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Parasitic Plants—Potential Vectors of Phytopathogens

Department of Biochemistry, Faculty of Biology, Sofia University “St. Kliment Ohridski”, 8 Dragan Tsankov blvd., 1164 Sofia, Bulgaria
Author to whom correspondence should be addressed.
Pathogens 2024, 13(6), 484;
Submission received: 10 May 2024 / Revised: 2 June 2024 / Accepted: 5 June 2024 / Published: 7 June 2024


Parasitic plants represent a peculiar group of semi- or fully heterotrophic plants, possessing the ability to extract water, minerals, and organic compounds from other plants. All parasitic plants, either root or stem, hemi- or holoparasitic, establish a vascular connection with their host plants through a highly specialized organ called haustoria. Apart from being the organ responsible for nutrient extraction, the haustorial connection is also a highway for various macromolecules, including DNA, proteins, and, apparently, phytopathogens. At least some parasitic plants are considered significant agricultural pests, contributing to enormous yield losses worldwide. Their negative effect is mainly direct, by the exhaustion of host plant fitness and decreasing growth and seed/fruit formation. However, they may pose an additional threat to agriculture by promoting the trans-species dispersion of various pathogens. The current review aims to summarize the available information and to raise awareness of this less-explored problem. We further explore the suitability of certain phytopathogens to serve as specific and efficient methods of control of parasitic plants, as well as methods for control of the phytopathogens.

1. Parasitism in Plants

1.1. Variety of Parasitic Plants

Parasitic flowering plants are a highly specialized group of vascular plants, which switch to a partially or fully heterotrophic lifestyle. Depending on the degree of loss of photosynthetic ability, they are commonly divided into hemiparasites (photosynthetic or partially photosynthetic) and holoparasites (non-photosynthetic), but the terms facultative and obligate are also used [1], and parasitic plants are also classified as obligate (cannot complete lifecycle without a host) and facultative. The other common classification is based on the site of vascular connection with the host plant, giving either root or stem parasitic plants. Of the nearly 5000 known species [2,3], the holoparasites account for about 10%, while the distribution between root and stem parasites is more even, 60 to 40%, respectively [1]. There are a total of 12 parasitic plant clades, corresponding to 12 independent evolutional events. However, over 90% of all parasitic plants fall into two nearly equal in species number (with over 2000 species) clades—the root hemi- and holoparasites of the family Orobanchaceae and the stem hemiparasites of the order Santalales [3]. The third in species number is the genus Cuscuta, family Convolvulaceae, with a little over 200 species. The other clades contain a few to several dozen species, often rare and highly specialized. Some exotic examples include the Rafflesiaceae, characterized by enormously large flowers, but a highly reduced vegetative part [4]. However, this list is far from exhaustive, and new species are described every year, mainly because root hemiparasites may remain unnoticed, or because some species are very difficult to distinguish morphologically [5].
Parasitic plants also differ significantly in their host preference. While some are highly specialized in a single, or several, host plant species, others are generalists and infect tens, or hundreds, of host plants from different families. Some notable examples of single-host specialists are Orobanche cumana Wallr., specifically infecting sunflower (Helianthus annuus L.) due to specific requirements of germination stimulants of the strigolactone group [6]. However, a switch in strigolactone responsiveness may expand the host range of O. cumana to other plant species [7]. The juniper dwarf mistletoe (Arceuthobium oxycedri (DC.) M. Bieb) is highly specific to several juniper species but is occasionally found on other tree species [8]. Therefore, it is highly unlikely that a parasitic plant is restricted to a single host species, but still, many are restricted to several closely related hosts. Well-known generalists are the members of the Cuscuta genus, as some were reported to have 200 (Cuscuta europaea L., Cuscuta campestris Yunck.) to nearly 350 (Cuscuta epithymum L.) host species in a single study [9]. At the individual level, Orobanchaceae and mistletoes tend to expand to a single or few host plant individuals; a single Cuscuta individual may simultaneously spread and infect multiple hosts. From a practical point of view, parasitic plants that are generalists and infect multiple hosts simultaneously would be more efficient pathogen vectors than host specialists, and such species that infect a single host (e.g., mistletoes).
Some parasitic plants are dangerous pathogens on their own. Probably the most devastating are the members of the Striga (witchweeds) genus (Orobanchaceae), namely Striga hermonthica (Delile) Benth., Striga asiatica (L.) Kuntze, and Striga gesnerioides (Willd.) Vatke. They cause nearly entire yield loss in various cultures, such as rice, sorghum, cowpea, maize, finger, and pearl millet in over 40 countries, mainly in Sub-Saharan Africa, causing over USD 10 billion in economic losses [10,11]. Broomrapes (Orobanche and Phelypanche genera) of the same family are probably second in terms of agricultural impact. At least nine species were reported as dangerous weeds in Europe, Asia, and North Africa [12]. They were reported to cause yield losses, anywhere from 10 to 100% in various legumes, carrots, tomatoes, etc. Out of over 200 Cuscuta species, relatively few are also considered important pests, with C. campestris being the top villain, causing significant losses in agriculture worldwide [13]. Some other parasitic plant species are also dangerous pathogens, but to a much lesser degree than the above-mentioned [13]. For the current overview, we focused mainly on representatives of Orobanchaceae and Cuscuta spp. (root and stem parasites, respectively), due to their economic impact.
The primary cause of the negative impact of parasitic plants on their hosts is the direct extraction of nutrients (both minerals and organic compounds) and water, thus decreasing the biomass and seed production of their hosts [14]. However, their impact may be much broader, including the modulation of below-ground communities [15], inhibition of host photosynthesis [16,17,18], etc. The most striking effect is the “bewitching” effect of Striga spp. on their hosts, consisting of wilting and chlorosis of the host in the very early stages of Striga infection [19]. This effect is not entirely understood, suggested to be caused by excessive amounts of exuded abscisic acid [20], and the disruption of other hormones [19], but is certainly far more devastating than the exhaustion of nutrients. Finally, parasitic plants may also represent a significant vector of other pathogens, such as viruses, phytoplasma, bacteria, and fungi, which they may acquire and transfer from one host to another, facilitated by the vascular connection.

1.2. Haustorium Properties

Regardless of their taxonomic position, host specificity, or evolutionary history, all parasitic plants require a vascular connection with their hosts, called the haustorium. Haustoria represent multicellular invasive organs, able to attach and penetrate host tissues, sometimes overcoming tissue incompatibilities, as many parasitic plants are able to parasitize non-related, taxonomically distant host species [21]. Haustoria of root parasites are divided into two types, lateral haustoria and terminal haustoria, which are specifically formed by the apical root meristem [22]. While lateral haustoria do not interfere with root tip elongation and allow the formation of multiple haustoria, the terminal haustoria lead to the termination of root growth [23]. Some plants can form both lateral and terminal haustoria, like it was shown in Phelipanche ramosa (L.) Pomel (=Orobanche ramosa L.) (Orobanchaceae), while others are capable of only lateral haustoria formation like Phtheirospermum japonicum (Thunb.) Kanitz (Orobanchaceae) [24]. To some extent, the presence of terminal haustoria is associated with obligate parasites, while lateral haustoria are present in hemiparasites and are regarded as evolutionarily older than terminal haustoria [25,26]. In the case of obligate parasites, like Striga, the terminal haustorium is formed first, immediately after germination, followed by the growth of adventitious roots and lateral haustoria [27]. In an older study [28], but still in use [27], terminal haustoria are also denoted as primary, while lateral haustoria are denoted as secondary. Haustoria of stem parasites are considered distinct from root haustoria and although similar in function, differ in molecular mechanisms of formation, like in Cuscuta spp. [26]. Other stem parasites may possess different haustoria in terms of structure and formation [1].
Roughly three distinct stages of haustorium formation can be defined—initiation (or prehaustorium), penetration into host tissues, and vascular connection establishment [21]. Before initiation, both root and stem parasites need to locate/recognize host tissue. In root parasites, this occurs through chemical signaling—the recognition of strigolactones, released in host root exudates to attract symbiotic mycorrhizal fungi [29,30,31]. Going further, obligate root parasites require strigolactone detection to germinate only in the presence of a suitable host [31], and are also definitive for the host specificity of certain root parasites [6,7]. Unlike them, members of the genus Cuscuta were not proven to require chemical stimuli for germination and employ light [32,33] and probably chemical [34] stimuli in host recognition. The initiation of the root haustorium (in Orobanchaceae) requires specific chemical compounds, released by the host, collectively called haustorium-inducing factors (HIFs), which were proved to be quite diverse [35,36]. Unlike them, haustorium initiation in Cuscuta further relies on specific light quality in combination with tactile stimuli [37]. The stage of penetration, regardless of the parasite taxonomy, involves a combination of hydrolytic and cell-wall-modifying enzymes [21,26,37]. Finally, the vascular connection is established, which is primarily xylem–xylem (xylem bridge), observed in all parasitic plants [21]. A linkage to the phloem of the host, however, seems to be less common and is observed in stem parasites Cuscuta spp., but also root parasites Orobanche spp., while not being presented in Striga spp. [38].
The main function of haustoria per se is the transfer of water, mineral nutrients, and photosynthates from the host to the parasite. It is often compared to grafting [39] and raises the problem of the complicated interaction between two genomes, connected via a non-interrupted connection [40]. In this context, the haustorium was reported on numerous occasions as a bidirectional transport highway for numerous different molecules, far more complicated than a simple source of nutrients for the parasite. In Cuscuta spp., this involves an open xylem–xylem connection, membrane-mediated phloem transport, and plasmodesmata connections between contacting parasite and host cell walls [41]. This allows extensive transport of macromolecules, including DNA [42], mRNAs [43,44], and proteins [45]. Similar results were reported and summarized for Orobanchaceae [46,47]. Concerning phytopathogens, there is an important question about the haustorial connection—whether it is a putative passage for such passengers, which it is [41,47], or if it could serve as a selective barrier, limiting their distribution. Although there is evidence that this connection may be selective at least for some chemical entities [48], overall, it is a putative passage for different phytopathogens.

