The L. japonicus homozygous ha1-1 mutant line with a LORE1 insertion in the HA1 gene (gene ID: LotjaGi3g1v0066100.1 in the Lotus Base Gifu v1.2) and the wild-type (WT) segregant were selected from a heterozygous LORE1 insertion line (plant ID: 30006854) that was obtained from Lotus Base.¹ Genotyping was performed using HA1-specific primers (Supplementary Table S1) combined with the P2 internal LORE1 primer (Fukai et al., 2012; Urbański et al., 2012). A two-compartment culture system consisting of root-hyphal (RHC) and hyphal (HC) compartments was used to cultivate plants (Figure 1). The two compartments were separated by a three-layered barrier (Kobae et al., 2014) comprising an RHC filter (57μM opening), a medial mesh (1mM in thickness; 2mM opening), and an HC filter (32μM opening), which prevented plant roots from passing through but allowed AM fungal hyphae to pass. The compartments were filled with autoclaved river sand (particle size, 0.5–2.0mM). L. japonicus seeds were surface sterilized with a hypochlorite solution, sown on a wetted filter paper in a Petri dish, and germinated at 26°C for 2days in dark and 3days in light. Seedlings were transplanted to RHC and inoculated with 500 spores of Rhizophagus irregularis DAOM 197198 (Mycorise, Premier Tech, Rivière-du-Loup, Canada) per plant. The inoculated and non-inoculated plants were grown in a growth chamber (photoperiod: 16h, temperature: 26°C, and photosynthetic photon flux density: 150μMolm⁻² s⁻¹) for 4weeks. RHC was supplied with a half-strength Hoagland’s solution containing a low concentration of KH₂PO₄ (100μM) every 2days to promote AM colonization. HC was provided with a high-nutrient solution with 500μM KH₂PO₄ to enhance P translocation from the HC via AM fungal hyphae. The amount of applied fertilizer that did not flow from the HC into the RHC was determined before cultivation.

Fresh mycorrhizal roots from at least three independent plants were cut into 5–10mM fragments, placed in plastic molds, and covered with the O.C.T. compound (Sakura Finetek, Tokyo, Japan). The roots were frozen in dry ice-isopentane and stored at −80°C. The frozen roots were sectioned longitudinally at a thickness of 25–30μM using a cryostat (Leica, Wetzlar, Germany). Cryosections were placed on an adhesive glass slide (Frontier, Matsunami, Osaka, Japan) and dried at room temperature for 60min. These sections were used for DAPI staining and enzyme cytochemistry of phosphatase activity.

Mycorrhizal roots from at least three independent plants of each genotype were cut into 5mM fragments using a razor blade in a Petri dish on ice. The root fragments were placed in aluminum specimen carriers with 0.05M sucrose as a cryoprotectant and then frozen using a Leica EM PACT2 high-pressure freezer. The frozen samples were temporarily maintained in liquid nitrogen and transferred into vials containing 1% glutaraldehyde in anhydrous acetone that was precooled to −80°C. The samples were freeze-substituted at −80°C for 48h and gradually warmed to 4°C over a period of 21h using a Leica EM AFS2. Roots were washed twice with anhydrous acetone for 15min and twice with propylene oxide for 5min at room temperature. The samples were infiltrated with Spurr’s resin: propylene oxide mixture of ratios 1:2, 1:1, 2:1, 3:1, and 4:1 for 1h in each mixture sequentially (Spurr’s resin from Polysciences, PA, United States). Final soak was thrice in pure resin for 3h, and samples were polymerized at 70°C for 12h. The embedded roots were trimmed and sectioned longitudinally using an ultramicrotome. For fluorescence microscopy, semithin sections, approximately 200nm thick, were cut with a knife (XAC, SYNTEC, Yokohama, Japan) and placed on an adhesive glass slide. For transmission electron microscopy (TEM), ultrathin sections of approximately 90nm thickness were cut with a diamond knife (DiATOME, PA, United States) and picked up on nickel grids. These sections were immediately processed for polyP labeling.

