SnRK2.6 phosphorylates sucrose transporter ZmSUT1 to enhance yield by modulating leaf senescence in maize

Tao Yang; Yunqin Huang; Zan Wu; Zhenglan Long; Hongwei Zhu; Shanshan Wang; Lei Tang; Ruirui Li; Zhengchen Liang; Haoyu Zhang; Yongcai Huang; Xiaoling Li; Xiaoduo Lu; Yongrui Wu

doi:10.1126/sciadv.aeb2472

. 2026 May 6;12(19):eaeb2472. doi: 10.1126/sciadv.aeb2472

SnRK2.6 phosphorylates sucrose transporter ZmSUT1 to enhance yield by modulating leaf senescence in maize

Tao Yang ^1,^2,^*,^†, Yunqin Huang ^1,^†, Zan Wu ^1,^†, Zhenglan Long ¹, Hongwei Zhu ¹, Shanshan Wang ², Lei Tang ¹, Ruirui Li ¹, Zhengchen Liang ¹, Haoyu Zhang ¹, Yongcai Huang ¹, Xiaoling Li ¹, Xiaoduo Lu ³, Yongrui Wu ^2,^*

PMCID: PMC13148305 PMID: 42090494

Abstract

The extended stay-green trait is beneficial to increase photosynthetic efficiency, thereby enhancing crop yield. However, factors and mechanisms affecting this process remain largely unknown. Here, we cloned a leaf precocious senescence gene (lps1-1) and created additional null alleles. All lps1 mutants exhibited accelerated leaf senescence with reduced chlorophyll contents and photosynthetic efficiency. The 100-kernel weight and storage reserve content were decreased in lps1 kernels, while the opposite was observed in overexpression lines. Lps1 encoded a protein kinase SnRK2.6 that phosphorylated the sucrose transporter ZmSUT1 in the BETL region, thereby enhancing protein stability, homodimer formation ability, and sucrose transport activity. Overexpression of ZmSUT1 delayed leaf senescence and enhanced photosynthetic capacity and kernel weight, while mutation of ZmSUT1 did the opposite. This mechanism appears to be conserved in grasses. Our findings provide insights for yield and quality improvement in crops through delaying leaf senescence.

SnRK2.6-phosphorylated ZmSUT1 enhances maize kernel yield and nutrient quality with delayed leaf senescence progress.

INTRODUCTION

With the growing population and decreasing arable land, agriculture development and world food security face great challenges. The highly efficient nutrient allocation from leaves to seeds is one of the most important determinants for genetic improvement of crop yield and nutrient quality (1). Leaf senescence is a genetically intricate programmed process, accompanied by nutrient remobilization from senescing leaves to seeds (2). Leaf senescence is an endogenous programmed and highly ordered process (3). Among them, a series of internal and environmental events orchestrate leaf senescence process, involving plant hormone, sugar signaling response, photosynthetic status, pathogenous reaction, and environmental stress (4, 5). After senescence initiation, chlorophyll degrades rapidly, reactive oxygen species accumulate, and programmed cell death is subsequently triggered, followed by nutrient remobilization into sink organs. A positive correlation between delayed leaf senescence and high yield has been reported and applied in crop breeding improvement (6–8). Maize is one of the most important crops worldwide, serving as the major supply for food and industrial materials. Since 2000, maize yield steadily improved, largely contributed by functionally delayed leaf senescence (8). Both high-yield and high-quality traits are desirable for maize breeding improvement, making it urgent for developing stay-green varieties. Therefore, understanding the molecular mechanism underlying leaf senescence is crucial for maize breeding.

Sucrose, the primary carbohydrate product from photosynthetic leaves, is transported to kernels, which is mediated by sugar transporters (9–14). During grain filling, abundant proteins and starch are rapidly synthesized, which requires a large amount of sucrose allocated from leaves to kernels through the basal endosperm transfer layer (BETL) region (15, 16). Sucrose is broken into glucose and fructose by cell wall invertase, transported into endosperm by Sugar Will Eventually Be Exporter Transporters, and then resynthesized into sucrose (17, 18). Sucrose can also be directly transported into endosperm by sucrose transporters (19). The first cloned sucrose transporter gene is ZmSUT1, which facilitates efficient phloem loading of sucrose into companion cells (20). Recently, spatial transcriptomics demonstrated that ZmSUT1 is highly expressed in the BETL region of the endosperm (11, 21). However, the detailed molecular mechanisms of ZmSUT1 governing kernel yield improvement and leaf senescence remain unknown.

Developing extended stay-green varieties are regarded as an effective way to enhance maize yield. In this study, we characterized a leaf precocious senescence mutant (lps1-1) that was induced by ethyl methanesulfonate (EMS). The lps1-1 leaves exhibited decreased chlorophyll contents and photosynthetic efficiency and reduced kernel weight and nutrient storage accumulation. Lps1-1 was determined to encode a protein kinase, SnRK2.6. The overexpression of SnRK2.6 resulted in delayed leaf senescence and yield improvement. Furthermore, SnRK2.6 interacted with and phosphorylated ZmSUT1 in the BETL region to enhance its protein stability, homodimer-forming capacity, and sucrose transport. The overexpression of ZmSUT1 displayed similarly extended stay-green phenotype and enhanced kernel weight as observed for the SnRK2.6 overexpression lines. This study provides previously unidentified insights and strategies for developing elite stay-green maize varieties with both high-yield and high-quality traits, advancing the field of crop breeding.

RESULTS

The lps1-1 mutant exhibits precocious leaf senescence, decreased kernel weight and storage reserve content

To decipher the genetic basis of leaf senescence and yield achievement in maize, we screened a large number of EMS-induced B73 mutants, resulting in identification of a precocious leaf senescence mutant lps1-1 (Fig. 1, A to C). Before 60 days after germination (DAG), although both wild-type and lps1-1 leaves exhibited a similar green color, the chlorophyll content of lps1-1 leaves was lower than wild type (fig. S1, A to C). At 70 DAG, the lps1-1 leaves begun to turn red, showing increased accumulation of anthocyanin and flavonoids, while the wild-type leaves remained green with higher chlorophyll contents (fig. S1, D to F). The lps1-1 plants showed lower stem weight and root weight, resulting in reduced plant biomass (fig. S2, A to C). The number of stem internodes was not affected, but the averaged internode length was significantly reduced in lps1-1 (fig. S2, D and E). At 80 DAG, while the wild-type leaves begun to lose their green color, the lps1-1 leaves had become greatly withered, exhibiting a marked reduction in chlorophyll content (fig. S1G). By 89 DAG, while the wild-type leaves had withered, the lps1-1 leaves had thoroughly died off, displaying a greatly lower chlorophyll level than wild type (fig. S1H). When harvested at 96 DAG, both wild-type and lps1-1 leaves had completely withered with extremely low chlorophyll contents (fig. S1I). Furthermore, we assessed the expression pattern of several important genes related to senescence (22). These genes showed significantly higher expression levels in lps1-1 leaves than wild type at 75 DAG, confirming early senescence occurring in lps1-1 (fig. S3, A to E).

The leaf tip regions exhibited the lowest chlorophyll content in comparison to both the middle and base regions. When comparing the wild-type and lps1-1 leaves at 75 DAG, a marked reduction in chlorophyll levels was observed across all leaf regions (Fig. 1, A to D). The photosynthetic efficiency in lps1-1 was 78.52% lower than that in wild type, presenting a notably decreased pattern (Fig. 1E). Accordingly, several key genes associated with photosynthesis were markedly down-regulated in lps1-1 leaves (fig. S3, F to J) (22). Abnormal reactive oxygen species (ROS) accumulation causes cell death, thereby resulting in accelerated leaf senescence (23). The 3,3′-diaminobenzidine (DAB) staining revealed a distinctly deeper brown color in lps1-1 leaves than wild type (Fig. 1F). Early senescence is also associated with accelerated programmed cell death (24, 25). Trypan blue staining displayed more dense blue spots in lps1-1 leaves than wild type, indicating accelerated programmed cell death development (Fig. 1F). Malondialdehyde (MDA), one of the lipid oxidation byproducts indicative cell death, accumulated at higher levels across all the regions of lps1-1 leaves than wild type (Fig. 1G). Previous studies have indicated that sugar functions as a signal molecule to trigger leaf senescence (23, 26). Consequently, we quantified the sugar contents to investigate the association between advanced leaf senescence and carbohydrate accumulation. At 75 DAG, the levels of soluble sugars and sucrose were significantly higher in lps1-1 leaves than wild type (Fig. 1, H and I).

At maturity, lps1-1 kernels displayed reduced kernel dimension and 100-kernel weight (Fig. 1, J and K, and fig. S4A). The fresh weight of the lps1-1 developing kernels was also consistently lighter than wild type from 6 to 30 days after pollination (DAP) (fig. S4B). Paraffin section observation also displayed reduced kernel size and area of the BETL region in lps1-1 at 18 DAP compared to wild type (fig. S4, C to E). In addition, we quantified the contents of different storage compounds, revealing that the levels of soluble sugars, sucrose, and starch in lps1-1 were significantly decreased compared to wild type (Fig. 1, L to N). The lower starch level was also observed via an iodine potassium iodide (I2-KI) staining assay (fig. S4F). SDS–polyacrylamide gel electrophoresis (SDS-PAGE) and protein quantification analysis showed that both zein and nonzein protein levels were notably decreased in lps1-1 kernels, leading to significantly decreased total protein accumulation (Fig. 1O and fig. S4, G to I). Collectively, lps1-1 exhibited accelerated leaf senescence with decreased photosynthetic efficiency, kernel weight, and storage reserve accumulation.

