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. 2020 Mar 11;183(1):250–262. doi: 10.1104/pp.20.00078

CASEIN KINASE2-Dependent Phosphorylation of PHOSPHATE2 Fine-tunes Phosphate Homeostasis in Rice1

Fei Wang a, Meiju Deng a, Jieyu Chen b, Qiuju He a, Xinye Jia a, Huaxing Guo a, Jiming Xu a, Yidong Liu c, Shuqun Zhang c, Huixia Shou a, Chuanzao Mao a,2,3
PMCID: PMC7210639  PMID: 32161109

The casein kinase II phosphorylates PHO2, causing it to be degraded more quickly, which affects its role in targeting phosphate transporter for degradation, thereby maintaining phosphate homeostasis in rice.

Abstract

Plants have evolved complex physiological and biochemical mechanisms to adapt to a heterogeneous soil phosphorus environment. PHOSPHATE2 (PHO2) is a phosphate (Pi) starvation-signaling regulator involved in maintaining Pi homeostasis in plants. Arabidopsis (Arabidopsis thaliana) PHO2 targets PHOSPHATE TRANSPORTER1 (PHT1) and PHO1 for degradation, whereas rice (Oryza sativa) PHO2 is thought to mediate PHOSPHATE TRANSPORTER TRAFFIC FACILITATOR1 degradation. However, it is unclear whether and how PHO2 is post-translationally regulated. Here, we show that in rice, the CASEIN KINASE2 (OsCK2) catalytic subunit OsCK2α3 interacts with OsPHO2 in vitro and in vivo in vascular tissues cells, and phosphorylates OsPHO2 at Ser-841. Phosphorylated OsPHO2 is degraded more rapidly than native OsPHO2 in cell-free degradation assays. OsPHO2 interacts with OsPHO1 and targets it for degradation through a multivesicular body-mediated pathway. PHO1 mutation partially rescued the pho2 mutant phenotype. Further genetic analysis showed that a nonphosphorylatable version of OsPHO2 rescued the Ospho2 phenotype of high Pi accumulation in leaves better than native OsPHO2. In addition to the previously established role of OsCK2 in negatively regulating endoplasmic reticulum exit of PHT1 phosphate transporters, this work uncovers a role for OsCK2α3 in modulating Pi homeostasis through regulating the phosphorylation status and abundance of OsPHO2 in rice.

Phosphorus (P) is an indispensable macronutrient for plant growth and development. Plants absorb P from the surrounding soil in the form of inorganic phosphate (Pi; Chiou and Lin, 2011), but due to its high chemical fixation rate and slow diffusion properties, Pi is commonly present at low concentrations even when P is abundant in soil (Raghothama, 1999). Therefore, plants have evolved intricate physiological and molecular regulatory mechanisms to increase the absorption, utilization, and remobilization of Pi to maintain P homeostasis (Ham et al., 2018; Srivastava et al., 2018; Wang et al., 2018; Pan et al., 2019).

PHOSPHATE2 (PHO2) is an important modulator of plant Pi homeostasis. The first pho2 mutant was identified in Arabidopsis (Arabidopsis thaliana), which overaccumulated Pi in leaves under Pi-replete conditions (Delhaize and Randall, 1995; Dong et al., 1998). PHO2 encodes a ubiquitin-conjugating (E2) enzyme (UBC24) that directly and/or indirectly degrades high-affinity PHOSPHATE TRANSPORTERs, which take up Pi from the soil, and PHO1, which loads Pi into the xylem for distribution from root to shoot in Arabidopsis (Liu et al., 2012; Huang et al., 2013; Park et al., 2014). AtPHO2 mainly localizes to the endoplasmic reticulum (ER) and Golgi (Huang et al., 2013), and AtPHO2 transcription is regulated by the microRNA miR399, which is induced by Pi starvation in the vascular cylinder (Aung et al., 2006).

Although much is known about the regulatory system of PHO2 in Arabidopsis, its role in rice (Oryza sativa) is largely unknown. OsPHO2/OsLTN1, the AtPHO2 homolog in rice, also plays an important role in Pi homeostasis downstream of miR399 and is involved in regulating multiple Pi starvation responses in rice (Hu et al., 2011; Cao et al., 2014). OsPHO2 transcripts are also detected mainly in vascular tissues (Hu et al., 2011). The Ospho2/ltn1 mutant shows enhanced Pi uptake and increased Pi translocation from root to shoot and displays leaf tip necrosis (Hu et al., 2011). OsPHO2 activity is modulated by thioredoxins (OsTrxh1 and OsTrxh4), which affect its function in rice (Ying et al., 2017). In a screen for suppressors of Ospho2 in rice, we previously identified the AtPHO1 homologous protein OsPHO1, which may be an OsPHO2 target (Hu et al., 2016). However, systematic work focusing on the control of OsPHO2 protein activity, as well as its targets, is lacking.

Protein phosphorylation, one of the most important post-translational modifications in eukaryotes, can act as a molecular switch or rheostat for protein function (Vlastaridis et al., 2017). We previously demonstrated that in rice, the ER-located CASEIN KINASE2 (OsCK2) holoenzyme (OsCK2α3/β3) phosphorylates the phosphate transporter OsPT8 at Ser-517 under Pi-sufficient conditions, thereby preventing it from interacting with PHOSPHATE TRANSPORTER TRAFFIC FACILITATOR1 (OsPHF1; Chen et al., 2015). By contrast, under Pi-deficient conditions, the OsCK2β3 subunit is degraded, and unphosphorylated OsPT8 is transported to the plasma membrane (PM) with the help of OsPHF1, thereby increasing Pi uptake and maintaining cellular Pi homeostasis (Chen et al., 2015). Thus, knockdown of OsCK2α3 or OsCK2β3 leads to significantly higher Pi concentrations in shoots and roots than in wild-type cv. Nipponbare (NIP; Chen et al., 2015). However, whether OsCK2 regulates components involved in the phosphate signaling pathway other than PTs is currently unknown.

In this study, we show that OsCK2α3 interacts with OsPHO2 in vitro and in vivo. OsCK2α3 phosphorylates OsPHO2 at Ser-841 via an atypical CK2 recognition motif, and the resulting phosphorylated OsPHO2 is degraded more rapidly than nonphosphorylated OsPHO2. PHO2S841A/pho2 transgenic lines had lower Pi concentrations and higher shoot biomass than PHO2/pho2 transgenic lines, whereas PHO2S841D/pho2 transgenic lines had higher Pi concentrations and lower shoot biomass than PHO2/pho2 lines, supporting the notion that phosphorylation of OsPHO2 Ser-841 is involved in regulating Pi homeostasis in rice. We also show that OsPHO2 targets OsPHO1 for degradation and regulates Pi homeostasis. Overall, OsCK2α3-dependent phosphorylation of OsPHO2 represents a previously unidentified mechanism in plants for fine-tuning internal phosphate homeostasis.

