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Proteomics 2014, 14, 579–592 579DOI 10.1002/pmic.201300240

REVIEW

Quantitative analysis of protein turnover in plants

Clark J. Nelson1,2, Lei Li1,2 and A. Harvey Millar1,2

1 ARC Centre of Excellence in Plant Energy Biology, University of Western Australia, WA, Australia2 Centre for Comparative Analysis of Biomolecular Networks, University of Western Australia, WA, Australia

Proteins are constantly being synthesised and degraded as plant cells age and as plants grow,develop and adapt the proteome. Given that plants develop through a series of events fromgermination to fruiting and even undertake whole organ senescence, an understanding ofprotein turnover as a fundamental part of this process in plants is essential. Both synthesisand degradation processes are spatially separated in a cell across its compartmented structure.The majority of protein synthesis occurs in the cytosol, while synthesis of specific componentsoccurs inside plastids and mitochondria. Degradation of proteins occurs in both the cytosol,through the action of the plant proteasome, and in organelles and lytic structures throughdifferent protease classes. Tracking the specific synthesis and degradation rate of individualproteins can be undertaken using stable isotope feeding and the ability of peptide MS to tracklabelled peptide fractions over time. Mathematical modelling can be used to follow the isotopesignature of newly synthesised protein as it accumulates and natural abundance proteins asthey are lost through degradation. Different technical and biological constraints govern thepotential for the use of 13C, 15N, 2H and 18O for these experiments in complete labelling andpartial labelling strategies. Future development of quantitative protein turnover analysis willinvolve analysis of protein populations in complexes and subcellular compartments, assessingthe effect of PTMs and integrating turnover studies into wider system biology study of plants.

Keywords:

Isotopic labelling / MS / Plant proteomics / Protein turnover / Synthesis

Received: June 18, 2013Revised: October 2, 2013

Accepted: October 14, 2013

1 Introduction

In the early 20th century biologists began to speculate thatperhaps proteins were being continuously synthesised anddegraded. Research in the 1930s pointed towards the idea ofcontinual protein synthesis in plants [1, 2] and then usingstable isotopes studies in tobacco and sunflower establishedthat continual protein turnover did occur [3,4]. Later studies,using radiolabelling, measured total protein turnover in var-ious plants tissues, obtaining more quantitative estimates oftotal protein turnover [5–9]. While such studies are useful and

Correspondence: Dr. A. Harvey Millar, The University of WesternAustralia, M316, 35 Stirling Highway, Crawley WA 6009, WesternAustralia, AustraliaE-mail: [email protected]: +61 8 64884401

Abbreviations: AAA, ATPases associated with diverse cellular ac-tivities; BN PAGE, blue native PAGE; OXPHOS, oxidative phos-phorylation; PS, photosystem

provide insight into whole plant physiology or allow a relativecomparison of genotypes or developmental stages, they arestill of limited utility in explaining physiological responsesof plants. More detailed information on turnover of specificproteins provides much greater insight into the biologicalprocesses at play in a tissue. The unique features of the au-totrophic life cycle of plants, such as germination from dryseeds, organ senescence and large temperature and hydrationcycles, provide new scenarios for analysis of protein turnovernot found in animals. The imperative to increase agricul-tural efficiency and product quality in the coming decadeswill require a thorough understanding of protein dynamicsas proteins represent a significant part of the cost of cell main-tenance, a key nitrogen storage product in plants and also aquality trait for nutrition and food processing. Central con-trollers of protein dynamics are likely to be targets in cropbreeding programs.

Early turnover studies of individual proteins focused onenzymes involved in primary metabolism such as nitrate re-ductase, phenylalanine ammonia lyase and invertase [10]. A

C© 2013 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim www.proteomics-journal.com

580 C. J. Nelson et al. Proteomics 2014, 14, 579–592

Figure 1. The role of protein synthe-sis and degradation in plant gene func-tion. With systems level approaches,global measurement of mRNA changesare measured using modern technolo-gies such as microarrays or RNAseq.Changes in protein abundance canbe measured using DIGE, iTRAQ andMRM. Below, synthesis and degrada-tion of transcripts can be monitored us-ing pulse labelling with 4-thiouridinecoupled to deep sequencing for tran-script counting. Using stable-isotopelabelling and MS biologists can mea-sure the relative isotope abundance(RIA) along a time course (T0, T1, T2,T3). By fitting an exponential decaycurve, protein degradation and proteinsynthesis rates can be calculated.

