germination of O. minor. Compound 185 was the most active with approximately 75% at 10 µM, while compounds 186 and 187 showed reduced activity, with stimulation activity below 30%.¹⁴²

Even though the previously described plant-derived furanones 181–187 have been reported as inhibiting agents of parasitic plants, other more complex structures bearing furanones (188 and 189) showed remarked stimulation activity in parasitic plant germination. Karrikin₁ (188), a compound identified in smoke from the combustion of plant materials, induced around 60% germination of O. minor and S. hermonthica at 0.1 µM, with the latter species showing germination levels comparable to those achieved in this study with GR24 at the same concentration.¹⁴³ At lower potency, compound 188 also stimulated the germination of P. aegyptiaca, P. ramosa, and other less studied parasitic species (belonging to genera Orobanche, Cistanche, Conopholis and Lathraea). Peagoldione (189), produced by P. sativum, selectively stimulated the germination of P. aegyptiaca seeds, inducing 49% germination at 2 mM, while germination of O. crenata and O. minor was negligible.¹⁰²

5.3 Quinones and other oxygenated cyclic ketones (190–196; Fig. 19)

This subsection comprises structurally diverse metabolites characterized by quinone or other highly oxygenated cyclic ketone moieties (190–196, Fig. 19).

Fig. 19 Bioactive quinones and other oxygenated cyclic ketones.

1,4-Benzoquinones are produced by higher plants (such as Sorghum spp), fungi, bacteria and animals and are involved in important biological processes. 1,4-Benzoquinone (190) and 2,6-dimethoxy-p-benzoquinone (191) inhibited radicle growth in O. cumana, O. minor and P. ramosa at 1–0.01 mM in a dose–response manner, with compound 190 showing stronger activity than compound 191 while offering mild inhibition (47%) against C. campestris seedling growth; both compounds induced necrosis in O. cumana radicles.^{90,92,144,145} Compound 191 triggered haustorium development in both S. asiatica and P. ramosa, indicating plant species-specific activity compared with O. cumana or O. minor. Sorgoleone (192) and some of its derivatives, differing in side-chain length and degree of unsaturation, were isolated from S. bicolor and stimulated the germination of S. asiatica, with no effect on haustorium development.¹⁴⁶

Cyclohexene derivatives produced by the fungal genus Diplodia have demonstrated both stimulatory and inhibitory effects on broomrape species. Sphaeropsidone (193) and epi-sphaeropsidone (194) were described as the first compounds exhibiting haustorial-inducing activity in broomrape species (O. crenata, O. cumana and S. hermonthica).¹⁴⁷ In addition, radicle growth inhibition in O. crenata, O. cumana, O. minor, and P. ramosa was observed at 100 µM for both compounds, possibly as a consequence of haustorium induction. Radicle growth inhibition was higher for compound 193, while induced germination inhibition was low and similar for both compounds. The fungal cyclohexene epoxide epi-epoformin (195) strongly inhibited O. cumana and O. crenata at 100 µM while showing weaker activity against O. minor and P. ramosa (approximately 50–70% inhibition relative to the control); it also almost completely inhibited the germination of O. cumana seeds and significantly decreased germination in O. crenata and P. ramosa.³⁷

It could be also highlighted that a study on the bioactivities of intermediates involved in the synthesis of strigol revealed several bioactive structures, including (7,7-dimethyl-1,4-dioxo-2,3,4,5,6,7-hexahydro-1H-inden-2-yl)acetic acid (196), which was shown to stimulate the germination of P. ramosa seeds by 34% at 100 µM, and whose occurrence as natural product has only been documented, to the best of our knowledge, in the essential oils of Myrtus communis L.^148,149

5.4 Carbohydrates and polyols (197–199; Fig. 20)

Although commonly associated with primary metabolism, carbohydrates and related polyols can also participate in biologically relevant interactions with parasitic weeds, influencing processes such as seed germination, early seedling development, and host–parasite establishment (197–199, Fig. 20). In particular, inositol (197) and sucrose (198) stimulated the germination of S. asiatica. Moreover, in narbon bean plants, increasing compound 198 concentrations in water reduced the germination of O. crenata seeds and significantly decreased the number of parasite attachments.^75,150

Fig. 20 Bioactive carbohydrates and polyols.

The plant trisaccharide planteose (199), described as a storage carbohydrate, plays a critical role during early germination stages of O. minor. Metabolization of this compound involves hydrolysis to sucrose upon strigolactone stimulation and could be targeted as a strategy for selective control of this parasitic species.¹⁵¹

Taken together, these findings suggest that carbohydrate-based and polyhydroxylated structures also deserve attention in parasitic plant research. This view is further supported by the bioactivity reported for the glycosylated or sugar-related natural products discussed above, including iridoids 39–43 (Fig. 6), the diterpenoids cotylenol and cotylenin A (53 and 54, Fig. 8), the flavonoid isoschaftoside (107, Fig. 11), the glycoalkaloid α-tomatine (146, Fig. 11), and the iminosaccharides 1-deoxynojirimycin and nojirimycin (147 and 148, Fig. 14).

6 Lipophilicity trends of the bioactive metabolites

After analyzing the set of 199 non-strigolactone natural products reported as active against parasitic weeds in the literature, a structure–property relationship can be established by considering molecular lipophilicity. In particular, the distribution of calculated partition coefficients (clog P values, calculated using ChemBioDraw Ultra 21.0)^152,153 provides insights into the physicochemical space most frequently associated with biological activity. Two main clog P intervals emerge as particularly enriched in bioactive structures (Fig. 21A): moderately lipophilic compounds with clog P values between 1.0 and 3.0 (93 compounds, 47%), and hydrophilic compounds with clog P values below 0 (47 compounds, 24%). These observations suggest that both moderate lipophilicity and pronounced hydrophilicity may represent more favorable ranges for discovering bioactive compounds active against parasitic weeds. Nevertheless, in view of the clog P values of germination-stimulant compounds (Fig. 21B) and metabolites with inhibitory activity in germination or seedling development (for simplicity, hereafter referred to as “inhibitors”; Fig. 21C), molecular lipophilicity alone does not provide clear ranges for distinguishing between stimulants and inhibitors.