RESULTS

OsCK2α3 Interacts with OsPHO2

We screened for OsPHO2-interacting proteins in a yeast two-hybrid (Y2H) assay and identified OsCK2α3. We then performed a split-ubiquitin Y2H assay using the DUALmembrane System (Dualsystems Biotech) to examine the possible interactions of OsPHO2 with OsCK2 subunits α2 and α3 or β1 and β3. Among these CK2 components, only OsCK2α3 interacted with OsPHO2 in yeast cells (Fig. 1A). To verify this result, we conducted a pull-down assay using recombinant glutathione s-transferase fused with OsCK2α3 (GST-α3) and HIS-PHO2 purified from Escherichia coli. The recombinant HIS-PHO2 protein bound to GST-α3, but not to GST (Fig. 1B), confirming that OsCK2α3 directly interacts with OsPHO2 in vitro.

Figure 1.

Figure 1.

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OsCK2α3 interacts with OsPHO2. A, Split-ubiquitin Y2H assay. Cub, C-terminal ubiquitin (Cub interacts with NubI but not with the NubG or NubG-CK2 subunits, which served as positive and negative controls, respectively); NubI and NubG, the wild-type and mutated N-terminal fragment of ubiquitin. Yeast cells were grown on control medium (SD/-Leu/-Trp [SD/LW]) or selective medium (SD/-Leu/-Trp/-His/-Ade [SD/LWHA]). B, In vitro pull-down assay. HIS-PHO2, GST-CK2α3 (GST-α3), and GST were expressed and purified in E. coli and subjected to GST pull-down assays. GST, GST-α3, and HIS-PHO2 were detected by immunoblotting using anti-GST and anti-HIS antibodies, respectively. C, Schematic diagrams of full-length PHO2 and its deletion derivatives used in the yeast-two hybrid assay with CK2α3. Numbers at the top refer to the positions of the first or last amino acid in each sequence. UBC, ubiquitin-conjugating domain. D, Interaction between CK2α3 and PHO2 truncations in a Y2H assay. SD/LHW, SD/-Leu-Trp-His; BD, pGBKT7 empty vector; AD, pGADKT7 empty vector.

To determine which part of OsPHO2 interacts with OsCK2α3, we generated three truncated versions of OsPHO2, including an N-terminal sequence without a known conserved domain (PHO2N), the ubiquitin-conjugating enzyme domain (PHO2M), and the C terminus (PHO2C), as shown in Figure 1C, and tested their interactions with OsCK2α3 using the Matchmaker GAL4 Yeast Two-Hybrid System (Clontech). Yeast cells cotransformed with PHO2N and OsCK2α3 grew well on SD/-Leu/-Trp/-His medium supplemented with 10 mm of 3-amino-1,2,4-triazole, indicating that PHO2N interacts with CK2α3 in yeast (Fig. 1D). However, full-length OsPHO2 failed to interact with OsCK2α3 in the nuclear Y2H system, likely because OsPHO2, an endomembrane-localized protein, was not correctly localized to the nucleus.

To investigate whether OsCK2α3 and OsPHO2 have similar subcellular localizations, we transiently expressed GFP-PHO2 and CK2α3-mCherry driven by the 35S promoter in Nicotiana benthamiana leaf epidermal cells. GFP-PHO2 showed reticular and punctate patterns like those of AtPHO2 (Huang et al., 2013), whereas CK2α3-mCherry displayed reticular patterns (Fig. 2A), which is consistent with previous results (Chen et al., 2015). The green fluorescent and red fluorescent signals strongly overlapped (Fig. 2A), indicating that OsCK2α3 colocalizes with OsPHO2.

Figure 2.

Figure 2.

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OsCK2α3 colocalizes with and interacts with OsPHO2. A, Colocalization of 35S-GFP-PHO2 and 35S-CK2α3-mCherry in N. benthamiana leaves. Scale bars = 20 μm. B, In situ PCR analysis of CK2α3, PHO2, and ACTIN in primary rice roots. Blue staining demonstrates the presence of cDNA, while brown indicates the absence of target cDNA within cells. ACTIN was used as a positive control, and a reaction mixture containing all reagents except primers was used as a negative control. ep, epidermis; ex, exodermis; co, cortex; st, stele. Scale bars = 50 μm. C, BiFC analysis of the in vivo interaction between CK2α3 and PHO2. CK2α3 and PHO2 were fused to the N terminus of the N- and C-terminal fragments of YFP (YFPN and YFPC), respectively. Combinations of YFPC or YFPN with the corresponding PHO2 or CK2α3 constructs were used as negative controls. Scale bars = 20 μm.

We performed reverse transcription quantitative PCR (RT-qPCR) analysis to investigate whether OsCK2α3 and OsPHO2 are expressed in the same plant tissues. Both genes were expressed in roots, stem bases, stems, leaf sheaths, leaf blades, and inflorescences, with the highest expression levels in roots and leaf sheaths (Supplemental Fig. S1). We further investigated the tissue-specific expression patterns of the two genes in roots via in situ PCR. OsCK2α3 showed blue staining in all root cells, from the epidermis to the stele, whereas staining of OsPHO2 was detected mainly in the stele (Fig. 2B).

To confirm the interaction between OsCK2α3 and OsPHO2 in vivo, we performed a bimolecular fluorescence complementation (BiFC) assay using N. benthamiana leaves. Coexpression of PHO2-YFPN and CK2α3-YFPC or CK2α3-YFPN and PHO2-YFPC produced strong GFP signals, whereas no detectable fluorescence was produced in the negative controls, such as YFPN/PHO2-YFPC and PHO2-YFPN/YFPC (Fig. 2C), indicating that OsCK2α3 interacts with OsPHO2 in vivo. These results demonstrate that OsPHO2 interacts with OsCK2α3 through its N-terminal domain in vascular tissues.

OsCK2α3 Phosphorylates OsPHO2 In Vitro and In Vivo

The finding that OsCK2α3 and OsPHO2 interact with each other raised the question of which protein affects the other. There are two possible relationships between OsCK2α3 and OsPHO2: (1) As a putative ubiquitin-conjugating enzyme, OsPHO2 might play a role in the degradation of OsCK2α3 or other OsCK2 subunits; and (2) as a kinase, OsCK2α3 might phosphorylate OsPHO2 and further affect its activity, for example, its localization or stability.

To test the first possibility, we performed an in vitro cell-free degradation assay. Equal amounts of purified GST, GST-α3, and GST-β3 proteins were incubated with protein extracts prepared from the roots of 10-d–old wild-type and pho2 (Supplemental Fig. S2) seedlings grown in P-sufficient conditions, followed by 7 d of P-deficiency treatment. OsCK2 protein levels were then assessed by immunoblot analysis. The GST-β3 was almost completely degraded at 60 min in both wild-type and pho2 extracts, while the degradation of GST-α3 was evident only after 180 min (Supplemental Fig. S3), which is consistent with our previous finding that CK2α3 protein is relatively stable while CK2β3 protein is degradable under Pi deficiency (Chen et al., 2015). As both α3 and β3 proteins degraded in the same manner after incubation with wild-type or pho2 plants extracts (Supplemental Fig. S3), we conclude that OsPHO2 does not affect the degradation of OsCK2α3 or OsCK2β3.

