Proteomics has emerged as a vital tool to identify the novel proteins and explore the cellular dynamics by employing highly proficient and innovative techniques introduced in the past few years. The expedition began with the emergence of gel-based 2DE approaches for evaluation of diverse proteins and differential protein analysis. Later, advancements in the field lead to the advent of gel-free approaches that render more accuracy and reliable results. In spite of introduction of high-throughput analysis offered by new techniques, the overall beneficence of gel-based approaches cannot be ignored. The combined effort of all these approaches can generate astonishing results. Protein analysis under specific conditions attempts to deduce the expression pathways and thus will help to expand our knowledge pertaining to metabolic and regulatory routes. Further, understanding the dynamics of these biomolecules and metabolites involved in the pathways can facilitate their desired manipulation for altering the gene expression. This will assist in clinical purposes for development of drugs and in systems biology to get the broad picture of metabolic processes and their regulation. This review highlights the trend with which progress in plant proteomics technology has been made right from gel based to gel free strategies with discussion of basic principles and procedures involved in each technique. Advances in this field can lead to precise interpretation of several biological processes.

Keywords: 2DE; DIGE; iTRAQ; ICAT; SILAC; plant proteomics; quantitative proteomics

Proteomics refers to the high throughput systematic analysis of protein expression and function with the aid of protein biochemistry, mass spectrometry and bioinformatics' tools. Most of the biological researches aim to decipher the metabolic regulatory pathways to gain deeper insight into various cellular processes and thus contributing to our understanding of biological systems. Although, there are ongoing efforts to gather information contained in the human genome sequence through genomics and transcriptomics approaches; elucidating the dynamic changes in proteins will provide a better picture of cellular genetics and its regulation. In this context, quantitative proteomics has been emerged recently as a fascinating platform to explore the hidden pathways and reveal fundamentals of biological systems. With the advent of the most feasible and spectacular techniques, quantitative proteomics is directed to exemplify the identification and quantification of diverse proteins that represent a rich source of biological information. Quantitative proteomics employs various techniques for separation of proteins based on their physicochemical properties (molecular mass, pH, charge etc) followed by their recognition and quantification by mass spectrometry methods utilizing data systems and software's. The major steps involved in these analytical processes are i) cell sampling ii) extraction, isolation and solubilisation of proteins iii) separation through electrophoresis (isotachophoresis, zonal, isoelectric focussing, off gel) and chromatography (ion exchange chromatography, 2- dimensional liquid chromatography, reverse phase) iv) quantification using gel based and gel free approaches and v) identification using mass spectrometry approaches. The chief principles and procedure of aforementioned steps are briefly illustrated in (a,b,c,d,e). For detailed understanding of MS protein identification principles, readers are advised to visit http://www.spectroscopynow.com/ ; http://www.ionsource. com/; http://www.asms.org/whatisms/index.html. Many advances have been made so far in the field of quantitative proteomics, however there is no technique that demonstrates the actual expression of these biological entities (proteins) in one go as in case of genomics (microarray analysis, genome sequencing). Proteomics studies require the collective efforts of different fields of sciences (biochemistry, physics, computer sciences and statistics) that collaborate to analyse structural and functional aspects of proteins. In this review, we have highlighted various efficient and reliable quantification methods that differ in many ways in terms of progression, sensitivity, robustness, accuracy and quality of data obtained. Quantitative mass spectrometry approaches can be mainly categorized into gel based (2- dimensional electrophoresis) and gel-free approaches. Later are further divided to label-based (chemical, enzymatic and metabolic tagging) and label-free (data independent and data dependent) methods. Each technique has its pros and cons and rather than replacement, these methods complement each other. The choice of methodology adopted for analysis depends on the biological sample taken under consideration and the information required. In order to make successful systematic and quantitative profiling, we require good quality separation and quantitative techniques accompanied with protein/ nucleotide databases that promise high accuracy to search peptide masses with the assistance of powerful automated bioinformatic tools and softwares. Here, we have summarized the basic strategies involved in the aforementioned techniques that expedite the functional analysis of proteins on global scale and thus assisting the conventional molecular biology methods.