number of studies also focused on the photochemical ma-chinery, photosystem I (PSI) and PSII, with the thrust ofmuch of the work being that protein D1 of PSII is the labilecomponent and that degradation and synthesis are correlatedwith stress and lighting conditions of plants [11–13]. How-ever, it was soon clear that other PSII subunits such as D2and CP43 could also turn over rapidly in response to envi-ronmental stimuli [14]. Chlorophyll was also found to be nec-essary to stabilise PSII after its formation [14–16]. Analysesof the carbon-fixing RUBISCO enzyme complex found thatthe nuclear-encoded small subunit and plastid-encoded largesubunit are maintained in tight stoichiometry with excess,uncomplexed proteins rapidly degraded [17].

With the advent of a systems biology approach and asso-ciated techniques, the way biologists have conducted theirinvestigations has changed dramatically. Today, investigatorstry to measure all components from the central dogma ofbiology from transcription to translation, and in the case ofprotein dynamics protein catabolism as well (Fig. 1). Microar-rays made transcriptomics the first robust systems level ap-proach, able to measure transcripts from tens of thousandsof genes in a single experiment and with the development ofnext generation sequencing, this approach is even more sen-sitive, fast and robust [18]. Attention has now turned fromtranscript abundance to considering transcript synthesis rate[19, 20], degradation rate [19–21] and isolation of RNA frompolysomal fractions to uncover the RNAs being translated[22].

Although high volume proteomics was slower to develop,given the dramatic improvements in proteomics technolo-gies including modern MS instrumentation, gel-based sepa-rations, isotopic-labelling techniques and software to analysesuch data it is now feasible to measure protein changes inthousands of protein in a single experiment [23]. For relative

quantitation, one can use gel-based methods such as differen-tial gel electrophoresis for quantifying proteins. Alternatively,there are MS-based techniques that are label free and mea-sure the number of times peptides from a respective proteinare observed (i.e. spectral counting) while other methods usestable-isotope labels and MS to compare changes in proteinabundance. These topics in plants have been relatively re-cently reviewed [24, 25]. In one noteworthy report to date inplants, 13 029 proteins were identified in Arabidopsis acrossseveral different plant tissues, which resulted in the charac-terisation of 4000 organ-specific proteins [26]. For absolutequantitation, stable isotope labelling of representative pep-tides [27] or entire proteins works well [28].

However, with any of these techniques as they are currentlyapplied in plants, comparisons are made between genotypesor treatments, and only the change in abundance of a pro-tein is measured. Because protein abundance is a balancebetween synthesis and degradation, it remains unclear if anychanges observed were the result of an alteration in synthesis,degradation, or both. To develop a complete understandingof biological phenomena, a firm grasp of these molecularmechanisms is necessary. New MS technologies, based onthe introduction of stable isotopes into biological systems,now make it feasible to isolate and measure both synthesisand degradation for proteins in situ (Fig. 1). In this review,we will briefly discuss the molecular machinery in plants in-volved in protein synthesis and degradation, summarisingwhat is known thus far regarding protein turnover in plants.We then review the stable isotope techniques available to plantbiologists while presenting advantages and disadvantages ofdifferent methodologies as well as additional plant-specificconsiderations for these types of experiments. Finally, wesuggest a few interesting new directions for this rapidly de-veloping sub-discipline of proteomics.


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Figure 2. Protein synthesis and degrada-tion within plant cells and their compart-ments. Protein synthesis processes areshown by red arrows. Proteins are synthe-sised by cytosolic ribosome and translo-cated into the ER or transported into or-ganelles including nucleus, peroxisome,mitochondria and plastids. A small num-ber of proteins are synthesised by mito-chondrial and chloroplast ribosome. Se-creted proteins, and some chloroplast pro-teins, are channelled through the ER andGolgi apparatus, where PTMs may occur.Proteins from the Golgi are transported toother sub-cellular locales via transportingvesicles. Protein degradation processesare marked by black arrows. Cytosolic andnuclear proteins are degraded by the ubiq-uitin/proteasome pathway. Compartment-specific proteases degrade organellarproteins. Damaged organelles or excessorganelles can be degraded by autophagy.The mTOR pathway is a regulator of thebalance between protein synthesis anddegradation.