2. Phytopathogens and Parasitic Plants

2.1. Major Phytopathogens

2.1.1. Viruses

The group of plant viruses is a constantly growing group of non-cellular infectious agents, containing DNA or RNA and a protein coat, that cause multiple diseases in plants, with visual expression as leaf discoloration, stunted growth, mottling, and necrosis. The estimated economic effect of plant viral diseases reaches USD 30 billion per year in crop yield losses [49]. The list of known plant viruses exceeds thousands [50] and will further expand in the future, but not all are equally devastating and agriculturally important. Some efforts to classify plant viruses resulted in surveys, like the “Top 10 plant viruses in molecular plant pathology” [51] and “Top 10 economically important plant viruses” [52], based on their importance as agricultural pests and molecular study objects. While some are comparatively restricted in the host range, others are characterized by an extremely wide host range and wide range of vectors—for Cucumber Mosaic Virus (CMV), Bromoviridae, over 1200 host plant species from over 100 families, and 80 aphid species from 33 genera, were reported [51].
Therefore, it is not surprising that viruses were reported in parasitic plants. Historically, Cuscuta spp. was reported to be infected and transmit over 50 different viruses [53]. More recent reports are comparatively scarce, but confirm dodders as usual hosts for a variety of plant viruses. Of the six enlisted viruses (Table 1), one is a DNA virus of the Geminiviridae family (TYLCV) and all others are positive-strand RNA viruses from four different families. Reports on viruses in Orobanchaceae are even more scarce, although some of the viruses, found in Cuscuta [54,55], were also proved to be acquired by Phelipanche aegyptiaca (Pers.) Pomel (=Orobanche aegyptiaca Pers.) from host plants [56]. Overall, the recent literature lacks substantial studies of virus distribution among parasitic plants, also shown by the fact that some viruses were discovered accidentally during transcriptomics analyses [57].

2.1.2. Phytoplasma

Phytoplasmas (Candidatus Phytoplasma: Mollicutes) are obligate intracellular bacteria that lack cell walls [63] and cause a variety of diseases [64]. They are phloem-mobile, e.g., they move within plants by the phloem traffic [65]. Common disease symptoms include little or yellow leaves, phyllody, witches’ brooms, etc., and could cause between 30% and 100% yield loss, depending on the crop plant [66]. Being phloem-mobile, it is not surprising that dodders are efficient reservoirs and vectors of several phytoplasma [67,68]. Phytoplasmas were also reported from several root parasites—Ph. ramosa as a host for tomato stolbur disease [69] and Orobanche spp. as a host for tomato big bud [70] and several Orobanche-specific phytoplasmas [71].

2.1.3. Bacteria

Phytopathogenic bacteria are among the most devastating disease-causing pests, contributing to enormous yield losses in crop plants [72]. Unlike viruses and phytoplasma, they are not fully dependent on hosts to survive, and more often are soilborne pathogens, infecting plants through wounds, and moving through the xylem vessels. Several of the most destructible pathovars belong to the Pseudomonas and Xanthomonas genera, as well as notorious pathogens like Ralstonia solanacearum (Smith 1896) Yabuuchi et al., 1995, single-handedly contributing to over USD 1 billion annual losses, and Erwinia amylovora (Burrill 1882) Winslow et al., 1920 [73]. Important pathogens among Actinomycetes include members of the Clavibacter [74] and Curtobacterium [75]. Most of the studies involving phytopathogenic bacteria and parasitic plants are related to methods of control of the parasitic plants [76] and will be further discussed in detail hereinafter.
Interestingly, extracts from the members of the genus Cuscuta were shown to possess strong antibacterial activity against major phytopathogenic bacteria. For example, water extracts of Cuscuta pedicellata Ledeb. were proven in vitro as efficient control agents for fruit lesions, caused by Xanthomonas campestris (Pammel 1895) Dowson 1939 [77]. Similarly, organic solvent extracts of Cuscuta reflexa Roxb. were effective in the inhibition of X. campestris and several human pathogenic bacteria [78]. Ethanol extracts of several Orobanche species were also shown as efficient antibacterial agents against Agrobacterium and Erwinia [79]. Cuscuta spp. are particularly rich in bioactive compounds [80], as well as members of Orobanchaceae [81,82], so their in vitro antibacterial activity is not surprising, and additionally, they are regarded as highly tolerant to phytopathogenic bacteria [79].

2.1.4. Fungi

The fourth major phytopathogenic group consists of fungi. According to some estimates, they account for nearly 80% of yield losses, caused by microbial pathogens [83]. Some of the most damaging species include Magnaporthe oryzae (T.T. Hebert) M.E. Barr, Botrytis cinerea Pers., and several Fusarium species, among others [84]. However, phytopathogenic fungi are a comparatively small portion of many fungi that live in symbiotic relations with plants, such as arbuscular mycorrhizal fungi, which are important and even critical for healthy plants [85], and endophytic fungi, which are also important players in combating other phytopathogens [86]. The relations between plants and fungi extend to the case of mycoheterotrophic plants, which are parasitizing fungi [87]. Both dodders [88,89] and Orobanchaceae [90,91,92] were shown to be rich in endophytic fungal biodiversity, and also several Fusarium spp. isolates showed promising results in terms of pathogenicity against the parasitic plants [93,94]. Phytopathogenic fungi were also the most exploited means of biological control of parasitic plants, which will be further discussed below.

2.2. Symptomatics

Since now, parasitic plants have been reported to be infected by multiple phytopathogens. However, they rarely exhibit visual symptoms of such infections, which is especially true for Cuscuta spp. and some holoparasitic members of Orobanchaceae, due to their simple, leafless morphology [1]. In terms of viral infections, they rarely experience visible phenotypic symptoms or are even reported as symptomless in various experiments [54,58,59]. Although some very early reports claim that Cuscuta spp. can suppress viruses from transmission to other plants [59], there is no recent confirmation of such inactivation and dodders can maintain high viral titer for years [59]. However, there are some reports that when parasitizing virus-infected hosts, some parasitic plants may exhibit a deformed phenotype, as in the case of Ph. aegyptiaca, infecting CMV-infected tobacco plants [95]. In terms of phytoplasma, Marcone [68] successfully employed Cuscuta spp. as vectors of several phytoplasmas, but did not report any visual symptoms in the dodder vector. Unlike them, Orobanche spp. showed clear evidence of phytoplasma infection, such as stem flattening and witches’ broom [71].
Fungal infection on Cuscuta gronovii Willd. was shown to cause observable symptoms, including the discoloration and shriveling of the stem, blighted portion, necrotic lesions, and tip necrosis [96]. Cuscuta pentagona Engelm. Alternaria destruens E.G. Simmons 1998 also causes necrotic spots and blights [97]. In Orobanche spp., different Fusarium isolates caused a variety of symptoms such as the inhibition of seed germination, wilting, and necrotic lesions on the stem and inflorescences [93,94]. Similar symptoms were also observed in Striga spp. as a result of Fusarium infection [98]. The seed germination of Ph. aegyptiaca was also found to be inhibited by Pseudomonas and Bacillus isolates [76]. Overall, symptomatics in parasitic plants seem to be much less pronounced and specific than in other plants, raising the necessity of molecular methods for the identification and screening of phytopathogens.

2.3. Detection Methods

General detection methods in plant pathology were recently summarized by Khakimov et al. [99] and could be roughly divided into traditional (e.g., visual inspection, microscopy, cultivation methods) and molecular (or modern, e.g., immunological, genetic, and mass-spectrometric). Visual observation highly relies on detectable symptoms, such as characteristic necrosis, chlorosis, etc., while microscopic methods may result in false identification, and also depend on the availability of certain pathogenic structures—for example, fungal spores or micellium [99]. Moreover, they are not always sufficient to provide the species identification of the pathogen and are somehow difficult to apply on parasitic plants, such as dodders and holoparasitic Orobancheacea, for the above-mentioned reasons. The most commonly employed modern methods for detection could be divided into immunological and molecular genetic methods. The immunological methods rely on the specific detection of an antigen of the pathogen by an antibody. Molecular methods employ PCR amplification and further sequencing of specific, taxon-discriminative DNA fragments. Phytopathogen detection in parasitic plants does not require any specific methods, differing from that established in the state-of-the-art. However, there are some peculiarities related to the scarce material of certain plant parts.
Plant virus detection is performed mainly by either commercialized ELISA kits [54] or quantitative RT-PCR with virus-specific primers, which also allow the determination of the viral titer [56,59]. Both approaches, however, allow the identification of particular viruses, selected in advance, which may underestimate the distribution of other viruses. Some more efficient methods, allowing the simultaneous detection of multiple, including unknown, viruses, as next generation sequencing (NGS)-based methods, loop-mediated isothermal amplification (LAMP), etc., were recently summarized by Mehetre et al. [100]. Being the most diverse group of phytopathogens, systematic viral detection would require a greater variety of methods. The detection of phytoplasmas commonly involves PCR amplification with phytoplasma-specific primers, of the DNA region, extending from the 5′ end of the 16S rRNA gene to the 5′ region of the 23S rRNA gene, and its amplicon may be either further analyzed by a Restriction Fragment Length Polymorphism (RFLP) analysis [68] or sequenced [101]. The typically low titers of phytoplasmas require the employment of nested PCR, or preferably quantitative real-time PCR, or LAMP.
The detection of bacterial and fungal pathogens is generally more straightforward than viruses and phytoplasmas. Most of them cause visible symptoms on parasitic plants (see above), are often visible by a common microscope, and could be cultivated on growth media in the absence of a host [102,103]. Moreover, the simultaneous NGS-based identification of thousands of bacterial and fungal taxa in a single sample, based on the sequencing of the variable regions of the 16S rRNA gene (for bacteria) and the nuclear ribosomal internal transcribed spacer (ITS) region (for fungi), is already a routine method [90].