To determine whether OsCK2α3 phosphorylates OsPHO2, we performed an in vitro phosphorylation assay using recombinant GST-α3 and HIS-PHO2. As shown in Figure 3A, OsPHO2 was phosphorylated in vitro in the presence of OsCK2α3. To identify the phosphorylation site(s) in OsPHO2, we cut out bands from gels containing PHO2 (HIS-PHO2 incubated with GST) and phosphorylated PHO2 (HIS-PHO2 incubated with GST-α3), extracted the proteins from the bands, and searched for their phosphorylation sites by liquid chromatography tandem mass spectrometry (LC-MS/MS). Three phosphorylation sites (Ser-513, Ser-518, and Ser-841) were detected in the PHO2 sample incubated with GST-α3, but no phosphorylation sites were detected in the PHO2 sample incubated with GST (Supplemental Fig. S4).

Figure 3.

Figure 3.

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CK2α3 phosphorylates PHO2 in vitro and in vivo. A, CK2α3 phosphorylates PHO2 in vitro. The input proteins HIS-PHO2 and GST-CK2α3 (GST-α3) were detected by Coomassie blue staining (right). Phosphorylation activity was detected by [γ-32P] ATP autoradiography (left). P-PHO2 indicates phosphorylated PHO2. B, In vitro phosphorylation assay indicating that Ser-841 is the site phosphorylated by CK2α3. The immunoblots were probed with anti-GST after Phos-Tag SDS-PAGE. Coomassie blue staining of proteins in a standard SDS-PAGE gel is shown as a loading control (lower). PHO2C indicates C-terminal 443 amino acids of PHO2. C, PHO2 is phosphorylated in vivo. Root proteins from 15-d–old PHO2pro-FLAG-PHO2 transgenic plants were separated by SDS-PAGE (top) or Phos-Tag SDS-PAGE (middle) and detected by immunoblot analysis using an anti-FLAG antibody. Open triangle indicates nonspecific bands. CB, Coomassie blue staining. D, PHO2 is phosphorylated by CK2α3 in vivo. Root proteins from 15-d–old α3Ri, α3Ri/PHO2pro-FLAG-PHO2, and PHO2pro-FLAG-PHO2 transgenic plants were separated by SDS-PAGE (top) or Phos-Tag SDS-PAGE (middle) and detected by immunoblot analysis using an anti-FLAG antibody. Open triangle indicates nonspecific bands. CB, Coomassie blue staining.

To validate these phosphorylation sites, we generated a truncated version of PHO2 with 443 C-terminal amino acids containing all three potential phosphorylation sites (designated PHO2C). We then mutated these PHO2C peptides, replacing Ser-513, Ser-518, or Ser-841 with Ala (producing peptides designated PHO2CS513A, PHO2CS518A, and PHO2CS841A, respectively, mimicking nonphosphorylatable forms of PHO2). A detectable phosphorylated protein band was absent in assays with PHO2CS841A, but not with PHO2CS513A or PHO2CS518A, suggesting that Ser-841 is the phosphorylation site of OsPHO2 mediated by OsCK2α3 (Fig. 3B). The phosphorylated band was also detected using a mutated PHO2 protein with both Ser-513 and Ser-518 replaced by Ala (designated PHO2CS513A/S518A), but not in the PHO2CS841A or λ-phosphatase (λ-PPase)-treated PHO2C or PHO2CS513A/S518A variants (Supplemental Fig. S5). These results indicate that OsCK2α3 directly phosphorylates OsPHO2 at Ser-841 in vitro.

To test the phosphorylation of OsPHO2 by OsCK2α3 in vivo, we separated protein extracts from rice plants expressing PHO2pro-FLAG-PHO2 in standard SDS-PAGE or Phos-Tag-containing SDS-PAGE gels. Two bands were detected on the Phos-Tag immunoblots but only one band on standard SDS-PAGE (Fig. 3C). The lower-mobility band was sensitive to λ-PPase treatment (Fig. 3C), suggesting that it contained the phosphorylated form of PHO2. We then generated the α3Ri/FLAG-PHO2 plants by crossing α3Ri with PHO2pro-FLAG-PHO2 plants to detect PHO2 phosphorylation status in a CK2α3 knockdown plant. Immunoblot results showed that the band corresponding to phosphorylated PHO2 was much weaker in the α3Ri background than in the wild-type CK2α3 background (Fig. 3D). These results suggest that OsCK2α3 phosphorylates OsPHO2 in vivo.

Phosphorylation of OsPHO2 Does Not Affect Its Interaction with OsCK2α3 or its Subcellular Localization

Protein phosphorylation is a multifunctional post-translational modification that may alter the structure, function, localization, molecular interactions, or stability of a protein (Nishi et al., 2014). A split-ubiquitin Y2H assay showed that, like native PHO2, PHO2S841A interacted with CK2α3 in yeast (Supplemental Fig. S6A). This result was confirmed by BiFC in N. benthamiana leaves. Coexpression of CK2α3-YFPN with PHO2-YFPC or PHO2S841A-YFPC led to the production of fluorescent signals in N. benthamiana leaves, while the empty vector controls showed no detectable fluorescence (Supplemental Fig. S6B). Analysis of leaves coexpressing CK2α3-YFPN with PHO2S513A-YFPC (as a control) showed that there was no difference in the interaction of CK2α3 with any of the three PHO2 variants (Supplemental Fig. S6). These results demonstrate that OsPHO2 interacts with OsCK2α3 regardless of whether it is phosphorylated by CK2α3. To investigate whether the phosphorylation of OsPHO2 affects its subcellular localization, we coexpressed 35S-GFP-PHO2, 35S-GFP-PHO2S841A, or 35S-GFP-PHO2S841D (phospho-mimicking) with the ER marker AtWAK2-mCherry in N. benthamiana leaves, respectively. The GFP fluorescent signals from GFP-PHO2, GFP-PHO2S841A, and GFP-PHO2S841D overlapped well with the mCherry signals, indicating that the phosphorylation of OsPHO2 does not alter its subcellular location (Supplemental Fig. S7).

Phosphorylated OsPHO2 Is Degraded More Rapidly than Native OsPHO2

In Arabidopsis, PHO2 transcript accumulation is suppressed by Pi limitation (Bari et al., 2006). To explore how OsPHO2 responds to Pi deficiency, we investigated its transcript and protein levels after various periods of Pi-starvation treatment. OsPHO2 transcript levels decreased with increasing Pi-starvation treatment (Supplemental Fig. S8, A and C). Similarly, OsPHO2 protein levels declined under Pi starvation in a time-course–dependent manner (Fig. 4A; Supplemental Fig. S8B). OsPHO2 protein levels began to decrease after 1 d of Pi starvation, and the protein band corresponding to FLAG-PHO2 was almost undetectable after 5 d of Pi starvation (Fig. 4A). Similar to transcript levels, OsPHO2 protein levels did not recover after resupplying the plants with Pi for 1 d (Supplemental Fig. S8, A and B). By contrast, OsPHO2 protein levels remained relatively stable under Pi-sufficient conditions (Fig. 4A). To determine whether the decrease in OsPHO2 protein levels was exclusively due to the reduced transcript levels, we added the de novo protein synthesis inhibitor cycloheximide (CHX) to the nutrient solution for various periods of time, extracted protein samples, and subjected them to immunoblotting. OsPHO2 protein levels gradually decreased with the extension of CHX treatment, and they decreased more rapidly under simultaneous Pi-starvation treatment (Fig. 4, B and C). The half-life of FLAG-PHO2 was ∼12.2 min under +P conditions and ∼6.9 min under −P conditions, demonstrating that OsPHO2 protein was unstable and that Pi starvation accelerated its degradation (Fig. 4). Both MG132, a 26S proteasome inhibitor, and E-64D, an endosomal protease inhibitor, could not inhibit PHO2 from degradation in vitro and in vivo (Supplemental Figs. S8B and S9), suggesting that the PHO2 is not degraded through the ubiquitin-mediated protein degradation pathway.

