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. 2024 Sep 24;67(2):294–310. doi: 10.1111/jipb.13778

A novel C2H2‐type zinc‐finger transcription factor, CitZAT4, regulates ethylene‐induced orange coloration in Satsuma mandarin flavedo (Citrus unshiu Marc.)

Quan Sun 1,2, Zhengchen He 1, Junli Ye 1, Ranran Wei 1, Di Feng 1, Yingzi Zhang 1, Lijun Chai 1, Yunjiang Cheng 1, Qiang Xu 1, Xiuxin Deng 1,3,✉
PMCID: PMC11814913  PMID: 39315817

ABSTRACT

Ethylene treatment promotes orange coloration in the flavedo of Satsuma mandarin (Citrus unshiu Marc.) fruit, but the corresponding regulatory mechanism is still largely unknown. In this study, we identified a C2H2‐type zinc‐finger transcription factor, CitZAT4, the expression of which was markedly induced by ethylene. CitZAT4 directly binds to the CitPSY promoter and activates its expression, thereby promoting carotenoid biosynthesis. Transient expression in Satsuma mandarin fruit and stable transformation of citrus calli showed that overexpressing of CitZAT4 inhibited CitLCYE expression, thus inhibiting α‐branch yellow carotenoid (lutein) biosynthesis. CitZAT4 overexpression also enhanced the transcript levels of CitLCYB, CitHYD, and CitNCED2, promoting β‐branch orange carotenoid accumulation. Molecular biochemical assays, including yeast one‐hybrid (Y1H), electrophoretic mobility shift (EMSA), chromatin immunoprecipitation quantitative polymerase chain reaction (ChIP‐qPCR), and luciferase (LUC) assays, demonstrated that CitZAT4 directly binds to the promoters of its target genes and regulates their expression. An ethylene response factor, CitERF061, which is induced by ethylene signaling, was found to directly bound to the CitZAT4 promoter and induced its expression, thus positively regulating CitZAT4‐mediated orange coloration in citrus fruit. Together, our findings reveal that a CitZAT4‐mediated transcriptional cascade is driven by ethylene via CitERF061, linking ethylene signaling to carotenoid metabolism in promoting orange coloration in the flavedo of Satsuma mandarin fruit. The molecular regulatory mechanism revealed here represents a significant step toward developing strategies for improving the quality and economic efficiency of citrus crops.

Keywords: α‐branch carotenoid, β‐branch carotenoid, CitZAT4, ethylene, orange coloration, Satsuma mandarin

The ethylene‐induced C2H2‐type zinc‐finger transcription factor CitZAT4 activates phytoene synthase expression to provide sufficient substrates for downstream carotenoid biosynthesis, CitZAT4 inhibits α‐branch carotenoid biosynthesis, promotes β‐branch carotenoid accumulation, and regulates carotenoid metabolic flow into the β‐branch, thus increasing the content of orange‐colored carotenoids in ethylene‐treated Satsuma mandarin flavedo.

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INTRODUCTION

Carotenoids are a subgroup of isoprenoids consisting of more than 750 members in nature (Yuan et al., 2015; Sun et al., 2018). In addition to providing vibrant colors to fruits and vegetables, carotenoids are important precursors of vitamin A and antioxidants when consumed by humans, helping to prevent chronic and degenerative diseases (Fraser and Bramley, 2004; Fiedor and Burda, 2014). The carotenoid biosynthesis pathway has been identified in many plant species, including citrus (Nisar et al., 2015; Yuan et al., 2015). The first is the condensation of two geranylgeranyl diphosphates (GGPPs) by phytoene synthase (PSY) to form phytoene, which is considered the major rate‐limiting step of this pathway. After a series of desaturation and isomerization reactions, the carotenoid metabolic pathway is divided into the α‐ and β‐branches. The α‐branch mainly consists of yellow lutein catalyzed by lycopene ε‐cyclase (LCYE), and the β‐branch consists of orange carotenoids (including β‐carotene, β‐cryptoxanthin, and violaxanthin) primarily catalyzed by lycopene β‐cyclase (LCYB), β‐carotene hydrolase (HYD), and 9‐cis‐epoxycarotenoid dioxygenases (NCEDs). Although most enzymes in the carotenoid biosynthetic pathway have been well characterized, little is known about the transcriptional regulatory mechanisms underlying carotenoid metabolism in citrus.

Transcription factors (TFs) are essential for the transcriptional regulation of the extensive variey of target genes by directly binding to their promoters. In recent years, several TFs have been reported to regulate carotenoid metabolism: for example, PIF1 and HY5, acting as a pair of antagonists, regulate carotenoid biosynthesis by binding to the PSY promoter in response to light and temperature cues (Gangappa and Botto, 2016; Llorente et al., 2016; Chenge‐Espinosa et al., 2018). In citrus, an ethylene response factor (ERF) TF CsERF061 and a basic helix‐loop‐helix (bHLH) TF CsTT8 positively regulate carotenoid accumulation by activating the expressions of carotenoid pathway genes (Zhu et al., 2021; Sun et al., 2023). CsMADS6 directly regulates the expression of CsLCYb1, thus inducing β‐branch orange carotenoid accumulation (Lu et al., 2018). CrMYB68 (an R2R3‐MYB TF) regulates the transformation of α‐branch and β‐branch carotenoids by negatively regulating the expression of CrBCH2 and CrNCED5 (Zhu et al., 2017). A recent study reported that the transcriptional regulatory cascade CsHB5‐CsBZIP44 synergistically regulates abscisic acid (ABA)‐mediated citrus fruit coloration by promoting carotenoid biosynthesis (Sun et al., 2024). These studies jointly indicate that transcriptional regulation is critical for carotenoid metabolism.

The zinc‐finger proteins comprise a large TF family in plants and are divided into subfamilies based on their secondary structures, such as Cys2/His2‐type (C2H2), C3H, C3HC4, C2HC5, C4HC3, C2HC, C4, C6, and C8 (Han et al., 2020a). Among these, C2H2‐type zinc‐finger proteins are the most important subfamily. C2H2‐type proteins can bind the cis‐elements known as A(G/C)T repeat sequences (Sakamoto et al., 2004; Kodaira et al., 2011), namely, the A[AG/CT]CNAC motif (Zhou et al., 2011; Han et al., 2020b), the TGCTANNATTG and TACAAT elements (Shi et al., 2014), and the CAGT and AGTGCACT binding motifs (Franco‐Zorrilla José et al., 2014), to regulate biological processes such as flowering and development (Zhou et al., 2011; Han et al., 2020b), the responses to biotic and abiotic stress (Sakamoto et al., 2004; Zhang et al., 2012; Shi et al., 2014), phytohormone signal transduction (Kodaira et al., 2011; Yang et al., 2021), and pigments metabolism (Asako et al., 2000; Wang et al., 2022b). Although C2H2‐type zinc‐finger proteins have been extensively reported to regulate biological processes in multiple species, their role in citrus carotenoid metabolism is poorly understood.