3. Putative Routes of Transmission

3.1. Plant-to-Plant Transmission

As already discussed, the haustorial connection between parasitic plants and their hosts represents a hotspot of bi-directional macromolecular trafficking [46]. Along with interspecific plasmodesmata between the host and parasite phloem [56], it is also the major route by which parasitic plants could infect their hosts with different phytopathogens, especially viruses and phytoplasma [55,67,104]. This was not confirmed, but also might be predicted for bacteria, either actively or passively moving through plant vascular tissues [105]. The penetration nature of haustorium establishment is also important for phytopathogens, ensuring an efficient invasion route, and overcoming the plant cuticle. However, the main purpose of the haustorium is to extract molecules from the host, suggesting that parasitic plants may easily acquire phytopathogens from their hosts (Figure 1), and further distribute them to other hosts. Especially for dodders, their role in transferring a variety of signaling molecules between plants was already established [106]. More importantly, their generalism and simultaneous infection of multiple hosts from different species and families [9] also ensure the efficient spreading of pathogens between species.
However, the haustorial connection does not mean that all possible endophytes (incl. pathogens) would be efficiently distributed between the parasitic plant and the host. For example, the overlapping of endophytic fungi in C. reflexa and different hosts varied between 25% and 37% [88]. Parasitism of C. campestris affected the endophytic microbiome of H. annuus, but also with poor overlapping between the parasitic plant and the host plant [107]. Although different viruses can be acquired from the host, as in the case of Ph. ramosa, not all can successfully replicate in the parasite, and therefore not all can be further transmitted to other hosts [56]. There could be even specificity of virus acquisition and transmission among closely related parasitic plants—when a single virus (GLRaV-7) was detected in three different Cuscuta species, but only two were shown as vectors, and it was host-specific, e.g., one Cuscuta species may transfer the virus to one host, but not to another [59]. A similar specificity of phytoplasma transmission by Cuscuta spp. was shown by Marcone [68], where transmission rates differed greatly depending on both the particular phytoplasma and the dodder species, all of these suggesting that parasitic plants may be efficient, but very specific, phytopathogen vectors.

3.2. Arthropod-to-Plant Transmission

Hemipteran insects (Hemiptera: Linnaeus, 1758), topped by aphids, whiteflies, and leafhoppers, are considered the most important viral vectors, accounting for over 50% of the transmissions of known plant viruses [108]. Other arthropods such as mites, most notably eriophyid mites (Eriophyidae: Nalepa, 1898), are also known vectors of viral diseases [109,110] and nematodes are also significant vectors [111]. Similarly, hemipterans are also the most important vectors of phytoplasma [112,113]. Unlike these two groups, phytopathogenic bacteria and fungi are less commonly but not unlikely to be transmitted by arthropods [114,115,116].
As much as any other plant, parasitic plants are also associated with multiple arthropods that either feed or parasitize on them (Table 2). As clearly shown, both dodders and Orobanchaceae are subjected to feeding and parasitism by different groups of arthropods, potential vectors of phytopathogens. Moreover, it was also proven that molecules, such as mRNAs, could be transferred from the arthropods to the host plant [117]. Although it seems that the arthropod–parasitic plant–host plant route of phytopathogen transfer is not significant, as the insects may directly feed and infect the host plants, it must be noted that some arthropods are more or less specific to parasitic plants [118,119,120], or might not feed equally on the multiple hosts of the parasitic plant, or feed preferentially on the parasitic plant [121] as in the case of Metcalfa pruinose (Say, 1830), which fed exclusively on C. campestris when presented, but also attacked the host plant in the absence of the parasite. This will at least improve the chance of a phytopathogen to infect multiple plant species.
Although there is no experimental evidence of parasitic plants, serving as mediators of phytopathogens from arthropods to other plants, this potential route of transmission (Figure 1) seems plausible and requires more studies to assess the impact of both dodders and root parasites.

3.3. Seed Transmission

Most seeds of parasitic plants are characterized by long persistence in soil, where they can stay dormant for decades, waiting for the proper host to appear. This is mostly true for Orobanchaceae [132], which in combination with a large number of seeds, produced by a single plant, up to 200,000 in Ph. ramosa [133], gives a potentially enormous, long-term reservoir for various pathogens. Cuscuta spp. are also prominent seed producers, and although they do not require specific chemical compounds for germination, they are characterized by soil longevity and continuous germination over decades [134]. Such an enormous and persisting seed bank also represents a potential reservoir of numerous seed-borne pathogens from virtually every group [135], including fungi, bacteria, and viruses. Phytoplasmas are also frequently found in seeds of infected parents; the seed transmission of phytoplasmas is considered unlikely [112].
Besides some scarce reports [54], some of which are quite old [136], the potential of parasitic plant seeds to serve as a reservoir of plant viruses, transmitted to generations, is highly unexplored. The bacterial and fungal microbiota of root parasitic plants are much better studied and were shown to contain opportunistic, or obligate, phytopathogens as in Ph. ramosa [103,137] and Cistanche phelypaea (L.) Cout. [138]. Such pathogens are mostly soilborne and host plants are in contact with them anyway. However, the role of parasitic plants may be in the facilitation of pathogen penetration, simultaneously with the haustorium formation.

4. Phytopathogens as Biocontrol Agents of Parasitic Plants

Control of economically important parasitic plants is among the important aspects of contemporary agricultural practice, often complicated by their similarities to their hosts [12,13]. Being plants of their own, there is also a possibility that chemical control agents will affect their hosts equally, or even to a greater extent than the target parasites [139]. To overcome these limitations, several phytopathogens were studied and successfully applied as biocontrol agents, assuming they are highly specific to the parasite and affect neither host crop plants nor other plant species in proximity.
Both bacteria and fungi were extensively studied as potential biocontrol agents, and mostly isolated from natural sources, e.g., parasitic plants with visual symptoms of a disease. In Cuscuta spp., several species were found to be effective. One promising and already patented bioherbicide, specifically for dodder control, is A. destruens Strain 059 [97,140]. It was shown to affect a variety of dodder species, but it is not infecting other plant species, thus being a promising biocontrol agent. Some other promising results were shown for Fusarium incarnatum (Desm.) Sacc., Alternaria dianthicola Neerg., and Curvularia pallescens Boedijn. isolates from C. gronovii [96]. Of the bacterial pathogens, several Bacillus species were also shown to inhibit Cuscuta seed germination [141].
The inhibitory effect of bacterial isolates on Orobanchaceae members seemed to be much more extensive. These include the inhibition of radical elongation in Ph. aegyptiaca and Orobanche cernua Loefl. by two Pseudomonas and two Bacillus species [142], and in Orobanche crenata Forssk. and Orobanche foetida Poir. by two Pseudomonas species [143]. Besides their inhibitory activity on the parasitic plants, they also showed a positive effect on the growth of the host plants. The isolation of particular pathogenic strains, however, involves laborious screening of hundreds of strains. However, the most promising results for control of root parasitic plants were acquired with fungal pathogens of the Alternaria and Fusarium genera in both Orobanche spp. and Striga spp. The most exploited is probably Fusarium oxysporum, whose highly specific strains are efficient selective control agents for both the germination and growth of Striga [144,145,146,147], but also Orobanche spp. [148]. Other Fusarium species were also shown to be effective and selective pathogens on Orobanche [93,102]. There are hundreds of reports on the isolation and application of such fungal isolates as biocontrol agents on root parasitic plants, underlining the importance of such pathogens for food production security, especially in the poorest regions of Africa. One major advantage of such a bioherbicide is the possibility to propagate without any special equipment, so it is readily available to farmers in distant and poor regions [147]. However, most wild strains of putative phytopathogens are insufficiently virulent to be effective bioherbicides. The inhibitory effect of phytopathogens may be exerted by the direct infection of the plants, but also by the secretion of exometabolites, which specifically inhibit seed germination. Such exometabolites may be specific compounds [149], but also common metabolites such as amino acids, produced in excess [150]. The selection of an efficient hypervirulent biocontrol agent may be enhanced by the identification of such exometabolites and screening of multiple strains for the excessive secretion of such metabolites.

5. Conclusions and Future Perspectives

Although there is certain evidence that parasitic plants may serve as efficient vectors of phytopathogenic viruses, phytoplasmas, bacteria, and fungi, this aspect of their biology is highly understudied. This is because they often lack specific symptoms, but also because most of the efforts in studying parasitic plant biology were devoted to mechanisms of parasitism and methods of control, rather than phytopathogens. The available information is fragmented and does not allow a complete picture of the transmission routes, molecular mechanisms, and overall distribution of phytopathogens among the most economically damaging parasites of the Cuscuta genus and Orobanchaceae family. Besides the additional harm that transmitted phytopathogens add to the negative effect of parasitic plants on crop plants, they also represent a promising means of biological control, and more research on the overall endophytic diversity in both root and stem parasites is needed, to identify potential candidates for bioherbicides. Contemporary molecular methods, such as NGS-based DNA barcoding, offer the necessary tools for the high-throughput characterization of microbial diversity, associated with parasitic plants.

