To examine whether the ferroxidases LPR1 and LPR2 are involved in the regulation of Fe nutrition, two T-DNA insertion lines of LPR1 and LPR2 (lpr1-1 and lpr2-1) and the double mutant lpr1lpr2 (generated by crossing lpr1-1 and lpr2-1) were analyzed. When 3-week-old seedlings were transferred to the nutrient solution with sufficient Fe [50 μM Fe(III)–EDTA] supply for 7 days, we found that the single mutants lpr1-1 and lpr2-1 did not differ from the wild type; however, the lpr1lpr2 double mutant exhibited strong stunted and higher chlorophyll levels, with less fresh biomass in shoots and roots (Figures 1A–C and Supplementary Figure 2). We also generated pLPR1:LPR1-YFP and pLPR2:LPR2-YFP transformants in the background of lpr1lpr2 double mutant for genetic complementation test. These complementation lines had a higher LPR1 and LPR2 expression than wild type did and were able to restore the defects of lpr1lpr2 double mutant in phenotypes (Figure 1A and Supplementary Figure 1). The above results indicate that LPR1 and LPR2 redundantly involve in maintaining normal growth under our growth conditions.

As both LPR1 and LPR2 have been identified as ferroxidases, we determined the concentrations of Fe and other elements in plants. Surprisingly, the Fe concentrations in the shoots and roots were both greatly higher in the lpr1lpr2 double mutant than in the wild type and single mutants. The Fe concentrations in the shoots and roots of the lpr1lpr2 were 1.8 and 1.5 times that of the wild type, respectively (Figures 1D,E). We also checked the levels of other nutrients (Supplementary Figure 3); although Zn, Mn, and Cu concentrations also increased significantly, they were not as high as Fe, suggesting that LPRs may be mainly involved in Fe nutrition. Accordingly, the seedlings were subsequently grown with two lower doses of Fe (1 μM and 0.2 μM Fe) to investigate whether the stunted growth of lpr1lpr2 double mutant is associated with the over-accumulation of Fe. As expected, both lower-Fe-dose treatments improved the growth of lpr1lpr2 (Figures 1B–E and Supplementary Figure 4). Particularly, the lpr1lpr2 displayed a similar growth and ionomic phenotypes to that of the wild type under 0.2 μM Fe condition. These results infer that LPR1 and LPR2 are functionally redundant and play an important role in maintaining Fe homeostasis for normal growth.

Given that soil pH has a great influence on the availability of Fe (Sarkar and Wynjones, 1982), we also compared the growth of wild type and lpr1lpr2 in the soils with different pH values. In the moderately acidified soil (pH 5.3), the shoots of lpr1lpr2 had significantly reduced biomass, but the Fe concentration of lpr1lpr2 is 4.72 times that of wild type. These differences were greatly minimized when the plants were grown in the soil with pH 6.2 (Supplementary Figure 5). These results provide a further support for that LPR1 and LPR2 are required for maintaining Fe homeostasis in plants.

To identify in which tissues LPR1 and LPR2 involve in maintaining Fe homeostasis, we used β-glucuronidase (GUS) as a reporter gene to investigate the expression patterns of LPR1 and LPR2. Histochemical staining of the plants expressing pLPR1:GUS and pLPR2:GUS revealed that both LPR1 and LPR2 were mainly expressed in the vascular tissues of roots, hypocotyls, leaves, lateral roots, stems, sepals, as well as repla to a lesser extent (Figures 2A,B and Supplementary Figure 6). To analyze the vascular tissues in more detail, we examined the expression patterns in the cross sections of roots, hypocotyls, and leaf veins. As shown in the insets of Figures 2A,B, LPR1 and LPR2 were mainly expressed in the parenchyma cells surrounding the xylem and phloem, indicating that LPRs may function in the vascular tissues.

The above expression patterns of LPRs in vascular tissues promoted us to test LPRs’ roles in Fe long-distance transport in A. thaliana. Under our growth condition with sufficient Fe [50 μM Fe(III)–EDTA], the total Fe concentration in the xylem and phloem saps was much higher in lpr1lpr2 than in wild type (Figures 2C,G), which was consistent with the Fe concentration of shoots (Figures 1D,E). Given that LPRs could oxidize Fe(II) to Fe(III) in vitro (Muller et al., 2015; Liu et al., 2022), we subsequently investigated the Fe valence states in the xylem sap based on the ferrozine method (Verschoor and Molot, 2013). Both Fe(II) and Fe(III) concentrations in the xylem saps were markedly increased in lpr1lpr2 compared with wild type (Figures 2D,E), and the proportion of Fe(II) in the total Fe of xylem sap of lpr1lpr2 was 2.2-fold the wild type (Figure 2F). These findings suggest that LPRs affect the transport of Fe in the vascular tissues via their Fe(II) oxidation activity.