Citrus fruits contain abundant nutritional components, such as vitamin C, vitamin E, and provitamin A, as well as the largest number of carotenoids (about 115 types) (Kato et al., 2004; Rodriguez‐Concepcion et al., 2018). The carotenoid composition is responsible for the attractive color of citrus fruits, which is a crucial feature of organoleptic quality and thus market value, making the regulation of carotenoid metabolism a particularly important economic trait in citrus (Matsumoto et al., 2007; Fanciullino et al., 2008). Ethylene accelerates flavedo coloration by promoting carotenoid accumulation, and ethylene treatments are widely commercially used to improve the appearance of fruits for sale (Rodrigo and Zacarias, 2007; Matsumoto et al., 2009; Zhou et al., 2010; Sun et al., 2021). Satsuma mandarin (Citrus unshiu Marc.) is a major citrus cultivar consumed worldwide and is rich in various carotenoids (Kato et al., 2004; Matsumoto et al., 2007; Fanciullino et al., 2008). During fruit ripening and/or upon ethylene treatment, a change from α‐branch carotenoid (mainly lutein) accumulation to β‐branch carotenoid (mainly β‐carotene, violaxanthin, and particularly β‐cryptoxanthin) accumulation is observed in the flavedo of Satsuma mandarin (Matsumoto et al., 2009; Zhou et al., 2010; Ma et al., 2012). This is accompanied by a reduction in α‐branch carotenogenic (CitLCYE) transcripts and an increase in β‐branch carotenogenic transcripts (including CitLCYB, CitNCED2, and especially CitHYD), which are responsible for the ethylene‐induced orange coloration in the flavedo of Satsuma mandarin (Matsumoto et al., 2009; Zhou et al., 2010; Ma et al., 2012). However, the transcriptional regulatory mechanisms involved in the above‐mentioned processes have not yet been elucidated in Satsuma mandarin.

In this study, we identified a C2H2‐type zinc‐finger TF, CitZAT4, as part of the ethylene‐induced carotenoid biosynthesis pathway. This protein was found to form a transcriptional cascade with the ethylene‐induced upstream activator CitERF061. CitZAT4 plays an essential role in ethylene‐promoted orange coloration in the flavedo of Satsuma mandarin fruit by up‐regulating the expression of CitPSY, CitLCYB, CitHYD, and CitNCED2 and down‐regulating CitLCYE. CitERF061 positively regulates CitZAT4‐mediated orange coloration in citrus fruit by activating its expression, linking ethylene signaling and CitZAT4 activity. The molecular regulatory mechanism revealed here is of great significance for enhancing the quality and nutritional value of citrus fruits.

RESULTS

Ethylene promotes orange coloration in the flavedo of Satsuma mandarin fruit by regulating carotenoid metabolic flow

To explore the role of ethylene in fruit coloration, we performed two different treatments. As shown in Figure 1A, ethylene treatment accelerated fruit coloration, transitioning from yellow‐green to full orange at 10 d after treatment (DAT), whereas no obvious changes were observed in the control and 1‐methylcyclopropene (1‐MCP) + ethylene groups. We further measured the citrus color index (CCI) to quantify changes in flavedo color. The CCI was significantly higher in the ethylene treatment group than in the control and 1‐MCP + ethylene groups, especially at 5 DAT (Figure 1B). These results indicate that ethylene significantly promotes the coloration of the flavedo of Satsuma mandarin fruit.

Figure 1.

Figure 1

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Effects of ethylene on the flavedo color of Satsuma mandarin fruit

(A) Phenotype of treated fruit under multiple treatments. Bars = 2 cm. Citrus fruits treated with distilled water were used as control. Control, ethylene, and 1‐methylcyclopropene + ethylene groups shared the same group of citrus fruits at 0 d after treatment (DAT). (B) Citrus color index (CCI). Contents of total carotenoid (C), phytoene (D), β‐carotene (E), β‐cryptoxanthin (F), violaxanthin (G), and lutein (H) in flavedo of treated fruit. (I) Percentage of α‐ and β‐branch in total carotenoids under multiple treatments. The α‐branch includes lutein and the β‐branch includes β‐carotene, β‐cryptoxanthin, and violaxanthin. Data represent means ± SD of three biological replicates. Asterisks indicate statistically significant differences was determined by Student's t‐test (**P < 0.01; n.s., no significant difference).

As the carotenoid content and composition directly determine citrus flavedo color, we investigated the effect of ethylene treatment on carotenoid metabolism. High‐performance liquid chromatography (HPLC) revealed that the total carotenoid, phytoene, β‐carotene, β‐cryptoxanthin, and violaxanthin contents were higher in the ethylene treatment group than in the control and 1‐MCP + ethylene groups (Figure 1C–G), while the lutein levels exhibited the opposite trend (Figure 1H). Among the carotenoids, β‐carotene, β‐cryptoxanthin, and violaxanthin belong to the β‐branch of orange carotenoids, while lutein belongs to the α‐branch of yellow carotenoids. The percentage of carotenoids from the β‐branch gradually increased in all treatments, but it was higher under the ethylene treatment than in the control and 1‐MCP + ethylene groups (Figure 1I). Conversely, the percentage of α‐branch carotenoids exhibited the opposite trend (Figure 1I). These results indicate that ethylene treatment promotes orange coloration in the flavedo of Satsuma mandarin fruit by inducing orange β‐branch carotenoid accumulation and inhibiting yellow α‐branch carotenoid accumulation.

To elucidate the molecular mechanisms by which ethylene regulates carotenoid metabolism, we further examined the expression of the carotenogenic genes. Quantitative reverse‐transcription polymerase chain reaction (qRT‐PCR) analyses revealed that the ethylene treatment significantly induced the expression of CitPSY, CitLCYB, CitHYD, and CitNCED2 compared with the control and 1‐MCP + ethylene treatment (Figure 2A–D). By contrast, the transcript level of CitLCYE was significantly lower under the ethylene treatment than under the control and 1‐MCP + ethylene treatment (Figure 2E). Based on the above results, we plotted the carotenoid metabolic pathway to examine the effect of ethylene treatment on carotenoid metabolism in the Satsuma mandarin flavedo. As shown in Figure 2F, ethylene treatment induced CitPSY expression to increase the phytoene content, thus providing sufficient substrate for downstream carotenoid biosynthesis. The ethylene treatment also up‐regulated the expression of CitLCYB, CitHYD (particularly), and CitNCED2, and down‐regulated CitLCYE expression, thereby promoting the accumulation of β‐branch orange carotenoids (especially β‐cryptoxanthin) and inhibiting α‐branch yellow carotenoid (lutein) biosynthesis. These results elucidate the ethylene‐induced formation of the orange flavedo in Satsuma mandarin fruit.

Figure 2.

Figure 2

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Effects of ethylene on the carotenoid metabolism of Satsuma mandarin fruit

Transcript levels of CitPSY (A), CitLCYB (B), CitHYD (C), CitNCED2 (D), and CitLCYE (E) in flavedo of treated fruit. Data represent means ± SD of three biological replicates. Asterisks indicate statistically significant differences which were determined by Student's t‐test (**P < 0.01; n.s., no significant difference). (F) Carotenoid metabolic pathway of Satsuma mandarin fruit under ethylene treatment. Red arrows and words indicate the high levels of carotenoid components and highly expressed carotenoid metabolism genes under ethylene treatment, and then blue indicates the opposite. Black bold arrows indicate the flow of carotenoids to the β‐branch.

Identification of the C2H2‐type zinc‐finger TF CitZAT4

Above, we showed that CitHYD and CitLCYE, as core carotenogenic genes, were mainly responsible for ethylene‐promoted orange coloration in Satsuma mandarin flavedo, which was consistent with previous findings (Zhou et al., 2010). To identify TFs regulating the ethylene‐induced orange coloration in citrus flavedo, we performed a yeast one‐hybrid (Y1H) screening with the CitHYD and CitLCYE promoters as bait. We identified four TFs in all Y1H screening assays. However, only one zinc‐finger family TF, CitZAT4, was found to target both CitHYD and CitLCYE promoters, with the others targeting one or the other. We therefore selected CitZAT4 as a candidate regulatory factor for further study.

