Abstract

Parasitic plants form intimate contacts with host tissue in order to gain access to host solutes. There are a variety of cell types within the host which parasitic plants could access to extract solutes. Depending on the degree to which the parasite has embraced the parasitic lifestyle, the extent of solute flux and the pathways used to transfer solutes from host to parasite will vary. To date, a variety of experimental approaches argue for diversity in the mechanisms and the routes by which parasites accumulate host solutes. Contact between host and parasite ranges from direct lumen‐to‐lumen links between host and parasite xylem and continuity between the sieve elements of host and parasite, to the involvement of transfer cells between host and parasite. Progress has been slow since Solms‐Laubach distinguished types of parasitic plants that fed from host phloem or xylem in 1867, but advances in clearly delineating the pathways that link host and parasite should now be possible using fluorescent proteins expressed and restricted to particular cell types of the host. This will initially necessitate using Arabidopsis, but should allow the types of connection, i.e. symplasmic or apoplasmic, to be determined and then the identification of parasite transporters responsible for solute flux.

Introduction

Parasitism among plants appears to have evolved multiple times during the radiation of the angiosperms. So far around 3000 species of parasitic angiosperm, distributed amongst 17 families have been documented (Parker and Riches, 1993). Whilst only a small number of these parasitic plants have been studied, our limited knowledge of their biology argues for diversity in the extent to which they rely on a host for growth, and also variation in the mechanisms by which they collect and attract host solutes. For example, some species of parasitic plant have functional roots (e.g. species of Rhinanthus and Olax) and therefore are able to take up inorganic nutrients from the soil, others have only vestigial ‘lumps’ (e.g. species of Orobanche), while others possess nothing that either resembles a root nor functions as one (e.g. the Cuscutaceae and the mistletoes). Thus there is a gradation in the extent to which parasitic plants must remove inorganic nutrients from a host. There is also a great deal of variation in terms of the photosynthetic capacity of parasitic plants. Some facultative parasitic plants such as Rhinanthus minor appear to have functional photosynthetic apparatus and can grow without a host providing reduced carbon. Others can not grow without the supply of photosynthate from a host either due to very low photosynthetic capacity (e.g. Cuscuta reflexa; Hibberd et al., 1998) or the complete inability to photosynthesize (e.g. Epifagus virginiana; dePamphlis and Palmer, 1990).

The degradation of nutrient uptake capacity and the ability to photosynthesize is associated with the production of specialized organs and mechanisms to steal resources from autotrophic plants. The transfer of host solutes into a parasitic plant relies on the formation of a bridge between the two organisms. This organ, the haustorium (from the Latin, haurire, to drink) is thus the defining feature of all parasitic plants. Parasitic plants can form haustoria within various host tissues, and this has led to convenient, yet unsatisfactory distinctions being made between a ‘shoot parasite’ and a ‘root parasite’. Although useful for descriptive purposes, these classifications provide no clue as to the mechanisms by which the parasites attach to the host, the cell types or apoplasmic spaces from which they collect solutes, the ways they attract host solutes, nor their impact on host nutrition.

Electron micrographs show that parasitic plants can form close contacts with host cells and, therefore, presumably set up symplasmic connections with those host cells (see below) but there is little definitive data on the mechanisms or routes by which parasitic plants remove solutes from the host in vivo. In this review, the anatomical evidence for associations between different cells of host and parasite will be summarized first, then the extent of current knowledge about solute flux through these pathways will be outlined and, lastly, an attempt will be made to identify the gaps in current knowledge that can be addressed with relatively simple experimental approaches.

Pathways and solute fluxes between hosts and parasitic plants

Parasitic plants have various means of removing solutes from the host. It appears that most make a bee‐line for their host's vascular system, where they are presumably able to tap into the large flux of amino acids, organic acids, ions, and water in the host xylem or the sugars, ions and amino acids of the host phloem. The options for contact include xylem vessels of host and parasite being adjacent to one another, direct lumenal contact between the xylem of host and parasite, symplasmic continuity between the phloem of host and parasite, or movement of either xylem or phloem solutes via specialized transfer cells into the vasculature of the parasite. These options are illustrated in Fig. 1. The evidence for these pathways from anatomical studies and then from studies where measurements of solutes in host and parasite have been measured, compared and interpreted, is addressed.