Fig. 21 Histograms showing the distribution of clog P values for (A) all the bioactive non-strigolactone metabolites reviewed (1–199), (B) only the germination stimulants, (C) only the metabolites with inhibitory activity in germination or seedling growth, and (D) only the sesquiterpene lactones with germination activity.

When focusing specifically on germination-stimulant sesquiterpene lactones, the most common clog P interval is 0.5–1.5 (44% of the germination-stimulant sesquiterpene lactones, Fig. 21D). Key conclusions can be drawn by comparing this lipophilicity range with that of the 23 canonical and 19 non-canonical strigolactones reported to date, to the best of our knowledge.^154,155 For canonical strigolactones, most exhibit clog P values in the range of 0.20 to 1.99 (Table 1), with only a few exceptions above or below this interval. This distribution is consistent with the finding of a majority of germination-stimulant sesquiterpene lactones whose clog P is in the range 0.5–1.5 (Fig. 21D), which may also help to rationalize the remarkable stimulant activity of sesquiterpene lactones.

Table 1 clog P Values of all the reported canonical and non-canonical strigolactones

Canonical strigolactones	clog P	Non-canonical strigolactones	clog P
5-Deoxystrigol and 4-deoxyorobanchol	2.28	Carlactone	4.74
Orobanchyl acetate and 4α-acetoxy-5-deoxystrigol	1.99	Methyl carlactonoate	4.39
Sorgolactone	1.76	Carlactonic acid	3.87
Solanacyl acetate	1.56	Hydroxymethyl carlactonoate	3.46
Fabacyl acetate	1.34	4-Hydroxycarlactone	2.86
Strigyl acetate	1.14	16-Hydroxycarlactone	2.76
Orobanchol and 4α-hydroxy-5-deoxystrigol	1.09	3-Hydroxycarlactone	2.66
Solanacol	0.73	Methyl-4-hydroxycarlactonoate	2.51
Medicaol	0.71	Methyl-16-hydroxycarlactonoate	2.41
Fabacol	0.49	Heliolactone	2.21
Sorgomol	0.30	18-Hydroxycarlactonoic acid	2.08
Strigone	0.26	4-Hydroxycarlactonoic acid	1.98
Strigol and 7β-hydroxy-5-deoxystrigol	0.20	16-Hydroxycarlactonoic acid	1.88
7-Oxoorobanchyl acetate	−0.14	3-Hydroxycarlactonoic acid	1.78
7α-Hydroxyorobanchol, 7β-hydroxyorobanchol, 7α-hydroxyorobanchyl acetate, and 7β-hydroxyorobanchyl acetate	−1.00	Vitislactone	1.58
		Zealactone (also known as methyl zealactonoate)	1.53
		Lotuslactone	1.27
		Zeapyranolactone	1.00
		Avenaol	−0.20

---

In contrast, non-canonical strigolactones display a much wider lipophilicity range, with log P values ranging from 4.74 to −0.20 (Table 1). This variation suggests that lipophilicity is a less critical determinant of their biological activity compared with canonical strigolactones and the bioactive non-strigolactone metabolites reviewed, and that other molecular features likely govern their specific modes of action and selectivity.

In relation to the compounds reviewed herein with inhibitory activity (either on germination or subsequent seedling growth), these are predominantly distributed within two clog P intervals: 1.5–2.25 and <0 (Fig. 21C). This pattern is attributed to the prevalence of aromatic metabolites (40% of the inhibitors, 51% of which have clog P values in the range of 1.5–2.5) and N-containing structures (35% of the inhibitors, 50% of which have clog P values of <−1.0).

7 Conclusions and prospects

A wide array of metabolites has been discovered as the germination stimulants, germination inhibitors, or growth inhibitors of parasitic plants, positioning them as valuable structural models for allelopathy research and bioherbicide development. Their classification in Sections 2–5 based on their chemical structures could be summarized in relation to each parasitic species and the type of bioactivity, as presented in Fig. 22.

Fig. 22 Classification of the reviewed metabolites (1–199) according to the parasitic species in which they showed significant activity. Compound numbers in red are terpenes and terpenoids (bold, sesquiterpene lactones), blue are aromatic metabolites, purple are N-containing metabolites, and green are miscellaneous.

Moreover, as depicted in Fig. 23, the structural class of the reviewed metabolites appears to be a key determinant of their type of biological activity on parasitic weeds. Germination-stimulant effects have been most frequently reported for terpenes and terpenoids (51.0%), mainly sesquiterpene lactones (34.7%), followed by N-containing compounds (23.5%). By contrast, inhibitory activities are predominantly associated with aromatic (41.7%) and N-containing (36.5%) structures. Notably, some metabolites have been reported to exhibit both stimulatory and inhibitory effects depending on the parasitic species studied (see as an example the case of α-costic acid, 45, or cyclopaldic acid, 99).

Fig. 23 Distribution of bioactive non-strigolactone natural products in relation to their structures and type of bioactivity on parasitic weeds. Inhibitors: compounds with inhibitory activity on germination or seedling growth, or other phytotoxic effects.