2 Protein synthesis

2.1 Synthesis in the plant cytosol

In plant cells, most polypeptides are synthesised by cytosolicribosomes and then transported to their respective destina-tions via targeting motifs (Fig. 2). The cytosolic ribosomeconsists of a large 60S subunit, a small 40S subunit and fourrRNA species and are conserved from yeast to mammals, withthe exact protein composition and macromolecular size oflarge and small subunits varying by taxa [29]. In Arabidopsis,more than 200 genes were found to encode for 81 cytosolicribosome proteins [30]. The synthesis rate (Ks) for a givenprotein can be regulated by various factors during translationon ribosomes including: initiation, elongation and termina-tion. Protein–protein and protein–RNA interactions betweentranslational factors and ribosomal proteins or rRNA can alsocontribute to this translational control [31]. Upstream ORFsin mRNA may also affect protein synthesis by influencingre-initiation of downstream ORFs [29]. In eukaryotes, trans-lation usually occurs in a cap-dependent manner, meaningthat a combination of proteins interacts with the 5′ modi-fied nucleotide of the transcript and this facilitates ribosomebinding. This is also true in plants, but there are notice-able differences in the cap-binding complex and the proteincomplement of this complex varies by developmental stagein plants [32]. Alternatively, translation can occur through acap-independent manner in which the ribosome can bypassscanning of the cap and enter directly at the start site when

the eIF2-� protein becomes phosphorylated in response tostress [33]. In plants, cap-independent initiation also occursin response to abiotic stresses but notably not all of the com-ponents in this machinery are conserved between animalsand plants [34].

All these factors in combination influence protein syn-thesis rate and thus protein abundance in vivo. In onestudy, a genome-wide scale investigation of mRNA and pro-tein turnover in mammalian cells established that transla-tion plays the dominant role in determining protein abun-dance under steady-state conditions [19]. It remains to bedetermined to what extent translation contributes to pro-tein abundance changes under non-steady-state systems andhow this might vary across taxa. A major regulator of cel-lular metabolism is the target of rapamycin (mTOR), a ser-ine/threonine kinase first characterised in mammals [35, 36]and known to function in plants [37]. Via its influence oninhibitory eIF4E-binding proteins, mTOR is a positive reg-ulator of initiation [38]. Protein synthesis can also be regu-lated in a negative but indirect manner by micro-RNAs andribosome-binding proteins, thereby affecting protein abun-dance [39, 40].

2.2 Synthesis in mitochondria and plastids

While 95% of the mitochondrial and plastid proteomes aresynthesised by the cytosolic ribosomes and imported viatranslocases [41] (Fig. 2), approximately 100 proteins are still



encoded and synthesised within these organelles [42,43]. Mi-tochondrial and plastidic ribosomes are distinct but relatedand consist of large 50S subunits and small 30S subunitsthat together form a bacterial-like 70S ribosome. While thereare many similarities in structure between organellar andbacterial ribosomes, noticeable differences exist. In bacte-ria, translation is often regulated in a negative manner [44],whereas plastid translation is influenced by the presenceof positive factors, increasing translational efficiency, withcontrol typically occurring at initiation or sometimes elonga-tion [44]. Environmental cues such as light can increase ordecrease translation via their effect on the plastid reducingpotential and associated PTMs [44]. Regulation of mitochon-drial translation is less clear. Unlike plastids, plant mitochon-dria have lost the bacterial Shine Dalgarno sequence, whichserves as a ribosome-binding site [45]. Besides the RNA edit-ing role of pentatricopeptide repeat proteins, these proteinshave also been implicated in translational regulation in mito-chondria [46].

3 Protein degradation in plant cells

3.1 Ubiquitin/proteasome pathway

Proteins within cells are continually being degraded, a pro-cess that is highly selective and precisely regulated (Fig. 2).The ubiquitin/proteasome pathway operates in the cytosoland was first discovered in mammals. It shows a high degreeof conservation across eukaryotic taxa, including plants. It isconsidered to be the primary pathway for protein degradationin eukaryotic cells [47,48]. The machinery consists of a seriesof enzymes (E1, E2, E3) which conjugate several ubiquitintags to proteins targeted for catabolism. Once target proteinsare tagged they are then rapidly degraded by the 26S protea-some complex. The importance of this pathway is signifiedby the fact that 6% of the Arabidopsis gene products are as-sociated with E1, E2, E3 subunits and the proteasome [49].This importance is also evident by the integral role theubiquitin/proteasome plays in several hormone signallingcascades [50].