Genes are considered to be the major repositories of biological information required for the processing of molecular mechanisms inside the cell. However, the information stored in the gene is in coded form and needs to be expressed and translated to proteins for performing vital functions of life. The genes are transcribed to mRNA in coded form of nucleotides that are further translated and decoded to amino acid sequence of proteins. Thus, for proper investigation of cellular processes, studies can be conducted at genomics (genes), transcriptomics (mRNA) and proteomics (protein) level. With the emergence of sequencing technique, it is possible to deduce the DNA sequence in short time but focus has now been shifted to predict the innumerable functions which are performed by the expression components of these genes i.e., Proteins. The study of cell at genomic and transcriptomic level provides a rough estimate of the expression and genome annotation. Thus study of proteins is important as these are the ultimate bio molecules that complement the structure and function of living systems. The concept of proteome was given by Marc Wilkins and his associates to describe the protein complement of the genome [1]. The large scale study of diverse proteins and precise measurement of their expression by utilizing powerful analytical techniques is referred to as "Proteomics" [2]. The wide acceptance of this recently emerged technology is attributed to advancements in the methodologies that attempt to accomplish accurate identification and quantification of proteins. The expression of genes can be detected by mRNA profiling but it neither reflects the correct amount of proteins nor their regulatory status [3]. Transcripts are highly unstable and are translated differently under varying environmental conditions producing different proteins. There is a dire need to analyse multiple protein products arising from single gene to figure out expression pathways thoroughly. Various post-translational modifications occur that affect the structure, localization and function of proteins and presents challenges in front of analytical tools to detect multiple forms of proteins [4]. A gene expressed under different set of environmental conditions generates separate transcripts which are further modified chemically and expressed in various ways to produce several forms of proteins. Thus, relying completely on genomics tools is not a good option. Expression studies should be carried out at proteome level to attain the complete knowledge of ongoing cellular processes. The first proteomic work in plants was published on evaluation of proteins in Arabidopsis thaliana using 2-dimensional gel electrophoresis [5]. The proteomic studies have made excellent progress in the past few years, right from the sub-cellular proteome expression analysis of various organelles [6-11]. And understanding the developmental process by spot comparison [12-14]. To deciphering the proteins involved in stress physiology [15-24] and plant pathogen interactions [25-28]. The detailed inspection of several proteins synthesized by various intriguing metabolic processes in relation to the external environment will help to assign functions to orphan genes and understanding their regulation ultimately serving the purpose. As proteins are directly involved in the molecular networks, accurate changes can be measured in response to alternating surroundings. Table 1 represents the key studies carried out on proteomic analysis of plants using advanced gel based and gel free methodologies. Thus, proteomics approaches can be considered as the essential key to unlock mysteries of life and its processes finally governing the efficient manipulation of pathways for improvement of crops.

Gel based protein separation and quantification is performed by integrating simple analytical methods of protein biochemistry with high throughput mass spectrometry analysis. The first report in this context was published about three decades ago by O'Farrel for analysis of complex proteins [68]. Since then, tremendous progress has been made in the field of protein studies due to continued development of methodologies in terms of precision and accuracy. This technique allows global analysis of thousands of protein isoforms expressing under specific set of conditions [4]. Gel based approach mainly employs 2D-PAGE (Two-dimensional polyacrylamide gel electrophoresis) and 2D-DIGE (Two dimensional differential gel electrophoresis) that efficiently separates proteins on the basis of their physicochemical properties, differentially analyses the spots obtained and finally identifies the components using mass spectrometry. Figure 2 (a, b) represents basic steps involved in gel based proteomics approach. In spite of huge advancements in the field of proteomics, these techniques have not been replaced and are routinely used for quantification of proteins.