Further insights in the field can be drawn from the specificity of parasitic species in relation to the type of structures capable of eliciting germination. For this purpose, Table 2 summarizes bioactive compounds that stimulate parasitic plant seed germination by more than 50% at concentrations up to 100 µM. Among the main Orobanche spp., O. cumana showed the best stimulation for sesquiterpene lactones, especially costunolide (1) and parthenolide (6), while O. minor showed the best stimulation for diverse terpenoids and miscellaneous compounds. Notably, no non-strigolactone natural product was found to be active at achieving 50% germination of O. crenata, in agreement with the pronounced specificity of this species towards strigolactones. On the other hand, Phelipanche spp. showed a wider variety of stimulants, i.e. dehydrocostuslactone (14, sesquiterpene lactone) and coumarin (92, aromatic) for P. aegyptiaca; and an array of N-containing and sesquiterpene lactones compounds for P. ramosa, with 4-pentenyl isothiocyanate (171) being the most active at the minimum concentration, followed by other isothiocyanates. In relation to Striga spp., it was found that sesquiterpene lactones are the main germination stimulants of S. asiatica, especially 4,5-epoxy-8-epi-inunolide (10), dihydrocostunolide (5), parthenolide (6) and costunolide (1); notably, compound 188 and different terpenoids (especially cotylenin A (54) and fusicoccin A (55)) were the most active metabolites reported for S. hermonthica germination.

Table 2 Specificity of each parasitic species to non-strigolactone natural products, illustrating the compounds that stimulate germination by over 50% at the minimum concentration up to 100 µM, in decreasing order of the stimulation level for each species

Parasitic species	Bioactive compound, minimum active concentration, and percentage stimulation
O. cumana	1 (sesquiterpene lactone), 0.1 µM, 70% (ref. 15)
	6 (sesquiterpene lactone), 1 µM, 75% (ref. 26)
	14 (sesquiterpene lactone), 10 µM, 60% (ref. 20)
	30 (sesquiterpene lactone), 100 µM, 75% (ref. 55)
	12 (sesquiterpene lactone), 100 µM, 60% (ref. 37)
O. minor	188 (miscellaneous), 0.1 µM, 58% (ref. 142)
	56 (sesterterpenoid), 0.1 µM, 50% (ref. 79)
	54 (diterpenoid), 10 µM, 79% (ref. 78)
	185 (miscellaneous), 10 µM, 75% (ref. 141)
	55 (diterpenoid), 10 µM, 56% (ref. 78)
P. aegyptiaca	14 (sesquiterpene lactone), 0.1 µM, 50% (ref. 46)
	92 (aromatic benzopyrone), 10 µM, 87% (ref. 93)
P. ramosa	171 (isothiocyanate), 0.001 µM, 50% (ref. 131)
	167, 168, 172 and 173 (isothiocyanates), 0.01 µM, 50% (ref. 131)
	14 (sesquiterpene lactone), 0.1 µM, 70% (ref. 45)
	26 (sesquiterpene lactone), 0.1 µM, 64% (ref. 55)
	30 (sesquiterpene lactone), 0.1 µM, 56% (ref. 55)
	29 (sesquiterpene lactone), 0.1 µM, 54% (ref. 55)
	56 (sesterterpenoid), 0.1 µM, 50%,^79 and 166 (isothiocyanate), 0.1 µM, 50% (ref. 131)
	27 (sesquiterpene lactone), 1 µM, 59% (ref. 55)
	146 (glycoalkaloid), 1 µM, 55% (ref. 114)
	28 (sesquiterpene lactone), 1 µM, 50% (ref. 55)
	16 (sesquiterpene lactone), 10 µM, 50% (ref. 48)
	32 (sesquiterpene lactone), 100 µM, 60% (ref. 48)
	17 (sesquiterpene lactone), 100 µM, 50% (ref. 48)
S. asiatica	10 (sesquiterpene lactone), 0.001 µM, 65% (ref. 31)
	5 (sesquiterpene lactone), 0.001 µM, 55% (ref. 31)
	6 (sesquiterpene lactone), 0.001 µM, 54% (ref. 31)
	1 (sesquiterpene lactone), 0.001 µM, 53% (ref. 31)
	7 (sesquiterpene lactone), 0.1 µM, 76% (ref. 32)
	21 (sesquiterpene lactone), 100 µM, 66% (ref. 31)
	23 (sesquiterpene lactone), 100 µM, 53% (ref. 31)
S. hermonthica	188 (miscellaneous), 0.1 µM, 61% (ref. 142)
	54 (diterpenoid), 10 µM, 79% (ref. 78)
	55 (diterpenoid), 10 µM, 57% (ref. 78)
	62 (limonoid terpenoid), 100 µM, 90% (ref. 82)
	63 (limonoid terpenoid), 100 µM, 89% (ref. 82)
	61 and 65 (limonoid terpenoids), 100 µM, 80% (ref. 82)
	180 (miscellaneous), 100 µM, 67% (ref. 138)

---

Although sesquiterpene lactones appear to predominate, largely because they include some of the most potent germination stimulants reported for several parasitic species, a true “gold standard” metabolite effective across all species has not yet been defined, either due to marked species specificity or because most studies do not evaluate activity across the full range of parasitic weeds. It is interesting to note that only a few compounds (mainly sesquiterpene lactones and isothiocyanates) are able to induce the germination of parasitic seeds at concentrations comparable to those of natural strigolactones. However, additional roles in haustorium initiation and host chemotropism underscore the need for in-depth investigations into all chemical signals involved in parasitic plant development.

8 Conflicts of interest

There are no conflicts to declare.

9 Data availability

No primary research results, software or code have been included, and no new data were generated or analyzed as part of this review.