3.2 Protein catabolism in organelles

Intracellular membranes protect organelle proteins fromproteasome-directed degradation. Degradation within or-ganelles such as mitochondria, plastids and peroxisomes is acombination of different degradation pathways. The AAA+(ATPases associated with diverse cellular activities) proteasesare proposed to be the primary players in organellar pro-tein degradation [51]. Multimeric ring structures [52, 53], theAAA+ proteases, including the Clp, FtsH and Lon subclasses,selectively bind, unfold and translocate their substrate in anATP-dependent manner into the proteolytic chamber, and di-gest the substrate into 3–30 amino acid-long peptides [54,55],

which are then degraded by peptidases such as PREP [56,57].In plants, the AAA+ proteases have been found to functionin tissue-specific protein degradation [58], maintenance of or-ganellar protein balance [59] and assembly/maintenance ofthe oxidative phosphorylation (OXPHOS) machinery [54,60].The ubiquitin/proteasome also plays a role in recycling of pro-teins from mitochondria in other systems, but to our knowl-edge there are no known cases in plants. There is one exam-ple of a RING-type E3 ligase regulating the developmentalshift of etioplasts to chloroplasts and chloroplasts to geronto-plasts, suggesting involvement of the ubiquitin/proteasomalmachinery in plastid development [61].

3.3 Autophagy in plant cells

In mammals, autophagy is a catabolic process involving thedegradation of the cell’s own components using the lysosomalmachinery [62, 63]. Organellar components, including mito-chondria, peroxisomes and ER, can also be recycled via au-tophagy and this process is important in protein turnover forthe day-to-day operations of the cell [64]. For example, the au-tophagic turnover of mitochondrial components (mitophagy)is the primary mechanism to degrade dysfunctional mito-chondria in stationary phase yeast [65,66]. A protein turnoverstudy in mice reported that proteins localised within specificorganelles, including the mitochondrion, nucleus and ER,had similar Kd and suggested that autophagy may play animportant role in defining the Kd of these compartments [67].However, a more detailed analysis of murine mitochondrialproteins revealed a much wider range of Kd values with theauthors proposing a smaller or perhaps subtler role for mi-tophagy [68].

Less research has been conducted on plant autophagy,leaving large gaps in our understanding of the mechanis-tic details [69], however, large portions of the recognitionmachinery appears to be conserved across eukaryotic phy-logeny including plants [70]. In contrast to animals, au-tophagy in yeast and plants occurs in the vacuole, not thelysosome [64,70]. The mTOR kinase is also important in reg-ulating autophagic processes in plants [70]. While historicallythe vacuole in plants was perceived as a non-selective dis-posal mechanism, recent studies have demonstrated a moredynamic role for vacuolar autophagy in plants, being involvedin rapid environmental responses. For example, autophagy isimportant in leaf senescence of darkened leaves [71] as well asin the whole plant response to starvation conditions [72] andresponds to the oxidative state of plant cells [73]. However, de-spite its presence, the quantitative importance of autophagyin defining protein turnover in plants is still not clear. Ina study of mitochondrial protein turnover from Arabidopsiscell culture, the authors concluded, similar to the mouse mi-tochondrial study [68], that mitochondria were probably notturned over en masse, suggesting a quantitatively smaller rolefor mitophagy [74].


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Figure 3. Complete and partial labellingstrategies for determining proteinturnover. Two different strategies can beemployed when using isotopic labellingto measure protein dynamics, a completelabelling strategy or a partial labellingstrategy. As examples, spectra for a 12merof averagine are provided. (A) In thecomplete labelling strategy, marked inblack is the natural abundance (NA) and incolour are spectra for 13C, 2H 18O, and 15N.For each spectrum the labelled populationis calculated as 50% enrichment for the re-spective heavy isotope. (B) For the partiallabelling strategy, the example peptideis shown as a natural abundance (NA)spectrum (top) and a theoretical spectrumwith 1.2% 2H label (middle). Using suchan approach, one generates measurablechanges in the isotopic envelope andby plotting the monoisotopic peak as afraction of the first few isotopic peaks(% M) versus time (bottom), a decay ratecan be calculated.

