**4. Microbiomes of parasitic plants and their hosts**

Microbiomes can expand the genomic potential of plants through efficient nutrients acquisition, promoting growth and development, and tolerance to biotic and abiotic stresses [65]. Endophytic microbial communities of parasitic plants may affect parasitism and influence host microbial composition. Microbiota or microbial communities within a parasite can be divided into core- and transientmicrobes. Core microbes are intrinsic to one or more developmental stages of a parasite that can vertically flow from parents to the offspring. Transient microbes are temporarily acquired by the parasite from their interacting hosts or environment [66]. A study on microbial communities of parasitic weed, *P. aegyptiaca*, showed that endophytic bacteria were present at different development stages (pre-haustorium, tubercle, and shoot) of the parasite [13]. It was observed that the presence of alpha- and gamma-proteobacteria (dominant species: *Sphingomonas* and *Acinetobacter*) were abundant during pre-haustorium formation (pre-attachment to the host). In the post-attachment stage, i.e., during attachment of tubercle of the parasite to the host, bacterial communities shifted to flavobacteria and betaproteobacteria, while during parasite shoot formation, an increase of Bacilli and Actinobacteria have been reported [13]. Besides bacterial communities, endophytic fungi also inhabit the inner tissues of parasitic plants. For instance, the rootparasitic plant *Cynomorium songaricum* parasitise *Nitraria tangutorum*, a flowering shrub from the *Nitrariaceae* family, harbours several fungal species assemblages belonging to the phylum, Ascomycota, Basidiomycota and Zygomycota [67]. Microbial communities play diverse roles during the growth and development and parasitism of parasitic plants on their host plants. For instance, some species of the genus, *Fusarium*, promote parasite seed germination, while symbiosis of arbuscular mycorrhizal fungi (*Glomus mosseae*) and rhizobia can alleviate plant host damage by root hemiparasites [67, 68]. Nitrogen-fixing bacteria associated with host plants may indirectly benefit parasitic plants through efficient N2-fixation and their availability to the parasite during host attachment [69, 70].

Microbial communities of parasitic plants overlapped extensively with their parasitised host while still maintaining taxonomically distinct communities [67, 71]. For instance, bacteria communities of the root holoparasite, *Orobanche hederae*, exhibit strong congruency with the host, *Hedera*; however, the individual bacterial taxa were differentially abundant between *Orobanche* and *Hedera* roots [72]. Transmission of microbiota through xylem tubes or apoplasts (intercellular spaces) may act as a mechanism for the shared microbial communities between the host plant and the parasite [13]. Studies have shown that host-associated microbes induce resistance against parasitic plants in many agriculturally important crop species. The induced resistance is mainly achieved via (i) microbe-mediated activation of the phenylpropanoid/isoflavonoid pathways leading to the production of toxic compounds, including phenolics and phytoalexins in the host plant against the parasite, (ii) reduced activity of host root exudates to inhibit parasite seed germination, and (iii) enhanced production of plant-derived peroxidase that causes tubercles necrosis of parasitic plants [73, 74]. Some *Fusarium* species can directly penetrate *Orobanche* cells leading to disintegration of cytoplasm without apparent damage to the host plant tomato [75]. Root-associated microbes can also modulate root physiology and architecture of host plants to prevent parasite seed germination and infection on hosts [76]. An example is colonisation by an arbuscular mycorrhizal fungus (*Glomus intraradices*) on tomato, which resulted in reduced root exudation of strigolactone (chemo-attractant for parasitic plants) and prevented germination of the *P. ramosa* seeds [77]. In another case study, the release of volatile organic compounds such as sesquiterpenes by ectomycorrhizal fungus, *Laccaria bicolor*, promoted lateral root formation in poplar and *Arabidopsis* plants [78]. Thus, changes in root architecture can potentially affect host infection by parasitic plants [76].

### **5. Mechanism of pathogen transmission**

Plant pathogens (mostly, viruses and phytoplasmas) are transmitted by parasitic plants by their twining stems. The parasite stem adheres to the host's stem by exuding cutin as it wraps tightly around the stem of the host plant. Few species of parasitic plants like *Cuscuta californica*, *C. campestris, C. subinclusa*, *C. europaea, C. epilinum* and *C. lupuliformis* are sometimes employed in various research areas for the transmission of viruses [9].

#### *Parasitic Plants as Vectors for Pathogens DOI: http://dx.doi.org/10.5772/intechopen.100187*

The parasitic plants attach to the host plant through haustoria which originates at the site of association between the parasite stem coil and the host stem or leaf. The haustoria vary among different parasitic plant species, considerably in their anatomy and function, mostly by whether they form connections exclusively to the xylem only or both xylem and phloem [40]. Initially, the haustorium enters the host tissue through the lower haustorium with the help of enzymes that break down cell wall connections. Cells then begin to elongate from the lower haustorium and traverse throughout the host tissue to reach the vascular system of the host which eventually leads to the formation of searching hyphae [79]. These cells, termed searching hyphae, as it grows through the host cells, formation of new host cell wall occurs over the parasite cell wall, which appears to encase the hyphae over their entire surface. This formation of a new host cell wall around the parasite cell wall forms a host–parasite interface similar to that of neighbouring cells of the same species. The searching hyphae may develop as a xylem element when connections are made with the host xylem or it may differentiate into cells that are similar to sieve elements after contacting the host phloem.

