The rice cultivar (Dongjin) seeds obtained from the Rural Development Administration (RDA, South Korea) were used in this study. Uniform and healthy seeds were surface-sterilized with 1% (v/v) sodium hypochlorite solution (NaOCl). Then, the seeds were germinated in a Petri dish for 3 days in the dark in a temperature-controlled room at 37°C. After germination, 3-day-old identical seedlings were selected and transferred to an aquaponic growing foam board container with half-strength Murashige and Skoog (½ MS) nutrient medium (Sigma-Aldrich, MO, United States). Then, the plants were grown in MS medium with HU and IAA (alone or combined, for 3 days. Four treatment groups of plants were maintained: control (CK), HU (1 mM), IAA (0.3 μM), and HU (1 mM) + IAA (0.3 μM). Plants were kept in a temperature-controlled growth chamber set at 28°C with 60% relative humidity and a photoperiod of 16/8 h (light/dark). The shoot and root samples were collected from 6-day-old seedlings and stored at –80°C for further experiments.

The method of Arnon (1949) was followed to measure the total chlorophyll, chlorophyll a, and chlorophyll b contents. Briefly, leaf samples of 200 mg were taken in test tubes and then added to 2 ml of 80% acetone. These samples were incubated at 65°C until the samples became colorless. The chlorophyll contents were read at 645 and 663 nm. The amount of chlorophyll content was calculated and expressed as μg/ml. Photosynthetic efficiency as the quantum yield of photosystem II (i.e., F_v/F_m) was measured as described by Chen et al. (2019) using an ultra-portable modulated chlorophyll fluorometer (Mini-PAM II Photosynthesis Yield Analyzer, WALZ, Germany). The minimal (F₀) and maximal fluorescence yield of the dark-adapted state (Fm) emissions were assessed in the leaves after 20 min of dark adaptation, and the maximum quantum yield of PSII was calculated as Fv/Fm = (Fm–F₀)/Fm.

The H₂O₂ superoxide generated inside the cell was observed using DAB staining. The root samples were immersed with DAB-hydrogen chloride (HCl) solution (0.5 mg/ml, pH 3.8) for 30 min with a mild shaking in an incubator at room temperature. The DAB reacted with H₂O₂ to develop the brown product in roots, which was further observed and captured using a light microscope. The cell death of the roots of the rice was observed using PI. The harvested roots around 15 mm were loaded with 20 μM of PI for 15 min at 25°C. The fluorescence was captured using the Olympus FluoView FV1,000 confocal microscope at an excitation of 535 nm and emission of 615 nm.

The H₂O₂ content was estimated according to the method previously described by Velikova et al. (2000). Root samples (0.25 g) were homogenized with 3 ml of 0.1% (w/v) trichloroacetic acid (TCA) in an ice bath. The supernatant was collected after centrifugation at 12,000 × g for 15 min. Next, to the 0.5 ml of 10 mM potassium phosphate buffer (pH 7.0) and 0.75 ml of 1 M KI, 0.5 ml aliquot of the supernatant was added. The H₂O₂ content was determined based on the absorbance of the supernatant at 390 nm, and it was expressed as nmol g^–1 fresh weight. Lipid peroxidation was measured by estimating MDA according to the method of Heath and Packer (1968). Root samples (0.25 g) were homogenized with 5% (w/v) TCA and centrifuged at 11,500 × g for 12 min. The supernatant was mixed with 20% TCA containing 0.5% TBA and heated at 95°C for 30 min. The MDA content was measured from the difference in absorbance at 532 and 600 nm, respectively, and expressed as nmol g^–1 fresh weight.

The electrolyte leakage (EC) was measured as previously described by Dionisio-Sese and Tobita (1998). Fresh leaf samples (200 mg) were cut into pieces (approximately 5 mm) and placed in test tubes containing 10 ml of deionized water. The test tubes were fixed with plastic caps and incubated in a water bath (32°C). After 2 h, the initial EC of the medium (EC₁) was quantified using the conductivity meter HI2300 (Hanna Instruments, Woonsocket, RI, United States). The samples were autoclaved at 121°C for 20 min, and the final EC (EC₂) was measured. Relative EC was calculated by the following formula:

Root samples (100 mg) were frozen in liquid nitrogen and homogenized in 200 μl of different extraction buffer: for ascorbate peroxidase (APX), catalase (CAT) (pH 7.0), peroxidase (POD) (pH 6.0), and superoxide dismutase (SOD) (pH 7.8). The homogenate was centrifuged at 12,000 rpm for 15 min at 4°C. The supernatant was collected to determine the activity of APX, CAT, POD, and SOD. Total proteins were determined with the same buffer solution, and the quantification was made using the Bradford method. The APX activity was quantified according to the decrease in the ascorbate content. The 1-ml reaction mixture contained 830 μl of 50 mM HEPES and 0.1 mM of EDTA buffer (pH 7.0), 50 μl 0.6 mM of ascorbic acid, 100 μl of 0.06 mM H₂O₂, and 20μl of crude enzyme solution. The APX activity was recorded at 290 nm, and the results were expressed as μmol g^–1 fresh weight (Nakano and Asada, 1981). The CAT activity was estimated upon the decrease of H₂O₂ content in a 1-ml reaction mixture containing 650 μl of 50 mM PBS (pH 7.0), 330 μl of 0.1% H₂O₂, and 20 μl of enzyme solution was prepared. The CAT activity was measured at 240 nm, and the results were expressed as μmol g^–1 fresh weight (Aebi, 1984). POD was estimated according to the production rate of pyrogallol. The 1-ml of the reaction mixture consists of 420 μl of 100 mM potassium phosphate buffer (pH 6.0), 320 μl of 5% pyrogallol, 6 μl of 30% H₂O₂, and 20 μl of enzyme solution. The change was recorded at 420 nm for 20 s after the reaction began and was expressed as μmol g^–1 fresh weight (Kwak et al., 1995). The SOD activity was detected by determining its ability to inhibit nitroblue tetrazolium (NBT) photochemical reduction. A 1-ml reaction mixture containing 500 μl of 50 mM PBS (pH 7.8), 100 μl and 20 μM of riboflavin, 100 μl and 150 mM of L-methionine, 100 μl and 750 μM of NBT, 100 μl and 100 μM of EDTA, 80 μl of H₂O, and 20 μl of crude enzyme solution was incubated for 20 min under a white fluorescent lamp. The SOD activity was measured at 560 nm (Beauchamp and Fridovich, 1971).

The antioxidant capacity of the samples was measured as previously described by Kumaran and Joel karunakaran (2006) based on the scavenging activity of the stable 1,1-diphenyl-2-picrylhydrazyl (DPPH) free radical. Briefly, various dilutions of the sample extracts were prepared. An aliquot of 500 μl of a diluted extract was added with 500 μl of freshly prepared 0.004% DPPH in methanol and incubated in the dark for 30 min at room temperature. The absorbance at 517 nm against blank was used to calculate [(A₀–A₁)/A₀] × 100, where A₀ is the absorbance of the control, and A₁ is the extract.

The glucose content was quantified using a commercial Starch Assay Kit (SA-20, Sigma-Aldrich, St. Louis, MO, United States), and all steps were performed strictly following the technical instruction. The absorbance of samples was measured at 340 nm and expressed as mg-g^–1dry weight. The sugar content was measured using Agilent 1100 series high-performance liquid chromatography (Agilent Technologies, CA, United States) with the RI -101, refractive index detector (shodex, Denmark). A total of 200 mg of the sample was finely powdered and dissolved in 1 ml of ethanol (80%) and incubated at 50°C for 30 min. Then centrifuged for 3,500 g for 15 min and was repeated twice to obtain all the available supernatant. Then the sample was loaded in a metacarp 87 H column and then eluted at 0.5 ml/min at 25°C. The mobile phase consisted of 0.005 M H₂SO₄. The standard curve was prepared by using glucose, fructose, and sucrose (mg/ml) at analytical purity. All the samples were repeated and measured three times.

A kit obtained from BioVision (Zurich, Switzerland) was used to measure apurinic/apyrimidinic (AP) sites in DNA (three replicates for each treatment group). According to the manufacturer’s instructions, fresh samples were harvested and ground into a fine powder using liquid nitrogen, and DNA was isolated using DNeasy Plant Mini Kit (Qiagen, Hilden, Germany). All genomic DNA samples were diluted to 0.1 μg/μl for accurate assay, and the absorbance of the sample was measured at 650 nm.