Considering LPR1 and LPR2 were expressed in both roots and shoots (Figure 2A), we decided to explore the relative contribution of LPRs in roots versus shoots. We first quantified the LPR1 and LPR2 transcripts of wild type. Quantitative real-time PCR (RT-qPCR) assay showed that the expression of LPR1 was overall lower than that of LPR2, and no significant difference was observed between roots and shoots. Nonetheless, a higher LPR2 expression was identified in shoots (Figure 3A), which prompted us to verify whether LPRs were involved in a shoot-based mechanism; hence, a reciprocal graft experiment under normal Fe growth conditions was employed. As the results shown, grafting wild type shoot scions onto lpr1lpr2 double mutant rootstocks (i.e., Col-0/lpr1lpr2-grafted plants) had normal growth in both shoots and roots (Figures 3B–D). However, the grafted plants with lpr1lpr2 scions (i.e., lpr1lpr2/Col-0- and lpr1lpr2/lpr1lpr2-grafted plants) grew smaller than the plants with that of wild type. The results indicate that LPRs in shoots rather than in roots are required for ensuring the normal growth phenotype of shoots.

We further measured the Fe level in the grafted plants. The Fe concentration in shoot scions of lpr1lpr2 double mutant, as expected, was significantly higher than that in wild type (Figure 3E). However, unlike the above growth phenotypes, the Fe concentration of rootstocks was consistent with their genotypes, which was in a shoot-independent manner (Figure 3F). As the translocation factor (TF) can be used to indicate the ability of substance transferred from roots to shoots (Takarina and Pin, 2017), we then calculated the TFs in the grafted plants. An increased TF was found in lpr1lpr2/Col-0-grafted plants (Figure 3G), while Col-0/lpr1lpr2-grafted plant showed a lower TF than that in other grafted plants. The above results suggest that LPRs in shoots and roots involve in Fe sequestration by their corresponding tissues.

The action of LPRs in maintaining Fe homeostasis under normal Fe growth conditions raises a question that how LPRs act under Fe-deficient conditions. When seedlings were grown in Fe-free nutrient solution, there was no noticeable difference in chlorosis between wild type and lpr1lpr2 double mutant (Supplementary Figure 7A). Leaf chlorosis was absent in wild type after 0.2 μM Fe resupply, while lpr1lpr2 still exhibited obvious yellowish color in the newly expanded leaves (Figure 4A). This appearance was also observed when plants were grown in a soil with alkalization treatment which lowered Fe availability (Supplementary Figure 7B). By comparing the alignment of leaves, we were surprised to find that the color of old leaves of lpr1lpr2 double mutant was greener than that of wild type (Figure 4B).

The above phenotype was consistent with the results of Fe concentration measured in different ages of leaves: The lpr1lpr2 had a lower Fe level in young leaf but a higher Fe level in old leaf compared with wild type plants (Figure 4C). We also measured the Fe concentration in different ages of leaves from the plants grown in Fe-free pre-grown plants resupplied with sufficient Fe (50 μM Fe). Under this Fe condition, although the Fe concentration in all leaves of lpr1lpr2 was higher than wild type, the difference was much greater in old leaves (Figure 4D). The calculation of relative Fe concentration (lpr1lpr2 vs. wild type) indicated that Fe accumulated preferentially in older leaves of the lpr1lpr2 double mutant compared with wild type, which was independent of Fe supplementation dose (Figure 4E).

The response of LPRs in the wild type to different Fe supplementation doses was then examined. Regardless of the Fe dose, the expression of LPRs in old leaves was higher than that in young leaves. It is also worth noting that Fe deficiency upregulated the expression of LPR1 and LPR2, especially in older leaves (Figures 4F–H). The calculation of relative expression of LRP1 and LPR2 (0.2 μM Fe vs. 50 μM Fe) showed that Fe deficiency preferentially stimulated these two genes in older leaves. Histochemical GUS staining further confirmed that the expression of LPRs in older vascular tissues was preferentially induced under low-Fe conditions (Supplementary Figures 8A,B). Taken together, these data indicate that LPRs may function mainly in older leaves to prevent Fe over-accumulation, which favors the Fe allocation to younger leaves under Fe-deficient conditions.