Figure 4.

Figure 4.

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Pi-dependent stability of OsPHO2. A, PHO2 protein levels under Pi deficiency treatment. Proteins from the roots of 15-d–old PHO2pro-FLAG-PHO2 transgenic plants were detected by immunoblot analysis using an anti-FLAG antibody. Equal amounts of protein (10 μg) were used for immunoblotting. Open triangle indicates nonspecific bands. Protein staining by Coomassie blue (CB) indicates that similar amounts of protein were loaded. B, The half-lives of PHO2 in vivo under +P (200 μm) and –P (0 μm) conditions. Immunoblot analysis shows the abundance of FLAG-PHO2 in 15-d–old seedling roots over a 60-min period of CHX (200 μm) treatment. Open triangles indicate nonspecific bands. C, The relative amount of FLAG-PHO2 remaining after CHX treatment was calculated and plotted on a semilog graph. PHO2 protein level was normalized to that of the 0-min control. Error bars represent ± sd (n = 3 independent experiments with eight plants for each experiment).

As the phosphorylation of OsPHO2 does not affect its localization or its interaction with OsCK2α3, we examined whether it might influence the stability of OsPHO2. We incubated HIS-PHO2, HIS-PHO2S841A, and HIS-PHO2S841D with total proteins extracted from seedling roots grown in high phosphate (HP, 200 μm) and low phosphate (LP, 10 μm) solution at various time points and subjected them to immunoblot analysis. When incubated with protein extracts from HP or LP plants, most HIS-PHO2S841D protein degraded within 15 min, HIS-PHO2 protein levels decreased after 45 min, and HIS-PHO2S841A degraded more slowly, as some was still present after 90 min of treatment (Fig. 5). Maltose Binding Protein (MBP, control) protein levels remained almost constant throughout incubation (Fig. 5). However, the levels of all proteins decreased more quickly when incubated with protein extracts from LP-versus-HP plants (Fig. 5). These results indicate that the phosphorylation of OsPHO2 at Ser-841 reduces its stability independently of Pi status in vitro. To address whether OsCK2α3-mediated OsPHO2 phosphorylation and degradation in vivo is relative to external Pi conditions, we extracted proteins from PHO2pro-FLAG-PHO2 and α3Ri/ FLAG-PHO2 lines subjected to Pi starvation for different periods of times. The immunoblot results showed that OsPHO2 protein abundance was higher and decreased slower in the α3Ri background than in the wild-type background under Pi-deficient conditions (Supplemental Fig. S10A). Upon Pi-deficiency treatment for 1 d, the phosphorylation of PHO2 increased. The PHO2 protein level decreased quickly and was almost undetectable after 5 d of Pi-deficiency treatment (Supplemental Fig. S10B). Combined with the in vitro experiment results demonstrating phosphorylation of OsPHO2 at Ser-841 impairs its stability, we inferred that Pi deficiency promotes OsPHO2 phosphorylation by OsCK2α3 and further promotes OsPHO2 degradation in wild type compared with that in α3Ri.

Figure 5.

Figure 5.

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Phosphorylated PHO2 is degraded more rapidly than unphosphorylated PHO2. A, Cell-free degradation of HIS-PHO2. Root proteins were extracted with cell-free degradation buffer from wild-type (NIP) plants cultured in HP (200 μm) or LP (10 μm) solution. The protein extracts were incubated with recombinant HIS-MBP, HIS-PHO2 (PHO2), HIS-PHO2S841A (PHO2S841A), and HIS-PHO2S841D (PHO2S841D) for the indicated time periods, and recombinant protein levels were determined by immunoblotting with anti-HIS antibody. HIS-MBP was used as a nondegraded loading control. B, Quantitation of PHO2 in a cell-free degradation assay. The level of PHO2 was normalized to that of the 0-min control (set to 1.0). Error bars represent ± sd (n = 3 independent experiments with eight plants used for each experiment).

The Phosphorylation Status of OsPHO2 at Ser-841 Affects Phosphate Homeostasis

To investigate whether the phosphorylation status of OsPHO2 at Ser-841 affects phosphate homeostasis in rice, we transformed pho2 plants with vectors PHO2pro-gPHO2, PHO2pro-gPHO2S841A, and PHO2pro-gPHO2S841D, containing the genomic sequences of PHO2, PHO2S841A, and PHO2S841D driven by its native promoter, to obtain complementation transgenic lines PHO2/pho2, PHO2S841A/pho2, and PHO2S841D/pho2, respectively. For each type of complementation transgenic line, three representative independent transgenic lines with similar PHO2 transcript levels were identified and used for further study (Supplemental Fig. S11). We investigated the phenotypes and cellular Pi concentrations of the wild-type NIP, pho2, and complementation lines (T2 plants). As shown in Figure 6A, the defective growth performance of pho2 was rescued to almost the level of wild-type NIP plants when transformed with constructs harboring PHO2, PHO2S841A, or PHO2S841D. The leaf necrosis of pho2 was also absent in the complementation lines, except for a few chlorotic areas on the leaf tip margins of the PHO2S841D/pho2 lines, especially in mature leaves (Fig. 6B). Notably, the shoot biomass of PHO2S841A/pho2 plants (∼1.9-g/plant) was higher than that of PHO2/pho2 plants (∼1.6-g/plant), whereas that of PHO2S841D/pho2 plants (∼1.4 g/plant) was lower than that of PHO2/pho2 plants (Fig. 6C).

Figure 6.

Figure 6.

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The phosphorylation status of PHO2 affects Pi homeostasis. A, Phenotypes of wild-type, complementation lines, and pho2 plants grown for 30 d in the presence of 200 μm of Pi. Scale bars = 10 cm. B, Necrosis symptoms on the second leaf tips of the plants shown in (A). Scale bars = 2 cm. C, Dry weights of wild-type, complementation lines, and pho2 plants grown for 30 d in the presence of 200 μm of Pi. D, Cellular Pi concentrations of wild-type, complementation lines, and pho2 plants grown for 30 d in the presence of 200 μm of Pi. Error bars represent + SD (n = 5 independent plants of each line in C, and n = 3 independent plants of each line in D). Different letters above the bars indicate significant differences between groups, as determined by ANOVA followed by the Tukey’s HSD test (P < 0.05). FW, fresh weight.

By contrast, the leaf Pi concentrations were the highest for PHO2S841D/pho2 plants (∼3.7 mg g−1) and the lowest for PHO2S841A/pho2 plants (∼1.5 mg g−1) among the three types of complementation plants (Fig. 6D). Consistently, the leaves of PHO2S841D/pho2 plants cultured in LP solution contained the highest Pi levels among the complementation plants, while the Pi levels of the two other complementation plants were similar to that of wild-type NIP (Supplemental Fig. S12). The root biomass of the complementation plants showed a similar trend to that of the shoots, while the root Pi concentration showed the opposite trend, although the differences among plants were small when grown under both HP and LP conditions (Fig. 6, C and D; Supplemental Fig. S12), suggesting that Pi translocation may differ in different transgenic lines. These results highlight the importance of phosphorylation of OsPHO2 at Ser-841 for Pi homeostasis in rice, which is in line with the impact of phosphorylation on OsPHO2 protein stability.