The quantitative analysis of phytohormones was performed as described by Pan et al. (2010). Fresh root samples of 50 mg were ground into a fine powder using liquid nitrogen and mixed with 500 μl of extraction solvent (2-propanol/H₂0/concentrated HCL 2:1:0.002, vol/vol/vol), vigorously shaken on a shaking bed for 30 min at 4°C. About 1 ml of dichloromethane was added and again kept in the shaker for 30 min at 4°C. And then centrifuged at 13,000 g for 5 min at 4°C. The supernatant was carefully transferred to a new 1.5-ml tube and concentrated using a nitrogen evaporator with nitrogen flow. The sample was redissolved in 0.1 ml methanol, and 50 μl of sample solution was injected into a C18 (reverse-phase) HPLC column for the (HPLC–ESI–MS/MS) analysis. Agilent 1100 HPLC (Agilent Technologies, CA, United States), Waters C18 column (150 × 2.1 mm, 5 μm), and API3000 MSMRM (Applied Biosystems, CA, United States) were used for the analysis. The elution gradient was carried out with a binary solvent system consisting of 0.1% formic acid in water (Solvent A) and 0.1% formic acid in methanol (solvent B) at a 0.5 ml/min flow rate. The content of the hormone (ABA, SA, JA, GA, CTK, and IAA) was determined using a standard external method (ng/g FW) with three biological replications.

Total RNA was isolated using the TRIzol reagent (Thermo Fisher Scientific, Waltham, MA, United States), according to the manufacturer’s instructions, and treated with RNase-free DNase I (Promega, WI, United States). The first-strand of complementary DNA (cDNA) was prepared by Revert Aid M-MuLV Reverse Transcriptase (Thermo Fisher Scientific, Waltham, MA, United States) according to the manufacturer’s guidelines. The primers used for quantitative real-time PCR (qRT-PCR) analysis are listed in Supplementary Table 1, Aliquots for qRT-PCR reactions included 20 μl of final volumes containing 1 μl of cDNA, 1 μl of each primer (10 μM), 7 μl of dH₂O, and 10 μl of SYBR Green PCR Master Mix kit (Toyobo, Japan). All reactions were performed three times in a 96-well reaction plate using the CFX96 Real-Time PCR detection system (Bio-Rad, CA, United States). Three biological replicates were performed for each sample. Ubiquitin gene from the rice was used to normalize, and the change in the transcripts was measured using the 2^–ΔΔ CT method.

Immunoblot analysis was performed according to the method of Li et al. (2011). About 200 mg of sample was finely powdered in liquid nitrogen. About 400 μl of aliquot of extraction buffer [62.5 mM TRIS-HCl (pH 7.4), 10% glycerol, 0.1% SDS, 2 mM EDTA, 1 mM phenylmethylsulphonyl fluoride (PMSF), and 5% (v/v) β-mercaptoethanol] was added; then, the sample mixture was vortexed and incubated on ice for 10 min. After two rounds of centrifugation at 12,000 × g for 10 min, at 4°C, the supernatants were collected and stored at –80°C until use. The protein concentrations of the samples of the rice tissues were obtained using the Bradford method. For immunoblot analysis, 30 μg of total protein samples were separated on SDS-PAGE 12.5% polyacrylamide gel and transferred to polyvinylidene fluoride (PVDF) membranes. Proteins were determined using a mouse anti-RbcL as a primary antibody (1:5,000; Agri sera AB, Vannas, Sweden) and horseradish peroxidase (HRP) conjugated anti-mouse as a secondary antibody (1:2,000). Using an ECL kit (Bio-Rad, Laboratories, United States), the antibody-protein complexes were visualized.

The result was shown as the mean of at least three replicates with standard deviation (SD). All the data were analyzed statistically by one-way ANOVA, and the mean differences were evaluated by Fisher’s post-test using the Minitab software (Minitab., Inc., State College, PA, United States). A significance threshold of p < 0.05 level was applied.

The 3-day-old rice seedlings were treated either with HU or IAA alone and in combination (HU + IAA) for 3 days to know how IAA influenced the HU stress. The results showed that the root length was significantly decreased under HU stress compared to the control, whereas HU + IAA mitigated the root length reduction caused by HU and improved the root length compared to HU-treated plants (Figures 1A,B). Similarly, plants treated with HU + IAA prevented the shoot growth and biomass decrease than the HU-treated plants (Figures 1A–C). Besides, the plants treated with HU significantly reduced chlorophyll content, whereas IAA-treated plants had a slight increase than the control (Figure 1D). When HU and IAA were applied together, chlorophyll content was significantly increased compared to HU-treated plants. Collectively, these results suggested that IAA reduced the effect of HU stress on plant growth.