Anatomical evidence

In 1867 the Count of Solms‐Laubach published a review on the anatomy and development of parasitic plants (Solms‐Laubach, 1867). At this stage, it was already known that different parasites came into very close contact with specific cell types of the host. For example, in Orobanche, close proximity between host and parasite xylem and phloem were already described. In contrast, in Rhinanthus, only the proximity between host and parasite xylem was evident. In addition, Solms‐Laubach made the distinction between parasites that withdraw carbon in complex, organic forms from the hosts, and the Loranthaceae, which derive only water and minerals from their hosts. This observation of variation between species has been vindicated. Work by subsequent authors has shown that contact between xylem of host and parasite ranges from close proximity between tracheids and a large degree of contact between parenchymatous cells of host and parasite (Olax phyllanthi; Pate et al., 1990), to direct lumenal contact between Striga hermonthica and hosts (Dörr, 1997) although in the latter study the proportion of lumen‐to‐lumen contact within the haustorium was not documented. In some species such as Olax phyllanthi, phloem appears to be absent in haustoria (Pate et al., 1990), while in others such as Orobanche and Cuscuta it is present (Kuijt and Toth, 1976; Dörr and Kollman, 1995). The extent to which the phloem of host and parasite communicate differs between species. It is reported that Striga asiatica possesses no phloem links to its hosts (although phloem‐like cells were seen in the nucleus of the haustorium) (Rogers and Nelson, 1962). In contrast, Striga gesneriodes and Pisum sativum appear to develop interspecific plasmodesmata (Dörr, 1996), and in Orobanche crenata it has been proposed that interspecific plasmodesmata develop into sieve pores between adjacent sieve elements of host and parasite (Dörr and Kollman, 1995). Transfer cells linking the phloem of host and parasite have also been reported, and in some cases these appear to be present in association with interspecific pores between host and parasite (Dörr, 1996).

Fig. 1.

Potential pathways via which parasitic plants could contact their hosts and access host solutes. (A) Contact between xylem of host and parasite. The xylem of parasite 1 (ParX 1) contacts the xylem of its host (HX), but there are no direct lumenal connections. The xylem of the second parasite (ParX 2), however, forms lumenal links with the host xylem. No connections are made to the host xylem parenchyma (host XP). (B) Transfer cells with fewer (ParX 3) or greater (ParX 4) degrees of cell membrane invagination of the parasite xylem parenchyma (ParXP) to facilitate solute flux, link parasite and host xylem. (C) The host sieve elements (HSE) of the phloem are lined by haustorial transfer cells (HauTC) of the parasite, which then allow unloading of host phloem solutes into the parasite haustorium. CC, companion cell; PAR, parenchyma. (D) Interspecific plasmodesmata or even interspecific sieve plates (ISSP) appear at the interface of host sieve elements (HSE) and parasite phloem sieve elements (PSE).

Fig. 1.

Potential pathways via which parasitic plants could contact their hosts and access host solutes. (A) Contact between xylem of host and parasite. The xylem of parasite 1 (ParX 1) contacts the xylem of its host (HX), but there are no direct lumenal connections. The xylem of the second parasite (ParX 2), however, forms lumenal links with the host xylem. No connections are made to the host xylem parenchyma (host XP). (B) Transfer cells with fewer (ParX 3) or greater (ParX 4) degrees of cell membrane invagination of the parasite xylem parenchyma (ParXP) to facilitate solute flux, link parasite and host xylem. (C) The host sieve elements (HSE) of the phloem are lined by haustorial transfer cells (HauTC) of the parasite, which then allow unloading of host phloem solutes into the parasite haustorium. CC, companion cell; PAR, parenchyma. (D) Interspecific plasmodesmata or even interspecific sieve plates (ISSP) appear at the interface of host sieve elements (HSE) and parasite phloem sieve elements (PSE).

Evidence from solutes

Since anatomical studies showed that parasitic plants are able to form intimate contacts with the host vascular system (see above), the question of whether parasitic plants are supplied with solutes that originate from the phloem or the xylem of the host arose. Although a simple question in itself, it has not proved easy to answer, and the developmental processes responsible for setting up close contact or even continuity between specific cell types of host and parasite have to date remained unapproachable.

Using the green fluorescent protein expressed specifically from companion cells of the phloem has provided good evidence that there is symplasmic continuity between the phloem of tobacco and the parasitic plant Cuscuta reflexa (Haupt et al., 2001). This approach needs to be refined and extended to other associations between parasitic plants and their hosts.

