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Review

Proteomic Analysis Dissects Molecular Mechanisms Underlying Plant Responses to Phosphorus Deficiency

by
Ming Zhou
1,†,
Shengnan Zhu
2,†,
Xiaohui Mo
1,†,
Qi Guo
1,
Yaxue Li
1,
Jiang Tian
1,* and
Cuiyue Liang
1,*
1
Root Biology Center, State Key Laboratory for Conservation and Utilization of Subtropical Agro-Bioresources, College of Natural Resources and Environment, South China Agricultural University, Guangzhou 510642, China
2
Life Science and Technology School, Lingnan Normal University, Zhanjiang 524048, China
*
Authors to whom correspondence should be addressed.
†
These authors contributed equally to this work.
Cells 2022, 11(4), 651; https://doi.org/10.3390/cells11040651
Submission received: 17 December 2021 / Revised: 2 January 2022 / Accepted: 5 January 2022 / Published: 14 February 2022
(This article belongs to the Special Issue Omics in Plant Genetics and Breeding)
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Abstract

:
Phosphorus (P) is an essential nutrient for plant growth. In recent decades, the application of phosphate (Pi) fertilizers has contributed to significant increases in crop yields all over the world. However, low efficiency of P utilization in crops leads to intensive application of Pi fertilizers, which consequently stimulates environmental pollution and exhaustion of P mineral resources. Therefore, in order to strengthen the sustainable development of agriculture, understandings of molecular mechanisms underlying P efficiency in plants are required to develop cultivars with high P utilization efficiency. Recently, a plant Pi-signaling network was established through forward and reverse genetic analysis, with the aid of the application of genomics, transcriptomics, proteomics, metabolomics, and ionomics. Among these, proteomics provides a powerful tool to investigate mechanisms underlying plant responses to Pi availability at the protein level. In this review, we summarize the recent progress of proteomic analysis in the identification of differential proteins that play roles in Pi acquisition, translocation, assimilation, and reutilization in plants. These findings could provide insights into molecular mechanisms underlying Pi acquisition and utilization efficiency, and offer new strategies in genetically engineering cultivars with high P utilization efficiency.

1. Introduction

Phosphorus (P) is an essential mineral nutrient for plants, accounting for up to 0.5% of plant dry weight depending on plant species [1]. It is not only an indispensable structural constituent of biomolecules, including deoxyribonucleic acid (DNA), proteins, and phospholipids, but also is a key signal factor that functions in mediating the phosphate (Pi)-signaling network [2,3,4,5]. Plants mainly take up P in the form of Pi, including HPO42− and H2PO4−, which is about 0.1–10 μM in soils [6]. Meanwhile, multiple factors exist in soils, limiting Pi availability by directly or indirectly chelating Pi into immobile forms, such as microbial activities and an abundance of cations (e.g., Al3+, Fe2+, Ca2+). For example, in acid soils, Pi easily reacts with Al3+ or Fe2+ and becomes sparingly soluble forms (i.e., Al-P and Fe-P) [6]. Low Pi availability and fluctuation severely limits crop yield by adversely affecting root growth, photosynthesis, respiration, and energy transduction [7,8]. To meet the requirement of P in crops, millions of tons of Pi fertilizers are profligately applied to farms worldwide every year. However, due to low P utilization efficiency in crops, applied Pi fertilizers are either fixed by soil particles or leached into the biosphere, leading to environmental pollution and biodiversity loss [4,9,10]. Therefore, a deeper understanding of the molecular mechanisms regarding plant tolerance to low Pi availability is required to develop cultivars with high Pi fertilizer utilization efficiency, thus ameliorating the sole reliance on excess Pi fertilizer applications to improve crop yield.
In recent decades, adaptive mechanisms underlying plant responses to P deprivation have been extensively investigated through forward and reverse genetic analysis, with the aid of the application of proteomics, transcriptomics, metabolomics, and ionomics [11,12,13,14]. Among these, proteomic analysis, as a powerful tool, has permitted us to identify numerous differentially accumulated proteins (DAPs) in response to Pi availability, which sheds light on molecular responses of plants to Pi starvation at the protein level, especially at protein modification levels (e.g., phosphorylation, succinylation) [13,15]. For example, a set of DAPs was identified at different P levels in many plant species, such as in Arabidopsis (Arabidopsis thaliana) [16,17], maize (Zea mays) [18,19], tomato (Solanum lycopersicum) [20], rice (Oryza sativa) [21,22], barley (Hordeum vulgare) [23], and soybean (Glycine max) [24,25] (Table 1). In this review, we mainly summarize recent advances in the identification and functional characterization of DAPs in response to Pi starvation in plants through proteomic analysis, and highlight the complex regulatory network underlying Pi acquisition, remobilization, and reutilization at protein levels. Meanwhile, we discuss the advantages and challenges of proteomic analysis to shed light on molecular mechanisms underlying plant adaptations to Pi starvation, and thus contribute to developing crop cultivars with high P efficiency in the future.

2. Morphological, Physiological, and Biochemical Responses of Plants to Pi Starvation

Phosphorus deficiency has many deleterious effects on plant growth, as reflected by a significant decrease in plant biomass and yield [4,45,46]. It is well known that plants have evolved a set of morphological, physiological, and molecular strategies to adapt to P deficiency conditions. These adaptive strategies could enhance plant capability to acquire Pi from soils and restrict Pi consumption in plant organisms or increase plant internal Pi reutilization [9,47,48,49]. Among plant morphological responses, modification of root morphology and architecture is generally considered to play a key role in controlling plant Pi acquisition efficiency, such as the formation of shallower root architecture and increases in both lateral root length and root hair density [50,51]. Interestingly, in white lupin and several species of the Proteaceae family, the formation of cluster roots termed “proteoid roots” was observed, which could enhance root–soil contact and thus facilitate plants to acquire more Pi from soils [52,53,54]. Accompanied by changes in root morphology and architecture, physiological and biochemical responses in plant roots could strengthen the capability of roots to mobilize sparingly soluble P in soils, including increased organic acid synthesis and exudation to mobilize inorganic-P forms (e.g., Al-P, Fe-P, Ca-P), and enhance root-associated purple acid phosphatase activity to hydrolyze organic-P forms (e.g., phytate-P, ATP) [9,55,56]. Meanwhile, plants could form beneficial symbiosis with soil microorganisms, such as arbuscular mycorrhizal (AM), to improve Pi acquisition [57,58,59,60,61].
In addition to changes in root traits, Pi availability also significantly triggers leaf/shoot morphology remodeling. For example, leaf angles (i.e., erectness, inclination) and tiller numbers are regulated by P deficiency in rice [62,63,64]. Moreover, accompanied by changes in shoot morphology, a group of physiological and biochemical processes in response to P deficiency was also influenced. For example, leaf color turned from green to purple or dark-green in most plants under low-P conditions, which was mainly caused by anthocyanin accumulations, along with the reduced chlorophyll contents [28,29,65]. Meanwhile, increased starch content and decreased sucrose content in leaves were widely observed in most plants [66,67]. Therefore, identification and functional characterization of the DAPs related to morphological, physiological, and biochemical responses in plants could help us to further improve Pi fertilizer utilization efficiency in crops [68,69].

3. DAPs Reveal Complex Repones of Plants to P Deficiency

3.1. Identification of DAPs in Plant Leaves/Shoots

3.1.1. Differential Proteins Related to Photosynthesis and Carbon Metabolism

Photosynthesis is a light-harvesting process whose intensity is mainly dependent on the generation rate of NADPH and ATP, as well as ribulose-1,5-diphosphate (RuBP) carboxylation [70,71,72]. Reduced accumulations of proteins associated with the photosynthesis process have been observed in plants under P deficiency conditions through differential proteomic analysis, such as proteins related to light energy absorption, electron transfer, and transformation to generate ATP and NADPH. For example, the differential accumulations of a series of proteins were identified, including chlorophyll a/b binding protein in ramie (Boehmeria nivea), ferredoxin (Fdx) in soybean, as well as ferredoxin-nitrite reductase (FNR) and alpha/beta subunit of the ATP synthetase in maize (Figure 1) [18,26,28,29]. Meanwhile, a set of DAPs exhibiting decreased accumulations were found to be involved in rubisco carboxylation reaction (RuBisco), the conversion of ATP and NADPH process, including NADP-malate dehydrogenase (NADP-MDH), pyruvate orthophosphate dikinase (PPDK), phosphoenolpyruvate carboxylase (PEPC), transketolase isoform, RuBisco activase A (RCA), and the large and small subunit of RuBisco [18,26,27,40,41]. Therefore, decreased accumulations of the proteins strongly suggest that Pi starvation could reduce photosynthesis in plants.
A set of proteins closely related to carbon and energy metabolism were found to be up- or down-regulated by Pi starvation in plant leaves (Figure 1). For example, some proteins associated with the glycolysis process were up-regulated, such as hexokinase (HXK) in ramie leaves, fructose-1,6-bisphosphate aldolase (FBP aldoase), glyceraldehyde-3-phosphate dehydrogenase (GAPDH), phosphoglycerate kinase (PGK), and enolase1 in leaves of both maize and Arabidopsis [18,26,43]. Moreover, the increased accumulations of proteins involved the tricarboxylic acid cycle (TCA) were also found to be affected by Pi deprivation in maize leaves, such as succinyl-CoA synthetase (SCS) and succinate dehydrogenase (SDH) [18]. Additionally, the abundance of MDH and isocitrate dehydrogenase (IDH) was found to be down-regulated in leaves of maize and ramie, respectively [26,27]. Meanwhile, other carbon metabolic processes were also influenced by Pi starvation in plants, as reflected by the identification of related DAPs (Figure 1). For example, the 6-phosphogluconolactonase (6-PGLS) and 6-phosphogluconate dehydrogenase (6-PGDG), associated with the pentose phosphate pathway (PPP), were up-regulated under P deficiency conditions in maize and barley [18,23]. In contrast, triose-phosphate isomerase (TPI) and 6-PGDG were down-regulated in leaves of rape and maize, respectively [18,27,41]. These results indicate that carbon and energy metabolic processes, like glycolysis, TCA cycle, and gluconeogenesis, are significantly influenced in plants under low-P conditions.