Author Contributions

L.Z. and S.S. conceptualized the review; B.M. prepared the figure and tables. All authors participated in the literature survey and writing of the manuscript. All authors have read and agreed to the published version of the manuscript.


This study is financed by the European Union-NextGenerationEU, through the National Recovery and Resilience Plan of the Republic of Bulgaria, project no. BG-RRP-2.004-0008 and grant KP-06-N31/10 and KP-06-COST/09 of the National Science Fund, Ministry of Education and Science, Bulgaria.

Institutional Review Board Statement

Not applicable.

Informed Consent Statement

Not applicable.

Data Availability Statement

Not applicable.

Conflicts of Interest

The authors declare no conflicts of interest.


  1. Heide-Jørgensen, H. Parasitic Flowering Plants; Brill: Leiden, The Netherlands, 2008. [Google Scholar]
  2. Nickrent, D.L. Parasitic angiosperms: How often and how many? Taxon 2020, 69, 5–27. [Google Scholar] [CrossRef]
  3. Teixeira-Costa, L.; Davis, C.C. Life history, diversity, and distribution in parasitic flowering plants. Plant Physiol. 2021, 187, 32–51. [Google Scholar] [CrossRef]
  4. Wicaksono, A.; Mursidawati, S.; Sukamto, L.A.; Teixeira da Silva, J.A. Rafflesia spp.: Propagation and conservation. Planta 2016, 244, 289–296. [Google Scholar] [CrossRef]
  5. Park, I.; Song, J.-H.; Yang, S.; Kim, W.J.; Choi, G.; Moon, B.C. Cuscuta species identification based on the morphology of reproductive organs and complete chloroplast genome sequences. Int. J. Mol. Sci. 2019, 20, 2726. [Google Scholar] [CrossRef]
  6. Fernández-Aparicio, M.; Yoneyama, K.; Rubiales, D. The role of strigolactones in host specificity of Orobanche and Phelipanche seed germination. Seed Sci. Res. 2011, 21, 55–61. [Google Scholar] [CrossRef]
  7. Dor, E.; Plakhine, D.; Joel, D.M.; Larose, H.; Westwood, J.H.; Smirnov, E.; Ziadna, H.; Hershenhorn, J. A new race of sunflower broomrape (Orobanche cumana) with a wider host range due to changes in seed response to strigolactones. Weed Sci. 2020, 68, 134–142. [Google Scholar] [CrossRef]
  8. Krasylenko, Y.A.; Janošíková, K.; Kukushkin, O.V. Juniper dwarf mistletoe (Arceuthobium oxycedri) in the Crimean Peninsula: Novel insights into its morphology, hosts, and distribution. Botany 2017, 95, 897–911. [Google Scholar] [CrossRef]
  9. Barath, K.; Csiky, J. Host range and host choice of Cuscuta species in Hungary. Acta Bot. Croat. 2012, 71, 215–227. [Google Scholar] [CrossRef]
  10. David, O.G.; Ayangbenro, A.S.; Odhiambo, J.J.; Babalola, O.O. Striga hermonthica: A highly destructive pathogen in maize production. Environ. Chall. 2022, 8, 100590. [Google Scholar] [CrossRef]
  11. Dafaallah, A.B. Biology and physiology of witchweed (Striga spp.): A review. Int. J. Acad. Multidiscip. 2019, 3, 42–51. [Google Scholar]
  12. Parker, C. Observations on the current status of Orobanche and Striga problems worldwide. Pest Manag. Sci. Former. Pestic. Sci. 2009, 65, 453–459. [Google Scholar] [CrossRef]
  13. Parker, C. Parasitic weeds: A world challenge. Weed Sci. 2012, 60, 269–276. [Google Scholar] [CrossRef]
  14. Jeschke, W.D.; Bäumel, P.; Räth, N.; Czygan, F.-C.; Proksch, P. Modelling of the flows and partitioning of carbon and nitrogen in the holoparasite Cuscuta reflexa Roxb. and its host Lupinus albus L. II. Flows between host and parasite and within the parasitized host. J. Exp. Bot. 1994, 45, 801–812. [Google Scholar] [CrossRef]
  15. Bardgett, R.D.; Smith, R.S.; Shiel, R.S.; Peacock, S.; Simkin, J.M.; Quirk, H.; Hobbs, P.J. Parasitic plants indirectly regulate below-ground properties in grassland ecosystems. Nature 2006, 439, 969–972. [Google Scholar] [CrossRef]
  16. Cameron, D.D.; Geniez, J.-M.; Seel, W.E.; Irving, L.J. Suppression of host photosynthesis by the parasitic plant Rhinanthus minor. Ann. Bot. 2008, 101, 573–578. [Google Scholar] [CrossRef]
  17. Gurney, A.L.; Press, M.C.; Ransom, J.K. The parasitic angiosperm Striga hermonthica can reduce photosynthesis of its sorghum and maize hosts in the field. J. Exp. Bot. 1995, 46, 1817–1823. [Google Scholar] [CrossRef]
  18. Shen, H.; Hong, L.; Ye, W.; Cao, H.; Wang, Z. The influence of the holoparasitic plant Cuscuta campestris on the growth and photosynthesis of its host Mikania micrantha. J. Exp. Bot. 2007, 58, 2929–2937. [Google Scholar] [CrossRef]
  19. Runo, S.; Kuria, E.K. Habits of a highly successful cereal killer, Striga. PLoS Pathog. 2018, 14, e1006731. [Google Scholar] [CrossRef]
  20. Fujioka, H.; Samejima, H.; Mizutani, M.; Okamoto, M.; Sugimoto, Y. How does Striga hermonthica Bewitch its hosts? Plant Signal. Behav. 2019, 14, 1605810. [Google Scholar] [CrossRef]
  21. Kokla, A.; Melnyk, C.W. Developing a thief: Haustoria formation in parasitic plants. Dev. Biol. 2018, 442, 53–59. [Google Scholar] [CrossRef]
  22. Yoshida, S.; Cui, S.; Ichihashi, Y.; Shirasu, K. The haustorium, a specialized invasive organ in parasitic plants. Annu. Rev. Plant Biol. 2016, 67, 643–667. [Google Scholar] [CrossRef] [PubMed]
  23. Ishida, J.K.; Wakatake, T.; Yoshida, S.; Takebayashi, Y.; Kasahara, H.; Wafula, E.; de Pamphilis, C.W.; Namba, S.; Shirasu, K. Local Auxin Biosynthesis Mediated by a YUCCA Flavin Monooxygenase Regulates Haustorium Development in the Parasitic Plant Phtheirospermum japonicum. Plant Cell 2016, 28, 1795–1814. [Google Scholar] [CrossRef]
  24. Cui, S.; Wakatake, T.; Hashimoto, K.; Saucet, S.B.; Toyooka, K.; Yoshida, S.; Shirasu, K. Haustorial hairs are specialized root hairs that support parasitism in the facultative parasitic plant Phtheirospermum japonicum. Plant Physiol. 2016, 170, 1492–1503. [Google Scholar] [CrossRef]
  25. Westwood, J.H.; Yoder, J.I.; Timko, M.P.; Depamphilis, C.W. The evolution of parasitism in plants. Trends Plant Sci. 2010, 15, 227–235. [Google Scholar] [CrossRef] [PubMed]
  26. Kirschner, G.K.; Xiao, T.T.; Jamil, M.; Al-Babili, S.; Lube, V.; Blilou, I. A roadmap of haustorium morphogenesis in parasitic plants. J. Exp. Bot. 2023, 74, 7034–7044. [Google Scholar] [CrossRef]
  27. Xiao, T.T.; Kirschner, G.K.; Kountche, B.A.; Jamil, M.; Savina, M.; Lube, V.; Mironova, V.; Al Babili, S.; Blilou, I. A PLETHORA/PIN-FORMED/auxin network mediates prehaustorium formation in the parasitic plant Striga hermonthica. Plant Physiol. 2022, 189, 2281–2297. [Google Scholar] [CrossRef]
  28. Weber, H.C. Evolution of the secondary haustoria to a primary haustorium in the parasitic Scrophulariaceae/Orobanchaceae. Plant Syst. Evol. 1987, 156, 127–131. [Google Scholar] [CrossRef]
  29. Tsuchiya, Y.; Yoshimura, M.; Sato, Y.; Kuwata, K.; Toh, S.; Holbrook-Smith, D.; Zhang, H.; McCourt, P.; Itami, K.; Kinoshita, T. Probing strigolactone receptors in Striga hermonthica with fluorescence. Science 2015, 349, 864–868. [Google Scholar] [CrossRef] [PubMed]
  30. Toh, S.; Holbrook-Smith, D.; Stogios, P.J.; Onopriyenko, O.; Lumba, S.; Tsuchiya, Y.; Savchenko, A.; McCourt, P. Structure-function analysis identifies highly sensitive strigolactone receptors in Striga. Science 2015, 350, 203–207. [Google Scholar] [CrossRef]
  31. Yoneyama, K.; Awad, A.A.; Xie, X.; Yoneyama, K.; Takeuchi, Y. Strigolactones as germination stimulants for root parasitic plants. Plant Cell Physiol. 2010, 51, 1095–1103. [Google Scholar] [CrossRef]
  32. Furuhashi, K.; Iwase, K.; Furuhashi, T. Role of Light and Plant Hormones in Stem Parasitic Plant (Cuscuta and Cassytha) Twining and Haustoria Induction. Photochem. Photobiol. 2021, 97, 1054–1062. [Google Scholar] [CrossRef] [PubMed]