The above findings led us to investigate the mechanism by which LPRs affect Fe homeostasis in plants. The Perls/DAB (diaminobenzidine) staining was first used to decipher the localization of Fe under normal growth conditions (50 μM Fe). Perls/DAB displayed heavy Fe accumulation in the root stele of lpr1lpr2 double mutant, whereas such a distribution was not observed in wild type (Figure 5A). In shoots, the vast majority of Fe was located within the vascular bundles of lpr1lpr2 in old leaves (Figure 5B), which was coincided with the LPRs expression. We also compared Fe staining in young and middle leaves, and a slight enhancement of staining color was found in lpr1lpr2 double mutant versus wild type.

To have a close-up view of Fe accumulation in the vascular bundles, the cross section of old leaves was examined. As shown in Figure 5C, we were surprised to find that Fe was mainly deposited at the periphery of xylem vessel, which is a cell wall matrix and thus coincides with the cell wall localization feature of either LPR1 or LPR2 (Muller et al., 2015; Liu et al., 2022). However, the cells adjacent to the xylem vessel of lpr1lpr2 also had deeper Fe staining than that of wild type. These data suggest more Fe unloading from the xylem vessel due to LPRs disruption, possibly because the available Fe in the xylem sap of lpr1lpr2 is higher (Figure 2B). The above notion was also supported by a higher Fe concentration in the phloem of lpr1lpr2 under sufficient Fe conditions (Figure 2F).

We also performed Perls/DAB staining under 0.2 μM Fe conditions. The Fe deposition in old leaves of lpr1lpr2 could still be observed in this low-Fe treatment (Supplementary Figure 9A). The greater Fe deposition in the cell wall matrix of xylem vessel could be expected to decrease Fe unloading from the xylem vessel under low-Fe condition. This assumption was confirmed by the observation that the Fe concentration in the phloem of lpr1lpr2 was less than that of wild type under low-Fe conditions (Supplementary Figure 10).

Given the Fe(II) oxidation by LPRs, we further tested the distribution characteristics of Fe(II) by Turnbull staining. As expected, Fe(II) was also found to be mainly located in the vascular tissues, particularly in the xylem vessels (Figures 5D–F and Supplementary Figure 9B). This was similar to the Perls/DAB staining above. Then, we asked why the disruption of LPRs results in abnormal Fe sequestration in the cell wall of xylem vessel? We therefore extracted the cell wall of A. thaliana to simulate the migration of Fe–citrate in this matrix. The adsorption kinetics analysis reveals that the cell wall had a stronger adsorption capacity for Fe(II)–citrate rather than that for Fe(III)–citrate (Figure 6), providing a reasonable explanation for the above question.

As Fe concentration in the xylem sap was increased due to the disruption of LPRs, we further investigated the gene expression related to Fe homeostasis by RT-qPCR assay. The results showed that under Fe-sufficient conditions, Fe uptake genes IRT1 and FRO2, as well as iron-loading into xylem genes FRD3 and FPN1, were induced to be significantly higher in the roots of lpr1lpr2 double mutant, but displayed a lower expression in the wild type (Figure 7). Under low-Fe conditions, the above genes were upregulated in wild type compared with that observed under normal Fe growth conditions, but were still expressed at higher levels in lpr1lpr2 except FRD3. These data exhibited the constitutive expression of Fe deficiency responses in lpr1lpr2 double mutant, independent of the Fe dose, which is consistent with the higher Fe concentration in lpr1lpr2 roots and shoots under normal Fe growth conditions (Figures 1D,E). The above results infer that LPRs are also required to maintain the appropriate expression of the genes related to Fe homeostasis in plants, which may be indirectly associated with the role of LPRs in preventing abnormal Fe sequestration in the vascular tissues.