OsPHO2 Targets OsPHO1 for Degradation

In screening for suppressors of Ospho2, we previously identified the Ospho1 mutant (Hu et al., 2016). This mutant, which harbors a C-to-T (2,374 bp from ATG) mutation, has lower leaf Pi levels and higher root Pi levels than wild-type NIP, while the Ospho1 mutation in the Ospho2 background partially rescued the phenotype of Pi toxicity and growth inhibition in Ospho2 (Supplemental Fig. S13). To determine whether OsPHO2 interacts with OsPHO1 in rice, we performed BiFC and split-ubiquitin Y2H assays. In the BiFC assay, GFP fluorescence was detected when OsPHO1 was cotransformed with a construct expressing OsPHO2C719A (the conserved catalytically active residue Cys-719 of OsPHO2 replaced with Ala), but not with wild-type OsPHO2 (Fig. 7, A and B). The split-ubiquitin Y2H assay showed that all OsPHO2 isoforms (in which the potential phosphorylation site Ser-513 or Ser-841 or the catalytically active residue Cys-719 was replaced with Ala) interacted with OsPHO1 (Supplemental Fig. S14). The different results obtained using OsPHO2 and OsPHO2C719A in the BiFC assay might have been due to the degradation of OsPHO1 by OsPHO2 in planta. Further Y2H analysis using truncated proteins indicated that the N terminus of OsPHO2 interacts with the N terminus of OsPHO1 (PHO1N), which contains the SPX domain (Fig. 7, C and D). These results indicate that OsPHO2 interacts with OsPHO1 through the N termini of both OsPHO2 and OsPHO1.

Figure 7.

Figure 7.

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OsPHO2 targets OsPHO1 for degradation. A, Sequence alignment of peptides from OsPHO2 and AtPHO2. Red rectangle indicates the conserved Cys catalytic, active residue, which is potentially important for ubiquitin-conjugating E2 enzyme activity. B, BiFC analysis of the interaction between PHO2 and PHO1. PHO2 and PHO1 were fused to the N terminus of the N- and C-terminal fragments of YFP (YFPN and YFPC), respectively. Scale bars = 20 μm. C, Schematic diagrams of full-length PHO1 and its deletion derivatives. Numbers refer to the positions of the first or last amino acid in each sequence. D, Interaction between PHO2 and PHO1 in a Y2H assay. AD, pGADKT7 empty vector; BD, pGBKT7 empty vector; SD/LW, SD/-Leu-Trp; SD/LWH, SD/-Leu-Trp-His; 3-AT, 3-amino-1,2,4-triazole. E, Cell-free degradation assay of HIS-PHO1N. Root proteins were extracted with cell-free degradation buffer from wild-type and pho2 plants. HIS-PHO1N recombinant protein was incubated with the extracts with or without 10 μm of E-64D or MG132, and protein abundance at the indicated time point was determined by immunoblotting. F, Cell-free degradation assay of HIS-PHO1N. Root proteins were extracted with cell-free degradation buffer from wild-type, complementation lines, and pho2 plants. HIS-PHO1N recombinant protein was incubated with the extracts, and protein abundance at the indicated time point was determined by immunoblotting.

We performed a cell-free degradation assay to investigate whether OsPHO2 targets OsPHO1 for degradation. We used the recombinant protein HIS-PHO1N in this assay, as we failed to purify the full-length membrane protein HIS-PHO1. We incubated recombinant HIS-PHO1N protein with protein extracts from wild-type NIP or pho2 plants. HIS-PHO1N protein levels decreased rapidly when incubated with wild-type NIP protein extracts compared to pho2 extracts or extraction buffer alone (Fig. 7E). The presence of E-64D, but not MG132, prevented the degradation of HIS-PHO1N (Fig. 7E), indicating that OsPHO2 degrades OsPHO1 via a multivesicular body-mediated vacuolar degradation pathway. We then examined the effects of different OsPHO2 isoforms (S841A or S841D) on the stability of OsPHO1. HIS-PHO1N levels declined most rapidly when incubated with PHO2S841A/pho2 root extracts; most of this protein degraded within 45 min. By contrast, HIS-PHO1N degraded at the slowest rate in PHO2S841D/pho2 extracts, with large amounts still not degraded after 90 min (Fig. 7F). These results are consistent with those shown in Figure 5, indicating that PHO2S841A and PHO2S841D display the highest and lowest protein stability, respectively.

DISCUSSION

Plants have evolved elaborate physiological and biochemical mechanisms to cope with ever-changing soil P conditions. PHO2, a ubiquitin-conjugating E2 enzyme, plays an important role in regulating Pi homeostasis in plants (Delhaize and Randall, 1995; Hu et al., 2011). In this study, we demonstrated that OsPHO2 is subjected to phosphorylation by the OsCK2 subunit OsCK2α3. The resulting phosphorylated OsPHO2 is degraded more quickly than nonphosphorylated OsPHO2, therefore affecting its functions, such as targeting OsPHO1 for degradation and (ultimately) Pi homeostasis. This CK2α3-dependent phosphorylation of PHO2 represents a previously unidentified mechanism used by plants to finely adjust internal phosphate levels.

OsPHO2 Is Regulated at the Post-Transcriptional and Post-Translational Levels

OsPHO2 transcript levels gradually decreased with increasing Pi-deprivation time, even after resupplying the plants with Pi for 1 d (Supplemental Fig. S8A). This finding is consistent with that for AtPHO2 (Aung et al., 2006). Thus, the post-transcriptional regulation of PHO2 is conserved between rice and Arabidopsis. We also identified a post-translational mechanism acting on OsPHO2 that controls its stability. Our study indicated that OsPHO2 is phosphorylated by OsCK2α3 and that phosphorylated OsPHO2 is degraded more rapidly than nonphosphorylated OsPHO2 (Fig. 5). MG132 or E-64D treatment did not inhibit the degradation of OsPHO2 in vitro and in vivo (Supplemental Figs. S8B and S9); therefore, it is unlikely that the degradation of this protein occurs through the ubiquitination-mediated degradation pathway. Whether OsPHO2 is degraded through the lysosome pathway or other pathways remains to be investigated. It has recently been reported that OsPHO2 is regulated by thioredoxins (OsTrxh1 and OsTrxh4; Ying et al., 2017), although whether thioredoxins affect OsPHO2 protein stability or enzyme activity remains to be investigated. Together, these findings highlight the central role of OsPHO2 in Pi homeostasis, and indicate that its activity is under a sophisticated control mechanism acting at different post-transcriptional levels.