4 Isotopic labelling for measuring proteinturnover

4.1 Use of stable-isotope labelled amino acids

When considering isotopic labelling approaches to study pro-tein turnover, it is desirable for the isotopic label in questionbe present in most if not all peptides and that the organ-ism in question be auxotrophic for the labelled molecule [75].For investigations of mammalian cells or auxotrophic yeastor bacteria lines, use of a SILAC approach [76] is easy toapply and readily adapted for turnover studies [77, 78]. Thisapproach has also been successfully applied in a study of pro-tein kinetics for salinity response in Chlamydomonas, whichwas auxotrophic for arginine [79]. In one proteomic studyof Arabidopsis cell culture a SILAC strategy achieved 80% la-belling of the arginine pool [80]. However, given the dynamicsof plant central metabolism in amino acid synthesis and thatnot all amino acids are transferred equally between sourceand sink tissues in plants, such an approach would be prob-lematic at least and perhaps uninterpretable in intact plants.The high cost of SILAC and the amount of reagents requiredto scale up for experiments on intact plants also makes thisapproach cost prohibitive.

4.2 Use of stable-isotope labelled elements

The autotrophic nature of plants allows researchers to labelthe plant proteome with stable isotopes through introducinglabel into the basic elements present in proteins (P, S, N, C,

O, H). Considering that P is not present in most peptides andthere are no stable isotopes, this element is not an option. ForS, there are stable isotopes but many peptides do not containS, leaving pragmatically only C, N, O and H as candidatelabels.

C represents approximately 30% of the atoms in peptides,and over 50% of their mass. 13C is used in pulse-chase exper-iments in plants to study metabolic labelling, but has beenrarely used for protein labelling over longer time periods. Thisapproach has been successfully applied in the form of labelledfoodstuffs to measure turnover in heterotrophic species [81]and could conceivably be applied to heterotrophic plant tis-sues. One challenge with this approach in autotrophic planttissues is that the label would need to be provided as a 13CO2

label, which is expensive and might be difficult to keep at highlevels due to the more abundant 12CO2 and the challenges in-herent in working with a gas phase label. Because of the largenumber of C atoms in a typical tryptic peptide, analysis ofsuch peptides would result in complicated labelled isotopicenvelopes that contain dozens of isotopes as demonstrated inFig. 3A, and has some associated disadvantages. With suchlarge isotopic envelopes, there is a higher chance that isotopicenvelopes from different peptides may overlap, complicatinganalysis. Additionally, when labelled peptides are of an inter-mediate labelling level, fewer peptides are identified becauseof the difficulties MS/MS search algorithms have with vari-able label incorporation [82]. The envelopes of peptide withintermediate labelling also reduce signal-to-noise because fora typical MS experiment there is a finite amount of a givenpeptide, so that the same amount of peptide is spread overmore isotopic peaks. However, due to the high mass accuracy,resolution, sensitivity and rapid data collection of modern MS



instrumentation this approach is viable. A proof of principlestudy used a custom-built growth enclosure to label planttissues and achieved 91% atom enrichment for the heavy iso-tope in proteins and 95% in metabolites [83], establishingthe viability of this technique. The use of this label mightbe particularly useful where multiple labelling schemes arenecessary or desired.

H is an abundant element, typically representing nearly50% of the atoms in peptides. Though only 7% of their mass,full labelling can increase peptide masses by >50 amu. Akey drawback with this label are the large complicated spec-tra resulting from a near complete 2H labelling strategy [84].Another significant drawback is the physiological changesassociated with high tissue levels of 2H2O [85]. In a priorstudy of Arabidopsis seedlings, 2H2O application changedtranscript expression, which limits its utility when used athigh levels [84]. Despite the disadvantages, there are somesignificant advantages associated with this technique, the firstof which is the relatively low cost of 2H2O. An additional ben-efit of using this label is that it rapidly equilibrates withintissues and sub-cellular compartments but decays slowly[68, 83].

Also worth consideration, O represents approximately 10%of the atoms in peptides. There are two stable isotopes, 17Oand 18O, to choose from but to our knowledge only the lat-ter in the form of H2

18O has been used in protein turnoverstudies, dating back more than 40 years [86]. In a more recentmammalian study investigators reported the 18O was incor-porated effectively and rapidly although not as quickly as for2H2O [87]. There are no known reports of this methodologyin plants but there is no apparent reason this methodologycannot be adopted. However, given the higher cost of H2

18Oas compared to 2H2O as well as the somewhat slower in-corporation into proteins, the deuterated label may be thebetter choice for plant studies if either label is suitable for theexperiment being considered.