2D-PAGE (Two Dimensional Polyacrylamide Gel Electrophoresis): Electrophoresis technique (separation of charged molecules under the influence of electric current) has been extensively utilized for the separation of biomolecules [69] based on their specific characteristics. Two dimensional approach attempts to separate proteins depending on two parameters i.e., pH and molecular mass. Former employs IEF (iso-electric focusing) technique and later utilizes polyacrylamide gels in electrolytic medium subjected to current under the influence of electric field. The second dimension represents better resolution and precision

to analyse complex proteins in the sample. First of all, the sample (tissue/cell) is collected and stored under suitable conditions and used to extract proteins free from impurities. Then proteins are solubilised using ionic/non-ionic detergents and reducing agents (DTT) and finally separated. Iso-electric focusing requires IPG strips (Immobilized pH gradient strips) that contain acrylamide polymerized with bis-acrylamide and immobilins covalently bind in gel. These immobilins were used to create stable pH gradient [70] inside the gel. These strips are dipped in the solubilisation buffer containing extracted proteins of the sample. Both positively and negatively charged proteins (amphoteric) have specific pI (iso-electric point) at which the charge of the protein is neutralized. Different amphoteric molecules get focused on the gel corresponding to their respective pI when subjected under electric field. The strips are further treated with SDS and chemicals like DTT, iodoacetamide for equilibration of their chemical properties and thus leaving proteins to be analysed only on the basis of their molecular masses. The hydrocarbon tail in the SDS (sodiumdodecyl sulphate) binds to the hydrophobic amino acids through non-covalent interactions and gets distributed over entire proteins imparting negative charge all over and causes denaturation [71]. The proteins then travel from porous polyacrylamide gel (PAGE) according to their molecular weight under the influence of electric current. Different pH of buffer systems present more accurate resolution due to isotachophoresis in which proteins are aligned on the stacking gel (less pH) and further separated onto resolving gel (high pH) thus giving a fair start to all proteins. The buffer systems possess leading and trailing ions whose electrophoretic mobility is affected by the external pH. Some of the most commonly used buffer systems are Tris-glycine, Bis-Tris, Tris-acetate and Tris-Tricine. Next step after resolution is the detection of the proteins which can be achieved by in-gel staining methods. The most commonly used chemicals for staining are coomassie brilliant blue, silver nitrate and fluorescent dyes that differ in terms of sensitivity, utility and suitability for MS analysis. Coomassie brilliant blue and fluorescent dyes bind noncovalently to the proteins and can be separated easily, thus suitable for MS analysis but silver staining involves covalent linking to the proteins thus not recommended for MS analysis in spite of its high sensitivity than other dyes [72]. Fluorescent dyes such as Deep Purple, SYPRO Ruby, SYPRO Red, SYPRO Orange, RuBPS, FlamingoTM, Krypton TM, ASCQ Ru, ProQdiamond, ProQemerald etc present very high sensitivity and accurate determination of protein isoforms in the sample due to their efficient compatibility with MS [73]. Stained gels are visualized using scanners/ detectors and differential spots are taken into consideration for further analysis (depicted in Figure 2a). The abundance of proteins is roughly assessed by the intensity of spots. However, for absolute detection, these spots are excised from the gel and subjected to enzymatic digestion producing variable peptides that undergo ionization in MS and analysed based on their different mass to charge ratios. The speed with which these peptides move towards detector is critically observed and fed to data systems. The data is then used to search matches in the databases (protein/ nucleotide) for peptide mass fingerprinting using powerful automated softwares for identification of proteins [74]. This technique ensures the utilization of cost-effective technologies for adequate resolution of complex proteins and can be used along with the advanced tandem MS approaches as the first fractionation step. However, it renders errors due to greater number of steps involved and low proteome coverage due to insolubility of highly acidic or basic, hydrophobic proteins. The detection of spot intensity permits inaccuracy due to resolution of diverse proteins in the same location and thus prevents the low abundant proteins to be detected [4,75-77]. In addition to this, different experimental treatment of samples and gel-to-gel variation also contribute to the poor reproducibility of 2D-PAGE. These drawbacks are overcome by 2D-DIGE technique that allows resolution on single gel rather than separate gels.