10 Acknowledgements

J. G. Z. thanks the Universidad de Cádiz for the postdoctoral support (grant Number: 2025-042/PU/POSTDOC-R3/CD).

11 References

1. M. Fernández-Aparicio, P. Delavault and M. P. Timko, Plants, 2020, 9, 1184.
2. M. Fernández-Aparicio, A. Pérez-de-Luque, E. Prats and D. Rubiales, Ann. Appl. Biol., 2008, 153, 117–126.
3. V. Dell'Oste, F. Spyrakis and C. Prandi, Molecules, 2021, 26, 4579.
4. X. Xie, K. Yoneyama and K. Yoneyama, Annu. Rev. Phytopathol., 2010, 48, 93–117.
5. M. Bürger, D. Peterson and J. Chory, iScience, 2024, 27, 12111491.
6. S. Ogawa, S. Cui, A. R. White, D. C. Nelson, S. Yoshida and K. Shirasu, Nat. Commun., 2022, 13, 4653.
7. A. Fajardo, C. Reyes-Bahamonde, F. E. Fontúrbel, F. I. Piper and R. Callaway, New Phytol., 2025, 247, 470–476.
8. K. Kawada, I. Takahashi, T. Saito, T. Inayama, Y. Seto, T. Nomura, Y. Sasaki, T. Asami, S. Yajima and S. Ito, J. Agric. Food Chem., 2025, 73, 15389–15398.
9. M. Z. Shimels, S. Rendine, C. Ruyter-Spira, P. J. Rich, G. Ejeta and H. J. Bouwmeester, Plants People Planet, 2025, 7, 318–330.
10. J. D. Rudolf, T. A. Alsup, B. Xu and Z. Li, Nat. Prod. Rep., 2021, 38, 905–980.
11. J. G. Zorrilla, A. Cala, C. Rial, F. J. R. Mejías, J. M. Molinillo, R. M. Varela and F. A. Macías, J. Agric. Food Chem., 2020, 68, 9636–9645.
12. F. A. Macías, M. D. Garcia-Diaz, A. Perez-de-Luque, D. Rubiales and J. C. Galindo, J. Agric. Food Chem., 2009, 57, 5853–5864.
13. Y. Jin, Y. Xie, P. Zhang, A. Khan, Z. Zhou and L. Liu, Chin. Herb. Med., 2024, 16, 70–81.
14. S. M. Adekenov, Fitoterapia, 2017, 121, 16–30.
15. F. M. Raupp and O. Spring, J. Agric. Food Chem., 2013, 61, 10481–10487.
16. A. Krupp, B. Bertsch and O. Spring, Front. Plant Sci., 2021, 12, 699068.
17. R. Kaur, P. Sharma, U. Bhardwaj and K. Rupinder, Discov. Chem., 2025, 2, 131.
18. M. M. Viveiros, M. C. Barreto and A. M. Seca, Separations, 2022, 9, 211.
19. S. Kasana, M. D. Dwivedi, P. L. Uniyal and A. K. Pandey, Phytotaxa, 2020, 450, 173–187.
20. K. Cao, W. Qian, Y. Xu, Z. Zhou, Q. Zhang and X. Zhang, Nat. Prod. Res., 2016, 30, 2160–2163.
21. H. K. Lee, H. E. Song, H. B. Lee, C. S. Kim, M. Koketsu, L. Thi My Ngan and Y. J. Ahn, PLoS One, 2014, 9, e95530.
22. C. A. L. Kassuya, A. Cremoneze, L. F. L. Barros, A. S. Simas, F. da Rocha Lapa, R. Mello-Silva, M. É. Alves Stefanello and A. R. Zampronio, J. Ethnopharmacol., 2009, 124, 369–376.
23. B. Ferrari, P. Castilho, F. Tomi, A. I. Rodrigues, M. do Ceu Costa and J. Casanova, Phytochem. Anal., 2005, 16, 104–107.
24. S. J. Wang, T. H. Wang, C. L. Kao, H. C. Yeh, H. T. Li and C. Y. Chen, Chem. Nat. Compd., 2023, 59, 1002–1004.
25. J. C. Galindo, A. P. de Luque, J. Jorrín and F. A. Macías, J. Agric. Food Chem., 2002, 50, 1911–1917.
26. A. P. De Luque, J. C. G. Galindo, F. A. Macías and J. Jorrín, Phytochemistry, 2000, 53, 45–50.
27. F. S. El-Feraly, Phytochemistry, 1984, 23, 2372–2374.
28. T. A. Hussien, A. A. Ahmed, M. A. El-Maghraby, A. M. Abou-Douh and J. Pharm, Med. Sci., 2021, 1, 49–54.
29. E. W. Coronado-Aceves, C. Velázquez, R. E. Robles-Zepeda, M. Jiménez-Estrada, J. Hernández-Martínez, J. C. Gálvez-Ruiz and A. Garibay-Escobar, Pharm. Biol., 2016, 54, 2623–2628.
30. L. V. Muschietti and J. L. Ulloa, Nat. Prod. Commun., 2016, 11, 1569–1578.
31. N. H. Fischer, J. D. Weidenhamer, J. L. Riopel, L. Quijano and M. A. Menelaou, Phytochemistry, 1990, 29, 2479–2483.
32. N. H. Fischer, J. D. Weidenhamer and J. M. Bradow, Phytochemistry, 1989, 28, 2315–2317.
33. J. S. Calderon, L. Quijano, F. Gómez-Garibay, D. M. Sanchez, T. Rios and F. R. Fronczek, Phytochemistry, 1987, 26, 1747–1750.
34. R. S. Dhillon, P. S. Kalsi, W. P. Singh, V. K. Gautam and B. R. Chhabra, Phytochemistry, 1987, 26, 1209–1210.