We amplified the coding sequence of CitZAT4 according to the reference genome of the Citrus Pan‐genome to Breeding Database (CPBD) (http://citrus.hzau.edu.cn/) and found that this gene comprised a 1,584‐bp coding sequence encoding a protein consisting of 527 amino acids. A Basic Local Alignment Search Tool (BLAST) search of the gene using the National Center for Biotechnology Information (NCBI) database revealed that this protein contained three C2H2‐type zinc‐finger conserved domains in the N‐ and C‐terminal regions (Figure S1A). To explore the potential function of CitZAT4, we conducted a phylogenetic comparison of this protein against all C2H2‐type zinc‐finger proteins in Arabidopsis (Arabidopsis thaliana (L.) Heynh), revealing that CitZAT4 was most similar to AT1G26610 and AT1G26590 (Figure S1B). A multiple sequence alignment showed that CitZAT4, AT1G26610, and AT1G26590 all contained three conserved C2H2‐type zinc‐finger domains (Figure S1C). AT1G26610 and AT1G26590 have not yet been fully characterized, suggesting they may have potentially novel functions.

CitZAT4, a nuclear TF, is involved in regulating ethylene‐induced orange coloration in the flavedo of Satsuma mandarin fruit

First, we examined the localization of CitZAT4 using a subcellular localization assay. As shown in Figure 3A, after a 3‐d incubation, green fluorescent protein (GFP)‐fused CitZAT4 (GFP–CitZAT4) was observed to co‐localize with the red fluorescent nuclear protein NF‐YA‐mCherry in the nucleus, indicating that CitZAT4 localizes to the nucleus.

Figure 3.

Figure 3

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Analysis of Citrus zinc‐finger transcription factor CitZAT4 transcriptional characteristics and ethylene response

(A) Subcellular localization of CitZAT4‐GFP (green fluorescent protein). Bars = 25 μm. CitZAT4‐GFP, GFP signal; NF‐YA4‐mCherry, red fluorescent protein (RFP) signal; Merged, combined GFP and RFP signals. (B) Mapping of the transactivation motif of CitZAT4 using a yeast assay. Yellow modules represent the C2H2‐type zinc‐finger conserved domain of CitZAT4 protein. (C) CitZAT4 transactivation activity assay. pGBKT7‐53 + pGADT7‐RecT and empty vector pGBKT7 were used as positive control (P. Control) and negative control (N. Control), respectively. (D) Expression of CitZAT4 under various treatments. Correlation analysis of CitZAT4 expression and CitLCYE expression (E) as well as CitHYD expression (F) under ethylene treatment. Data represent means ± SD of three biological replicates. Asterisks indicate statistically significant differences which were determined by Student's t‐test (**P < 0.01; n.s., no significant difference).

Subsequently, we performed a yeast two‐hybrid (Y2H) experiment with AH109 yeast cells to determine the transcriptional activity of CitZAT4 in vivo. We divided CitZAT4 into four fragments based on the positions of conserved domains (Figure 3B). These four fragments were cloned into the pGBKT7 vector and transformed into yeast cells. The yeast cells harboring the pGBKT7–CitZAT41–460/CitZAT4 constructs and the positive control pGBKT7‐53 + pGADT7‐RecT grew well on synthetic defined (SD)/‐Trp‐His‐Ade plates, but the N‐terminal fragments pGBKT7‐CitZAT41–34, CitZAT41–124, and CitZAT41–264 and the negative control did not grow (Figure 3C), indicating that the C terminus of CitZAT4 is necessary for its transcriptional activity.

In addition to the typical zinc‐finger domain, most C2H2‐type zinc‐finger proteins also contain an ERF‐associated amphiphilic repression (EAR) motif near the C terminus, which functions as a transcriptional repressor domain (Yanover and Bradley, 2011; Wang et al., 2022b). Our BLAST search for the EAR motif using the PlantEAR functional analysis platform for plant EAR motif‐containing proteins showed that a typical EAR motif was present at the C terminus of CitZAT4 (Figure S2). This result implies that CitZAT4 might repress the transcription of downstream target genes.

To further explore the CitZAT4‐mediated regulation of ethylene‐induced orange coloration in the Satsuma mandarin flavedo, we examined the expression of this gene throughout the storage period after an ethylene treatment. The qRT‐PCR results showed that CitZAT4 expression was significantly higher under the ethylene treatment than under the control and 1‐MCP + ethylene treatment, especially at 5 DAT (Figure 3D). A correlation analysis showed that CitZAT4 expression was significantly negatively and positively correlated with the expression levels of CitLCYE and CitLCYB, respectively, during the ethylene‐induced orange coloration of the Satsuma mandarin flavedo (Figure 3E, F). These findings reveal that CitZAT4 positively responds to the ethylene signal and participates in regulating ethylene‐induced orange flavedo formation in Satsuma mandarin fruit.

CitZAT4 positively regulates β‐branch carotenoid biosynthesis and negatively regulates α‐branch carotenoid accumulation in citrus calli

To evaluate the possible function of CitZAT4, we stably overexpressed CitZAT4 in citrus calli and selected three independent CitZAT4‐overexpressing lines (OE1, OE6, and OE9) with high CitZAT4 transcription levels using qRT‐PCR (Figure S3). As shown in Figure 4A, the CitZAT4‐overexpressing (PH7‐CitZAT4) citrus calli exhibited a more yellow color than the control (empty PH7 vector). Next, we examined the carotenoid contents in control and CitZAT4‐overexpressing calli. A HPLC analysis revealed that the three independent CitZAT4‐overexpressing lines exhibited higher levels of phytoene, β‐carotene, violaxanthin, and total carotenoids than the control but lower lutein levels (Figure 4B). Similarly, the percentage of β‐branch carotenoids (β‐carotene and violaxanthin) was significantly higher in the CitZAT4‐overexpressing calli than in the control, whereas the percentage of α‐branch lutein was remarkably lower (Figure 4C). Compared with their expression in the control, CitPSY and the β‐branch carotenogenic genes (CitLCYB, CitHYD, and CitNCED2) were up‐regulated in the CitZAT4‐overexpressing calli, while CitLCYE expression was down‐regulated (Figure 4D). These results indicate that overexpressing CitZAT4 promotes β‐branch carotenoid biosynthesis and inhibits α‐branch carotenoid accumulation in citrus calli.

Figure 4.

Figure 4

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Stable overexpression of CitZAT4 in citrus calli

(A) Phenotypes of stable transformation of CitZAT4 in citrus calli. PH7‐CitZAT4 indicates overexpressing CitZAT4, and empty vector PH7 as control. Bars = 2 cm. (B) Carotenoid content. (C) The percentage of α‐ and β‐branch carotenoid content in the CitZAT4‐overexpressing citrus calli. (D) The expression levels of carotenoid metabolism genes. Data represent means ± SD of three biological replicates. Asterisks indicate statistically significant differences which were determined by Student's t‐test (**P < 0.01).