This modified gel based technique offers few advantages over 2D-PAGE by minimizing the errors caused by gel–to-gel variation and need for analysis of more than one gel thereby reducing manual error. The protein samples are pre-stained by using cyanine-based fluorescent dyes. The NHS (N-hydroxysuccinimidyl) ester group and maleimide derivatives of these dyes react with the amino and thiol groups of the protein respectively. The labelling is performed in two ways- minimal labelling and saturation labelling [78]. Former deals with the labelling of N'- terminal of lysine residues by amide linkage to NHS ester group (required for maintaining the multiple charges on the surface of protein thereby preventing in solubilisation) and later facilitates the binding of thiol groups of cysteine residues to the maleimide derivatives of dyes (recommended for low abundant proteins due to high sensitivity) leading to comparatively wide proteome coverage. The cyanine based dyes should be tagged such that they might not influence the mobility of proteins when subjected to electrophoresis [79]. The labelling of different samples with resolvable fluorescent cyanine based dyes allows differential expression studies. The protein samples to be quantified are labelled with Cy3 and Cy5 dyes that impart different colours when visualized by fluorescence scanner and thus depict the amount of protein within the sample by measurement of spot intensity (Illustrated in Figure 2b). These labelled proteins along with the internal standard Cy2 (representing presence of both the samples) is used for normalization of the ratios of intensities retrieved from different samples paving way for accurate quantification of proteins. The labelled protein samples are mixed and subjected to electrophoretic separation in a single gel thereby eliminating the need to analyse more gels and reducing the experimental error. The stained images are captured, scanned, digitalized and the fluorescence intensities of variable samples are analysed and the data is fed to efficient softwares such as DeCyder, Proteom weaver PDQuest and Progenesis [70] for comparison of spot intensities and useful information is generated depicting to the abundance of specific proteins. DIGE technique is more appealing as compared to 2D-PAGE in terms of sensitivity, reproducibility, reliability, accuracy, automation, and more suitability for more diverse proteins & MS analysis. The gel based techniques have been applied for differential protein expression studies in plants [17,80-84]. And few research findings emphasized the equivalent need of gel based techniques to accomplish the task of protein annotation [85-88]. Regardless of so many advantages of 2D-DIGE, it is unable to beat some of the immanent drawbacks of gel based approaches like narrow-coverage of proteins due to tagging of lysine and cysteine amino acids only, insolubility of some of the membrane proteins and sample preparation variation.

Advancements in the field of proteomics have led to the emergence of gel-free proteomics approach that addresses the issues such as reproducibility, low-proteome coverage, quality of data obtained that are observed in case of gel-based methods. Gelfree methods are mainly dependent on LC-MS/MS technique and instead of examining one spot at a time, it takes into consideration all the peptides generating from the proteins proving to be more robust and extremely informative high-throughput strategy.

Gel free methods can be considered as a direct consequence of the numerous innovative developments in the past two decades. These methods eliminate few of the major experimental errors observed in case of gel based approaches. Gel free approaches can be mainly classified into two categories i.e., label-based approaches and label free approaches [89]. These methods utilize the proficient liquid chromatography and mass spectrometry tools to generate the quantification data thereby promising efficient protein studies. The gel free strategy is quite simple and engages few steps in which proteins are isolated, digested (labelled/non-labelled), eluted on liquid chromatography and detected to be analyzed by mass spectrometry approaches for absolute or relative quantification. Wide methodologies pertaining to label-based and label-free approaches are discussed in the following sections.