3.1.2. Pi Starvation Responsive Proteins Related to Remodeling Lipid Membranes

One of the vital adaptive strategies to low Pi availability in plants is to improve internal P utilization efficiency by remodeling lipid membranes [73]. Under low-Pi stress, phospholipids, accounting for 30% of total organophosphate in plants, were replaced by specific non-phosphorous lipids, such as sulfoquinovosyl diacylglycerol (SQDG) [74]. Consistently, a set of DAPs was identified to function in remodeling lipid membranes. For example, in maize and Arabidopsis leaves, increased accumulations of UDP-sulfoquinovose synthase (SQD1) were observed, which were suggested to function in releasing sulfoquinovose for SQDG production [18,27,75]. Meanwhile, increased accumulations of SQDG synthase (SQD2) were also observed under low-P conditions, and SQDs were suggested to accelerate the replacement of phospholipid glycerate (PG) by SQDG in plant cell plasma membranes [17,75]. Additionally, some proteins, including lipid-transfer protein, lipoxygenase, and 3-phosphoglycerate dehydrogenase, were also observed to be up-regulated in Arabidopsis leaves [17]. These results support the hypothesis that changes in the phospholipid metabolism could further accelerate the conversion of intracellular organic P forms, and thus improve P utilization efficiency under low-P conditions.

3.1.3. Proteins Involved Anthocyanin, Polyamine, and Reactive Oxygen Species (ROS) Metabolisms

It is generally believed that increases in anthocyanin synthesis in shoots or leaves could protect plants from ROS damage under low-P conditions [76,77,78,79]. Anthocyanins are products generated from the phenylpropanoid and flavonoid metabolic pathways, regulated by multiple enzymes, such as chalcone synthase (CHS) and dihydroflavonol-4-reductase (DFR) [80]. DAPs functioning in anthocyanin synthesis have been found in Arabidopsis leaves through proteomic analysis, including 4-coumarate:CoA ligase 1 (4-CL1), DFR, leucoanthocyanidin dioxygenase (LDOX), anthocyanidin synthase (ANS), anthocyanidin 5-O-glucosyltransferase (AA5GT), phenylalanine ammonia-lyases (PALs), and glutathione-S-transferase F12 (GSTF12) [17,75] (Figure 1). The results strongly suggest that P deficiency could increase accumulations of enzymes functioning in anthocyanin synthesis, and thus enhance anthocyanin accumulations in leaves.
Polyamines are aliphatic amines and play an important role in plant adaptation to biotic and abiotic stresses [81]. Interestingly, several DAPs controlling polyamines biosynthesis were found in maize leaves under P deficiency conditions, such as spermidine synthase (SPDS) and arginine decarboxylase (ADC) [18]. Although the detailed functions of polyamines in plant adaptation to Pi starvation remained fragmentary, identification of DAPs mediating its biosynthesis strongly suggests that polyamines could participate in plant adaption to P deficiency, which merits further study.
Phosphorus deficiency also leads to the elevation of ROS in plant leaves [82]. Therefore, increased accumulations of antioxidation-related proteins were generally observed in plant leaves under P deficiency conditions, including glutathione S-transferase (GST) in maize and barley [18,23], ascorbate peroxidase (APX) in maize [18], 2-cysteine peroxiredoxin B in ramie [26], non-specific lipid-transfer protein 1, and extra-large G protein 3 in tomato [20], as well as proteins belonging to peroxidase superfamily in Arabidopsis [17]. Increased accumulations of these proteins may help plants against ROS damage under low-P stress.

3.2. Identification of DAPs in Plant Roots

Recently, DAPs controlling root responses to P deficiency have been identified in different plants, such as soybean [25], rape [39], and maize [19]. Functions of the DAPs have been suggested to involve a series of adaptive strategies in plant roots, including changes in root development, regulating organic acid synthesis and secretion, increasing activities of root-associated purple acid phosphatases (PAPs). Furthermore, several studies were conducted to elucidate complex responses of protein modifications to P deficiency in plant roots, especially for protein phosphorylation and succinylation.

3.2.1. Pi Starvation-Responsive Proteins Participated in Root System Remodeling

It is well known that root architecture remodeling involves changes in hormones, such as auxin, ethylene, cytokinin (CK), or jasmonic acid (JA), which react differently but coordinately regulate root growth [83,84,85,86]. Thus, a set of DAPs was found to involve hormone synthesis, transport, and distribution in root response to P deficiency [19,31,32,33,34,35] (Figure 2). For example, in several plant species, such as rice and masson pine (Pinus massoniana), comparative proteomic studies were conducted to find that abundance of several ethylene-precursor synthetases, including 1-aminocyclopropane-1-carboxylate oxidase (ACCO) and SAMS, was up-regulated by P deficiency, even though ACCO and SAMS were down-regulated in maize and Arabidopsis, respectively [30,33,34,42]. The results strongly suggest that ethylene participated in regulating root growth under P-deprivation conditions, which was probably attributed to changes in its biosynthesis [87,88]. Consistently, in Arabidopsis, the number of lateral roots was significantly higher in aco1-1 mutant compared to that in wild-type, strongly suggesting that ethylene could regulate lateral root development [89]. In addition to ethylene, auxin is considered to play a key role in mediating the development of lateral root and root hair under Pi-starvation conditions [85,90]. Consistently, a set of DAPs for the auxin signaling pathway was observed to be up-regulated in Arabidopsis and maize, including auxin-response proteins (ARFs), phosphatase 2A (PP2A), and nonspecific phospholipase C4 (NPC4) [16,19,32,41]. Furthermore, overexpression of ZmPP2AA1, encoding a subunit of phosphatase 2A, significantly increased the lateral root density and promoted root growth under P-deficiency conditions in maize, strongly suggesting that auxin could regulate lateral root development [91,92]. Additionally, root growth in response to P deficiency was also found to be mediated by the JA pathway, supported by the evidence that allene oxide synthase (AOS), allene oxide cyclase 1 (AOC1), and jacalin-related lections (e.g., JAL5/23/31) for controlling JA metabolism were up-regulated in Arabidopsis under low-P conditions [17,75]. Interestingly, knockout of atwrky6-1, a mediator in the Pi-signaling network, led to decreases in lateral root formation, but increased accumulations of AOS, strongly suggesting the JA metabolism might participate in regulating Arabidopsis root growth [17,93]. Meanwhile, decreased accumulations of tRNA isopentenyl transferase (tRNA IPT) and beta-glucosidease (BGL) involved CK synthesis were observed in long-term P-deficient maize roots, suggesting that CK could regulate plant root growth [31,32]. Moreover, several studies reported that the application of CK could disturb the expression of PIN genes and prevent the formation of the auxin gradient, which is required for lateral root primordia development. In contrast, the biosynthesis of CK was rapidly suppressed by auxin via the isopenteneyadenosine-50-monophosphate independent pathway [94,95,96]. Several studies also reported that ethylene could regulate root growth via stimulating auxin biosynthesis and transport in root apex, but DAPs involved in both auxin- and ethylene-signaling pathways were scarcely identified [96,97,98]. These results suggest that P deficiency-induced changes in root morphology and architecture may be modulated through sophisticated interactions among different phytohormones.
In addition to the identification of phytohormone-related DAPs, a set of DAPs involved in cell cycle, division, and expansion processes was also suggested to regulate root architecture plasticity in response to low-P stress [99,100] (Figure 2). For example, translationally controlled tumor protein (TCTP), functioning in fundamental biological processes (e.g., mitotic, cell proliferation, cytoprotective, and anti-apoptotic), was observed to be up-regulated by Pi starvation in Arabidopsis [40,101,102]. Further genetic analysis showed that reduced AtTCTP1 expression could result in significant inhibition of lateral root formation [103]. Additionally, accumulations of cell proliferation-related proteins, including GTP-binding nuclear protein RAN-B1 (Ran GTPase), cell division cycle protein 48 (CDC48), and mini-chromosome maintenance protein 6 (MCM6), were increased by Pi starvation in maize [19,31]. Contrastingly, an increased abundance of actin and tubulin was observed in plants in response to low-P stress in plants, including soybean, maize, rice, and barley, while decreased abundance was observed in Arabidopsis [19,23,31,35,36]. Interestingly, a loss of rmd mutant, encoding a rice actin-binding protein, exhibited steeper growth angles for crown roots in rice, strongly suggesting that cell proliferation-related proteins could play a role in root growth [104].

3.2.2. Pi Starvation-Responsive Proteins Related to Root Exudates

Plant root exudates mainly include a class of metabolites and proteins with low- or high-molecular weight, such as organic acids (OAs), sugars, flavonoids, and phosphatases [100,105]. It is well known that root exudates play crucial roles in increasing Pi uptake by desorbing immobile P in soils [100,106]. Consistently, differential proteomic analysis led to identifying a set of DAPs controlling root exudates, which contributes to deepening understandings of the molecular mechanisms underlying rhizosphere P mobilization [100,105] (Figure 2). For example, increased OA exudation was found to be associated with activities of several differential proteins related to OA synthesis, including citrate synthase (CS) in Arabidopsis, maize and soybean, IDH in maize, MDH in rape, soybean, and maize, as well as decreased abundance of aconitase in rice [21,30,32,35,41]. Consistently, overexpression of CS from Daucus carota (DcCS) improved Arabidopsis growth under P deficiency conditions due to enhanced citrate synthesis and exudation from the roots [107]. These results strongly indicate that increased OA synthesis and exudation play an important role for plants when scavenging Pi in soils.
In addition to OA synthesis and exudation, some secreted proteins, such as purple acid phosphatases (PAPs) and ribonucleases, are critical for plants to utilize organic-P pools in soils. Among PAPs, root-associated PAPs were widely found to be involved in the activation and utilization of extracellular organic-P sources, such as ATP, deoxy-ribonucleotide triphosphate (dNTPs), and phytate-P [56,108]. For example, several PAPs were purified and functionally characterized to mediate organic-P utilization, such as SgPAP23 in stylo (stylosanthes guianensis), GmPAP7a/b in soybean, OsPAP10a in rice, and LeSAP1/2 in tomato [25,47,109,110,111]. Meanwhile, a variety of low P-induced PAPs was identified in plants through proteomic analysis, such as AtPAP12/26 in Arabidopsis, GmPAP1-like, and GmPAP22-like proteins in soybean [25,44]. In Arabidopsis, atpap12/atpap26 mutant exhibited impaired growth coupling the decreases in root secretory APase activity and total Pi concentration in rosettes [112,113]. Overexpressing GmPAP1-like enhanced Arabidopsis growth and P content when dNTP was supplied as the sole external P source [25]. In addition, some DAPs functions in cell wall carbohydrate metabolism were also identified to be up-regulated in Arabidopsis suspension cells or root cell walls under P-deficient conditions, such as polygalacturonase (PG) and xyloglucan endotransglycosylase (XET), which was suggested to release cell wall-bound Pi under low-P conditions [25,112,114]. Additionally, since changes in polysaccharides in cell wall metabolism could influence the rhizosphere size, which was suggested to be positively correlated with P accumulation in plants, DAPs related to cell wall metabolism might affect plant P efficiency partially through changing rhizosphere size [115].

