Proteomic dissection of the chloroplast: Moving beyond photosynthesis

Journal Pre-proof Proteomic dissection of the chloroplast: Moving beyond photosynthesis

Nilesh Vikram Lande, Pragya Barua, Dipak Gayen, Sunil Kumar, Subhra Chakraborty, Niranjan Chakraborty PII:

S1874-3919(19)30314-8

DOI:

https://doi.org/10.1016/j.jprot.2019.103542

Reference:

JPROT 103542

To appear in:

Journal of Proteomics

Received date:

26 June 2019

Revised date:

15 September 2019

Accepted date:

3 October 2019

Please cite this article as: N.V. Lande, P. Barua, D. Gayen, et al., Proteomic dissection of the chloroplast: Moving beyond photosynthesis, Journal of Proteomics (2019), https://doi.org/10.1016/j.jprot.2019.103542

This is a PDF file of an article that has undergone enhancements after acceptance, such as the addition of a cover page and metadata, and formatting for readability, but it is not yet the definitive version of record. This version will undergo additional copyediting, typesetting and review before it is published in its final form, but we are providing this version to give early visibility of the article. Please note that, during the production process, errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.

© 2019 Published by Elsevier.

Journal Pre-proof

Proteomic dissection of the chloroplast: moving beyond photosynthesis Nilesh Vikram Lande, Pragya Barua, Dipak Gayen, Sunil Kumar, Subhra Chakraborty, Niranjan Chakraborty* [email protected] National Institute of Plant Genome Research, Jawaharlal Nehru University Campus, Aruna Asaf Ali Marg, New Delhi 110067, India. *

Corresponding author at: National Institute of Plant Genome Research, Jawaharlal Nehru

University Campus, Aruna Asaf Ali Marg, New Delhi-110067, India.

of

Abstract

ro

Chloroplast, the photosynthetic machinery, converts photoenergy to ATP and NADPH, which

-p

powers the production of carbohydrates from atmospheric CO2 and H2O. It also serves as a major production site of multivariate pro-defense molecules, and coordinate with other

re

organelles for cell defense. Chloroplast harbors 30-50% of total cellular proteins, out of which

lP

80% are membrane residents and are difficult to solubilize. While proteome profiling has illuminated vast areas of biological protein space, a great deal of effort must be invested to understand the proteomic landscape of the chloroplast, which plays central role in

na

photosynthesis, energy metabolism and stress-adaptation. Therefore, characterization of chloroplast proteome would not only provide the foundation for future investigation of

ur

expression and function of chloroplast proteins, but would open up new avenues for modulation

Jo

of plant productivity through synchronizing chloroplastic key components. In this review, we summarize the progress that has been made to build new understanding of the chloroplast proteome and implications of chloroplast dynamicsing generate metabolic energy and modulating stress adaptation.

Keywords: Chloroplast; Differentially accumulated proteins; Kranz regulators; Photosynthetic machinery; Proteome landscape; Stress adaptation

1. Introduction

Journal Pre-proof Photosynthesis is the most essential biochemical process that occurs in a specialized intracellular organelle, the chloroplast. During the process of photosynthesis, CO2 and light energy are utilized to make sugar molecules and oxygen as by-products. As early as 1883, Andreas F. W. Schimper defined this organelle as “chloroplastids” and later Eduard Strasburger used the word “chloroplast”. Chloroplasts are believed to be originated 1.2 billion years ago through cyanobacterial endosymbiosis in eukaryotes [1]. That is why chloroplasts, although characteristic of plant and green algae, show numerous prokaryotic features. In evolution, cyanobacterial genome was enormously decreased within chloroplast due to gene rearrangement and loss [2].

of

The chloroplast genome harbors ~120-130 genes, the majority of which are associated with

ro

photosynthetic apparatus.

Inheritance of chloroplast is typically uniparental like that of mitochondria with some exception

-p

like Passiflora [3]. Most angiosperms inherit chloroplasts maternally via female gamete, whereas gymnosperms inherit paternally via male pollen. The chloroplasts are guarded by double

re

membrane, which is suggestive of their emergence via endosymbiosis. Most of the

lP

photosynthetic apparatus viz., photosynthetic pigments and electron carriers are part of internal organized membrane sacs, called thylakoids [4] and often numerous thylakoids are stacked,

na

called grana. The thylakoid membrane envelops a central aqueous region designated thylakoid lumen. The space between the inner membrane and the thylakoid membrane is filled with stroma, which is the site of carbohydrate synthesis. Since most of the chloroplast genome had

ur

been lost during evolution, the function and maintenance of the chloroplast thus required to

Jo

import protein from cytosol. Proteins that are synthesized by cytosolic ribosome and translocated to chloroplast must have chloroplast transit peptide. These transit peptides are recognized by translocons present at the outer and inner chloroplast envelope membranes (TOC and TIC) [5]. The precursor proteins that are resident of thylakoid membrane and lumen must have one more additional signal sequence to chloroplast transit peptide. After translocation, transit peptide is cleaved by stromal signal peptidase, which eventually exposes the thylakoid targeting signal. These hydrophilic thylakoid signal sequences then initiate next translocation of proteins to the thylakoids. In contrast to transit peptide mediated transport to chloroplast, several proteins such as quinone oxidoreductase (ceQORH) and Tic32 are imported to the chloroplast in an independent manner [6,7]. Chloroplast proteins of import machinery, such as OEP34, are translocated to chloroplast outer envelope membrane without any transit peptide.

Journal Pre-proof

To increase photosynthetic activity, the chloroplasts orient themselves in such way that more photo light can be absorbed by chlorophyll. In low-light conditions, they spread over the peripheral surface of cell towards direction of light, but they align in vertical columns to protect themselves from photooxidative damage under high-light intensity, [8]. This helps the land plants to evolve in such a way that the cells can accommodate many small chloroplasts than a single large chloroplast. In higher plants, the chloroplast relocation movement is regulated by the

of

blue light receptors, the phototropins.

ro

Chloroplasts alone synthesize almost all of a plant cell's amino acids in the stroma except sulfurcontaining ones, albeit some part of methionine and cysteine biosynthesis also takes place in

-p

chloroplast [9, 10]. In plants, O-phosphohomoserine acts as common precursor of threonine and methionine. Most of methionine biosynthesis reaction takes place in chloroplast except final

re

step, which takes place in cytosol [11]. Methionine serves as a precursor for protein and S-

lP

adenosyl methionine (SAM) biosynthesis, which plays important role in biosynthesis of polyamines and phytohormone ethylene [10]. The metabolic process of sulfur assimilation takes

na

place in more than one subcellular compartment, presumably because cysteine has difficulty in crossing double-envelope membrane. Chloroplasts also synthesize nitrogenous bases, purines and pyrimidines. Nitrite reductase catalyzes the 6e- reduction from NO2– to NH4+ found in

ur

plastids, which contributes to make amino acids and nucleotides. Chloroplast also carries out

Jo

protein biotinylation reaction as mitochondria does. Biotin dependent carboxylases in plant such as acetyl-CoA carboxylase, geranyl-CoA carboxylases and methylcrotonoyl-CoA carboxylases are residents of cytosol, chloroplast and mitochondria, respectively [12].

Unlike animals, plants do not have any specified cells for immune reaction and thus every plant cells take part in immune response. Chloroplast plays a major role in plant response along with nucleus, mitochondria, plasma membrane and endoplasmic reticulum [13]. It is one of two prime sources of reactive oxygen species (ROS) production [14] and responsible for producing the pathogen-response signaling molecules, salicylic acid (SA) and jasmonic acid (JA). Chloroplast triggers both biotic and abiotic stress responses by changing the levels of ROS. During plantpathogen interactions, chloroplast induces the production of H2O2 that directly kills the invading

Journal Pre-proof pathogen. On the other hand, lower concentration of H2O2 promotes systemic acquired resistance effective against a wide range of phytopathogens. The structural complexity and fundamental functions accomplished by the chloroplast makes its constituent proteins promising targets for molecular intervention and crop improvement. The evolution of C4 photosynthesis particularly involved specialized distinct leaf anatomy, known as Kranz where compartmentalized metabolic reactions take place. So far, many groups had attempted to develop C4 pathways in rice plant, but fragmented information about regulatory mechanism of Kranz anatomy hindered such efforts [15]. The expression of candidate Kranz regulators, ZmSHR1, involved in vascular development

of

and bundle sheath cell specification failed to induce C4-like leaf anatomy [16]. However,

ro

constitutive expression of ZmGLK1 in rice could induce an anatomy which, to some extent, was similar to C4 plant [17]. Notably, such regulators can be identified through screening of

-p

chloroplast proteome. During the past decade, chloroplast research, particularly on photosynthesis and photorespiration, had reached an exemplary advanced stage [18]. Much

re

attention has been focused on delineating proteome landscape of chloroplast to understand the

lP

biological and defense related functions of the protein components, beyond photosynthesis.

na

2. Cell fractionation and structural integrity of chloroplast

The foremost and crucial step of chloroplast isolation is the breaking of cell wall without

ur

hampering the vacuoles. Notably, this problem does not occur in case of chloroplast isolated

Jo

from the protoplast. However, the yield of chloroplasts isolated from protoplast is very less and not sufficient for global proteomics analysis. To prevent the activity of hydrolytic enzymes and secondary metabolites from vacuoles, protecting agents such as bovine serum albumin (BSA), polyvinyl pyrrolidone (PVP) along with reducing agents more specifically ascorbate, cysteine and 2-mercaptoethanol are used in extraction buffer [19]. Chelating agents (EDTA/EGTA) are also used to capture divalent metal cations (Ca2+, Mg2+ and Zn2+) to rescue envelope membrane from the activity of ion-dependent lipases and phospholipases. It is very important to have osmotic agents in chloroplast extraction buffer to avoid hypotonic shock as chloroplast, in living cells, is surrounded by high osmotic pressure. The osmotic pressure is controlled by adding a variety of sugar and sugar alcohols such as mannitol, sorbitol and sucrose. The osmotic agents can be used separately or in mixture of two and more [20].

Journal Pre-proof

2.1. Isolation of intact chloroplasts

Successful isolation of chloroplast has previously been reported from model plants and many crop species of economic importance including barley, chickpea, maize, pea and wheat [21-25]. The widely used chloroplast isolation techniques in various crops across laboratories are compared and listed in Table 1. Most homogenization buffers have 0.33 M sorbitol as osmoticum and have an osmotic potential of - 1.0 MPa, which is the osmotic potential of the cell

of

sap of Arabidopsis chickpea and spinach. It is important to determine the osmotic potential of the

ro

cell sap and ensure that the osmotic potential of homogenisation buffer is equal to that of the tissues under study. The amount and growth stage of tissues used for isolation, vary from 1 g to

-p

140 g and 7 days to 4 months, respectively. This variation in tissue dictates the Percoll density used, and the force and duration of density gradient centrifugation step (Table 1). Percoll or

re

sucrose gradients are most commonly used to purify chloroplasts. The disadvantage of sucrose

lP

gradient is that chloroplast might burst by high sucrose concentrations [26]. Percoll is expensive but well suited for density gradient experiment because it possesses a less viscosity as compared

na

to sucrose and low osmolarity, and no toxicity towards cells. Also, it does not interfere with majority of the enzyme assays. However, it reacts with Fohn-Ciocalteau protein assay and several nucleic acid and polysaccharide assays [27]. Both step and continuous Percoll density

Jo

centrifugation speed.

ur

gradients are widely used in chloroplast isolation with different Percoll percentage and

To develop a method for chloroplast isolation from a particular plant, some points should be taken into consideration. This includes the type of experimental tissue and the effective gradient composition. The quantity and quality of the isolated chloroplast varies with tissues used. It is important to use optimized concentration of agar and sucrose in MS medium while developing experimental tissues. If the medium is too soft, it might come out with harvested tissues and interfere with homogenization. Starch grains in chloroplast generally break down during centrifugation and affect the isolation of intact of chloroplast. Therefore, to reduce starch amount plants should be kept in dark before tissue harvesting. After the selection of tissue, cell lysis is an important step to release chloroplast. Inadequate homogenization results in incomplete cell lysis or chloroplast rupture. As discussed above, different types of gradients are used for chloroplast

Journal Pre-proof isolation and thus which type of gradient would perform better has to be standardized for individual cases. Rice chloroplast are difficult to isolate by Percoll gradient because they highly aggregate in such gradient and hence Nycodenz gradient is used [28]. To prepare step gradient expertise hand or a peristaltic pump is required to avoid mixing. In step gradient, intact chloroplasts form a layer at the interface, which is difficult to extract than pellet or lower band formed in continuous gradient. It is better to collect the interface of intact chloroplasts after removal of the upper layer of gradient to reduce contamination. In case of continuous gradient, chloroplast pellet remains loosely attached to the bottom of tube. Therefore, it is suitable to

ro

-p

2.2 Purity and enrichment of chloroplast proteins

of

remove the supernatant by aspiration to avoid losses.