Cryosections of fresh roots and semithin sections of resin-embedded roots were used for DAPI staining. Cryosections were washed gently with 70% ethanol, 25% ethanol, and distilled water (DW) sequentially for 3min in each solution. Semithin sections were etched with 0.2% sodium ethoxide in ethanol for 5min and then washed twice with 100% ethanol, 70% ethanol, and 25% ethanol sequentially for 3min in each solution. Root samples were stained with 80μgml⁻¹ DAPI in 70% ethanol for 60min at room temperature in the dark. After washing with 70% ethanol and DW, the sections were mounted in DW and observed within 1h. Fluorescence microscopy was performed using an Axio Imager D1 microscope (Carl Zeiss, Jena, Germany). DAPI-polyP fluorescence was excited with UV light, and the emitted fluorescence was detected using a long-pass filter, LP420. Digital images were captured with a digital CCD camera (AxioCam MRc5, Carl Zeiss) operated with AxioVision (Carl Zeiss). After randomly selecting images of mature arbuscules, the areas showing the DAPI-polyP signal in the arbuscules were measured using ImageJ software (Schneider et al., 2012).

PolyP labeling using E. coli PPBD was performed according to the method described by Saito et al. (2005) and Kuga et al. (2008) with some modifications. For fluorescence microscopy, semithin sections of the resin-embedded roots were etched with 0.2% sodium ethoxide in ethanol for 5min and then washed with 0.05% sodium ethoxide followed by 100% ethanol, 50% ethanol, and DW wash. The sections were blocked with 1% bovine serum albumin (BSA) in Tris-buffered saline (TBS) containing low concentrations of salts (TBS-low salt; 25mM Tris-HCl pH 7.4, 13.7mM sodium chloride, and 0.27mM potassium chloride) for 10min. Sections were first incubated in a mixture of 20μgml⁻¹ PPBD, 10μgml⁻¹ mouse anti-Xpress epitope antibody (#R910-25, Thermo Fisher Scientific, MA, United States), TBS-low salt, and 1% BSA for 2h at room temperature. The sections were then washed with TBS-low salt buffer containing 0.05% Triton X-100, followed by TBS-low salt. Samples were incubated with a goat anti-mouse IgG antibody conjugated with Alexa Fluor 488 (#A28175, Thermo Fisher Scientific) diluted to ratio 1:100 in TBS-low salt containing 1% BSA for 1h at room temperature. The sections were washed with TBS-low salt containing 0.05% Triton X-100, followed by TBS-low salt and DW. Alexa Fluor 488 fluorescence was excited with a band-pass filter BP470/40, and the emitted fluorescence was detected with BP525/50 using an epifluorescence microscope Axio Imager D1. Digital images were captured with a digital CCD camera (AxioCam MRm, Carl Zeiss) operated with AxioVision.

For TEM observation, ultrathin sections of the resin-embedded roots were etched with 0.2% sodium ethoxide in ethanol for 5min. They were then washed with 0.02% sodium ethoxide, followed by 100% ethanol, 50% ethanol, and DW wash. Specimens were blocked with 1% BSA in phosphate-buffered saline (PBS) containing low concentrations of salts (PBS-low salt; 10mM phosphate buffer pH 8.4, 13.7mM sodium chloride, and 0.27mM potassium chloride). They were then incubated in a mixture of 20μgml⁻¹ PPBD, 10μgml⁻¹ mouse anti-Xpress epitope antibody, PBS-low salt, and 1% BSA, and washed with PBS-low salt buffer containing 0.05% Triton X-100, followed by PBS-low salt. Samples were incubated with a goat anti-mouse IgG antibody conjugated with 6nm colloidal gold (#115-195-146, Jackson ImmunoResearch Laboratories, PA, United States) diluted 1:20 in PBS-low salt buffer containing 1% BSA for 1h at room temperature. The samples were washed with PBS-low salt containing 0.05% Triton X-100, followed by PBS-low salt and DW wash. Specimens were stained with 50% TI-Blue (Nisshin EM, Tokyo, Japan) for 10min, followed by lead citrate staining for 5min, and observed using a TEM (JEOL, Tokyo, Japan) at an accelerating voltage of 80kV. Negative controls were prepared by incubating sections without PPBD. Another negative control was prepared by incubating sections with an excessive competitor (100mM tripolyphosphate) during the first reaction.