3.2.3. Response of Symbiotic Association to Pi Starvation in Plants

Although it is well known that the formation of a symbiotic association between plants and arbuscular mycorrhiza (AM) is a typical plant strategy in response to P deficiency, no proteomic analysis was conducted to identify DAPs in the symbiotic association at different P levels. However, several studies have highlighted that Pi acquisition efficiency in legume crops could be improved through formation of the symbiotic association between roots and rhizobia, which can directly fix atmosphere N2 in nodules and supply nitrogen for plant growth. For example, rhizobium inoculation led to enhancing the capability of soybean in utilization of sparingly soluble P forms (e.g., Ca-P, Al-P, Fe-P), as reflected by higher P content and plant biomass [116]. Furthermore, overexpression of two nodule preferring Pi transporters, GmPT7 and GmPT5 promoted soybean nodulation, P content, as well as fresh weight [117,118]. To date, only one study has elucidated differential protein profiles in soybean nodules at two P levels through two-dimensional electrophoresis combined with matrix-assisted laser desorption ionization (MALDI)-time of flight (TOF)/TOF mass spectrometry (MS) analysis. A total of 44 DAPs were identified in soybean nodules, which were mainly involved in stress response and carbon and amino acid metabolism, strongly indicating that a complex but precise module exists in soybean nodules to sustain mutualistic symbiosis to maximize Pi uptake [24]. Among them, GmMDH12 was found to be up-regulated in soybean nodules by Pi starvation, which was consistent with the increased malate concentration in nodules at low-P levels. Furthermore, overexpression of GmMDH12 significantly increased malate concentrations and inhibited nodule size in soybean, strongly suggesting that Pi-starvation increased malate, which might have multiple functions except for the enhancement of malate exudation and the activation of sparingly soluble P in soils [119].

4. Identification of DAPs Exhibiting Post-Transcriptional Modifications

Post-transcriptional modifications (PTMs) play crucial roles in regulating protein activity, longevity, and localization, including phosphorylation, ubiquitination, succinylation, glycosylation, and SUMOylation modification [15,38,120]. Recently, several studies were conducted to investigate changes in protein profiles with PTMs in plant roots at different P levels, such as phosphoproteome in soybean, maize, and rice [15,22,38]. In soybean roots, a total of 427 phosphoproteins were found to be regulated by Pi starvation, including 213 up-regulated and 214 down-regulated [15]. For example, auxin efflux transporter GmPIN2 and ethylene-insensitive protein GmEIN2 were identified to be up-regulated by P deprivation, strongly suggesting participation of auxin and ethylene in modulating root architecture in response to P deficiency [15]. Meanwhile, in rice roots, a total of 554 phosphoproteins were found to exhibit differential accumulations, with 546 down-regulated and 8 up-regulated proteins. Several proteins, including four mitogen-activated protein kinases (MAPKs), five calcium-dependent protein kinases (CDPKs), and OsCK2β3, were observed to be decreased in response to Pi starvation, especially for OsCK2, which was suggested to regulate phosphorylation of OsPT2 and OsPT8 and thus influence their subcellular localization [22,121]. Meanwhile, in maize roots, about 51 phosphoprotein spots were observed to exhibit differential accumulations by low-P treatment; these were involved in a group of cellular and metabolic pathways, such as signal transduction and carbon metabolism [32]. Among them, one auxin receptor, ABP1, was found to be up-regulated in maize roots under low-P conditions [32]. Recently, overexpression of AtABP1 has been found to influence primary root growth, root bending, and lateral root development [122].
In addition to phosphorylation analysis, succinylated-proteomic analysis was also conducted to identify differential proteins with succinylation in barley roots under P-deficiency conditions [38]. A total of 120 succinylation sites were identified across 83 proteins, including 79 increased and 4 decreased succinylated proteins at 48 h of Pi-starvation treatment, respectively [38]. Moreover, these differentially succinylated proteins were enriched in ribosome pathways, glycolysis, and RNA degradation pathways. For example, 60S/50/40S/30 ribosomal protein was suggested to involve ribosome pathways, while TPI, GAPDH, and FBP aldolase were predicted to participate in glycolysis pathways [38]. However, functions for most of the identified proteins with succinylation remain unknown, which merits further study.

5. Perspectives

In recent decades, with the aid of proteomic analysis, identification and functional characterization of DAPs have opened the way to elucidate molecular mechanisms underlying plant adaptation to Pi starvation. For example, the increased abundances of SPX1, PHT1;4, and PHF1 proteins were observed in Arabidopsis roots, and other proteins, such as TaPHT1;9-4B, TaPHT1;3-5B, and TaPHT1;6-5B, were also observed in wheat roots, acting as the main proteins mediating Pi homeostasis [16,37]. However, the functions of most of the DAPs remain largely unknown and require furthered investigation via reverse and forward genetic analysis. For example, a histone chaperone (nap1;2) was found to be up-regulated under Pi starvation via proteomic analysis in Arabidopsis, and its functions in maintaining Pi homeostasis were clarified via analysis of triple nap1;1 nap1;2 nap1;3 mutant lines [13]. Meanwhile, although the method of differential proteomics based on mass spectrometry was established and successfully used, there is still a lack of a more effective method for protein separation and identification in order to qualitatively analyze plant proteins. In order to solve these problems, multiple technologies should be combined to develop a more effective method for protein extraction and separation. Meanwhile, in addition to commonly used quantitative techniques, such as label-free quantitation [123], isobaric tags for relative and absolute quantitation (iTRAQ) [124], tandem mass tags (TMT) [125], and stable isotope labeling by amino acids in cell culture (SILAC) [126], application of a relatively new technique, termed sequential window acquisition of all theoretical mass spectra (SWATH) [127], together with data-independent acquisition (DIA) [128], might be very helpful to identify more DAPs. Furthermore, the vigorous development and wide application of new technologies (e.g., microfluidic chip, reverse micelles, magnetic nanoparticles) will break the bottleneck of protein separation, enrichment, and detection in single cells and different organelles [129].
Although many questions remain to be solved, we believe that the application of proteomics integrating multiple omics, as well as genetic analysis, will help us to elucidate molecular mechanisms underlying plant adaptation to P deficiency and develop cultivars with high P efficiency in the future.

Author Contributions

M.Z. and S.Z. wrote the manuscript; J.T. and C.L. conceived the idea; M.Z., Q.G. and Y.L. prepared tables and figures; J.T., C.L., M.Z., S.Z. and X.M. reviewed the manuscript and provided input; J.T. and C.L. provided suggestions to improve the manuscript, tables, and figures. All authors have read and agreed to the published version of the manuscript.

Funding

This work was supported by National Key Research and Development Program of China (2021YFF1000500), the Major Program of Guangdong Basic and Applied Research (2019B030302006), National Natural Science Foundation of China (32172658 and 32172659), the Natural Science Foundation of Guangdong Province (2021A1515010826).

Institutional Review Board Statement

Not applicable.

Informed Consent Statement

Not applicable.

Conflicts of Interest

The authors declare that there is no conflict of interests regarding the publication of this manuscript.

