Paxillus involutus-Facilitated Cd²⁺ Influx through Plasma Membrane Ca²⁺-Permeable Channels Is Stimulated by H₂O₂ and H⁺-ATPase in Ectomycorrhizal Populus × canescens under Cadmium Stress

Abstract

Using a Non-invasive Micro-test Technique, flux profiles of Cd²⁺, Ca²⁺, and H⁺ were investigated in axenically grown cultures of two strains of Paxillus involutus (MAJ and NAU), ectomycorrhizae formed by these fungi with the woody Cd²⁺-hyperaccumulator, Populus × canescens, and non-mycorrhizal (NM) roots. The influx of Cd²⁺ increased in fungal mycelia, NM and ectomycorrhizal (EM) roots upon a 40-min shock, after short-term (ST, 24 h), or long-term (LT, 7 days) exposure to a hydroponic environment of 50 μM CdCl₂. Cd²⁺ treatments (shock, ST, and LT) decreased Ca²⁺ influx in NM and EM roots but led to an enhanced influx of Ca²⁺ in axenically grown EM cultures of the two P. involutus isolates. The susceptibility of Cd²⁺ flux to typical Ca²⁺ channel blockers (LaCl₃, GdCl₃, verapamil, and TEA) in fungal mycelia and poplar roots indicated that the Cd²⁺ entry occurred mainly through Ca²⁺-permeable channels in the plasma membrane (PM). Cd²⁺ treatment resulted in H₂O₂ production. H₂O₂ exposure accelerated the entry of Cd²⁺ and Ca²⁺ in NM and EM roots. Cd²⁺ further stimulated H⁺ pumping activity benefiting NM and EM roots to maintain an acidic environment, which favored the entry of Cd²⁺ across the PM. A scavenger of reactive oxygen species, DMTU, and an inhibitor of PM H⁺-ATPase, orthovanadate, decreased Ca²⁺ and Cd²⁺ influx in NM and EM roots, suggesting that the entry of Cd²⁺ through Ca²⁺-permeable channels is stimulated by H₂O₂ and H⁺ pumps. Compared to NM roots, EM roots exhibited higher Cd²⁺-fluxes under shock, ST, and LT Cd²⁺ treatments. We conclude that ectomycorrhizal P. × canescens roots retained a pronounced H₂O₂ production and a high H⁺-pumping activity, which activated PM Ca²⁺ channels and thus facilitated a high influx of Cd²⁺ under Cd²⁺ stress.

Introduction

The presence of highly toxic cadmium (Cd²⁺) in the environment is a serious threat to human health as heavy metals can be enriched in plants and eventually enter the human body through the food chain (Nawrot et al., 2006; Kaplan et al., 2011). The genus Populus spp. is of particular interest for phytoremediation of Cd²⁺ pollution (Sell et al., 2005; Krpata et al., 2008, 2009; Kieffer et al., 2009; He et al., 2011, 2013, 2015; Ma Y. et al., 2014), due to its widespread distribution, rapid growth, and genotypic differences in response to ion-specific stress (Chen and Polle, 2010; Polle et al., 2013; Chen et al., 2014; Polle and Chen, 2015). Populus tremula (Kieffer et al., 2009) and Populus × canescens (He et al., 2011) have been recently identified as woody Cd²⁺-hyperaccumulators. Cd²⁺ enrichment in these poplars (Kieffer et al., 2009; He et al., 2011; Ma Y. et al., 2014) exceed the threshold of 100 μg Cd²⁺ g^-1 DW that has commonly been defined for hyperaccumulation (Milner and Kochian, 2008; Krämer, 2010). He et al. (2013) demonstrated that P. × canescens could detoxify Cd²⁺ by its sequestration in the bark.

In nature, poplar roots form symbioses with mycorrhizal fungi (Danielsen et al., 2012, 2013). For example, colonization of P. × canescens roots with the ectomycorrhizal fungus Paxillus involutus improves growth, primes for increased stress tolerance, increases nutrition, and regulates the ion balance under salt stress (Schützendübel and Polle, 2002; Gafur et al., 2004; Langenfeld-Heyser et al., 2007; Luo et al., 2009, 2011; Li J. et al., 2012; Ma X. et al., 2014). A notable finding was that Paxillus involutus ectomycorrhizas enhance both Cd²⁺ uptake and tolerance in P. × canescens (Ma Y. et al., 2014). Thus, ectomycorrhizal poplar plants offer a great potential for phytoremediation of Cd²⁺-polluted soils (Sell et al., 2005; Krpata et al., 2008, 2009; Luo et al., 2014; Ma Y. et al., 2014).

Cd²⁺ is generally believed to enter plant cells through high affinity transporters responsible for the uptake of divalent cations (Cu²⁺, Co²⁺, Fe²⁺, Ca²⁺, Mn²⁺, and Zn²⁺; Liu et al., 1997; Clemens et al., 1998; Cohen et al., 1998; Hirschi et al., 2000; Thomine et al., 2000; Zhao et al., 2002; Cosio et al., 2004; Clemens, 2006; Roth et al., 2006). Cd²⁺ can even induce nutrient deficiencies by competing with the uptake of essential elements (Zhao et al., 2006; Papoyan et al., 2007; DalCorso et al., 2008; Gallego et al., 2012; Baliardini et al., 2015). On the other hand, elevated Ca²⁺ levels suppress Cd²⁺ uptake in different ecotypes of Sedum alfredii also supporting competition of Cd²⁺ uptake with nutrient cations (Lu et al., 2010). Transcript levels of the transporters involved in Cd²⁺ uptake and transport have been investigated in herbaceous and woody species (Kim et al., 2006; Plaza et al., 2007; Krämer, 2010; Migeon et al., 2010; Mendoza-Cózatl et al., 2011; Lin and Aarts, 2012). In poplar plants, a variety of heavy metal transporters, such as ZRT-IRT-like proteins (ZIP2, ZIP6.2), natural resistance associated macrophage proteins (NRAMP1.1, NRAMP1.3), ATP-binding cassette transporter C1 (ABCC1), heavy metal ATPase 4 (HMA4), ATP-binding cassette transporter in mitochondria (ATM3), have been suggested to play pivotal roles in Cd²⁺ transport and detoxification (Ma Y. et al., 2014; He et al., 2015). In addition to these heavy metal transporters, ion channels in the plasma membrane (PM) that are permeable to Cd²⁺ contribute the Cd²⁺ uptake (Li et al., 2012a; Sun et al., 2013a,b; He et al., 2015). High external Cd²⁺ concentrations establish a large electrochemical gradient facilitating the rapid movement of Cd²⁺ ions through Cd²⁺-permeable channels. Perfus-Barbeoch et al. (2002) suggested that Cd²⁺ enters root cells via plasma membrane (PM) Ca²⁺ channels.

Ca²⁺ channels in the PM have been characterized by electrophysiological measurements involving incorporation of plasma-membrane vesicles into planar lipid bilayers (PLB, White, 2000) and patch clamping (Perfus-Barbeoch et al., 2002). According to their electrophysiological properties, the channels can be divided into depolarisation-, hyperpolarisation-, elicitor-activated, and voltage-insensitive channels (Thuleau et al., 1998; White, 2000). These channels display different sensitivities to typical inhibitors of Ca²⁺ channels, such as La³⁺, Gd³⁺, TEA, and verapamil. Specifically, verapamil and TEA inhibit depolarisation-activated Ca²⁺ channels, such as the wheat root channel rca (Piñeros and Tester, 1997; White, 1998), and rye root voltage-dependent cation channel 2, VDCC2 (White, 1998). La³⁺ shares a high similarity to another trivalent cation, Gd³⁺. Both cations are able to inhibit three distinct classes of Ca²⁺ channels, including depolarisation-activated Ca²⁺ channels, rca (Piñeros and Tester, 1997; White, 1998), hyperpolarisation-activated Ca²⁺ channels (HACCs) in onion bulb epidermis (Pickard and Ding, 1993), voltage-insensitive channels such as Arabidopsis root epidermal non-selective cation channels (NSCCs; Demidchik et al., 2002), and large-conductance elicitor-activated channel (LEAC) in parsley cell suspension (Zimmermann et al., 1997). Ca²⁺ channels in the PM are permeable to divalent (including Ca²⁺, Mg²⁺, Ba²⁺, Sr²⁺, Co²⁺, Zn²⁺, Mn²⁺, Ni²⁺, Cu²⁺; Cosgrove and Hedrich, 1991; Ping et al., 1992; Pickard and Ding, 1993; Thuleau et al., 1994a,b; Gelli and Blumwald, 1997; Zimmermann et al., 1997; White, 1998; Grabov and Blatt, 1998, 1999) and monovalent cations (Na⁺, K⁺, Cs⁺, Li⁺, Rb⁺; Cosgrove and Hedrich, 1991; Pickard and Ding, 1993; Zimmermann et al., 1997; Piñeros and Tester, 1997; White, 1998). In accordance with the suggestion that Cd²⁺ ions can be transported into cells through Ca²⁺ channels (Perfus-Barbeoch et al., 2002; Gallego et al., 2012; Li et al., 2012b) the permeability for Cd²⁺ through wheat VDCC2 was detected when the plasma membrane derived from root cells was incorporated into PLB (White, 1998). Using the whole-cell patch-clamp technique, Perfus-Barbeoch et al. (2002) confirmed that Cd²⁺ permeates through the PM Ca²⁺ channels in Arabidopsis guard cells. The Cd²⁺ influx was effectively blocked by Ca²⁺ channel blockers, e.g., LaCl₃ and verapamil in Suaeda salsa (Li et al., 2012a), Populus euphratica (Sun et al., 2013b), and P. tremula × P. alba (He et al., 2015), further indicating that Cd²⁺ ions penetrate into plant cells through Ca²⁺-permeable channels.

It is possible that hydrogen peroxide (H₂O₂) stimulates the entry of Cd²⁺ through PM Ca²⁺ channels as the activity of these channels has been shown to be stimulated by H₂O₂. Pei et al. (2000) found that H₂O₂ activates the PM Ca²⁺ channels, leading to a subsequent rise of cytosolic Ca²⁺ in Arabidopsis guard cells. Demidchik et al. (2007) observed a transient increase of Ca²⁺ influx in the root epidermis when exogenous H₂O₂ was applied to Arabidopsis thaliana. In NaCl-stressed P. euphratica cells, Ca²⁺ influx through Ca²⁺ channels was activated by H₂O₂ (Sun et al., 2010). Recently, H₂O₂ was shown to accelerate Cd²⁺ influx in P. euphratica cells, while the H₂O₂-stimulated Cd²⁺ influx was blocked by LaCl₃ (Sun et al., 2013b; Han et al., 2016). Moreover, the application of a H₂O₂ scavenger, catalase, lowered the Cd²⁺ influx across the PM in Cd²⁺-stressed P. euphratica cells (Sun et al., 2013b). In Cd²⁺-treated P. euphratica cells, hydrogen sulfide was found to reduce Cd²⁺ influx through down-regulation of H₂O₂-stimulated Cd²⁺ transport across the PM Ca²⁺ channels (Sun et al., 2013b). H₂O₂ is not only produced in Cd²⁺-stressed poplar cells (Sun et al., 2013b; Han et al., 2016) and roots (Ma Y. et al., 2014; He et al., 2015), but is also massively enriched in Populus × canescens–Paxillus involutus ectomycorrhizal associations (Gafur et al., 2004; Langenfeld-Heyser et al., 2007). Thus, it can be speculated that the fungal-elicited H₂O₂ accelerates the entry of Cd²⁺ through PM Ca²⁺ channels. However, this hypothesis needs to be clarified by further electrophysiological investigations.