Acid phosphatase (ACP) and neutral phosphatase (NTP) activity was detected using TEM with a cerium-based method (Dreyer et al., 2008) with some modifications. Root fragments (5–10mM) were pre-fixed with 2.5% glutaraldehyde in 100mM cacodylate buffer (pH 7.0) for 2h on ice. The fragments were washed twice with cacodylate buffer for 30min. Roots were further cut into 0.5mM fragments. The fragments were sequentially incubated in a pre-incubation buffer [2mM CeCl₃ in 100mM acetate buffer (pH 4.6) and 100mM Tris-HCl buffer (pH 7.4) for acid and NTP activity, respectively] for 1h, a reaction buffer (1mM β-glycerophosphate and 2mM CeCl₃ in acetate or Tris-HCl buffer) for 30min, a pre-incubation buffer for 15min, and an acetate or Tris-HCl buffer for 15min at 37°C. Samples were post-fixed with 1% OsO₄ in cacodylate buffer for 2h on ice, followed by washing with DW thrice for 5min. Roots were dehydrated with an ethanol series (20, 50, 70, 90, and 95% once and 100% thrice) for 20min each at 4°C and incubated twice with propylene oxide for 5min at room temperature. The samples were infiltrated with Spurr’s resin:propylene oxide (1:2, 1:1, 2:1, 3:1, and 4:1) for 1h in each step and thrice with pure resin for 3h, and polymerized at 70°C for 12h. Ultrathin sections on copper grids were stained with gadolinium acetate (Nakakoshi et al., 2011) and lead citrate and observed using a TEM at an accelerating voltage of 80kV. Controls were prepared by incubating sections without β-glycerophosphate in the reaction buffer and without 2mM CeCl₃ in the pre-incubation and reaction buffers.

The localization of phosphatase activity was also analyzed by fluorescence microscopy. Cryosections of mycorrhizal roots were fixed with 0.25% glutaraldehyde in PBS (pH 7.4) for 8min at 4°C. They were then permeabilized by immersion in 25% ethanol in 100mM acetate buffer (pH 4.6) and 100mM Tris-HCl buffer (pH 7.4) for several seconds for ACP and NTP activity, respectively, and washed twice with the same buffer. The sections were incubated with 25μM ELF97 phosphatase substrate (Thermo Fisher Scientific) in either acetate or Tris-HCl buffer for 30min at room temperature in the dark and washed with the same buffer without the substrate. Fluorescence of the ELF97 reaction product was excited with UV, and the emitted fluorescence was detected with a long-pass filter, LP420, using an epifluorescence microscope Axio Imager D1 microscope.

For dual labeling of polyP accumulation and phosphatase activity, cryosections of mycorrhizal roots were sequentially washed with 70% ethanol, 25% ethanol, and DW. First, sections were labeled with 80μgml⁻¹ DAPI in either 100mM acetate buffer (pH 4.6) or 100mM Tris-HCl buffer (pH 7.4) for 60min at room temperature in the dark. The samples were then gently washed in either acetate buffer or Tris-HCl buffer. Subsequently, sections were incubated with 25μM ELF97 in acetate buffer or Tris-HCl buffer for 30min at room temperature in the dark. After the second labeling, the sections were mounted in DW, covered with a glass coverslip, and observed within 30min. The fluorescence of the DAPI-polyP complex and the ELF97 reaction product was excited with UV, and the emitted fluorescence was detected with a long-pass filter, LP420, using the Axio Imager D1 microscope. Digital images were captured using an AxioCam MRc5 operated with AxioVision.

Roots were cleared in a 10% (w/v) KOH solution at 90°C for 10min, acidified with a 2% (v/v) HCl solution for 5min, and then stained with 0.05% trypan blue in lactic acid at 90°C for 10min. AM colonization was assessed using the method described by Trouvelot et al. (1986) with some modifications. Approximately 10 root fragments per plant were mounted on a slide and observed under a light microscope. The intensity of AM fungal colonization and arbuscule abundance in a field of view (diameter: 1.28mM) was categorized into six and three classes, respectively. AM colonization parameters (F%, M%, m%, A%, and a%) were calculated based on the scores in 100 fields of view. To visualize the fine structure of AM fungal colonization, roots were also stained with wheat germ agglutinin (WGA) conjugated with Oregon Green 488 (Kojima et al., 2014). Fluorescent images of the mycorrhizal roots were captured using an Axio Imager D1 microscope equipped with a digital CCD camera AxioCam MRc5. The length of the major axis of arbuscules was measured with the AxioVision software.

The plants were divided into shoots and roots and dried at 70°C for 48h and were weighed after cooling in a desiccator. Dried samples were digested with sulfuric acid and hydrogen peroxide at 200°C for 2h. P concentrations in the samples were determined using the acid-molybdate blue method (Watanabe and Olsen, 1965). The P content was calculated by multiplying each sample’s dry weight by their P concentrations.