We also generated RNA interference (RNAi) lines to silence the expression of CitZAT4 in citrus calli. As shown in Figure S4A, there was no significant color difference between the CitZAT4‐interfering calli and the control. qRT‐PCR showed that the expression of CitLCYB and CitHYD in the CitZAT4‐interfering calli was significantly lower than in the control, but there was no significant difference in the expression of other carotenogenic genes between these genotypes (Figure S4B). Similarly, HPLC showed that the interference of CitZAT4 significantly reduced the β‐carotene content but did not significantly affect the other carotenoids or the total carotenoids (Figure S4C). Moreover, the percentage of β‐branch carotenoids (β‐carotene and violaxanthin) among the total carotenoids was lower in the CitZAT4‐interfering calli than in the control, whereas α‐branch lutein was higher than that in control (Figure S4D). These results further support the conclusion that CitZAT4 positively regulates β‐branch carotenoid accumulation in citrus calli.

CitZAT4 is essential for ethylene‐induced orange coloration in the Satsuma mandarin flavedo

As β‐cryptoxanthin was difficult to detect due to its extremely low levels in citrus calli, we further performed transient injection experiments with citrus fruit to explore the function of CitZAT4. Agrobacterium tumefaciens cells containing the CitZAT4‐overexpression construct PK7‐CitZAT4 were infiltrated into Satsuma mandarin fruit. The tissue surrounding the injection site turned orange in the fruit transformed with the CitZAT4‐overexpression construct, whereas no coloration was detected in the control (injected with an empty PK7 vector) (Figure 5A), indicating that the expression of CitZAT4 was responsible for flavedo coloration. HPLC revealed that the CitZAT4‐overexpressing fruit had higher levels of phytoene, β‐carotene, β‐cryptoxanthin, violaxanthin, and total carotenoids than the control, but less lutein (Figure 5B). Likewise, the percentage of β‐branch carotenoids (β‐carotene and violaxanthin) was significantly higher in CitZAT4‐overexpressing fruit than in the control, whereas the α‐branch lutein percentage was remarkably lower than in the control (Figure 5C). qRT‐PCR showed that CitPSY and the β‐branch carotenogenic genes (CitLCYB, CitHYD, and CitNCED2) were up‐regulated in CitZAT4‐overexpressing fruit, while the α‐branch carotenogenic gene (CitLCYE) expression was down‐regulated (Figure 5D). The above results jointly revealed that overexpressing CitZAT4 significantly promotes orange coloration in the Satsuma mandarin flavedo.

Figure 5.

Figure 5

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Transient expression of CitZAT4 in citrus fruit

(A) Phenotypes of transient expression of CitZAT4 in citrus fruit. Empty vector PK7 and RNA interference (RNAi) as control. PK7‐CitZAT4 and RNAi‐CitZAT4 indicate overexpressing and interfering CitZAT4, respectively. Ethylene treatment was performed after 3 d of infiltration. Bars = 3 cm. (B) Carotenoid content. (C) The percentage of α‐ and β‐branch carotenoid content in Satsuma mandarin fruit transiently expressing CitZAT4. (D) The expression levels of CitZAT4 and carotenoid biosynthesis genes. Data represent means ± SD of three biological replicates. Asterisks indicate statistically significant differences which were determined by Student's t‐test (**P < 0.01).

To further determine the role of CitZAT4 in ethylene‐induced flavedo coloration, we transformed Satsuma mandarin fruit by injecting an A. tumefaciens solution containing the CitZAT4‐interfering construct or an empty RNAi vector as a control. The CitZAT4‐interfering and control fruit were treated with an ethylene solution and the coloration changes around the injection site were examined. Orange coloration was observed around the injection site in the flavedo of control fruit; however, the injection site of the CitZAT4‐interfering fruit remained pale yellow (Figure 5A). HPLC analysis showed that the contents of phytoene, β‐branch orange carotenoids (β‐carotene, β‐cryptoxanthin, and violaxanthin), and total carotenoids in the CitZAT4‐interfering fruit were significantly lower than in the control, while the α‐branch lutein content was significantly higher (Figure 5B, C). Moreover, the expression patterns of CitPSY and β‐branch carotenogenic genes (CitLCYB, CitHYD, and CitNCED2) were consistent with the changes in phytoene and β‐branch orange carotenoid (β‐carotene, β‐cryptoxanthin, and violaxanthin) contents, and the expression pattern of the α‐branch carotenogenic gene CitLCYE was in line with the change in the α‐branch lutein content (Figure 5D). qRT‐PCR further indicated that the expression of CitZAT4 was responsible for the above changes (Figure 5D). In summary, we found that CitZAT4 is an essential regulator of ethylene‐promoted orange coloration in the flavedo of Satsuma mandarin fruit.

Ethylene signaling enhances the regulatory effect of CitZAT4 on downstream carotenoid metabolism genes

Based on the finding that the expression of carotenogenic genes was significantly changed in the transgenic citrus calli and fruit, we speculated that carotenogenic genes might be the targets of CitZAT4. To test this, we first performed a Y1H assay to investigate the interactions between CitZAT4 and the promoters of CitPSY, CitLCYB, CitHYD, CitNCED2, and CitLCYE. All transformed yeast cells were spotted on the SD/‐Leu and SD/‐Leu/AbAx culture medium. After 3 d of culture, the yeast cells co‐transformed with the positive control (PGADT7‐Recp53 + p53‐AbAi) and the experimental group (PGADT7‐CitZAT4 + pAbAi‐proCitPSY, proCitLCYB, proCitHYD, proCitNCED2, proCitLCYE) survived on the selective medium (SD/‐Leu/AbAx), while the negative control (empty PGADT7 vector PGADT7 + pAbAi‐promoter) did not (Figure 6A). These results suggest that CitZAT4 directly interacts with the CitPSY, CitLCYB, CitHYD, CitNCED2, and CitLCYE promoters.

Figure 6.

Figure 6

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Analysis of the binding interaction between Citrus zinc‐finger transcription factor CitZAT4 and promoters of carotenoid metabolism genes

(A) Yeast one‐hybrid assay identified interactions of CitZAT4 with target gene promoters. Empty PGADT7 + pAbAi‐proCBG and PGADT7‐Rec‐p53 + p53‐AbAi as the negative and positive controls (P. Control), respectively. CBG, carotenoid biosynthesis genes. Aureobasidin A (AbA), as a yeast cell growth inhibitor. SD/‐Leu/AbAx, synthetic defined/‐Leu medium supplemented with 200 ng/mL as the basal concentration of proCitLCYB and proCitLCYE. SD/‐Leu/AbAx, SD/‐Leu medium supplemented with 150 ng/mL as the basal concentration of proCitHYD and proCitNCED2. (B–F) Chromatin immunoprecipitation – polymerase chain reaction(ChIP‐PCR) assays showed the interaction of CitZAT4 with several regions in the promoters of CitPSY, CitLCYB, CitLCYE, CitHYD, and CsNCED2, respectively. Asterisks indicate statistically significant differences which were determined by Student's t‐test (*P < 0.05; **P < 0.01; n.s., no significant difference).