In addition to H₂O₂, the PM H⁺-ATPase plays a crucial role in accelerating Cd²⁺ transport in poplar roots (Ma Y. et al., 2014; He et al., 2015). He et al. (2015) demonstrated that the net Cd²⁺ influx was pH-dependent in poplar roots and effectively blocked by inhibitors of H⁺-pumps. Ma Y. et al. (2014) showed that the active PM H⁺-ATPase-driven Cd²⁺ uptake is a major factor for increased Cd²⁺ accumulation in ectomycorrhizal (EM) poplar plants. They suggested that the EM-induced transcripts of HA2.1 and AHA10.1 genes, encoding PM H⁺-ATPases in P. × canescens, may result in H⁺-pump-stimulated Cd²⁺ enrichment (Ma Y. et al., 2014). In agreement with this suggestion transgenic poplars that were more Cd²⁺ tolerant by overexpression of γ-glutamylcysteine synthetase, showed upregulated transcript levels of VHA1.1, HA2.1 and AHA10.1 and a high Cd²⁺ uptake rate (He et al., 2015). The PM H⁺-ATPases maintain a H⁺ gradient across the membrane to promote active transport of essential elements across the PM (Beritognolo et al., 2007; Ma et al., 2010; Sun et al., 2010; Luo et al., 2013). Increased H⁺-pumping activities have been well characterized in arbuscular mycorrhizal associations (Ramos et al., 2005; Rosewarne et al., 2007) and in ectomycorrhizal associations formed by Paxillus involutus (strains MAJ and NAU) with Populus × canescens (Li J. et al., 2012). We have previously shown that the upregulated H⁺-pumping activities in Paxillus involutus-Populus × canescens symbiosis resulted in enhanced Ca²⁺ uptake and enrichment (Li J. et al., 2012). Demidchik et al. (2002) proposed that voltage modulation of the co-existing NSCC/HACC by PM H⁺-ATPase would be a potent regulator for Ca²⁺ entry to the root cell cytoplasm. The high H⁺-pumping activity leads to hyperpolarization of the PM and, thus, may increase Cd²⁺ influx through hyperpolarisation-activated Ca²⁺ channels. However, it is unknown whether the PM H⁺-ATPases could stimulate the entry of Cd²⁺ through Ca²⁺-permeable channels in ectomycorrhizal plants.

The two P. involutus strains, MAJ and NAU, form different colonization structures with P. × canescens roots (Gafur et al., 2004). Strain MAJ forms a typical hyphal mantle and Hartig net with roots of P. × canescens, while NAU is unable to intrude between the host cells and forms only a hyphal mantle ensheathing the root tips (Gafur et al., 2004). The colonization of P. × canescens roots with the competent strain MAJ results in enriched Cd²⁺ levels under Cd²⁺ stress (Ma Y. et al., 2014). Whether the incompatible fungal isolate NAU also affects the Cd²⁺ entry into P. × canescens host plants needs to be clarified.

In this study, we used a non-invasive micro-test technique (NMT) to measure fluxes of Cd²⁺, Ca²⁺ and H⁺ in Cd²⁺-stressed roots of non-mycorrhizal (NM) and ectomycorrhizal P. × canescens plants colonized with Paxillus involutus strains, MAJ and NAU. The aim was to elucidate whether the Cd²⁺ influx through Ca²⁺-permeable channels is stimulated by H₂O₂ and H⁺-ATPase in ectomycorrhizal roots since the ectomycorrhizas exhibit enhanced H₂O₂ production and upregulated H⁺-pumping activity. NMT microelectrodes measure the ion fluxes on the surface of the tissues, which are either the plant root cells for the NM plants or the fungal hyphae forming the mantle structure ensheathing the roots. To discriminate between potentially different Cd²⁺ effects on fungus and plant roots, fluxes of Cd²⁺, Ca²⁺ and H⁺ were examined for pure fungal mycelia of the two P. involutus isolates, MAJ and NAU, in addition to flux recordings on NM and EM roots. Furthermore, flux profiles of Cd²⁺ and Ca²⁺ were recorded in P. involutus-inoculated roots after 7 days of co-culture. The aim was to determine whether flux profiles of mature EM associations resemble the pattern of those from host roots at early stages of fungal colonization when the host is known to activate transient defense responses in contrast to the mature ectomycorrhizal symbioses (Duplessis et al., 2005).

Materials and Methods

Fungus and Plant Cultures for EM Colonization

The Paxillus involutus isolates MAJ and NAU, obtained from the Büsgen Institute: Institute of Forest Botany and Tree Physiology (Göttingen University, Germany), were grown on 2% modified Melin Norkrans (MMN) agar medium (g⋅L^-1): KH₂PO₄ 0.5, (NH₄)₂SO₄ 0.25, MgSO₄⋅7H₂O 0.15, CaCl₂⋅2H₂O 0.05, NaCl 0.025, FeCl₃⋅6H₂O 0.01, thiamine HCl 0.0001, glucose 10, malt extract 3, pH 5.2 (Gafur et al., 2004; Li J. et al., 2012). Prior to the colonization, the fungi were pre-grown on the agar culture medium for 1 week in petri dishes (diameter 90 mm) and kept in darkness at 23°C.

Plantlets of Populus × canescens (a hybrid of Populus tremula × Populus alba) were propagated by micropropagation as described by Leple et al. (1992). Regenerated P. × canescens plants were grown for 3–4 weeks on Murashige and Skoog (MS) medium (Murashige and Skoog, 1962). Uniform plants with sufficient roots were used for ectomycorrhization. The colonization of P. × canescens with Paxillus involutus strains MAJ and NAU was followed the procedures described by Gafur et al. (2004). In brief, rooted plantlets from sterile culture were placed on the MMN agar medium in the presence or absence of EM mycelium. After fungal inoculation, the petri dishes were sealed with Parafilm and covered with aluminum foil to keep the roots in darkness. During the period of incubation, the temperature in the climate chamber was maintained at 23°C with a light period of 16 h (6:00 AM–22:00 PM). Photosynthetic active radiation (PAR) of 200 μmol m^-2 s^-1 was supplied by cool white fluorescent lamps. After 1 month of inoculation, EM and NM root tips for anatomical investigations were embedded, stained, and photographed as described previously (Gafur et al., 2004). EM and NM plants with similar height and growth performance were used for CdCl₂ treatment.

Liquid Culture of Fungi

Liquid culture of P. involutus was grown as previously described (Ott et al., 2002; Langenfeld-Heyser et al., 2007; Li J. et al., 2012). In brief, mycelium from the agar plate was homogenized, transferred into 100 mL of liquid medium (pH 4.8) in flasks, and incubated on a rotary shaker in darkness (150 rpm, 23°C). P. involutus in submerged culture grew in the form of compact spherical masses of mycelium (pellets). For Cd²⁺ shock treatment, sterile filtered CdCl₂ solutions were added to achieve final concentrations of 50 μM. After ST (24 h) or LT (7 days) treatment, axenic cultures of MAJ and NAU were used for steady flux measurements of Cd²⁺, H⁺, and Ca²⁺.

Cadmium Treatment

Ectomycorrhizal and non-mycorrhizal plants were carefully removed from MMN agar medium. Rooted plantlets were cultivated in individual pots containing hydroponic MS nutrient solution (MS medium without agar and sucrose) (Murashige and Skoog, 1962). Plants were covered with plastic bags to reduce the rapid water loss in a growth room. NM and EM plantlets were subjected to 50 μM CdCl₂ for a short-term (ST) exposure, 24 h or a long-term (LT) exposure for 7 days. The required amount of CdCl₂ was added to the MS nutrient solution. Control plants were treated in the same manner without the addition of CdCl₂. The plants were maintained at 23°C with a light period of 16 h (6:00 AM–22:00 PM) and PAR was 200 μmol m^-2 s^-1. Plants were continuously aerated by passing air to hydroponic MS nutrient solution, which was regularly renewed. Steady fluxes of Cd²⁺, Ca²⁺ and H⁺ in NM and EM roots were examined after 24 h and 7 days of CdCl₂ treatment. In addition, ST-induced alterations of Cd²⁺ and Ca²⁺ fluxes were also examined in non-inoculated and P. involutus-inoculated roots after 7 days of co-culture.

Measurements of Net Cd²⁺, Ca²⁺, and H⁺ Fluxes

Preparations of Ion-Selective Microelectrodes

Non-invasive Micro-test Technique (NMT-YG-100, Younger USA LLC, Amherst, MA01002, USA) with ASET 2.0 (Sciencewares, Falmouth, MA 02540, USA) and iFluxes 1.0 Software (Younger USA, LLC, Amherst, MA 01002, USA) was used to monitor fluxes of Cd²⁺, Ca²⁺ and H⁺ in EM and NM roots (Sun et al., 2009a,b; Sun et al., 2013a,b; Ma X. et al., 2014). Ion-selective electrodes were prepared as described in Sun et al. (2009a, 2013a) and Ma X. et al. (2014). Briefly, pre-pulled and silanized glass micropipettes (diameter 4–5 μm, XY-DJ-01; Xuyue (Beijing) Science and Technology Co. Ltd., Beijing, China) were back-filled with backfilling solution [Cd²⁺ microelectrodes: 10 mM Cd(NO₃)₂ and 0.1 mM KCl; Ca²⁺ microelectrodes: 100 mM CaCl₂; H⁺ microelectrodes: 40 mM KH₂PO₄ and 15 mM NaCl, pH 7.0] to a length of 1.0 cm from the tip. Then the micropipettes were front-filled with 15 μm columns of selective liquid ion exchange cocktails (LIXs) (Cd: Fluka 20909, Sigma–Aldrich, St Louis, MO, USA; Ca: Fluka 21048; H: Fluka 95293 Fluka Chemie GmbH, Buchs, Switzerland). An Ag/AgCl wire electrode holder (XYEH01-1; Xuyue Sci. and Tech. Co., Ltd.) was inserted in the back of the electrode to create an electrical contact with the electrolyte solution. DRIREF-2 (World Precision Instruments, Inc., Sarasota, FL, USA) was used as the reference electrode (CMC-4). Prior to the measurements, ion-selective microelectrodes for the target ions were calibrated by the following standard solution:

• simple (1)

  Cd²⁺: 0.01, 0.05, 0.1 mM (Cd²⁺ concentration was 0.05 mM in the measuring solution);

• simple (2)

  Ca²⁺: 0.1, 0.5, 1.0 mM (Ca²⁺ was 0.2 mM in the measuring buffer);

• simple (3)

  H⁺: pH 4.2, 5.2, 6.2 (pH of the measuring solution was adjusted to 5.2 with KOH and HCl for root samples).

Electrodes were used when the Nernstian slopes in ranges of 29 ± 3 mV/decade (Cd²⁺, Ca²⁺) and 58 ± 5 mV/decade (H⁺). The flux rate was calculated on the basis of Fick’s law of diffusion:

J = -D (dc/dx),

where J is the ion flux in the x direction, D is the ion diffusion coefficient in a particular medium, dc represents the ion concentration difference, dx is the microelectrode movement between two positions, and dc/dx represents the ion concentration gradient. As part of the NMT system, ASET software [Science Wares (East Falmouth, MA, USA) and Applicable Electronics], was used for data and image acquisition, preliminary processing, control of three-dimensional electrode positioner and stepper-motor-controlled fine focus of the microscope stage.