  33. Johnson, B.I.; De Moraes, C.M.; Mescher, M.C. Manipulation of light spectral quality disrupts host location and attachment by parasitic plants in the genus Cuscuta. J. Appl. Ecol. 2016, 53, 794–803. [Google Scholar] [CrossRef]
  34. Runyon, J.B.; Mescher, M.C.; De Moraes, C.M. Volatile chemical cues guide host location and host selection by parasitic plants. Science 2006, 313, 1964–1967. [Google Scholar] [CrossRef] [PubMed]
  35. Goyet, V.; Wada, S.; Cui, S.; Wakatake, T.; Shirasu, K.; Montiel, G.; Simier, P.; Yoshida, S. Haustorium inducing factors for parasitic Orobanchaceae. Front. Plant Sci. 2019, 10, 1056. [Google Scholar] [CrossRef]
  36. Fernández-Aparicio, M.; Masi, M.; Cimmino, A.; Evidente, A. Effects of benzoquinones on radicles of Orobanche and Phelipanche species. Plants 2021, 10, 746. [Google Scholar] [CrossRef] [PubMed]
  37. Jhu, M.-Y.; Sinha, N.R. Cuscuta species: Model organisms for haustorium development in stem holoparasitic plants. Front. Plant Sci. 2022, 13, 1086384. [Google Scholar] [CrossRef]
  38. Teixeira-Costa, L. A living bridge between two enemies: Haustorium structure and evolution across parasitic flowering plants. Braz. J. Bot. 2021, 44, 165–178. [Google Scholar] [CrossRef]
  39. Okayasu, K.; Aoki, K.; Kurotani, K.-I.; Notaguchi, M. Tissue adhesion between distant plant species in parasitism and grafting. Commun. Integr. Biol. 2021, 14, 21–23. [Google Scholar] [CrossRef]
  40. Gaut, B.S.; Miller, A.J.; Seymour, D.K. Living with two genomes: Grafting and its implications for plant genome-to-genome interactions, phenotypic variation, and evolution. Annu. Rev. Genet. 2019, 53, 195–215. [Google Scholar] [CrossRef]
  41. Kim, G.; Westwood, J.H. Macromolecule exchange in Cuscuta–host plant interactions. Curr. Opin. Plant Biol. 2015, 26, 20–25. [Google Scholar] [CrossRef]
  42. Mower, J.P.; Stefanović, S.; Hao, W.; Gummow, J.S.; Jain, K.; Ahmed, D.; Palmer, J.D. Horizontal acquisition of multiple mitochondrial genes from a parasitic plant followed by gene conversion with host mitochondrial genes. BMC Biol. 2010, 8, 150. [Google Scholar] [CrossRef] [PubMed]
  43. Park, S.-Y.; Shimizu, K.; Brown, J.; Aoki, K.; Westwood, J.H. Mobile host mRNAs are translated to protein in the associated parasitic plant Cuscuta campestris. Plants 2021, 11, 93. [Google Scholar] [CrossRef] [PubMed]
  44. Westwood, J.H.; Kim, G. RNA mobility in parasitic plant–host interactions. RNA Biol. 2017, 14, 450–455. [Google Scholar] [CrossRef] [PubMed]
  45. Haupt, S.; Oparka, K.J.; Sauer, N.; Neumann, S. Macromolecular trafficking between Nicotiana tabacum and the holoparasite Cuscuta reflexa. J. Exp. Bot. 2001, 52, 173–177. [Google Scholar] [CrossRef] [PubMed]
  46. Aly, R. Trafficking of molecules between parasitic plants and their hosts. Weed Res. 2013, 53, 231–241. [Google Scholar] [CrossRef]
  47. LeBlanc, M.; Kim, G.; Westwood, J.H. RNA trafficking in parasitic plant systems. Front. Plant Sci. 2012, 3, 203. [Google Scholar] [CrossRef] [PubMed]
  48. Förste, F.; Mantouvalou, I.; Kanngießer, B.; Stosnach, H.; Lachner, L.A.M.; Fischer, K.; Krause, K. Selective mineral transport barriers at Cuscuta-host infection sites. Physiol. Plant. 2020, 168, 934–947. [Google Scholar] [CrossRef] [PubMed]
  49. Sastry, K.S.; Zitter, T.A. Management of virus and viroid diseases of crops in the tropics. In Plant Virus and Viroid Diseases in the Tropics: Volume 2: Epidemiology and Management; Springer: Dordrecht, The Netherlands, 2014; pp. 149–480. [Google Scholar]
  50. Bhat, A.I.; Rao, G.P. Isolation and Diagnosis of Virus Through Indicator Hosts. In Characterization of Plant Viruses; Springer Protocols Handbooks; Humana: New York, NY, USA, 2020; pp. 23–27. [Google Scholar]
  51. Scholthof, K.B.G.; Adkins, S.; Czosnek, H.; Palukaitis, P.; Jacquot, E.; Hohn, T.; Hohn, B.; Saunders, K.; Candresse, T.; Ahlquist, P. Top 10 plant viruses in molecular plant pathology. Mol. Plant Pathol. 2011, 12, 938–954. [Google Scholar] [CrossRef]
  52. Rybicki, E.P. A Top Ten list for economically important plant viruses. Arch. Virol. 2015, 160, 17–20. [Google Scholar] [CrossRef]
  53. Hosford, R.M. Transmission of plant viruses by dodder. Bot. Rev. 1967, 33, 387–406. [Google Scholar] [CrossRef]
  54. Teofanova, D.; Lozanova, Y.; Lambovska, K.; Pachedjieva, K.; Tosheva, A.; Odjakova, M.; Zagorchev, L. Cuscuta spp. populations as potential reservoirs and vectors of four plant viruses. Phytoparasitica 2022, 50, 555–566. [Google Scholar] [CrossRef]
  55. Birschwilks, M.; Haupt, S.; Hofius, D.; Neumann, S. Transfer of phloem-mobile substances from the host plants to the holoparasite Cuscuta sp. J. Exp. Bot. 2006, 57, 911–921. [Google Scholar] [CrossRef] [PubMed]
  56. Gal-On, A.; Naglis, A.; Leibman, D.; Ziadna, H.; Kathiravan, K.; Papayiannis, L.; Holdengreber, V.; Guenoune-Gelbert, D.; Lapidot, M.; Aly, R. Broomrape can acquire viruses from its hosts. Phytopathology 2009, 99, 1321–1329. [Google Scholar] [CrossRef] [PubMed]
  57. Choi, D.; Shin, C.; Shirasu, K.; Hahn, Y. Two novel poty-like viruses identified from the transcriptome data of purple witchweed (Striga hermonthica). Acta Virol. 2021, 65, 365. [Google Scholar] [CrossRef] [PubMed]
  58. Dikova, B. Establishment of Tobacco rattle virus (TRV) in weeds and Cuscuta. Biotechnol. Biotechnol. Equip. 2006, 20, 42–48. [Google Scholar] [CrossRef]
  59. Mikona, C.; Jelkmann, W. Replication of Grapevine leafroll-associated virus-7 (GLRaV-7) by Cuscuta species and its transmission to herbaceous plants. Plant Dis. 2010, 94, 471–476. [Google Scholar] [CrossRef] [PubMed]
  60. Jelkmann, W.; Hergenhahn, F.; Berwarth, C. Transmission of Little cherry virus-1 (LChV-1) by Cuscuta europea to herbaceous host plants. In Proceedings of the 21st International Conference on Virus and other Graft Transmissible Diseases of Fruit Crops, Neustadt, Germany, 5–10 July 2009. [Google Scholar]
  61. Vachev, T.; Ivanova, D.; Minkov, I.; Tsagris, M.; Gozmanova, M. Trafficking of the Potato spindle tuber viroid between tomato and Orobanche ramosa. Virology 2010, 399, 187–193. [Google Scholar] [CrossRef] [PubMed]
  62. Ivanova, D.; Vachev, T.; Baev, V.; Minkov, I.; Gozmanova, M. Identification of Potato Spindle Tuber Viroid Small RNA in Orobanche ramosa by Microarray. Biotechnol. Biotechnol. Equip. 2010, 24, 144–146. [Google Scholar] [CrossRef]
  63. Lee, I.-M.; Davis, R.E.; Gundersen-Rindal, D.E. Phytoplasma: Phytopathogenic mollicutes. Annu. Rev. Microbiol. 2000, 54, 221–255. [Google Scholar] [CrossRef]
  64. Bertaccini, A.; Duduk, B. Phytoplasma and phytoplasma diseases: A review of recent research. Phytopathol. Mediterr. 2009, 48, 355–378. [Google Scholar]
  65. IRPCM Phytoplasma/Spiroplasma Working Team–Phytoplasma Taxonomy Group. ‘Candidatus Phytoplasma’, a taxon for the wall-less, non-helical prokaryotes that colonize plant phloem and insects. Int. J. Syst. Evol. Microbiol. 2004, 54, 1243–1255. [Google Scholar] [CrossRef] [PubMed]
  66. Kumari, S.; Nagendran, K.; Rai, A.B.; Singh, B.; Rao, G.P.; Bertaccini, A. Global status of phytoplasma diseases in vegetable crops. Front. Microbiol. 2019, 10, 441347. [Google Scholar] [CrossRef] [PubMed]
  67. Přibylová, J.; Špak, J. Dodder transmission of phytoplasmas. Phytoplasma Methods Protoc. 2013, 938, 41–46. [Google Scholar]