To further determine whether this interaction also occurs in vivo, we performed a chromatin immunoprecipitation (ChIP) assay followed by quantitative PCR (ChIP‐qPCR). The fusion construct CitZAT4‐GFP was overexpressed in citrus calli with the empty GFP vector as a control. The results of our PCR‐based assay showed that CitZAT4 was substantially enriched on the promoters of the carotenogenic genes, indicating that CitZAT4 directly binds to these promoters in vivo (Figures S5, 6B–F). We also conducted electrophoretic mobility shift assays (EMSAs) using the purified CitZAT4‐maltose‐binding protein (MBP) fusion protein, with empty MBP protein as the control. The fragments with the strongest binding signal in the ChIP‐qPCR assay were selected to synthesize the probes. Shifted bands were identified when using the recombinant CitZAT4‐MBP protein containing the hot probe, while the shifted bands were not present in the recombinant CitZAT4‐MBP protein containing the mutated hot probe (Figure 7A–E). The increasing addition of unlabeled DNA probes led to a clear weakening of the shifted bands (Figure 7A–E). These data indicate that CitZAT4 directly binds to the promoters of carotenogenic genes in vitro.

Figure 7.

Figure 7

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Analysis of the regulatory effect of Citrus zinc‐finger transcription factor CitZAT4 on carotenoid metabolism genes

(A–E) Electrophoretic mobility shift assay (EMSA) indicated that CitZAT4 directly binds the promoters of CitPSY, CitHYD, CitNCED2, CitLCYB, and CitLCYE, respectively. Purified maltose‐binding protein (MBP) was used as a negative control. Black arrows point to the position of biotin‐labeled promoter fragment (hot probe). Red arrows point to the positions of protein–DNA complexes or free probes. Red letters represent the binding motifs, and blue letters indicate their corresponding mutant motifs. “+” and “−” indicate the presence and absence of the probe or protein, respectively. The unlabeled DNA fragments (cold probe) were added (50‐, 100‐, and 150‐fold) as competitors. (F) Schematic representation of reporter and effector constructs used in dual‐luciferase assay. CBG, carotenoid biosynthesis genes. Dual‐luciferase assays indicated that CitZAT4 activates these promoters and this activation was induced by ethylene. ACC (1‐aminocyclopropane‐1‐carboxylate) represents these materials used for luciferase assays under ethylene treatment. Data represent means ± SD of three biological replicates. Asterisks indicate statistically significant differences which were determined by Student's t‐test (**P < 0.01).

To investigate the regulatory effect of CitZAT4 on the target genes, we co‐transformed the Pro35S:CitZAT4 and ProCBG:LUC constructs into Nicotiana benthamiana leaves (Figure 7F). We found that CitZAT4 significantly enhanced the activities of the target genes except for CitLCYE, which was markedly suppressed by CitZAT4 (Figure 7F). Furthermore, the luciferase (LUC) intensity of the target gene promoters was further strengthened or weakened by an ethylene treatment (Figure 7F). Taken together, our findings reveal that ethylene‐induced CitZAT4 directly activates the expression of CitPSY and the β‐branch carotenogenic genes (CitLCYB, CitHYD, and CitNCED2), and suppresses the expression of the α‐branch carotenogenic gene CitLCYE.

The transcriptional cascade CitERF061–CitZAT4 positively regulates ethylene‐induced orange coloration in citrus flavedo

In previous studies, we identified a citrus ERF, ERF061, which is induced by ethylene and positively regulates carotenoid accumulation by activating the expression of nine carotenogenic genes involved in the β‐branch of the carotenoid pathway (Zhu et al., 2021). We therefore investigated whether ERF061 acts as an upstream regulatory factor of CitZAT4, bridging the ethylene signaling pathway and the regulation of CitZAT4 expression. A PlantCARE analysis showed there are two typical ERF‐binding motifs on the CitZAT4 promoter, which further strengthened our speculation (Figure S6). To test our hypothesis, we first performed a Y1H assay to investigate the interactions between CitERF061 and the CitZAT4 promoter. As shown in Figure 8A, after a 3‐d culture, the yeast cells co‐transformed with PGADT7‐CitERF061 + pAbAi‐proCitZAT4 survived on the selective medium (SD/‐Leu/AbA200), while those expressing the negative control (PGADT7 + pAbAi‐proCitZAT4) did not, indicating that CitERF061 directly binds to the CitZAT4 promoter. In addition, we used LUC experiments to clarify the regulatory effect of CitERF061 on CitZAT4, revealing that CitERF061 significantly enhances the activity of the CitZAT4 promoter (Figure 8B). Furthermore, an ethylene treatment significantly enhanced this effect (Figure 8C). These results show that ethylene‐induced CitERF061 directly activates CitZAT4 expression.

Figure 8.

Figure 8

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Analysis of Citrus ethylene response factor 061 (CitERF061) regulation of Citrus zinc‐finger transcription factor CitZAT4‐mediated carotenoid metabolism

(A) Yeast one‐hybrid assay identified interactions of CitERF061 with CitZAT4 promoter. Empty PGADT7 + pAbAi‐proCitZAT4 was the negative control. Abscisic acid (ABA) was a yeast cell growth inhibitor. Synthetic defined (SD)/‐Leu/AbA200, SD/‐Leu medium supplemented with 200 ng/mL AbA was the basal concentration of proCitZAT4. (B, C) Schematic representation of CitZAT4 promoter used in dual‐luciferase assay. Dual‐luciferase assays indicated that CitERF061 activates CitZAT4 promoter and this activation was induced by ethylene. ACC (1‐aminocyclopropane‐1‐carboxylate) represents these materials used for luciferase assays under ethylene treatment. (D–G) Stable transformation of CitERF061 in citrus calli. (D) Phenotypes. PH7‐CitERF061 indicates overexpressing CitERF061 and empty vector PH7 as control. (E) CitERF061 expression. (F) CitZAT4 expression. (G) Carotenoid content. (H–J) Transient expression of CitERF061 in citrus fruit. (H) Phenotypes. PK7‐CitERF061 indicates overexpressing CitERF061, and empty vector PK7 as control. Bars = 3 cm. (I) The expression of CitZAT4 and CitERF061. (J) Carotenoid content. Data represent means ± SD of three biological replicates. Asterisks indicate statistically significant differences which were determined by Student's t‐test (*P < 0.05; **P < 0.01; n.s., no significant difference).

In order to clarify the function of CitERF061 in CitZAT4‐regulated carotenoid metabolism, we determined the calli coloration, CitZAT4 expression and carotenoid content of CitERF061‐overexpressing calli. As shown in Figure 8D–G, CitERF061‐overexpressing calli had a yellowish color, and both the CitZAT4 expression and carotenoid content were significantly higher than in the control. In addition, we generated RNAi lines silencing CitERF061 expression in citrus calli. As shown in Figure S7A, there was no significant color difference between the CitERF061‐interfering calli and the control. qRT‐PCR showed that CitLCYB and CitHYD expression was significantly lower in the CitERF061‐interfering calli than in the control, although there were no significant differences in the expression levels of the other carotenogenic genes or CitZAT4 (Figure S7B). Similarly, an HPLC analysis showed that the RNAi of CitERF061 significantly reduced the β‐carotene content but had no significant effect on the other carotenoids or the total carotenoid abundance (Figure S7C). No significant differences were detected in the percentages of the β‐branch carotenoids (β‐carotene and violaxanthin) or the α‐branch carotenoid (lutein) between the control and the CitERF061‐interfering calli (Figure S7D). These results further confirm that CitERF061 positively modulates the regulatory activity of CitZAT4 on the flux of carotenoid metabolism in citrus calli. Furthermore, we performed transient expression injection experiments in citrus fruits. Orange coloration, higher carotenoid contents, and increased CitZAT4 expression were found in CitERF061‐overexpressing fruit around the injection site, whereas no orange coloration was detected in the control fruit (Figure 8H–J). These results jointly reveal that CitERF061 positively regulates CitZAT4‐mediated orange coloration in citrus fruits.