Nitrogen is the final element to consider for metabolic la-belling of proteins in plants, and is approximately 10% of theatoms in the average peptide. As with the other labels, thereare drawbacks with the methodology. One potential compli-cation is that labelled peptide envelopes with intermediatelabelling may have large, complex spectra although not aslarge as for C given that there are a third fewer N than Catoms in amino acids. The same difficulties discussed withregards to C labelling apply to N but to a lesser degree, includ-ing the reduction in peptide identifications. However, in onestudy, investigators circumvented the peptide identificationdifficulty by using precise protein-level electrophoretic sepa-rations in combination with peptide accurate mass and LC re-tention time information from analysis of unlabelled samplesto cross extract data from labelled samples in order to quantifylabelled peptide populations [67]. A significant advantage of Nis that the label is easily delivered to plants as inorganic saltsof 15NO3 and/or 15NH4, which can be provided in a measur-able and regulated fashion, and has been applied in multiple

studies of turnover in the last few years [67,74,88–91]. An ad-ditional advantage of this label is that it is as easily applied tocell culture [74,88–90], as to hydroponically grown plants [92].As a more physiologically relevant substrate, sand has alsoproved to be an effective medium for 15N labelling becauseunlabelled nutrients can be easily removed by washing [93],conceivably allowing pulse/chase analysis.

4.3 Partial versus complete labelling

Beyond the choice of element, the second issue to resolve isthe nature of the label swap in terms of the amount of labelapplied, with two different strategies possible. In the first ap-proach, a label is applied that, at least in theory, replaces theentire unlabelled amino acid pool, that is, a complete labelswap. This is the only strategy applied thus far in proteinturnover studies of plants. With this approach, two separateisotopic envelopes are anticipated that can be quantified andcompared to calculate a decay rate for proteins [75, 94, 95] asdemonstrated in Fig. 3A. In reality, the label swap is typicallynot complete and labelled populations have an intermediateand variable state of labelling, with labelling increasing withtime [67,83,88]. A second strategy does not rely on a completelabel swap but rather aims for only a small shift in the ratiobetween two versions of amino acids, a light or unlabelled ver-sion and a heavy or labelled version that contains stable heavyisotopes of a given element. This partial-labelling approachresults, not in two separable envelopes, but rather in sub-tle, measurable modifications to a single envelope (Fig. 3B).By tracking this shift in the labelled envelope, the kinetics ofsynthesis and degradation can be calculated. The partial strat-egy has significant advantages including simplified spectra,which should make for easier quantitation. Additionally, withthe partial-labelling strategy, valuable instrument time con-ducting MS/MS experiments would not be used for peptidesof an intermediate labelling state, which are not likely to yieldpeptide identifications given the unknown and variable stateof labelling in many cases. However, a drawback with thisapproach is that multiple time points are required to ade-quately fit a curve, demanding more MS machine time. Anunknown with the partial-labelling strategy is how it will per-form in non-steady-state systems or with rapidly turning overproteins in plants. From a technical perspective, it is feasi-ble as partial labelling has been successfully applied for allof the isotopic labels discussed. An early study demonstratedthe feasibility of 13C partial labelling for proteomics in thephotosynthetic algae Synechocystis [82]. A partial labelling ap-proach with 2H2O, as demonstrated in two studies, avoidedthe negative consequences associated with a highly enricheddeuterium label while tapping into the advantages of this iso-tope [68,87], with the latter study also establishing the efficacyof H2

18O as a dilute tracer [87]. Finally, a partial-labelling with15N has also been reported as a viable tool in Arabidopsis [96]for standard proteomic analysis.


Proteomics 2014, 14, 579–592 585

4.4 Practical considerations for analysing labelled

proteins in plants

The single most important factor that needs evaluation iswhether the system in question is in a steady or a non-steady-state when evaluating protein turnover. For a thor-ough discussion of these concepts, associated models andmathematics for turnover, readers should see the followingreviews [75,94,95]. As a basic definition, a system is in steadystate when synthesis equals degradation and any changes inprotein abundance are due to growth with relative proteinabundances not changing [75]. Alternatively, in a non-steady-state system, where degradation and synthesis are not equaland protein abundance changes through time these abun-dance changes need to be accounted for when calculating Ks

and Kd. The state of the system will vary depending on theplant tissue, the developmental stage and the time over whichthe labelling is undertaken. Plants are continuously growingand developing so a simple steady state with no growth sce-nario will rarely exist in plants; however, there may be inter-vals in vegetative growth where plant tissues may be reason-ably approximated by a steady-state assumption and changesin protein abundance are simply due to cell duplication andexpansion. For plant biologists, perturbed systems are fre-quently of interest, such as biotic or abiotic stresses, whichwill require that investigators consider changes in proteinabundance when measuring and reporting Ks and Kd. Also,if an experiment occurs across more than one developmen-tal stage it is likely that relative protein abundances wouldchange, requiring that protein abundances be considered.