References

  1. Cashikar, A.G.; Kumaresan, R.; Rao, N.M. Biochemical characterization and subcellular localization of the red kidney bean purple acid phosphatase. Plant Physiol. 1997, 114, 907–915. [Google Scholar] [CrossRef] [Green Version]
  2. Raghothama, K.G. Phosphate acquisition. Annu. Rev. Plant Physiol. Plant Mol. Biol. 1999, 50, 665–693. [Google Scholar] [CrossRef] [PubMed]
  3. Plaxton, W.C.; Shane, M.W. The role of post-translational enzyme modifications in the metabolic adaptations of phosphorus-deprived plants. Annu. Plant Rev. 2015, 48, 99–123. [Google Scholar]
  4. Oldroyd, G.; Leyser, O. A plant’s diet, surviving in a variable nutrient environment. Science 2020, 368, eaba0196. [Google Scholar] [CrossRef] [PubMed]
  5. Ham, B.K.; Chen, J.; Yan, Y.; Lucas, W.J. Insights into plant phosphate sensing and signaling. Curr. Opin. Biotechnol. 2018, 49, 1–9. [Google Scholar] [CrossRef]
  6. Hinsinger, P. Bioavailability of soil inorganic P in the rhizosphere as affected by root-induced chemical changes: A review. Plant Soil 2001, 237, 173–195. [Google Scholar] [CrossRef]
  7. Medici, A.; Szponarski, W.; Dangeville, P.; Safi, A.; Dissanayake, I.M.; Saenchai, C.; Emanuel, A.; Rubio, V.; Lacombe, B.; Ruffel, S.; et al. Identification of molecular integrators shows that nitrogen actively controls the phosphate starvation response in plants. Plant Cell 2019, 31, 1171–1184. [Google Scholar] [CrossRef] [PubMed]
  8. Cho, H.; Bouain, N.; Zheng, L.; Rouached, H. Plant resilience to phosphate limitation: Current knowledge and future challenges. Crit. Rev. Biotechnol. 2021, 41, 63–71. [Google Scholar] [CrossRef]
  9. Lambers, H.; Finnegan, P.M.; Jost, R.; Plaxton, W.C.; Shane, M.W.; Stitt, M. Phosphorus nutrition in Proteaceae and beyond. Nat. Plants 2015, 1, 15109. [Google Scholar] [CrossRef] [PubMed]
  10. Hallama, M.; Pekrun, C.; Lambers, H.; Kandeler, E. Hidden miners-the roles of cover crops and soil microorganisms in phosphorus cycling through agroecosystems. Plant Soil 2019, 434, 7–45. [Google Scholar] [CrossRef] [Green Version]
  11. Abdelrahman, M.; El-Sayed, M.A.; Hashem, A.; Abd, A.E.; Alqarawi, A.A.; Burritt, D.J.; Tran, L.P. Metabolomics and transcriptomics in legumes under phosphate deficiency in relation to nitrogen fixation by root nodules. Front. Plant Sci. 2018, 9, 922. [Google Scholar] [CrossRef] [Green Version]
  12. Deng, Q.; Luo, X.; Chen, Y.; Zhou, Y.; Zhang, F.; Hu, B.; Xie, K. Transcriptome analysis of phosphorus stress responsiveness in the seedlings of Dongxiang wild rice (Oryza rufipogon Griff.). Biol. Res. 2018, 51, 7. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  13. Iglesias, J.; Trigueros, M.; Rojas-Triana, M.; Fernández, M.; Albar, J.P.; Bustos, R.; Paz-Ares, J.; Rubio, V. Proteomics identifies ubiquitin-proteasome targets and new roles for chromatin-remodeling in the Arabidopsis response to phosphate starvation. J. Proteom. 2013, 94, 1–22. [Google Scholar] [CrossRef]
  14. Watanabe, T.; Urayama, M.; Shinano, T.; Okada, R.; Osaki, M. Application of ionomics to plant and soil in fields under long-term fertilizer trials. Springerplus 2015, 4, 781. [Google Scholar] [CrossRef] [PubMed]
  15. Jiang, W.; He, P.; Zhou, M.; Lu, X.; Chen, K.; Liang, C.; Tian, J. Soybean responds to phosphate starvation through reversible protein phosphorylation. Plant Physiol. Biochem. 2021, 167, 222–234. [Google Scholar] [CrossRef] [PubMed]
  16. Lan, P.; Li, W.; Schmidt, W. Complementary proteome and transcriptome profiling in phosphate-deficient Arabidopsis roots reveals multiple levels of gene regulation. Mol. Cell. Proteom. 2012, 11, 1156–1166. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  17. Wang, Z.Q.; Zhou, X.; Dong, L.; Guo, J.; Chen, Y.; Zhang, Y.; Wu, L.; Xu, M. iTRAQ-based analysis of the Arabidopsis proteome reveals insights into the potential mechanisms of anthocyanin accumulation regulation in response to phosphate deficiency. J. Proteom. 2018, 184, 39–53. [Google Scholar] [CrossRef] [PubMed]
  18. Zhang, K.; Liu, H.; Tao, P.; Chen, H. Comparative proteomic analyses provide new insights into low phosphorus stress responses in maize leaves. PLoS ONE 2014, 9, e98215. [Google Scholar]
  19. Li, K.; Xu, C.; Li, Z.; Zhang, K.; Yang, A.; Zhang, J. Comparative proteome analyses of phosphorus responses in maize (Zea mays L.) roots of wild-type and a low-P-tolerant mutant reveal root characteristics associated with phosphorus efficiency. Plant J. 2008, 55, 927–939. [Google Scholar] [CrossRef] [PubMed]
  20. Muneer, S.; Jeong, B.R. Proteomic analysis provides new insights in phosphorus homeostasis subjected to Pi (inorganic phosphate) starvation in tomato plants (Solanum lycopersicum L.). PLoS ONE 2015, 10, e0134103. [Google Scholar]
  21. Sepideh, T.; Matthias, W.; Manzar, H.; Mohammad-Reza, N.; Gilany, K.; Mohammad-Reza, H.; Mansoor, O.; Yazdi-Samadi, B.; Abdelbagi, M.I. A comparative proteome approach to decipher the mechanism of rice adaptation to phosphorous deficiency. Proteomics 2009, 9, 159–170. [Google Scholar]
  22. Yang, J.; Xie, M.; Yang, X.; Liu, B.; Lin, H. Phosphoproteomic profiling reveals the importance of CK2, MAPKs and CDPKs in response to phosphate starvation in rice. Plant Cell Physiol. 2019, 60, 2785–2796. [Google Scholar] [CrossRef] [PubMed]
  23. Nadira, U.A.; Ahmed, I.M.; Zeng, J.; Wu, F.; Zhang, G. Identification of the differentially accumulated proteins associated with low phosphorus tolerance in a Tibetan wild barley accession. J. Plant Physiol. 2016, 198, 10–22. [Google Scholar] [CrossRef] [PubMed]
  24. Chen, Z.; Cui, Q.; Liang, C.; Sun, L.; Tian, J.; Liao, H. Identification of differentially expressed proteins in soybean nodules under phosphorus deficiency through proteomic analysis. Proteomics 2011, 11, 4648–4659. [Google Scholar] [CrossRef]
  25. Wu, W.; Lin, Y.; Liu, P.; Chen, Q.; Tian, J.; Liang, C. Association of extracellular dNTP utilization with a GmPAP1-like protein identified in cell wall proteomic analysis of soybean roots. J. Exp. Bot. 2018, 69, 603–617. [Google Scholar] [CrossRef] [Green Version]
  26. Deng, G.; Liu, L.J.; Zhong, X.Y.; Lao, C.Y.; Wang, H.Y.; Wang, B.; Zhu, C.; Shah, F.; Peng, D.X. Comparative proteome analysis of the response of ramie under N, P and K deficiency. Planta 2014, 239, 1175–1186. [Google Scholar] [CrossRef] [PubMed]
  27. Zhang, K.; Liu, H.; Song, J.; Wu, W.; Li, K.; Zhang, J. Physiological and comparative proteome analyses reveal low-phosphate tolerance and enhanced photosynthesis in a maize mutant owing to reinforced inorganic phosphate recycling. BMC Plant Biol. 2016, 16, 129. [Google Scholar] [CrossRef] [Green Version]
  28. Chu, S.; Li, H.; Zhang, X.; Yu, K.; Chao, M.; Han, S.; Zhang, D. Physiological and proteomics analyses reveal low-phosphorus stress affected the regulation of photosynthesis in soybean. Int. J. Mol. Sci. 2018, 19, 1688. [Google Scholar] [CrossRef] [Green Version]
  29. Cheng, L.; Min, W.; Li, M.; Zhou, L.; Hsu, C.; Yang, X.; Jiang, X.; Ruan, Z.; Zhong, Y.; Wang, Z.; et al. Quantitative proteomics reveals that GmENO2 proteins are involved in response to phosphate starvation in the leaves of Glycine max L. Int. J. Mol. Sci. 2021, 22, 920. [Google Scholar] [CrossRef]
  30. Chevalier, F.; Rossignol, M. Proteomic analysis of Arabidopsis thaliana ecotypes with contrasted root architecture in response to phosphate deficiency. J. Plant Physiol. 2011, 168, 1885–1890. [Google Scholar] [CrossRef]
  31. Li, K.; Xu, C.; Zhang, K.; Yang, A.; Zhang, J. Proteomic analysis of roots growth and metabolic changes under phosphorus deficit in maize (Zea mays L.) plants. Proteomics 2007, 7, 1501–1512. [Google Scholar] [CrossRef]
  32. Li, K.; Xu, C.; Fan, W.; Zhang, H.; Hou, J.; Yang, A.; Zhang, K. Phosphoproteome and proteome analyses reveal low-phosphate mediated plasticity of root developmental and metabolic regulation in maize (Zea mays L.). Plant Physiol. Biochem. 2014, 83, 232–242. [Google Scholar] [CrossRef]
  33. Jiang, H.; Zhang, J.; Han, Z.; Yang, J.; Ge, C.; Wu, Q. Revealing new insights into different phosphorus-starving responses between two maize (Zea mays) inbred lines by transcriptomic and proteomic studies. Sci. Rep. 2017, 7, 44294. [Google Scholar] [CrossRef] [Green Version]