DISCUSSIONS

Citrus plants produce non‐climacteric fruit, and ABA plays a crucial role in their coloration (Rodrigo et al., 2006; Wu et al., 2014; Romero et al., 2019). Moreover, the ABA content exhibits continuous increases, activating the expression of the carotenoid biosynthesis genes to drive peel coloration during citrus fruit ripening (Sun et al., 2024). Although citrus is defined as non‐climacteric fruits, they undergo climacteric‐like endogenous ethylene production during fruit ripening, which is induced by the genes responsible for 1‐aminocyclopropane‐1‐carboxylate (ACC) synthesis (including CsACS1, CsACS6, and CsACS9) and ACC oxidase (mainly CsACO1 and CsACO4) (Ding et al., 2015; Feng et al., 2021). In addition, ethylene treatment has been widely reported to significantly promote fruit coloration by regulating the carotenoid biosynthesis pathway in various citrus species (Rodrigo and Zacarias, 2007; Matsumoto et al., 2009; Sun et al., 2021). Here, we showed that ethylene promotes orange coloration in the Satsuma mandarin flavedo by mediating the carotenoid metabolic flow (Figures 1A, 2F), highlighting the importance of ethylene in the coloration of citrus fruits.

Carotenoid concentration and composition vary greatly across citrus varieties (Xu et al., 2006; Wang et al., 2008); hence, their carotenoid accumulation during ethylene‐induced flavedo coloration differs as well (Zhou et al., 2010). The red color of ethylene‐treated tangerines, tangelos, and oranges is mainly due to the accumulation of β‐citraurin (Rodrigo and Zacarias, 2007; Sun et al., 2021), while the yellow color of ethylene‐treated lemons, limes, and pummelos is mainly due to the accumulation of lutein and violaxanthin (Lucchesi et al., 1983; Nishikawa et al., 2002; Zhang and Zhou, 2019). The orange color of ethylene‐treated Satsuma mandarins is mainly determined by the accumulation of β‐cryptoxanthin (Fujii et al., 2007; Rodrigo and Zacarias, 2007; Matsumoto et al., 2009).

In this study, we systematically analyzed the effect of ethylene treatment on carotenoid metabolism in the Satsuma mandarin flavedo. We found that ethylene promoted phytoene accumulation by inducing CitPSY expression (Figures 1D, 2A), thus providing sufficient substrates for downstream carotenoid biosynthesis. Our results are consistent with previous reports in other citrus species, including tangerines and oranges, after ethylene treatment (Rodrigo and Zacarias, 2007; Sun et al., 2021). Moreover, a postharvest ethylene application clearly increased the phytoene content of tomato (Solanum lycopersicum L.) and apricot (Prunus armeniaca L.) fruits by stimulating the expression of PSY (Picton et al., 1993; Hoeberichts et al., 2002; Marty et al., 2005). The above findings suggest that the rapid upregulation of early carotenogenic genes appears to be a common response to ethylene in climacteric or non‐climacteric fruits, which promotes the accumulation of carotenoid precursors and their flux into the carotenoid pathway.

It should be noted that CitLCYE was the only down‐regulated carotenogenic gene under the ethylene treatment. The striking decline in its expression might be responsible for the decrease in α‐branch carotenoid biosynthesis (mainly lutein) (Figures 1H, 2E). Conversely, the β‐branch carotenogenic genes (particularly CitHYD) were significantly up‐regulated by the ethylene treatment, thus promoting the accumulation of β‐branch carotenoids (especially β‐cryptoxanthin) (Figures 1F, 2C). Changes in the CitLCYE and CitHYD expression levels in ethylene‐treated citrus fruit played a key role in the shift of carotenoid biosynthesis from the α‐branch to the β‐branch (Figure 2C, E, F). Similar ethylene‐induced changes in carotenoid metabolic flux (α‐branch to β‐branch) have also been reported in other citrus varieties, such as oranges and tangerines (Rodrigo and Zacarias, 2007; Zhou et al., 2010). Differently, ethylene promotes the accumulation of β‐citraurin (a red carotenoid produced from the β‐branch) by activating CCD4b expression, thus resulting in flavedo reddening in oranges and tangerines (Sun et al., 2021). Taken together, these findings suggest that ethylene treatment dramatically promotes the flux of carotenoids into the β‐branch, eventually leading to flavedo coloration, which is ubiquitous in different citrus varieties. The specific accumulation of different carotenoids contributes to the formation of distinctive flavedo colors, which are specific to ethylene‐treated citrus fruits.

As transcription activators or repressors, C2H2‐type zinc‐finger TFs play indispensable roles in regulating physiological processes in plants; for instance, AtZP1, a transcriptional repressor, contains an EAR motif and negatively regulates root hair initiation and elongation by suppressing the transcription of bHLH TF genes (Han et al., 2020b). Other C2H2‐type zinc‐finger TFs, such as ZPT2‐related proteins, also contain an EAR motif, and function as transcriptional repressors involved in plant growth in response to abiotic stress (Sakamoto et al., 2004; Kodaira et al., 2011). In addition, C2H2‐type zinc‐finger TFs containing an EAR motif have been reported to possess dual functions of transcriptional activation and inhibition. The overexpression of MdZAT17 and MdZAT5 increases the anthocyanin content by inducing the expression of anthocyanin biosynthesis genes and improves the sensitivity to salt stress by inhibiting the expression of salt‐stress‐related genes in apple (Malus pumila Mill.) (Wang et al., 2022a, 2022b). C2H2‐type zinc‐finger TFs thus play important roles in biological processes such as stress responses, phytohormone signal transduction, and secondary metabolism.

In the current study, we identified an ethylene‐induced C2H2‐type zinc‐finger family protein, CitZAT4, which contains three C2H2‐type zinc‐finger conserved domains at the N and C terminals as well as one EAR motif and one transactivation motif at the C terminus (Figures S1, S2). The expression pattern of CitZAT4 was closely related to the orange coloration of the citrus flavedo and the accumulation of β‐branch carotenoids (especially β‐cryptoxanthin). CitZAT4 expression was similar to the expression patterns of the β‐branch carotenogenic genes (particularly CitHYD) but contrasted with that of the α‐branch carotenoid metabolism (CitLCYE expression and lutein content) (Figure 3D–F). These results suggest that CitZAT4 contributes to ethylene‐induced orange coloration in the Satsuma mandarin flavedo. Our molecular biochemical experiments and transgenic assays further confirmed that ethylene promotes CitZAT4 to regulate the shift of carotenoid metabolic flow from the α‐branch to the β‐branch by directly activating the expression of β‐branch carotenogenic genes (particularly CitHYD) and inhibiting the transcription of the α‐branch carotenogenic gene CitLCYE (4, 5, 6, 7). This resulted in the accumulation of orange carotenoids dominated by β‐cryptoxanthin, finally forming the orange flavedo of Satsuma mandarin fruit (Figures 1, 2, and 5). Our results reveal the dual functions of C2H2‐type zinc‐finger TFs, namely, transcriptional activation and transcriptional repression, especially in regulating carotenoid metabolism.