Label-based approaches utilize specialized isotope/isobar tags having specific groups that label proteins and peptides chemically/metabolically or enzymatically [90]. The steps undertaken for label-based quantification are depicted in Figure 3 a & b. The separation of these labelled peptides on LC and further analyses by highly sensitive MS technique yields highly informative data for retrieving precise results by comparing the relative abundances of heavy and light samples. Numerous experiments have been conducted using label based strategies to understand the mechanisms of various developmental stages, stress physiology by differential expression studies [26,28,91- 95]. The label based approaches assure automated highthroughput quantitative proteome analysis of unknown proteins with reliable automated and multiplexing abilities. Proteins/

peptides are labelled either chemically (using isotopes or isobars) or metabolically to introduce mass shifts for distinguishing the relative intensities of the proteins in the samples.

Chemical labelling: The method involves utilization of chemically synthesized tags that incorporate variable isotopes and isobars which introduce mass difference within the labelled proteins (ICAT) and peptides (ICPL, iTRAQ, TMT) for their differential expression studies based on the abundance of peptides detected by their peak intensities (based on m/z) and MS/MS fragmentation. This strategy offers accuracy and simultaneous comparison of more than two samples. It has been further sub-divided into isotopic (ICAT & ICPL) and isobaric (iTRAQ & TMT) labelling.

Isotope-Coded Affinity Tags (ICAT): The first in vitro method that permits tagging of proteins and peptides of all types of biological samples using stable isotopes was developed by Gygi and his associates in 1999 [96]. This technique employs ICAT reagents that consist of mainly iodo-acetamide group or N-ethymaleimide, a spacer or linker arm and biotin. Iodoacetamide group/N-ethymaleimide is highly specific chemical reactive group that alkylates thiol group of cysteine residues in the protein sample. A spacer or linker arm is meant for introduction of mass shift by incorporation of different isotopes in different samples. Isotopes used for this purpose are proton (H)/ deuterium (D), ¹²C/¹³C, ¹⁵N/¹⁶N that introduce mass difference of upto 8Da due to presence of these elements in the labelled or unlabelled amino acid residues producing light and heavy tags. Biotin group assists purification of labelled peptides by affinity chromatography in biotin-avidin/streptavidin systems and thus captures all the cysteine containing peptides from the mixture. The strategy employs isolation of protein from two samples followed by synthetic chemical labelling using ICAT reagents; one with heavy and other with light isotopes. The labelled samples are then pooled, enzymatically digested to peptides and subjected to affinity purification by biotin moiety to reduce sample complexity. The next step involves the removal of biotin as it decreases the resolution efficiency of mass spectrometry [96] using acid-cleavable (ALICE/ introduction of disulphide bond in linker) or photo-cleavable linkers (UV light) [97,98]. The peptides are then allowed to resolve on liquid chromatographic separations and further analysed by tandem mass spectrometric techniques. Relative peak intensities obtained in MS spectra directly correlates to the abundance of respective peptides in the sample whereas tandem MS allows peptide mass fingerprinting by detection of product ions generated during peptide fragmentation which leads to identification of proteins. The technique offers accuracy as samples are similarly treated by protease preventing experimental variations and reduced sample complexity due to tagging of only cysteine residues but is also associated with loss of information leading to lower proteome coverage. To overcome these limitations, other tags have been developed that allow multiplexing as well.