  34. Kim, S.G.; Wang, Y.; Lee, C.H.; Mun, B.G.; Kim, P.J.; Lee, S.Y.; Kim, Y.C.; Kang, K.Y.; Rakwal, R.; Agrawal, G.K.; et al. A comparative proteomics survey of proteins responsive to phosphorous starvation in roots of hydroponically-grown rice seedlings. J. Korean Soc. Appl. Biol. Chem. 2011, 54, 667–677. [Google Scholar] [CrossRef]
  35. Vengavasi, K.; Pandey, R.; Abraham, G.; Yadav, R. Comparative analysis of soybean root proteome reveals molecular basis of differential carboxylate efflux under low phosphorus stress. Genes 2017, 8, 341. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  36. Zhao, H.; Yang, A.; Kong, L.; Xie, F.; Wang, H.; Ao, X. Proteome characterization of two contrasting soybean genotypes in response to different phosphorus treatments. AoB Plants 2021, 13, plab019. [Google Scholar] [CrossRef] [PubMed]
  37. Wang, P.; Li, G.; Li, G.; Yuan, S.; Wang, C.; Xie, Y.; Guo, T.; Kang, G.; Wang, D. TaPHT1;9-4B and its transcriptional regulator TaMYB4-7D contribute to phosphate uptake and plant growth in bread wheat. New Phytol. 2021, 231, 1968–1983. [Google Scholar] [CrossRef] [PubMed]
  38. Wang, J.; Ma, Z.; Li, C.; Ren, P.; Yao, L.; Li, B.; Meng, Y.; Ma, X.; Si, E.; Yang, K.; et al. Dynamic responses of barley root succinyl-proteome to short-term phosphate starvation and recovery. Front. Plant Sci. 2021, 12, 649147. [Google Scholar] [CrossRef]
  39. Chen, S.; Luo, Y.; Ding, G.; Xu, F. Comparative analysis of Brassica napus plasma membrane proteins under phosphorus deficiency using label-free and MaxQuant-based proteomics approaches. J. Proteom. 2016, 133, 144–152. [Google Scholar] [CrossRef] [PubMed]
  40. Li, L.; Huang, L.; Pan, G.; Liu, L.; Wang, X.; Lu, L. Identifying the genes regulated by AtWRKY6 using comparative transcript and proteomic analysis under phosphorus deficiency. Int. J. Mol. Sci. 2017, 18, 1046. [Google Scholar] [CrossRef] [Green Version]
  41. Yao, Y.; Sun, H.; Xu, F.; Zhang, X.; Liu, S. Comparative proteome analysis of metabolic changes by low phosphorus stress in two Brassica napus genotypes. Planta 2011, 233, 523–537. [Google Scholar] [CrossRef]
  42. Fan, F.H.; Ding, G.J.; Wen, X.P. Proteomic analyses provide new insights into the responses of Pinus massoniana seedlings to phosphorus deficiency. Proteomics 2016, 16, 504–515. [Google Scholar] [CrossRef]
  43. Tran, H.T.; Plaxton, W.C. Proteomic analysis of alterations in the secretome of Arabidopsis thaliana suspension cells subjected to nutritional phosphate deficiency. Proteomics 2008, 8, 4317–4326. [Google Scholar] [CrossRef]
  44. Mehta, D.; Ghahremani, M.; Pérez Fernández, M.; Tan, M.; Schläpfer, P.; Plaxton, W.C.; Uhrig, R.G. Phosphate and phosphite have a differential impact on the proteome and phosphoproteome of Arabidopsis suspension cell cultures. Plant J. 2021, 105, 924–941. [Google Scholar] [CrossRef]
  45. Uzokwe, V.N.E.; Asafo-Adjei, B.; Fawole, I.; Abaidoo, R.; Odeh, I.O.A.; Ojo, D.K.; Dashiell, K.; Sanginga, N. Generation mean analysis of phosphorus-use efficiency in freely nodulating soybean crosses grown in low-phosphorus soil. Plant Breed. 2017, 136, 139–146. [Google Scholar] [CrossRef] [Green Version]
  46. Sims, L.; Pastor, J.; Lee, T.; Dewey, B. Nitrogen, phosphorus and light effects on growth and allocation of biomass and nutrients in wild rice. Oecologia 2012, 170, 65–76. [Google Scholar] [CrossRef] [PubMed]
  47. Tian, J.; Wang, C.; Zhang, Q.; He, X.; Whelan, J.; Shou, H. Overexpression of OsPAP10a, a root-associated acid phosphatase, increased extracellular organic phosphorus utilization in rice. J. Integr. Plant Biol. 2012, 54, 631–639. [Google Scholar] [CrossRef]
  48. Liang, C.; Wang, J.; Zhao, J.; Tian, J.; Liao, H. Control of phosphate homeostasis through gene regulation in crops. Curr. Opin. Plant Biol. 2014, 21, 59–66. [Google Scholar] [CrossRef] [PubMed]
  49. Dissanayaka, D.M.S.B.; Nishida, S.; Tawaraya, K.; Wasaki, J. Organ-specific allocation pattern of acquired phosphorus and dry matter in two rice genotypes with contrasting tolerance to phosphorus deficiency. Soil Sci. Plant Nutr. 2018, 64, 282–290. [Google Scholar] [CrossRef]
  50. Lopez-Bucio, J.; Cruz-Ramirez, A.; Herrera-Estrella, L. The role of nutrient availability in regulating root architecture. Curr. Opin. Plant Biol. 2003, 6, 280–287. [Google Scholar] [CrossRef]
  51. Peret, B.; Desnos, T.; Jost, R.; Kanno, S.; Berkowitz, O.; Nussaume, L. Root architecture responses: In search of phosphate. Plant Physiol. 2014, 166, 1713–1723. [Google Scholar] [CrossRef] [Green Version]
  52. Horsham, V. Proteoid root morphology and function in lupinus albus. Plant Soil 1981, 60, 143–147. [Google Scholar]
  53. Neumann, G.; Massonneau, A.; Langlade, N.; Dinkelaker, B.; Hengeler, C.; Roemheld, V.; Martinoia, E. Physiological aspects of cluster root function and development in phosphorus-deficient white lupin (Lupinus albus L.). Ann. Bot. 2000, 85, 909–919. [Google Scholar] [CrossRef]
  54. Lambers, H.; Finnegan, P.M.; Laliberte, E.; Pearse, S.J.; Ryan, M.H.; Shane, M.W.; Veneklaas, E.J. Update on phosphorus nutrition in proteaceae. Phosphorus nutrition of proteaceae in severely phosphorus-impoverished soils: Are there lessons to be learned for future crops? Plant Physiol. 2011, 156, 1058–1066. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  55. Chen, Z.C.; Liao, H. Organic acid anions: An effective defensive weapon for plants against aluminum toxicity and phosphorus deficiency in acidic soils. J. Genet. Genom. 2016, 43, 631–638. [Google Scholar] [CrossRef] [PubMed]
  56. Dissanayaka, D.; Plaxton, W.C.; Lambers, H.; Siebers, M.; Marambe, B.; Wasaki, J. Molecular mechanisms underpinning phosphorus-use efficiency in rice. Plant Cell Environ. 2018, 41, 1483–1496. [Google Scholar] [CrossRef] [PubMed]
  57. Tarafdar, J.C.; Marchner, H. Efficiency of VAM hyphae in utilisation of organic phosphorus by wheat plants. Soil Sci. Plant Nutr. 1994, 40, 593–600. [Google Scholar] [CrossRef]
  58. Sa, T.M.; Israel, D.W. Energy status and functioning of phosphorus-deficient soybean nodules. Plant Physiol. 1991, 97, 928–935. [Google Scholar] [CrossRef] [Green Version]
  59. Drevon, J.; Hartwig, U.A. Phosphorus deficiency increases the argon-induced decline of nodule nitrogenase activity in soybean and alfalfa. Planta 1997, 201, 463–469. [Google Scholar] [CrossRef]
  60. Hodge, A.; Berta, G.; Doussan, C.; Merchan, F.; Crespi, M. Plant root growth, architecture and function. Plant Soil 2009, 321, 153–187. [Google Scholar] [CrossRef]
  61. Hiruma, K.; Gerlach, N.; Sacristan, S.; Nakano, R.T.; Hacquard, S.; Kracher, B.; Neumann, U.; Ramirez, D.; Bucher, M.; O’Connell, R.J.; et al. Root endophyte colletotrichum tofieldiae confers plant fitness benefits that are phosphate status dependent. Cell 2016, 165, 464–474. [Google Scholar] [CrossRef] [Green Version]
  62. Umehara, M.; Hanada, A.; Magome, H.; Takeda-Kamiya, N.; Yamaguchi, S. Contribution of strigolactones to the inhibition of tiller bud outgrowth under phosphate deficiency in rice. Plant Cell Physiol. 2010, 51, 1118–1126. [Google Scholar] [CrossRef] [Green Version]
  63. Mghase, J.J.; Shiwachi, H.; Takahashi, H.; Irie, K. Nutrient deficiencies and their symptoms in upland rice. J. ISSAAS 2011, 17, 59–67. [Google Scholar]
  64. Ruan, W.; Guo, M.; Xu, L.; Wang, X.; Zhao, H.; Wang, J.; Yi, K. An SPX-RLI1 module regulates leaf inclination in response to phosphate availability in rice. Plant Cell 2018, 30, 853–870. [Google Scholar] [CrossRef] [Green Version]
  65. Mo, X.; Zhang, M.; Zhang, Z.; Lu, X.; Liang, C.; Tian, J. Phosphate (Pi) starvation up-regulated GmCSN5A/B participates in anthocyanin synthesis in Soybean (Glycine max) dependent on Pi availability. Int. J. Mol. Sci. 2021, 22, 12348. [Google Scholar] [CrossRef] [PubMed]
  66. Ticconi, C.A.; Abel, S. Short on phosphate: Plant surveillance and countermeasures. Trends Plant Sci. 2004, 9, 548–555. [Google Scholar] [CrossRef] [PubMed]
  67. Yuan, H.; Liu, D. Signaling components involved in plant responses to phosphate starvation. J. Integr. Plant Biol. 2008, 50, 849–859. [Google Scholar] [CrossRef]
  68. Liang, C.; Tian, J.; Liao, H. Proteomics dissection of plant responses to mineral nutrient deficiency. Proteomics 2013, 13, 624–636. [Google Scholar] [CrossRef] [PubMed]
  69. Motte, H.; Vanneste, S.; Beeckman, T. Molecular and environmental regulation of root development. Annu. Rev. Plant Biol. 2019, 70, 465–488. [Google Scholar] [CrossRef] [Green Version]