Lps1-1 encodes a protein kinase SnRK2.6

Through reciprocal crossing between lps1-1 and the B73 inbred line, both resulting F₁ plants exhibited green leaves at 74 DAG, while the lps1-1 leaves were notably withered with a marked chlorophyll content reduction (fig. S5, A and B). The F₂ plants segregated the normal and lps1-1 phenotypes at a 3:1 ratio (normal: senescence =100: 29, X² = 2.94 < 3.84), indicating that lps1-1 resulted from a monogenic mutation.

We further used the MutMap strategy to clone the lps1-1 gene. In the segregating F₂ populations, leaf samples were harvested from 96 normal and mutant plants each, which were then pooled separately to construct the DNA libraries (27). The comparison of the single-nucleotide polymorphism index revealed a peak on chromosome 4, encompassing an interval ranging from 116 to 126 M (Fig. 2, A and B). This interval contained 48 genes as listed in table S1. Genomic sequencing revealed that lps1-1 contained a G-to-A nucleotide transition in the splice acceptor site of the first intron of the gene Zm00001eb181940, encoding a protein kinase SnRK2.6 and causing frameshift termination of the protein translation (Fig. 2C and fig. S6, A to D). The homozygous mutation was uniformly detected in early-senescence plants, but not in stay-green plants.

Fig. 2. — (A) Mapping by sequencing of *lps1-1*. SNP index distribution along chromosomes based on the MutMap analysis. (B) Peak detail on chromosome 4. (C) Schematic representation of the *SnRK2.6* structure with the mutant alleles indicated. Black rectangles and lines indicate exons and introns, respectively. The red triangles indicate the mutation sites in *lps1-1*, *lps1-2*, *lps1-3*, and *lps1-4* alleles. gRNA1, guide RNA 1. (D) Plant phenotypes of the cross of *lps1-1* × *lps1-2* at 70 DAG. (E and F) Photosynthetic efficiency (E) and chlorophyll content (F) of the *lps1-1* × *lps1-2* leaves at 70 DAG. (G and H) 100-kernel weight (G) and starch contents (H) of the *lps1-1* × *lps1-2* mature kernels. (I) Plant phenotypes of *lps1-3* and *lps1-4* at 74 DAG. (J and K) Photosynthetic efficiency (J) and chlorophyll contents (K) of *lps1-3* and *lps1-4* at 74 DAG. (L) Ear phenotypes of *lps1-3* and *lps1-4*. Scale bar, 1 cm. (M and N) Starch (M) and total protein (N) levels of the *lps1-3* and *lps1-4* mature kernels. ***P < 0.001.

To confirm SnRK2.6 mutation responsible for the precocious leaf senescence phenotype, we searched the maize EMS mutant library (http://maizeems.qlnu.edu.cn/) and identified a different splicing defective mutant (EMS4-20024b) in SnRK2.6 (lps1-2). A single G-to-A substitution at the right splice site of the first intron resulted in the intron retention and thus premature termination (fig. S6, A to D). Same as the observation in lps1-1, reduced chlorophyll contents were detected in lps1-2 leaves compared to wild type at 50 DAG, although lps1-2 displayed similar green leaves as wild type (fig. S7A). At 80 DAG, lps1-2 leaves were withered with markedly low chlorophyll contents compared to wild type (fig. S7B). Cross of lps1-1 and lps1-2 failed to complement their mutant phenotypes. The F₁ leaves begun to turn red and withered with significantly decreased photosynthetic efficiency and chlorophyll level and enhanced anthocyanin and flavonoid contents at 70 DAG, similar to the observation in lps1-1 leaves (Fig. 2, D to F, and fig. S7, C to E). The F1 mature kernels also detected decreased 100-kernel weight, starch, soluble sugar, zein, nonzein, and total protein levels (Fig. 2, G and H, and fig. S7, F to J).

We also created null mutations of SnRK2.6 in the B104 inbred line through CRISPR-Cas9. Two independent transgenic lines were generated and designated as lps1-3 and lps1-4, respectively (fig. S6, E to G). Lps1-3 harbored a 197–base pair (bp) deletion, and lps1-4 carried a 4-bp insertion in the first exon. The latter was also found with a 118-bp deletion occurring in the second intron (fig. S6, E to G). Both lps1-3 and lps1-4 plants exhibited early senescence at 74 DAG with low photosynthetic efficiency, reduced chlorophyll contents, accelerated programmed cell death development, hyperaccumulated ROS, sucrose, and soluble sugar levels (Fig. 2, I to K, and fig. S8, A to C). Same as in lps1-1, the expression levels of several genes associated with senescence and photosynthesis were significantly increased and decreased in lps1-3 and lps1-4, respectively (fig. S9, A to G). The kernel dimensions in lps1-3 and lps1-4 were markedly reduced compared to wild type, leading to decreased 100-kernel weight (Fig. 2L and fig. S8, D and E). Sugar and protein accumulation exhibited markedly decreased levels in lps1-3 and lps1-4, encompassing sucrose, soluble sugar, starch, zein, nonzein, and total proteins (Fig. 2, M and N, and fig. S8, F to K). Together, these results confirm that mutation in SnRK2.6 is responsible for the precocious leaf senescence phenotype in lps1.

RNA in situ hybridization showed that SnRK2.6 was expressed throughout the developing kernel at 20 DAP with strong staining intensity in the BETL region and the embryo (fig. S10, A and B). In leaves at 60 DAG, SnRK2.6 transcripts were broadly detectable across the tissue, with pronounced signals in vascular bundles, particularly in the lateral veins. Confocal imaging further revealed that the SnRK2.6 protein was located in both cytoplasm and nuclei in Nicotiana benthamiana leaf cells (fig. S10C).

Overexpression of SnRK2.6 delays leaf senescence with enhanced kernel weight and storage reserve synthesis

To further elucidate the roles of SnRK2.6 in leaf senescence and yield improvement, the SnRK2.6 overexpression lines were generated driven by the Ubiquitin promoter, and two independent overexpression lines (OE1 and OE2) were selected for further analysis. The two OE lines detected markedly elevated SnRK2.6 transcript levels in the leaves at 60 DAG and developing kernels at 20 DAP (Fig. 3A and fig. S11A). Immunoblot analysis using the anti-Flag antibody detected distinct target bands in both overexpression lines, but not in wild type (Fig. 3B and fig. S11B). Both OE lines and wild type exhibited green leaves at 50 DAG, while OE lines had higher chlorophyll contents (fig. S11, C and D). At 74 DAG, we observed delayed leaf senescence in OE lines, which exhibited enhanced photosynthetic efficiency and chlorophyll contents compared to wild type (Fig. 3, C to E). Consistent with this observation, transcript levels of several key senescence-associated genes were down-regulated in the OE leaves at 74 DAG, while those of photosynthesis-related genes were up-regulated in comparison with wild type (fig. S12, A to H). The sucrose and soluble sugar levels were lower in OE than wild type at 74 DAG (Fig. 3F and fig. S11E). By 84 DAG, the wild-type plants became markedly senescent, exhibiting extremely reduced chlorophyll contents, whereas the OE leaves showed partial leaf senescence (fig. S11, F and G).

The OE mature kernels exhibited increased dimension and 100-kernel weight (Fig. 3, G and H, and fig. S13, A and B). Paraffin section also showed increased kernel area in OE at 18 DAP compared to wild type (fig. S13C). We further quantified the contents of different storage compounds, displaying higher soluble sugars, sucrose, starch, zein, nonzein, and total proteins in the OE kernels (Fig. 3, I to K, and fig. S13, D to G). In addition, the complementation test was conducted by crossing lps1-1 with OE1, followed by two rounds of self-pollination. The resulting progeny plants closely resembled wild type in terms of photosynthetic efficiency and chlorophyll content, further substantiating the role of SnRK2.6 in regulating leaf senescence (fig. S14, A to C).

OsSAPK6 was the SnRK2.6 homolog in rice with 90.19% protein sequence similarity (fig. S15). To investigate whether OsSAPK6 and SnRK2.6 share a conserved function in regulation of leaf senescence and yield achievement, OsSAPK6 was overexpressed in Zhonghua11 and designated as SAPK6-OE. Both wild-type and SAPK6-OE plants displayed green leaves at 60 DAG with similar chlorophyll contents (fig. S16, A and B). At 80 DAG, the wild-type leaves were detected with lower chlorophyll levels than SAPK6-OE, although both plants still exhibited green leaves (fig. S16, C and D). From 90 to 100 DAG, higher chlorophyll contents were still detected in SAPK6-OE leaves than wild type (Fig. 3, I to M, and fig. S16, E to I). DAB and trypan blue staining assays revealed paler reddish brown and fewer blue spots in SAPK6-OE than wild type at 100 DAG (ig. S16, J to L). Notably, SAPK6-OE exhibited greater panicle length and higher grain number per panicle than wild type (fig. S16, M to O). The SAPK6-OE mature kernels displayed larger kernel dimension with enhanced 1000-kernel weight (Fig. 3, N and O). Both starch and soluble sugar contents were significantly elevated in SAPK6-OE kernels (Fig. 3P and fig. S16P). SDS-PAGE analysis revealed that the synthesis of major storage protein components—including glutelin precursors, glutelin acid and basic subunits, α-globulin, and prolamin—were pronouncedly enhanced in SAPK6-OE kernels, resulting in increased total protein accumulation (Fig. 3Q and fig. S16Q).