A major challenge in organelle proteomics is to obtain the organelle per se as pure as possible,

re

and the issue of contamination during organelle enrichment has triggered the development of

lP

different approaches. These approaches majorly rely upon repetitive assessment of the organelle and enrichment of the organelle-specific markers with respect to total protein content.

na

Chloroplast is usually quantified in terms of unit mg of chlorophyll [29]. Chloroplast integrity is assessed by monitoring the release of oxygen from the chloroplasts. Ferricyanide reacts with electron transport system of thylakoid membrane, accepts electron from water and leads to the

ur

release of oxygen, which can be quantified by an oxygen electrode [30]. Chloroplast integrity

Jo

can also be determined by detecting ferricyanide photoreduction with and without osmotic shock inflicted to the chloroplasts. Photoreduction of ferricyanide can be detected by estimating the time-dependent change in absorbance at 410 nm. Further, the purity of chloroplast fraction can be determined from stained SDS-PAGE using the abundant marker protein RuBisCo (54 kDa RbcL) [31]. Also, enrichment of chloroplast proteins can be examined by immunoblot analysis against different marker proteins, notably LHC II (Light-harvesting complex II of thylakoid membrane), CPN 60 (chaperonin 60 of stroma) and TOC 75. Chloroplast fraction is likely to be contaminated with other cell organelles, notably peroxisomes, mitochondria and vacuoles. Therefore, antibodies against catalase, cytochrome oxidase II and V-ATPase are widely used to check peroxisome, mitochondria and vacuolar contaminants, respectively. Furthermore, the purity of chloroplasts is widely screened by assessing activities of maker enzyme viz., UDPase

Journal Pre-proof and COX in both whole cell lysate and chloroplast fraction to inspect cytoplasmic and mitochondrial contamination, respectively [32].

2.3 Sub-fractionation of chloroplast

To study protein localization and import system, the chloroplast can be further sub-fractionated. Sequential isolation sub-chloroplast fraction such as stroma, thylakoid membrane and lumen are well established in several plant species [33]. Using a combination of osmotic shock and

of

differential centrifugation, chloroplasts can be sub-fractionated. Chloroplast membrane is

ro

ruptured either by osmotic shock in hypotonic buffer or by freeze-thaw in hyperosmotic buffer [34]. After lysis of chloroplasts is complete, the membrane fraction including both thylakoids

-p

and envelope membranes are retrieved by centrifugation, while the stromal fraction remains as supernatant. Sucrose step gradient is more predominately used to separate thylakoids from

re

envelop fraction as compared to Percoll gradient. Isolation of inner and outer envelope

lP

membrane of pea and spinach chloroplast has recently been elaborated by Block et al. [35].

na

3. Advances in proteomic techniques and MS analysis Over the last few decades, mass spectrometry technique has gradually developed and become a powerful tool for identifying thousands of proteins. A major breakthrough in MS analysis was

ur

the introduction of label-based and label-free techniques for quantitative proteomics. Relative

Jo

protein quantification is based on the intensity-based quantification of its corresponding peptides in each sample, and data from multiple peptides are integrated to get a ratio for the parent protein [36]. It involves comparison of the peaks of the same peptide, with the same ionization efficiency in different samples. However, ionization efficiencies of different peptides are different and thus it is not possible to directly compare peaks of different peptides to determine their abundance in a sample [37]. Absolute quantification involves addition of standard with known concentration. Such experiments become limited to smaller number of samples due to high cost of standard used [38]. Analyses of thousands of samples in label-free quantification came as a major advantage over labelling approach [39]. Missing value problem is one of the major limitation of label-free approach [40]. Friso et al. [41] described workflow of label-free spectral counting for quantitative chloroplast proteome analysis in Arabidopsis and maize. In

Journal Pre-proof label-based quantitative proteomics, different samples are co-analysed and relative quantification is carried out within single runs, which leads to much higher reproducibility and avoids missing value problem [42]. In Data-Dependent Acquisition (DDA) method, the instrument selects the largest peak from MS1 spectrum for further MS2 analysis for peptide identification [43]. In Data-Independent Acquisition (DIA), all the spectra generated in specific m/z range from MS1 spectra, are selected and used for fragmentation [44]. The problem of missing value in label-free DDA approach is not faced in DIA because it covers all peptides and every peptide is fragmented multiple times.

of

DIA is more frequently used in comparing large number of samples. Fragmentation of a number

ro

of peptides results in generation of very complex series of MS2 spectra, which is more difficult to analyse. Notably SWATH-MS approach is advantageous to analyze these complex spectra

-p

using prior knowledge of peptides chromatograph [45].

Dunkley et al. [46] developed localization tool for organellar proteins by isotope tagging

re

(LOPIT) used for simultaneous analysis of multiple subcellular proteins of Golgi, endoplasmic

lP

reticulum (ER), plasma membrane, mitochondria, and chloroplast. It does not require absolute purification of organelle. It is based on principle that proteins from similar organelle exhibit

na

similar distributions in the density gradient. Proteins distributed to different gradient fractions are analysed by relative quantitation. LOPIT has been applied to study the subcellular proteomes of Arabidopsis [47-49]. Recently, an improved version of LOPIT method, hyperplexed localisation

ur

of organelle Proteins by Isotope Tagging (hyperLOPIT) and Localisation of Organelle Proteins

Jo

by Isotope Tagging after Differential ultra-Centrifugation (LOPIT-DC) have been developed, integrating novel approaches for sample preparation and MS data acquisition. HyperLOPIT achieves high-resolution separation of organelles and sub-organelle compartments, but is relatively time- and resource-intensive when compared to LOPIT-DC [50].

4. Characterization of chloroplast proteomes using classical and emerging techniques

During the past decades, chloroplast proteomes have been analyzed from a wide range of crop species, besides model plant, Arabidopsis. In Arabidopsis alone, more than 10 chloroplast proteomes have been documented, highlighting various techniques in mapping the chloroplast proteome and discovery of proteins (Table 2). Several proteomics tools, starting from SDS-

Journal Pre-proof PAGE to gel-free and novel peptide labeling techniques, have been employed to generate the chloroplast proteome.

4.1 Development of chloroplast reference proteome map Chloroplast proteome of Arabidopsis has been extensively characterized by gel-based and gelfree strategies (Table 2). Kleffmann et al. [51] reported a list of 690 chloroplast proteins using shotgun proteomic technique. This dataset includes proteins that did not have chloroplast transit peptide and showed homology with those in cyanobacteria, highlighting cyanobacterial origin of

of

chloroplast. Behrens et al. [52] identified 1841 and 436 non-redundant chloroplast proteins

ro

belonging to eight different protein complex separated by 2-D blue native (BN)/PAGE.

-p

Using a traditional approach, Fan et al. [53], and identified 15 chloroplast proteins from Arabidopsis and Salicornia europaea. A comprehensive analysis of chickpea chloroplast

re

proteome led to the identification of 2451 proteins including several novel candidates [22]. In

lP

silico prediction showed more than 50% of the identified proteins to be localized to the chloroplast. Yuan et al. [54] identified more than 100 proteins form chloroplast isolated from

na

poplar. Chloroplasts from submerged marine plant, Posidoniaoceanica, revealed 72 proteins [55]. The proteomic analysis of submerged plant showed high abundance of distinct group of proteins when compared with land plants. These DAPs in submerged plant are likely to modulate

ur

photosynthetic activity and glucose metabolisms to adapt to low light condition.

Jo

A comparative analysis of chloroplasts purified from bundle sheath and mesophyll cells of maize revealed 125 DAPs predominantly associated with lipid biosynthesis, nitrogen import, and tetrapyrrole and isoprenoid biosynthesis in mesophyll, whereas sulfur import and starch synthesis in bundle sheath chloroplasts, respectively [23]. This comparative data is available in Plastid Proteome Database (PPDB) at http://ppdb.tc.cornell.edu/. The deciphered proteome of bundle sheath and mesophyll chloroplasts differed both qualitatively and quantitatively. Plastid protein expression and protein import system was found to be significantly reduced in bundle sheath chloroplasts, which also showed upregulation of remodeling proteins such as ClpB and HSP90 [56]. Proteomic analysis of membrane-depleted pea chloroplast revealed 179 proteins, of which very small number of proteins belonged to envelope (23%) and thylakoid membrane (9%) [57]. To identify low abundant soluble chloroplast proteins Bayer et al. [58] initially depleted

Journal Pre-proof RuBisCo by size-exclusion chromatography, followed by affinity chromatography to reduce complexity. In a separate study, chloroplast proteome of wheat single seed descent line revealed 16 DAPs when compared to parental line [59]. Protoplast isolation process imposes stress on plant cells, which triggers change in chloroplast proteome. The comparison of moss chloroplasts isolated from tissue and protoplasts, using DIA SWATH technology, led to the identification of 503 proteins, 479 of which were annotated as chloroplast proteins [60].

of

4.2 Chloroplast subproteomes

ro

The chloroplast subproteomics includes analysis of stroma [61-63] thylakoid membranes [61, 6364], envelopes [61,67, 68] and thylakoid lumen [23, 69]. As discussed above, envelope

-p

membrane contains less than 5% of the total chloroplast membrane and thus development of envelope proteome is often hampered by non-availability of sufficient quantity of intact

re

chloroplasts. An earlier study on Arabidopsis envelope membranes (inner and outer), using

lP

traditional 2-DE coupled with LC-MS/MS led to the identification of 392 nonredundant proteins [68]. Yet another proteomics study of Arabidopsis chloroplast envelope enlisted >100 proteins

na

[67], involved in ion transport and metabolism followed by chloroplast import system and lipid metabolism. A combined proteomics analysis of envelope membrane from two model species, Arabidopsis and Medicago, and pea led to the identification of 341 proteins [70]. Out of 247

ur

unique proteins identified, only 39 were found to be common in, at least, two species. Next an

Jo

alternative approach was adopted in pea employing 2-DE coupled with label-free LC-MS/MS shotgun proteomics. Out of 546 proteins identified, 321 showed differential distribution with 180 and 141 being enriched in inner and outer envelope, respectively [71]. In a separate study, 1269 proteins were identified in purified chloroplast envelope [72], of which 462 were assigned to chloroplast envelope by calculating enrichment of each protein in envelope fraction to that of crude cell extracts along with manual annotation and online prediction tools.