35. A. Rustaiyan, Z. Sharif, A. Tajarodi and A. S. Sadjadi, Phytochemistry, 1987, 26, 2856–2857.
36. A. R. de Vivar, C. A. L. Perez, N. C. Leon and G. Delgado, Phytochemistry, 1980, 21, 2905–2908.
37. A. Cimmino, M. Fernández-Aparicio, A. Andolfi, S. Basso, D. Rubiales and A. Evidente, J. Agric. Food Chem., 2014, 62, 10485–10492.
38. A. Andolfi, N. Zermane, A. Cimmino, F. Avolio, A. Boari, M. Vurro and A. Evidente, Phytochemistry, 2013, 86, 112–120.
39. A. Moeini, M. Masi, M. C. Zonno, A. Boari, A. Cimmino, O. Tarallo, M. Vurro and A. Evidente, Org. Biomol. Chem., 2019, 17, 2508–2515.
40. N. Serino, A. Boari, G. Santagata, M. Masi, M. Malinconico, A. Evidente and M. Vurro, Pest Manage. Sci., 2021, 77, 646–658.
41. J. G. Zorrilla, M. Innangi, A. Cala Peralta, G. Soriano, M. T. Russo, M. Masi, M. Fernández-Aparicio and A. Cimmino, Plants, 2024, 13, 178.
42. F. Scribe, Justus Liebigs Ann. Chem., 1842, 44, 298–299.
43. M. Suchý, V. Benešová, V. Herout and F. Šorm, Chem. Ber., 1960, 93, 2449–2456.
44. M. Suchý, V. Herout and F. Šorm, Collect. Czech. Chem. Commun., 1965, 30, 2863–2864.
45. C. Rial, S. Tomé, R. M. Varela, J. M. Molinillo and F. A. Macías, J. Chem. Ecol., 2020, 46, 871–880.
46. D. M. Joel, S. K. Chaudhuri, D. Plakhine, H. Ziadna and J. C. Steffens, Phytochemistry, 2011, 72, 624–634.
47. R. Kaur, R. Kaur, R. Singh, U. Bhardwaj and P. Sharma, J. Biol. Act. Prod. Nat., 2024, 14, 551–566.
48. D. M. Cárdenas, J. Bajsa-Hirschel, C. L. Cantrell, C. Rial, R. M. Varela, J. M. Molinillo and F. A. Macías, Pest Manage. Sci., 2022, 78, 4240–4251.
49. W. C. Mar and J. H. Shin, Kor. Pat., KR101418458B1, 2014.
50. Z. Zeng, H. Huang, H. He, L. Qiu, Q. Gao, Y. Li and W. Ding, Molecules, 2022, 27, 5915.
51. J.-J. Qin, J.-X. Zhu, Q. Zeng, X.-R. Cheng, Y. Zhu, S.-D. Zhang, L. Shan, H.-Z. Jin and W.-D. Zhang, J. Nat. Prod., 2011, 74, 1881–1887.
52. M. Vasquez, L. Quijano, L. E. Urbatsch and N. H. Fischer, Phytochemistry, 1992, 31, 2051–2054.
53. A. B. Gutiérrez and W. Herz, Planta Med., 1990, 56, 295–297.
54. F. Bohlmann, K. Umemoto and J. Jakupovic, Phytochemistry, 1985, 24, 1017–1019.
55. J. G. Zorrilla, C. Rial, R. M. Varela, J. M. Molinillo and F. A. Macías, Phytochem. Lett., 2019, 31, 229–236.
56. A. A. Elnour and N. H. Abdurahman, Heliyon, 2024, 10, e37790.
57. H. I. Lu, K. L. Chen, C. Y. Yen, C. Y. Chen, T. M. Chien, C. Shu and H. W. Chang, Pharmaceuticals, 2024, 17, 230.
58. J. H. Oh, J. Kim, F. Karadeniz, H. R. Kim, S. Y. Park, Y. Seo and C. S. Kong, Molecules, 2021, 26, 3585.
59. J. Zhang, Q. Xu, H. Y. Yang, M. Yang, J. Fang and K. Gao, Front. Mol. Biosci., 2021, 8, 710676.
60. F. S. El-Feraly, Y. M. Chan and D. A. Benigni, Phytochemistry, 1979, 18, 881–882.
61. K. Michalska, K. Pieron and A. Stojakowska, Nat. Prod. Commun., 2018, 13, 1934578X1801300403.
62. C. Y. Chen, S. T. Huang, H. M. D. Wang, S. L. Liu, M. J. Cheng and H. T. Li, Chem. Nat. Compd., 2025, 61, 698–701.
63. H. Mansilla and J. A. Palenzuela, Phytochemistry, 1999, 51, 995–997.
64. A. Cala, J. M. Molinillo, M. Fernández-Aparicio, J. Ayuso, J. A. Álvarez, D. Rubiales and F. A. Macías, Org. Biomol. Chem., 2017, 15, 6500–6510.
65. M. Masi, M. Fernández-Aparicio, R. Zatout, A. Boari, A. Cimmino and A. Evidente, Molecules, 2019, 24, 3479.
66. G. Soriano, A. Siciliano, M. Fernández-Aparicio, A. Cala Peralta, M. Masi, A. Moreno-Robles, M. Guida and A. Cimmino, Toxins, 2022, 14, 559.
67. P. L. Rodriguez and A. Albert, J. Exp. Bot., 2026, 77, 227–230.
68. M. Jamil, Y. Alagoz, J. Y. Wang, G. T. E. Chen, L. Berqdar, N. M. Kharbatia, J. C. Moreno, H. N. J. Kuijer and S. Al-Babili, Plant J., 2024, 117, 1305–1316.