The rice seedlings treated with HU, IAA, and HU + IAA were analyzed for photosynthesis-related parameters to see if IAA maintains photosynthesis performance under HU stress. The photochemical efficiency (Fv/Fm) of Photosystem II was decreased under HU stress compared to the control, while HU + IAA significantly increased the photochemical efficiency of the plants treated with HU (Figure 2A). The photosynthetic efficiency is directly linked to the Ribulose-1,5- bisphosphate carboxylase/oxygenase (Rubisco) enzyme activity. The HU stress significantly downregulated the Rubisco expression in transcript and protein levels compared to the IAA and control. However, it was significantly upregulated by HU + IAA (Figure 2B). Moreover, chloroplast synthesis and development are regulated by the integrated expression of plastid and nuclear genes. Thus, we examined the transcript level of the positive and negative regulators of the genes involved in chloroplast biogenesis. The HU stress suppressed the transcript levels of psbA, RbcL, and DVR genes that are key positive regulators of chloroplast formation and photoautotrophic growth. However, when IAA was combined with HU, the transcript levels of these genes were upregulated (Figure 2C). In contrast, the transcript levels of rpoA, rpoB, and rpoC1 genes related to the negative regulation of chloroplast formation were significantly upregulated under HU stress while they were downregulated following HU + IAA treatment (Figure 2D). These observations suggest that HU-induced adverse effects on chloroplast formation and development were enhanced upon HU + I AA treatment.

As shown in Figures 3A–C, the HU inhibits the activities of the RNRs enzyme that are highly temperature-sensitive. Compared to IAA and control plants, the HU-treated plants showed a chlorosis phenotype in leaves at low temperatures (20 and 23°C). However, this effect was slightly improved under favorable conditions (28°C) compared to control. Notably, HU + IAA-treated plant showed a recovery from chlorosis phenotype at all the temperatures. These results suggest that phenotypic development upon HU + IAA is closely associated with increased RNR activity under ambient temperatures.

To elucidate the potential role of IAA on the DNA damage induced by HU stress, the HU-induced DNA damage and the resulting alterations in purine/pyrimidine and carbohydrate metabolisms and their transporter activity were evaluated. Interestingly, in the HU-treated plants, the DNA damage in response to H₂O₂ was more evident with a substantial upregulation in the accumulation of apurinic/apyrimidinic (AP) sites than the control and IAA-treated plants (Figure 4A). Furthermore, the accumulation of sugar was stimulated in response to HU treatment. Compared to the control and the other treatments, sucrose, glucose, and fructose contents were significantly increased following the HU treatment (Figures 4B,C). Also, the transcript levels of sucrose transporter genes (SUT1, SUT2, and SWEET15) decreased in HU-treated plants compared to the control (Figure 4D). In contrast, a strong alleviating effect of IAA on HU was observed in HU + IAA treatment; the transcript levels of these genes were increased. It provided significant protection against DNA damage. Overall, these results indicate that IAA alleviated DNA damage and sugar transport inhibition, resulting in the decreased accumulation of sugars in the chloroplast.

The HU stress is known to disturb the RNR activity that negatively regulates the replication of chloroplast DNA. In contrast, auxin plays a prominent role in regulating both cell proliferation and cell expansion. Thus, to ascertain the specific cell cycle role of IAA in response to HU, we analyzed the transcription level of several genes related to the cell cycle. The RNR of large and small subunits (RNRL1 and RNRS2) are related to DNA duplication. Cyclin-dependent kinase (CDKA2) primarily expressed in the dividing zone of roots throughout the cell cycle, and their regulatory partner cyclins (CYCD3; 1 and CYCA1; 1) are required for the normal regulation of G1/S and G2/M transitions. Transcription factor 2 (E₂F₂) regulates cell cycle entry and progression in which the transcript levels of all these genes were significantly downregulated in HU treatment relative to the control (Figure 5A). Importantly, IAA and HU + IAA combination treatment displayed a remarkable upregulation of the RNRs, E₂F₂, and CDK-CYCs genes. In contrast, the CDK inhibitor, KRP3, negatively affects cell division, the transcript levels in both IAA and HU + IAA were decreased compared to HU and the control. Furthermore, the accumulation of dead cells triggered by HU treatment was reduced in the HU + IAA treatment (Figure 5B). These results indicate that the addition of IAA alleviated the HU-induced cell cycle arrest by stimulating the RNR.