All plant lines used in this study were in A. thaliana ecotype Col-0 background. The mutant seeds of lpr1-1, lpr2-1, and lpr1-1lpr2-1 were previously described (Dong et al., 2017; Liu et al., 2022). The pLPR1:GUS/Col-0 transgenic line was a kind gift from Jian-Li Yang (Xu et al., 2020). The seeds were surface-sterilized with 25% NaClO and vernalized at 4°C for 3 days. Then, the seeds were germinated on a nylon mesh and then transferred to the nutrient solution as previously described (Fang et al., 2016). The seedlings were grown under the controlled environment with a 12-h light/12-h dark photoperiod and 100 μmol m^–2s^–1 light intensity at 22°C. Considering the lpr1lpr2 double mutant had developmental defects in Fe-sufficient media, the seedlings were precultured in the nutrient solution with 0.2 μM Fe(III)–EDTA. The other nutrients in the solution were KNO₃ (1.5 mM), CaCl₂ (1 mM), NaH₂PO₄ (0.5 mM), MgSO₄ (0.25 mM), (NH₄)₂SO₄ (0.25 mM), H₃BO₃ (10 μM), MnSO₄ (0.5 μM), ZnSO₄ (0.5 μM), CuSO₄ (0.1 μM), and (NH₄)₆Mo₇O₂₄ (0.1 μM) at pH 5.8. The nutrient solution was refreshed every 4 days. After 21-day preculture, the seedlings were transferred to different doses of Fe(III)–EDTA for further experiments.

For pot experiment, seedlings were germinated on nylon net and then transferred to the peat soil with pH 5.3 (moderately acidified soil) or 6.2 (normal soil) or 7.3 (alkaline soil). The pH of peat soils was adjusted by adding different amounts of CaO. The seedlings were watered weekly with iron-free nutrient solution and harvested after 4 weeks.

To construct genetic complementation lines, a 5.9-kb genomic DNA fragment of LPR1 and a 4.3-kb genomic DNA fragment of LPR2 (each fragment includes 2.9- and 2.2-kb promoters, respectively) were designed and cloned into the pEarleyGate 101 vector containing a YFP reporter as previously described (Liu et al., 2022). The constructs were transformed into the Agrobacterium tumefaciens strain GV3101 and then introduced into lpr1lpr2 double mutant by the floral dip method. For the construction of the pLPR2:GUS/Col-0 transgenic line, a 2.8-kb promoter of LPR2 was cloned into the pCAMBIA 1,301 vector. The resulting construct was transformed into Col-0. All primers used in this study are listed in Supplementary Table 1. Homozygous T3 transgenic plants were used in subsequent studies.

Total RNA in various parts of plant was extracted using RNAiso Plus regent (code no. 9109, TaKaRa, China) following the instruction (MacRae, 2007), and the RNA was quantified and assessed for degradation using a micro-spectrophotometer (Nano-400A, Allsheng, China). For real-time PCR (RT-PCR), 800 ng RNA was used to synthesize the first-strand cDNA by HiScript II Q Select RT SuperMix for qPCR (+gDNA wiper) (code no. R223-01, Vazyme, China). For RT-qPCR, the reaction mixture contained 1 μl cDNA template, 0.5 μl forward primer, 0.5 μl reverse primer, 8 μl ddH₂O, and 10 μl Taq Pro Universal SYBR qPCR Master Mix (code no. Q712-02, Vazyme, China). We performed RT-qPCR analysis on a StepOnePlus Real-Time PCR System (code no. 4376600, Thermo Fisher Scientific, Carlsbad, CA, United States). The specificity of primers was confirmed by melting curve analysis. EF1α and UBQ10 were used as the internal control. The primers used for RT-qPCR are listed in Supplementary Table 1.

To analyze GUS activity, the plant materials were stained as described in the study of Ge et al. (2022). In brief, pLPR1:GUS/Col-0 and pLPR2:GUS/Col-0 plants were incubated in staining buffer (50 mM PBS at pH 7.0, 0.1% Triton X-100, 2 mM K₃Fe[CN]₆, 2 mM K₄[Fe(CN)₆]⋅3H₂O, 10 mM Na₂EDTA⋅2H₂O, 2 mM X-Gluc) at 37°C overnight. Then, the plants were incubated in 70% ethanol at 37°C for decolorization and optically cleared with chloral hydrate. The stained samples were observed using a microscope (Nikon, Tokyo, Japan).

Grafting experiment was performed as previously described with some modifications (Marsch-Martinez et al., 2013; Ye et al., 2021). The seeds were surface-sterilized and sown on 1/2 MS medium (Coolaber) containing 1% agar and 0.5% sucrose at pH 5.8. After 6 days of germination, seedlings were transversely cut in the middle of the hypocotyl. Afterward, shoot scions and rootstocks were joined on the new plate with the same medium. Ten days later, the adventitious roots were removed, and plants were transferred to hydroponic culture with low-Fe (0.2 μM Fe) nutrient solution for another 7 days of preculture. Finally, the grafted plants were harvested 7 days after transferring to Fe-sufficient (50 μM Fe) nutrient solution.