Experimental Protocols for Steady-State Flux Measurements

Cd²⁺, Ca²⁺, and H⁺ fluxes were non-invasively measured by moving the ion-selective microelectrode between two positions close to the materials in a preset excursion (30 μm for excised roots and fungal mycelia) at a programmable frequency in the range of 0.3–0.5 Hz. P. involutus mycelia, EM and NM roots from the ST and LT CdCl₂ treatments were rinsed with re-distilled water for 2–3 times, and then incubated in the basic measuring solution to equilibrate for 25 min. The concentration gradients of Cd²⁺, Ca²⁺, and H⁺ were measured as previously described (Li J. et al., 2012; Lu et al., 2013; Sun et al., 2013a,b).

• simple (1)

  Cd²⁺ measuring solutions: 0.1 mM KCl, 0.1 mM MgCl₂, 0.05 mM CaCl₂ and 0.05 mM CdCl_2, pH was adjusted to 5.2 with KOH and HCl;

• simple (2)

  Ca²⁺ measuring solutions: 0.1 mM NaCl, 0.1 mM MgCl₂, 0.1 mM KCl, and 0.2 mM CaCl₂, pH was adjusted to 5.2 with KOH and HCl;

• simple (3)

  H⁺ measuring solutions: 0.1 mM NaCl, 0.1 mM MgCl₂, 0.1 mM CaCl₂ and 0.5 mM KCl, pH 5.2 was adjusted with KOH and HCl.

The steady fluxes of roots were then recorded 100 μm from the apex and conducted along the root axis until 2300 μm at intervals of 200–300 μm. The fluxes of each measuring point in apical regions were continuously recorded for 6–8 min. For P. involutus mycelia, Cd²⁺, Ca²⁺, and H⁺ fluxes were measured around the surface of pelleted hyphae over a recording period of 30 min.

Transient Flux Recording

Paxillus involutus fungal mycelia and roots sampled from EM and NM plants were immobilized in the measuring solutions of Cd²⁺ (0.1 mM KCl, 0.1 mM MgCl₂, 0.05 mM CaCl₂, pH 5.2); Ca²⁺ (0.1 mM NaCl, 0.1 mM MgCl₂, 0.1 mM KCl, and 0.2 mM CaCl₂, pH 5.2) and H⁺ (0.1 mM NaCl, 0.1 mM MgCl₂, 0.1 mM CaCl₂ and 0.5 mM KCl, pH 5.2) for 25 min equilibration. Then the steady-state fluxes in fungal mycelia and the root apical region (100 μm from the root apex) were continuously recorded for 5 min prior to the CdCl₂ shock. CdCl₂ stock (100 μM) was slowly added to the measuring solution using a pipette until the final Cd²⁺ concentration reached 50 μM. Afterward, transient kinetics of Cd²⁺, Ca²⁺, and H⁺ were restarted and continued for 40 min. The data measured during the first 1–2 min was discarded, due to the effects of the diffusing stock solution. The high flux of Cd²⁺, Ca²⁺, and H⁺ during the following 2 min was defined as peaking values.

Effects of H₂O₂ on CdCl₂-altered transient kinetics of Cd²⁺ and Ca²⁺ were also examined in NM and EM roots. Following the CdCl₂ shock (50 μM) as described above, H₂O₂ (1.0 mM) was introduced to the measuring solution and transient kinetics of Cd²⁺ and Ca²⁺ were recorded for 20 min.

Fungal mycelia were exposed to 50 μM CdCl₂ to induce a shock. Cd²⁺, Ca²⁺, and H⁺ fluxes were monitored over a continuous recording period of 40 min. For transient flux kinetics, the data measured during the first 1–2 min were discarded due to the diffusion effects of stock addition.

Effects of Ca²⁺ on Sensitivity of Cd²⁺ Electrodes

To determine whether Ca²⁺ ions compete with Cd²⁺ to penetrate across PM Ca²⁺-permeable channels, the effects of additional Ca²⁺ ions on Cd²⁺ electrodes was examined. Cd²⁺ calibrating solutions were added with 0, 0.01, 0.025, 0.05, 0.1, 0.2, 0.5, 1.0, or 2.0 mM Ca²⁺. Then Cd²⁺ microelectrodes were calibrated in Ca²⁺-supplemented solutions as described above. Moreover, the Nernst slope and intercept of the Cd²⁺ electrodes were calibrated in the measuring solution containing 0.1 mM KCl, 0.1 mM MgCl₂, and 0.05 mM CaCl₂.

Flux Oscillations

Oscillations in membrane-transport activity are ubiquitous in plant response to salinity, temperature, osmotic, hypoxia, and pH stresses (Shabala et al., 2006). In our study, rhythmic (ultradian) flux oscillations in NM and EM P. × canescens roots were not noticeable as that observed in herbaceous species (Shabala et al., 1997, 2003, 2006; Shabala and Knowles, 2002). This finding is presumably due to a lower growth rate of woody roots compared with crop species (Li J. et al., 2012). The flux oscillations of the measured ions, e.g., H⁺, Ca²⁺, and Cd²⁺, were more like fluctuations as previously reported in poplar roots (e.g., Na⁺, K⁺, H⁺, and Ca²⁺; Li J. et al., 2012). In this study, H⁺, Ca²⁺, and Cd²⁺ fluxes were recorded for 6–8 min at each point, which is long enough to cover oscillatory periods of measured ions.

Inhibitor and Stimulator Treatment

In this study, the effects of Ca²⁺, pH, H₂O₂, and PM transporter and channel inhibitors on Cd²⁺-altered ion flux profiles were examined in fungal mycelia and roots (NM and EM). Briefly,

Series 1: Ca²⁺ channel inhibitors. NM and EM roots were pre-treated with or without LaCl₃ (5 mM; Sun et al., 2010; Li et al., 2012b), GdCl₃ (500 μM, Demidchik et al., 2007, 2009; Sun et al., 2012), TEA (50 μM, White, 1998; Li J. et al., 2012), or verapamil (20 μM, Li et al., 2012a; He et al., 2015) for 24 h in the presence and absence of 50 μM CdCl₂. Fungal mycelia of the two P. involutus isolates, MAJ and NAU, were subjected to 0 or 5 mM LaCl₃ treatment for 24 h supplemented with or without 50 μM CdCl₂.

Series 2: Ca²⁺. After being subjected to Cd²⁺ stress (CdCl₂, 50 μM) for 24 h, NM and EM roots were then exposed to 25, 50, or 100 μM CaCl₂ for flux recordings in the presence of CdCl₂.

Cd²⁺ and Ca²⁺ fluxes in Series 1 and 2 were measured along root axes, 100–2,300 μm from the apex, at intervals of 200–300 μm. In P. involutus mycelia, Cd²⁺ and Ca²⁺ fluxes were continuously measured around the surface of pelleted hyphae over a recording period of 30 min.

Series 3: Hydrogen peroxide. NM and EM roots were sampled and immobilized in Cd²⁺ or Ca²⁺ measuring solutions for transient flux recordings in the apical region (100 μm from the root apex). The steady-state fluxes were continuously recorded for 10–20 min prior to the CdCl₂ shock. CdCl₂ stock (100 μM) was slowly added to the measuring solution until the final Cd²⁺ concentration reached 50 μM and transient kinetics of Cd²⁺ and Ca²⁺ were continuously for 20–30 min. Afterward, H₂O₂ (1.0 mM) was slowly added to the measuring solution and transient kinetics of Cd²⁺ and Ca²⁺ were restarted and continued for 20 min.

Series 4: ROS scavenger. NM and EM roots were pre-treated with or without 1, 3-Dimethyl-2-thiourea (DMTU, 5 mM, Chung et al., 2008; Sun et al., 2010) for 24 h in the presence and absence of 50 μM CdCl₂. Then Cd²⁺ and Ca²⁺ fluxes were measured along root axes, 100–2,300 μm from the apex, at intervals of 200–300 μm.

Series 5: External pH. NM and EM roots were pre-treated with 50 μM CdCl₂ for 24 h prior to flux measurements. Cd²⁺ and Ca²⁺ fluxes along root axes (100–2,300 μm from the apex) were recorded in Cd²⁺ or Ca²⁺ measuring solutions at pH 5.2, 6.2, or 7.2, respectively.

Series 6: PM H⁺-ATPase inhibitor. NM and EM roots were pre-treated with or without sodium orthovanadate (500 μM, Sun et al., 2010; Lu et al., 2013) for 24 h in the presence and absence of 50 μM CdCl₂. Then H⁺, Cd²⁺, and Ca²⁺ fluxes were measured along root axes, 100–2,300 μm from the apex, at intervals of 200–300 μm. P. involutus isolates, MAJ and NAU, were exposed to 0 or 500 μM sodium orthovanadate for 24 h prior to a 30-min of continuous recording of H⁺ flux.

Measurements of Net H₂O₂ Fluxes

An H₂O₂-sensititive microelectrode [tip diameter 2–3 μm, XY-DJ-502, Xuyue (Beijing) Science and Technology Co. Ltd., Beijing, China] was used to monitor H₂O₂ fluxes in EM and NM roots. H₂O₂ microelectrodes were prepared according to the method described by Twig et al. (2001). Before the measurement, H₂O₂ microelectrode was polarized at +0.60 V against an Ag/AgCl reference electrode. Thereafter, the microelectrodes were calibrated by the standard solution: 0.01, 0.1 and 1 mM H₂O₂. Roots sampled from control and CdCl₂ (50 μM,30 min)-treated EM and NM plants were immobilized in the measuring solution (0.1 mM NaCl, 0.1 mM MgCl₂, 0.1 mM CaCl₂ and 0.5 mM KCl, pH was adjusted to 5.2 with KOH and HCl) and equilibrated for 25 min. The fluxes were recorded 100 μm from the apex and conducted along the root axis until 2300 μm, at intervals of 200–300 μm, and then calculated.

Data Analysis

Ionic fluxes were calculated using the program JCal V3.2.1, a free MS Excel spreadsheet, which was developed by the Yue Xu¹. The experimental data were subjected to SPSS (SPSS Statistics 17.0, 2008) for statistical tests and analyses. Unless otherwise stated, P < 0.05 was considered as significant.

Results

Cd²⁺-Altered Ion Flux Profiles in Paxillus involutus, and Roots of NM and EM Poplar

Cd²⁺ Fluxes

We recorded transient Cd²⁺ kinetics upon Cd²⁺ shock at the root apex (100 μm from the root tip; Figure 1A), where a vigorous ion flux (e.g., Na⁺, K⁺, Ca²⁺, Cd²⁺, Cl^-) is usually observed in woody and herbaceous plants (Sun et al., 2009a,b; Li J. et al., 2012; Lu et al., 2013; Han et al., 2014). The addition of CdCl₂ (50 μM) caused an immediate Cd²⁺ influx in both EM and NM roots which declined with increasing duration of Cd²⁺ exposure (40 min; Figure 1A). The peak and mean flux rate of Cd²⁺ in EM roots with MAJ were significantly (13.5 and 38.8%) higher than in NM roots or NAU-colonized roots (Figure 1A). Similar to the Cd²⁺ kinetics in EM roots, an instantaneous increase in the Cd²⁺ influx was detected in pure P. involutus mycelia after CdCl₂ exposure (50 μM; Figure 1A). However, the fungal Cd²⁺ influx remained constant over the recording period (40 min; Figure 1A) with significantly higher flux rates in MAJ (75.4 pmol cm^-2 s^-1) than in NAU (25.9 pmol cm^-2 s^-1).