Isotope-Coded Protein Labelling (ICPL): The ICPL strategy, referred to as modified version of ICAT approach was developed by Schmidt and co-workers that solved major shortcomings of confined sequence coverage and low throughput [99]. It utilizes amine-reactive N-nicotinoyloxy-succinimide tags that cause derivatization of free amino-terminal groups and ɛ-amino groups of lysine residues and introduce mass difference of 4 Da and ~6 Da in case of H/D and ¹²C₆/¹³C₆ labelling respectively. The first step involves extraction, reduction and alkylation of protein samples to ensure uniform labelling of all free amino terminals of lysine residues. The heavy and light labelled protein samples are then pooled and subjected to enzymatic digestion. Since lysine residues are labelled, treatment with trypsin will generate longer fragments, thus Glu-C endoproteinases in addition to trypsin are used for digestion to obtain shorter fragments. ICPL tags are hydrophilic in nature and help to maintain the intrinsic characteristics of peptides that allow efficient quantification. The isotopes used in ICPL tags can be used in varied combinations for multiplexing (triplex and quadruplex) and differentiates the sample by even 2 Da. This method permits labelling of lysine containing peptides that are found abundant as compared to cysteine residues in most of the proteins and eliminates the confined sequence coverage of proteome to some extent but not completely as lysine is also absent in some of the proteins. To overcome this issue, post-digest ICPL approach was developed that allows labelling of peptides after enzymatic digestion. All the free N-terminals of peptides are labelled uniformly and thus the results obtained are non-biased and considered to be more accurate [100]. The digested labelled peptides are separated on liquid chromatography and finally analysed on MS for quantification and identification of proteins. ICPL strategy is observed to be very suitable for efficient MS/MS fragmentation and detection of peak intensities and also offers the analysis of post-translational modifications and isoforms [101]. Despite of advantages over ICAT method, post digest ICPL involves labelling after digestion which can lead to manual error and the tags can interfere in the mobility of peptides when subjected to liquid chromatography [102]. In addition to isotopes, isobars have been employed for the quantification purpose that can introduce mass difference of even 1 Da and efficient multiplexing (iTRAQ & TMT).

Isobaric Tags for Relative and Absolute Quantification (iTRAQ): ITRAQ strategy has been utilized efficiently to explore the diverse molecular mechanisms occurring in plants. The strategy involves isobaric labelling of peptides; introduced by Ross and his associates in 2004 [103]. The isobaric tags bind covalently to the N-terminal of peptides; introduce a mass shift of even 1Da and thus paves way for multiplexing. The tag is comprised of three main functional elements i) Reporter group that introduces mass shifts ii) Balance group that is required to maintain the overall mass of isobaric tag iii) NHS group which specifically binds to the peptide. Chemically, reporter group is N-methylpiperizine which provides a mass shift range; balance group is mainly carbonyl group whose mass is adjusted according to the reporter group so that all tags have same mass for combined reporter and balance group and NHS ester group is amine reactive. The isobars are employed in varied combinations leading to efficient comparison of two, four and eight samples simultaneously which is otherwise not possible in case of ICAT strategy. Reporter groups have a mass range varying from 114-117 Da and 113-121 Da that are compensated by balance group having a range from 28-31 Da and 184-192 Da in 4-plex (mass tag = 145 Da) and 8-plex (mass tag = 305 Da) respectively. Thus, the overall mass of reporter and balance group remains constant (as shown in Figure 3 a). The strategy includes labelling of enzymatically digested protein samples in which NHS-group binds covalently to all the N-termini of peptides equally. The labelled peptides are then further analysed by LC-MS/MS. All the labelled peptides are resolved by chromatographic separation and detected by MS to generate spectra. However, the peptides remain unresolved as second round fragmentation of peptides is necessary for producing product ions associated with release of reporter ions. For this purpose high quality MS with triple quadrupole is used. The ions entering are fragmented by less collision energies to produce precursor ions which cannot be distinguished and presented as a single peak. These precursor ions are again introduced under the influence of high collision energies to give product ions that are detected and resolved to generate spectra for simultaneous protein quantification and identification when retrieved information is searched against available protein and nucleotide databases. Although iTRAQ strategy has been utilized extensively in the past years due to its high accuracy and multiplexing abilities, it is associated with the need of high-throughput data acquisition system and modified versions of MS that can read slight mass differences.

Tandem mass tag: This technique shares the same principle as iTRAQ strategy with slight variation in the chemical structure of the tag used for labelling. ¹³C/¹⁵N isotopomers are mainly used in varying proportion to create mass difference. The tag is comprised of reporter region (creates mass difference), linker region (conjugates reporter to balance group and is easily cleavable), balance group (maintains constant mass) and protein reactive group (binds to amine/cysteine/carbonyl) [104]. The reporter group mass varies from 126-131 Da which is maintained to a constant mass of 230 Da by balance group having mass ranging from 99-104 Da which leads to 6-plex analysis. Similar to iTRAQ, this technique involves efficient uniform labelling of peptides (amine-) and cysteine residues (cys TMT) after enzymatic digestion of proteins. The reporter ions are released at the time of peptide fragmentation in MS/MS, produce spectra that is recorded and finally the abundance of peptides is interpreted that allows protein identification and relative quantification.