SnRK2.6 interacts with the sucrose transporter ZmSUT1

Given that lps1 encodes a protein kinase to carry out subsequent substrate phosphorylation, we further performed quantitative DIA-based phosphoproteomics to dissect the specific phosphorylation events mediated by SnRK2.6 using the lps1-1 and wild-type leaves at 75 DAG. Totally, we obtained 14,889 concrete phosphopeptides containing 6205 phosphosites, responsible for 2518 phosphoproteins (fig. S17A and table S2). Among them, 757 phosphoproteins displayed lower abundance in lps1-1, whereas 756 exhibited higher accumulation with the threshold of log₂ fold change ≥ 1 and P < 0.05. These identified phosphorylation sites were distributed in serine (37.4%), threonine (43.6%), and tyrosine (19.0%), which are the most conserved residues in plants (fig. S17B). We also identified the substrate motif (fig. S17C), in which the motif (R××S) was markedly defined with the canonical SnRK2 substrate motif (28–30).

ZmSUT1 was the first cloned sucrose transporter from C4 species, responsible for sucrose allocation from source to sink (20). The mutation of ZmSUT1 displays stunted plant growth and reproductive development defects, along with decreased photosynthetic efficiency, reduced chlorophyll accumulation, and accelerated leaf senescence, similar with lps1-1 (31, 32). The ZmSUT1 expression pattern showed no significant alterations in lps1-1 and SnRK2.6-OE leaves at 70 DAG and BETL regions at 20 DAP in comparison with wild type (fig. S18, A to E). However, immunoblot analysis revealed a notably decreased ZmSUT1 protein accumulation in the leaves and BETL region of lps1-1 (Fig. 4A and fig. S18B). The phosphorylation level of ZmSUT1 was much lower in lps1-1 leaves than wild type through the phosphoproteome analysis (table S2 and fig. S17D). Accordingly, we speculated that ZmSUT1 was a substrate of SnRK2.6 to play a role in leaf senescence and yield improvement. Through immunoprecipitation followed by liquid chromatography–mass spectrometry (LC-MS) analysis of the SnRK2.6-OE leaf and kernel mixtures, ZmSUT1 was identified to be one of the potential interactors with SnRK2.6 (table S3). Pull-down, split luciferase complementation imaging (LCI), and biomolecular fluorescence complementation (BiFC) assays were carried out, which demonstrated that SnRK2.6 and ZmSUT1 had strong protein-protein interaction (Fig. 4, B to D). We also performed semi-in vitro pull-down assay to validate their interaction, in which ZmSUT1, purified from the maize leaf protoplast cells, was targeted by the recombinant SnRK2.6-MBP protein (Fig. 4E). Co-immunoprecipitation assays in N. benthamiana leaves revealed that both SnRK2.6–hemagglutinin (HA) and SnRK2.6–green fluorescent protein (GFP) were detected in the immunoprecipitated ZmSUT1-Flag complex, further demonstrating their interaction in vivo (Fig. 4, F and G). These data collectively confirmed that SnRK2.6 physically interacted with ZmSUT1 in vitro and in vivo.

Fig. 4. — (A) Immunoblot analysis detecting the protein levels of ZmSUT1 and ZmSUT7 in the BETL region of the *lps1-1* kernels at 20 DAP. The protein levels were quantified using the ImageJ software. (B) In vitro pull-down assay showing the interaction between SnRK2.6 and ZmSUT1. (C) BIFC assay showing the interaction between SnRK2.6 and ZmSUT1 in *N. benthamiana* leaf cells. Scale bars, 100 μm. (D) Split LCI assay showing the interaction between SnRK2.6 and ZmSUT1 in *N. benthamiana* leaf cells. (E) Semi in vitro pull-down assay detecting the interaction between SnRK2.6 and ZmSUT1. (F and G) Co-immunoprecipitation assay detecting the interaction between SnRK2.6 and ZmSUT1. The indicated plasmid mixtures were cotransformed into the *N. benthamiana* leaf cells for transient protein expression. IP, immunoprecipitation.

ZmSUT1 mutation results in decreased kernel weight and accelerated leaf senescence

Previous studies showed that ZmSUT1 mutation leads to functional disruption in plant growth, pollen fertility, and ear development (31, 32). It is impractical to investigate the impact on kernel yield and nutrient quality achievements through constructing the zmsut1 knockout mutants. Accordingly, the zmsut1 RNA interfering specifically targeting ZmSUT1 and its duplicate copy ZmSUT7 was created (fig. S19), which was driven by the BETL-specific expressed BETL9 promoter. Two independently transgenic lines were recovered, which were designated as zmsut1/7 RNAi-1 and zmsut1/7 RNAi-2 (Fig. 5A). Both ZmSUT1 and ZmSUT7 transcript and protein levels were markedly decreased in the BETL region of the zmsut1/7 RNAi kernels at 20 DAP (fig. S20, A to C). Phenotypic analysis revealed that zmsut1/7 RNAi mature kernels exhibited reduced dimension with markedly decreased 100-kernel weight (Fig. 5, A and B). Paraffin section observation further revealed that the zmsut1/7 RNAi developing kernels at 18 DAP displayed decreased size (fig. S20D). The fresh kernel weight from 10 to 30 DAP was also significantly lower in zmsut1/7 RNAi-1 than wild type (fig. S20E). The lower contents of sucrose, soluble sugars, and starch were detected in zmsut1/7 RNAi in comparison with wild type (Fig. 5C and fig. S20, F and G). I2-KI staining assay detected lighter coloration in zmsut1/7 RNAi endosperm at 18 DAP compared to wild type (Fig. 5D). SDS-PAGE and protein quantification analysis revealed that both zein and nonzein protein levels were significantly decreased in zmsut1/7 RNAi kernels, resulting in total protein accumulation reduction (Fig. 5E and fig. S20, H to J).

Fig. 5. — (A) Ear phenotype of *zmsut1/7 RNAi*. Scale bar, 1 cm. (B and C) One hundred–kernel weight (B) and starch contents (C) of the *zmsut1/7 RNAi* mature kernels. (D) I2-KI staining assay showing the starch contents in the developing *zmsut1/7 RNAi* kernels at 18 DAP. Scale bar, 1 mm. (E) SDS-PAGE analysis of zein and nonzein protein contents. γ27, 27-kDa γ-zein; α22, 22-kDa α-zein; α19, 19-kDa α-zein; γ16, 16-kDa γ-zein; β15, 15-kDa β-zein; δ10, 10-kDa δ-zein. (F) Plant phenotypes of the *zmsut1/7 RNAi* lines at 64 DAG. (G and H) Photosynthetic efficiency (G) and chlorophyll contents (H) in the *zmsut1/7 RNAi* leaves at 64 DAG. (I) DAB and trypan blue staining assays of the *zmsut1/7 RNAi* leaves. Scale bars, 1 mm. (J and K) Sucrose (J) and soluble sugar (K) contents of the *zmsut1/7 RNAi* leaves at 64 DAG. (L and M) Plant (L) and ear leaf (M) phenotypes of the *zmsut1/7 RNAi-1*; *lps1-1* double mutant at 80 DAG. Scale bar, 1 cm. (N and O) Photosynthetic efficiency (N) and chlorophyll contents (O) of the *zmsut1/7 RNAi-1*; *lps1-1* double mutant at 80 DAG. *P < 0.05, **P < 0.01, and ***P < 0.001.

At 64 DAG, the zmsut1/7 RNAi plants displayed accelerated leaf senescence, exhibiting reduced photosynthetic efficiency and chlorophyll contents (Fig. 5, F to H, and fig. S21A). The expression levels of key genes related to photosynthesis and senescence were markedly decreased and elevated in zmsut1/7 RNAi, respectively (fig. S22, A to N). Intriguingly, the ZmSUT1 transcript and protein levels were also decreased in leaves, indicating the regulatory feedback from kernels to leaves (fig. S21, B and C). In addition, DAB and trypan blue staining assays revealed that the zmsut1/7 RNAi leaves exhibited deeper coloration than wild type, indicating higher ROS accumulation and accelerated programmed cell death occurring in zmsut1/7 RNAi leaves (Fig. 5I and fig. S21, D and E). The MDA contents were also higher across all regions of zmsut1/7 RNAi leaves than wild type (fig. S21F). Contrary to the decreased sugar accumulation in zmsut1/7 RNAi kernels, the zmsut1/7 RNAi leaves accumulated higher contents of sucrose and soluble sugars than wild type at 64 DAG (Fig. 5, J and K). The advanced leaf senescence phenotype in zmsut1/7 RNAi was further validated in Sichuan (2023) with reduced chlorophyll contents (fig. S21, G and H).