Total RNA was isolated from roots using RNAiso Plus (Takara Bio, Shiga, Japan) combined with Fruit-mate (Takara Bio) according to the manufacturer’s instructions. Contaminating genomic DNA was eliminated using a TURBO DNA-free kit (Thermo Fisher Scientific). cDNA was synthesized using a ReverTra Ace qPCR RT Kit (Toyobo, Osaka, Japan). Quantitative PCR was conducted using a StepOne Real-Time PCR System (Thermo Fisher Scientific) with a THUNDERBIRD SYBR qPCR Mix (TOYOBO). The primers used for qRT-PCR are listed in Supplementary Table S1. The L. japonicus elongation factor 2 gene and R. irregularis EF1β (Kobae et al., 2015) genes were used as internal controls for the expression analysis of plant and AM fungal genes, respectively. Melting curve analysis confirmed the presence of single peaks. Relative expression levels were calculated using the 2^−ΔCt method (Schmittgen and Livak, 2008). All the reactions were performed using three biological replicates. RT-PCR was performed using HA1-specific primers (Supplementary Table S1) and 10ng of cDNA template to confirm homozygous LORE1 insertion lines at the transcript level, and the amplified products were sequenced.

All statistical analyses were performed using the software R (version 4.0.0). The mycorrhizal colonization rate and relative gene expression values were logit-transformed (ln[p/1−p]) and log-transformed, respectively, to avoid violating normality and homoscedasticity assumptions. To test the differences in mycorrhizal colonization and gene expression between WT and ha1-1 mutants, data were analyzed using the Student’s t-test. Tukey’s HSD test was performed for multiple comparisons of P contents of plants. Relative frequency densities of arbuscule size and the percentage of arbuscule area showing DAPI-polyP signal were calculated using the MASS package in the software. The size distributions of WT and ha1-1 mutants were compared using a two-sample Kolmogorov–Smirnov test. The proportion of mature and collapsed arbuscules in WT and ha1-1 was analyzed using Fisher’s exact test.

First, we investigated the effect of the mutation of HA1 on P acquisition through the mycorrhizal pathway. A L. japonicus homozygous line carrying a LORE1 insertion in exon 8 of the HA1 gene was selected to obtain a ha1 mutant, ha1-1 (Figure 1A). RT-PCR analysis revealed that although HA1 transcripts were observed in AM roots of the ha1-1 mutant (Figure 1B), they had indels caused by putative abnormal splicing that created a frameshift and a premature stop codon 13 amino acids after the insertion (Figure 1C). To determine whether the HA1 mutation affects P acquisition via the mycorrhizal pathway, we examined the P nutrition of the ha1-1 mutant using a two-compartment system consisting of RHC and HC (Figure 1D). Under uninoculated conditions, plant’s P nutrition did not differ between WT and ha1-1 plants (Figure 1E). In the presence of AM fungi, P content increased in both genotypes. However, the positive effects on shoot P content were lower in the ha1-1 mutant (average increase of 268%) than in the WT (average increase of 492%). These data demonstrate that P transfer from the AM fungus to the host via the mycorrhizal pathway was partially impaired in the ha1-1 mutant. To examine the effects of ha1-1 on hyphal development within roots, we assessed AM fungal colonization by staining mycorrhizal roots with trypan blue and WGA–Oregon Green 488. Quantitative analysis revealed that arbuscule densities (A% and a%) were moderately reduced in the mutant relative to the WT 4weeks post-inoculation (Figure 1F). Furthermore, the arbuscules formed in ha1-1 mutant roots were slightly smaller than those in WT roots (Figure 1G). However, the range of arbuscule sizes almost overlapped between the two genotypes. These results suggest that the HA1 mutation induced a phase transition of the arbuscule life cycle from mature to degrading stage.

We analyzed polyP accumulation in arbuscules by DAPI staining. Once DAPI binds to polyP, the complex emits yellow fluorescence under UV irradiation (Tijssen et al., 1982). Mycorrhizal root cryosections were DAPI-stained after polyP fixation and cell permeabilization with ethanol. We observed arrays of fully-developed arbuscules (mature arbuscules) and the patchy colonization of degenerating arbuscules which are small arbuscules with shrunken fine branches (Figure 2A). In most mature arbuscules formed in cortical cells, the fluorescent DAPI-polyP complex signal was confined to the central region, irrespective of the plant’s genotype. The other region within the arbuscules, containing fine branches, exhibited non-specific blue fluorescence. In contrast, the entire hyphae of the degenerating arbuscules were stained with DAPI and fluoresced yellow. This type of arbuscules accounted for 57% of the total arbuscules formed in ha1-1 mutants, which was 1.4-fold higher than that in the WT (Figure 2B). This result prompted us to examine whether the shift in size distribution toward small arbuscules in the mutant, as observed by WGA staining was due to the increased number of degenerating arbuscules. We measured the size of mature and degenerating arbuscules distinguished by the DAPI staining pattern. Overall, WGA staining reduced arbuscule size relative to DAPI, possibly due to cell shrinkage caused by fixation with Farmer’s solution which contained ethanol and acetic acid (Figure 1G and Figure 2C). In both arbuscule types, the size distribution almost overlapped between the WT and the ha1-1 mutant, but the median length was 5–8μM shorter in the mutant (Figure 2C). Our DAPI staining data also suggested that HA1 mutations caused early arbuscule degeneration.