The host–parasite cell wall is perforated by both simple and branched plasmodesmata, complete with desmotubules typical of normal plasmodesmata [80]. The plant pathogens, mostly viruses are transmitted to the host plant through these plasmodesmata. The virus transmission through the plasmodesmata is felicitated by non-structural proteins, called movement proteins, which act to facilitate the movement of virus particles from cell to cell through these plasmodesmata [81].

Another mechanism of transmission of the virus from the infected parasite to the host is through the sieve element. The virus after being acquired from the vascular bundles of the infected host plant by the haustoria is transmitted in the food stream of the parasitic plant. After translocation through the parasite phloem, the virus is introduced to the next plant by the new parasite haustoria produced in contact with the vascular bundles of the inoculated plant. The parasitic plant absorbs phloem contents from the host, the searching hyphae of the parasite that contact host sieve elements grow around the element with finger-like projections. The parasite cell then differentiates like a sieve element, but with extensive development of smooth endoplasmic reticulum (ER) near the host cell and grows around the phloem cells of the host [82]. These parasite cells then differentiate in a manner consistent with the development of sieve elements, although they also contain an elaborate network of smooth ER proximal to the host cell, a feature of transfer cells [83]. In contrast to *Cuscuta*, direct connections between sieve elements of *Orobanche crenata* and those of its host *Vicia narbonensis* have been imaged using electron microscopy [82]. Host–parasite connections for *Orobanche* are less controversial in that direct connections between host and parasite sieve elements have been documented by electron microscopy. Plasmodesmata between these species have also been documented and are proposed to lead to the formation of sieve pores between adjacent sieve elements. Because sieve pores are much larger than plasmodesmata openings, the path for pathogens from host to parasite would seem to be relatively unobstructed.

### **6. Management**

The management of parasitic plants is difficult because there are few sources of crop resistance and is challenging to selectively kill the parasitic plants without damaging the host, as they are physically and biochemically attached to the host. The efficiency of the management of parasitic plants is also obstructed due to

#### *Parasitic Plants*

the dispersal efficiency, persistent seed bank, and quick responses to changes in agricultural practices. These qualities of the parasitic plants allow them to adapt to new hosts and manifest aggressively against new resistant cultivars. However, the management strategies of parasitic plants or crop resistance to parasitic plant infection can be classified as pre-attachment or post-attachment resistance according to whether the resistance occurs before or after the haustorium attaches to the host surface [84].

Mostly, the pre-attachment resistance or management includes the mechanisms that can be adopted by a host plant to prevent or avoid parasite attachment, this includes (*i*) prevent germination of the seed by reduced production of germination stimulant(s); (*ii*) production of germination inhibitors; (*iii*) delay, reduction, or complete inhibition of haustorium formation leading to attachment incompetence; and (*iv*) to impede the attachment on the host surface by formation of preformed mechanical or structural barriers which include enhanced cell wall lignification, suberization, or other modifications and structures (hairs or other outgrowths) that retard attachment to the host [5].

Post-attachment resistance occurs when the attached parasite haustorium attempts to penetrate host tissues to make connections with the vascular system. Substantial experimental evidence demonstrates that parasitic plants connect to the endodermis by activating the expression of genes encoding various cell wall degrading/softening enzymes such as pectate lyases, pectin methylesterase, polygalacturonase, endocellulase, β-xylanase, expansins. The expression of these enzymes assists the parasitic plants to penetrate the host endodermis through the epidermis and cortex [85]. During this intrusive process, the host can succumb passively, rely on constitutively expressed general defence responses, or activate specific innate immune response cascades to fend off parasitic progress [86]. Innate immunity can present as (*i*), the synthesis and release of cytotoxic compounds (e.g., phenolic acids, phytoalexins), by the challenged host root cells; (*ii*) rapid formation of physical barriers to prevent possible pathogen progress and growth (e.g., lignification and other forms of cell wall modification at the host–parasite interface); (*iii*) release of reactive oxygen species and activation of programmed cell death in the form of a hypersensitive response at the point of parasite attachment to limit parasite development and retard its penetration; and (*iv*) prevention of the parasite establishing the essential functional vascular continuity (i.e., xylem-to-xylem and/or phloem-to-phloem connections) with the host, delaying parasite growth followed by parasite developmental arrest and eventual death [5, 87].

#### **6.1 Use of herbicides as a strategy for parasitic plant control**

The use of herbicides for management needs to be specifically designed depending on the target combination of the parasite-crop species and on the information available on the specific herbicide and the optimum herbicidal doses that have been proved to be sub lethal for the crop, on the other hand, it can be applied as lethal doses to the parasite, and the availability of crop varieties with herbicide resistance.

The systemic herbicide is applied to the crop foliage and delivered to the shoot or root parasites either via the haustorium or through exudation to the rhizosphere from the crop roots [88]. The systemic herbicides used for parasitic weeds include inhibitors of aromatic (glyphosate) or branched-chain amino acid synthesis (imidazolinones and sulfonylureas), inhibitors of the vitamin folic acid (asulam), inhibitors of glutamine synthetase (glufosinate), or hormonal herbicides (2,4-D and dicamba) [89, 90].

Rationale and most effective control of parasitic plant disease is possible only if