To further decipher the genetic relevance of SnRK2.6 and ZmSUT1, the zmsut1/7 RNAi; lps1-1 double mutant was generated. Phenotypic analysis revealed that the double mutant displayed severely advanced leaf senescence traits, largely resembling zmsut1/7 RNAi as assessed by chlorophyll content and photosynthetic efficiency (Fig. 5, L to O). The transcript levels of key genes associated with photosynthesis and senescence were detected, displaying comparably decreased and enhanced patterns in zmsut1/7 RNAi and zmsut1/7 RNAi; lps1-1 leaves at 80 DAG, respectively (fig. S23, A to N).

Overexpression of ZmSUT1 enhances kernel weight and delays leaf senescence

To further determine the genetic significance of ZmSUT1 in kernel yield improvement, we specifically overexpressed ZmSUT1 at the BETL region using the BETL9 promoter. Two independent lines were recovered and used for subsequent analysis (ZmSUT1-OE1 and ZmSUT1-OE2). The reverse transcription quantitative polymerase chain reaction (RT-qPCR) analysis detected higher transcript levels of ZmSUT1 in the BETL region of ZmSUT1-OE at 18 DAP than wild type (fig. S24A). A distinct band was detected in the BETL region of ZmSUT1-OE kernels using the Flag antibody, but not in wild type (fig. S24B).

The ZmSUT1-OE kernel area was increased, resulting in elevated 100-kernel weight along with greater extent of cell wall ingrowth in the BETL region (Fig. 6, A and B, and fig. S24, C to E). The levels of soluble sugars, sucrose, and starch in ZmSUT1-OE kernels were consistently increased compared to wild type (Fig. 6, C to E). Although the zein content was significantly elevated in ZmSUT1-OE kernels, the nonzein contents were markedly decreased, leading to similar total protein accumulation between ZmSUT1-OE and wild type (Fig. 6F and fig. S24, F to H).

Fig. 6. — (A) Ear phenotypes of the *ZmSUT1-OE* lines. Scale bar, 1 cm. (B to E). 100-kernel weight (B), soluble sugar (C), sucrose (D), and starch (E) levels of the *ZmSUT1-OE* mature kernels. (F) SDS-PAGE analysis of the zein and nonzein contents. γ27, 27-kDa γ-zein; α22, 22-kDa α-zein; α19, 19-kDa α-zein; γ16, 16-kDa γ-zein; β15, 15-kDa β-zein; δ10, 10-kDa δ-zein. (G) Plant and ear leaf phenotypes of the *ZmSUT1-OE* lines at 84 DAG. Scale bar, 1 cm. (H to K) Photosynthetic efficiency (H), chlorophyll (I), soluble sugar (J), and sucrose levels (K) of the *ZmSUT1-OE* leaves at 84 DAG. **P < 0.01 and ***P < 0.001.

At 50 DAG, both wild-type and ZmSUT1-OE plants exhibited green leaves with similar chlorophyll contents (fig. S25, A and B). By 74 DAG, the ZmSUT1-OE leaves detected significantly higher chlorophyll levels than wild type (fig. S25, C and D). At 84 DAG, the wild-type plants became withered and senescent. In contrast, the ZmSUT1-OE leaves remained green, detecting relatively higher photosynthetic efficiency and chlorophyll contents (Fig. 6, G to I). Elevated ZmSUT1 transcripts and protein accumulation were detected in ZmSUT1-OE leaves at 84 DAG, indicating a potential feedback response (fig. S25, E and F). In addition, soluble sugar and sucrose levels were lower in ZmSUT1-OE leaves than wild type (Fig. 6, J and K). By 94 DAG, the wild-type leaves were nearly completely senescent, while the ZmSUT1-OE leaves were just beginning to senesce and contained higher chlorophyll levels than wild type (fig. S25, G and H).

SnRK2.6 phosphorylates ZmSUT1 at the residue S516

In general, kinases interact with substrates to catalyze phosphorylation, regulating various physiological and developmental processes (33–36). Considering the interaction between SnRK2.6 and ZmSUT1, we subsequently investigated potential phosphorylation of ZmSUT1 mediated by SnRK2.6. The coexpression of ZmSUT1-Flag and SnRK2.6-HA in N. benthamiana leaves revealed a more intense phosphorylation signal of ZmSUT1 compared to the control groups (Fig. 7A). Stronger ZmSUT1 phosphorylation intensity was also observed when ZmSUT1-Flag was incubated with SnRK2.6-GFP (Fig. 7B). Furthermore, leaf protoplast assay demonstrated that SnRK2.6 enhanced the phosphorylation level of ZmSUT1 (Fig. 7C). Conversely, the phosphorylation level was substantially decreased in lps1-1 (Fig. 7C). However, the cotransformation of SnRK2.6 and ZmSUT1 in lps1-1 increased the phosphorylation pattern of ZmSUT1 (Fig. 7C). We also performed in vitro kinase assay to test the phosphorylation of ZmSUT1 mediated by SnRK2.6. Immunoblot analysis revealed that the recombinant SnRK2.6-MBP proteins detected strong autophosphorylation activity by the anti-phosSer/Thr antibody (fig. S26). Previous works reveal that mutations in the two conserved residues in SnRK2, D160 and S175, abolish the SnRK2 activity (37–39). Phosphorylation-deficient substitutions of the conserved sites D160 and S175 (SnRK2.6^Mu) completely abolished the phosphorylation signal (fig. S26). A retarded ZmSUT1 band was observed when the mixture of recombinant ZmSUT1 and SnRK2.6 proteins was analyzed using the PhosTag gel (Fig. 7D). In contrast, the retarded band was absent when ZmSUT1 was incubated with SnRK2.6^Mu.

Fig. 7. — (A and B) SnRK2.6-HA (A) and SnRK2.6-GFP (B) elevated ZmSUT1-Flag phosphorylation levels. Total proteins from the *N. benthamiana* leaves were immunoprecipitated using the Flag agarose beads. (C) Phosphorylation level detection of ZmSUT1 in wild-type and *lps1-1* leaf protoplast cells. (D) In vitro kinase assay detecting the ZmSUT1 phosphorylation mediated by SnRK2.6. (E) In vivo kinase assay detecting ZmSUT1 phosphorylation mediated by SnRK2.6 in the *N. benthamiana* leaves. (F) In vitro kinase assay detecting SnRK2.6-mediated ZmSUT1 phosphorylation at S516. (G) Immunoblot assay detecting the elevated ZmSUT1 protein levels mediated by SnRK2.6 in the leaf protoplast cells. (H to J) Cell-free degradation assay showing the protein degradation rate of the three ZmSUT1 versions mediated by SnRK2.6. (K) LCI assay showing the elevated ZmSUT1 homodimer-forming intensity mediated by SnRK2.6. (L) Immunoblot assay detecting the elevated ZmSUT1 homodimer-forming signal mediated by phosphorylation. The protein levels were quantified using the ImageJ software.

Next, we identified the specific amino acids in ZmSUT1 phosphorylated by SnRK2.6, which recognizes the conserved phosphorylation motif (R××S) in their substrates (fig. S17C). ZmSUT1 has 12 membrane-spanning regions, with a 31–amino acid N terminus and a 15–amino acid C terminus located on the cytoplasmic side (fig. S27A). We analyzed the ZmSUT1 protein sequence and identified that only the residue serine-516 (S516) in the C-terminal region was predicted as a potential phosphorylation site that could be recognized by SnRK2.6 (fig. S27, B and C). LC-MS analysis further identified S516 as the phosphorylation site of ZmSUT1 after incubation with SnRK2.6 in maize protoplast cells (fig. S27D). Furthermore, S516 was substituted into alanine to mimic the phosphorylation-deficient pattern (ZmSUT1^A). Immunoblot detection coupled with immunoprecipitation in N. benthamiana leaves showed that ZmSUT1^A exhibited markedly weaker phosphorylation intensity than ZmSUT1 (Fig. 7E). The ZmSUT1^A phosphorylation intensity was also slightly altered when incubation with SnRK2.6, while SnRK2.6 notably elevated the ZmSUT1 phosphorylation level. Moreover, in vitro kinase assay showed a markedly decreased intensity of the retarded phosphorylation band for ZmSUT1^A induced by SnRK2.6 in comparison with ZmSUT1 (Fig. 7F). Collectively, these data indicate that S516 is a major site in ZmSUT1 phosphorylated by SnRK2.6, but likely not the only relevant one.