FIGURE 1.

Effects of CdCl₂ on transient kinetics of Cd²⁺, Ca²⁺, and H⁺ in Populus × canescens roots and Paxillus involutus strains MAJ and NAU. Cd²⁺ (A), Ca²⁺ (B), and H⁺ (C) kinetics were recorded before and after the required amount of 50 μM CdCl₂ was introduced into the measuring chamber. Prior to the CdCl₂ shock, steady-state fluxes of Cd²⁺, Ca²⁺, and H⁺ in ectomycorrhizal (MAJ and NAU) and non-mycorrhizal (NM) P. × canescens roots (measuring site was ca. 100 μm from the root tip) and P. involutus isolates were monitored for approximately 5 min. Transient kinetics of Cd²⁺, Ca²⁺, and H⁺ were recorded after the required amount of 50 μM CdCl₂ was introduced into the measuring solution. Inserted sections show the peaking and/or mean values of Cd²⁺, Ca²⁺, and H⁺ flux before (-Cd) and after (+Cd) the addition of CdCl₂. Columns represent the mean of four to five individual plants or axenic EM cultures (pelleted hyphae), and bars represent the standard error of the mean. Different letters, a, b, c, d, e, and f, indicate significant difference at P < 0.05 between treatments.

After ST (24 h) or LT (7 days) exposure to 50 μM CdCl₂ in hydroponic conditions, steady-state Cd²⁺ flux was recorded along root axis (100–2,300 μm from the apex) at intervals of 200–300 μm (Figure 2). In NM roots, ST and LT stress caused a net Cd²⁺ influx with an overall mean of 28.9 pmol cm^-2 s^-1 along the whole measured distance; LT treatment resulted in a higher flux rate at the region 100–1,000 μm from the apex than at more distant root positions (Figure 2). A similar trend was observed in the Cd²⁺-stressed EM roots, though mean Cd²⁺ fluxes in MAJ- and NAU-ectomycorrhizal roots were 43.1 and 32.0% higher than those of the NM roots under ST and LT stress (Figure 2). The mycelia of the two P. involutus strains, MAJ and NAU, exhibited a stable Cd²⁺ influx under ST and LT stress, although the CdCl₂-induced Cd²⁺ influx was typically higher under LT conditions, 68.9 pmol cm^-2 s^-1, compared with ST treatment, 27.1 pmol cm^-2 s^-1 (Figure 2). Cd²⁺-induced alterations of Cd²⁺ flux were also examined in non-inoculated and P. involutus-inoculated roots after 7 days of co-culture. NAU- and MAJ-colonized roots showed larger flux rates than non-inoculated roots after ST Cd²⁺ stress (Supplementary Figure S1A).

FIGURE 2.

Effects of CdCl₂ on steady Cd²⁺ fluxes in Populus × canescens roots and Paxillus involutus strains MAJ and NAU. P. involutus isolates, ectomycorrhizal (MAJ and NAU) and non-mycorrhizal (NM) P. × canescens plants were subjected to short-term (ST, 24 h) and long-term (LT, 7 d) exposure to 50 μM CdCl₂, respectively. Control roots and axenic mycelia were well fertilized but treated without CdCl₂. Cd²⁺ fluxes in poplar roots were measured along root axis, 100–2,300 μm from the apex, at intervals of 200–300 μm. Cd²⁺ fluxes of P. involutus isolates MAJ and NAU were measured along the surface of pelleted hyphae over a recording period of 30 min. Inserted sections show the Cd²⁺ fluxes in P. involutus isolates after short-term (ST, 24 h) or long-term (LT, 7 days) CdCl₂ treatment. Each point is the mean of 4–5 individual plants or axenic EM cultures (pelleted hyphae), and bars represent the standard error of the mean. Asterisks denote significant difference at P < 0.05 between treatments.

Our data show that P. involutus mycelia and EM roots both exhibited an enhanced Cd²⁺ uptake upon Cd²⁺ shock, ST, or LT treatment (Figures 1A and 2). Unexpectedly, the Cd²⁺ influx in EM roots did not show a high correlation to the flux rate of Cd²⁺ in fungal hyphae under various treatments (shock, ST, or LT, Supplementary Figure S2). However, a relatively high correlation between EM and NM roots was observed especially in response to Cd²⁺ shock (Supplementary Figure S2). This result supports that in the ectomycorrhizal symbioses the continuous Cd²⁺ entry detected by NMT microelectrodes depends on the uptake capacity of inner root cells and that in the plant–fungal interaction divergent regulation of fungal Cd²⁺ transport compared with pure mycelium must take place.

Ca²⁺ Fluxes

In the absence of Cd²⁺ stress, poplar roots exhibited a net Ca²⁺ influx, with a greater flux rate in MAJ- and NAU-ectomycorrhizal roots, 26.9 pmol cm^-2 s^-1, than in NM roots, 9.6 pmol cm^-2 s^-1 (Figure 1B). Similarly, the mycelia of the two strains exhibited a stable and steady influx of Ca²⁺ (162.3 pmol cm^-2 s^-1), which is ca. 6.0-fold higher than that detected in EM roots (Figure 1B). CdCl₂ shock (50 μM) caused a transient Ca²⁺ efflux in NM and EM roots with maximum values ranging from 10.9 to 14.8 pmol cm^-2 s^-1 (Figure 1B). Thereafter, the direction shifted toward an influx and the mean flux over the recording period then declined in EM roots, or displayed a net efflux in NM roots (Figure 1B). In contrast to NM and EM roots, Cd²⁺ addition markedly increased the Ca²⁺ influx in the hyphae of pure mycelium, typically with higher flux rates in strain NAU than in MAJ in the first 20 min of Cd²⁺ application (Figure 1B). Under ST and LT treatment, Cd²⁺ stress caused a marked decline of Ca²⁺ influx along the root axis (Figure 3). MAJ- and NAU-ectomycorrhizal roots maintained 40.5 and 20.6% higher Ca²⁺ fluxes than NM roots under ST and LT stress (Figure 3). In the hyphae of the two fungal strains, the Ca²⁺ influx was enhanced by ST and LT treatments (Figure 3), similar to the shock treatment (Figure 1B). We observed that the flux rate in the two strains declined with increasing duration of hydroponic culture regardless of control and Cd²⁺ treatments (Figure 3). Non-inoculated P. × canescens roots exhibited a net Ca²⁺ influx under unstressed control conditions and the Ca²⁺ influx was stimulated by 7 days inoculation with MAJ and NAU (Supplementary Figure S1B). ST-treated P. involutus-inoculated roots retained higher Ca²⁺ influx than non-inoculated roots although the Ca²⁺ influx in poplar roots was lowered by Cd²⁺ stress (Supplementary Figure S1B).

FIGURE 3.

Effects of CdCl₂ on steady Ca²⁺ fluxes in Populus × canescens roots and Paxillus involutus strains MAJ and NAU. P. involutus isolates, ectomycorrhizal (MAJ and NAU) and non-mycorrhizal (NM) P. × canescens plants were subjected to short-term (ST, 24 h) and long-term (LT, 7 days) exposure to 50 μM CdCl₂, respectively. Control roots and axenic mycelia were well fertilized but treated without CdCl₂. Ca²⁺ fluxes in poplar roots were measured along root axis, 100–2,300 μm from the apex, at intervals of 200–300 μm. Ca²⁺ fluxes of P. involutus isolates MAJ and NAU were measured along the surface of pelleted hyphae over a recording period of 30 min. Inserted sections show the Ca²⁺ fluxes in P. involutus isolates after short-term (ST, 24 h) or long-term (LT, 7 days) CdCl₂ treatment. Each point is the mean of 4–5 individual plants or axenic EM cultures (pelleted hyphae), and bars represent the standard error of the mean. Asterisks denote significant difference at P < 0.05 between treatments.

It has been suggested that the Ca²⁺ enrichment in EM roots was associated with the P. involutus fungal hyphae exhibiting a high capacity for Ca²⁺ uptake (Figures 1B and 3; Li J. et al., 2012; Ma X. et al., 2014). However, the Ca²⁺ influx in EM roots was not evidently correlated to the flux rate of Ca²⁺ in fungal hyphae under Cd²⁺ shock, ST, or LT (Supplementary Figure S3). Unexpectedly, the Ca²⁺ flux in EM roots was even negatively correlated to the flux rate of Ca²⁺ in fungal hyphae after a shock treatment (Supplementary Figure S3). The observed correlation of Ca²⁺ fluxes between EM roots and NM roots (Supplementary Figure S3) supports that the Ca²⁺ flow was mainly the consequence of host roots in the Cd²⁺-stressed ectomycorrhizal symbioses.

Correlations between Cd²⁺ and Ca²⁺ Fluxes

We analyzed the correlation between Cd²⁺ and Ca²⁺ fluxes as NM and EM roots took up these elements with a similar flux rate (Figures 1A,B, 2 and 3). Under ST and LT stress conditions, the total flux rates of Cd²⁺ and Ca²⁺ in the presence of Cd²⁺ (=Σ_Ca²⁺_+Cd²⁺ with a molar ratio of Cd²⁺ to Ca²⁺ of 1:1) were 37.8–77.4 (NM), 54.1–96.2 (MAJ), and 53.7–122.1 pmol cm^-2 s^-1 (NAU), as calculated on the basis of Figures 2 and 3 (Supplementary Figure S4). The relationships between Σ_Ca²⁺_+Cd²⁺ and Ca²⁺ flux in the absence of CdCl₂ [Σ_Ca²⁺_(-Cd²⁺₎] were highly significant and close to 1 for NM and MAJ colonized roots and slightly increased to 1.4 for NAU colonized roots (Figure 4). These suggest that the entry of Cd²⁺ and Ca²⁺ is mainly through the same pathway in NM and EM roots, mostly likely through Ca²⁺-permeable channels in the PM (see below).

FIGURE 4.

The correlation between total fluxes of Ca²⁺ and Cd²⁺ (Σ_Ca²⁺_+Cd²⁺) in the presence of CdCl₂ (50 μM, +Cd) and Ca²⁺ flux [Σ_Ca²⁺_(-Cd²⁺₎] in the absence of CdCl₂ (-Cd) in roots of ectomycorrhizal (MAJ and NAU) and non-mycorrhizal (NM) Populus × canescens. Ectomycorrhizal (MAJ and NAU) and NM P. × canescens plants were subjected to short-term (ST, 24 h) and long-term (LT, 7 days) exposure to 50 μM CdCl₂, respectively. Control roots were well fertilized but treated without CdCl₂. Ca²⁺ and Cd²⁺ fluxes were measured along root axis, 100–2,300 μm from the apex, at intervals of 200–300 μm. Each point is the mean of 4–5 individual plants.

H⁺ Fluxes

In the absence of CdCl₂, EM roots showed a typical H⁺ efflux at the apex, which was 7.6-fold higher than that in NM roots (Figure 1C). CdCl₂ (50 μM) shock stimulated H⁺ efflux in both NM and EM plants with a stronger response in EM than in NM roots (Figure 1C). Pure MAJ and NAU mycelia exhibited a net H⁺ efflux under control conditions similar to that observed for MAJ- and NAU-colonizing roots (Figure 1C). However, in pure mycelia the fluxes were 4.8-fold higher than in EM roots (Figure 1C). After exposure to CdCl₂ (50 μM), hyphae exhibited a transient increase in the H⁺ efflux, which then remained constant during the period of recording (40 min; Figure 1C). Compared with strain MAJ, strain NAU exhibited higher H⁺ efflux irrespective of control or CdCl₂ shock treatments (Figure 1C).