69. Y. Ding, A. Huffaker, T. G. Köllner, P. Weckwerth, C. A. Robert, J. L. Spencer, A. E. Lipka and E. A. Schmelz, Plant Physiol., 2017, 175, 1455–1468.
70. M. Rey, M. S. Kruse, R. N. Magrini-Huamán, J. Gómez, M. J. Simirgiotis, A. Tapia, G. E. Feresin and H. Coirini, Metabolites, 2021, 11, 579.
71. D. M. Cárdenas, A. Cala, F. J. R. Mejías, J. G. Zorrilla and F. A. Macías, in Handbook of Dietary Phytochemicals, ed. J. Xiao, S. Sarker and Y. Asakawa, Springer, Singapore, 2020,  DOI:10.1007/978-981-13-1745-3_16-1.
72. R. W. Doskotch and M. S. Flom, Tetrahedron, 1972, 28, 4711–4717.
73. Y. Sugimoto, N. E. Ahmed, N. Yasuda and S. Inanaga, Weed Sci., 2002, 50, 658–661.
74. R. El-Kassas, Z. K. El-Din, M. H. Beale, J. L. Ward and R. N. Strange, Weed Res., 2005, 45, 212–219.
75. A. I. Hsiao, A. D. Worsham and D. E. Moreland, Ann. Bot., 1988, 62, 17–24.
76. W. G. Edwards, R. P. Hiron and A. I. Mallet, Z. Pflanzenphysiol., 1976, 80, 105–111.
77. Y. Takeuchi, Y. Omigawa, M. Ogasawara, K. Yoneyama, M. Konnai and A. D. Worsham, Plant Growth Regul., 1995, 16, 153–160.
78. K. Yoneyama, Y. Takeuchi, M. Ogasawara, M. Konnai, Y. Sugimoto and T. Sassa, J. Agric. Food Chem., 1998, 46, 1583–1586.
79. M. Fernández-Aparicio, A. Andolfi, A. Cimmino, D. Rubiales and A. Evidente, J. Agric. Food Chem., 2008, 56, 8343–8347.
80. A. Evidente, A. Cimmino, M. Fernández-Aparicio, D. Rubiales, A. Andolfi and D. Melck, Pest Manag. Sci., 2011, 67, 1015–1022.
81. G. Leubner-Metzger, in Brassinosteroids: bioactivity and crop productivity, ed. S. Hayat and A. Ahmad, Springer, Dordrecht, 2003, pp. 119–128.
82. J. K. Rugutt, K. J. Rugutt and D. K. Berner, J. Nat. Prod., 2001, 64, 1434–1438.
83. M. S. Rajab, J. K. Rugutt, F. R. Fronczek and N. H. Fischer, J. Nat. Prod., 1997, 60, 822–825.
84. P. M. Dewick, Medicinal natural products: a biosynthetic approach, John Wiley & Sons, 2nd edn, 2002.
85. M. Chang, D. H. Netzly, L. G. Butler and D. G. Lynn, J. Am. Chem. Soc., 1986, 108, 7858–7860.
86. A. Cala Peralta, G. Soriano, J. G. Zorrilla, M. Masi, A. Cimmino and M. Fernández-Aparicio, Molecules, 2022, 27, 7421.
87. M. Fernández-Aparicio, A. Cimmino, A. Evidente and D. Rubiales, J. Agric. Food Chem., 2013, 61, 9797–9803.
88. T. Lurthy, S. Perot, F. Gerin-Eveillard, M. Rey, F. Wisniewski-Dyé, J. Vacheron and C. Prigent-Combaret, Microb. Biotechnol., 2023, 16, 2313–2325.
89. A. Moreno-Robles, A. Cala Peralta, J. G. Zorrilla, G. Soriano, M. Masi, S. Vilariño-Rodríguez, A. Cimmino and M. Fernández-Aparicio, Plants, 2023, 12, 697.
90. A. Moreno-Robles, A. Cala Peralta, G. Soriano, J. G. Zorrilla, M. Masi, S. Vilariño-Rodríguez, A. Cimmino and M. Fernández-Aparicio, Agriculture, 2022, 12, 1746.
91. A. Moreno-Robles, A. C. Peralta, J. G. Zorrilla, G. Soriano, M. Masi, S. Vilariño-Rodríguez, A. Cimmino and M. Fernández-Aparicio, Plants, 2022, 11, 2846.
92. M. Fernández-Aparicio, M. Masi, A. Cimmino, S. Vilariño and A. Evidente, Plants, 2021, 10, 543.
93. N. Bar Nun and A. M. Mayer, Isr. J. Plant Sci., 2005, 53, 97–101.
94. C. Kehelpannala, G. N. Rathnayake, D. Dissanayake, D. Kanatiwela, N. S. Kumar, N. Adikaram, L. Jayasinghe, H. Araya and Y. Fujimoto, Chem. Pap., 2021, 75, 2233–2235.
95. N. E. Ahmed, Y. Sugimoto, A. G. Babiker, O. E. Mohamed, Y. Ma, S. Inanaga and H. Nakajima, Weed Sci., 2001, 49, 354–358.
96. C. Veronesi, E. Billard, P. Delavault and P. Simier, Pest Manage. Sci., 2025, 81, 720–726.
97. G. Soriano, C. Petrillo, M. Masi, M. Bouafiane, A. Khelil, A. Tuzi, R. Isticato, M. Fernández-Aparicio and A. Cimmino, Toxins, 2022, 14, 311.
98. A. M. Hooper, M. K. Tsanuo, K. Chamberlain, K. Tittcomb, J. Scholes, A. Hassanali, Z. R. Khan and J. A. Pickett, Phytochemistry, 2010, 71, 904–908.