Comparative proteomics of chloroplast envelope membranes is widely used as a strategy to dissect chloroplast-mediated adaptation mechanism. In a classical study, proteome landscape of chloroplast envelope membranes of C3 plant, pea and mesophyll chloroplast envelopes of C4 plant, maize were systematically [73]. The results showed several fold increases in transport of

Journal Pre-proof triosephosphate and phosphoenolpyruvate in maize mesophyll chloroplast envelope as compared to pea. In a similar study, Manandhar-Shrestha et al. [74] identified 21 and 36 DAPs in bundle sheath and mesophyll cells, respectively. In yet another study, membrane composition of etioplast and chloroplasts was analyzed using DIGE and mass spectrometry. The inner membranes of etioplasts and chloroplasts were found to share at least 8 protein complexes [75]. Ferro et al. [61] attempted a novel strategy targeting various chloroplast compartments and developed a database, AT_CHLORO dedicated to chloroplasts subproteomes. The databases were employed to identify proteins from three subcompartments: envelope, stroma and

of

thylakoids. The analysis revealed 337 proteins in stromal fraction, 163 in envelope and 113 in

ro

thylakoids. In an attempt to develop a native state stromal proteome, Peltier et al. [62] separated intact Arabidopsis chloroplasts by colorless native (CN)-PAGE and SDS-PAGE and identified

-p

241 non-redundant proteins. Two-thirds of the identified proteins were assigned to metabolic pathways such as glycolysis, Calvin cycle and pentose phosphate pathway, and only 10% being

re

involved in chloroplast protein synthesis and biogenesis. Another multiprotein complex study of

lP

tic56-3 mutant revealed that assembly of 1-MDa inner envelope translocase (TIC) gets markedly affected [76]. Comparative proteomics of wild-type and tic56-3 mutant led to the identification

na

of 17 DAPs, among which 6 subunits of the chloroplast ATPase, 3 subunits of the 1-MDa TIC complex were dominant. In a stromal proteomic analysis, multiplex mass spectrometry method coupled with ion mobility separation (HD-MSE) recognized 1011 stromal proteins [77]. Some of

ur

them were not previously identified in any large-scale proteomics analyses, suggesting HD-MSE

Jo

as a suitable complementary tool for discovery proteomics. In yet another study, stromal multiprotein complex was fractionated through size exclusion chromatography with a size range up to 5 MDa along with high accuracy tandem mass spectrometry, which revealed 1081 proteins [78]. Thylakoid membranes harbor machinery of light-dependent photosynthetic reactions on the stromal side. To gain in-depth information about proteins that regulate and maintain photosynthetic process, both in-gel and in-solution techniques are widely used. Yin et al. [79] identified 58 proteins, which includes transporters and ABC proteins. Enzymes involved in protein folding and pigment biosynthesis along with photosystem, besides ATP synthase were also identified in cyanobacterium thylakoid membranes [80]. A semi-quantitative proteomics analysis in Arabidopsis revealed 300 thylakoid proteins, being enriched in either grana or stroma lamellae [81], indicating that photosynthetic machineries are unevenly distributed throughout

Journal Pre-proof thylakoids, grana and stroma lamellae. Peltier et al. [69] examined lumenal and peripheral thylakoid fraction of pea chloroplast using mass spectrometry and N-terminal Edman sequencing approach. A classical study of chloroplast protein complex of wheat revealed a total of 18 protein complexes [82]. Thylakoid fraction contained 9 protein complexes, while four complexes were observed in envelope fraction. While procedures for isolation and purification of plastid nucleoids and transcriptionally active chromosome (TCA) are well established, their proteomics characterization is scarce. Melonek et al. [83] reported the proteome landscape of chloroplast TCA fraction of Arabidopsis/mustard and

of

spinach, as well as nucleoid fractions of Arabidopsis, maize and pea [83]. The results showed

ro

that nucleoids and/or TAC might serve as docking station for proteins involved in key metabolic processes. Majeran et al. [84] analyzed nucleoid proteome of proplastids and mature chloroplasts

-p

of maize and identified 2460 proteins. Proteomic analysis of chloroplast plastoglobules (PGs) of Arabidopsis and bell pepper showed that PGs consist of seven fibrillins along with 25 proteins

re

presumably involved in metabolism of isoprenoid-derived molecules and lipids and carotenoid

lP

cleavage [85]. Quantitative differences in protein composition of PG in response to high light stress suggested that it acts as functional metabolic link between the inner envelope and

na

thylakoid membranes and play a role in oxidative stress tolerance.

ur

4.3 Stress-adaptive responses of chloroplast proteome

Jo

Plants encounter diverse environmental stresses during their life cycle, and thus modulate the proteome landscape to survive the adverse conditions. During the adaptation process, proteins are more active than other biological molecules. Chloroplasts act as delicate environmental sensors, since they harbor numerous metabolic pathways that are readily imbalanced by environmental fluctuations. This drives plant biologists to focus on chloroplast proteome and subproteomes in deciphering molecular adaptation to stress. Hypersalinity-responsive chloroplast proteomics of wheat recognized 65 DAPs, most of which were found to be upregulated, while ATP synthase and V-type proton ATPase were downregulated [25]. Furthermore, the comparative salinity-responsive proteomics of wheat cv. SR3 and JN177 revealed 26 DAPs, which include only 8 stress-responsive proteins, and the remaining eighteen exhibited cultivarspecific expressions [86]. The differentially regulated proteins associated with photosynthesis

Journal Pre-proof were found to contribute to capacity of redox maintenance and carbon assimilation. Yet another study compared hypersalinity-responsive chloroplast proteomes of two contrasting brown mustard genotypes and identified 12 DAPs [87]. The analysis of chloroplast proteome in black locust enlisted 61 DAPs [88], indicating that the tetraploid genotypes are better adapted to hypersalinity than the diploid ones. Interestingly the effect of salt stress on mesophyll and bundle sheath chloroplast of amaranth revealed 77 DAPs [89]. Screening of tomato chloroplast proteome indicated that multiprotein complexes, PSI and PSII, respond to slight changes in hypersalinity [90]. However, the abundance of cytochrome b6/f and ATP-synthase complex were

of

found to be reduced, while there was significant increase in light harvesting complex trimers and

ro

monomers. A quantitative proteomic study on a mangrove species, revealed 1030 proteins, out of which 76 proteins were found to be expressed in chloroplast in response to hypersalinity [91]. In

-p

a separate study, using a different genotype, this group identified 46 DAPs [92]. Chloroplast is highly sensitive even to slight change in light intensity. Upon exposure to high-

re

light stress, 52 Arabidopsis chloroplast proteins were found to be differentially modulated [93].

lP

Most of the DAPs were downregulated, while heat shock proteins (HSPs), dehydroascorbate reductase (DHAR) and superoxide dismutase (SOD) were upregulated. Interestingly, 81

na

Arabidopsis chloroplast proteins were found to be differentially acumulated in response to prolonged dark conditions [94]. In another chloroplast proteomics study, proteins associated with photosynthesis, carbon metabololism and plastid mRNA processing were found to be

ur

differentially accumulated under high-light stress in two T-DNA insertional knockout lines,

Jo

EX1and EX2 [95]. A UV-B induced alterations in wheat chloroplast proteome identified 28 DAPs [96]. In both high-light and dark conditions, most DAPs associated with photosynthesis were found to be downregulated. Besides light stress, chloroplast proteome is most affected by dehydration. A recent iTRAQ-based quantitative chloroplast proteomics of rice revealed 40 dehydration-responsive proteins (DRPs) in chloroplast. These DRPs were found to be involved in a wide range of metabolic processes including energy metabolism, photosynthesis and defense response [97]. In yet another study on tomato, 31 chloroplast proteins were found to be differentially regulated under dehydrated condition [98]. One of the interesting observations was that while metabolome indicated a recovery of plants, chloroplast displayed an active adjustment of the proteome [98]. The comparative chloroplast proteomics of sensitive and tolerant tall fescue genotypes under dehydration revealed 10 DAPs [99]. A similar study on chloroplast

Journal Pre-proof proteome, under freezing stress in two contrasting genotypes of pea, showed different acclimation capabilities of chloroplast [100]. During cold acclimation in pea, out of total identified DAPs only 6 were observed with similar change in abundance in both genotypes, 39 DAPs showed genotype-specific accumulation [101]. Simultaneous use of IEF-SDS PAGE and BN-PAGE techniques led to the identification of 56 thylakoid DAPs in beetroot, suggesting that BN-PAGE as a superior technique to resolve highly hydrophobic membrane proteins [102]. Ozone (O3) causes significant changes in chloroplast proteome. Proteomic analysis of soybean chloroplast proteome exposed to high concentration of O3 revealed 32 DAPs [103]. Proteins

of

associated with photosynthesis and carbon assimilation were shown to be downregulated, while

ro

proteins involved in antioxidant defense and carbon metabolism were upregulated. Label-free, isotope labelling and BN-PAGE approaches altogether led to the identification of 119 DAPs in

-p

chloroplast proteome of isoprene-emitting and non-isoprene-emitting poplars [104]. In recent years, some research groups has initiated investigation on chloroplast-virus interaction.

re

Significantly, tomato blistering mosaic virus infection to tobacco, recognized 20 DAPs, some of

lP

which were found to be directly interacting with viral proteins such as ATP synthase β subunit,

na

RNA polymerase beta-subunit, 50S ribosomal protein L6 and Trigger factor-like protein [105].

4.4 Posttranslational modifications (PTM) of chloroplast proteome

ur

PTM is a covalent modification resulting from the addition of a functional group to amino acid

Jo

side chains. Thus far, more than 200 PTMs have been identified and characterized in both prokaryotes and eukaryotes. In Arabidopsis, TiO2, IMAC and MudPIT strategies have been widely used which led to the identification of 965 phosphorylation sites on 712 proteins, out of which 181 phosphorylation sites were predicted on 125 proteins [106]. Another phosphoproteomic analysis of Arabidopsis chloroplast identified 2597 phosphopeptides with 2172 unique phosphorylation sites from 1346 phosphoproteins [107]. Further, Reiland et al. [108] analysed phosphoproteome of Arabidopsis and identified 1,429 phosphoproteins and 3,029 unique phosphopeptides. Investigation of global lysine crotonylation using immune-affinity purification coupled with LCMS/MS recognized 2044 lysine crotonylation sites on 637 proteinsof tobacco [109]. Most of these crotonylated proteins were predicted to be chloroplast localized (37%), followed by cytosol

Journal Pre-proof (30%), nucleus (12%) and mitochondria (5%). Among 236 crotonylted chloroplast proteins, a total of 72 were found to be involved in photosystem II complex, cytochrome b6f complex, photosystem I complex, ferredoxin-NADP reductase and ATP synthesis complex. Additionally, rice crotonylome analysis identified 1265 crotonylation site on 690 proteins. Localization predictions revealed 51% of the proteins to be chloroplast residents, which is higher than tobacco [110]. Proteome-wide analysis of histone deacetylases signature sequences revealed, 91 new acetylated proteins other than histones, and majority of the protein targets have functions in photosynthesis [111].

of

Bienvenut et al. [112] developed a Stable-Isotope Protein N-terminal Acetylation Quantification

ro

(SILProNAQ) workflow to determine the N-terminal acetylation yield of mature proteins for a clearly defined localization. SILProNAQ involved both wet-lab techniques and data analysis.