Steady-state recordings on EM roots showed that the pattern of H⁺ flux in ST-stressed roots (50 μM CdCl₂, 24 h) differed from those subjected to LT Cd²⁺ exposure (50 μM CdCl₂, 7 days). Under ST conditions, CdCl₂ (50 μM) stimulated H⁺ efflux in EM plants, whereas under LT conditions, EM roots showed a pronounced H⁺ influx (Figure 5). In NM roots, CdCl₂ (50 μM) decreased H⁺ influx upon ST exposure or shifted it to a net H⁺ efflux under LT stress conditions (Figure 5). The pattern of H⁺ flux in the fungal mycelia differed from that in EM roots under ST stress (Figure 5). ST treatment reduced the efflux of H⁺ from the two fungal strains, which is contrast to EM roots where an enhanced H⁺ efflux was observed (Figure 5). LT stress caused a pronounced shift of H⁺ efflux to influx into pure mycelia of the two strains, similar to the finding in LT-stressed EM roots (Figure 5).

FIGURE 5.

Effects of CdCl₂ on steady H⁺ fluxes in Populus × canescens roots and Paxillus involutus strains MAJ and NAU. P. involutus isolates, ectomycorrhizal (MAJ and NAU) and non-mycorrhizal (NM) P. × canescens plants were subjected to short-term (ST, 24 h) and long-term (LT, 7 days) exposure to 50 μM CdCl₂, respectively. Control roots and axenic mycelia were well fertilized but treated without CdCl₂. H⁺ fluxes in poplar roots were measured along root axis, 100–2,300 μm from the apex, at intervals of 200–300 μm. H⁺ fluxes of P. involutus isolates MAJ and NAU were measured along the surface of pelleted hyphae over a recording period of 30 min. Inserted sections show the H⁺ fluxes in P. involutus isolates after short-term (ST, 24 h) or long-term (LT, 7 days) CdCl₂ treatment. Each point is the mean of 4–5 individual plants or axenic EM cultures (pelleted hyphae), and bars represent the standard error of the mean. Asterisks denote significant difference at P < 0.05 between treatments.

Cd²⁺-Altered Flux Profiles of H₂O₂ in EM Roots

H₂O₂-sensitive microprobes were used to detect the H₂O₂ response to Cd²⁺ exposure in NM and EM roots. In the absence of Cd²⁺, NM roots exhibited a stable H₂O₂ efflux (0.7–1.5 pmol cm^-2 s^-1) along the root axis; the mean flux rate increased 2.4-fold in response to Cd²⁺ treatment (50 μM CdCl₂, 30 min, Figure 6). Ectomycorrhization of poplar roots with P. involutus stains, MAJ and NAU, resulted in a significant increase of H₂O₂ efflux along the roots (Figure 6). However, upon CdCl₂ exposure EM roots displayed decreased H₂O₂ efflux in contrast to NM roots (Figure 6).

FIGURE 6.

Effects of CdCl₂ on steady H₂O₂ flux in roots of ectomycorrhizal (MAJ and NAU) and non-mycorrhizal (NM) Populus × canescens plants. Ectomycorrhizal (MAJ and NAU) and NM P. × canescens plants were subjected to 50 μM CdCl₂ for 30 min. Control roots were well fertilized but treated without CdCl₂. H₂O₂ flux was measured along root axis, 100–2,300 μm from the apex, at intervals of 200–300 μm. Each point is the mean of 4–5 individual plants, and bars represent the standard error of the mean. Asterisks denote significant difference at P < 0.05 between treatments.

Effects of Ca²⁺, H₂O₂, pH, and PM Transporter and Channel Inhibitors on Cd²⁺-Altered Ion Flux Profiles in EM Roots

Ca²⁺ and Ca²⁺ Channel Inhibitors

Here, pharmacological experiments were carried out to test whether putative Ca²⁺ channels inhibitors could inhibit Cd²⁺ influx in poplar roots. Four typical Ca²⁺ channels inhibitors, LaCl₃, GdCl₃, verapamil, and TEA effectively inhibited Ca²⁺ influx in NM and EM roots, regardless of Cd²⁺ treatments (Figure 7A, Supplementary Figures S5A, S6A, and S7A). LaCl₃ restricted Cd²⁺ influx in CdCl₂-treated NM and EM roots (Figure 7B). This suggests that Cd²⁺ is taken up through Ca²⁺-permeable channels because La³⁺ is able to block various types of Ca²⁺-permeable channels, including depolarisation-, hyperpolarisation-, elicitor-activated, and voltage-insensitive channels (Weiss, 1974; Pickard and Ding, 1993; Gelli and Blumwald, 1997; Piñeros and Tester, 1997; Zimmermann et al., 1997; White, 1998, 2000). Moreover, the other three Ca²⁺-permeable channel inhibitors, GdCl₃, verapamil, and TEA, diminished Cd²⁺ influx to a similar extent as LaCl₃-treated plants (Figure 7B, Supplementary Figures S5B, S6B, and S7B). Similarly, in the pure P. involutus mycelia, LaCl₃ also effectively restricted influx of Ca²⁺ and Cd²⁺ or induced net efflux (Supplementary Figure S8).

FIGURE 7.

Effects of LaCl₃ and external Ca²⁺ on steady Cd²⁺ and/or Ca²⁺ fluxes in roots of ectomycorrhizal (MAJ and NAU) and non-mycorrhizal (NM) Populus × canescens plants under Cd²⁺ stress. (A,B) Ectomycorrhizal (MAJ and NAU) and NM P. × canescens plants were subjected to 50 μM CdCl₂ for 24 h in the presence and absence of 5 mM LaCl₃. Control roots were well fertilized but treated without CdCl₂ or LaCl₃. (C) Ectomycorrhizal (MAJ and NAU) and NM P. × canescens plants were subjected to 50 μM CdCl₂ for 24 h prior to Cd²⁺ flux recordings in the presence of CaCl₂ (25 μM, 50 μM, or 100 μM; the ratio of Ca²⁺:Cd²⁺ was 1:2; 1:1, and 2:1). Ca²⁺ (A) and Cd²⁺ (B,C) fluxes were measured along root axes, 100–2,300 μm from the apex, at intervals of 200–300 μm. Each point is the mean of 4–5 individual plants and bars represent the standard error of the mean. Inserted sections show the mean flux rates and different letters, a, b, c, and d, indicate significant difference at P < 0.05 between treatments.

Additionally, a co-application of Cd²⁺ and Ca²⁺ suppressed the entry of Cd²⁺ in NM and EM roots, and the restriction increased with the increasing fraction of Ca²⁺ in the mixture (Ca²⁺: Cd²⁺ = 1:2, 1:1, 2:1; Figure 7C). The mean Cd²⁺ flux decreased by 95.7% (NM), 72.1% (MAJ), and 45.5% (NAU) at a ratio of Ca²⁺:Cd²⁺ = 2:1, compared to a those with a higher Cd²⁺ fraction, Ca²⁺:Cd²⁺ = 1:2 (Figure 7C). These results suggest that the divalent cations, Cd²⁺ and Ca²⁺, competitively permeated the plasma membrane through Ca²⁺ channels. The lower reduction in Cd²⁺ influx in EM than in NM roots in the presence of Ca²⁺ (Figure 7C) reflects the high flow of Cd²⁺ through the activated Ca²⁺ channels.

We observed that the presence of Ca²⁺ in the measuring solution marginally lowered the Cd²⁺ signals (14.7–26.0%) detected by the Cd²⁺ microelectrodes filled with Cd²⁺ liquid ion exchanger (LIX) (Supplementary Table S1). In the absence of Ca²⁺, the working voltage of microelectrodes and the detected Cd²⁺ signals in Cd²⁺-treated roots were unstable and fluctuated greatly during the period of recording (data not shown). This behavior is presumably caused by the plant response to nutrient deficiency in the root medium (Li J. et al., 2012). In our study, Cd²⁺ electrodes exhibited higher sensitivity at 0.05 mM Ca²⁺ in the absence and presence of 0.1 mM K⁺ and 0.1 mM Mg²⁺ (Supplementary Table S1). The presence of nutrients, K⁺, Ca²⁺, and Mg²⁺, did not affect the accuracy of our conclusions relating to Cd²⁺ fluxes in NM and EM roots.

H₂O₂ and ROS Scavenger

To investigate whether Cd²⁺ entry through Ca²⁺-permeable channels is activated by H₂O₂, we examined the effects of hydrogen peroxide and the ROS (reactive oxygen species) scavenger DMTU on Cd²⁺ and Ca²⁺ fluxes. Transient kinetic recordings showed that Cd²⁺ shock caused an immediate increase of Cd²⁺ influx but enhanced Ca²⁺ efflux in NM and EM roots (Figure 8). The flux rates of Cd²⁺ and Ca²⁺ decreased with prolonged exposure time (Figure 8). Notably, Cd²⁺ influx markedly increased upon H₂O₂ shock (1.0 mM) in both NM and EM roots (Figure 8A). However, the Cd²⁺-elicited Ca²⁺ efflux was reduced by H₂O₂ in EM roots or shifted to a net influx in NM roots (Figure 8B). These results suggest that H₂O₂ stimulated the entry of Cd²⁺ and Ca²⁺, presumably through the plasma membrane Ca²⁺ channels of the roots.

FIGURE 8.

Effects of CdCl₂ and H₂O₂ on transient kinetics of Cd²⁺ and Ca²⁺ in roots of ectomycorrhizal (MAJ and NAU) and non-mycorrhizal (NM) Populus × canescens. Cd²⁺ (A) and Ca²⁺ (B) kinetics were recorded before and after the required amount of 50 μM CdCl₂ or 1.0 mM H₂O₂was introduced into the measuring chamber. Prior to the CdCl₂ shock, steady-state fluxes of Cd²⁺ and Ca²⁺ were monitored at the apex (measuring site was ca. 100 μm from the root tip) for approximately 10–20 min. Transient kinetics of Cd²⁺ and Ca²⁺ were recorded after the required amount of 50 μM CdCl₂ was introduced into the measuring solution. After 20–30 min continuous recording of Cd²⁺ and Ca²⁺ fluxes, Cd²⁺ and Ca²⁺ kinetics were recorded for 20 min after 1.0 mM H₂O₂ was introduced into the measuring solution. Each point represents the mean of 4–5 individual plants and bars represent the standard error of the mean.

Ca²⁺ influx in NM and EM roots were suppressed by the ROS scavenger, DMTU (5 mM), irrespective of the presence and absence of Cd²⁺ (Figure 9A). Similarly, the supplement of DMTU significantly reduced the influx of Cd²⁺ in NM and EM roots (Figure 9B). These data indicated that H₂O₂ play a crucial role in accelerating the influx of Ca²⁺ and Cd²⁺, which is accordance to the results obtained by direct H₂O₂ applications (Figure 8).

FIGURE 9.

Effects of DMTU on steady Ca²⁺ and Cd²⁺ fluxes in roots of ectomycorrhizal (MAJ and NAU) and non-mycorrhizal (NM) Populus × canescens under Cd²⁺ stress. Ectomycorrhizal (MAJ and NAU) and NM P. × canescens plants were subjected to 0 or 50 μM CdCl₂ for 24 h in the presence or absence of 5 mM DMTU. Ca²⁺ (A) and Cd²⁺ (B) fluxes were measured along root axes, 100–2,300 μm from the apex, at intervals of 200–300 μm. Each point is the mean of 4–5 individual plants and bars represent the standard error of the mean. Inserted sections show the mean flux rates and different letters, a, b, c, and d, indicate significant difference at P < 0.05 between treatments.