  68. Marcone, C.; Hergenhahn, F.; Ragozzino, A.; Seemüller, E. Dodder transmission of pear decline, European stone fruit yellows, rubus stunt, picris echioides yellows and cotton phyllody phytoplasmas to periwinkle. J. Phytopathol. 1999, 147, 187–192. [Google Scholar] [CrossRef]
  69. Ãzdemir, N.; Saygili, H.; Sahin, F.; Karsavuran, Y.; Bayrak, O.; Oral, B. Host range and genetic characterization of a phytoplasma causing tomato stolbur disease in Turkey. Acta Hortic. 2007, 808, 255–262. [Google Scholar] [CrossRef]
  70. Shokri, M.; Jafary, H.; Moghadam, M.A. Molecular detection and survey of tomato big bud phytoplasma in Zanjan province. Genet. Eng. Biosaf. J. 2023, 12, 123–130. [Google Scholar]
  71. Sarab, R.T.; Bakhsh, M.S.; Motlagh, M.A. First report of a phytoplasma associated with Orobanche spp. in Iran. In Proceedings of the 22nd Iranian Plant Protection Congress, Karaj, Iran, 27–30 August 2016. [Google Scholar]
  72. Borkar, S.G.; Yumlembam, R.A. Bacterial Diseases of Crop Plants; CRC Press: Boca Raton, FL, USA, 2016. [Google Scholar]
  73. Mansfield, J.; Genin, S.; Magori, S.; Citovsky, V.; Sriariyanum, M.; Ronald, P.; Dow, M.; Verdier, V.; Beer, S.V.; Machado, M.A. Top 10 plant pathogenic bacteria in molecular plant pathology. Mol. Plant Pathol. 2012, 13, 614–629. [Google Scholar] [CrossRef] [PubMed]
  74. Eichenlaub, R.; Gartemann, K.-H.; Burger, A. Clavibacter michiganensis, a group of Gram-positive phytopathogenic bacteria. In Plant-Associated Bacteria; Springer: Dordrecht, The Netherlands, 2006; pp. 385–421. [Google Scholar]
  75. Evseev, P.; Lukianova, A.; Tarakanov, R.; Tokmakova, A.; Shneider, M.; Ignatov, A.; Miroshnikov, K. Curtobacterium spp. and Curtobacterium flaccumfaciens: Phylogeny, genomics-based taxonomy, pathogenicity, and diagnostics. Curr. Issues Mol. Biol. 2022, 44, 889–927. [Google Scholar] [CrossRef] [PubMed]
  76. Iasur Kruh, L.; Lahav, T.; Abu-Nassar, J.; Achdari, G.; Salami, R.; Freilich, S.; Aly, R. Host-parasite-bacteria triangle: The microbiome of the parasitic weed Phelipanche aegyptiaca and tomato-Solanum lycopersicum (Mill.) as a host. Front. Plant Sci. 2017, 8, 269. [Google Scholar] [CrossRef]
  77. Ali, A.; Haider, M.S.; Muhammad, A. Biological control of fruit lesions caused by Xanthomonas campestris pathovars from Cuscuta pedicellata Ledeb. in vitro. J. Pure Appl. Microbiol. 2013, 7, 3149–3153. [Google Scholar]
  78. Islam, R.; Rahman, M.S.; Rahman, S.M. GC-MS analysis and antibacterial activity of Cuscuta reflexa against bacterial pathogens. Asian Pac. J. Trop. Dis. 2015, 5, 399–403. [Google Scholar] [CrossRef]
  79. Saadoun, I.M.; Hameed, K.M.; Al-Momani, F.; Ababneh, Q. Effect of three Orobanche spp. extracts on some local phytopathogens, Agrobacterium and Erwinia. Turk. J. Biol. 2008, 32, 113–117. [Google Scholar]
  80. Ahmad, A.; Tandon, S.; Xuan, T.D.; Nooreen, Z. A Review on Phytoconstituents and Biological activities of Cuscuta species. Biomed. Pharmacother. 2017, 92, 772–795. [Google Scholar] [CrossRef] [PubMed]
  81. Genovese, C.; D’Angeli, F.; Attanasio, F.; Caserta, G.; Scarpaci, K.S.; Nicolosi, D. Phytochemical composition and biological activities of Orobanche crenata Forssk.: A review. Nat. Prod. Res. 2021, 35, 4579–4595. [Google Scholar] [CrossRef] [PubMed]
  82. Augustine, R.; Maikasuwa, B.; Etonihu, A. Phytochemical Screening, Thin Layer Chromatography and Antibacterial Activity of the Leaf Extracts of Striga hermonthica. Chem. Res. J. 2020, 5, 192. [Google Scholar]
  83. Moore, D.; Robson, G.D.; Trinci, A.P. 21st Century Guidebook to Fungi; Cambridge University Press: Cambridge, UK, 2020. [Google Scholar]
  84. Dean, R.; Van Kan, J.A.; Pretorius, Z.A.; Hammond-Kosack, K.E.; Di Pietro, A.; Spanu, P.D.; Rudd, J.J.; Dickman, M.; Kahmann, R.; Ellis, J. The Top 10 fungal pathogens in molecular plant pathology. Mol. Plant Pathol. 2012, 13, 414–430. [Google Scholar] [CrossRef] [PubMed]
  85. Gosling, P.; Hodge, A.; Goodlass, G.; Bending, G. Arbuscular mycorrhizal fungi and organic farming. Agric. Ecosyst. Environ. 2006, 113, 17–35. [Google Scholar] [CrossRef]
  86. Akram, S.; Ahmed, A.; He, P.; He, P.; Liu, Y.; Wu, Y.; Munir, S.; He, Y. Uniting the role of endophytic fungi against plant pathogens and their interaction. J. Fungi 2023, 9, 72. [Google Scholar] [CrossRef]
  87. Leake, J.R. The biology of myco-heterotrophic (‘saprophytic’) plants. New Phytol. 1994, 127, 171–216. [Google Scholar] [CrossRef]
  88. Suryanarayanan, T.; Senthilarasu, G.; Muruganandam, V. Endophytic fungi from Cuscuta reflexa and its host plants. Fungal Divers. 2000, 4, 117–123. [Google Scholar]
  89. Khare, E.; Vishwakarma, A.; Maurya, V.; Kaistha, S.D. Endophytic fungi from parasitic-plant Cuscuta, and their potential for producing L-asparaginase of pharmaceutical significance. Environ. Sustain. 2024, 7, 93–101. [Google Scholar] [CrossRef]
  90. Ruraż, K.; Przemieniecki, S.W.; Piwowarczyk, R. Interspecies and temporal dynamics of bacterial and fungal microbiomes of pistil stigmas in flowers in holoparasitic plants of the Orobanche series Alsaticae (Orobanchaceae). Sci. Rep. 2023, 13, 6749. [Google Scholar] [CrossRef] [PubMed]
  91. Gafar, N.Y.; Hassan, M.M.; Ahmed, M.M.; Osman, A.G.; Abdelgani, M.E.; Babiker, A.E. In vitro study of endophytic bacteria, carbohydrates and their combination on early developmental stages of Striga hermonthica (Del.) Benth. Adv. Environ. Biol. 2016, 10, 66–74. [Google Scholar]
  92. Lombard, L.; van Doorn, R.; Groenewald, J.; Tessema, T.; Kuramae, E.; Etolo, D.; Raaijmakers, J.; Crous, P. Fusarium diversity associated with the Sorghum-Striga interaction in Ethiopia. Fungal Syst. Evol. 2022, 10, 177–215. [Google Scholar] [CrossRef] [PubMed]
  93. Dor, E.; Hershenhorn, J.; Andolfi, A.; Cimmino, A.; Evidente, A. Fusarium verticillioides as a new pathogen of the parasitic weed Orobanche spp. Phytoparasitica 2009, 37, 361–370. [Google Scholar] [CrossRef]
  94. Gibot-Leclerc, S.; Guinchard, L.; Edel-Hermann, V.; Dessaint, F.; Cartry, D.; Reibel, C.; Gautheron, N.; Bernaud, E.; Steinberg, C. Screening for potential mycoherbicides within the endophyte community of Phelipanche ramosa parasitizing tobacco. FEMS Microbiol. Ecol. 2022, 98, fiac024. [Google Scholar] [CrossRef] [PubMed]
  95. Ibdah, M.; Dubey, N.K.; Eizenberg, H.; Dabour, Z.; Abu-Nassar, J.; Gal-On, A.; Aly, R. Cucumber Mosaic Virus as a carotenoid inhibitor reducing Phelipanche aegyptiaca infection in tobacco plants. Plant Signal. Behav. 2014, 9, e972146. [Google Scholar] [CrossRef]
  96. Bangar, V.; Joshi, M.; Valvi, H. Isolation and pathogenicity of pathogen causing diseases on dodder (Cuscuta gronovii). IJCS 2019, 7, 1769–1773. [Google Scholar]
  97. Cook, J.C.; Charudattan, R.; Zimmerman, T.W.; Rosskopf, E.N.; Stall, W.M.; MacDonald, G.E. Effects of Alternaria destruens, glyphosate, and ammonium sulfate individually and integrated for control of dodder (Cuscuta pentagona). Weed Technol. 2009, 23, 550–555. [Google Scholar] [CrossRef]
  98. Idris, A.E.; Abouzeid, M.A.; Boari, A.; Vurro, M.; Evidente, A. Identification of phytotoxic metabolites of a new Fusarium sp. inhibiting germination of Striga hermonthica seeds. Phytopathol. Mediterr. 2003, 42, 65–70. [Google Scholar]
  99. Khakimov, A.; Salakhutdinov, I.; Omolikov, A.; Utaganov, S. Traditional and current-prospective methods of agricultural plant diseases detection: A review. IOP Conf. Ser. Earth Environ. Sci. 2022, 951, 012002. [Google Scholar] [CrossRef]