-p

TAILS method of labelling and MS/MS-based identification revealed that many nucleusencoded proteins accumulated in chloroplast with two or three different N-termini amino acids

re

[113]. Proteomic data analysis showed that small, apolar, or hydroxylated residues (Ala, Val,

lP

Ser, and Thr) are more prominent to N-terminal amino acids of stromal proteins. It seems that chloroplast protein degradation mostly occurs through cleavage between Arg and Thr. Yet

na

another bioinformatics approach concerning lysine and arginine methylation network of Arabidopsis identified 31 methylation sites of 23 chloroplastic proteins, associated with

ur

processes like photosynthesis, chloroplast biogenesis and maintenance [114].

Jo

5. Chloroplast protein databases

During past few years, several proteomics studies, in different laboratories, aimed at addressing subcellular fractions of chloroplast and quantified thousands proteins. These large proteomic data are available in public web databases. The AT_CHLORO database contains information about 1,856 protein groups of chloroplast subfractions of Arabidopsis. It contains 487, 483, 220 and 1290 proteins in envelope, stroma, thylakoid and grana/stroma lamellae, respectively. The AT_CHLORO database provides knowledge about MS data, function and subplastidial localization, besides links to other public databases and references [115, 116]. Chloroplast Function Database contains 1369 chloroplast proteins linked to phenotype of 3,246 transposon or T-DNA mutant lines [117]. The Chloroplast Function Database II is a large-scale collection of 2495 nucleus-encoded chloroplast proteins linked to homozygous single gene knockout

Journal Pre-proof phenotypic T-DNA mutant lines [118]. Yet another Plastid Protein Database (plprot) is a largescale collection of 2,043 proteins of etioplasts, chloroplasts, chromoplasts and undifferentiated proplastid-like organelles of tobacco BY2 cells [119]. The database has two modules, one provides blast search option and the other gives information of the plastid proteomes. The Plant Proteomics Database (PPDB) was initially dedicated to disseminating chloroplast proteome, designated Plastid Proteome DB. Now PPDB holds all cellular proteins along with their posttranslational modifications identified from Arabidopsis and maize [120]. Interestingly, ChloroKB, is a web application useful for the analysis of chloroplast metabolic network and

of

related cellular pathway [121]. The latest ChloroKB database contains 1147 proteins (559*, 68**

ro

and 520*** proteins localised to plastid*, dual-localised to plastid** and outside plastid***, respectively) in 125 interconnected metabolic networks].

-p

Sun et al. [122] carefully collected previously published datasets of chloroplast sub proteomes and observed that Cys content is much lower in thylakoid than inner envelope membrane, with

re

significant differences in protein length, pI and TMD distribution.

Based on Arabidopsis

lP

genome analysis, it is predicted that 4255 proteins (520- inner envelope or thylakoid membrane, 576 soluble lumen, 57 intergal membrane and 3387 localized proteins) have chloroplast transient

na

peptides. Using these parameters, an investigator can predict plastid subproteomes and identify low abundant proteins from fully annotated genome. Yet another genome analysis of apple predicted roughly 40% of its total proteins to be plastid-targeted, which was 57% less than that

ur

of Arabidopsis, suggesting difference in plastid-targeted proteome [123]. Support Vector

Jo

Machine (SVM) is an online prediction system of genome-wide identification and classification of plastid proteins. This system initially differentiates the plastid from non-plastid protein, and then categorizes them to chloroplast, chromoplast, etioplast and amyloplast [124]. Schlfiff et al. [125] had analysed chloroplast outer envelope membrane proteome by in silico as well as proteomic approach and developed an algorithm for prediction of outer envelope β barrel proteins. Based on chloroplast evolutionary relationship with Gram-negative bacteria, they hypothesized that envelope membrane should have large number of β barrel proteins and validated this hypothesis by calculating the probability for the existence of β-sheet and hairpin structures among all proteins. The chloroplast proteomic data generated from MS/MS analysis becomes more authenticated by prediction of the subcellular localization of the identified proteins to chloroplast and detection of

Journal Pre-proof the presence of cleavable chloroplast transit peptides. A considerable number of localization prediction tools are freely available online, some of which are documented in Table 3 [126-135]. Further databases such as PRIDE or NCBI accept submission of raw files generated from mass spectrometry analysis, which can be downloaded and further analysed.

6. Comparative analysis of chloroplast proteomes To have a comprehensive overview of the chloroplast proteomes, reported thus far, we carried out a sequence-based comparative analysis. First, we made an exhaustive lists of the chloroplast

of

proteins and catalogued them according to individual plant species, and then prepared separate

ro

non-redundant sets. Table 2 enlists the individual proteomics studies reported in different species using different tissue types and methodologies. In all, 9626 chloroplast proteins were identified

-p

from plants and green algae, which summed up to non-redundant set of 7400 proteins. Table 4 provides the non-redundant sets of proteins identified from individual plant species. Of them, a

re

total non-redundant set of 3629 proteins were identified in Arabidopsis from 18 different

lP

chloroplast proteomes (Table 2). In summary, the largest non-redundant set was from Arabidopsis (3629), followed by chickpea (2451), maize (1438) and pea (892) (Table 4,

na

Supplementary Table 1).

To calculate the number of overlaps between chloroplast proteins identified from different plants, sequence-based BLAST analyses were performed. Only 377 proteins were found to be

ur

common across Arabidopsis, chickpea, maize and pea nr datasets with more than 60% identity.

Jo

Amongst them, some evolutionarily conserved proteins associated with photosynthesis and carbon assimilation reaction were recognized. While comparing between Arabidopsis and chickpea, 1592 proteins were found to be common, while 2039 were exclusive to Arabidopsis and 859 were unique to chickpea. Berglund et al. [136] has earlier reported 40 proteins that are encoded by nuclear genome, translated in cytosol, and are transported to both chloroplast and mitochondria. Of them, 23 proteins were listed in the Arabidopsis nr set of chloroplast proteins (Supplementary Table 2), while remainings came from proteomic analysis. TAIR and NCBI databases predicted 88 and 75 proteins to be encoded by chloroplast genome of Arabidopsis and chickpea, respectively. Notably, investigations of Arabidopsis and chickpea chloroplast proteomes provided experimental validation of 61 and 56 of these predicted plastid-encoded proteins, respectively

Journal Pre-proof (Supplementary Table 1). While proteins synthesized on cytosolic ribosomes are imported to the chloroplast with the help of transit peptide, chloroplast encoded proteins do not require any such signal peptide. Significantly, these studies not only encompass chloroplast proteome landscape, but also provide experimental validation of predicted chloroplast-encoded proteins. The chloroplast subproteomes of different plant species estimated a total of 3392, 2670, 2027, 81 and 448 proteins enriched in chloroplast envelope, stroma, thylakoids, plastoglobuli and nucleoid, respectively (Table 2). In all, about 1000 chloroplast proteins were shown to be differentially accumulated in response to

of

multivariate stress conditions, among them 386 were hypersalinity-responsive, followed by 162

ro

high-light and 95 dehydration-responsive (Table 2). Till date only 4 reports are available on chloroplast proteome of C4 plants, maize and amaranths [23, 56, 74, 89]. These studies involved

-p

analysis of bundle sheath and mesophyll chloroplast proteins (Table 2). which are likely to be potential candidates for crop improvement through molecular breeding and genetic engineering

lP

re

approaches.

na

8. Conclusion

This review encompasses a comprehensive literature survey of more than 80 publications on

ur

chloroplast proteomes and subproteomes. These proteome data provide basic knowledge for crop

Jo

improvement through biotechnological application, which rely on information generated from various omics technology. Genome always remains static in nature, while proteome has capability to alter with change in environmental conditions. The knowledge about the key proteins that play crucial roles in photosynthesis and carbon assimilation are critical to develop plants with improved photosynthetic performance and enhanced nutritional value. A search of the chloroplast proteome research literature illustrates more than 1000 stress-responsive proteins, which can eventually be placed in the pipeline for crop improvement programs. The DAPs of bundle sheath and mesophyll chloroplast are valuable to elucidating proteins that direct the mechanism of Kranz anatomy. These key regulators important for engineering C3 crops, for example rice, to use C4 pathway are currently held back by lack of information. The characterization of the chloroplast proteomes from various systems reveals their importance and

Journal Pre-proof biological relevance. We highlight the advances in proteomic techniques, contributing to increased identification of chloroplast proteins, would help to better understand their diverse functional role in cell beyond photosynthesis.

Conflict of interest The authors declare no conflict of interest.

ro

of

Acknowledgments

This work was supported by the Council of Scientific and Industrial research (CSIR)

-p

[38/1385/14/EMR-II], Govt. of India and Department of biotechnology (DBT) [BT/AGR/CGPhase II/01/2014]. We thank CSIR, UGC and Department of Science and Technology (DST) for

re

providing research fellowship to NVL, SK, PB and DG, respectively. We sincerely apologize to

lP

all plant proteomics groups whose work could not be cited because of space constraints.

na

References

ur

1. V. Zimorski, C. Ku, W.F. Martin, S.B. Gould, Endosymbiotic theory for organelle origins, Curr. Opin. Microbial. 22 (2014) 38-48.

Jo

2. W. Martin, B. Stoebe, V. Goremykin, S. Hansmann, M. Hasegawa, K.V. Kowallik, Gene transfer to the nucleus and the evolution of chloroplasts, Nature 393 (1998) 162-165. 3. A.K. Hansen, L.K. Escobar, L.E. Gilbert, R.K. Jansen, Paternal, maternal, and biparental inheritance of the chloroplast genome in Passiflora (Passifloraceae): implications for phylogenetic studies, Am. J. Bot. 94 (2007) 42-46. 4. E. Shimoni, H.O. Rav, I. Ohad, V. Brumfeld, Z. Reich, Three-dimensional organization of higher-plant chloroplast thylakoid membranes revealed by electron tomography, Plant Cell 17 (2005) 2580-2586. 5. P. Jarvis, J. Soll, Toc, Tic, and chloroplast protein import, Biochim. Biophys. Acta. 1541 (2001) 64-79.

Journal Pre-proof 6. S. Miras, D. Salvi, M. Ferro, D. Grunwald, J. Garin, J. Joyard, N. Rolland, Non-canonical transit peptide for import into the chloroplast, J. Biol. Chem. 277 (2002) 47770-47778. 7. A. Nada, J. Soll, Inner envelope protein 32 is imported into chloroplasts by a novel pathway, J. Cell Sci. 117 (2004) 3975-3982. 8. T. Kagawa, T. Sakai, N. Suetsugu, K. Oikawa, S. Ishiguro, T. Kato, S. Tabata, K. Okada, M. Wada, Arabidopsis NPL1: a phototropin homolog controlling the chloroplast highlight avoidance response, Science 291 (2001) 2138-2141. 9. B. Miflin, P. Lea, Amino acid metabolism, Annu. Rev. Plant Biol. 28 (1977) 299-329.

of

10. S. Ravanel, B. Gakière, D. Job, R. Douce, The specific features of methionine

ro

biosynthesis and metabolism in plants Proc. Natl. Acad. Sci. 95 (1998) 7805-7812. 11. H. Hesse, R. Hoefgen, Molecular aspects of methionine biosynthesis. Trends Plant Sci. 8

-p

(2003) 259-262.

12. V. Pinon, S. Ravanel, R. Douce, C. Alban, Biotin synthesis in plants. The first committed

re

step of the pathway is catalyzed by a cytosolic 7-keto-8-aminopelargonic acid synthase.

lP

Plant physiol., 139 (2005), 1666-1676.