Next, we focused on the distribution of polyP in mature arbuscules. It should be noted that the size of the DAPI-stained region in the arbuscules varied (Figure 3A). Quantitative analysis revealed that the distribution of the yellow fluorescent area relative to the entire mature arbuscule significantly differed between the WT and ha1-1 (Figure 3B). In the WT, the most abundant size class was 0–5%; arbuscules with large DAPI-stained regions were infrequent. In contrast, the size distribution in the ha1-1 mutant peaked at 10–15% with a higher right tail than the WT, indicating an expansion of the DAPI-polyP positive region.

To obtain more precise polyP localization in mature arbuscules, we analyzed semithin sections of resin-embedded AM roots. Arbuscules develop with repeated branching and narrowing of hyphal diameter from a thick trunk hypha to fine branches. Here, we define the fine branch module as a set of connected branches (Figure 3C). For arbuscules generated in WT roots, yellow fluorescence was detected in some fine branch modules located in the vicinity of the trunk hyphae (Figure 3D). The fluorescent signal was also visible around the periphery of the trunk hyphae where the cell wall was localized, but not inside these hyphae. Similarly, the ha1-1 mutant displayed DAPI-polyP signals in fine branch modules in the central region of the arbuscule and cell walls of the trunk hyphae.

We further analyzed the subcellular localization of polyP using enzyme-affinity labeling with PPBD, which binds specifically to long-chain polyP (Saito et al., 2005), because the yellow fluorescence of DAPI is not specific only to polyP. PolyP was labeled in semithin sections of mycorrhizal roots with recombinant PPBD containing an epitope tag. The tag was then detected by indirect immunocytochemistry with an Alexa Fluor 488-conjugated secondary antibody and observed under a fluorescence microscope. In arbuscules formed in WT roots, a strong polyP signal was observed at the cell periphery of the trunk hyphae, possibly at the cell wall (Figure 4). Weak fluorescence was detected in plant and fungal nuclei, likely due to the low-affinity DNA-binding capability of PPBD (Saito et al., 2005). There was no or very weak polyP signal in the fine branches throughout the cortical cells harboring arbuscules. This localization differed from the DAPI-polyP signal, which was associated with fine branch modules in the central region of arbuscules. These contradictory observations may result from a difference in polyP binding properties as DAPI can bind to short-chain polyP with at least 14 residues (Smith et al., 2018), but PPBD has an extremely low affinity for polyP shorter than 35 residues (Saito et al., 2005). In addition, polyP detected by DAPI was immobilized in ethanol. Conversely, the PPBD enzyme-affinity method included incubation in an aqueous solution that might have eluted some polyP from the sections. In the ha1-1 mutant, the labeling pattern of fungal polyP was similar to that of the WT, in which the cell periphery of trunk hyphae was labeled (Figure 4). However, we found that polyP signals were occasionally present at the cell periphery of some fine branches.

PolyP localization was visualized by TEM using the enzyme-affinity method with gold-coupled secondary antibodies. In arbuscules formed in WT, the polyP signal was evenly distributed in the trunk cell wall, as observed with fluorescence microscopy (Figures 5A–C). Sparse labeling was observed sporadically within vacuoles of trunk hyphae (Figure 5D). However, there was no polyP signal in the fine branches (Figures 5E–F). In ha1-1 roots, polyP localization was similar to that in the WT, with presence in the trunk hyphae fungal cell wall (Figures 5G–H). Interestingly, polyP signals were sometimes observed on the fine branch cell walls (Figures 5I–J). In particular, fine branches cut obliquely against the longitudinal axis showed prominent signals in their cell walls or their surrounding PAS (Figures 5K–L). In the negative controls, where sections of mycorrhizal roots were incubated in a reaction mixture without PPBD, no polyP signals were detected (Supplementary Figure S1). There was also no polyP signal when the PPBD polyP-binding site was masked with a high concentration of tripolyphosphate. In summary, enzyme-affinity labeling using PPBD revealed that in the WT, relatively long-chain polyP was mainly distributed in the trunk hyphae cell walls and absent in the fine branches. However, the HA1 mutation led to polyP accumulation in some fine branch cell walls.