  100. Mehetre, G.T.; Leo, V.V.; Singh, G.; Sorokan, A.; Maksimov, I.; Yadav, M.K.; Upadhyaya, K.; Hashem, A.; Alsaleh, A.N.; Dawoud, T.M. Current Developments and Challenges in Plant Viral Diagnostics: A Systematic Review. Viruses 2021, 13, 412. [Google Scholar] [CrossRef] [PubMed]
  101. Nair, S.; Manimekalai, R. Phytoplasma diseases of plants: Molecular diagnostics and way forward. World J. Microbiol. Biotechnol. 2021, 37, 102. [Google Scholar] [CrossRef] [PubMed]
  102. Boari, A.; Vurro, M. Evaluation of Fusarium spp. and other fungi as biological control agents of broomrape (Orobanche ramosa). Biol. Control 2004, 30, 212–219. [Google Scholar] [CrossRef]
  103. Durlik, K.; Żarnowiec, P.; Piwowarczyk, R.; Kaca, W. Culturable endophytic bacteria from Phelipanche ramosa (Orobanchaceae) seeds. Seed Sci. Res. 2021, 31, 69–75. [Google Scholar] [CrossRef]
  104. Kamińska, M.; Korbin, M. Graft and dodder transmission of phytoplasma affecting lily to experimental hosts. Acta Physiol. Plant. 1999, 21, 21–26. [Google Scholar] [CrossRef]
  105. van der Wolf, J.; De Boer, S.H. Phytopathogenic bacteria. In Principles of Plant-Microbe Interactions: Microbes for Sustainable Agriculture; Springer: Cham, Switzerland, 2014; pp. 65–77. [Google Scholar]
  106. Hettenhausen, C.; Li, J.; Zhuang, H.; Sun, H.; Xu, Y.; Qi, J.; Zhang, J.; Lei, Y.; Qin, Y.; Sun, G. Stem parasitic plant Cuscuta australis (dodder) transfers herbivory-induced signals among plants. Proc. Natl. Acad. Sci. USA 2017, 114, E6703–E6709. [Google Scholar] [CrossRef]
  107. Avila, A.T.; Van Laar, T.A.; Constable, J.V.; Waselkov, K. 16S rRNA Gene Diversity of Bacterial Endophytes in Parasitic Cuscuta campestris and Its Helianthus annuus Host. Microbiol. Resour. Announc. 2020, 9, e00968-20. [Google Scholar] [CrossRef] [PubMed]
  108. Hogenhout, S.A.; Ammar, E.-D.; Whitfield, A.E.; Redinbaugh, M.G. Insect vector interactions with persistently transmitted viruses. Annu. Rev. Phytopathol. 2008, 46, 327–359. [Google Scholar] [CrossRef]
  109. Sarwar, M. Mite (Acari Acarina) vectors involved in transmission of plant viruses. In Applied Plant Virology; Elsevier: Amsterdam, The Netherlands, 2020; pp. 257–273. [Google Scholar]
  110. de Lillo, E.; Freitas-Astúa, J.; Watanabe Kitajima, E.; Ramos-González, P.L.; Simoni, S.; Tassi, A.D.; Valenzano, D. Phytophagous mites transmitting plant viruses: Update and perspectives. Entomol. Gen. 2021, 41, 439. [Google Scholar] [CrossRef]
  111. Singh, S.; Awasthi, L.; Jangre, A.; Nirmalkar, V.K. Transmission of plant viruses through soil-inhabiting nematode vectors. In Applied Plant Virology; Elsevier: Amsterdam, The Netherlands, 2020; pp. 291–300. [Google Scholar]
  112. Weintraub, P.G.; Beanland, L. Insect vectors of phytoplasmas. Annu. Rev. Entomol. 2006, 51, 91–111. [Google Scholar] [CrossRef] [PubMed]
  113. Alma, A.; Lessio, F.; Nickel, H. Insects as phytoplasma vectors: Ecological and epidemiological aspects. In Phytoplasmas: Plant Pathogenic Bacteria-II: Transmission and Management of Phytoplasma-Associated Diseases; Springer: Singapore, 2019; pp. 1–25. [Google Scholar]
  114. Bini, A.P.; Serra, M.C.D.; Pastore, I.F.; Brondi, C.V.; Camargo, L.E.A.; Monteiro-Vitorello, C.B.; van Sluys, M.-A.; Rossi, G.D.; Creste, S. Transmission of Xanthomonas albilineans by the spittlebug, Mahanarva fimbriolata (Hemiptera: Cercopidae), in Brazil: First report of an insect vector for the causal agent of sugarcane leaf scald. J. Insect Sci. 2023, 23, 28. [Google Scholar] [CrossRef] [PubMed]
  115. Kisera Kwach, J.; Nyakomitta, P.S. Diversity of Insects Associated with Banana in Banana Xanthomonas Wilt Epidemic Areas of Western Kenya. Asian J. Agric. Hortic. Res. 2022, 9, 1–12. [Google Scholar] [CrossRef]
  116. Esquivel, J.F.; Bell, A.A. Acquisition and Transmission of Fusarium oxysporum f. sp. vasinfectum VCG 0114 (Race 4) by Stink Bugs. Plant Dis. 2021, 105, 3082–3086. [Google Scholar] [PubMed]
  117. Song, J.; Bian, J.; Xue, N.; Xu, Y.; Wu, J. Inter-species mRNA transfer among green peach aphids, dodder parasites, and cucumber host plants. Plant Divers. 2022, 44, 1–10. [Google Scholar] [CrossRef] [PubMed]
  118. Piwowarczyk, R.; Mielczarek, Ł.; Panek-Wójcicka, M.; Ruraż, K. First report of Melanagromyza cuscutae (Diptera: Agromyzidae) from Poland. Fla. Entomol. 2020, 103, 124–126. [Google Scholar] [CrossRef]
  119. Bayram, Y.; Cikman, E. Efficiency of Pytomyza orobanchia Kaltenbach (Diptera: Agromyzidae) on Orobanche crenata Forsk. (Orobanchaceae) in Lentil Fields at Diyarbakır and Mardin Provinces, Turkey. Egypt. J. Biol. Pest Control 2016, 26, 365–371. [Google Scholar]
  120. Aistova, E.; Bezborodov, V. Weevils belonging to the Genus Smicronyx Schönherr, 1843 (Coleoptera, Curculionidae) affecting dodders (Cuscuta Linnaeus, 1753) in the Russian Far East. Russ. J. Biol. Invasions 2017, 8, 184–188. [Google Scholar] [CrossRef]
  121. Zagorchev, L.; Albanova, I.; Lozanova, Y.; Odjakova, M.; Teofanova, D. Chitinase Profile of Arabidopsis, Subjected to the Combined Stress of Soil Salinity, Cuscuta campestris Parasitism and Herbivores. Proc. Bulg. Acad. Sci. 2022, 75, 835–844. [Google Scholar] [CrossRef]
  122. Cagáň, L.; Komáromyová, E.; Barta, M.; Tóth, P. Cuscuta lupuliformis Krocker (Cuscutaceae)—A new host for bean aphid, Aphis fabae Scopoli (Homoptera, Aphididae). Acta Fytotech. Zootech. 2001, 4, 84. [Google Scholar]
  123. Azami-Sardooei, Z.; Shahreyarinejad, S.; Rouzkhosh, M.; Fekrat, F. The first report on feeding of Oxycarenus hyalinipennis and Aphis fabae on dodder Cuscuta campestris in Iran. J. Crop Prot. 2018, 7, 121–124. [Google Scholar]
  124. Zhuang, H.; Li, J.; Song, J.; Hettenhausen, C.; Schuman, M.C.; Sun, G.; Zhang, C.; Li, J.; Song, D.; Wu, J. Aphid (Myzus persicae) feeding on the parasitic plant dodder (Cuscuta australis) activates defense responses in both the parasite and soybean host. New Phytol. 2018, 218, 1586–1596. [Google Scholar] [CrossRef]
  125. Piwowarczyk, R.; Guzikowski, S.; Depa, Ł.; Kaszyca, N. First report of Smynthurodes betae (Hemiptera: Aphididae) on Phelipanche ramosa (Orobanchaceae). Fla. Entomol. 2018, 101, 339–341. [Google Scholar] [CrossRef]
  126. Boukhris-Bouhachem, S.; Youssef, S.B.; Kharrat, M. First report of Geoica utricularia (Hemiptera: Aphididae) population on parasitic broomrape Orobanche foetida. Fla. Entomol. 2011, 94, 343–344. [Google Scholar] [CrossRef]
  127. Tóth, P.; Černý, M.; Cagáň, L. First records of Melanagromyza cuscutae Hering, 1958 (Diptera: Agromyzidae) from Slovakia and its new host plant. Entomol. Fenn. 2004, 15, 48–52. [Google Scholar] [CrossRef]
  128. Klein, O.; Kroschel, J. Biological control of Orobanche spp. with Phytomyza orobanchia, a review. Biocontrol 2002, 47, 245–277. [Google Scholar] [CrossRef]
  129. Al-Eryan, M.; Altahtawy, M.; El-Sherief, H.; Abu-Shall, A. Efficacy of Phytomyza orobanchia Kalt. in reduction of Orobanche crenata Forsk. seed yield under semi-field conditions. Egypt. J. Biol. Pest Control 2004, 14, 237–242. [Google Scholar]
  130. Zhekova, E.; Petkova, D.; Ivanova, I. Smicronyx smreczynskii F. Solari, 1952 (Insecta: Curculionidae): Possibilities for biological control of two Cuscuta species (Cuscutaceae) in district of Ruse. Acta Zool. Bulg. 2014, 66, 431–432. [Google Scholar]