ZmSUT1 phosphorylation enhances its protein stability, sucrose transport activity, and homodimer-forming capacity

To illustrate the biological relevance of ZmSUT1 phosphorylation, we first examined whether phosphorylation influenced the interaction between SnRK2.6 and ZmSUT1. Co-immunoprecipitation assay revealed that ZmSUT1, ZmSUT1^D (phosphorylation-mimic form), and ZmSUT1^A (phosphorylation-deficient form) consistently interacted with SnRK2.6, indicating that phosphorylation did not affect the interaction between SnRK2.6 and ZmSUT1 (fig. S28, A to C). The effect of ZmSUT1 phosphorylation was further characterized using heterologous expression in the sucrose-uptake deficient yeast strain. Growth curve estimation revealed that yeast cells expressing ZmSUT1^D displayed faster growth at the 2% sucrose growth condition than ZmSUT1 and ZmSUT1^A (fig. S28, D and E). Sucrose uptake efficiency of the three complemented yeast cells was investigated using the sucrose-analog esculin and live-cell microscopy. Notably, esculin uptake for the yeast cells expressing ZmSUT1^D was stronger than ZmSUT1 and ZmSUT1^A, whereas yeast cells expressing ZmSUT1^A were the lowest (fig. S28, F and G). The phosphorylation effect on the kinetics of ZmSUT1-mediated transport was further determined by measuring esculin uptake using the complemented yeast cells. ZmSUT1^D displayed the lowest K_m (Michaelis constant) than ZmSUT1 and ZmSUT1^A, while ZmSUT1^A was the highest (fig. S28H). In contrast, V_max was the highest in ZmSUT1^D and lowest in ZmSUT1^A (fig. S28I). In addition, sucrose feeding assay was performed to test the phosphorylation effect on ZmSUT1 activity. The sucrose uptake contents were higher in protoplasts overexpressing ZmSUT1^D than ZmSUT1 and ZmSUT1^A (fig. S28J). These data collectively support the notion that phosphorylation promotes ZmSUT1-mediated sucrose transport.

We then investigated whether SnRK2.6-mediated ZmSUT1 phosphorylation affected its protein stability. Protoplast transformation assay revealed that the ZmSUT1 protein level was notably increased when incubation with SnRK2.6-HA compared to the control (Fig. 7G). Cell-free degradation assays further demonstrated that SnRK2.6 markedly elevated the protein stability of ZmSUT1, displaying delayed protein degradation rate of ZmSUT1 in the presence of SnRK2.6 (Fig. 7H). The ZmSUT1^D protein stability was also increased upon incubation with SnRK2.6, whereas the protein degradation rate was accelerated when ZmSUT1^A was co-incubated with SnRK2.6 (Fig. 7, I to J). The elevated protein degradation rate for ZmSUT1^A was more evident when the three recombinant ZmSUT1 protein versions were separately incubated with the total protein extracts from the B73 leaves at 74 DAG with the addition of 1 mM adenosine 5′-triphosphate (ATP) (fig. S29A). Furthermore, higher ZmSUT1 proteins were detected in SnRK2.6-OE leaves than wild type (Fig. 3B and fig. S29B). Overall, these data demonstrate that ZmSUT1 phosphorylation at S516 enhances its protein accumulation, sucrose transport, and protein stability.

The sucrose transporter activity depends on the interaction with the cytoplasm membrane-localized proteins (40, 41). We wondered whether ZmSUT1 forms homodimers to perform it function. Both LCI and co-immunoprecipitation assays demonstrated that ZmSUT1 formed homodimers as indicated by visible luciferase signals and immunoprecipitated ZmSUT1-HA proteins, respectively (fig. S30, A and B). The BiFC assay further confirmed the interaction with the reconstructed yellow fluorescent protein (YFP) signal observed in the cytoplasmic membrane (fig. S30C). In addition, stronger luciferase signal was observed in the mixture of ZmSUT1-nLUC and ZmSUT1-cLUC with the addition of SnRK2.6 (Fig. 7K). Furthermore, with the addition of SnRK2.6-GFP, a higher amount of ZmSUT1-HA protein was immunoprecipitated by ZmSUT1-Flag in a dosage-dependent manner, indicating that SnRK2.6 enhanced the ZmSUT1 homodimer-forming capacity (fig. S30D). We further explored whether SnRK2.6-mediated phosphorylation affects the ZmSUT1 homodimer-forming activity. More intense YFP signals were observed when ZmSUT1-nYFP (n-terminal region of YFP) was co-infiltrated with ZmSUT1^D-cYFP (c-terminal region of YFP), whereas weaker YFP intensity was detected in the mixture of ZmSUT1-nYFP and ZmSUT1^A-cYFP (fig. S30E). In addition, a great amount of ZmSUT1^D-HA proteins and markedly reduced ZmSUT1^A-HA proteins were immunoprecipitated by ZmSUT1-Flag compared to ZmSUT1-HA (Fig. 7L). Collectively, these data indicate that SnRK2.6-mediated ZmSUT1 phosphorylation enhances the capacity of ZmSUT1 homodimer formation.

DISCUSSION

Leaf senescence is a highly coordinated process, extensively reported in Arabidopsis and rice, yet remains largely unknown in maize (42). It is an intricated program strictly controlled by multiple regulatory layers, including transcriptional, posttranscriptional, translational, and posttranslational aspects (43). Exploring the molecular mechanism underlying senescence is advanced through characterization of senescence-regulatory genes using forward and reverse genetic approaches (4, 44, 45). It is generally regarded as an important breeding strategy to delay the onset and progression of senescence, thereby enhancing crop yield and nutrient storage accumulation (4, 26). In this study, we screened an accelerated senescence mutant lps1-1, resulting from the disruption of a protein kinase-encoding gene, SnRK2.6. The lps1-1 mutants exhibited reduced photosynthetic efficiency and decreased kernel weight and nutrient reserve accumulation compared to wild type. Given that SnRK2.6 was globally expressed in kernels and leaves, the premature leaf senescence occurred in lps1 is contributed to disrupted roles in both source leaves and sink kernels. A positive correlation is widely accepted between photosynthetic capacity and grain yield (5, 6, 46). Precise manipulation of photosynthetic efficiency could serve as a beneficial strategy to enhance grain yield and nutrient storage accumulation. We found that the overexpression of SnRK2.6 exhibited increased photosynthetic efficiency, extended stay-green period, and enhancements in kernel weight and nutrient reserve content, resulting from the complicated regulatory roles of SnRK2.6 in both leaves and kernels. In Arabidopsis, AtSnRK2.6/OST1 is closet to SnRK2.8 in maize and OsSAPK8 in rice, playing crucial roles in stress response and stomatal movements (47, 48). However, it is unknown whether SnRK2.6 similarly responses to external biotic and abiotic stresses, a question that warrants further investigation. The function of SnRK2.6 appears to be conserved across grasses. The overexpression of OsSAPK6 in rice also led to noticeably delayed leaf senescence with higher chlorophyll contents, resulting in increased kernel dimension and nutrient storage accumulation. The squamosa promoter binding-like protein (SPL) transcription factor Ideal Plant Architecture1 (IPA1) was reported to shape ideal plant architecture and balance the trade-off between yield and disease resistance (49). OsSAPK6 phosphorylates IPA1 and stabilizes this protein, thereby enhancing chilling tolerance through the OsSAPK6-IPA1-OsCBF signaling pathway (50). OsSAPK6 positively regulates both chilling tolerance and stay-green process, providing a potential target for genetic improvement in rice. Previous research demonstrates that feedback of photosynthesis, endoplasmic reticulum stress, and ROS signaling initiates different senescence processes (51). Notably, sugar hyperaccumulation essentially influences senescence initiation and progression (23). We observed that sugar levels were elevated in lps1 leaves, concomitant with decreased photosynthetic efficiency and chlorophyll contents. We propose that the excessive sugar accumulation may act as a specific signal to trigger the rapidity of senescence.

Sucrose, the primary photoassimilate, is translocated through the BETL region of the endosperm, which serves as a gateway for nutrient transfer from maternal tissue into kernels (15). The lps1-1 kernels displayed reduced BETL region area, diminished kernel size, weight, and nutrient storage accumulation. Sugar loading is facilitated by symplasmic pathways in conjunction with concentration gradients and apoplasmic steps mediated by sucrose transporters (19, 52, 53). In maize, the first cloned sucrose transporter gene, ZmSUT1, participates in sucrose loading in source leaves (11, 20, 21). The zmsut1 null mutants exhibit dwarfed plant growth, precocious leaf senescence, and defective reproductive development (23). Recent work demonstrates that ZmSUT1 is highly expressed in the BETL region of the endosperm (11). However, the detailed molecular and genetic mechanisms of ZmSUT1 controlling kernel yield and nutrient reserve accumulation remain unclear. In this study, the RNAi approach was used to specifically knockdown the expression of ZmSUT1 and ZmSUT7 in the BETL region, which aimed to block sucrose transport from leaves to endosperm. As a result, significantly small kernels were observed in zmsut1/7 RNAi, while large kernels were observed in ZmSUT1 overexpression lines. Moreover, higher sugar levels were detected in zmsut1/7 RNAi leaves with comparatively lower concentration measured in their kernels. These results further support the role of ZmSUT1 in sucrose transport from leaves to kernels via the BETL region (11). In addition, we observed apparent syndromes of advanced leaf senescence in zmsut1/7 RNAi with reduced photosynthetic efficiency and chlorophyll contents, similar with the zmsut1 null mutant (31, 32). We propose that any disruption in this sucrose transport chain causes hyperaccumulated sucrose in leaves instead of being transported to kernels, resulting in early leaf senescence.

