Background

Phosphorus (P) is one of the essential macronutrients for plant growth and development. P is involved in the processes of photosynthesis, respiration, energy metabolism and signal transduction in plants [1]. Furthermore, P is also an important structural component of various biomolecules in plant cells, including adenosine triphosphate (ATP), phospholipids, DNA and RNA [2]. Although total P is abundant in soils, P is easily immobilized by soil components into unavailable forms that cannot be directly utilized by plants [3]. Thus, low phosphate (Pi) availability is considered a major limiting factor for crop growth, especially in acid soils that occupy about 50% of the world’s arable land [3, 4]. To obtain high yields in traditional agricultural systems, farmers need to apply excessive quantities of P fertilizer, potentially leading to soil deterioration and eutrophication problems [5]. Furthermore, P fertilizer is derived from mined phosphate rock, which is a finite resource that is slowly depleting [6]. Therefore, improving the absorption and utilization of soil P can be an effective way for increasing crop yield and reducing fertilizer P application. Such improvements aim for the development of a more sustainable and environmentally sound agriculture.

To cope with low-P stress, plants have improved Pi uptake and homeostasis through a wide range of morphological, physiological and molecular changes, such as modifying root morphology and architecture, increasing secretion of organic acid and acid phosphatases, enhancing expression of high-affinity Pi transporters, develo** symbioses with arbuscular mycorrhizal fungi, and regulating complex P signaling networks in plant cells [7, 8]. It has been well demonstrated that plants display plasticity in root growth under P deficiency by changing root morphology and architecture, and thus increasing acquisition of P from the soil [9, 10]. For example, increase in root length and root/shoot ratio is observed in maize (Zea mays) [11], faba bean (Vicia faba) [12], rapeseed (Brassica napus) and wheat (Triticum aestivum) [13] in response to low P supply. Furthermore, acid phosphatase activities are up-regulated by Pi deprivation in many plants, such as rice (Oryza sativa), soybean (Glycine max) and chickpea (Cicer arietinum), which could contribute to increased organic P utilization [14,15,16].

To date, a variety of P responsive genes have been identified to participate in Pi uptake and homeostasis [8, 17]. For example, PHOSPHATE STARVATION RESPONSE 1 (AtPHR1) in Arabidopsis and OsPHR2 in rice, encoding the MYB transcription factor, are the central regulators involved in Pi signaling pathway [8]. AtPHR1 is demonstrated to regulate a set of Pi starvation induced (PSI) genes through binding to the P1BS cis element of target genes [18]. Furthermore, a negative regulatory role for protein containing the SYG1/PHO81/XPR1 (SPX) domain in rice is documented where OsSPX1 suppresses the transcripts of several PSI genes, such as Pi transporters (OsPT2 and OsPT6) and purple acid phosphatases (OsPAP10) [7]. A group of Pi transporters have been functionally characterized to be involved in Pi uptake and/or translocation in many plants; examples include: AtPT1 and AtPT2 from Arabidopsis [19, 20], OsPT1/9/10 from rice [21, 22] and GmPT5/7 from soybean [23, 24]. In addition, numerous purple acid phosphatase (PAP) homologues have also been demonstrated to function in Pi release from organic P, including AtPAP10/12/26 from Arabidopsis [25], OsPHY1 from rice [

Methods

Plant growth and treatments

In this study, two stylo (Stylosanthes guianensis) cultivars, ‘RY2’ and ‘RY5’, were used, which were widely grown in South China [33]. The stylo seeds were provided by the Institute of Tropical Crop Genetic Resources (TCGRI), Chinese Academy of Tropical Agricultural Sciences (CATAS), Hainan, China. Experiments were performed in a greenhouse at temperatures of 25 °C to 32 °C under natural sunlight with a photoperiod of about 13 h at the TCGRI, CATAS, Hainan, China (19°30′N, 109°30′E). Seeds were germinated for 3 d, and stylo seedlings were then transferred to a modified Hoagland nutrient solution containing 250 μM KH2PO4 for 14 d as previously described [71]. After that, seedlings were separately transplanted into nutrient solution supplied with 0, 100 and 250 μM KH2PO4, which were regarded as low (Pi deprivation), moderate and high P supply treatments, respectively. The nutrient solution was adjusted to a pH value of 5.8 and refreshed weekly. After 21 d of P treatments, shoots and roots were separately harvested for further analysis. An individual hydroponic box containing three seedlings of each stylo cultivar was set as one biological replicate. Each treatment included three biological replicates.