13. H. Nomura, T. Komori, S. Uemura, Y. Kanda, K. Shimotani, K. Nakai, T. Furuichi, K.

na

Takebayashi, T. Sugimoto, S. Sano, Chloroplast-mediated activation of plant immune signalling in Arabidopsis, Nat. Commun. 3 (2012) 1926. 14. K. Asada, Production and scavenging of reactive oxygen species in chloroplasts and their

ur

functions, Plant physiol. 141, (2006) 391-396.

Jo

15. O.V. Sedelnikova, T.E. Hughes, J.A. Langdale, Understanding the genetic basis of C4 Kranz anatomy with a view to engineering C3 crops. Annual review of genetics, 52 (2018) 249-270.

16. M.L. Schuler, O.V. Sedelnikova, B.J. Walker, P. Westhoff, J.A. Langdale, SHORTROOT-mediated increase in stomatal density has no impact on photosynthetic efficiency. Plant Physiol. 176 (2018) 757–72. 17. P. Wang, R. Khoshravesh, S. Karki, R. Tapia, C.P. Balahadia, A.Bandyopadhyay, et al, Re-creation of a key step in the evolutionary switch from C3 to C4 leaf anatomy. Curr. Biol. 27 (2017) 3278–87 18. U. Armbruster, P. Pesaresi, M. Pribil, A. Hertle, D. Leister, Update on chloroplast research: new tools, new topics, and new trends, Mol. Plant 4 (2011) 1-16.

Journal Pre-proof 19. J.M. Gualberto, H. Handa, J.M. Grienenberger, Isolation and fractionation of plant mitochondria and chloroplasts: specific examples, Methods Mol. Biol, 50 (1995) 161175. 20. D.A. Walker, Z.G. Cerovic, S.P. Robinson, Isolation of intact chloroplasts: General principles and criteria of integrity, Methods Enzymol. 148 (1987) 145-157. 21. M. Ploscher, V. Reisinger, L.A. Eichacker, Proteomic comparison of etioplast and chloroplast protein complexes. J Proteomics 74 (2011) 1256-1265. 22. N.V. Lande, P. Subba, P. Barua, D. Gayen, T.K. Prasad, S. Chakraborty, N. Chakraborty,

of

Dissecting the chloroplast proteome of chickpea (Cicer arietinum L.) provides new

ro

insights into classical and non-classical functions, J. Proteomics 65 (2017) 11-22. 23. W. Majeran, Y. Cai, Q. Sun, K.J. van Wijk, Functional differentiation of bundle sheath

-p

and mesophyll maize chloroplasts determined by comparative proteomics, Plant Cell 17 (2005) 3111-3140.

re

24. J.B. Peltier, G. Friso, D.E. Kalume, P. Roepstorff, F. Nilsson, I. Adamska, K.J. van

lP

Wijka, Proteomics of the chloroplast: systematic identification and targeting analysis of lumenal and peripheral thylakoid proteins, Plant Cell 12 (2000) 319-341.

na

25. A. von Zychlinski, W. Gruissem, Preparation and analysis of plant and plastid proteomes by 2DE. Methods Mol Biol. 519 (2009) 205-220. 26. A.H.M. Kamal, K. Cho, D.E. Kim, N. Uozumi, K.Y. Chung, S.Y. Lee, J.S. Choi, S.W.

ur

Cho, C.S. Shin, S.H. Woo, Changes in physiology and protein abundance in salt-stressed

Jo

wheat chloroplasts, Mol. Biol. Rep. 39 (2012) 9059-9074. 27. A.H. Millar, A. Liddell, C.J. Leaver, Isolation and subfractionation of mitochondria from plants. Methods Cell Biol. 65 (2001) 53-74. 28. P.T. Sharpe, Methods of cell separation. Elsevier, 18 (1988). 29. .A. von Zychlinski, W. Gruissem,. Preparation and analysis of plant and plastid proteomes by 2DE. Methods Mol. Biol. 519 (2009) 205-220. 30. D.I. Arnon, Copper enzymes in isolated chloroplasts. Polyphenoloxidase in Beta vulgaris, Plant physiol. 24 (1949) 1-15. 31. R. Lilley, M. Fitzgerald, K. Rienits, D. Walker, Criteria of intactness and the photosynthetic activity of spinach chloroplast preparations, New Phytol. 75 (1975) 1-10.

Journal Pre-proof 32. R. Zrenner, L. Willmitzer, U. Sonnewald, Analysis of the expression of potato uridinediphosphate-glucose pyrophosphorylase and its inhibition by antisense RNA. Planta 190 (1993) 247-252. 33. M. Hall, Y. Mishra, W. P. Schröder, 2011. Preparation of stroma, thylakoid membrane, and lumen fractions from Arabidopsis thaliana chloroplasts for proteomic analysis. Chloroplast Research in Arabidopsis in Methods Mol. Biol, 775 (2011) 207-222. 34. L. Moyet, D. Salvi, M. Tomizioli, D. Seigneurin-Berny, N. Rolland, 2018. Preparation of Membrane Fractions (Envelope, Thylakoids, Grana, and Stroma Lamellae) from

of

Arabidopsis Chloroplasts for Quantitative Proteomic Investigations and Other Studies.

ro

Methods Mol. Biol. 1696 (2018) 117-136.

35. M.A. Block, 2018. Isolation of Inner and Outer Membranes of the Chloroplast Envelope

-p

from Spinach and Pea. Methods Mol. Biol. 1829 (2018) 137-144 36. M. Bantscheff, M. Schirle, G. Sweetman, J. Rick, B. Kuster, Quantitative mass

re

spectrometry in proteomics: a critical review. Anal. Bioanal. Chem. 389 (2007) 1017-

lP

1031.

37. P.V. Bondarenko, D. Chelius, T.A. Shaler, 2002. Identification and relative quantitation

na

of protein mixtures by enzymatic digestion followed by capillary reversed-phase liquid chromatography tandem mass spectrometry. Anal. Chem. 74 (2002) 4741-4749. 38. S.A. Gerber, J. Rush, O. Stemman, M.W. Kirschner, S.P. Gygi, Absolute quantification

ur

of proteins and phosphoproteins from cell lysates by tandem MS. Proc. Natl. Acad. Sci.

Jo

100 (2003), 6940-6945.

39. J. Cox, M.Y. Hein, C.A. Luber, I. Paron, N. Nagaraj M. Mann, Accurate proteome-wide label-free quantification by delayed normalization and maximal peptide ratio extraction, termed MaxLFQ. Mol. Cell proteomics, 13 (2014), 2513-2526. 40. B.J.M. Webb-Robertson, H.K. Wiberg, M.M. Matzke, J.N. Brown, J. Wang, J.E. McDermott, et al, Review, evaluation, and discussion of the challenges of missing value imputation for mass spectrometry-based label-free global proteomics. J. Proteome rResearch, 14 (2015) 1993-2001. 41. G. Friso, P.D.B. Olinares, K.J. van Wijk, The workflow for quantitative proteome analysis of chloroplast development and differentiation, chloroplast mutants, and protein

Journal Pre-proof interactions by spectral counting. Chloroplast research in Arabidopsis Methods Mol Biol. 775 (2011) 265-282. 42. N. Pappireddi, L. Martin, M. Wühr, A Review on Quantitative Multiplexed Proteomics. ChemBioChem, 20 (2019), 1210-1224. 43. D.C. Stahl, K.M. Swiderek, M.T. Davis, T.D. Lee, Data-controlled automation of liquid chromatography/tandem mass spectrometry analysis of peptide mixtures. J. Am. Soc. Mass Spectrom, 7(1996) 532-540. 44. B.C. Searle, L.K. Pino, J.D. Egertson, Y.S. Ting, R.T. Lawrence, B.X. MacLean, J.

of

Villén, and M.J. MacCoss, Chromatogram libraries improve peptide detection and

ro

quantification by data independent acquisition mass spectrometry. Nat. Commun. 9 (2018) 5128.

-p

45. L.C. Gillet, P. Navarro, S. Tate, H. Röst, N. Selevsek, L. Reiter, R. Bonner, R. Aebersold, Targeted data extraction of the MS/MS spectra generated by data-independent

re

acquisition: a new concept for consistent and accurate proteome analysis. Mol. & Cell.

lP

Proteomics, 11(2012), O111-016717.

46. T.P. Dunkley, R. Watson, J.L Griffin, P. Dupree, K.S. Lilley, Localization of organelle

na

proteins by isotope tagging (LOPIT). Mol. Cell. Proteomics, 3 (2004), 1128-1134. 47. T.P. Dunkley, S. Hester, I.P. Shadforth, J. Runions, T. Weimar, S.L. Hanton, J.L Griffin, C. Bessant, F. Brandizzi, C. Hawes, R.B. Watson, R.B. Mapping the Arabidopsis

ur

organelle proteome. Proc. Natl. Acad. Sci. 103 (2006) 6518-6523.

Jo

48. N. Nikolovski, D. Rubtsov, M.P. Segura, G.P. Miles, T.J. Stevens, T.P. Dunkley, S. Munro, K.S. Lilley, P. Dupree. Putative glycosyltransferases and other plant Golgi apparatus proteins are revealed by LOPIT proteomics. Plant Physiol. 160 (2012) 10371051. 49. A. .

roen,

. Sancho-Andr s,

. .

rec els,

.

atto,

. Aniento,

.S.

illey,

Identification of trans-Golgi network proteins in Arabidopsis thaliana root tissue. J. Proteome Res., 13 (2014), 763-776. 50. A.

elada i, N. .

O.M.

Crook,

L.

ritovše , . . Gatto,

K.S.

rec els, T.S. Smith, O. . Vennard, C. . Lilley,

Combining

LOPIT

with

ulvey,

differential

ultracentrifugation for high-resolution spatial proteomics. Nat. Commun. 10 (2019) 331.

Journal Pre-proof 51. T. Kleffmann, D. Russenberger, A. von Zychlinski, W. Christopher, K. Sjölander, W. Gruissem, S. Baginsky, The Arabidopsis thaliana chloroplast proteome reveals pathway abundance and novel protein functions, Curr. Biol. 14 (2004) 354-362. 52. C. Behrens, C. Blume, M. Senkler, H. Eubel, C. Peterhänsel, H.P.

raun, The ‘protein

complex proteome’ of chloroplasts in Arabidopsis thaliana, . Proteomics 91 (2013) 7383. 53. P. Fan, X. Wang, T. Kuang, Y. Li, An efficient method for the extraction of chloroplast proteins compatible for 2‐ DE and MS analysis. Electrophoresis 30 (2009) 3024-3033.

of

54. H.M. Yuan, K.L. Li, R.J. Ni, W.D. Guo, Z. Shen, C.P. Yang, B.C. Wang, G.F. Liu, C.H.

method, Mol. Biol. Rep. 38 (2011) 3045-3054.

ro

Guo, J. Jiang, A systemic proteomic analysis of Populus chloroplast by using shotgun

-p

55. A. Piro, I.A. Serra, A. Spadafora, M. Cardilio, L. Bianco, G. Perrotta, R. Santos, S. Mazzuca, Purification of intact chloroplasts from marine plant Posidonia oceanica

re

suitable for organelle proteomics, Proteomics 15 (2015) 4159-4174.

lP

56. G. Friso, W. Majeran, M. Huang, Q. Sun, K.J. Van Wijk, Reconstruction of metabolic pathways, protein expression, and homeostasis machineries across maize bundle sheath

na

and mesophyll chloroplasts: large-scale quantitative proteomics using the first maize genome assembly, Plant physiol. 152 (2010).1219-1250. 57. B.S. Phinney, J.J. Thelen, Proteomic characterization of a triton-insoluble fraction from

ur

chloroplasts defines a novel group of proteins associated with macromolecular structures.