The more standard experimental systems that have been used to investigate the question of whether a particular parasite receives solutes from the host phloem or xylem or both revolve mainly around sampling the xylem and phloem of host and parasite and comparing their solute compositions. Radiotracers can also be used to test whether there is the potential for a solute to move from one plant to another. A number of studies (Hibberd et al., 1999; Jeschke et al., 1994b; Tennakoon et al., 1997) have used modified versions of the approach first pioneered by John Pate to quantify fluxes of solutes in the phloem and xylem in whole plants (Pate et al., 1979). The approach relies on measurements of solute composition and concentrations within each pathway and the molar increments of elements of a tissue during a defined experimental period. If measurements from only the xylem are possible, then the flux through the phloem can be calculated from the difference between xylem supply and the rate at which the concentration of elements increase in the tissue of interest (Jeschke et al., 1994a; Hibberd et al., 1999). In some cases, the relative phloem immobility of calcium is also used as a method to predict elemental fluxes into parasites via the xylem (Jeschke et al., 1994a). To sample xylem sap from parasites that attach to roots of the host, a modified pressure vessel based on the original design of John Passioura (Passioura, 1980) was designed and used so that xylem could be sampled from both host and parasite (Hibberd et al., 1999; Seel and Jeschke, 1999).

However, direct measurements of xylem and phloem contents are not always possible. For those plants where phloem or xylem sampling is problematic, the possibility of using sap‐sucking insects such as the xylem feeding spittlebug, Philaenus spumarius (Malone et al., 1999), or the generalist phloem feeder Mysus persicae should be investigated. Markers to check the purity of samples extracted will have to be obtained, and care taken that the insect itself does not induce alterations in phloem or xylem content. The general conclusions that can be made about solute supply to those parasitic plants that appear to depend primarily on phloem or xylem of the host are summarized next.

Phloem feeders

Striking similarities in terms of how host physiology is altered exist between the parasites Cuscuta reflexa parasitizing a variety of hosts and Orobanche cernua parasitizing tobacco, despite the fact that one penetrates the stem and the other the root of the host (Jeschke et al., 1994a, b, 1997; Jeschke and Hilpert, 1997; Hibberd et al., 1999). Figure 2 presents as an example the flows of carbon and nitrogen between Lupinus albus and Cuscuta reflexa. Due to the lack of roots in both parasites, all mineral nutrients have to be supplied from the host, and most seem to come from the host phloem rather than the xylem. In Orobanche all carbon has to be supplied by the host as the parasite lacks chlorophyll and is non‐photosynthetic. In Cuscuta, although it retains functional photosynthetic apparatus in a ring of cells around its vascular system (Hibberd et al., 1998), 99% of the carbon that Cuscuta uses comes from the host (Jeschke et al., 1994b). In both species almost 100% of the carbon they take from the host derives from the host phloem (Hibberd et al., 1999; Jeschke et al., 1994b). It appears that with hosts infected by either Cuscuta or Orobanche the additional sink generated by the parasite induces an increase in host photosynthesis (Hibberd et al., 1999; Jeschke et al., 1994b; see Fig. 2). In addition, when Cuscuta parasitises Lupinus albus, atmospheric nitrogen fixation by the nitrogen‐fixing host is severely depressed (Fig. 2). Cuscuta stimulated photosynthesis in young leaves 20–30 d after the parasite attached (Jeschke and Hilpert, 1997), whilst Orobanche led to delayed senescence in older leaves by around 40 d after infection (Hibberd et al., 1999). In fact, in Cuscuta and Orobanche, the phloem supplies the majority of most nutrients, even minerals such as nitrogen, magnesium and potassium whose fluxes are larger in the host xylem. To supply Cuscuta with nitrogen from the phloem, this necessitates a massive transfer of nitrogen from host xylem to phloem, (see the asterisk in Fig. 2).

Despite the fact that it can be calculated that most solutes taken by Cuscuta and Orobanche must be supplied by host phloem, and also that microscopical evidence shows host and parasite phloem abutting one another, there is little clear evidence as to which cell types of the host are specifically involved in the transfer of solutes. An exception is the recent demonstration that symplasmic continuity is found between tobacco and Cuscuta reflexa (Haupt et al., 2001), whether the parasite also removes solutes from the apoplast has not yet been clarified. For most associations between host and parasitic plant it is not clear whether solute transfer is symplasmic, or if it involves unloading into the apoplasm and then active uptake via transporters.

Fig. 2.