An interesting feature of polyP localization was its almost complete absence from fine branches at the periphery of cortical cells in WT and ha1-1 (Figure 3). Dreyer et al. (2008) reported that ACP activity localizes at the interface between fungal cell walls and PAM. Generally, ACPs have broad substrate specificities for phosphate esters. To elucidate the spatial relationship between polyP and ACP in cells with arbuscules, we investigated the localization of ACP activity at the ultrastructural level (Figure 6). ACP activity was detected by incubating mycorrhizal roots in an acidic buffer containing the reaction substrate (β-glycerophosphate) and the co-precipitant (cerium salt) and observing the black precipitates of the reaction product (cerium phosphate) with TEM. High ACP activity was frequently detected in PAS around the fine branches in both WT and ha1-1, which was consistent with previous observations (Dreyer et al., 2008). Notably, most of the dense precipitates localized along the host-derived PAM and in small vesicles resembling the intramatrix compartment type I (IMC-I) or apoplastic vesicular structures (AVS; Ivanov et al., 2019; Roth et al., 2019). In contrast, only a few signals were associated with PAS around the trunk hyphae in both WT and ha1-1 cells. ACP activity was sometimes observed in fungal vacuoles. No dense cerium phosphate precipitate was observed in the control experiment lacking cerium salt (Supplementary Figure S2). In the control reaction without β-glycerophosphate, a few precipitates were observed in the fungal vacuoles, possibly due to a reaction with endogenous vacuole substrates. Next, we investigated the localization of NTP activity in the ha1-1 mutant because the acidification around arbuscules diminishes in ha1 cortical cells (Krajinski et al., 2014; Liu et al., 2020). NTP activity localization was similar to ACP activity with signals along the PAM and in small vesicles present in the PAS surrounding fine branches (Figure 7), as previously reported (Jeanmaire et al., 1985).

To further study the relationship between phosphatase activity and polyP accumulation, we detected phosphatase activity and polyP signals simultaneously by enzyme cytochemistry using the ELF97 phosphatase substrate and DAPI staining, respectively. ELF97 forms fine precipitates after hydrolysis of its phosphate ester bond by non-specific phosphatases, emitting yellow-green fluorescence at the site of phosphatase activity (Haugland, 2005). First, ACP and NTP activities were visualized in a typical mature arbuscule by single ELF97 staining in WT and ha1-1 roots, respectively (Figure 8A). In the WT, ACP activity was present throughout the arbuscule but was excluded from its central region. Similarly, NTP activity was detected in arbuscules in the mutant but the central region without phosphatase activity was larger than that in the WT. Next, we performed double labeling of phosphatase activity and polyP. The localization of phosphatase activity (green) and polyP (yellow) was distinct based on different emission colors using a long-pass filter, albeit showing weak and different color signals compared to the single staining (Figure 8B). Double labeling showed that polyP was present in the center of arbuscules, and phosphatase activity localized in the surrounding regions. Thus, polyP and phosphatase activity showed opposite localization in both WT and ha1-1 cells.

We investigated whether the HA1 mutation affected gene expression related to plant Pi uptake and fungal polyP metabolism during AM symbiosis. The levels of the phosphate transporter PT1, likely to function in the direct pathway due to its downregulation during mycorrhization (Maeda et al., 2006), were slightly elevated in ha1-1 roots compared to the WT (Figure 9A), which may reflect a partial block in P translocation via the mycorrhizal pathway. Transcripts of other phosphate transporters, including AM-specific PT4 (Guether et al., 2009a; Takeda et al., 2009), accumulated equally in the mycorrhizal roots of both genotypes. Similarly, the ha1-1 mutation did not affect the expression of AM marker genes, AM-specific ammonium transporter AMT2;2 (Guether et al., 2009b) and glycerol-3-phosphate acyltransferase RAM2 (Wang et al., 2012). Fungal endopolyphosphatase genes PPN1, PPN2, and PPN3 were downregulated in AM fungi colonizing the mutant, whereas the VTC genes involved in polyP synthesis were not affected (Figure 9B).