Jo

J Proteome Res, 4 (2005) 497-506. 58. R.G. Bayer, S. Stael, E. Csaszar, M. Teige, Mining the soluble chloroplast proteome by affinity chromatography. Proteomics 11 (2007) 1287-1299. 59. Z.H. He, H.W. Li, Y. Shen, Z.S. Li, H. Mi, Comparative analysis of the chloroplast proteomes of a wheat (Triticum aestivum L.) single seed descent line and its parents. J Plant Physiol. 170 (2013) 1139-1147. 60. I. Fesenko, A. Seredina, G. Arapidi, V. Ptushenko, A. Urban, I. Butenko, S. Kovalchuk, K. Babalyan, A. Knyazev, R. Khazigaleeva, E. Pushkova, The Physcomitrella patens chloroplast proteome changes in response to protoplastation. Front. Plant Sci. 7 (2016) 1661.

Journal Pre-proof 61. M. Ferro, S. Brugière, D. Salvi, D. Seigneurin-Berny, L. Moyet, C. Ramus, S. Miras, M. Mellal, S. Le Gall, S. Kieffer-Jaquinod, AT_CHLORO, a comprehensive chloroplast proteome database with subplastidial localization and curated information on envelope proteins, Mol. Cell. Proteomics 9 (2010) 1063-1084. 62. J.B. Peltier, Y. Cai, Q. Sun, V. Zabrouskov, L. A. Giacomelli, Rudella, A.J. Ytterberg, H. Rutschow, K.J. van Wijk, The oligomeric stromal proteome of Arabidopsis thaliana chloroplasts, Mol. Cell. Proteomics 5 (2006) 114-133. 63. H. Rutschow. A.J. Ytterberg, G. Friso, R. Nilsson, K.J. van Wijk, Quantitative

of

proteomics of a chloroplast SRP54 sorting mutant and its genetic interactions with

ro

CLPC1 in Arabidopsis, Plant Physiol. 148 (2008) 156-175.

64. G. Friso, L. Giacomelli, A.J. Ytterberg, J.B. Peltier, A. Rudella, Q. Sun, K.J. van Wijk,

-p

In-depth analysis of the thylakoid membrane proteome of Arabidopsis thaliana chloroplasts: new proteins, new functions, and a plastid proteome database, Plant Cell 16

re

(2004) 478-499.

lP

65. L. Giacomelli, A. Rudella, K.J. van Wijk, High light response of the thylakoid proteome in Arabidopsis wild type and the ascorbate-deficient mutant vtc2-2. A comparative

na

proteomics study, Plant Physiol, 141 (2006) 685-701. 66. J.B. Peltier, A.J. Ytterberg, Q. Sun, K.J. van Wijk, New functions of the thylakoid membrane proteome of Arabidopsis thaliana revealed by a simple, fast, and versatile

ur

fractionation strategy, J. Biol. Chem. 279 (2004) 49367-49383

Jo

67. M. Ferro, D. Salvi, S. Brugière, S. Miras, S. Kowalski, M. Louwagie, J. Garin, J. Joyard, N. Rolland, Proteomics of the chloroplast envelope membranes from Arabidopsis thaliana, Mol. Cell. Proteomics 2 (2003) 325-345. 68. J.E. Froehlich, C.G. Wilkerson, W.K. Ray, R.S. McAndrew, K.W. Osteryoung, D.A. Gage, B.S. Phinney, Proteomic study of the arabidopsis thaliana chloroplastic envelope membrane utilizing alternatives to traditional two-dimensional electrophoresis, J. Proteome Res. 2 (2003) 413-425. 69. M. Schubert, U.A. Petersson, B.J. Haas, C. Funk, W.P. Schröder, T. Kieselbach, Proteome map of the chloroplast lumen of Arabidopsis thaliana, J. Biol. Chem. 277 (2002) 8354-8365.

Journal Pre-proof 70. S. Simm, D.G. Papasotiriou, M. Ibrahim, M.S. Leisegang, B. Müller, T. Schorge, M. Karas, O. Mirus, M.S. Sommer, E. Schleiff, Defining the core proteome of the chloroplast envelope membranes, Front. Plant Sci. 4 (2013) 11. 71. E. Gutierrez-Carbonell, D. Takahashi, G. Lattanzio, J. Rodríguez-Celma, J. Kehr, . Soll, . Philippar,

.

emura, . Abad a, A. .

pe - ill n, The distinct functional roles of

the inner and outer chloroplast envelope of Pea (Pisum sativum) as revealed by proteomic approaches. J. Proteome Res. 13 (2014) 2941-2953. 72. I. Bouchnak, S. Brugiere, L. Moyet, S. Le Gall, D. Salvi, M. Kuntz, M. Tardif, N.

of

Rolland, Unravelling hidden components of the chloroplast envelope proteome:

ro

opportunities and limits of better MS sensitivity. Mol. Cell. Proteomics (2019) 118. 73. A. Brautigam, S. Hoffmann-Benning, A.P. Weber, Comparative proteomics of

-p

chloroplast envelopes from C3 and C4 plants reveals specific adaptations of the plastid envelope to C4 photosynthesis and candidate proteins required for maintaining C4

re

metabolite fluxes, Plant Physiol 148 (2008) 568-579.

lP

74. K. Manandhar-Shrestha, B. Tamot, E.P. Pratt, S.M. Saitie, A. Braeutigam, A.P. Weber, S. Hoffmann-Benning, Comparative proteomics of chloroplasts envelopes from bundle

na

sheath and mesophyll chloroplasts reveals novel membrane proteins with a possible role in C4-related metabolite fluxes and development, Frot. Plant Sci. 4 (2013) 65. 75. M. Ploscher, V. Reisinger, L.A. Eichacker, Proteomic comparison of etioplast and

ur

chloroplast protein complexes. J Proteomics 74 (2011) 1256-1265.

Jo

76. P. Schafer, S. Helm, D. Kohler, B. Agne, S. Baginsky, Consequences of impaired 1-MDa TIC complex assembly for the abundance and composition of chloroplast high-molecular mass protein complexes. PLoS One, 14 (2019) 0213364. 77. S. Helm, D. Dobritzsch, A. Rodiger, B. Agne, S. Baginsky, Protein identification and quantification by data-independent acquisition and multi-parallel collision-induced dissociation mass spectrometry (MSE) in the chloroplast stroma proteome. J Proteomics 98 (2014) 79-89. 78. P. Dominic, B. Olinares, L. Ponnala, K.J. van Wijk, Megadalton complexes in the chloroplast

stroma

of

Arabidopsis

thaliana

characterized

by

size

exclusion

chromatography, mass spectrometry, and hierarchical clustering. Mol. & Cell. Proteomics, 9 (2010) 1594-1615.

Journal Pre-proof 79. L. Yin, A.V. Vener, C. Spetea, The membrane proteome of stroma thylakoids from Arabidopsis thaliana studied by successive in‐solution and in‐gel digestion, Physiol. Plant. 154 (2015) 433-446. 80. R. Srivastava, T. Pisareva, B. Norling, Proteomic studies of the thylakoid membrane of Synechocystis sp. PCC 6803, Proteomics 5 (2005) 4905-4916. 81. M. Tomizioli, C. Lazar, S. Brugière, T. Burger, D. Salvi, L. L. Gatto, Moyet, L.M. Breckels, A.M. Hesse, K.S. Lilley, Deciphering thylakoid sub-compartments using a mass spectrometry-based approach, Mol. Cell Proteomics 13 (2014) 2147-2167.

of

82. Q. Meng, L. Rao, X. Xiang, C. Zhou, X. Zhang, Y. Pan, A systematic strategy for

ro

proteomic analysis of chloroplast protein complexes in wheat. Biosci. Biotechnol. Biochem. 75 (2011) 2194-2199.

-p

83. J. Melonek, S. Oetke, K. Krupinska, Multifunctionality of plastid nucleoids as revealed by proteome analyses. Biochim Biophys Acta. 1864 (2016) 1016-1038.

re

84. W. Majeran, G. Friso, Y. Asakura, X. Qu, M. Huang, L. Ponnala, K.P. Watkins, A.

lP

Barkan, K.J. Van Wijk, Nucleoid-enriched proteomes in developing plastids and chloroplasts from maize leaves: a new conceptual framework for nucleoid functions.

na

Plant Physiol. 158 (2012) 156-189.

85. A.J. Ytterberg, J.B. Peltier, K.J. Van Wijk, Protein profiling of plastoglobules in chloroplasts and chromoplasts. A surprising site for differential accumulation of

ur

metabolic enzymes. Plant Physiol. 140 (2006) 984-997.

Jo

86. W. Xu, H. L. M. Zhao, Y. Li, Y. Qi, Z. Peng, G. Xia,. M. Wang, Proteomic comparison reveals the contribution of chloroplast to salt tolerance of a wheat introgression line. Sci. Rep. 6 (2016) 32384.

87. P.Y. Yousuf, A. Ahmad, I. M. Aref, M. Ozturk, A.H. Ganie, M. Iqbal, Salt-stressresponsive chloroplast proteins in Brassica juncea genotypes with contrasting salt tolerance and their quantitative PCR analysis, Protoplasma 253 (2016) 1565-1575. 88. F. Meng, Q. Luo, Q. Wang, X. Zhang, Z. Qi, F. Xu, X. Lei, Y. Cao, W.S. Chow, G. Sun, Physiological and proteomic responses to salt stress in chloroplasts of diploid and tetraploid black locust (Robinia pseudoacacia L.), Sci. Rep. 6 (2016). 89. A. Joaquín-Ramos, J.A. Huerta-Ocampo, A. Barrera-Pacheco, A. De León-Rodríguez, S. Baginsky, A.P.B. de la Rosa, Comparative proteomic analysis of amaranth mesophyll and

Journal Pre-proof bundle sheath chloroplasts and their adaptation to salt stress. J Plant Physiol. 171 (2014) 1423-1435. 90. S. Muneer, Y.G. Park, A. Manivannan, P. Soundararajan, B.R. Jeong, Physiological and proteomic analysis in chloroplasts of Solanum lycopersicum L. under silicon efficiency and salinity stress, Int. J. Mol. Sci. 15 (2014) 21803-21824. 91. L. Wang, W. Liang, J. Xing, F. Tan, Y. Chen, L. Huang, C. L. Cheng, W. Chen, Dynamics of chloroplast proteome in salt-stressed mangrove Kandelia candel (L.) Druce, J. Proteome Res. 12 (2013) 5124-5136.

of

92. L. Wang, D. Pan, J. Li, F. Tan, B.S. Hoffmann, W. Liang, W. Chen, Proteomic analysis

salt tolerance, Plant Sci. 231 (2015) 159-172.

ro

of changes in the Kandelia candel chloroplast proteins reveals pathways associated with

-p

93. B.K. Phee, J.H. Cho, S. Park, J.H. Jung, Y.H. Lee, J.S. Jeon, S.H. Bhoo, T.R. Hahn, Proteomic analysis of the response of Arabidopsis chloroplast proteins to high light

re

stress, Proteomics, 4 (2004) 3560-3568.

lP

94. J. Wang, Q. Yu, H. Xiong, S. Chen, Z. Yang, S. Dai, Proteomic Insight into the Response of Arabidopsis Chloroplasts to Darkness, PLoS One 11 (2016) 0154235.

na

95. E. Uberegui, M. Hall, O. Lorenzo, W.P. Schröder, M. Balsera, An Arabidopsis soluble chloroplast proteomic analysis reveals the participation of the Executer pathway in response to increased light conditions. J Exp. Bot. 66 (2015) 2067-2077.

ur

96. L. Gao, X. Wang, Y. Li, R. Han, 2019. Chloroplast proteomic analysis of Triticum

Jo

aestivum L. seedlings responses to low levels of UV-B stress reveals novel molecular mechanism associated with UV-B tolerance, Environ. Sci. Pollut. Res. (2019) 1-13. 97. D. Gayen, P. Barua, N.V. Lande, S. Varshney, S. Sengupta, S. Chakraborty, N. Chakraborty, Dehydration-responsive alterations in the chloroplast proteome and cell metabolomic profile of rice reveals key stress adaptation responses, Environ. Exper. Bot. 160 (2019) 12-24. 98. R. Tamburino, M. Vitale, A. Ruggiero, M. Sassi, L. Sannino, S. Arena, A. Costa, G. Batelli, N. Zambrano, A. Scaloni, Chloroplast proteome response to drought stress and recovery in tomato (Solanum lycopersicum L.), BMC Plant Biol. 17 (2017) 40.