External pH and H⁺-ATPase Inhibitor

Fluxes of Cd²⁺ and Ca²⁺ depend on external pH. An acidic environment accelerated Cd²⁺ and Ca²⁺ influxes in both NM and EM roots with the strongest influx at pH 5.2 and the lowest at pH 6.2 or a neutral pH, 7.2 (Figure 10). Moreover, we noticed that the pH effects on fluxes of Cd²⁺ and Ca²⁺ were more pronounced in NM roots than in EM roots (Figure 10). Compared to an acidic environment (pH 5.2), the mean flux rate of the divalent cations decreased by 45.8% (Ca²⁺) and 38.8% (Cd²⁺) in EM roots under pH 6.2–7.2 (Figure 10). In NM roots, the increasing pH lowered Cd²⁺ influxes by 56.5% or even reversed the rectifications of Ca²⁺ (influx → efflux) at a neutral pH, 7.2 (Figure 10). The less reduced influx of Ca²⁺ and Cd²⁺ in EM roots at pH 6.2 or 7.2 was due to the high H⁺-pumping activity in the PM (see below).

FIGURE 10.

Effects of pH on steady Cd²⁺ and Ca²⁺fluxes in roots of ectomycorrhizal (MAJ and NAU) and non-mycorrhizal (NM) Populus × canescens under Cd²⁺ stress. Ectomycorrhizal (MAJ and NAU) and NM P. × canescens plants were subjected to 50 μM CdCl₂ for 24 h prior to flux recordings at pH 5.2, 6.2 or 7.2. Cd²⁺ (A) and Ca²⁺ (B) fluxes were measured along root axes, 100–2,300 μm from the apex, at intervals of 200–300 μm. Each point is the mean of four to five individual plants and bars represent the standard error of the mean. Inserted sections show the mean flux rates and different letters, a, b, and c, indicate significant difference at P < 0.05 between treatments.

Sodium orthovanadate (500 μM), the specific inhibitor of PM H⁺-ATPase, increased the H⁺ influx in NM roots slightly, but caused a drastic shift from H⁺ efflux toward influx in both EM roots and P. involutus mycelia, irrespective of Cd²⁺ treatment (Figure 11A, Supplementary Figure S9). Sodium orthovanadate significantly reduced the Cd²⁺ influx along the roots in Cd²⁺-treated NM and EM plants (Figure 11B). In the absence of Cd²⁺, the PM H⁺-ATPase inhibitor reduced Ca²⁺ influx in NM and MAJ-ectomycorrhizal roots or shifted to efflux in NAU-ectomycorrhizal roots (Figure 11C). The inhibition of Ca²⁺ influx by sodium orthovanadate was more pronounced in the presence of Cd²⁺: the H⁺-pump inhibitor reversed the rectifications of Ca²⁺ from influx to efflux in NM and EM roots (Figure 11C).

FIGURE 11.

Effects of sodium orthovanadate on steady H⁺, Cd²⁺, and Ca²⁺ fluxes in roots of ectomycorrhizal (MAJ and NAU) and non-mycorrhizal (NM) Populus × canescens under Cd²⁺ stress. Ectomycorrhizal (MAJ and NAU) and NM P. × canescens plants were subjected to 0 or 50 μM CdCl₂ for 24 h in the presence and absence of 500 μM sodium orthovanadate. H⁺ (A), Cd²⁺ (B), and Ca²⁺ (C) fluxes were measured along root axes, 100–2,300 μm from the apex, at intervals of 200–300 μm. Each point is the mean of 4–5 individual plants and bars represent the standard error of the mean. Inserted sections show the mean flux rates and different letters, a, b, c, and d, indicate significant difference at P < 0.05 between treatments.

Discussion

Colonization of P. × canescens Roots with Paxillus involutus Stimulates Cd²⁺ Uptake under Cd²⁺ Stress

The woody Cd²⁺-hyperaccumulator P. × canescens exhibited a vigorous Cd²⁺ uptake after a 50 μM CdCl₂ shock (40 min), ST (24 h), and LT (7 days) treatment (Figures 1A and 2). The result is consistent with previous findings where P. × canescens roots exhibited a high Cd²⁺ uptake after 40 days of CdSO₄ exposure (50 μM, Ma Y. et al., 2014). Similarly, a high entry of Cd²⁺ was recorded in hyperaccumulating ecotypes of Sedum alfredii (Lu et al., 2010; Sun et al., 2013a) and Suaeda salsa under Cd²⁺ stress (Li et al., 2012a). An important result was that EM roots exhibited higher Cd²⁺ influx than NM roots irrespective of Cd²⁺ stress conditions, shock, ST, and LT (Figures 1A and 2). Substantial evidence indicates that Cd²⁺ can be enriched in ectomycorrhizal plants (Sell et al., 2005; Baum et al., 2006; Krpata et al., 2008, 2009; Sousa et al., 2012; Ma Y. et al., 2014). The enhanced Cd²⁺ uptake in EM roots is partly due to the capacity of the fungus to take up Cd²⁺ because CdCl₂ shock resulted in a net Cd²⁺ influx in the mycelia of the two P. involutus strains and the flux rate increased with the prolonged duration of CdCl₂ treatment from 24 h to 7 days (Figures 1A and 2). In liquid cultures, P. involutus cultures also showed high capacities for Cd²⁺ accumulation (Ott et al., 2002). P. involutus could bind Cd²⁺ onto the cell walls or accumulate the metal in the vacuolar compartment (Blaudez et al., 2000; Ott et al., 2002). Moreover, the ectomycorrhizal fungus appears to detoxify high concentrations of Cd²⁺ by (i) the chelation of metal ions in the cytosol with thiol-containing compounds, e.g., glutathione, phytochelatins, or metallothioneins (Courbot et al., 2004; Jacob et al., 2004), and (ii) activation of antioxidative defense system (Jacob et al., 2001; Ott et al., 2002). Our pharmacological data revealed that Cd²⁺ entered the fungal hyphae mainly through PM Ca²⁺ channels because the influx was suppressed by LaCl₃, a Ca²⁺ channel blocker (Supplementary Figure S8B). Therefore, Cd²⁺ enriched by ectomycorrhizal hyphae is thought to be transferred to the host roots, probably through the apoplastic space during the period of Cd²⁺ stress.

There were marked differences between the two strains in Cd²⁺ uptake given the shock treatment (Figure 1A). Pure fungal mycelium of MAJ accumulated Cd²⁺ with a higher rate than NAU (Figure 1A). In the P. involutus-ectomycorrhizal symbioses, the incompatible fungal isolate NAU is unable to induce a functional ectomycorrhizae while MAJ forms a typical Hartig net with the roots of P. × canescens (Gafur et al., 2004). Thus, in MAJ-colonized roots the host cells might have been more accessible to Cd²⁺. In accordance, MAJ roots exhibited a higher influx than NAU roots after the onset of CdCl₂ shock (Figure 1A). However, Cd²⁺ influx into NAU-colonized roots was similar to that of MAJ-colonized roots during ST or LT Cd²⁺ treatment (Figure 2). This was likely due to (i) similar capacities for Cd²⁺ uptake of MAJ and NAU hyphae during a 24-h or 7-days of Cd²⁺ exposure (Figure 2), or (ii) similar uptake capacity of the fungus-ensheathed inner root cells (Figure 2). The observed correlation between EM and NM roots showed that the continuous Cd²⁺ flow was mainly the consequence of host roots in the Cd²⁺-stressed ectomycorrhizal symbioses during a prolonged period of Cd²⁺ exposure (24 h to 7 days; Supplementary Figure S2).

Paxillus involutus-Ectomycorrhizas Enhance Cd²⁺ Influx through Ca²⁺-Permeable Channels in the Plasma Membrane

Our data revealed that the entry of Cd²⁺ is likely mediated through PM Ca²⁺ channels in the fungal hyphae and poplar roots, and P. involutus-ectomycorrhizas facilitated the channel-mediated Cd²⁺ influx under Cd²⁺ stress. The experimental evidence for these conclusions is briefly listed below.

• simple (1)

  The addition of Cd²⁺ resulted in an immediate influx of Cd²⁺ in NM roots, and the flux was more pronounced in EM roots (Figure 1A). Rapid entry of Cd²⁺ is generally through PM ion channels that are permeable to Cd²⁺ (The first 1–2 min flux recordings were discarded to diminish the diffusion effect of stock addition in roots and fungal mycelia). Our pharmacological data revealed that the net Cd²⁺ influx in CdCl₂-stressed NM and EM roots was strongly suppressed by typical Ca²⁺ channel blockers, such as LaCl₃, GdCl₃, verapamil, and TEA (Figure 7B, Supplementary Figures S5B, S6B, and S7B). Moreover, in P. involutus mycelium the CdCl₂-elicited influx of Cd²⁺ was also inhibited by LaCl₃ (Supplementary Figure S8B). These results suggest that under CdCl₂ stress Cd²⁺ enters fungal and root tissues through PM Ca²⁺ channels.

• simple (2)

  Cd²⁺ treatments (shock, ST, and LT) affected the uptake of Ca²⁺ in poplar roots (Figures 1B and 3), while the influx of Cd²⁺ declined with increasing the concentration of Ca²⁺ when NM and EM roots were subjected to the concomitant application of Cd²⁺ and Ca²⁺ (Figure 7C). Similarly, the Cd²⁺ influx was affected by the presence of Ca²⁺ in two contrasting (hyperaccumulating and non-hyperaccumulating) Sedum alfredii ecotypes (Lu et al., 2010). It was suggested that Ca²⁺ and Cd²⁺ ions compete for the binding sites of transporters (Gussarsson et al., 1996; Rodríguez-Serrano et al., 2009). Our transient kinetics showed that Cd²⁺ exposure blocked the Ca²⁺ influx and caused an immediate change in the rectification of Ca²⁺ from influx to efflux (Figures 1B and 8B). This suggests that Cd²⁺ ions competed with Ca²⁺ to penetrate across PM Ca²⁺ channels that are permeable to divalent cations (Perfus-Barbeoch et al., 2002).

• simple (3)

  In ST- and LT-stressed NM and EM roots, the total flux rates of Cd²⁺ and Ca²⁺ in the presence of Cd²⁺ (Σ_Ca²⁺_+Cd²⁺) were nearly equal to the flux rate of Ca²⁺ in the absence of Cd²⁺ stress (Σ_Ca²⁺_(-Cd²⁺₎; Supplementary Figure S4). Moreover, the correlations between Σ_Ca²⁺_+Cd²⁺ and Σ_Ca²⁺_(-Cd²⁺₎ (Figure 4) suggest that Cd²⁺ ions enter NM and EM roots mainly through Ca²⁺-permeable channels in the PM.

Collectively, under CdCl₂ stress Cd²⁺ ions could penetrate the PM Ca²⁺ channels in fungal hyphae and in P. × canescens roots. At present we cannot exclude the possibility that Cd²⁺ penetrated the PM through transporters for Cd²⁺ (Ma Y. et al., 2014; He et al., 2015) or other nutritional ions (Gussarsson et al., 1996; Cohen et al., 1998; Zhao et al., 2002; Cosio et al., 2004; Clemens, 2006), because (1) the four types of Ca²⁺ channel inhibitors applied here were not able to fully block the Cd²⁺ influx in NM and EM roots (Figure 7B, Supplementary Figures S5B, S6B and S7B), and (2) the total flux of Cd²⁺ and Ca²⁺ (Σ_Ca²⁺_+Cd²⁺, molar ratio of Cd²⁺ to Ca²⁺ is 1:1) under ST and LT Cd²⁺ stress was 10.9–27.7% higher than the flux rate of Ca²⁺ under non-Cd²⁺ conditions (Figure 4, Supplementary Figure S4). This implies that a small fraction of Cd²⁺ ions penetrated the PM through other channels and transporters.