The protein kinase CK2 is an evolutionarily conserved Ser/Thr kinase found in all eukaryotes and has been studied for more than 30 years (Montenarh and Götz, 2018). Plant CK2 is involved in various physiological processes, such as light signaling, the circadian clock, phytohormone responses, and so on (Portolés and Mas, 2010; Bu et al., 2011; Vilela et al., 2015). In many organisms, the CK2 holoenzyme is a heterotetramer composed of two types of subunits: two catalytic subunits (CK2α) and two regulatory subunits (CK2β; Litchfield, 2003). However, increasing evidence indicates that both the CK2α subunits and CK2β subunits can also function as monomers and exert independent functions (Filhol et al., 2004). The OsCK2α3/β3 holoenzyme phosphorylates OsPT8 and regulates its PM localization. However, OsCK2α3 cannot interact with OsPT8 in the absence of OsCK2β3 (Chen et al., 2015). In this study, Y2H and BiFC assays showed that OsCK2α3 directly binds to the N terminus (635 amino acids) of OsPHO2 without an existing β-subunit (Fig. 1), while the catalytic subunit OsCK2α2 and the regulatory subunits OsCK2β1 and OsCK2β3 failed to interact with OsPHO2 in yeast (Fig. 1A). Further study is required to determine whether a β-subunit is needed for the phosphorylation of OsPHO2 by OsCK2α3 in vivo.

CK2 is acidophilic in nature, and its phosphorylation site has been defined as -Ser/Thr-Xaa-Xaa-Asp/Glu- (Ser is the most common phosphor acceptor, and Xaa represents any amino acid; Johnson and Wu, 2016). However, we found that OsCK2α3 phosphorylated OsPHO2 at the C-terminal Ser-841 in the -Ser-Asp-Asp-Gly- motif, which does not conform with the previous report. Analysis of the corresponding Ser-841 and the nearby amino acids of OsPHO2 among monocotyledons and dicotyledons revealed that most monocots contain Ser at the corresponding site of OsPHO2, whereas this site is not conserved in dicots (Supplemental Fig. S15). This finding suggests that monocots (especially Graminaceae) and dicots evolved different regulatory mechanisms of PHO2 activity. In vivo analyses demonstrated that OsCK2α3 interacts with OsPHO2 in the ER (Fig. 2, A and C), suggesting that OsPHO2 is phosphorylated in the ER. It is interesting that both PHO2 mRNA and protein levels did not increase after Pi was resupplied for 1 d (Supplemental Fig. S8). These results suggest that PHO2 expression suffers other post-transcriptional regulation than low Pi-responsive miR399 cleavage.

OsCK2 Together with OsPHO2 Fine-tunes Pi Homeostasis

In this study, we found that OsCK2α3 phosphorylates OsPHO2, making OsPHO2 easier to be degraded. The nonphosphorylatable version of OsPHO2 (PHO2S841A) in plants was more stable than the wild-type protein, and PHO2S841A/pho2 transgenic plants accumulated reduced levels of Pi in leaves. On the contrary, the phosphorylation-mimic version of OsPHO2 (PHO2S841D) in plants was more fragile and PHO2S841D/pho2 transgenic plants accumulated more Pi in leaves compared to the controls (Fig. 6). Although the differences of Pi concentrations among wild-type NIP, pho2, and the complementation lines roots were slight, the Pi concentrations of PHO2S841D/pho2 roots were the lowest compared to PHO2/pho2 and PHO2S841A/pho2 plants (Fig. 6D; Supplemental Fig. S12). These results demonstrate that OsCK2α3 at least partially exerts its function on Pi homeostasis through OsPHO2. Unfortunately, the complementation lines did not fully recover the phenotype of pho2 to that of the wild type. In a previous study, complementation with a genomic sequence 322-bp longer than that used in this study also failed to completely restore the pho2 phenotype (Ying et al., 2017; Supplemental Fig. S16). A recent study reported that OsPHO2 might produce two or more alternative transcripts with longer 5ʹ untranslated regions or shorter coding sequences (CDSs) than the standard transcript in both shoot and root (Dong et al., 2018). Whether the incomplete complementation of Ospho2 observed in this study was due to the insufficient length of the genomic OsPHO2 sequence or the involvement of other mechanisms requires further study.

In this article, we demonstrated that OsCK2α3 has a positive effect on Pi levels by negatively regulating the stability of OsPHO2. A previous study indicated that OsCK2α3/β3 phosphorylates OsPTs and inhibits the exit of PTs from the ER to avoid excessive Pi uptake under Pi-replete conditions (Chen et al., 2015). The effect of OsCK2α3 on OsPTs requires the regulatory subunit OsCK2β3, whereas its effect on OsPHO2 does not appear to require the participation of the regulatory subunit. The differential regulation of the β- and the α-subunit (e.g. the β-subunit is much more unstable under Pi starvation than the α-subunit) provides an additional means for partially uncoupling the activity of OsCK2α3 on OsPTs and OsPHO2. We speculate that in high Pi conditions, CK2α3 suppresses excessive Pi absorption from soil through inhibiting the PM localization of PTs and promotes Pi translocation from root to shoot for storage in vacuoles through the OsPHO2-OsPHO1 pathway; while in low Pi conditions, OsCK2α3 is released from interacting with PTs as OsCK2β3 is degraded, the PM localization of PTs are promoted for Pi absorption, and more OsCK2α3 is free to phosphorylate OsPHO2, which further promotes Pi translocation from root to shoot for the Pi requirement of shoot growth. The dual roles of OsCK2α3 on Pi absorption and root-to-shoot translocation may reflect an elaborated regulatory mechanism for plants to adapt to environmental Pi fluctuations. In α3Ri lines, the OsPHO2 protein is more stable to degrade OsPHO1 and inhibit Pi translocation, although more PTs are unphosphorylated and transported to PM for Pi acquisition and translocation. A similar phenomenon exists in Arabidopsis WRKY42, which promotes the expression of AtPHT1;1 and suppresses the expression AtPHO1, thereby promoting Pi uptake and suppressing root-to-shoot translocation, respectively (Su et al., 2015).

The Pi excess phenotype displayed by pho2 can be attributed to the misregulation of Pi homeostasis at three different levels: (1) enhanced Pi uptake, (2) increased root-to-shoot translocation of Pi, and (3) retention of Pi in older leaves (Aung et al., 2006). Y2H and BiFC assays indicated that OsPHO2 binds to the N terminus of OsPHO1 (Fig. 7, B–D). Moreover, E-64D, an endosomal protease inhibitor, inhibited OsPHO2-mediated OsPHO1 degradation (Fig. 7E). These findings indicate that the PHO2-PHO1 regulatory pathway is conserved between rice and Arabidopsis. However, a mutation of OsPHO1 in the Ospho2 background only partially recovered the defective phenotype of this mutant (Supplemental Fig. S13A), pointing to the existence of other potential PHO2 downstream target(s). In Arabidopsis, PHO2, along with NITROGEN LIMITATION ADAPTATION, also targets PT2 for degradation (Park et al., 2014). However, in rice, OsPHO2 does not interact with OsPT2, OsPT6, or OsPT8, but it interacts with PHF1 (Ying et al., 2017). Therefore, perhaps OsPHO2 also targets OsPHF1 for degradation in rice.