Journal Pre-proof 99. A.

osmala, D. Perli ows i, I. Pawłowic ,

. Rapac , Changes in the chloroplast

proteome following water deficit and subsequent watering in a high-and a low-droughttolerant genotype of Festuca arundinacea, J. Exp. Bot. 63 (2012) 6161-6172. 100.

F. Grimaud, J. Renaut, E. Dumont, K. Sergeant, A. Lucau-Danila, A.S. Blervacq,

H. Sellier, N. Bahrman, I. Lejeune-Hénaut, B. Delbreil, Exploring chloroplastic changes related to chilling and freezing tolerance during cold acclimation of pea (Pisum sativum L.), J. Proteomics 80 (2013) 45-159. 101.

F. Grimaud, J. Renaut, E. Dumont, K. Sergeant, A. Lucau-Danila, A.S. Blervacq,

of

H. Sellier, N. Bahrman, I. Lejeune-Hénaut, B. Delbreil, E. Goulas, Exploring

ro

chloroplastic changes related to chilling and freezing tolerance during cold acclimation of pea (Pisum sativum L.). J Proteomics 80 (2013) 145-159.

S. Andaluz, A.F. López-Millán, J. De las Rivas, E.M. Aro, J. Abadía, A. Abadía,

-p

102.

Photosyn. Res. 89 (2006) 141-155.

N. Ahsan, Y. Nanjo, H. Sawada, Y. Kohno, S. Komatsu, Ozone stress‐induced

lP

103.

re

Proteomic profiles of thylakoid membranes and changes in response to iron deficiency,

proteomic changes in leaf total soluble and chloroplast proteins of soybean reveal that

10 (2010) 2605-2619. 104.

na

carbon allocation is involved in adaptation in the early developmental stage, Proteomics

V. Velikova, A. Ghirardo, E. Vanzo, J. Merl, S.M. Hauck, J.P. Schnitzler, Genetic

ur

manipulation of isoprene emissions in poplar plants remodels the chloroplast proteome. J

105.

Jo

Proteome Res. 13 (2014) 2005-2018. E. Megias, L.S.T. do Carmo, C. Nicolini, L.P. Silva, R. Blawid, T. Nagata, A.

Mehta, 2018. Chloroplast proteome of Nicotiana benthamiana infected by tomato blistering mosaic virus. Protein J. 37(2018) 290-299. 106.

K. Lohrig, B. Müller, J. Davydova, D. Leister, D.A. Wolters, Phosphorylation site

mapping of soluble proteins: bioinformatical filtering reveals potential plastidic phosphoproteins in Arabidopsis thaliana. Planta 229 (2009) 1123-1134. 107.

N. Sugiyama, H. Nakagami, K. Mochida, A. Daudi, M. Tomita, K. Shirasu,

Ishihama, Y. Large‐ scale phosphorylation mapping reveals the extent of tyrosine phosphorylation in Arabidopsis. Mol. Syst. Biol. 4 (2008).

Journal Pre-proof 108.

S. Reiland, G. Messerli, K. Baerenfaller, B. Gerrits, A. Endler, J. Grossmann, W.

Gruissem, S. Baginsky, Large-scale Arabidopsis phosphoproteome profiling reveals novel chloroplast kinase substrates and phosphorylation networks. Plant Physiol. 150 (2009) 889-903. 109.

H. Sun, X. Liu, F. Li, W. Li, J. Zhang, Z. Xiao, L. Shen, Y. Li, F. Wang, J. Yang,

First comprehensive proteome analysis of lysine crotonylation in seedling leaves of Nicotiana tabacum. Sci. Rep. 7(2017) 3013. 110.

S. Liu, C. Xue, Y. Fang, G. Chen, X. Peng, Y. Zhou, C. Chen, G. Liu, M. Gu, K.

of

Wang, K. W. Zhang, Global involvement of lysine Crotonylation in protein modification

111.

ro

and transcription regulation in Rice. Mol. Cell. Proteomics 17 (2018) 1922-1936. M. Hartl, M. Fubl, P.J. Boersema, J.O. Jost, K. Kramer, A. Bakirbas, J.

-p

Sindlinger, M. Plochinger, D. Leister, G. Uhrig, G.B. Moorhead, Lysine acetylome profiling uncovers novel histone deacetylase substrate proteins in Arabidopsis. Mol. Syst.

W.V. Bienvenut, C. Giglione, T. Meinnel, SILProNAQ: a convenient approach

lP

112.

re

Biol. 13 (2017).

for proteome-wide analysis of protein N-termini and N-terminal acetylation quantitation.

113.

na

Methods Mol Biol. 1574 (2017) 17-34.

E. Rowland, J. Kim, N.H. Bhuiyan, K. J. van Wijk, K.J. The Arabidopsis

chloroplast stromal N-terminome: complexities of amino-terminal protein maturation and

C. Alban, M. Tardif, M. Mininno, S. Brugière, A. Gilgen, S. Ma, M. Mazzoleni,

Jo

114.

ur

stability. Plant Physiol. 169 (2015) 1881-1896.

O. Gigarel, J. Martin-Laffon, M. Ferro, S. Ravanel, Uncovering the protein lysine and arginine methylation network in Arabidopsis chloroplasts. PLoS One 9 (2014) 95512. 115.

D. Salvi, S. Bournais, L. Moyet, I. Bouchnak, M. Kuntz, C. Bruley, N. Rolland,

AT_CHLORO: the first step when looking for information about subplastidial localization of proteins. Methods Mol Biol. 1829 (2018) 395-406. 116.

C. Bruley, V. Dupierris, D. Salvi, N. Rolland, M. Ferro, AT_CHLORO: a

chloroplast protein database dedicated to sub-plastidial localization. Front. Plant Sci. 3 (2012) 205. 117.

F. Myouga, K. Akiyama, R. Motohashi, T. Kuromori, T. Ito, H. Iizumi, R.

Ryusui, T. Sakurai, K. Shinozaki, The Chloroplast Function Database: a large‐ scale

Journal Pre-proof collection of Arabidopsis Ds/Spm‐ or T‐ DNA‐ tagged homozygous lines for nuclear‐ encoded chloroplast proteins, and their systematic phenotype analysis. Plant J 61(2010) 529-542. 118.

F. Myouga, K. Akiyama, Y. Tomonaga, A. Kato, Y. Sato, M. Kobayashi, N.

Nagata, T. Sakurai, K. Shinozaki, The Chloroplast Function Database II: a comprehensive collection of homozygous mutants and their phenotypic/genotypic traits for nuclear-encoded chloroplast proteins. Plant Cell Physiol. 54 (2013) 2-e2. 119.

T. Kleffmann, M. Hirsch-Hoffmann, W. Gruissem, S. Baginsky, plprot: A

of

comprehensive proteome database for different plastid types. Plant Cell physiol. 47

120.

ro

(2006) 432-436.

Q. Sun, B. Zybailov, W. Majeran, G. Friso, P.D.B. Olinares, K.J. van Wijk,

121.

-p

PPDB, the plant proteomics database at Cornell. Nucleic Acids Res 37 (2008) 969-974. P. Gloaguen, S. Bournais, C. Alban, S. Ravanel, D. Seigneurin-Berny, M.

re

Matringe, M Tardif, M. Kuntz, M. Ferro, C. Bruley, N. Rolland, ChloroKB: a web

lP

application for the integration of knowledge related to chloroplast metabolic network. Plant Physiol. 174 (2017) 922-934.

Q. Sun, O. Emanuelsson, K.J. van Wijk, Analysis of curated and predicted plastid

na

122.

subproteomes of Arabidopsis. Subcellular compartmentalization leads to distinctive proteome properties. Plant Physiol.135 (2004) 723-734. S. Schaeffer, A. Harper, R. Raja, P. Jaiswal, A. Dhingra, Comparative analysis of

ur

123.

Jo

predicted plastid-targeted proteomes of sequenced higher plant genomes. PLoS One 9 (2014), 112870. 124.

R. Kaundal, S.S. Sahu, R. Verma, T. Weirick, October. Identification and

characterization of plastid-type proteins from sequence-attributed features using machine learning. BMC Bioinformatics 14 (2013) S7. 125.

E. Schleiff, L.A. Eichacker, K. Eckart, T. Becker, O. Mirus, T. Stahl, J. Soll,

Prediction of the plant β‐ barrel proteome: a case study of the chloroplast outer envelope. Protein Sci. 12 (2003) 748-759. 126.

I. Small, N. Peeters, F. Legeai, C. Lurin, Predotar: a tool for rapidly screening

proteomes for N‐ terminal targeting sequences. Proteomics 4 (2004) 1581-1590.

Journal Pre-proof 127.

O. Emanuelsson, H. Nielsen, G. Von Heijne, ChloroP, a neural network-based

method for predicting chloroplast transit peptides and their cleavage sites. Protein Sci. 8 (1999) 978-984. 128.

P. Horton, K.J. Park, T. Obayashi, N. Fujita, H. Harada, C. J. Adams-Collier, K.

Nakai, WoLF PSORT: protein localization predictor. Nucleic Acids Res. 35 (2007) 585587. 129.

W.L. Huang, C.W. Tung, S.W. Ho, S.F. Hwang, S.Y. Ho, ProLoc-GO: utilizing

informative Gene Ontology terms for sequence-based prediction of protein subcellular

H.N. Lin, C.T. Chen, T.Y. Sung, S.Y. Ho, W.L. Hsu, December. Protein

ro

130.

of

localization. BMC Bioinformatics 9 (2008) 80.

subcellular localization prediction of eukaryotes using a knowledge-based approach.

131.

-p

BMC Bioinformatics 10 (2009) S8.

T. Blum, S. Briesemeister, O. Kohlbacher, MultiLoc2: integrating phylogeny and

132.

lP

Bioinformatics 10 (2009) 274.

re

Gene Ontology terms improves subcellular protein localization prediction. BMC

S. Briesemeister, T. Blum, S. Brady, Y. Lam, O. Kohlbacher, H. Shatkay,

na

SherLoc2: a high-accuracy hybrid method for predicting subcellular localization of proteins. J Proteome Res. 8 (2009) 5363-5366. 133.

M. Ryngajllo, L. Childs, M. Lohse, F.M. Giorgi, A. Lude, J. Selbig, B. Usadel,

ur

SLocX: Predicting Subcellular Localization of Arabidopsis Proteins Leveraging Gene

134.