Plasma membrane Ca²⁺ channels in P. involutus hyphae maybe more permeable to Cd²⁺ compared to the channels in P. × canescens roots as the fungal mycelium displayed a typical higher Ca²⁺ influx than poplar roots under control and Cd²⁺-stress conditions (Figures 1B and 3). We cannot discriminate between the channels of the fungus and those of the plant in the ectomycorrhizal symbiosis, but the Cd²⁺ and Ca²⁺ fluxes in EM roots appear to mainly reflect the response of the host plants to Cd²⁺ stress because (1) EM roots exhibited a different pattern from the P. involutus mycelia in enhancing Ca²⁺ and Cd²⁺ uptake under hydroponic Cd²⁺ conditions. Cd²⁺-shocked MAJ and NAU fungal strains usually displayed a stable Cd²⁺ influx with the exception of an initial transient increase (Figure 1A). However, EM roots showed a declined Cd²⁺ influx over the duration of Cd²⁺ exposure, similar to the Cd²⁺ kinetics in NM roots (Figure 1A). Moreover, the Cd²⁺ influx in the mycelia of the two P. involutus strains increased with the prolonged CdCl₂ exposure from 24 h to 7 days (from 23.9 to 72.7 pmol cm^-2 s^-1; Figure 2). In contrast, the Cd²⁺ fluxes in EM roots were relatively stable under ST (43.7 ± 8.4 pmol cm^-2 s^-1) and LT treatments (35.9 ± 6.0 pmol cm^-2 s^-1; Figure 2). ST, LT, and Cd²⁺ shock increased the Ca²⁺ influx in P. involutus mycelia, while the Ca²⁺ influx in EM roots was declined by these Cd²⁺ treatments (Figures 1B and 3). (2) NMT data showed that ion fluxes in mature P. × canescens–P. involutus symbiotic associations bear a striking resemblance to the ST inoculated roots (Supplementary Figure S1). Similar findings have been previously reported in a salt stress study where P. × canescens roots were inoculated with P. involutus for 10 and 20 days (Ma X. et al., 2014). At early stages of fungal co-culture the Cd²⁺ and Ca²⁺ influx is mostly the result of host properties. Therefore, the Cd²⁺ and Ca²⁺ stimulation in P. involutus-ectomycorrhizal roots reflects the enhanced root uptake ability. (3) The correlation analyses revealed that Cd²⁺ and Ca²⁺ influxes in EM roots show a significant relationship with NM roots but not with fungal mycelia under various Cd²⁺ treatments (shock, ST, and LT; Supplementary Figures S2 and S3). Taken together, these data suggest that the continuous flow of Cd²⁺ and Ca²⁺ in EM roots detected by NMT microelectrodes was largely driven by the host and that the fungal partner enhanced fluxes leading to enriched Cd²⁺ and Ca²⁺ concentrations.

The observed patterns of Cd²⁺ and Ca²⁺ fluxes upon Cd²⁺ exposure could be explained by channel-mediated ion fluxes. NMT data show that the Ca²⁺ flux in EM roots was negatively correlated with the Ca²⁺ influx in fungal hyphae upon Cd²⁺ shock treatment (Supplementary Figure S3). This is presumably the result of Cd²⁺-Ca²⁺ competition across the Ca²⁺ channels in the root PM. After being exposed to Cd²⁺ shock, Ca²⁺ entry was enhanced in the hyphae (Figure 1B). However, the fungal hyphae which were enriched in Ca²⁺ ions, were unable to deliver Ca²⁺ to the root cells because the Cd²⁺ ions competitively inhibited the entry of Ca²⁺ through the PM channels. As a result, the high influx of Ca²⁺ through fungal hyphae led to an apparently greater Ca²⁺ efflux in Cd²⁺-exposed EM roots (Figures 1B and 8B).

Paxillus involutus colonization enhanced the uptake of Cd²⁺ under shock, ST, and LT stress, compared to NM roots (Figures 1A and 2). The increased entry of Cd²⁺ is likely due to the activation of PM Ca²⁺ channels in the ectomycorrhizas. The stimulated Ca²⁺ influx by P. involutus inoculation revealed the activation of PM Ca²⁺ channels since the ectomycorrhiza-enhanced entry of Ca²⁺ was suppressed by Ca²⁺ channel blockers (LaCl₃, GdCl₃, verapamil, or TEA; Figure 7A, Supplementary Figures S5A, S6A, and S7A). The activated PM Ca²⁺ channels allowed the entry of Cd²⁺ in addition to Ca²⁺ under Cd²⁺ stress (Figures 1A and 2).

Hydrogen Peroxide Induced by CdCl₂ and Fungal Colonization Stimulates Cd²⁺ Influx through PM Ca²⁺ Channels

After being subjected to CdCl₂ exposure, NM roots displayed an increased H₂O₂ efflux along the root axis (Figure 6). It is well documented that Cd²⁺ induced accumulation of H₂O₂ in pine roots (Schützendübel et al., 2001), P × canescens roots (Schützendübel et al., 2002), and in suspension cultures of tobacco (Piqueras et al., 1999) and P. euphratica (Sun et al., 2013b; Han et al., 2016). H₂O₂ efflux was evident in EM roots irrespective of the presence or absence of Cd²⁺ treatments (Figure 6). Our results suggest that the Cd²⁺ influx through PM Ca²⁺ channels is stimulated by H₂O₂ in NM and EM roots. The experimental evidence and explanations are briefly listed here.

• simple (1)

  H₂O₂ (1.0 mM) exhibited an enhancement on Ca²⁺ influx in NM and EM roots (Figure 8B). Pei et al. (2000) showed that H₂O₂ (0.05–5.0 mM) activates Ca²⁺ currents through PM Ca²⁺ channels of Arabidopsis thaliana guard cells. Moreover, H₂O₂ increased Ca²⁺ influx across the PM in P. euphratica cells (Sun et al., 2010), roots of A. thaliana (Demidchik et al., 2007) and mangroves (Lu et al., 2013). Furthermore, the H₂O₂-stimulated entry of Ca²⁺ in P. euphratica cells was inhibited by LaCl₃ (Sun et al., 2010). In this study, Cd²⁺ influx in NM and EM roots was significantly enhanced after exposure to 1.0 mM H₂O₂ (Figure 8A). This finding is in agreement with Sun et al. (2013b) and Han et al. (2016), who found that H₂O₂ (3.0 mM) stimulated entry of Cd²⁺ into P. euphratica cells. In addition, the Cd²⁺ influx was blocked by LaCl₃ in CdCl₂-stressed roots (Figure 7B). These results suggest that H₂O₂ stimulates the influx of Cd²⁺ and Ca²⁺ through Ca²⁺-permeable channels in the PM.

• simple (2)

  The H₂O₂ induction of Cd²⁺ resembles the pattern of Ca²⁺ kinetics in response to H₂O₂ (Figure 8). Moreover, Cd²⁺ and Ca²⁺ influx in NM and EM roots were both suppressed by the ROS scavenger, DMTU (Figure 9). Similarly, Sun et al. (2013b) showed that the entry of Cd²⁺ into P. euphratica cells was reduced when a H₂O₂ scavenger, catalase, was applied.

Taken together, these results suggest that Cd²⁺ and Ca²⁺ ions enter NM and EM roots by the same pathway involving PM Ca²⁺ channels that are activated by Cd²⁺-elicited H₂O₂.

The high Cd²⁺ influx in EM roots resulted from the pronounced activation of PM Ca²⁺ channels that were stimulated, at least in part, by the fungal-elicited H₂O₂. Compared to NM roots, MAJ- and NAU-ectomycorrhizal roots displayed a significant higher H₂O₂ efflux in the absence of Cd²⁺ stress (Figure 6), suggesting that the inoculation with P. involutus caused a strong production of H₂O₂ in EM roots. This finding agrees with Gafur et al. (2004) and Langenfeld-Heyser et al. (2007), who detected strong H₂O₂ accumulation in the outer hyphae mantle of compatible (MAJ) and incompatible (NAU) interactions. H₂O₂ production in the hyphae is suggested to regulate host’s root growth, defense against other invading microbes, and increasing plant-innate immunity (Salzer et al., 1999; Gafur et al., 2004). In our study, H₂O₂ produced in the ectomycorrhizae accelerated the influx of Ca²⁺ in the absence of Cd²⁺, whereas it increased entry of Cd²⁺ in the presence of high external Cd²⁺ (Figure 8). ROS scavenging by DMTU simultaneously decreased Ca²⁺ and Cd²⁺ influxes along the root axis of EM plants (Figure 9). These observations suggest that H₂O₂ produced in compatible (MAJ) and incompetent (NAU) ectomycorrhizal associations activated Ca²⁺ permeable channels, which allowed the entry of Cd²⁺ under Cd²⁺ stress.

We noticed that the H₂O₂ efflux in MAJ and NAU-ectomycorrhizal roots was lowered by Cd²⁺ stress (Figure 6). This reduction may have resulted from the activation of antioxidant enzymes and increased amounts of ROS scavengers produced as a defense response. It has been repeatedly shown that the antioxidant enzyme activities are activated under heavy metal stresses (Schützendübel et al., 2001, 2002; Rozpądek et al., 2014; Chen et al., 2015; Tan et al., 2015). The enhanced activities of superoxide dismutase, peroxidase, catalase, and ascorbate peroxidase play an important role in scavenging the Cd²⁺-elicited H₂O₂ in plants (Garg and Aggarwal, 2012; Anjum et al., 2015; Tan et al., 2015). To combat Cd²⁺-induced superoxide and H₂O₂, P. × canescens plants were found to rely mainly on antioxidant enzymes and the formation of the potential radical scavenging molecules, such as proline, sugar alcohols and soluble phenolics (He et al., 2011). However, the lowered H₂O₂ efflux in EM roots (Figure 6) did not reduce the Cd²⁺-elicited entry of Cd²⁺, because (i) the fungal-elicited H₂O₂ had already activated the Ca²⁺-channels before the Cd²⁺ addition, and/or (ii) the H₂O₂ level is still high enough to activate the channels under Cd²⁺ stress. The observation that stressed EM roots still contain high concentrations of H₂O₂ in the hyphae (Langenfeld-Heyser et al., 2007) supports these speculations.