Protein phosphorylation plays pivotal roles in plant development and abiotic response by modulating protein activity, subcellular localization, and stability (34, 54). Protein phosphorylation also precisely regulates the timing of senescence process (45, 55). ZmSUT1 was identified to be a target of SnRK2.6, and the phosphorylation site S516 was identified to positively affect protein stability, sucrose transport, and homodimer-forming capacity of ZmSUT1. The BETL-specific expressed zmsut1/7 RNAi and ZmSUT1 overexpression lines not only exhibited marked alterations in kernel development but also were accompanied by accelerated leaf senescence and stay-green phenotypes, respectively. Given that SnRK2.6 and ZmSUT1 are coexpressed in the BETL region, we suggest that altered SnRK2.6 expression levels in zmsut1/7-RNAi leaves may trigger accelerated leaf senescence through activation of downstream senescence pathways, a mechanism that warrants further investigation. In addition, Krugel et al. (56) reported that StSUT1 from Solanum tuberosum forms homodimers in a redox-dependent manner. It is unknown about the relationship between redox and phosphorylation to orchestrate the dimerization state of ZmSUT1, which needs further exploration. Collectively, our findings demonstrate that SnRK2.6 phosphorylates ZmSUT1 in the BETL region to enhance sucrose transport from leaves to kernels, thereby promoting grain filling and nutrient storage. Concurrently, the elevated expression of SnRK2.6 and ZmSUT1 is associated with delayed leaf senescence, offering a promising strategy for developing new high-yield and high-quality maize varieties (Fig. 8).

Fig. 8. — In wild-type plants (Left), SnRK2.6 interacts with and phosphorylates ZmSUT1 in the BETL region, enhancing sucrose transport from leaves into kernels (Middle, top). This promotes starch and protein accumulation, leading to increased kernel weight and nutrient storage. Conversely, in the *snrk2.6*/*lps1* mutants (Right), ZmSUT1-mediated sucrose transport activity in the BETL region is compromised, resulting in reduced starch and protein levels in the kernel (Middle, bottom). This impaired sucrose flux is accompanied by accelerated leaf senescence, likely resulting from altered *SnRK2.6* expression and the consequent activation of downstream senescence pathways.

MATERIALS AND METHODS

Plant materials and growth conditions

The lps1-1 mutant was created by EMS-induced mutagenesis in the B73 background according to previously published protocol (57). To purge most irrelevant mutations, the lps1-1 mutant was recurrently backcrossed to B73 for six generations. The resulting heterozygous plants were self-pollinated to generate the homozygous lps1-1 kernels. The lps1-2 mutant was obtained from the public EMS mutant library database (http://maizeems.qlnu.edu.cn/) with the accession number EMS4-20024b.

The CRISPR-Cas9 constructs were designed using two guide RNA sequences “CGACAGGATCTGCAGCGCC” and “GGGAATTGGATTTCGCGTA.” The transgenic plants were backcrossed to the B104 inbred line twice to isolate the Cas9 constructs. The resulting heterozygous plants were self-pollinated to generate null mutants.

To create the SnRK2.6 overexpression lines, the SnRK2.6 coding sequence fused with the Flag tags and driven by the Ubiquitin promoter was inserted into the pCAMBIA3300 plasmid for transformation in the B104 inbred line.

The zmsut1/7 RNAi knockdown plants were generated in the KN5585 background using the conserved sequences of ZmSUT1 and its duplicated gene ZmSUT7 to create the silencing cassette. The RNA interference (RNAi) cassette is driven by the BETL-specific gene BETL9 promoter (11). To specifically overexpress ZmSUT1 in the BETL region, the coding sequence of ZmSUT1 was inserted into the pTF102 plasmid and driven by the BETL-specific gene BETL9 promoter. The resulting construct was transformed into the KN5585 inbred line (58). All the used primers were listed in table S4.

All the genetic materials were grown in the experimental fields in Sichuan (30.5°N, 103.6°E) and Sanya (18.2°N, 109.3°E), China. The mutant and overexpression plants were grown together with wild type in the same row to compare their corresponding agronomic traits. The N.benthamiana plants were grown with 22°C and 70% humidity under 16-hour light and 8-hour dark.

Genetic mapping

The lps1-1 and B73 plants were crossed, and the resulting F₁ generation was self-pollinated. The F₂ plants were phenotyped by observing the occurrence of advanced leaf senescence. Individual leaves were collected for DNA extraction. For each sample pool, a total of 96 samples exhibiting either the normal or lps1 phenotype were combined with equal proportions. Genomic DNA was prepared for the sequencing platform according to the standard next-generation sequencing (MGI sequencer) process. The MutMap method was used to identify the genomic intervals linked to the precocious senescence trait as described previously (27). The genomic DNA samples were further genotyped through PCR and sequencing using the corresponding primers. All the primers used in this study are listed in table S4.

Agronomic trait and biochemical analysis

The kernel weight was examined by a seed phenotyping system (Jielaimei, Sichuan, China). The longitudinal sections of the mature kernels were cut in the middle of the embryo along the vertical axe, and the transverse sections were cut above the embryo region and imaged on a Leica stereomicroscope.

For the paraffin observation, longitudinal sections of the harvested samples were cut, fixed using the 4% paraformaldehyde solution (m/v), and vacuumed for 30 min. After dehydration with gradient ethanol series, the samples were embedded in paraffin. These sections were cut into 8-μm slices and rehydrated with a series of ethanol gradients. Then, the sections were stained using 0.1% toluidine blue solution and photographed using the Leica microscope.

The chlorophyll contents were determined using the middle region of the ear leaves in maize and flag leaves in rice according to the method described previously (59). Briefly, 50 mg of fresh leaves was extracted with 1.5 ml of 95% ethanol for 1 day in dark. The supernatants were separated and measured at the absorbance of 470, 649, and 665 nm using a microplate reader (Varioskan LUX, Thermo Fisher Scientific). Photosynthetic efficiency was determined in the middle region of the ear leaf using a LI6400 Portable Photosynthesis System with a red/blue light-emitting diode light source (6400-02B; Li-Cor). The photosynthesis measurement details were performed on sunny day from 9:00 a.m. to 11:30 a.m. in the field with the following settings: 400–parts per million reference CO₂ and a flow rate of 500 μmol s⁻¹.

The anthocyanin and flavonoid contents were determined as previously described (60). A total of 50 mg of fresh samples was used to extract anthocyanin by adding 0.5 ml of extraction buffer (methanol with 1% HCl). The extracted samples were incubated overnight at 4°C at dark condition. The extracts were centrifuged twice and measured at an absorbance of 530 and 657 nm for anthocyanin contents using the formula (A₅₃₀-0.25 × A₆₅₇/fresh weight). The flavonoid content was quantified with the standards and evaluated at an absorbance of 350 nm.

The soluble sugar levels were determined according to previously published protocol (61). Briefly, per kernel was weighted and extracted using 80% ethanol for 2 hours and then repeated twice. The extracted supernatants were further used for the anthranone-mediated reaction process. The soluble sugar levels were measured at an absorbance of 620 nm. The sucrose and starch levels were measured using the Plant Sucrose Content Assay Kit (BC2465, Solarbio) and Starch Content Assay Kit (BC0705, Solarbio), respectively. The detailed experiment process was performed according to the manufacturer’s instructions.

For the iodine staining assays, the harvested samples were cut in the middle of the embryo and then used for staining using Lugo iodine solution (L769703, MACKLIN) for 10 min. Then, the samples were destained with water for several times before imaging using the Leica stereomicroscope.

The DAB staining assay was performed as previously described (24). Briefly, the harvested samples were soaked in 0.1% DAB solution (36201ES03, Yeasen). The samples were vacuum infiltrated for 30 min, stained in the dark overnight, and decolorized for 3 days using 95% ethanol before observation. For the trypan blue staining assay, the harvested leaves were submerged in the Trypan blue solution [25% lactic acid (m/v), 23% water-saturated phenol (v/v), 25% glycerol (v/v), and 0.25% Trypan blue (m/v)] and boiled for 30 min. After cooling, the samples were further vacuum infiltrated for 30 min and incubated overnight in the dark. Then, the stained samples were decolorized for 3 days using 95% ethanol before imaging using the Leica stereomicroscope.

The MDA contents were measured using the MDA Content Assay Kit (BC0025, Solarbio) and performed according to the manufacturer’s instruction. The extracted samples were measured at an absorbance of 532 and 600 nm with the microreader (Varioskan LUX, Thermo Fisher Scientific).

The zein and nonzein components were determined according to previously described (62). Briefly, per kernel for each independent samples were weighed and added with zein extraction buffer [70% ethanol, 2% 2-mercaptoethanol (v/v), 3.75 mM sodium borate (pH 10.0), and 0.3% SDS] and then incubated overnight. After extracting zein proteins, the remained sediments were further used for nonzein extraction by adding nonzein extraction buffer (12.5 mM sodium borate, 5% SDS, and 2% 2-mercaptoethanol) and then kept on the bench for additional 2 hours. The diluted zein and nonzein proteins were individually detected on a 15% SDS-PAGE gel and stained with Coomassie brilliant blue. The Protein Assay Kit (P0006C, Beyotime) was used to quantify the protein contents.

Subcellular localization

The CDS fragments of SnRK2.6 were inserted into the pCAMBIA1300-35S-eGFP plasmid. The construct was further infiltrated and transiently expressed in the N. benthamiana leaf cells. After 48-hour incubation, the GFP fluorescence signal was observed using a Leica confocal microscope.