Although chemical labelling presents accuracy in determination of peptide abundance and overcomes in-gel experimental variation, it requires careful sample preparation methods and highly efficient mass spectrometric analysis which can discriminate peptides varying by even 1 Da mass. Multiplex versions are undoubtedly presenting proficient comparison but are associated with complicated data analysis due to its inability to select peptides after one round of fragmentation. The major drawback of these techniques is that pooling of samples is done just before LC-MS/MS analysis which creates space for experimental biasness and inaccuracy. To overcome the limitations of in vitro techniques, metabolic labelling came in limelight that incorporates the tags in the samples from the very beginning.

Enzymatic labelling: This method employs substitution of natural (16O) and isotopic oxygen (18O) in the carboxyl groups of amino acid residues. The isolated proteins are digested by proteases that target serine, lysine and arginine residues in presence of heavy water (H₂¹⁸O) and light water (H₂¹⁶O). Later, Hcl was used to serve as catalyst for labelling of carboxyl terminal residues along with water (H₂¹⁸O/ H₂¹⁶O) referred to as acid mediated oxygen substitution [105]. During the enzymatic digestion, amide bond is broken and one isotopic oxygen atom is substituted in carboxyl group. The cleaved peptide undergoes one more substitution in place of second oxygen of carboxyl group in presence of enzyme creating three possibilities for mass differences i.e., ¹⁶O/¹⁶O (0Da), ¹⁶O/¹⁸O (2Da), ¹⁸O/¹⁸O (4Da) when compared to unlabelled peptides. The mass differences are detected to depict the relative abundance of peptides by comparing their ionic intensities. This method allows efficient incorporation of tags but permits side reactions that interferes with accurate data analysis. However, to inhibit undesirable reactions, suitable buffers and esterification is performed methanol and deuterated methanol [106].

Metabolic labelling: Metabolic labelling strategy employs biological incorporation of isotopic amino acids and elements through cell culture in plants and dietary food in animals. It surpasses the major drawback of in vitro labelling and eliminates experimental error to a great extent. After the intake of labelled amino acids inside the body, the cells are rapidly multiplied and undergo vast array of cellular processes that ensures efficient incorporation of isotopes. Metabolic labelling can be carried out by either isotopic essential amino acids (arginine, lysine, leucine, and tyrosine) or isotopic elements (¹³C, ¹⁵N, ²H, ¹⁸O). Despite of so many advantages, it lacks applicability for all biological samples and is somewhat tedious and expensive.