To collect the xylem sap, the plants were grown in hydroponic culture with low-Fe nutrient solution for 3 weeks and transferred to Fe-sufficient nutrient solution for another 2 weeks. After rinsing the hypocotyl with ultrapure water, the hypocotyl was transversely cut, and four plants were used as a sample. The xylem sap was collected within 0.5 h with some modifications (Schuler et al., 2012; Chao et al., 2021). The first drop of the xylem sap was discarded, and the subsequent sap was collected in a 1.5-ml tube with 100 μl 1 M sodium acetate (pH 5.0). After collection, the total volume of each sample was recorded, and they were then divided into two equal parts for subsequent determination of ferrous iron and total iron as described (Verschoor and Molot, 2013).

The plants were grown in hydroponic culture for 5 weeks. After rinsing with ultrapure water, eight leaves of each two plants were detached at the petiole. Phloem exudates were collected within 1 h by using EDTA-facilitated method as described (Zhai et al., 2014; Chao et al., 2021). After collection, the fresh weight of leaves was recorded, and the phloem collections were diluted with 1.5 ml of 5% HNO₃ for subsequent detection by inductively coupled plasma MS (ICP-MS, Thermo Fisher Scientific, United States).

Harvested plants were first rinsed with 5 mM CaCl₂ for 5 min and then rinsed three times with ultrapure water. Afterward, the samples were dried in an oven at 65°C for 72 h, and the dried weight was recorded. The plant samples were then digested using HNO₃ at 140°C until the mixture became clear. After dilution with ultrapure water and filtering with a 0.22-μm filter, the Fe and other element concentrations were analyzed by a microwave plasma-atomic emission spectroscope (MP-AES, Agilent Technologies, Santa Clara, CA, United States).

Both Fe(II) and Fe(III) were detected by the Perls/DAB (3,3′-diaminobenzidine tetrahydrochloride) method as described in the study of Roschzttardtz et al. (2013). In brief, plant tissues were rinsed with 2 mM CaSO₄ and 10 mM EDTA for 5 min and washed three times with ultrapure water. Then, they were fixed in fixation solution [methanol/chloroform/glacial acetic acid (6:3:1)] for 1 h. After rinsing, the materials were incubated in staining solution [2% K₄Fe(Cn)₆ and 2% HCl] for 1 h. These samples were then incubated in the prepared solution (0.01 M NaN₃ and 0.3% H₂O₂ in methanol) for 1 h. Finally, the samples were incubated in intensification solution (0.025% DAB, 0.005% H₂O₂ in 0.1 M phosphate buffer, pH 7.0) for 5–15 min. For Turnbull staining, the plant samples were incubated for 1 h in 2% K₃Fe(CN)₆ and 2% HCl as described (Sperotto et al., 2010). After staining, the samples were imbedded in 4% agarose. Approximately 150-μm cross sections were cut with a vibratome (ZQP-86, Shanghai Zhixin Instrument, Shanghai, China). The stained samples were observed using a microscope (Nikon, Tokyo, Japan).

The cell wall was extracted as described (Zhu et al., 2020). Then, the Fe adsorption kinetics of cell wall were analyzed as described earlier (Jin et al., 2007). In brief, 10 mg dried cell wall powder was loaded into a 2-ml column with a filter at both ends. The composition of adsorption solution consisted of 0.1 m sodium acetate (pH 5.0) solution with 50 μM Fe(II)–citrate [prepared from Fe(NH₄)₂⋅(SO₄)₂⋅6H₂O and citrate in equimolar proportions] or 50 μM Fe(III)–citrate (prepared from FeCl₃ and citrate in equimolar proportions). The solution was pumped by a peristaltic pump at a speed of 4.4 ml min^–1 passing through the column. Then, the collected solution was reduced with 2% ascorbic acid and determined by 0.2% 2,2-bipyridyl at 520 nm (Heaney and Davison, 1977).

The data were analyzed by one-way and two-way analysis of variance (ANOVA) with Tukey’s test using SPSS Statistics version 20.0. A p-value <0.05 was considered statistically significant.