99. H. Ueda and Y. Sugimoto, Biosci. Biotechnol. Biochem., 2010, 74, 1662–1667.
100. A. Scurria, M. Sciortino, L. Albanese, D. Nuzzo, F. Zabini, F. Meneguzzo, R. Alduina, A. Presentato, M. Pagliaro, G. Avellone and R. Ciriminna, ChemistryOpen, 2021, 10, 1055–1058.
101. A. Evidente, A. Cimmino, M. Fernandez-Aparicio, A. Andolfi, D. Rubiales and A. Motta, J. Agric. Food Chem., 2010, 58, 2902–2907.
102. A. Evidente, M. Fernández-Aparicio, A. Cimmino, D. Rubiales, A. Andolfi and A. Motta, Tetrahedron Lett., 2009, 50, 6955–6958.
103. H. S. Nzioki, F. Oyosi, C. E. Morris, E. Kaya, A. L. Pilgeram, C. S. Baker and D. C. Sands, Front. Plant Sci., 2016, 7, 1121.
104. M. Fernández-Aparicio, A. Bernard, L. Falchetto, P. Marget, B. Chauvel, C. Steinberg, C. E. Morris, S. Gibot-Leclerc, A. Boari, M. Vurro, D. A. Bohan, D. C. Sands and X. Reboud, Front. Plant Sci., 2017, 8, 842.
105. K. Tsuzuki, T. Suzuki, M. Kuruma, K. Nishiyama, K. I. Hayashi, S. Hagihara and Y. Seto, Plant Cell Physiol., 2024, 65, 1377–1387.
106. M. Vurro, A. Boari, A. L. Pilgeram and D. C. Sands, Biol. Control, 2006, 36, 258–265.
107. S. Gibot-Leclerc, F. Dessaint, M. Connault and R. Perronne, J. Plant Dis. Prot., 2024, 131, 91–99.
108. T. Dadon, N. B. Nun and A. M. Mayer, Isr. J. Plant Sci., 2004, 52, 83–86.
109. W. He, Y. Li, W. Luo, J. Zhou, S. Zhao and J. Xu, AMB Express, 2022, 12, 52.
110. A. de Zélicourt, P. Letousey, S. Thoiron, C. Campion, P. Simoneau, K. Elmorjani, D. Marion, P. Simier and P. Delavault, Planta, 2007, 226, 591–600.
111. H. Thomas, A. Heller, J. Sauerborn and D. Müller-Stöver, Ann. Bot., 1999, 83, 453–458.
112. M. Aytbeke, B. Sen and S. Ökten, J. Basic Microbiol., 2014, 54, S93–S101.
113. X. Hu, X. Cao, Q. Zhao, X. Zeng, Y. Wei, Z. Yao and S. Zhao, Plants, 2025, 14, 1591.
114. C. Rial, E. Gómez, R. M. Varela, J. M. Molinillo and F. A. Macías, J. Agric. Food Chem., 2018, 66, 4638–4644.
115. H. Lee, M. J. Seo and S. Jang, Biotechnol. Bioprocess Eng., 2024, 29, 1–12.
116. K. Harada, Y. Kurono, S. Nagasawa, T. Oda, Y. Nasu, T. Wakabayashi, Y. Sugimoto, H. Matsuura, S. Muranaka, K. Hirata and A. Okazawa, J. Pestic. Sci., 2017, 42, 166–171.
117. T. Wakabayashi, B. Joseph, S. Yasumoto, T. Akashi, T. Aoki, K. Harada, S. Muranaka, T. Bamba, E. Fukusaki, Y. Takeuchi, K. Yoneyama, T. Muranaka, Y. Sugimoto and A. Okazawa, J. Exp. Bot., 2015, 66, 3085–3097.
118. N. A. Garas, C. M. Karssen and J. Bruinsma, Z. Pflanzenphysiol., 1974, 71, 108–114.
119. A. G. T. Babiker, L. G. Butler, G. Ejeta and W. R. Woodson, Physiol. Plant., 1993, 89, 21–26.
120. D. C. Logan and G. R. Stewart, Plant Physiol., 1991, 97, 1435–1438.
121. K. Uematsu, M. Nakajima, I. Yamaguchi, K. Yoneyama and Y. Fukui, J. Plant Growth Regul., 2007, 26, 245–254.
122. A. Evidente, M. Fernández-Aparicio, A. Andolfi, D. Rubiales and A. Motta, Phytochemistry, 2007, 68, 2487–2492.
123. B. Bouizgarne, H. El-Maarouf-Bouteau, K. Madiona, B. Biligui, M. Monestiez, A. M. Pennarun, Z. Amiar, J. P. Rona, Y. Ouhdouch, I. El Hadrami and F. Bouteau, Mol. Plant-Microbe Interact., 2006, 19, 550–556.
124. M. C. Zonno, M. Vurro, R. Capasso, A. Evidente, A. Cutignano, J. Sauerborn and H. Thomas, Phytotoxic metabolites produced by Fusarium nygamai from Striga hermonthica, in Proceedings of the IX International Symposium on Biological Control of Weeds, Stellenbosch, South Africa, 1996, pp. 223–226.
125. Y. Yun, X. Zhou, S. Yang, Y. Wen, H. You, Y. Zheng, J. Norvienyeku, W. B. Shim and Z. Wang, Curr. Genet., 2019, 65, 773–783.
126. M. E. Amer, M. I. Abou-Shoer, M. S. Abdel-Kader, A. El-Shaibany and N. A. Abdel-Salam, J. Braz. Chem. Soc., 2004, 15, 262–266.