PM H⁺-ATPase Activated by Cd²⁺ and Fungal Colonization Stimulates Cd²⁺ Influx through PM Ca²⁺ Channels

In addition to H₂O₂, PM H⁺-ATPase activated by Cd²⁺ and enhanced by fungal colonization also accelerated Cd²⁺ influx through PM Ca²⁺ channels in NM and EM roots. PM H⁺-ATPases pump protons into the external medium to maintain an electrochemical H⁺ gradient across the PM (Blumwald et al., 2000; Zhu, 2003). Krämer (2010) suggested that H⁺-ATPases play an important role in adaptation of plants to heavy metal stress. The finding that the net H⁺ efflux in fungal mycelia and EM roots was markedly reduced by a specific inhibitor of PM H⁺-ATPase (sodium orthovanadate) in the presence and absence of Cd²⁺ stress (Figure 11A, Supplementary Figure S9) supports that the vigorous H⁺ efflux is the consequence of H⁺-ATPase activity. Accordingly, the increased H⁺ efflux upon Cd²⁺ shock (NM, MAJ and NAU roots; Figure 1C), ST (MAJ and NAU roots; Figure 5) and LT stress (NM roots; Figure 5) indicates the activated H⁺-pumping activity. In NM and EM roots, Cd²⁺ exposure led to a marked upregulation of HA2.1 and AHA10.1, two important genes encoding PM H⁺-ATPases (Ma Y. et al., 2014). The activation of PM H⁺-ATPase by Cd²⁺ is likely associated with the Cd²⁺-elicited H₂O₂, since (i) H₂O₂ increased H⁺ pumping activity in P. euphratica callus cells (Sun et al., 2010), in roots of P. euphratica (Sun et al., 2010) and secretor and non-secretor mangrove species (Lu et al., 2013; Lang et al., 2014), and (ii) the expression of genes encoding PM H⁺-ATPase are stimulated by H₂O₂ in Cucumis sativus roots (Janicka-Russak and Kabala, 2012; Janicka-Russak et al., 2012).

The activated PM H⁺-ATPase enabled NM and EM roots to maintain an acidic environment, which favors the entry of Cd²⁺ across the PM (Figure 10A). Similarly, He et al. (2015) showed that pH 5.5 accelerates Cd²⁺ influx into poplar roots compared to pH 4.0 or pH 7.0. Moreover, the Cd²⁺ influx was markedly suppressed by the application of sodium vanadate, an inhibitor of PM H⁺-ATPase (Figure 11B). These results indicate that the PM H⁺-pumps play a crucial role in enhancing the entry of Cd²⁺ (Ma Y. et al., 2014). Accordingly, NMT profiles of NM and EM roots showed that the maximum influx of Ca²⁺ was observed at pH 5.2 (Figure 10B), and that Ca²⁺ influx was blocked by sodium vanadate (Figure 11C). Therefore, we infer that Cd²⁺ activated H⁺-pumping in the PM, which led to hyperpolarization of the PM and increased Cd²⁺ influx through hyperpolarization-activated Ca²⁺ channels (HACCs). However, at present we cannot exclude the possibility that Cd²⁺ ions also penetrated through depolarization-activated (DACCs) and voltage-independent Ca²⁺ channels (VICCs), because the inhibitor of PM H⁺-ATPase, sodium vanadate, could not fully block the Cd²⁺ influx in NM and EM roots (Figure 11B). It has been shown that NSCCs co-exist with HACCs in the root cell plasma membrane to mediate the entry of Ca²⁺, but the two Ca²⁺ influx routes differ in their voltage sensitivity (Demidchik et al., 2002).

Ectomycorrhizal Populus × canescens show highly activated H⁺-pumping activity in the PM, which favors the Cd²⁺ influx through HACCs. Our NMT data showed that colonization of P. × canescens with P. involutus caused a marked H⁺ efflux (Figures 1C, 5, and 11A), suggesting that the fungal colonization could activate the PM H⁺-ATPase in ectomycorrhizas. This is consistent to our previous studies (Li J. et al., 2012; Ma X. et al., 2014). It has been documented that some host PM H⁺-ATPase isoforms show high activity in arbuscular mycorrhizal associations (Ramos et al., 2005; Rosewarne et al., 2007). Obviously, H⁺-pumping activity was activated by Cd²⁺ shock and ST exposure, as the H⁺ efflux in MAJ- and NAU-ectomycorrhizal roots were significantly higher than the NM roots (Figures 1C and 5). Increased abundance of HA2.1 and AHA10.1 encoding PM H⁺-ATPase in ectomycorrhizas compared to NM roots of P. × canescens were suggested to lead to higher activities of PM H⁺-ATPases (Ma Y. et al., 2014). The highly activated PM H⁺-ATPase, on the one hand maintains a more suitable acidic environment to promote the Cd²⁺ and Ca²⁺ influx across the PM (Figure 10) and on the other hand, provides an electrochemical H⁺ gradient for PM hyperpolarization, thus increasing Cd²⁺ influx via HACCs. Accordingly, the Cd²⁺-stimulated Cd²⁺ and Ca²⁺ in the P. involutus mycelia (Figures 1–3) was associated with the activated H⁺ pumps since Cd²⁺ treatment markedly upregulated the transcription of PM H⁺-ATPase 1 (Jacob et al., 2004).

Importantly, the H₂O₂ produced in the ectomycorrhizal associations may accelerate the Cd²⁺ through the PM H⁺-ATPase-mediated HACCs. Whole-cell patch clamp recordings of Arabidopsis guard cells showed that the PM hyperpolarization only activates Ca²⁺ currents in the presence of H₂O₂ (50 μM to 5 mM), and the Ca²⁺ current amplitudes increase with increasing H₂O₂ concentrations (Pei et al., 2000). Demidchik et al. (2007) showed that application of H₂O₂ (10 mM) to the external PM face of elongation zone epidermal protoplasts resulted in the appearance of a hyperpolarization-activated Ca²⁺ permeable conductance. In mature epidermal protoplasts, PM HACCs were activated only when H₂O₂ was present at the intracellular membrane face, and channel opening probability increased with intracellular H₂O₂ concentrations at hyperpolarized voltages (Demidchik et al., 2007). A massive presence of H₂O₂ was demonstrated in the outer hyphae mantle of P. involutus symbiosis (Gafur et al., 2004; Langenfeld-Heyser et al., 2007) and obviously could be released from the hyphae into the surrounding medium (Figure 6). Therefore, we suppose that in ectomycorrhizal P. × canescens, H₂O₂ elicited by fungal colonization stimulated Cd²⁺ influx through the HACCs that had been activated by P. involutus colonization. In addition, we found that Cd²⁺ influx in NAU-roots was less restricted than in MAJ-roots by DMTU and sodium orthvanadate (Figures 9B and 11B). The difference in the sensitivity to antagonists of H₂O₂ and PM H⁺-ATPase indicates the involvement of voltage-independent Ca²⁺ channels (VICCs) in the mediation of Cd²⁺ uptake in NAU-roots, in addition to the dominant Cd²⁺ entry through HACCs.

We noticed that LT stress in hydroponic conditions caused a pronounced shift of H⁺ efflux toward an influx in EM roots (Figure 5). LT-stressed P. involutus mycelia exhibited a trend similar to that in EM roots (Figure 5). These results imply that ectomycorrhization activated an H⁺/Cd²⁺ antiport to reduce excessive Cd²⁺ uptake and accumulation under prolonged stress conditions (Sun et al., 2013b). Similarly, we have previously shown that NaCl-treated P. euphratica roots retain an active PM Na⁺/H⁺ antiport to avoid the excessive buildup of Na⁺ when exposed to LT salinity (Sun et al., 2009a,b). Here, the rate of H⁺/Cd²⁺ antiport could not be determined, because our NMT data only show the net flux of the target element across the PM, instead of an unidirectional flux. In addition, EM roots were able to avoid the ROS burst in Cd²⁺ environments (Figure 6), probably because these roots were characterized by elevated H₂O₂ production (Gafur et al., 2004). Therefore, EM roots are likely to control the Cd²⁺ influx through the H₂O₂-activated PM Ca²⁺ channels, thus avoiding an excessive accumulation of the heavy metal ions under prolonged period of Cd²⁺ stress.

Conclusion

High external Cd²⁺ facilitates the rapid movement of Cd²⁺ along its electrochemical gradient into fungal and plant cells. Based on pharmacological evidence, we conclude that Cd²⁺ ions mainly penetrated the ectomycorrhizal fungal hyphae and poplar roots through PM Ca²⁺ channels. Because the entry of Cd²⁺ could not be fully blocked by various Ca²⁺ channel inhibitors (LaCl₃, GdCl₃, verapamil, and TEA), our results indicate that Cd²⁺ ions also entered the root and fungal cells through other metal transporters or channels. Our flux measurements show that the Cd²⁺-permeable Ca²⁺ channels were activated by H₂O₂ and H⁺-pumping activity. Altogether based on the current and literature data, we propose a signaling pathway that triggers Ca²⁺-channel-mediated Cd²⁺ influx in NM P. × canescens roots and explains the pronounced Cd²⁺ stimulation in ectomycorrhizal associations under Cd²⁺ stress. As shown in Figure 12, the Cd²⁺-elicited H₂O₂ and active H⁺-pumps favored the Cd²⁺ influx through Ca²⁺ channels in NM roots and P. involutus-ectomycorrhiza, while these channels mediate Ca²⁺ influx in the absence of Cd²⁺ stress. In ectomycorrhizas, Cd²⁺ enriched in hyphae is thought to be delivered to the host roots. Moreover, the colonization of P. × canescens roots with the fungal strains MAJ and NAU stimulates H₂O₂ production and increases H⁺-pumping activity, and thus accelerates Cd²⁺ entry through Ca²⁺ channels, in particular through HACCs, under excessive Cd²⁺. Cd²⁺ ions competitively enter Ca²⁺ channels, and thus diminish the entry of Ca²⁺, leading to a marked Cd²⁺ enrichment in ectomycorrhizal roots under Cd²⁺ stress.

FIGURE 12.

Schematic models showing Cd²⁺ influx through plasma membrane (PM) Ca²⁺ channels that stimulated by H₂O₂ and H⁺-ATPase in Paxillus involutus-ectomycorrhizal (MAJ and NAU) and non-mycorrhizal (NM) Populus × canescens roots under Cd²⁺ stress. High external Cd²⁺ facilitates the rapid movement of Cd²⁺ along its electrochemical gradient into fungal and plant cells. Cd²⁺ ions penetrated the ectomycorrhizal fungal hyphae and poplar roots through PM Ca²⁺ channels and other metal transporters or channels. The PM Ca²⁺ channels mediate the entry of Ca²⁺ in the absence of Cd²⁺ (-Cd) while allow the entry of Cd²⁺ in the presence of Cd²⁺ ions (+Cd). The Cd²⁺-permeable Ca²⁺ channels were activated by H₂O₂ and H⁺-pumping activity. Thus the Cd²⁺-elicited H₂O₂ and active H⁺-pumps favored the Cd²⁺ influx through Ca²⁺ channels in NM roots and P. involutus-ectomycorrhizas. In ectomycorrhizas, Cd²⁺ enriched in hyphae is thought to be delivered to the host roots. Moreover, the colonization of P. × canescens roots with the fungal strains MAJ and NAU stimulates H₂O₂ production and increases H⁺-pumping activity, and thus accelerates Cd²⁺ entry through Ca²⁺ channels under excessive Cd²⁺. Cd²⁺ ions competitively enter Ca²⁺ channels, and thus diminish the entry of Ca²⁺, leading to a marked Cd²⁺ enrichment in ectomycorrhizal roots under Cd²⁺ stress.

Author Contributions

YhZ and SC conceived the original screening and research plans; SC supervised the experiments; YhZ, GS, YnZ, ZZ, and NL performed most of the experiments; SD, JS, JL, JY, NZ, RZ, and XM provided technical assistance to YhZ, GS and YnZ; YhZ designed the experiments and analyzed the data; YhZ conceived the project and wrote the article with contributions of all the authors; SC and AP supervised and complemented the writing. All authors have read and approved the manuscript.

Conflict of Interest Statement

The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Supplementary Material

The Supplementary Material for this article can be found online at: http://journal.frontiersin.org/article/10.3389/fpls.2016.01975/full#supplementary-material