  70. Noctor, G.; Foyer, C.H. Homeostasis of adenylate status during photosynthesis in a fluctuating environment. J. Exp. Bot. 2000, 51, 347–356. [Google Scholar] [CrossRef] [Green Version]
  71. Rochaix, J.D. Role of thylakoid protein kinases in photosynthetic acclimation. FEBS Lett. 2007, 581, 2768–2775. [Google Scholar] [CrossRef]
  72. Carmo-Silva, E.; Scales, J.C.; Madgwick, P.J.; Parry, M.A. Optimizing Rubisco and its regulation for greater resource use efficiency. Plant Cell Environ. 2015, 38, 1817–1832. [Google Scholar] [CrossRef] [PubMed]
  73. Nakamura, Y. Phosphate starvation and membrane lipid remodeling in seed plants. Prog. Lipid Res. 2013, 52, 43–50. [Google Scholar] [CrossRef] [PubMed]
  74. Preiss, J. Bacterial glycogen synthesis and its regulation. Annu. Rev. Microbiol. 1984, 38, 419–458. [Google Scholar] [CrossRef] [PubMed]
  75. Carrera, D.N.Ï.; Oddsson, S.; Grossmann, J.; Trachsel, C.; Streb, S. Comparative proteomic analysis of plant acclimation to six different long-term environmental changes. Plant Cell Physiol. 2018, 59, 510–526. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  76. Jiang, C.; Gao, X.; Liao, L.; Harberd, N.P.; Fu, X. Phosphate starvation root architecture and anthocyanin accumulation responses are modulated by the Gibberellin-DELLA signaling pathway in Arabidopsis. Plant Physiol. 2007, 145, 1460–1470. [Google Scholar] [CrossRef] [Green Version]
  77. Yin, Y.; Borges, G.; Sakuta, M.; Crozier, A.; Ashihara, H. Effect of phosphate deficiency on the content and biosynthesis of anthocyanins and the expression of related genes in suspension-cultured grape (Vitis sp.) cells. Plant Physiol. Biochem. 2012, 55, 77–84. [Google Scholar] [CrossRef]
  78. Manna, M.; Islam, T.; Kaul, T.; Reddy, C.S.; Fartyal, D.; James, D.; Reddy, M.K. A comparative study of effects of increasing concentrations of phosphate and phosphite on rice seedlings. Acta Physiol. Plant 2015, 37, 258. [Google Scholar] [CrossRef]
  79. Tominaga-Wada, R.; Masakane, A.; Wada, T. Effect of phosphate deficiency-induced anthocyanin accumulation on the expression of Solanum lycopersicum GLABRA3 (SlGL3) in tomato. Plant Signal. Behav. 2018, 13, e1477907. [Google Scholar] [CrossRef] [Green Version]
  80. Sakuta, M. Diversity in plant red pigments: Anthocyanins and betacyanins. Plant Biotechnol. Rep. 2014, 8, 37–48. [Google Scholar] [CrossRef]
  81. Lutts, S.; Hausman, J.F.; Quinet, M.; Lefèvre, I. Polyamines and their roles in the alleviation of ion toxicities in Plants. In Ecophysiology and Responses of Plants under Salt Stress; Parvaiz, A., Azooz, M.M., Prasad, M.N.V., Eds.; Springer: New York, NY, USA, 2013; pp. 315–353. [Google Scholar]
  82. Niu, Y.F.; Chai, R.S.; Jin, G.L.; Wang, H.; Tang, C.X.; Zhang, Y.S. Responses of root architecture development to low phosphorus availability: A review. Ann. Bot. 2013, 112, 391–408. [Google Scholar] [CrossRef] [PubMed]
  83. Borch, K.; Bouma, T.J.; Lynch, J.P.; Brown, K.M. Ethylene: A regulator of root architectural responses to soil phosphorus availability. Plant Cell Environ. 1999, 22, 425–431. [Google Scholar] [CrossRef]
  84. Zhang, Y.J.; Lynch, J.P.; Brown, K.M. Ethylene and phosphorus availability have interacting yet distinct effects on root hair development. J. Exp. Bot. 2003, 54, 2351–2361. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  85. Talboys, P.J.; Healey, J.R.; Withers, P.J.; Jones, D.L. Phosphate depletion modulates auxin transport in Triticum aestivum leading to altered root branching. J. Exp. Bot. 2014, 65, 5023–5032. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  86. Khan, G.A.; Vogiatzaki, E.; Glauser, G.; Poirier, Y. Phosphate deficiency induces the jasmonate pathway and enhances resistance to insect herbivory. Plant Physiol. 2016, 171, 632–644. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  87. Song, L.; Yu, H.; Dong, J.; Che, X.; Jiao, Y.; Liu, D. The molecular mechanism of ethylene-mediated root hair development induced by phosphate starvation. PLoS Genet. 2016, 12, e1006194. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  88. Zhu, X.F.; Zhu, C.Q.; Zhao, X.S.; Zheng, S.J.; Shen, R.F. Ethylene is involved in root phosphorus remobilization in rice (Oryza sativa) by regulating cell-wall pectin and enhancing phosphate translocation to shoots. Ann. Bot. 2016, 118, 645–653. [Google Scholar] [CrossRef] [Green Version]
  89. Park, C.H.; Roh, J.; Youn, J.; Son, S.; Park, J.H.; Kim, S.Y.; Kim, T.; Kim, S. Arabidopsis ACC oxidase 1 coordinated by multiple signals mediates ethylene biosynthesis and is involved in root development. Mol. Cells 2018, 41, 923–932. [Google Scholar] [PubMed]
  90. Bhosale, R.; Giri, J.; Pandey, B.K.; Giehl, R.F.H.; Hartmann, A.; Trainil, R.; Truskina, J.; Leftley, N.; Hanlon, M.; Swarup, K.; et al. A mechanistic framework for auxin dependent Arabidopsis root hair elongation to low external phosphate. Nat. Commun. 2018, 9, 1409. [Google Scholar] [CrossRef] [Green Version]
  91. Su, Y.; Li, M.; Guo, L.; Wang, X. Different effects of phospholipase Dζ2 and non-specific phospholipase C4 on lipid remodeling and root hair growth in Arabidopsis response to phosphate deficiency. Plant J. 2018, 94, 315–326. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  92. Wang, J.; Pei, L.; Jin, Z.; Zhang, K.; Zhang, J. Overexpression of the protein phosphatase 2A regulatory subunit a gene ZmPP2AA1 improves low phosphate tolerance by remodeling the root system architecture of maize. PLoS ONE 2017, 12, e0176538. [Google Scholar] [CrossRef] [Green Version]
  93. Chen, Y.F.; Li, L.Q.; Xu, Q.; Kong, Y.H.; Wang, H.; Wu, W.H. The WRKY6 transcription factor modulates PHOSPHATE1 expression in response to low Pi stress in Arabidopsis. Plant Cell 2009, 21, 3554–3566. [Google Scholar] [CrossRef] [Green Version]
  94. Nordstrom, A.; Tarkowski, P.; Tarkowska, D.; Norbaek, R.; Astot, C.; Dolezal, K.; Sandberg, G. Auxin regulation of cytokinin biosynthesis in Arabidopsis thaliana: A factor of potential importance for auxin-cytokinin-regulated development. Proc. Natl. Acad. Sci. USA 2004, 101, 8039–8044. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  95. Laplaze, L.; Benkova, E.; Casimiro, I.; Maes, L.; Vanneste, S.; Swarup, R.; Weijers, D.; Calvo, V.; Parizot, B.; Herrera-Rodriguez, M.B.; et al. Cytokinins act directly on lateral root founder cells to inhibit root initiation. Plant Cell 2007, 19, 3889–3900. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  96. Ruzicka, K.; Ljung, K.; Vanneste, S.; Podhorska, R.; Beeckman, T.; Friml, J.; Benkova, E. Ethylene regulates root growth through effects on auxin biosynthesis and transport-dependent auxin distribution. Plant Cell 2007, 19, 2197–2212. [Google Scholar] [CrossRef] [Green Version]
  97. Swarup, R.; Perry, P.; Hagenbeek, D.; Straeten, D.V.D.; Beemster, G.T.S.; Sandberg, G.; Bhalerao, R.; Ljung, K.; Bennett, M.J. Ethylene upregulates auxin biosynthesis in Arabidopsis seedlings to enhance inhibition of root cell elongation. Plant Cell 2007, 19, 2186–2196. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  98. Stepanova, A.N.; Yun, J.; Likhacheva, A.V.; Alonso, J.M. Multilevel interactions between ethylene and auxin in Arabidopsis roots. Plant Cell 2007, 19, 2169–2185. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  99. Guo, W.; Zhao, J.; Li, X.; Qin, L.; Yan, X.; Liao, H. A soybean beta-expansin gene GmEXPB2 intrinsically involved in root system architecture responses to abiotic stresses. Plant J. 2011, 66, 541–552. [Google Scholar] [CrossRef]
  100. Lopez-Arredondo, D.L.; Leyva-Gonzalez, M.A.; Gonzalez-Morales, S.I.; Lopez-Bucio, J.; Herrera-Estrella, L. Phosphate nutrition: Improving low-phosphate tolerance in crops. Annu. Rev. Plant Biol. 2014, 65, 95–123. [Google Scholar] [CrossRef] [PubMed]
  101. Bommer, U.A.; Thiele, B.J. The translationally controlled tumour protein (TCTP). Int. J. Biochem. Cell Biol. 2004, 36, 379–385. [Google Scholar] [CrossRef]
  102. Tao, J.J.; Cao, Y.R.; Chen, H.W.; Wei, W.; Li, Q.T.; Ma, B.; Zhang, W.K.; Chen, S.Y.; Zhang, J.S. Tobacco translationally controlled tumor protein interacts with ethylene receptor tobacco histidine kinase1 and enhances plant growth through promotion of cell proliferation. Plant Physiol. 2015, 169, 96–114. [Google Scholar] [CrossRef] [PubMed]