127. T. Wakabayashi, B. Joseph, S. Yasumoto, T. Akashi, T. Aoki, K. Harada, S. Muranaka, T. Bamba, E. Fukusaki, Y. Takeuchi, K. Yoneyama, T. Muranaka, Y. Sugimoto and A. Okazawa, J. Exp. Bot., 2015, 66, 3085–3097.
128. L. D. Hohenschutz, E. A. Bell, P. J. Jewess, D. P. Leworthy, R. J. Pryce, E. Arnold and J. Clardy, Phytochemistry, 1981, 20, 811–814.
129. R. Nogami, M. Nagata, R. Imada, K. Kai, T. Kawaguchi and S. Tani, J. Pestic. Sci., 2024, 49, 22–30.
130. M. Masi, J. G. Zorrilla and S. Meyer, Toxins, 2022, 14, 588.
131. A. Cimmino, M. Fernández-Aparicio, F. Avolio, K. Yoneyama, D. Rubiales and A. Evidente, Phytochemistry, 2015, 109, 57–65.
132. B. Auger, J. B. Pouvreau, K. Pouponneau, K. Yoneyama, G. Montiel, B. Le Bizec, K. Yoneyama, P. Delavault, R. Delourme and P. Simier, Mol. Plant-Microbe Interact., 2012, 25, 993–1004.
133. H. Miura, R. Ochi, H. Nishiwaki, S. Yamauchi, X. Xie, H. Nakamura, K. Yoneyama and K. Yoneyama, Plants, 2022, 11, 606.
134. L. Du, X. Li, Y. Ding, D. Ma, C. Yu and L. Duan, J. Agric. Food Chem., 2024, 72, 8944–8954.
135. M. J. Van Hezewijk and J. A. C. Verkleij, Weed Res., 1996, 36, 395–404.
136. A. de Zélicourt, G. Montiel, J. B. Pouvreau, S. Thoiron, S. Delgrange, P. Simier and P. Delavault, Planta, 2009, 230, 1047–1055.
137. S. Castaldi, J. G. Zorrilla, C. Petrillo, M. T. Russo, P. Ambrosino, M. Masi, A. Cimmino and R. Isticato, J. Fungi, 2023, 9, 326.
138. B. Zwanenburg, A. S. Mwakaboko and C. Kannan, Pest Manage. Sci., 2016, 72, 2016–2025.
139. K. Yoneyama, M. Ogasawara, Y. Takeuchi, M. Konnai, Y. Sugimoto, H. Seto and S. Yoshida, Biosci. Biotechnol. Biochem., 1998, 62, 1448–1450.
140. G. Soriano, D. Arnodo, M. Masi, M. Fernández-Aparicio, B. B. Landa, C. Olivares-García, A. Cimmino and C. Prandi, J. Agric. Food Chem., 2024, 72, 4737–4746.
141. E. G. Padilla Suarez, J. G. Zorrilla, M. Spampinato, T. Pannullo, F. Esposito, M. Fernández-Aparicio, G. Libralato, A. Siciliano, M. Masi and A. Cimmino, Toxins, 2025, 17, 169.
142. A. Okazawa, H. Samejima, S. Kitani, Y. Sugimoto and D. Ohta, J. Pestic. Sci., 2021, 46, 242–247.
143. M. I. Daws, H. W. Pritchard and J. Van Staden, J. S. Afr. Bot., 2008, 74, 116–120.
144. M. Fernández-Aparicio, M. Masi, A. Cimmino and A. Evidente, Plants, 2021, 10, 746.
145. M. Chang and D. G. Lynn, J. Chem. Ecol., 1986, 12, 561–579.
146. D. H. Netzly, J. L. Riopel, G. Ejeta and L. G. Butler, Weed Sci., 1988, 36, 441–446.
147. M. Fernández-Aparicio, M. Masi, L. Maddau, A. Cimmino, M. Evidente, D. Rubiales and A. Evidente, J. Agric. Food Chem., 2016, 64, 5188–5196.
148. S. Saaidpour and A. Jahannamaie, J. Anal. Chem., 2019, 74, 756–763.
149. S. L. Vail, O. D. Dailey, E. J. Blanchard, A. B. Pepperman and J. L. Riopel, J. Plant Growth Regul., 1990, 9, 77–83.
150. S. Nadal, C. I. González-Verdejo, J. R. Guzmán, M. J. Suso and B. Román, Afr. J. Biotechnol., 2009, 8, 3027–3030.
151. M. Onitsuka, T. Wakabayashi, T. Ogawa, Y. Sugimoto, D. Ohta and A. Okazawa, J. Pestic. Sci., 2025, 50, 40–46.
152. C. Vraka, L. Nics, K. H. Wagner, M. Hacker, W. Wadsak and M. Mitterhauser, Nucl. Med. Biol., 2017, 50, 1–10.
153. H. Hlaili, J. G. Zorrilla, M. M. Salvatore, M. Abassi, M. T. Russo, M. I. Martínez-González, M. DellaGreca, A. Cimmino, F. A. Macías, A. Andolfi, R. M. Varela and M. Masi, Phytochem. Anal., 2025, 36, 2374–2384.
154. F. J. Soto-Cruz, J. G. Zorrilla, C. Rial, R. M. Varela, J. M. Molinillo, J. M. Igartuburu and F. A. Macías, Agronomy, 2021, 11, 2174.
155. V. Lailheugue, I. Merlin, S. Boutet, F. Perreau, J. B. Pouvreau, S. Delgrange, P. H. Ducrot, B. Cottyn-Boitte, G. Mouille and V. Lauvergeat, Phytochemistry, 2023, 215, 113837.

---