Jo

Expression Data. Front. Plant Sci., 2 (2011) 43. O. Emanuelsson, H. Nielsen, S. Brunak, G. von Heijne, Predicting subcellular

localization of proteins based on their N-terminal amino acid sequence. J Mol. Biol. 300 (2000) 1005-1016 135.

R. Kaundal, G.P.S. Raghava, RSLpred: an integrative system for predicting

subcellular localization of rice proteins combining compositional and evolutionary information. Proteomics 9 (2009) 2324-2342. 136.

A.K. Berglund, E. Spånning, H. Biverståhl, G. Maddalo, C. Tellgren-Roth, L.

Mäler, E. Glaser, Dual targeting to mitochondria and chloroplasts: characterization of Thr–tRNA synthetase targeting peptide. Mol. Plant 2 (2009) 1298-1309.

Jo

ur

na

lP

re

-p

ro

of

Journal Pre-proof

Journal Pre-proof Table 1: Optimized percentage gradients used for isolation of intact chloroplasts across different species. Osmotic agents in HB

Gradient

20-day old

Sorbitol

45 & 85% *

55-day old

Sorbitol

40 & 85% *

20 g

Sorbitol

10, 30 & 50%*

3g

Sorbitol

50%#

8 week old

Sorbitol

50%#

Pea

8 day old

Sorbitol

40 & 80%*

Pea

7 day old

Sorbitol

50%#

Sorbitol

40 & 80%*

Sucrose

40 & 80%*

Wheat Wheat Maize Amaranthus cruentus Chickpea

Rice Tomato

Sorbitol

10000g, 10 min 3750g, 10 min 8000g, 20 min 2000g, 10 min 10700g, 6 min 10000g 10 min 8000g, 10 min 10000g 10 min 3000g, 5min

Chloroplast collected from Interface Interface 30-50 Interface Pellet

40 & 85%*

Pellet

ro

-p

re

lP

Wheat

na

Wheat

8-9 day old Flag leaves 3 leaf stage (10 g) 14 day old (15-20 g) 12 day old 50 g 10 day old 3rd leaf (80-140 g) Fresh leaves (30 g) 21 day old seedling (100 g) 21 day old seedling (100 g) 21 day old (30 g)

Interface pellet Interface

References Simm et al., 2013 Dominic et al., 2010 Fan et al., 2008 Kley et al., 2010 Bayer et al., 2011 Simm et al., 2013 Phinney et al., 2004 Bayer et al., 2011

Interface

He et al., 2013

47400g, 15 mim

Interface

Xu et al., 2016 Meng et al., 2014 Kamal et al., 2012

Sorbitol

20 & 45%*

-

Lower band

Sorbitol

40%#

8000g, 10 min

Pellet

Sorbitol

88 & 35%*

4740g, 15 min

Interface

Majeran et al., 2012

Sorbitol

80 & 40%*

2700g, 15 min

Interface

Ramos et al., 2014

Sorbitol

40%#

1700g, 7 min

Pellet

Lande et al., 2017

Sorbitol

40%#

1700g, 7 min

Pellet

Gayen et al., 2019

Sorbitol

50%#

13000g, 10min 3200g, 15 min

ur

Pea

Jo

Arabidopsis thaliana Arabidopsis thaliana Arabidopsis thaliana Arabidopsis thaliana Arabidopsis thaliana

Centrif ugation

of

Sample (Leaves)

Plant

Pellet

Tobacco

30 day old

Sorbitol

40 & 80%*

Interface

Brassica juncea Medicago sativa

20 day old (10 g)

Sorbitol

40 & 85%*

4100g

Interface

20 day old

Sorbitol

42 & 82%*

10000g, 10 min

Interface

Tamburino et al., 2017 Megias et al., 2018 Yousu et al., 2016 Simm et al., 2013

Journal Pre-proof Black locust

30 g

Sucrose

10, 40, 70 & 90%*

Populas

100 g

Mannitol

40%#

Kandelia candel

10 g

Sorbitol

40 & 80%*

Sorbitol

40 & 80%@

Sorbitol

30 & 50%*

10 g

Sorbitol

10, 30, & 50%*

1g

Sorbitol

60%#

Festuca arundinacea Salicornia europaea Posidonia oceanica

4 month old 10 g Fully expanded

40-70 Interface Pellet Interface Interface Interface 30-50 Interface Lower band

of

Poplar

40000g, 30 min 3000g, 6 min 16000g, 20 min 10000g, 10 min 3200g, 15 min 8000g, 20 min 13300g, 10 min

Meng et al., 2016 Yuan et al., 2011 Wang et al., 2013 Velikova et al., 2005 Kosmala et al., 2012 Fan et al., 2008 Piro et al., 2015

Jo

ur

na

lP

re

-p

ro

H : Homogeni ation buffer ‘*’represents Percoll step gradient; ‘#’ represents Percoll continuous gradient; ‘@’ represent sucrose step gradient

Journal Pre-proof Table 2: A list of published papers on chloroplast proteomics accomplished through gel-based and gel-free techniques.

Fraction

Stress

Chloroplast Light

-p

Stroma

re

Arabidopsis

lP

Stroma thylakoids

ur

Stroma, thylakoids & envelope

1-DE

na

Thylakoid

Jo

Envelope Plastoglobule Chloroplast

Pea

Wheat

1-DE 2-DE 2-DE BN-PAGE BN-PAGE 2-DE DIGE In-solution 1-DE In-solution 2-DE In-Solution Tricine-SDSPAGE 1-DE In-gel & Insolution

Number of proteins 690 154 15 436 381 52 29 81 128 110 392 1011

Kleffmann et al., 2004 Kley et al., 2010 Fan et al., 2009 Behrens et al., 2013 Schafer et al., 2019 Phee et al., 2004 Uberegui et al., 2015 Wang et al., 2016 Ferro et al., 2003 Simm et al., 2013 Froehlich et al., 2003 Helm et al., 2014

241

Peltier et al., 2006

1081

Dominic et al., 2010

58

Yin et al., 2015

ro

Chloroplast envelope

Methods used

of

Species

Cold

Chloroplast envelope Inner & outer envelope Lumenal & peripheral Thylakoid

In-gel & Insolution 1-DE In-Sol 1-DE 1-DE DIGE In-solution 1-DE In-solution 1-DE, 2-DE

Chloroplast Salt UV-B

Tricine-SDSPAGE 2-DE BN-PAGE 2-DE 2-DE 2-DE

1295(65grana, 450 stroma) 1323 (337stroma, 113thylakoid, 163-envelope) 1269 54 448 179 6, 39 87-IE, 73-OE 322 546(180IE,141-OE)

Reference

Tomizioli et al., 2014

Ferro et al., 2010 Bouchnak et al., 2019 Ytterberg et al., 2006 Bayer et al., 2011 Phinney et al., 2004 Grimaud et al., 2013 Simm et al., 2013 Brautigam et al., 2008 Carbonell et al., 2014

200-lumenal, 230-peripheral

Peltier et al., 2000

767

Kamal et al., 2012

16 20 65 26 28

He et al., 2013 Meng et al., 2011 Kamal et al., 2012 Xu et al., 2016 Gao et al., 2019

Journal Pre-proof

Envelope Chloroplast Chloroplast Chloroplast BS&M chloroplast Etioplast & chloroplast Thylakoid membranes Chloroplast Chloroplast Chloroplast

Beta vulgaris Brassica juncea Soybean Black locust Capsicum annuum Kandelia candel Physcomitrella patens Posidonia oceanica Festuca arundinacea Cyanobacterium

280 (21-BS, 36 M )

Shrestha et al., 2013

1-DE BN-PAGE

448 23

Majeran et al., 2012 Muneer et al., 2014

2-DE

31, 54

Tamburino et al., 2017

119

Yuan et al., 2011

119

Velikova et al., 2006

Dehydration Biotic

In-solution BN-PAGE, Insolution In-solution 1-DE In-solution 2-DE

Salt

Abiotic

Fedeficiency Salt Ozone Salt

Plastoglobule Chloroplast

Salt

ur

Barley

1-DE spectral count Salt Waterdeficit

71 2451 40 20

Simm et al., 2013 Lande et al., 2017 Gayen et al., 2019 Megias et al., 2018

2-D, BN/PAGE

77

Ramos et al., 2014

DIGE

68

Ploscher et al., 2011

55

Andaluz et al., 2006

12 32 61

Yousuf et al., 2016 Ahsan et al., 2010 Meng et al., 2016

In-Sol

27

Ytterberg et al., 2006

2-DE

46 1030 (76 DEPs) 479 (219 DEPs)

Wang et al., 2015

2-DE, BN/PAGE 2-DE 2-DE 2-DE

In-solution

Wang et al., 2013

Chloroplast

In-solution

Chloroplast

1-DE

72

Piro et al., 2015

2-DE

10

Kosmala et al., 2012

1-DE, 2-DE

76

Srivastava et al., 2005

Jo

Amaranthus

Majeran et al., 2005 Friso et al., 2010

of

Medicago sativa Chickpea Rice Tobacco

221, 100, 305 954,882

ro

Chloroplast

2-DE BN-PAGE

-p

Poplar

Brautigam et al., 2008

re

Chloroplast

231

lP

Tomato

1-DE

na

Maize

Chloroplast envelopes BS & M chloroplast BS & M chloroplast envelope Nucleoid

Chloroplast Thylakoid membranes

Waterdeficit

Fesenko et al., 2016

1-DE- One-dimensional electrophoresis, 2-DE- Two-dimensional electrophoresis, BN-PAGE- Blue Native Polyacrylamide gel electrophoresis, DIGE- Differential gel electrophoresis, IE- inner envelope, OE- Outer envelope, BS- Bundle Sheath, M- Mesophyll, DEP- Differentially expressed protein

Jo

ur

na

lP

re

-p

ro

of

Journal Pre-proof

Journal Pre-proof Table 3. A list of published chloroplast protein localization prediction tools.

Reference Small et al., 2004 Emanuelsson et al., 1999 Horton et al., 2007 Huang et al., 2008 Lin et al., 2009 Blum et al., 2009 Briesemeister et al., 2009 Ryngajllo et al., 2011 Emanuelsson et al., 2000 Kaundal et al., 2009

ur

na

lP

re

-p

ro

of

Localization prediction Chloroplastic or non chloroplastic Chloroplastic or non chloroplastic different organelles different organelles different organelles different organelles different organelles different organelles Extracellular, mitochondrial, chloroplast Chloroplast, cytoplasm, mitochondria, nucleus

Jo

Tool Name Predotar ChloroP WoLF PSORT ProLoc-GO KnowPredsite MultiLoc2 SherLoc2 SlocX TargetP RSLPred

Journal Pre-proof Table 4. Number of proteins in the non-redundant chloroplast protein datasets of individual plant species, curated from the reports available so far.

of

ro

re

Arabidopsis Chickpea Maize Pea Physcomitrella patens Poplar Kandelia candel Wheat Tomato Amaranthus Medicago sativa Posidonia oceanica Black locust Common beet

Number of nonredundant proteins 3629 2451 1438 892 369 119 79 93 79 70 65 57 56 55

-p

Organism

Jo

ur

na

lP

Barley Rice Capsicum annuum Tobacco Soybean Brassica Festuca arundinacea

46 40 27 20 19 12 10

Journal Pre-proof Graphical abstract:

Highlights This review summarizes strategies for chloroplast isolation and sub-fractionation.



Described classical and emerging techniques for chloroplast proteome profiling.



The proteome inventory revealed known and novel uncharacterized proteins.

Jo

ur

na

lP

re

-p

ro

of