  103. Branco, R.; Masle, J. Systemic signalling through translationally controlled tumour protein controls lateral root formation in Arabidopsis. J. Exp. Bot. 2019, 70, 3927–3940. [Google Scholar] [CrossRef] [PubMed]
  104. Huang, G.; Liang, W.; Sturrock, C.J.; Pandey, B.K.; Giri, J.; Mairhofer, S.; Wang, D.; Muller, L.; Tan, H.; York, L.M.; et al. Rice actin binding protein RMD controls crown root angle in response to external phosphate. Nat. Commun. 2018, 9, 2346. [Google Scholar] [CrossRef] [Green Version]
  105. Chai, Y.N.; Schachtman, D.P. Root exudates impact plant performance under abiotic stress. Trends Plant Sci. 2021, 27, 80–91. [Google Scholar] [CrossRef] [PubMed]
  106. Tian, J.; Wang, X.; Tong, Y.; Chen, X.; Liao, H. Bioengineering and management for efficient phosphorus utilization in crops and pastures. Curr. Opin. Biotechnol. 2012, 23, 866–871. [Google Scholar] [CrossRef]
  107. Koyama, H.; Kawamura, A.; Kihara, T.; Hara, T.; Takita, E.; Shibata, D. Overexpression of mitochondrial citrate synthase in Arabidopsis thaliana improved growth on a phosphorus-limited soil. Plant Cell Physiol. 2000, 41, 1030–1037. [Google Scholar] [CrossRef] [PubMed]
  108. Wang, L.; Liu, D. Functions and regulation of phosphate starvation-induced secreted acid phosphatases in higher plants. Plant Sci. 2018, 271, 108–116. [Google Scholar] [CrossRef]
  109. Liu, P.; Cai, Z.; Chen, Z.; Mo, X.; Ding, X.; Liang, C.; Liu, G.; Tian, J. A root-associated purple acid phosphatase, SgPAP23, mediates extracellular phytate-P utilization in Stylosanthes guianensis. Plant Cell Environ. 2018, 41, 2821–2834. [Google Scholar] [CrossRef]
  110. Zhu, S.; Chen, M.; Liang, C.; Xue, Y.; Lin, S.; Tian, J. Characterization of purple acid phosphatase family and functional analysis of GmPAP7a/7b involved in extracellular ATP utilization in soybean. Front. Plant Sci. 2020, 11, 661. [Google Scholar] [CrossRef] [PubMed]
  111. Bozzo, G.G.; Dunn, E.L.; Plaxton, W.C. Differential synthesis of phosphate-starvation inducible purple acid phosphatase isozymes in tomato (Lycopersicon esculentum) suspension cells and seedlings. Plant Cell Environ. 2006, 29, 303–313. [Google Scholar] [CrossRef] [PubMed]
  112. Tran, H.T.; Qian, W.; Hurley, B.A.; She, Y.M.; Wang, D.; Plaxton, W.C. Biochemical and molecular characterization of AtPAP12 and AtPAP26: The predominant purple acid phosphatase isozymes secreted by phosphate-starved Arabidopsis thaliana. Plant Cell Environ. 2010, 33, 1789–1803. [Google Scholar] [CrossRef] [PubMed]
  113. Robinson, W.D.; Park, J.; Tran, H.T.; Del, V.H.; Ying, S.; Zins, J.L.; Patel, K.; McKnight, T.D.; Plaxton, W.C. The secreted purple acid phosphatase isozymes AtPAP12 and AtPAP26 play a pivotal role in extracellular phosphate-scavenging by Arabidopsis thaliana. J. Exp. Bot. 2012, 63, 6531–6542. [Google Scholar] [CrossRef] [Green Version]
  114. Wu, W.; Zhu, S.; Chen, Q.; Lin, Y.; Tian, J.; Liang, C. Cell wall proteins play critical roles in plant adaptation to phosphorus deficiency. Int. J. Mol. Sci. 2019, 20, 5259. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  115. Brown, L.K.; George, T.S.; Thompson, J.A.; Wright, G.; Lyon, J.; Dupuy, L.; Hubbard, S.F.; White, P.J. What are the implications of variation in root hair length on tolerance to phosphorus deficiency in combination with water stress in barley (Hordeum vulgare)? Ann. Bot. 2012, 110, 319–328. [Google Scholar] [CrossRef] [PubMed]
  116. Qin, L.; Jiang, H.; Tian, J.; Zhao, J.; Liao, H. Rhizobia enhance acquisition of phosphorus from different sources by soybean plants. Plant Soil 2011, 349, 25–36. [Google Scholar] [CrossRef]
  117. Chen, L.; Qin, L.; Zhou, L.; Li, X.; Chen, Z.; Sun, L.; Wang, W.; Lin, Z.; Zhao, J.; Yamaji, N.; et al. A nodule-localized phosphate transporter GmPT7 plays an important role in enhancing symbiotic N2 fixation and yield in soybean. New Phytol. 2018, 221, 2013–2025. [Google Scholar] [CrossRef] [PubMed]
  118. Qin, L.; Zhao, J.; Tian, J.; Chen, L.; Sun, Z.; Guo, Y.; Lu, X.; Gu, M.; Xu, G.; Liao, H. The high-affinity phosphate transporter GmPT5 regulates phosphate transport to nodules and nodulation in soybean. Plant Physiol. 2012, 159, 1634–1643. [Google Scholar] [CrossRef] [Green Version]
  119. Zhu, S.; Chen, Z.; Xie, B.; Guo, Q.; Chen, M.; Liang, C.; Bai, Z.; Wang, X.; Wang, H.; Liao, H.; et al. A phosphate starvation responsive malate dehydrogenase, GmMDH12 mediates malate synthesis and nodule size in soybean (Glycine max). Environ. Exp. Bot. 2021, 189, 104560. [Google Scholar] [CrossRef]
  120. Pan, W.; Wu, Y.; Xie, Q. Regulation of ubiquitination is central to the phosphate starvation response. Trends Plant Sci. 2019, 24, 755–769. [Google Scholar] [CrossRef]
  121. Chen, J.; Wang, Y.; Wang, F.; Yang, J.; Gao, M.; Li, C.; Liu, Y.; Liu, Y.; Yamaji, N.; Ma, J.F.; et al. The rice CK2 kinase regulates trafficking of phosphate transporters in response to phosphate levels. Plant Cell 2015, 27, 711–723. [Google Scholar] [CrossRef] [Green Version]
  122. Gelova, Z.; Gallei, M.; Pernisova, M.; Brunoud, G.; Zhang, X.; Glanc, M.; Li, L.; Michalko, J.; Pavlovicova, Z.; Verstraeten, I.; et al. Developmental roles of auxin binding protein 1 in Arabidopsis thaliana. Plant Sci. 2021, 303, 110750. [Google Scholar] [CrossRef] [PubMed]
  123. Washburn, M.; Wolters, D.; Yates, J. Large-scale analysis of the yeast proteome by multidimensional protein identification technology. Nat. Biotechnol. 2001, 19, 242–247. [Google Scholar] [CrossRef]
  124. Unwin, R.; Pierce, A.; Watson, R.; Sternberg, D.; Whetton, A. Quantitative proteomic analysis using isobaric protein tags enables rapid comparison of changes in transcript and protein levels in transformed cells. Mol. Cell. Proteom. 2005, 4, 924–935. [Google Scholar] [CrossRef] [Green Version]
  125. Thompson, A.; Schafer, J.; Kuhn, K.; Kienle, S.; Schwarz, J.; Schmidt, G.; Neumann, T.; Hamon, C. Tandem mass tags: A novel quantification strategy for comparative analysis of complex protein mixtures by MS/MS. Anal. Chem. 2003, 75, 1895–1904. [Google Scholar] [CrossRef]
  126. Ong, S.; Blagoev, B.; Kratchmarova, I.; Kristensen, D.; Steen, H.; Pandey, A.; Mann, M. Stable isotope labeling by amino acids in cell culture, SILAC, as a simple and accurate approach to expression proteomics. Mol. Cell. Proteom. 2002, 1, 376–386. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  127. Messner, C.; Demichev, V.; Bloomfield, N.; Yu, J.; White, M.; Kreidl, M.; Egger, A.; Freiwald, A.; Ivosev, G.; Wasim, F.; et al. Ultra-fast proteomics with scanning SWATH. Nat. Biotechnol. 2021, 39, 846–854. [Google Scholar] [CrossRef] [PubMed]
  128. Zhang, H.; Liu, P.; Guo, T.; Zhao, H.; Bensaddek, D.; Aebersold, R.; Xiong, L. Arabidopsis proteome and the mass spectral assay library. Sci. Data 2019, 6, 278. [Google Scholar] [CrossRef] [Green Version]
  129. Liu, S.; Li, Z.; Yu, B.; Wang, S.; Shen, Y.; Cong, H. Recent advances on protein separation and purification methods. Adv. Colloid Interface Sci. 2020, 284, 102254. [Google Scholar] [CrossRef] [PubMed]