To know the protective role of IAA in plant growth under HU stress, we examined the oxidative stress indicators (H₂O₂, MDA, and EL) and antioxidant enzyme activities. The H₂O₂ accumulation was visualized through histochemical staining with DAB reagent in root samples, and a solid brown color deposit was observed in the HU than in the control and IAA-treated plants. Furthermore, the H₂O₂ content significantly increased under HU stress, which was consistent with qualitative results. On the other hand, plants treated with HU + IAA showed a reduction in brown color deposit and H₂O₂ levels compared to HU-treated plants (Figures 6A,B). Likewise, MDA and EL levels were also decreased in HU + IAA-treated plants than in the HU-treated plants (Figure 6C and Supplementary Figure 1A). Moreover, under HU stress, the activities of all the enzymes (SOD, APX, CAT, and POD) were decreased (Figures 6D–F and Supplementary Figure 1B). In particular, the CAT activity was significantly reduced when compared to the control. Conversely, HU + IAA significantly increased the activities of all antioxidant enzymes and their radical scavenging capacity than in the HU-treated plants (Supplementary Figure 4). These results indicated that IAA promoted antioxidant enzyme activity, scavenging the excess H₂O₂ accumulated in plant cells under HU stress.

The inhibition of root growth by HU stress was reversed by HU + IAA treatment. Thus, we examined the role of IAA under HU stress using auxin biosynthesis genes (TAA2, TAA4, YUCCA, YUCCA3, and YUCCA6). The HU + IAA and IAA treatment showed a significant increase in the transcript levels of all the biosynthetic genes than the control. A slight increase was observed in HU, indicating that hormone levels were altered upon the stress treatment (Figure 7A). In addition, we examined the transcript level of the genes encoding UDP (UGT90A1) and auxin-specific (IAGT1) glucosyltransferases responsible for mediating the glycosylation reaction. The transcript levels of both the genes (UGT90A1 and IAGT1) were significantly upregulated in HU + IAA than in the other treatments (Figure 7B). Importantly, the auxin transporter gene (PIN5B) is vital for exporting and regulating free auxin and their conjugates between the cytoplasm and endoplasmic reticulum (ER). The HU + IAA and IAA displayed a significant upregulation in the transcript level of the PIN5B gene (Supplementary Figure 2). This result indicates the transportation of IAA and HU + IAA-induced hormone conjugates from chloroplast to ER. We also investigated the impact of these treatments on the root growth of the putative glycosyltransferase, a csld1 mutant. Treatment with either HU or HU + IAA promoted the root growth of csld1 mutant compared to IAA and untreated condition (Figure 7C). Moreover, the transcript levels of cellulose synthase genes (CSLD4, CesA4) in HU and HU + IAA were also significantly upregulated compared to IAA and control (Supplementary Figure 2). These results indicated that the sugar accumulation triggered by HU treatment involves hormone conjugation upon HU + IAA treatment. The recovery of glycosylation activity in the csld1 mutant enhanced the plant growth and development.

Plant hormones are essential molecular signals that stimulate plant growth and induce plant tolerance under stress conditions. Therefore, we investigated whether IAA-glucose conjugation is involved in mitigating the plant architecture by triggering the phytohormones. So, we carried out HPLC-MS/MS analysis to quantify the hormone content in the roots under CK, HU, IAA, and HU + IAA treatments. The HU-treated plants showed a significant increase in JA, SA, GA, ABA, and IAA levels compared to the control and IAA-treated plants, with decreasing the CTK levels. While the ABA, SA, CTK, and GA levels were upregulated in HU + IAA compared to all other treatments (Table 1). Transcript-level analysis of SA marker genes (NPR2, NPR3, and WRKY45) also showed a significant upregulation in HU + IAA than other treatments (Supplementary Figure 3). These results indicate that combined treatment (HU + IAA) induced hormone conjugation, which regulates plant growth by integrating multiple hormone pathways.

The phytohormone conjugates do not function as signaling molecules; instead, they fulfill other functions, such as transport and storage. Thus, we performed a transcript level analysis using auxin signaling and downstream marker genes (OsARF1,OsARF4, OsARF5, OsARF7, OsARF8, OsIAA7, OsGH3-1, and OsGH3-5) to determine the role of IAA-glucose conjugation in auxin signal transduction. The transcript level was significantly downregulated in the IAA-glucose conjugation-induced HU + IAA treatment than the other treatments (Figure 8), which shows that auxin signaling and homeostasis was perturbated in HU + IAA glycosylated plants.