Combining this data with previous findings, we propose a model for how OsCK2, together with OsPHO2, regulates Pi homeostasis in rice (Fig. 8). Under Pi-replete conditions, the phosphorylation of OsPHO2 by OsCK2α3 promotes its degradation to maintain OsPHO2 and its target, OsPHO1 (and perhaps OsPHF1), at appropriate levels to ensure Pi uptake and translocation from root to shoot to promote shoot growth. On the other hand, OsCK2α3, together with OsCK2β3, phosphorylates OsPTs and inhibits their exit from the ER (trafficking to the PM) to avoid excess Pi uptake and to maintain cellular Pi homeostasis. Under Pi deprivation, OsCK2β3 is degraded and OsCK2α3 stops phosphorylating PTs, allowing them to traffic to the PM to promote Pi uptake. In the meantime, more OsCK2α3 is free to phosphorylate OsPHO2 to promote its degradation. Therefore, the OsPHO2 target, OsPHO1 (and perhaps OsPHF1), is upregulated to promote Pi uptake and translocation from root to shoot. In summary, we uncovered a new regulatory mechanism underlying Pi homeostasis involving the phosphorylation of OsPHO2 by OsCK2α3. This finding should facilitate the dissection of the Pi regulatory network to improve the efficiency of Pi uptake and Pi use in crops via breeding.

Figure 8.

Figure 8.

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Working model of the role of the OsCK2α3-OsPHO2 module in monitoring rice Pi homeostasis. Phosphorylation of OsPHO2 protein by OsCK2α3 promotes (bold arrow) its degradation (dashed oval) to maintain OsPHO2, along with its target, OsPHO1 (and perhaps OsPHF1, as indicated by a dashed line), at an appropriate level to maintain Pi uptake and translocation from root to shoot. Meanwhile, OsCK2α3, together with OsCK2β3, phosphorylates OsPTs and inhibits their exit from the ER (trafficking to the PM) to avoid excessive Pi uptake, thereby maintaining cellular Pi homeostasis.

MATERIALS AND METHODS

Plant Materials and Growth Conditions

Wild-type rice (Oryza sativa ‘Nipponbare’ [NIP]) and transgenic plants were grown hydroponically in a greenhouse as described in Chen et al. (2015). For +P and −P conditions, the concentration of KH2PO4 was adjusted to 200 and 0 μm, respectively, unless specified otherwise (HP, 200 μm; LP, 10 μm). Nicotiana benthamiana plants were cultivated in growth chambers as described in Lv et al. (2014). The pho2 mutant (Tos17 insertion line NE8536) was purchased from the Rice Tos17 Insertion Mutant Database (https://tos.nias.affrc.go.jp). The primers used are listed in Supplemental Table S1. The CK2α3 RNA interference plants (designated as α3Ri) were described in Chen et al. (2015). The PHO2pro-FLAG-PHO2 plants (designated as FLAG-PHO2) were described by Ying et al. (2017). The pho1 plants were described by Hu et al. (2016).

Construction of Vectors and Generation of Transgenic Plants

For the complementation lines, a 9,478-bp DNA fragment containing the promoter sequence (4,777 bp upstream sequence from ATG), the genomic sequence, and the 569-bp 3ʹ untranslated region of PHO2 were amplified from wild-type NIP genomic DNA and cloned into the pCAMBIA2300 vector, designated as PHO2pro-gPHO2. The mutated PHO2 genes were generated using the multifragment fusion method according to the manufacturer’s instructions (C113-02; Vazyme Biotech); the vectors containing the mutated PHO2 genes were named PHO2pro-gPHO2S841A and PHO2pro-gPHO2S841D.

All primers used are listed in Supplemental Table S1. The constructs PHO2pro-gPHO2, PHO2pro-gPHO2S841A, and PHO2pro-gPHO2S841D were transformed into calli induced from mature rice pho2 embryos to produce the complementation lines PHO2/pho2, PHO2S841A/pho2, and PHO2S841D/pho2, respectively. All transgenic plants were produced via Agrobacterium tumefaciens (strain EHA105)-mediated transformation, as described in Wang et al. (2014).

Y2H Assays

To identify OsPHO2 interaction partners, standard Y2H assay was performed using the N-terminal of OsPHO2 as bait to screen a rice complementary DNA (cDNA) library using the Matchmaker GAL4 Two-hybrid System (Clontech). Split-ubiquitin Y2H assays were performed following the instructions provided with the DUALmembrane Pairwise Interaction kit (Dualsystems Biotech). For the split-ubiquitin Y2H assays, the full-length OsPHO2 CDS without the stop codon (2,628 bp) was cloned into the BD domain of the pDHB1 vector, and the OsPHO1 CDS (2,448 bp) was amplified and fused in frame to the 3ʹ terminus of the pPR3-N vector. For the standard Y2H assays, the full-length and truncated OsPHO2 (PHO2N, PHO2M, and PHO2C) CDSs were cloned into the pGBKT7 vector, and the truncated OsPHO1 (PHO1N and PHO1C) CDS was cloned into the pGADT7 vector. All primers used are listed in Supplemental Table S1. All prey vectors of OsCK2 used in the assays were described in Chen et al. (2015). SD/-Leu-Trp-His supplemented with 10 mm of 3-amino-1,2,4-triazole and SD/-Leu-Trp-His-Ade were used for selection.

Pull-Down Assays

MBP sequence was cloned into pET-28a (+; Novagen) with NdeI and EcoRI to obtain MBP-tag vector, and then full-length PHO2 was cloned in-frame into the 3ʹ terminus of the MBP-tag with EcoRI, named HIS-PHO2; the primers used are listed in Supplemental Table S1. The fusion HIS-PHO2 construct was transformed into Escherichia coli BL21 cells (CD901-01; TransGen Biotech), and the transformed cells were treated with 0.5 mm of isopropyl-b-d-thiogalactoside for 16 h at 20°C to induce the expression of the fusion proteins. Recombinant protein produced in E. coli was purified using a Ni-NTA column according to the manufacturer’s instructions (Qiagen). The recombinant proteins GST, GST-α3, and GST-β3 were purified as described in Chen et al. (2015). The purified fusion proteins GST-α3 and HIS-PHO2 or GST and HIS-PHO2 were incubated with Glutathione Sepharose 4 Fast Flow (17-5132-01; GE Healthcare) in 1× PBS buffer with 1 mm of phenylmethylsulfonyl fluoride for 3 h. After washing five times with 1× PBS buffer, the beads were suspended in 60 μL of elution buffer. After a brief centrifugation, the supernatant was collected and subjected to immunoblot analysis as described in Lv et al. (2014). The mouse anti-His (Abcam) and mouse anti-GST antibodies (HT601-02; TransGen Biotech) were diluted 1:5,000. The secondary antibody, rabbit anti-mouse IgG peroxidase antibody (AP160P; Sigma-Aldrich) was diluted 1:10,000.

RNA Isolation and RT qPCR Analysis

Total RNA was isolated from rice samples using TRIzol reagent (Invitrogen), followed by treatment with DNase I (Qiagen) before RT-qPCR to eliminate genomic DNA contamination. cDNA was synthesized from 1 µg of total RNA using the Reverse Transcription System (A3500; Promega). RT-qPCR was performed using SYBR Green I Master (Roche) on the LightCycler 480 Real-Time PCR system (Roche) according to the manufacturer’s instructions. Relative expression levels were normalized to that of the housekeeping gene OsACTIN1. The primers used for RT-qPCR are listed in Supplemental Table S1.