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Figure 1. A model of integration of different adaptive strategies to P deficiency regulated by DAPs in shoots or leaves. The DAPs identified by proteomic analysis associated with different metabolic pathways. Red color indicates proteins with increased accumulations in plants under low-P conditions; blue color indicates proteins with decreased accumulations in plants under low-P conditions; brown color suggests proteins exhibiting either up-regulated or down-regulated accumulations under low-P conditions; dashed lines indicate multiple steps; AA5GT, anthocyanidin 5-O-glucosyltransferase; ACC, 1-aminocyclopropane-1-carboxylate; ADP, adenosine diphosphate; ANS, anthocyanidin synthase; APX, ascorbate peroxidase; Arg, arginine; ADC, arginine decarboxylase; ASA, ascorbic acid; ATP, adenosine triphosphate; 1,3-BPG, 1,3-bisphosphoglycerate; 4-CL1, 4-coumaroyl-CoA ligase 1; DFR, dihydroflavonol-4-reductase; DHAR, dehydroascorbate reductase; E4P, erythritos-4-phosphate; F6P, fructose-6-phosphate; FBP aldoase, fructose-1,6-bisphosphate aldolase; FBP, fructose-1,6-bisphosphate; FNR, ferredoxin-nitrite reductase; Fdx, ferredoxin; G1P, glucose-1-phosphate; G3P, glyceraldehyde-3-phosphate; G6P, glucose-6-phosphate; 6-PGDG, 6-phosphogluconate dehydrogenase; GAPDH, glyceraldehyde-3-phosphate dehydrogenase; GR, glutathione reductase; GSH, reduced glutathione; GSSG, oxidized glutathione; GST, glutathione-S-transferase; GSTF12, glutathione-S-transferase F12; HXK, hexokinase; LDOX, leucoanthocyanidin dioxygenase; MDH, malate dehydrogenase; MDHA, monodehydroascorbate; MDHAR, monodehydroascorbate reductase; Met, methionine; NADP-MDH, NADP-malate dehydrogenase; OAA, oxaloacetic acid; PEP, phosphoenolpyruvate; PEPC, phosphoenolpyruvate carboxylase; 2-PG, 2-phosphoglycerate; 3-PGA, 3-phosphoglycerate; 6-PG-δ-lactone, 6-phosphoglucono-δ-lactone; 6PGLS, 6-phosphogluconolactonase; PGK, phosphoglycerate kinase; Pi, phosphate; PPDK, pyruvate orthophosphate dikinase; RCA, ribulose bisphosphate carboxylase/oxygenase activase; Ru5P, Ribulose-5-phosphate; RuBP, Ribulose-1,5-diphosphate; SAM, s-adenosyl methionine; SCS, succinyl-CoA synthetase; SDH, succinate dehydrogenase; SOD, superoxide dismutase; SPDS, spermidine synthase; SQD1, UDP-sulfoquinovose synthase; SQD2, sulfoquinovosyl diacylglycerol synthase; SQDG, sulfoquinovosyl diacylglycerol; TCA cycle, tricarboxylic acid cycle; UDPG, uridine diphosphoglucose; UDP-SQ, UDP-sulfoquinovosyl.
Figure 1. A model of integration of different adaptive strategies to P deficiency regulated by DAPs in shoots or leaves. The DAPs identified by proteomic analysis associated with different metabolic pathways. Red color indicates proteins with increased accumulations in plants under low-P conditions; blue color indicates proteins with decreased accumulations in plants under low-P conditions; brown color suggests proteins exhibiting either up-regulated or down-regulated accumulations under low-P conditions; dashed lines indicate multiple steps; AA5GT, anthocyanidin 5-O-glucosyltransferase; ACC, 1-aminocyclopropane-1-carboxylate; ADP, adenosine diphosphate; ANS, anthocyanidin synthase; APX, ascorbate peroxidase; Arg, arginine; ADC, arginine decarboxylase; ASA, ascorbic acid; ATP, adenosine triphosphate; 1,3-BPG, 1,3-bisphosphoglycerate; 4-CL1, 4-coumaroyl-CoA ligase 1; DFR, dihydroflavonol-4-reductase; DHAR, dehydroascorbate reductase; E4P, erythritos-4-phosphate; F6P, fructose-6-phosphate; FBP aldoase, fructose-1,6-bisphosphate aldolase; FBP, fructose-1,6-bisphosphate; FNR, ferredoxin-nitrite reductase; Fdx, ferredoxin; G1P, glucose-1-phosphate; G3P, glyceraldehyde-3-phosphate; G6P, glucose-6-phosphate; 6-PGDG, 6-phosphogluconate dehydrogenase; GAPDH, glyceraldehyde-3-phosphate dehydrogenase; GR, glutathione reductase; GSH, reduced glutathione; GSSG, oxidized glutathione; GST, glutathione-S-transferase; GSTF12, glutathione-S-transferase F12; HXK, hexokinase; LDOX, leucoanthocyanidin dioxygenase; MDH, malate dehydrogenase; MDHA, monodehydroascorbate; MDHAR, monodehydroascorbate reductase; Met, methionine; NADP-MDH, NADP-malate dehydrogenase; OAA, oxaloacetic acid; PEP, phosphoenolpyruvate; PEPC, phosphoenolpyruvate carboxylase; 2-PG, 2-phosphoglycerate; 3-PGA, 3-phosphoglycerate; 6-PG-δ-lactone, 6-phosphoglucono-δ-lactone; 6PGLS, 6-phosphogluconolactonase; PGK, phosphoglycerate kinase; Pi, phosphate; PPDK, pyruvate orthophosphate dikinase; RCA, ribulose bisphosphate carboxylase/oxygenase activase; Ru5P, Ribulose-5-phosphate; RuBP, Ribulose-1,5-diphosphate; SAM, s-adenosyl methionine; SCS, succinyl-CoA synthetase; SDH, succinate dehydrogenase; SOD, superoxide dismutase; SPDS, spermidine synthase; SQD1, UDP-sulfoquinovose synthase; SQD2, sulfoquinovosyl diacylglycerol synthase; SQDG, sulfoquinovosyl diacylglycerol; TCA cycle, tricarboxylic acid cycle; UDPG, uridine diphosphoglucose; UDP-SQ, UDP-sulfoquinovosyl.
Cells 11 00651 g001
Cells 11 00651 g002 550
Figure 2. A model of integration of adaptive strategies of plant roots to Pi starvation regulated by DAPs. Up or down arrows indicate DAPs exhibiting increased or decreased accumulations; AM, arbuscular mycorrhiza; ACCO, 1-aminocyclopropane-1-carboxylate oxidase; AOC1, allene oxide cyclase 1; AOS, the allene oxide synthase; BGL, beta-glucosidase; CDC48, cell division cycle protein 48; CS, citrate synthase; JALs, jacalin-related lections; MCM6, mini-chromosome maintenance protein 6; MDH, malate dehydrogenase; PAPs, purple acid phosphatases; PP2A, phosphatase 2A; NPC4, non-phospholipase C4; Ran GTPase, GTP-binding nuclear protein RAN-B1; SAMS, S-adenosyl methionine synthase; TCTP, translationally controlled tumor protein; tRNA IPT, tRNA isopentenyl transferase.
Figure 2. A model of integration of adaptive strategies of plant roots to Pi starvation regulated by DAPs. Up or down arrows indicate DAPs exhibiting increased or decreased accumulations; AM, arbuscular mycorrhiza; ACCO, 1-aminocyclopropane-1-carboxylate oxidase; AOC1, allene oxide cyclase 1; AOS, the allene oxide synthase; BGL, beta-glucosidase; CDC48, cell division cycle protein 48; CS, citrate synthase; JALs, jacalin-related lections; MCM6, mini-chromosome maintenance protein 6; MDH, malate dehydrogenase; PAPs, purple acid phosphatases; PP2A, phosphatase 2A; NPC4, non-phospholipase C4; Ran GTPase, GTP-binding nuclear protein RAN-B1; SAMS, S-adenosyl methionine synthase; TCTP, translationally controlled tumor protein; tRNA IPT, tRNA isopentenyl transferase.
Cells 11 00651 g002
Table
Table 1. A list of proteomic analyses of plant responses to phosphorus deficiency.
Table 1. A list of proteomic analyses of plant responses to phosphorus deficiency.
Plant SpeciesOrgan/TissuesCulture Time before Treatment (d)Treatment Time (d)Protein Separation MethodTotal Protein Number (#)Number of DAPs (#)Protein Number
Identified by MS Analysis		References
Up-RegulatedDown-Regulated
Arabidopsis thalianaLeaves107SCX iTRAQ LC-MS/MS510615610650[17]
Zea maysLeaves4252-DE MALDI TOF MS/TOF1342200ndnd[26]
Zea maysLeaves a18252-DE MALDI-TOF/MALDI-TOF-TOF MS680/59229/719/2055/16[27]
Glycine maxLeaves5142D-IEF/SDS-PAGE MALDI-TOF MS5517710[28]
Glycine maxLeaves314SDS-PAGE Gel Digestion LC-MS/MS4219707267440[29]
Solanum lycopersicumLeavesnd102-DE MALDI-TOF MS/MS/MS600463115[20]
Arabidopsis thalianaRoots1032-DE MALDI
TOF MS		45630ndnd[30]
Arabidopsis thalianaRoots1032-DE iTARQ
LC-MS		13,298356199157[16]
Arabidopsis thalianaRootsnd142-DIGE MALDI-TOF/TOF1420301416[13]
Zea maysRoots24172-DE MALDI
TOF MS		13002547630[31]
Zea maysRoots101/3/7/112-DE MALDI-TOF-MS85091ndnd[32]
Zea maysRoots a24172-DE MALDI
TOF MS		282273/9525/2412/6[19]
Zea maysRoots and102-DE MALDI-TOFnd83/32530/24653/79[33]
Oryza sativaRoots3802-DE MALDI TOF MS66934ndnd[21]
Oryza sativaRoots7212-DE MALDI-TOF MS1401028[34]
Glycine maxRootsnd202-DIGE3251056144[35]
Glycine maxRoots a39SDS-PAGE TMT4216660/133656/1274/6[36]
Glycine maxRootsnd10iTRAQ LC-MS/MSnd713041[25]
Glycine maxRoots514iTRAQnd427213214[15]
Triticum aestivumRoots148iTRAQ6842323ndnd[37]
Hordeum vulgareRoots100.25/2SDS PAGE LC-MS/MSnd697ndnd[38]
Brassica napusRoots a2032-phase LC/MS-MS82831/408/2823/12[39]
Arabidopsis thalianaLeaves/roots732-DIGE MALDI TOF/TOF MS88ndndnd[40]
Hordeum vulgareLeaves/roots17212-DE MALDI-TOF/TOF-MSnd31ndnd[23]
Brassica napusLeaves/roots20262-DE MALDI TOF MS1000324/1213/3[41]
Pinus massonianaSeedlings10582-DE MALDI-TOF/TOF MSnd984454[42]
Arabidopsis thalianaSuspension cells772-DE MALDI TOF MS11046266[43]
Arabidopsis thalianaSuspension cells92SDS-PAGE LFQ5013/18811169/994ndnd[44]
Glycine maxNodules5252-DE MALDI TOF MSnd441727[24]
a Two genotypes were used in the studies. nd, not described.
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Zhou, M.; Zhu, S.; Mo, X.; Guo, Q.; Li, Y.; Tian, J.; Liang, C. Proteomic Analysis Dissects Molecular Mechanisms Underlying Plant Responses to Phosphorus Deficiency. Cells 2022, 11, 651. https://doi.org/10.3390/cells11040651

AMA Style

Zhou M, Zhu S, Mo X, Guo Q, Li Y, Tian J, Liang C. Proteomic Analysis Dissects Molecular Mechanisms Underlying Plant Responses to Phosphorus Deficiency. Cells. 2022; 11(4):651. https://doi.org/10.3390/cells11040651

Chicago/Turabian Style

Zhou, Ming, Shengnan Zhu, Xiaohui Mo, Qi Guo, Yaxue Li, Jiang Tian, and Cuiyue Liang. 2022. "Proteomic Analysis Dissects Molecular Mechanisms Underlying Plant Responses to Phosphorus Deficiency" Cells 11, no. 4: 651. https://doi.org/10.3390/cells11040651

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