Besides protein degradation, an additional factor that con-tributes to a reduction in the relative amount of an unlabelledpeptide population through time is new protein synthesisfrom growth. This can be a sizable proportion of the changeseen in the spectra of peptides from slowly turning over pro-teins and requires the subtraction of a dilution factor (Kdil).Depending on the ratio of Kd to Kdil, it is conceivable therewill be very slowly degraded proteins where accurate mea-surement of Kd will be difficult over the period of a typicalturnover experiment. In one experiment Arabidopsis plantswere labelled for up to 8 weeks [83], but most studies in plantcells are 1 week or less [74,84,89,90]. Histones in plants andanimals are known to decay at very slow rates with half-liveson the order of months. In the green algae, Ostreococcus tauri,histones were the slowest turning over proteins with reportedrates of 0.017–0.024 per day [91]. Reports from other systemsare even slower with values of less than 0.01 per day observedin radiolabelling experiments [97]. It is not yet know if pro-teins such as histones and nuclear pore proteins, which havebeen reported to degrade slowly in animals have similar Kd

values in land plants [97–99]. For these slowly turning overproteins, experiments would probably need to run for severaldays and perhaps even weeks in order to make accurate mea-surements of protein dynamics. This should be feasible aslong as the plant is not transitioning between developmentalstages.

At the other end of the spectrum are the difficulties associ-ated with measuring synthesis and degradation in rapidlyturning over proteins. As mentioned, most of the stable-isotope labels that can be provided to whole plants require thatthe label be applied to the roots and transported throughoutthe plant. This can make for a lag associated with the inter-val of transport from root to tissue of interest. This lag phasewould be expected to be maximal with leaf tissue when consid-ering 2H, 18O and 15N labels given that it is the furthest tissuefrom the roots. For proteins with fast kinetics, this lag mayresult in an underestimate of Ks and Kd. This issue might befurther exacerbated under stress conditions when stomata areclosed [100,101], which reduces the transport of water and ni-trogenous salts and intake of CO2, thereby most likely extend-ing the lag of label delivery and making accurate measure-ment of Kd that much more challenging. However, assumingthat these technical hurdles can be accounted for, multipledata points within the first day will be needed in order to getaccurate measurement of synthesis and degradation rate offast turning over proteins. For very fast turnover proteins suchas D1 of PSII, where protein half-life may be as short as 30 minin the light [102], accurate measurements will be challenging.

Potentially further complicating experimental designand/or analysis is the existence of subcellular pools of aminoacids and their precursors and the degree of exchange be-tween these pools. In many plant tissues, the vacuole com-prises the majority of the cell volume [103] and is a large reser-voir for several metabolites including amino acids [104, 105].An analysis of vacuolar metabolites in barley reported thatapproximately half of the amino acids were of comparableor higher levels in the vacuole relative to the protoplast as awhole, while several organic acids were at higher levels in thevacuole [105]. Depending on the degree of exchange betweenthese large vacuolar pools and other sub-cellular pools, pro-tein turnover measurements can be influenced. Older studieshave confirmed that plants have translationally active and in-active pools of amino acids and organic acids [7–9, 106, 107].However, for the handful of turnover studies for plants wherethe ratio of label in amino acid pools and peptides have bothbeen calculated [83, 84, 88], this has not been an issue so far.Given that a small set of proteins are synthesised in mitochon-dria and chloroplasts, adequate labelling of these organellaramino acid pools is also of concern in interpreting these data.In elegant 13C labelling experiments in developing Brassicanapus embryos, investigators compared isotopic enrichmentof amino acids derived from the large subunit of RUBISCO,which is synthesised in the plastid, and the small subunit,which is synthesised in the cytosol. The study reported thatalanine, serine and glycine in plastids and the cytosol haddifferent metabolic origins and that the different amino acidpools were not in equilibrium relative to plant growth but dif-ferences were not extreme [108]. These biases would probablyalso vary depending on the label used.