Stable Isotopic Labelling of Amino Acids in Cell Culture (SILAC): SILAC is a simple in vivo technique that was first developed in 2002 [107]. Later on, the method was efficiently introduced in eukaryotic organisms as well. Isotopic lysine (C₆H₁₄N₂O₂) and arginine (C₆H₁₄N₄O₂) amino acid tags are mainly used in this strategy. Looking at the chemical structure, it can be estimated that isotopic and normal amino acid will have a mass difference of 6Da (¹²C/¹³C), 2 Da (¹⁴N/¹⁵N) and likewise varied combinations of isotopic amino acids will lead to simultaneous high-throughput analysis of more samples. First of all, a cell culture medium is prepared and cells whose proteome is to be analysed are grown. The medium is divided into two sections, one is provided with labelled amino acids and other with unlabelled amino acids (as shown in Figure 3b). The cells are allowed to divide for four to five generations after subculturing in presence of similar growth regulators and conditions to confirm the unbiased incorporation of amino acid residues. To validate the efficient labelling of cells with heavy isotopes, mass spectrometric analysis is performed. The cells from both the culture medium are then pooled and proteins are extracted, treated with reducing agents and alkylated using iodoacetamide. The reduced proteins are then treated with trypsin for digestion to peptides. This can also be performed after separation of reduced proteins on SDS-PAGE. The mixture of peptides is co-eluted in liquid chromatography (reverse phase or strong ion exchange chromatography). The separated fractions are then further analysed by mass spectrometric techniques. This technique ensures less chance of biasness and handling errors with 100% incorporation of tags if grown for sufficient time [108]. SILAC strategy can provide absolute quantification of proteins provided there is no undesirable metabolic conversion of labelled amino acids to other by products (eg arginine is converted to proline). Except the labelled amino acids, all other amino acids present in sample should be non-isotopic and present in sufficient amount to eliminate the probability of side reactions. The technique has been modified to identify post-translational modifications, especially methylation by using heavy-methyl SILAC approach [109]. The technique has certain limitations associated to suitability of biological material in question and laborious steps involving highly proficient tools and expensive synthesises of tags.

¹⁵N labelling: The first metabolic labelling study was performed using isotopically enriched media containing 15N and is observed to be appropriate mainly for prokaryotes. The technique utilizes same strategy as SILAC with the difference of incorporating isotopic elements instead of amino acids. The labelled and unlabelled elemental nitrogen is provided to the growing cells by introducing inorganic salts in the culture medium (as shown in figure 3b). The cells are grown separately in the medium containing all the essential components required for growth. One sample is provided with the isotopic labelled element (heavy ¹⁵N) and other is grown in presence of natural Nitrogen element (light ¹⁴N). Multiple division of cells is allowed for few generations and then samples are mixed, protein is extracted, reduced, alkylated and digested by trypsin. The labelled and unlabelled peptides are then eluted on chromatographic separation and finally analysed by MS to generate ion chromatograms depicting intensities that are directly proportional to their relative abundances. The technique facilitates incorporation of ~98% tags but lacks in precision due to variation in number of nitrogen in peptides and also peptide sequences. Thus it makes the analysis and interpretation complicated.

Culture-Derived Isotope Tags (CDITs): This technique provides absolute and relative quantification of proteomes by using labelled internal standards. This strategy has been derived from SILAC technique as it is also associated with in vivo introduction of isotopes in cultured cell [110]. The cells are chosen from the tissue (taken under consideration for protein quantification). These cells are grown in suitable culture medium in presence of isotopes, thus referred to as culture-derived isotope tags (CDITs). The CDITs are mixed with the tissue sample which is to be analysed. The strategy further allows combined extraction of proteins followed by reduction and digestion to peptides. Now, the peptides from labelled (CDITs) and unlabelled (tissue sample) are analysed by mass spectrometry. Ion chromatograms are generated that depict m/z ratio of labelled and unlabelled peptides, isotopic distribution of peptides is observed and thus the quantity of the peptide in question is estimated. The calculated ratio of sample (m/z of same sequence of labelled peptides serving as reference and unlabelled peptides of tissue) depicts the absolute abundance of respective peptide. In case of more than one sample, same CDITs are added to all the different samples serving as internal standard. Protein from different samples (T₁/T₂/T₃/T₄ + CDITs) is extracted and digested separately. Mass to charge ratio for all the samples having internal standards are calculated which shows isotopic distribution between CDITs and tissue sample. The number of ratios calculated is equal to the number of samples addressed and these calculated ratios are finally compared to estimate the relative abundance of peptides (as depicted in Figure 3b).

To date, a lot of experiments have been conducted and published for biological studies in plants using label-based approaches due to wide applicability and accuracy. However, this method involves many steps and the samples that can be analysed are limited. Moreover, handling errors could lead to incomplete incorporation of tags and give way to side reactions. These constraints have been taken away by more advanced labelfree approaches that allow direct analysis on LC and comparison on the basis of mass spectrometric data.