ZmSUT1/7 antibody production and immunoblot analysis

A partial sequence of the ZmSUT1 protein, spanning from 164 to 291 amino acids was used to immunize rabbits. For immunoblot analysis, the resulting antibodies were diluted at a ratio of 1:2000. The total proteins were extracted using the native lysis buffer [50 mM tris-MES (pH 8.0), 1 mM MgCl₂, 0.5 mM sucrose, 10 mM EDTA, 2 mM dithiothreitol (DTT), 1 mM PMSF, cocktail, and 1% NP-40], incubated at 98°C for 10 min, followed by adding 4× SDS loading buffer, and detected with the anti-ZmSUT1/7 antibody. All assessments of protein levels via immunoblot analysis were performed at least three times with independent biological replicates.

Phosphoproteomic analysis

The phosphoproteomic analysis was performed with three independent biological replicates as previously described (63). Briefly, the harvested leaves were ground into fine powder and extracted using the homogenization buffer (50 mM Hepes, 250 mM sucrose, 10 mM EDTA, 5% glycerol, 3 mM DTT, 0.5% insoluble polyvinylpyrrolidone, cocktail, 1 mM PMSF, protein phosphatase inhibitors, 25 mM sodium fluoride, 1 mM sodium molybdate, and 50 mM sodium pyrophosphate decahydrate). The extracted protein concentration was determined using the Pierce Detergent Compatible Bradford Assay (23246, Thermo Fisher Scientific). A total of 20 mg of proteins per sample was used for enzyme digestion. Subsequently, 5 mM DTT was added and incubated with 50°C for 1 hour, and 50 mM indole-3-acetic acid was added for additional 1 hour. After ammonium bicarbonate treatment, trypsin (V5111, Promega) with a ratio of 1:50 was added, and proteins were digested overnight at 37°C. The digested peptides were enriched using the High-Selected TiO₂ Phosphopeptide Enrichment Kit (A32993, Thermo Fisher Scientific) according to the provided instrument. The enriched peptides were taken for specific library construction by DIA, and the rest of the samples were used for Data-dependent acquisition (DDA) proteomic analysis. These samples were analyzed using the timsTOF Pro2 protein Mass Spectrometer (Bruker). These recovered data were processed and analyzed using Spectronaut X with default parameters. The rmotifx package in R was employed for the enriched motif analysis with a statistical significance (P < 1 × 10⁻⁶) (64).

RT-qPCR and RNA in situ hybridization assays

Total RNA was extracted using TRIzol regent (Invitrogen). A total of one μg RNA was used for reverse-transcription into cDNA using HiScript RT SuperMix (R423-01, Vazyme). The subsequent RT-qPCR reactions were performed with at least three biological replicates using the Hieff qPCR SYBR Green Master Mix (11145ES, Yeasen).

The developing leaves at 60 DAG and kernels at 20 DAP were used for RNA in situ hybridization. The amplified SnRK2.6 fragments were inserted into the pSPT18 plasmid. The RNA probes were individually generated using the SP6 and T7 RNA polymerases (Roche). The detailed procedures with 8-μM sections were performed as described previously (65, 66).

Protein-protein interaction assays

For in-vitro pull down assay, a total of five μg of the purified SnRK2.6-MBP and one μg of ZmSUT1-Halo-Tag proteins were incubated in the pull-down buffer [40 mM KCl, 1 mM EDTA, 25 mM HEPES (pH 7.5)] for 4 hours at 4°C. Then, 50 μl of MBP agarose beads (SA077005, Smart Lifesciences Biotech) were added and further incubated for another 2 hours. The reaction mixture was washed with pull-down buffer for five times with 5 min for each time. Subsequently, the proteins were boiled with 50 μl of 2× SDS loading buffer and detected using the anti-MBP (M20051, Abmart) and anti-Halo-Tag (G9281, Promega) antibodies, respectively.

For co-immunoprecipitation assay, the ZmSUT1-Flag and SnRK2.6-HA or SnRK2.6-GFP plasmid constructs were co-infiltrated in N. benthamiana leaves. After growing for 72 hours, the infiltrated leaves were harvested, and total proteins were extracted using the native lysis buffer for 1 hour on ice. Then, the homogenous mixture was incubated with Flag agarose beads (20018, Abmart) with rotation at 4°C for 5 hours. After washing the beads with native lysis buffer for five times, the immunoprecipitates were detected using the anti-Flag (F1804, Sigma-Aldrich), anti-HA (H6533, Sigma-Aldrich), and anti-GFP (M20004, Abmart) antibodies, respectively. To detect the ZmSUT1 homodimer-forming capacity, the ZmSUT1-Flag construct was coexpressed with ZmSUT1-HA, ZmSUT1^D-HA, and ZmSUT1^A-HA in leaf protoplast cells, respectively. The immunoprecipitated mixtures were detected using anti-Flag (F1804, Sigma-Aldrich) and anti-HA (H6533, Sigma-Aldrich) antibodies, respectively.

For semi in vitro pull-down assay, the ZmSUT1-Flag proteins were immunoprecipitated from the maize protoplast cells and then eluted from the Flag agarose beads using the Flag peptide (F3290, Sigma-Aldrich). The SnRK2.6-MBP proteins were incubated with the eluted ZmSUT1-Flag protein for 2 hours. The protein mixtures were separated on the 10% (v/v) SDS-PAGE gel and detected using the anti-MBP (M20051, Abmart) and anti-Flag (F1804, Sigma-Aldrich) antibodies, respectively.

For BiFC assays, the construct combinations were co-infiltrated into N. benthamiana leaves. After incubation for 72 hours, the fluorescent signal was detected using the Leica confocal microscope.

For LCI assays, the CDS fragments of SnRK2.6 and native ZmSUT1 were individually cloned into the pCAMBIA1300-35S-cLUC and pCAMBIA1300-35S-nLUC. The constructs were co-infiltrated into N. benthamiana leaves. After growing for 72 hours, 0.5 mM luciferin was infiltrated into the leaves and then imaged using the Multi-functional Imaging system (CheniDoc, Bio-Rad).

Yeast complementation and sucrose uptake assays

The coding sequences of the three ZmSUT1 variants were inserted into the plasmid pDR196. The resulting constructs were transformed into the sucrose uptake-deficient yeast strain SUSY7/ura3. The transformed yeast was grown in the synthetic-defined and uracil-deficient (SD/-ura) media supplemented with 2% glucose and 2% sucrose for 4 days at 30°C. The cell number was calculated using previously published calibration curve through yeast strain intensity determination on a spectrofluorometer with an absorbance of 600 nm at the indicated times (67).

The esculin uptake assay was performed as described previously with some modifications (68). Briefly, the complemented yeast cells were centrifuged at 2000 rpm for 5 min. After the medium was removed, 1 mM esculin in phosphate buffer [25 mM Na₂HPO₄ (pH 4.0)] was added to each sample and then incubated at 30°C for 2 hours with shaking. Then, the cells were centrifuged at 2000 rpm for 5 min and washed with the phosphate buffer for five times for confocal image observation by the Leica confocal microscope.

For the kinetic analysis, these complemented yeast strains were grown in liquid SD/-Ura medium containing glucose with an optical density at 600 nm (OD₆₀₀) of 0.6. The yeast cells were centrifugated at 2000 rpm for 5 min, washed with 25 mM sodium phosphate buffer (pH 5.0), and resuspended to a final concentration to OD₆₀₀ of 20.0. Subsequently, 2 mM esculin was added and incubated at 30°C, and the samples were harvested to determine the fluorescence intensity on a spectrofluorometer with excitation at 367 nm and emission at 454 nm at the indicated times.

For the sucrose uptake assay in protoplast cells, the extracted protoplast cells were individually transfected with 10 μg of the indicated plasmids for 30 min. After transfection, these protoplasts were incubated overnight in the incubation solution (20 mM MES, 20 mM KCl, 0.2 CaCl₂, and 0.2 mM mannitol) at room temperature. In the following day, the protoplasts were centrifuged at 100g for 10 min, resuspended in the incubation solution, and incubated with 10 mM sucrose. The incubated protoplast cells were harvested at the indicated times, centrifugated at 100g for 5 min, washed with the incubation buffer for five times, frozen in nitrogen, and used for the sucrose level quantification. All the harvested samples were used for sucrose level determination using the LC-MS analysis as previously described (33).