The ERFs are the primary ethylene‐responsive TFs and activate the expression of genes in the ethylene response pathway, ultimately regulating various biological processes (Fujimoto et al., 2000; Alonso Jose and Stepanova Anna, 2004; Lin et al., 2009; An et al., 2018). Our qRT‐PCR results showed that CitZAT4 significantly responded to ethylene induction (Figure 3D). Moreover, we found multiple potential ERF‐binding elements on the CitZAT4 promoter (Figure S6). We therefore speculated that the ERF TFs might respond to ethylene signaling and promote CitZAT4 expression to regulate carotenoid metabolism. Our previous studies reported that an ethylene‐induced ERF TF, ERF061, is involved in regulating carotenoid metabolism in citrus (Zhu et al., 2021). Notably, ERF061 positively regulates carotenoid accumulation by activating the expression of nine carotenogenic genes involved in the β‐branch of the carotenoid pathway. This suggests that ERF061 may serve as an upstream regulator of CitZAT4 expression in response to ethylene signaling, participating in the regulatory role of CitZAT4 in carotenoid metabolic pathway flux. In this study, molecular biochemical and transgenic experiments jointly demonstrated that ethylene‐induced CitERF061 directly binds to the CitZAT4 promoter and activates its expression, thus forming the transcriptional cascade CitERF061–CitZAT4 and ultimately regulating ethylene‐promoted orange coloration in the Satsuma mandarin flavedo. While CitERF061 positively regulates CitZAT4‐mediated orange coloration in citrus fruit, there was no significant difference in CitZAT4 expression after CitERF061 was silenced (Figure S7B). As shown in Figure 8D–G, we found that, although the CitZAT4 expression, calli color, and carotenoid content of the CitERF061‐overexpressing calli were significantly higher than those of the control, this was the result of an approximately 20,000‐fold increase in CitERF061 expression. These results imply that the expression of CitERF061 in wild‐type citrus calli was very low, which is the reason why there was no significant difference in callus color or the expression of the target genes (including CitZAT4, CitPSY, CitNCED2, and CitLCYE) between the CitERF061‐interfering calli and the control (Figure S7).

In summary, we propose a regulatory model in which ethylene‐induced CitZAT4 regulates the formation of the orange flavedo in Satsuma mandarin fruit (Figure 9). First, CitERF061, induced by ethylene signaling, directly binds to the CitZAT4 promoter and activates its expression. Subsequently, CitZAT4 activates the expression of CitPSY to provide sufficient substrates for downstream carotenoid biosynthesis. Furthermore, CitZAT4 regulates the shift of the carotenoid metabolic flow from the α‐branch to the β‐branch by inhibiting the expression of CitLCYE and inducing the expression of CitLCYB, CitHYD, and CitNCED2. This promotes the accumulation of orange carotenoids (especially β‐cryptoxanthin), ultimately forming the orange flavedo in ethylene‐treated Satsuma mandarin fruit. Our findings provide a new target for improving citrus fruit quality.

Figure 9.

Figure 9

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Model in which CitZAT4 regulates ethylene‐induced orange‐colored flavedo in Satsuma mandarin fruit

First, Citrus ethylene response factor 061 (CitERF061), induced by ethylene signaling, directly binds to the CitZAT4 promoter and activates CitZAT4 expression. Subsequently, Citrus zinc‐finger transcription factor CitZAT4 directly activates CitPSY expression to provide sufficient substrates (phytoene) for downstream carotenoid metabolism. Second, CitZAT4 significantly inhibits the biosynthesis of α‐branch carotenoid (yellow‐colored lutein) by down‐regulating CitLCYE expression, and promotes the accumulation of β‐branch carotenoid (orange‐colored β‐carotene, β‐cryptoxanthin, and violaxanthin). Finally, ethylene‐induced CitZAT4 promotes the carotenoid metabolism flow into the β‐branch, thus increasing the accumulation of orange‐colored carotenoids, eventually leading to orange coloration in ethylene‐treated Satsuma mandarin flavedo. “+” represents promotion. “−” represents inhibition.

MATERIALS AND METHODS

Plant materials and treatments

Citrus calli were cultured on solid Murashige Tucker (MT) medium in a growth chamber (25°C; dark) and were subcultured on new MT medium every 20 d.

Harmless Satsuma mandarin (Citrus unshiu Marc.) fruits were randomly selected at 180 DAFB (days after full blossom) in the National Center of Citrus Breeding, Huazhong Agricultural University, Wuhan, China, and then immediately transferred to the laboratory. Citrus fruit was treated with 500 mg/L ethephon for 2 min as ethylene treatment; ethylene treatment was performed after treatment with 5 mg/L 1‐MCP (EthylBloc, Floralife Inc., USA) in plastic chambers for 16 h as 1‐MCP + ethylene treatment; citrus fruits were treated with distilled water for 2 min as control. Subsequently, all fruits were placed in a phytotron (temperature: 23°C–25°C; dark) after natural drying for 10 d. The flavedo was collected along the equatorial plane of citrus fruit every 5 d from 0 d after treatment (DAT) to 10 DAT. A total of 84 Satsuma mandarin fruits were used for treatment throughout the experiment. At 0 DAT, 12 fruits each were in the control, ethylene and ethylene + 1‐MCP treatment groups. At 5 and 10 DAT, the control, ethylene, and ethylene + 1‐MCP treatment groups contained 12 fruits, respectively, for a total of 72 fruits. All flavedo samples were immediately frozen in liquid nitrogen and stored at −80°C for further analysis.

Gene isolation, phylogenetic tree construction, and multiple sequence alignment

The coding sequence (CDS) of CitZAT4 from Satsuma mandarin fruit flavedo was amplified based on the reference genome of CPBD (http://citrus.hzau.edu.cn/). Multiple sequence alignments and phylogenetic tree were performed using NCBI and MEGA 7.0. Analysis of EAR motif was conducted as previously described (Yang et al., 2018). All primers were designed using Primer 5.0 and are listed in Table S1.

Determinations and analyses of CCI and carotenoid content

The CCI was quantified according to the formula: CCI = 1,000 a × L−1 × b−1, and was used to indicate citrus peel color. Peel color parameters of L, a, and b were measured using the KONICA MINOLTA CR‐400 (Japan) following the manufacturer's instructions.

Carotenoid extraction, determination, and analysis of flavedo and calli were performed as previously described (Sun et al., 2023, 2024). Nine citrus fruits were randomly divided into triplicate as three biological replicates. Each set of flavedo was used as one biological replicate. One replicate was determined three times.

Total RNA isolation and qRT‐PCR

All samples were ground to powder with liquid nitrogen, and then RNAs from these samples were extracted using TRIzol RNA. qRT‐PCR was performed with a Roche LightCycler 480 system using the 2 × LightCycler 480 SYBR Green master mix (Roche, Basel, Switzerland); the specific operation is as described previously (Sun et al., 2023, 2024). Three biological replicates from independent samples were determined, and each biological replicate was performed in triplicate. Gene‐specific primers used for qRT‐PCR are shown in Table S1.

Subcellular localization and transcriptional activation analysis in yeast cells

For subcellular localization, transient co‐expression of CitZAT4‐GFP and the nuclear marker NF‐YA4‐mCherry into N. benthamiana leaves via Agrobacterium tumefaciens‐mediated transformation. After incubation for 3 d, fluorescence was observed and imaged under a confocal scanning microscope (TCS SP8, Leica, Germany).