Empirical model of the net flows and the incorporation of carbon (green) and nitrogen (red) in the xylem (filled arrows) and in the phloem (dashed arrows) in nitrogen‐fixing Lupinus albus and between Lupinus and the parasitic plant Cuscuta reflexa. The width of the arrows and the height of the histogram bars are proportional to the rates of flows or the rates of incorporation, respectively, of carbon and nitrogen. The area of green circles is proportional to the rate of respiration. The figures refer to mmol C (black) or N (red) plant−1 (12 d)−1 over the period 43–55 d after sowing of lupin and 4–16 d after attachment of Cuscuta. (For details see Jeschke et al., 1994b.) Note: (a) the stimulated photosynthesis, (b) the drastically depressed N2 fixation and (c) the strongly increased xylem‐to‐phloem transfer (*) in the parasitized host.

Fig. 2.

Empirical model of the net flows and the incorporation of carbon (green) and nitrogen (red) in the xylem (filled arrows) and in the phloem (dashed arrows) in nitrogen‐fixing Lupinus albus and between Lupinus and the parasitic plant Cuscuta reflexa. The width of the arrows and the height of the histogram bars are proportional to the rates of flows or the rates of incorporation, respectively, of carbon and nitrogen. The area of green circles is proportional to the rate of respiration. The figures refer to mmol C (black) or N (red) plant−1 (12 d)−1 over the period 43–55 d after sowing of lupin and 4–16 d after attachment of Cuscuta. (For details see Jeschke et al., 1994b.) Note: (a) the stimulated photosynthesis, (b) the drastically depressed N2 fixation and (c) the strongly increased xylem‐to‐phloem transfer (*) in the parasitized host.

Xylem feeders

In Odontites verna, a facultative parasite where direct xylem‐to‐xylem links have been reported (Govier et al., 1968), both 14C and 32P will move from host to parasite (Govier et al., 1967). 14CO2 fed to the host shoot moved down towards the parasite, but was not found as sucrose in the parasite itself, implying its conversion either prior to, or very soon after, transfer to the parasite. When Rhinanthus minor (another facultative parasite) is attached to a host its growth is stimulated. The reason appears to be related to increased access to xylem constituents, both higher water fluxes into the parasite and greater supplies of nitrogen and phosphorus have been reported (Seel and Jeschke, 1999). These authors noted that removing tillers from the barley host stimulated parasite growth, implying that to some extent competition between host and parasite for resources determined growth of the parasite. This competition could simply be for elements such as nitrogen and phosphorus, supplied to the parasite via the host xylem, that would otherwise be allocated to the developing tillers of the host.

The South‐Western Australian hemiparasite, Olax phyllanthi, possesses few xylem connections to the host (Pate et al., 1990), so transfer cells appear to be critical to move solutes from host to parasite. The contents of the parasite xylem depends on the species of host it parasitizes, the extent to which conversion of host metabolites occurs, and also the degree to which the parasite is selective in its uptake of solutes (Pate et al., 1994; Tennakoon and Pate, 1996). Olax phyllanthi is also able to parasitize nodules formed on the leguminous host Acacia littorea, but this appears to be a relatively minor route by which it removes nitrogen from the host (Tennakoon et al., 1997). Figure 3 provides an example of the flows between the nitrogen‐fixing Acacia littorea and Olax phyllanthi showing that apart from a strong increase in host root and a decrease in host shoot growth, the repercussions of this xylem feeder on its host were not as serious as with phloem feeders.

The other well‐known xylem tapping parasites are the mistletoes. It appears that the mistletoes that tap into host xylem can receive between 5% and 63% of their carbon from the host (Richter and Popp, 1992; Marshall et al., 1994; Marshall and Ehleringher, 1990). There is little full modelling data available on interactions between mistletoes and their hosts.

Xylem feeders tend to be hemiparasites, using the host xylem to bolster their own resources. There are fewer complete quantitative assessments of fluxes between host and parasite for both xylem and phloem in xylem feeders than for the predominantly phloem feeding parasites.

Fig. 3.

Empirical model of the net flows and the incorporation of carbon (green) and nitrogen (red) in the xylem (filled arrows) and in the phloem (dashed arrows) in nitrogen‐fixing Acacia littorea and between Acacia and the hemiparasite Olax phyllanthi. For further details see Fig. 2. The figures refer to mmol C (black) or N (red) plant−1 (4 months)−1. (Modified from Tennakoon et al., 1997.) The withdrawal of C and N by direct attachment of haustoria to nodules has been omitted as a minor source for nutrients. Note: (a) the strong competition of the parasite for nitrogen and (b) the substantial shift in the incorporation of C and N in favour of the host root.