In vivo and in vitro phosphorylation assays

For in vivo phosphorylation assay, ZmSUT1-Flag and SnRK2.6-HA or SnRK2.6-GFP were cotransferred into in N. benthamiana leaves. The protoplast cells from the B73 inbred line or lps1-1 mutant leaves were also extracted. The constructed plasmids were transformed into the maize mesophyll protoplasts using the polyethylene glycol-CaCl₂ method. After incubation for 20 hours, protoplast cells were centrifuged at 500g for 10 min. The total proteins were further extracted using the native lysis buffer for 1 hour on ice. Then, the homogenous samples were incubated with Flag agarose beads (20018, Abmart) with rotation at 4°C for 5 hours. After washing the beads with native lysis buffer for five times, the immunoprecipitated mixtures were detected using the anti-Flag (F1804, Sigma-Aldrich), anti-HA (H6533, Sigma-Aldrich), anti-GFP (M20004, Abmart), and anti–phos-Ser/Thr (9631S, Cell Signaling Technology) antibodies, respectively. To confirm the phosphorylation site, ZmSUT1-Flag or ZmSUT1^A-Flag was co-infiltrated with SnRK2.6-HA in N. benthamiana leaves, respectively. The immunoprecipitates were further detected using the anti-Flag (F1804, Sigma-Aldrich), anti-HA (H6533, Sigma-Aldrich), and anti–phos-Ser/Thr (9631S, Cell Signaling Technology) antibodies, respectively.

For in vitro kinase assays, the recombinant ZmSUT1-Halo-Tag and ZmSUT1^A-Halo-Tag proteins were individually incubated with SnRK2.6-MBP or SnRK2.6^Mu-MBP proteins in the kinase reaction buffer [50 mM tris-HCl (pH 7.5), 10 mM MgCl₂, 1 mM DTT, and 2 mM ATP] at 30°C for 1 hour. The reaction mixture was terminated by adding 4× SDS loading buffer and loaded on the Phos-Tag and SDS-PAGE gels, respectively. Then, the protein bands were detected using the anti-Halo-Tag and anti-MBP antibodies, respectively.

Cell-free degradation assay

For the in vitro cell-free degradation assay, the CDS regions of ZmSUT1, ZmSUT1^D, and ZmSUT1^A were individually cloned into the pFN19K-HaloTag plasmid, and the recombinant proteins were expressed using the TNT SP6 High-Yield Wheat Germ Protein Expression System (L3260, Promega). The recombinant proteins were incubated with a total of 100 μg of total protein extracts from the leaf samples in the B73 inbred line and incubated at the indicated time points. Then, all the incubated proteins were boiled at 95°C for 10 min and detected using the anti-Halo-Tag (G9281, Promega) and anti-ACTIN (M20009, Abmart) antibodies, respectively.

For in vivo cell free degradation assays, ZmSUT1, ZmSUT1^D-Flag, and ZmSUT1^A-Flag were separately cotransformed with SnRK2.6-HA in the leaf protoplast cells. After incubation for 20 hours at room temperature, the proteins were extracted using the cell-free lysis buffer [25 mM tris-HCl (pH 7.5), 10 mM NaCl, 4 mM PMSF, 5 mM DTT, 10 mM MgCl₂, and 1 mM ATP] for 1 hour on ice. The lysates were incubated at room temperature and sampled at the indicated time points. The immunoblot analysis was performed and detected using the anti-Flag (F1804, Sigma-Aldrich), anti-HA (H6533, Sigma-Aldrich), and anti-ACTIN (M20009, Abmart) antibodies, respectively.

Statistical analysis

The raw data and detailed statistical analysis were tested in table S5. Protein quantification analysis was performed by tracing out the individual band using the ImageJ software (https://imagej.net/ij/). Significant differences between two groups were evaluated based on the two-sided Student’s t test analysis on Microsoft Excel (2022).

Accession numbers

Gene sequences from this article can be found in the MaizeGDB database under accession numbers: ZmSUT1, Zm00001d027854; SnRK2.6, Zm00001d050723; 15-kD β-zein, Zm00001d035760; 16-kD γ-zein, Zm00001d005793; 22-kD α-zein, Zm00001d048809; 27-kD γ-zein, Zm00001d020592; 50-kD γ-zein, Zm00001d020591; CASP, Zm00001d016363; Cys1, Zm00001d002065; Cys2, Zm00001d020636; Peroxiredoxin, Zm00001d046682; Ccp14, Zm00001d005391; Chl, Zm00001d015385; ZmMDH, Zm00001d031899; ZmPEPC, Zm00001d046170; ZmNADPME, Zm00001d037961; and ZmRBSC, Zm00001d051485.

Acknowledgments

We thank J. Li and H. Yu from Yazhou Bay Seed Laboratory, China for sharing the OsSAPK6 overexpression lines, X. Chen from Sichuan Agricultural University for assistance with rice related assays, H. Qing from Sichuan Agricultural University for rice planting, Y. Feng from Sichuan Agricultural University for confocal image, and Y. Gao from the CAS center for Excellence in CEMPS for sucrose level identification.

Funding:

This research was supported by the National Natural Science Foundation of China (32472122 to T.Y. and 31925030 to Y.W.), the Young Scientist Program of National Key Research and Development Program of China (2023YFD1201800), the STI 2030-Major Project (2023ZD0406901), the Sichuan Provincial General Project (2024NSFSC0315), and the Open Project Program and Biological Breeding Program of State Key Laboratory of Crop Gene Exploration and Utilization in Southwest China (SKL-ZY202211).

Author contributions:

Conceptualization: T.Y., Y.H., and Y.W. Methodology: T.Y., Yunqin.Huang., Z.W.., Z.Long, H.Zhu, Z.Liang, S.W., L.T., R.L., H.Zhang., Y.H., and X. Li. Investigation: T.Y., Yongcai.Huang., and Y.W., and X. Lu. Visualization: T.Y. and Y.H. Funding acquisition: T.Y. and Y.W. Resource, data curation, validation, supervision, formal analysis, project administration, writing–original draft, writing–review and editing: T.Y., Y.H., and Y.W.

Competing interests:

The authors declare that they have no competing interests.

Data, code and materials availability:

All data and code needed to evaluate and reproduce the results in the paper are present in the paper and/or the Supplementary Materials. The raw data for the phosphoproteome in this paper has been deposited at iProX database with the accession number PXD071793. The lps1 materials can be provided by X. Lu pending scientific review and a completed material transfer agreement with X. Lu. Requests for lps1 should be submitted to X. Lu. at lu.xiaoduo@163.com. This study did not generate new materials.

Supplementary Materials

The PDF file includes:

Figs. S1 to S30

Legends for tables S1 to S5

sciadv.aeb2472_sm.pdf^{(10.3MB, pdf)}

Other Supplementary Material for this manuscript includes the following:

Tables S1 to S5

sciadv.aeb2472_tables_s1_to_s5.zip^{(1.2MB, zip)}

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Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Figs. S1 to S30

Legends for tables S1 to S5

sciadv.aeb2472_sm.pdf^{(10.3MB, pdf)}

Tables S1 to S5

sciadv.aeb2472_tables_s1_to_s5.zip^{(1.2MB, zip)}

Data Availability Statement

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PERMALINK

SnRK2.6 phosphorylates sucrose transporter ZmSUT1 to enhance yield by modulating leaf senescence in maize

Tao Yang

Yunqin Huang

Zan Wu

Zhenglan Long

Hongwei Zhu

Shanshan Wang

Lei Tang

Ruirui Li

Zhengchen Liang

Haoyu Zhang

Yongcai Huang

Xiaoling Li

Xiaoduo Lu

Yongrui Wu

Roles

Abstract

INTRODUCTION

RESULTS

The lps1-1 mutant exhibits precocious leaf senescence, decreased kernel weight and storage reserve content

Fig. 1. Plant and kernel phenotypes of the lps1-1 mutant.

Lps1-1 encodes a protein kinase SnRK2.6

Fig. 2. Gene cloning and genetic verification of lps1–1.

Overexpression of SnRK2.6 delays leaf senescence with enhanced kernel weight and storage reserve synthesis

Fig. 3. Phenotypic and biochemical analyses of SnRK2.6 and OsSAPK6 overexpression lines.

SnRK2.6 interacts with the sucrose transporter ZmSUT1

Fig. 4. Protein-protein interaction assays of SnRK2.6 and ZmSUT1.

ZmSUT1 mutation results in decreased kernel weight and accelerated leaf senescence

Fig. 5. Kernel and plant phenotypes of the zmsut1/7 RNAi lines.

Overexpression of ZmSUT1 enhances kernel weight and delays leaf senescence

Fig. 6. Kernel and plant phenotypes of the ZmSUT1-OE lines.

SnRK2.6 phosphorylates ZmSUT1 at the residue S516

Fig. 7. SnRK2.6 phosphorylates ZmSUT1 to enhance its protein stability and homodimer-forming capacity.

ZmSUT1 phosphorylation enhances its protein stability, sucrose transport activity, and homodimer-forming capacity

DISCUSSION

Fig. 8. A proposed model of the SnRK2.6-ZmSUT1 module in regulating grain filling and leaf senescence.

MATERIALS AND METHODS

Plant materials and growth conditions

Genetic mapping

Agronomic trait and biochemical analysis

Subcellular localization

ZmSUT1/7 antibody production and immunoblot analysis

Phosphoproteomic analysis

RT-qPCR and RNA in situ hybridization assays

Protein-protein interaction assays

Yeast complementation and sucrose uptake assays

In vivo and in vitro phosphorylation assays

Cell-free degradation assay

Statistical analysis

Accession numbers

Acknowledgments

Funding:

Author contributions:

Competing interests:

Data, code and materials availability:

Supplementary Materials

The PDF file includes:

Other Supplementary Material for this manuscript includes the following:

REFERENCES

Associated Data

Supplementary Materials

Data Availability Statement

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