Determination of root morphology and P concentration

Plant fresh roots were scanned using an Epson 12000XL scanner (Epson, Japan) with a resolution of 300 dpi, and the obtained image was saved as JEPG format. Total root length, root surface area and root volume were analyzed with WinRhizo Pro software (Regent Instruments Inc., Quebec, Canada). After that, shoot and root samples were oven dried at 75 °C for 7 d, and the dry mass of shoots and roots was further determined. For P concentration analysis, approximately 0.07 g dry samples were burned to ash at 600 °C in a muffle furnace. The sample of ash was absolutely dissolved in 100 mM HCl, and the supernatant was then used for P concentration analysis as previously described [72].

Analysis of APase activity

APase activities in stylo leaf and root were analyzed as previously described [38] with some modification. Approximately 0.15 g of leaf and root samples were ground in 1.5 mL of 45 mM Na-acetate buffer (pH 5.0) at 4 °C. After centrifugation at 12,000 rpm for 15 min at 4 °C, the supernatants were mixed with 2 mL of 45 mM Na-acetate buffer (pH 5.0) containing 1 mM ρ-nitrophenyl phosphate (Sigma, Saint Louis, MO, USA). After incubation at 37 °C for 15 min, the reaction was terminated by the addition of 1 mL of 1 M NaOH. APase activity was spectrophotometrically detected at 405 nm and expressed as micromoles of ρ-nitrophenyl phosphate hydrolyzed per mg protein per min. Protein concentration in the extracts was analyzed using the Coomassie Brilliant Blue staining method [73].

RNA extraction and sequencing

Total RNA from roots of RY2 at 0 (LP) and 250 (HP) μM KH2PO4 treatments was isolated using Trizol reagent (Invitrogen, Carlsbad, CA, USA) following the manufacturer's instructions. RNA purity and integrity were assessed by Nanodrop 2000c Spectrophotometer (Thermo Fisher Scientific, Waltham, MA, USA) and Agilent 2100 (Agilent Technologies, Palo Alto, CA, USA), respectively. RNA sequencing analysis was conducted by Novogene Bioinformatics Technology Co., Ltd. (Bei**g, China). RNA-seq libraries were constructed using the NEBNext® UltraTM RNA Library Prep kit (NEB, Beverly, MA, USA), and the cDNA libraries were sequenced using an Illumina HiseqTM platform (Illumina, San Diego, CA, USA). The 150-bp paired-end reads (PE150) were generated.

Transcriptomic analysis

RNA-seq raw data were obtained using the Casava v.1.8 program. The raw reads in FASTQ format were processed, and then the high-quality clean reads were obtained after removing adaptor, ploy-N and low-quality sequences. The final clean reads were assembled using Trinity software (version 2). For annotation, all assembled unigenes were searched against a number of public databases, including the National Center for Biotechnology Information (NCBI) non-redundant protein sequences (Nr), the non-redundant nucleotide sequences (Nt), the Protein Family (Pfam), the Clusters of Orthologous Groups of protein database (COG), the Swiss-Prot protein database, the GO and the KEGG databases.

The expression level of each gene was analyzed and represented by the expected number of fragments per kilobase of transcript sequence per millions base pairs (FPKM) using RSEM software with default settings [74]. Differentially expressed genes between two P treatments were identified using DESeq2 [75]. Genes with q-value <0.05 and |log2(fold change)| ≥1 were assigned as differentially expressed. GO and KEGG enrichment analyses of DEGs were performed as previously described [71, 76]. The interaction networks were analyzed by Cytoscape (version 3.8.0). The raw data were deposited in the Gene Expression Omnibus under GEO series number GSE171448.

Validation of DEGs by qRT-PCR analysis

A total of 13 DEGs were selected to assess the accuracy of RNA-seq data using qRT-PCR method. qRT-PCR analysis was performed according to SYBR Green Master Mix kit (Vazyme, China), and was monitored on a QuantStudio™ 6 Flex Real-Time System (Thermo Fisher Scientific, Waltham, MA, USA). Specific primers of the tested genes are listed in Additional file 12. The relative expression of candidate gene was calculated relative to the expression of reference gene SgEF-1a as previously described [37]. Three biological replicates were included in the qRT-PCR analysis.

Statistical analysis

Data analysis was performed for the mean and standard error calculation using Microsoft Excel 2003 (Microsoft Company, USA). One-way ANOVA and Student’s t-test analyses were performed with the SPSS program (SPSS Institute, USA, v. 13.0).