Fig. 3.

Empirical model of the net flows and the incorporation of carbon (green) and nitrogen (red) in the xylem (filled arrows) and in the phloem (dashed arrows) in nitrogen‐fixing Acacia littorea and between Acacia and the hemiparasite Olax phyllanthi. For further details see Fig. 2. The figures refer to mmol C (black) or N (red) plant−1 (4 months)−1. (Modified from Tennakoon et al., 1997.) The withdrawal of C and N by direct attachment of haustoria to nodules has been omitted as a minor source for nutrients. Note: (a) the strong competition of the parasite for nitrogen and (b) the substantial shift in the incorporation of C and N in favour of the host root.

Mechanisms of solute attraction

In 1967 Pate and co‐workers showed a flux of radiotracers from host to parasite and said ‘these results suggest that certain, as yet unspecified, properties of the hemiparasite facilitate attraction and retention of a large share of the metabolites and ions circulating within the host plant’ (Govier et al., 1967). Present knowledge is no greater, either for the xylem‐tapping hemiparasites or the phloem‐dependent holoparasites.

In the case of hemiparasites that rely on solutes from the host transpiration stream, it is clear that a higher conductance of parasite xylem and stomata sustains water flux from the host. The mechanisms by which the stomata of the parasite remain more open than those of the host are not understood, although it has been suggested that for Striga hermonthica they relate to high potassium concentrations found in the leaves of the parasite (Smith and Stewart, 1990). It would be interesting to test whether the rate of parasite transpiration per unit leaf area can be used as a predictor for dependence on contents of the host xylem.

For phloem feeders the mechanisms controlling contact with the host and the ability to sequester host solutes are also poorly understood. It is believed that in the association between Cuscuta europaea and Vicia faba, the parasite induces unloading from the phloem in the host stem, prior to uptake by the parasite (Wolswinkel et al., 1984). However, by labelling host phloem with the green fluorescent protein and the phloem‐mobile probe carboxyfluorescein, symplasmic continuity has been demonstrated between Cuscuta reflexa and its tobacco host (Haupt et al., 2001). Although these results seem incompatible, it is possible that Cuscuta uses both symplasmic and apoplasmic routes to access host solutes. As sucrose transporters have been cloned from a variety of plants (Reismeier et al., 1992; Hirose et al., 1997; Bürkle et al., 1998), it should be possible to determine whether parasite sucrose transporters are involved in moving sugars via an apoplasmic route from host to parasite, and also whether parasitism induces down‐regulation of host sucrose transporters that normally retrieve sucrose from the apoplast.

Opportunities for advances

In terms of identifying the pathways for solute transfer, the most compelling way unequivocally to mark and identify the pathways that connect host and parasite would be to use Arabidopsis, and parasites that are able to attach to it, for example, Orobanche aegyptiaca (Goldwasser et al., 2000), and Cuscuta reflexa (JM Hibberd, unpublished results). Specific cell types can be marked with the green fluorescent protein from the jellyfish Aequorea victoria, and in addition, symplasmic domains within plants can be mapped out (Imlau et al., 1999; Oparka et al., 1999). This approach has been used to demonstrate symplasmic continuity between the phloem of tobacco and the parasitic plant Cuscuta reflexa (Haupt et al., 2001), although connections to other host cells with which the parasite comes into close contact have not yet been demonstrated. This experimental approach would ascertain the cell types of the host that the parasite contacts, and whether symplasmic transport takes place. An extension of this would be to mark the xylem with a fluorophore to investigate the extent of lumenal contact between partners.

To identify and quantify fluxes, recent developments also offer the opportunity for advances. For example, sampling from sap‐sucking insects could be used, and more automated methods with higher resolution to sample xylem sap are being developed (Schurr, 1998). Mutant or transgene expressing Arabidopsis will also be used to dissect which host transporters and solutes are critical to the establishment of the host‐parasite association as well as the subsequent growth of the parasite. As long as the results gained from this approach are not extended to all host–parasites associations with unblinking faith, the near future should herald significant advances in the understanding of solute fluxes between hosts and parasitic plants.

3

To whom correspondence should be addressed. Fax: +44 1223 333953. E‐mail: julian.hibberd@plantsci.cam.ac.uk

We thank the Deutsche Forschungsgemeinschaft (SFB 251) for financial support, and JMH thanks the BBSRC for a Sir David Phillips Research Fellowship.

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