  131. Otoidobiga, L.C.; Vincent, C.; Stewart, R.K. Relationship between Smicronyx spp. population and galling of Striga hermonthica (Del.) Benth. Int. J. Trop. Insect Sci. 1998, 18, 197–203. [Google Scholar] [CrossRef]
  132. Rubiales, D.; Fernández-Aparicio, M.; Wegmann, K.; Joel, D. Revisiting strategies for reducing the seedbank of Orobanche and Phelipanche spp. Weed Res. 2009, 49, 23–33. [Google Scholar] [CrossRef]
  133. Prider, J. The reproductive biology of the introduced root holoparasite Orobanche ramosa subsp. mutelii (Orobanchaceae) in South Australia. Aust. J. Bot. 2015, 63, 426–434. [Google Scholar]
  134. Goldwasser, Y.; Miryamchik, H.; Rubin, B.; Eizenberg, H. Field dodder (Cuscuta campestris)—A new model describing temperature-dependent seed germination. Weed Sci. 2016, 64, 53–60. [Google Scholar] [CrossRef]
  135. Chalam, V.C.; Deepika, D.; Abhishek, G.; Maurya, A. Major seed-borne diseases of agricultural crops: International trade of agricultural products and role of quarantine. In Seed-Borne Diseases of Agricultural Crops: Detection, Diagnosis & Management; Springer: Singapore, 2020; pp. 25–61. [Google Scholar]
  136. Bennett, C. Latent virus of dodder and its effect on sugar beet and other plants. Phytopathology 1944, 34, 77–91. [Google Scholar]
  137. Huet, S.; Pouvreau, J.-B.; Delage, E.; Delgrange, S.; Marais, C.; Bahut, M.; Delavault, P.; Simier, P.; Poulin, L. Populations of the parasitic plant Phelipanche ramosa influence their seed microbiota. Front. Plant Sci. 2020, 11, 1075. [Google Scholar] [CrossRef] [PubMed]
  138. Petrosyan, K.; Thijs, S.; Piwowarczyk, R.; Ruraż, K.; Kaca, W.; Vangronsveld, J. Diversity and potential plant growth promoting capacity of seed endophytic bacteria of the holoparasite Cistanche phelypaea (Orobanchaceae). Sci. Rep. 2023, 13, 11835. [Google Scholar] [CrossRef] [PubMed]
  139. Nadler-Hassar, T.; Shaner, D.L.; Nissen, S.; Westra, P.; Rubin, B. Are herbicide-resistant crops the answer to controlling Cuscuta? Pest Manag. Sci. Former. Pestic. Sci. 2009, 65, 811–816. [Google Scholar] [CrossRef] [PubMed]
  140. Bewick, T.; Porter, J.; Ostrowski, R. Smolder™: A bioherbicide for suppression of dodder (Cuscuta spp.). In Proceedings of the Annual Meeting-Southern Weed Science Society, Tulsa, OK, USA, 24–26 January 2000. [Google Scholar]
  141. Hadizadeh, F.; Mehrvarz, S.S.; Mirpour, M.S. Effect of Bacillus spp. on seed germination of selected species of the genus Cuscuta (Convolvulaceae). Mod. Phytomorphol. 2014, 6, 97. [Google Scholar]
  142. Barghouthi, S.; Salman, M. Bacterial inhibition of Orobanche aegyptiaca and Orobanche cernua radical elongation. Biocontrol Sci. Technol. 2010, 20, 423–435. [Google Scholar] [CrossRef]
  143. Zermane, N.; Souissi, T.; Kroschel, J.; Sikora, R. Biocontrol of broomrape (Orobanche crenata Forsk. and Orobanche foetida Poir.) by Pseudomonas fluorescens isolate Bf7-9 from the faba bean rhizosphere. Biocontrol Sci. Technol. 2007, 17, 483–497. [Google Scholar] [CrossRef]
  144. Elzein, A.; Kroschel, J. Fusarium oxysporum Foxy 2 shows potential to control both Striga hermonthica and S. Asiat. Weed Res. 2004, 44, 433–438. [Google Scholar] [CrossRef]
  145. Ndambi, B.; Cadisch, G.; Elzein, A.; Heller, A. Colonization and control of Striga hermonthica by Fusarium oxysporum f. sp. strigae, a mycoherbicide component: An anatomical study. Biol. Control 2011, 58, 149–159. [Google Scholar]
  146. Oula, D.A.; Nyongesah, J.M.; Odhiambo, G.; Wagai, S. The effectiveness of local strains of Fusarium oxysporium f. Sp. Strigae to control Striga hermonthica on local maize in western Kenya. Food Sci. Nutr. 2020, 8, 4352–4360. [Google Scholar] [CrossRef]
  147. Baker, C.S.; Sands, D.C.; Nzioki, H.S. The Toothpick Project: Commercialization of a virulence-selected fungal bioherbicide for Striga hermonthica (witchweed) biocontrol in Kenya. Pest Manag. Sci. 2024, 80, 65–71. [Google Scholar] [CrossRef] [PubMed]
  148. Müller-Stöver, D.; Kohlschmid, E.; Sauerborn, J. A novel strain of Fusarium oxysporum from Germany and its potential for biocontrol of Orobanche ramosa. Weed Res. 2009, 49, 175–182. [Google Scholar] [CrossRef]
  149. Anteyi, W.O.; Klaiber, I.; Rasche, F. Diacetoxyscirpenol, a Fusarium exometabolite, prevents efficiently the incidence of the parasitic weed Striga hermonthica. BMC Plant Biol. 2022, 22, 84. [Google Scholar] [CrossRef]
  150. Sands, D.C.; Pilgeram, A.L. Methods for selecting hypervirulent biocontrol agents of weeds: Why and how. Pest Manag. Sci. 2009, 65, 581–587. [Google Scholar] [CrossRef]
Figure 1. Putative transmission routes for phytopathogens, hypothesized for Cuscuta spp. as a vector. The parasitic plant acquires phytopathogens through the haustorial connection with an infected host plant (1) or from arthropods feeding on it (2). Subsequently, the phytopathogens are transmitted through the seed bank of Cuscuta (3) or directly (4) to a non-infected host plant.
Figure 1. Putative transmission routes for phytopathogens, hypothesized for Cuscuta spp. as a vector. The parasitic plant acquires phytopathogens through the haustorial connection with an infected host plant (1) or from arthropods feeding on it (2). Subsequently, the phytopathogens are transmitted through the seed bank of Cuscuta (3) or directly (4) to a non-infected host plant.
Pathogens 13 00484 g001
Table 1. Recent examples of plant viruses, detected in parasitic plants.
Table 1. Recent examples of plant viruses, detected in parasitic plants.
SpeciesParasitic PlantReference
TYLCV, CMVCuscuta campestris[54]
TRVCuscuta spp.[58]
GLRaV-7Cuscuta spp.[59]
LChV-1Cuscuta europaea[60]
PVYCuscuta reflexa[55]
SaPlV1/2Striga hermonthica[57]
CMV, ToMV, PVY, TYLCVPhelipanche aegyptiaca[56]
PSTVdOrobanche ramosa[61,62]
Abbreviations: TYLCV—Tomato Yellow Leaf Curl Virus; CMV—Cucumber Mosaic Virus; TRV—Tobacco Rattle Virus; GLRaV-7—Grapevine Leafroll-Associated Virus-7; LChV-1—Little Cherry Virus-1; PVY—Potato Virus Y; SaPlV—Striga-Associated Poty-Like Virus; ToMV—Tomato Mosaic Virus; PSTVd—Potato Spindle Tuber Viroid.
Table 2. Non-exhaustive list of arthropods, associated with different parasitic plants.
Table 2. Non-exhaustive list of arthropods, associated with different parasitic plants.
GroupSpeciesParasitic PlantReference
Hemiptera: Aphididae
Hemiptera: Flatidae
Hemiptera: Lygaeidae
Diptera: Agromyzidae
Coleoptera: Curculionidae
Aphis fabae
Myzus persicae
Smynthurodes betae
Geoica utricularia
Metcalfa pruinosa
Oxycarenus hyalinipennis
Melanagromyza cuscutae
Phytomyza orobanchia
Smicronyx spp.
Cuscuta lupuliformis
Cuscuta campestris
Cuscuta australis
Phelipanche ramosa
Orobanche foetida
Cuscuta campestris
Cuscuta campestris
Cuscuta spp.
Orobanche spp.

Cuscuta spp.
Striga spp.
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Savov, S.; Marinova, B.; Teofanova, D.; Savov, M.; Odjakova, M.; Zagorchev, L. Parasitic Plants—Potential Vectors of Phytopathogens. Pathogens 2024, 13, 484.

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Savov S, Marinova B, Teofanova D, Savov M, Odjakova M, Zagorchev L. Parasitic Plants—Potential Vectors of Phytopathogens. Pathogens. 2024; 13(6):484.

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Savov, Stefan, Bianka Marinova, Denitsa Teofanova, Martin Savov, Mariela Odjakova, and Lyuben Zagorchev. 2024. "Parasitic Plants—Potential Vectors of Phytopathogens" Pathogens 13, no. 6: 484.

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