In Situ PCR

In situ PCR was performed as described in Athman et al. (2014). Briefly, the roots were sliced into 50-μm–thick sections using a Vibratome (Leica). Then the samples were transferred into 87.5 μL of sterile water with 100 U of RNaseOUT, to which was added 10 μL of 10× Turbo DNA-free buffer (Ambion) and 2.5 μL 1 U μL−1 DNase I (Qiagen) to give a final volume of 100 μL in a 0.2-mL PCR tube. After incubation at 37°C for 45 min to eliminate the genomic DNA, the reaction was stopped by adding 3.3 μL of 0.5 m EDTA at pH 8.0 and heating to 75°C for 10 min. Then reverse transcription and PCR were conducted using an in situ PCR system, and the cDNAs were amplified with gene-specific primers (Supplemental Table S1). After the PCR amplification, the samples were washed twice for 5 min with PBS buffer, blocked for 30 min in 0.1% (w/v) BSA, and incubated for 1 h with 1.5 U of Anti-DIG-AP antibody (Roche). The samples were then washed twice for 15 min with washing buffer (0.1 m of Tris-HCl and 0.15 m of NaCl [ pH 9.5]), and stained with BM Purple AP Substrate (Roche) for 40 min, after which they were washed twice with water and imaged using an ECLIPSE 90i microscope (Nikon).

Subcellular Localization Analysis and BiFC Assays

To produce the 35S-CK2α3-mCherry and 35S-GFP-PHO2 fusion constructs, the CDSs of CK2α3 (996 bp) without the stop codon was amplified from NIP cDNA with KOD-Plus (Toyobo) and fused in-frame to the 5ʹ terminus of mCherry in the modified pCAMBIA1300-35S-mCherry plasmid. The PHO2 CDS (2631 bp) was amplified and fused in-frame to the 3ʹ terminus of eGFP in the modified pCAMBIA1300-35S-eGFP plasmid. For the BiFC assays, full-length CK2α3, PHO2, and PHO1 were cloned into pCB301 vectors and fused in-frame with either the C-terminal or N-terminal fragment of Yellow Fluorescent Protein (YFP) as described in Lv et al. (2014). Primers used are listed in Supplemental Table S1. The resulting constructs, 35S-GFP-PHO2 with 35S-CK2α3-mCherry and 35S-AtWAK2-mCherry, were transiently transformed into N. benthamiana leaves by Agrobacterium-mediated infiltration as described in Walter et al. (2004). Observations were made with a laser-scanning microscope (Axiovert LSM 710; Zeiss). A 25× water immersion objective was used for confocal imaging. For excitation of fluorescent proteins, the following lines of an argon ion laser were used: 488 nm for GFP and YFP and 543 nm for mCherry. Fluorescence was detected at 493 to 542 nm for GFP, 500 to 542 nm for YFP, and 578 to 625 nm for mCherry.

Cell-free Degradation Assay

The PHO1N sequence (1–1,254 bp) was cloned into pET28a (+; Novagen), and HIS-PHO1N was purified as described above for HIS-PHO2. Ten-d–old seedling roots of wild-type NIP grown in standard nutrient solution followed by 7 d of growth under +P/−P conditions were harvested and ground into a fine powder in liquid nitrogen. Total proteins were extracted from the samples and prepared as described in Lv et al. (2014). Recombinant GST, GST-α3, GST-β3, HIS-MBP, HIS-PHO2, HIS-PHO2S841A, or HIS-PHO1N (100 ng) was incubated with 40 μL of protein extracts containing 100 μg of protein per reaction. The reactions were incubated at 28°C, and samples were collected at the indicated intervals and subjected to immunoblot analysis. All experiments were repeated three times, and representative results are shown.

In Vitro Phosphorylation Assay

Approximately 4 µg of kinase (GST or GST-α3) and 1 μg of substrate (HIS-PHO2) proteins were combined with 1× kinase buffer (100 mm Tris-HCl [pH 8.0], 5 mm DTT, 5 mm EGTA, 5 mm MgCl2, and 100 μm ATP, with 0.5 of μCi γ-32P-ATP for the autoradiography assay) in a total volume of 30 μL. The reactions were incubated at 30°C for 30 min and stopped by adding 5× loading buffer and boiling for 5 min. An aliquot of each assay was resolved by SDS-PAGE, and the gels were stained and dried. Phosphorylation was detected by autoradiography using a Typhoon-8600 scanner (Molecular Dynamics). For LC-MS/MS analysis, the corresponding bands in the gel were sliced, digested with trypsin, and analyzed using an LC-MS/MS system. For λ-PPase (0753S; New England Biolabs) treatment and the Phos-Tag SDS-PAGE (Phos-tag AAL-107; Wako) assay, the reactions were separated into two duplicates after a 30-min incubation: one was combined with 50 U of λ-PPase and the other was combined with DMSO as a control. Other components were added according to the manufacturer’s instructions (P0753S; New England Biolabs). The reactions were incubated at 30°C for 30 min and separated on 7.5% Phos-Tag acrylamide gels.

Immunoblotting

To extract total proteins, the rice root samples were ground in liquid nitrogen and resuspended in IP lysis buffer (50 mm Tris-HCl [pH 7.4], 150 mm NaCl, 1 mm EDTA, 1% [v/v] Triton X-100, and 10% [v/v] glycerol) with freshly added 1 mm of phenylmethylsulfonyl fluoride and 1× protease inhibitor cocktail (Roche). Protein concentrations were determined by a protein assay (Bio-Rad). For the in vivo phosphorylation assay, the protein extracts were treated with 100 U of λ-PPase in a total volume of 30 μL and incubated at 30°C for 30 min. For the control, an equal volume of DMSO (the solvent used to prepare the inhibitor stock solutions) was added to the sample instead. The reactions were stopped by adding 5× SDS loading buffer and boiling. Each sample (20 μg of total protein) was loaded onto a 10% SDS-PAGE gel or 7.5% Phos-Tag acrylamide gel. The mouse anti-FLAG antibody (F1804; Sigma-Aldrich) was diluted 1:3,000. The secondary antibody, rabbit anti-mouse IgG peroxidase antibody (AP160P; Sigma-Aldrich), was diluted 1:10,000.

Measurement of Cellular Pi Concentration

Cellular Pi concentration was measured as described in Chen et al. (2011).

Statistical Analyses

Statistical analysis was conducted using the program SPSS Statistics v22.0 (IBM). Data were analyzed by ANOVA followed by the Tukey’s Honest Significant Difference test (P < 0.05 as the level of significance).

Accession Numbers

Sequence data from this article can be found in the Michigan State University Rice Genome Annotation Project Database (http://rice.plantbiology.msu.edu/index.shtml) under the following accession numbers: LOC_Os07g02350 (CK2α2), LOC_Os03g10940 (CK2α3), LOC_Os10g41520 (CK2β1), LOC_Os07g31280 (CK2β3), LOC_Os03g50885 (ACTIN), LOC_Os05g48390 (PHO2), and LOC_Os02g56510 (PHO1).

Supplemental Data

The following supplemental materials are available.

Acknowledgments

We thank Javier Paz-Ares and Laurent Nussaume for critical reading and comments.

Footnotes

1

This work was supported by the National Key Research and Development Program of China (grant no. 2016YFD0100700), the National Natural Science Foundation of China (grant nos. 31972486 and 31572187), the Ministry of Agriculture of China (grant no. 2016ZX08001003–009), the Natural Science Foundation of Zhejiang Province, China (grant no. LZ17C020001), and the Ministry of Education and Bureau of Foreign Experts of China (grant no. B14027).

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