For exploring the transcriptional activation domain of CitZAT4, the full‐length or truncated CDSs of CitZAT4 were cloned into the PGBKT7 vector, and then the construct was transformed into yeast strain (AH109) according to the yeast protocols handbook (TaKaRa). The empty vector PGBKT7 and PGBKT7‐53 + PGADT7‐T plasmids were used as negative and positive controls, respectively. The growth status of yeast cells was used to evaluate the transcriptional activation domain of CitZAT4.

Yeast one‐hybrid analysis and EMSA

For Y1H assay, the promoters of target genes (TGs) were cloned into pABAi vector and the CDSs of CitZAT4 and CitERF061 were cloned into PGADT7 vector. Empty PGADT7 + pAbAi‐proTGs and PGADT7‐Rec‐p53 + p53‐AbAi acted as the negative (N. Control) and positive controls (P. Control), respectively. The Y1H assay was performed according to the manufacturer's instructions (Matchmaker® Gold Y1H Library Screening System User Manual; TaKaRa).

The LightShiftTM Chemiluminescent EMSA kit (Thermo Fisher Scientific, Waltham, MA, USA) was used in EMSA according to the manufacturer's instructions. Purified MBP was used as control. Biotin‐labeled and mutant hot probes were synthesized by Sangon Biotech, and an unlabeled probe was used as the competitor. The whole process was operated according to the manufacturer's instructions. All probe sequences are shown in Figure 7A–E.

Transformation of citrus calli and injection of citrus fruit

The stable transformation of citrus calli was conducted following previously described methods (Sun et al., 2023, 2024). Briefly, the A. tumefaciens infection solution containing PH7‐CitZAT4 overexpression construct was incubated with citrus calli for 30 min, and then the transformed citrus calli were then screened on resistance MT medium. The empty PH7WG2D (PH7) vector was used as control. Three independently transgenic lines as one biological replicate and one biological replicate were repeated three times for further determining.

The transiently infiltration of Satsuma mandarin fruit was performed as previously described (Sun et al., 2023, 2024). Briefly, overexpression (PK7‐CitZAT4) and interference (RNAi‐CitZAT4) constructs were transformed into A. tumefaciens GV3101 strain. Empty PK7 and RNAi vectors were used as control. Flavedo from around the injection site was sampled and uniformly divided into three portions for further analysis. Three portions acted as three biological replicates and each biological replicate was performed in triplicate.

Dual‐luciferase reporter assay

PK7‐CitZAT4 and PK7‐CitERF061 vectors were effectors. The promoters of TGs were cloned into pGreenII 0800 vector as the reporter. Luciferase activities were measured with Dual‐Luciferase® Reporter Assay Kit (Promega, Madison, WI, USA) according to the manufacturer's instruction. Fluorescence intensities were determined by the dual‐luciferase reporter assay system (Promega) with an Infinite200 Pro microplate reader (Tecan, Männedorf, Switzerland).

Chromatin immunoprecipitation‐qPCR assay

The ChIP assay was performed as previously described (Sun et al., 2023, 2024). A SimpleChip Plus Sonication Chromatin IP Kit (Cell Signaling Technology, Danvers, MA, USA) was used in the ChIP experiment according to the manufacturer's instructions. Chromatin extracts were prepared from transgenic citrus calli (overexpression of CitZAT4‐GFP fusion construct). The chromatin was sheared using a Vibra‐cell VCX 150 sonicator (Sonics & Materials, USA) and immunoprecipitated with an anti‐GFP antibody (Roche, USA). The enriched chromatin fragments were analyzed by qRT‐PCR. Three independently transgenic calli acted as three biological replicates and each replicate was performed in triplicate. The specific primers used in this ChIP‐qPCR assay are shown in Table S1.

Statistical analyses

The data are presented means ± SD of at least three biological replicates. Data analyses were conducted with Microsoft Excel 2019. Figures were drawn with GraphPad Prism 8 and Adobe IIIustrator 2020. Asterisks indicate statistically significant differences which were determined by Student's t‐test (*P < 0.05; **P < 0.01; n.s., no significant difference).

Accession numbers

Sequence data from this study can be found in the reference genome of CPBD (http://citrus.hzau.edu.cn/). All accession numbers for this study are shown in Table S1.

CONFLICTS OF INTEREST

The authors declare no conflict of interest.

AUTHOR CONTRIBUTIONS

X.D. supervised the research; Q.S. and X.D. designed the experiments; Q.S. and Z.H. performed the experiments with contributions from R.W., D.F., and Y.Z.; Q.S. wrote the manuscript; Q.S., J.Y., and X.D. revised the manuscript; L.C., Y.C., and Q.X. provided critical comments on manuscript editing. All authors read and approved of the contents of this paper.

Supporting information

Additional Supporting Information may be found online in the supporting information tab for this article: http://onlinelibrary.wiley.com/doi/10.1111/jipb.13778/suppinfo

Figure S1. Sequence alignment and phylogenetic analysis of CitZAT4

Figure S2. Analysis of the ethylene response factor (ERF)‐associated amphiphilic repression (EAR) motif (amino acid sequence: IDLNLP) in the amino acid sequence of CitZAT4

Figure S3. Expression of CitZAT4 in the CitZAT4‐overexpressing of citrus calli

Figure S4. Stable interference of CitZAT4 in citrus calli

Figure S5. Potential binding elements of CitZAT4 on the promoters of carotenoid biosynthesis genes

Figure S6. Potential binding elements of ethylene response factor (ERF) transcription factors (TFs) on the CitZAT4 promoter

Figure S7. Stable interference of CitERF061 in citrus calli

JIPB-67-294-s001.docx (44MB, docx)

Table S1. Genes and primers used in this study

JIPB-67-294-s002.xlsx (17.9KB, xlsx)

ACKNOWLEDGEMENTS

This work was supported by the National Key R&D Program of China (2023YFD2300600), the National Natural Science Foundation of China (No. 31930095) and the National Modern Agricultural (Citrus) Technology Systems of China (No. CARS‐27).

Biographies

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graphic file with name JIPB-67-294-g005.gif

Sun, Q. , He, Z. , Ye, J. , Wei, R. , Feng, D. , Zhang, Y. , Chai, L. , Cheng, Y. , Xu, Q. , and Deng, X. (2025). A novel C2H2‐type zinc‐finger transcription factor, CitZAT4, regulates ethylene‐induced orange coloration in Satsuma mandarin flavedo (Citrus unshiu Marc.). J. Integr. Plant Biol. 67: 294–310.

Edited by: Diqiu Yu, Yunnan University, China

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Associated Data

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

Supplementary Materials

Additional Supporting Information may be found online in the supporting information tab for this article: http://onlinelibrary.wiley.com/doi/10.1111/jipb.13778/suppinfo

Figure S1. Sequence alignment and phylogenetic analysis of CitZAT4

Figure S2. Analysis of the ethylene response factor (ERF)‐associated amphiphilic repression (EAR) motif (amino acid sequence: IDLNLP) in the amino acid sequence of CitZAT4

Figure S3. Expression of CitZAT4 in the CitZAT4‐overexpressing of citrus calli

Figure S4. Stable interference of CitZAT4 in citrus calli

Figure S5. Potential binding elements of CitZAT4 on the promoters of carotenoid biosynthesis genes

Figure S6. Potential binding elements of ethylene response factor (ERF) transcription factors (TFs) on the CitZAT4 promoter

Figure S7. Stable interference of CitERF061 in citrus calli

JIPB-67-294-s001.docx (44MB, docx)

Table S1. Genes and primers used in this study

JIPB-67-294-s002.xlsx (17.9KB, xlsx)

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