An additional challenge for 15N labelling in photosynthetictissues is variation in photorespiration and the consequencesfor generation of NH3 and/or the assimilation of NO3

−. In



leaves, NH3 generated by photorespiration and reincorpo-rated back into amino acids via glutamine synthetase candramatically exceed the NH3 from new primary assimila-tion [109]. This will slow label incorporation in leaves rel-ative to other tissues such as roots, potentially leading tounderestimates of Kd and Ks for rapidly turning over pro-teins in illuminated green tissues. This issue may be furtherexacerbated under stress conditions such as salinity or coldwhen photorespiration increases as a protective mechanismfrom oxidative stress [110]. As an interesting counterpoint,under high CO2 levels both photorespiration and NO3 assim-ilation decrease, suggesting that malate synthesised duringphotorespiration, which can generate NADH when oxidised,is shuttled from plastid to the cytosol to facilitate nitrate re-duction [111,112]. These challenges should be considered byexperimentalists in interpreting labelling studies, especiallywith regards to the interplay and close relationship betweenC and N metabolism in plants.

Finally, care must also be taken in comparing turnovermeasurements for a protein between different species, tissuetypes, and even for a protein in the same tissue but of differentages. This warning may be most applicable when comparingartificial systems such as callus and cell culture to those ofintact plants. Given the paucity of proteomic turnover studiesin plants it is difficult to make definitive comparisons. How-ever, some comparisons of whole protein turnover studiesmay provide preliminary insight. Representative Kd measure-ments from intact plants include: 0.60 per day in expandingsoybean hypocotols, 0.11 per day in developing secondarywheat leaves, 0.06 per day in developed primary wheat leaves,0.04 per day in developing tobacco leaves and immeasurablein mature tobacco leaves [10]. Measurement of turnover inLemna callus was 0.09 per day while tobacco culture was fasterat 0.26 per day [10]. In a proteomic study from Arabidopsiscell culture, the median Kd value for measured proteins was0.24 per day [88]. Measurements from whole plants tended tobe slower than the undifferentiated growth systems but alsoshowed a wide range so care must be used in interpreting datafrom any turnover study but particularly as the study in ques-tion deviates from the conditions of interest, emphasising theneed for more baseline protein turnover studies.

5 Future directions

While there are most certainly challenges associated with la-belling plant tissues as discussed, there are also benefits inworking with plants. Because of the short generation timeand large collection of knock out lines available for model or-ganisms like Arabidopsis [113], plant biologists have powerfultools with which to unravel the dynamics of protein turnoverand its interplay with other biochemical processes. For exam-ple, comparison between mutant and wild-type backgroundswill allow researchers to disentangle the complex phenotypesthat appear to be the knock on consequences from loss of spe-cific proteases [59,60], or removal of single proteins in protein

complexes, metabolic and signalling pathways [114, 115]. Inplants it is also easy to apply experimental designs for la-belling that might be more challenging in animal systems.

Most of the turnover studies to date have used shotgun pro-teomics of crude lysates to determine a single Kd for a proteinin a given tissue [67, 77, 78]. Such studies are useful and pro-vide large data sets that are a good starting point for assess-ing global protein turnover, however, a Kd is contextual anddepends not only on tissue type but also on sub-cellular local-isation and even the protein complex. A single Kd value froma crude lysate will be an average of all the sub-populations fora given protein within a tissue so that a significant amount ofbiological information is lost through this averaging process.Imaging MS would be the interesting solution to this prob-lem as it would minimise sample handling and eliminate theneed for separation of tissues and sub-cellular fractions whilepotentially providing tissue level, sub-cellular level and evencomplex level information. Additionally, using imaging of se-quential tissue sections would even allow the assembly of 3Dprotein profiles. There has been noticeable progress made inthis area with MALDI MS imaging being the most frequentlyapplied technique for proteomics [116] while other new tech-niques and technologies are also being tested [117]. Prelim-inary work with imaging via MALDI MS in plants has beenpromising, however, two prominent technical hurdles were(i) difficulty collecting intense enough tandem mass spectrawith which to make confident protein identifications, and (ii)reliable sample preparation including matrix application thatwould be necessary for protein turnover studies [118].

Until imaging MS becomes a robust methodology spatialand protein interaction information can be accessed by ap-plying various fractionation techniques prior to analysis byLC-MS/MS. In one large-scale study of cell culture, inves-tigators applied sub-cellular fractionation prior to analysisand observed that a large fraction of proteins with multiplesub-cellular localisations had variable Kd values in differentcompartments [119]. In detailed analysis of CI and CV ofOXPHOS in Arabidopsis, investigators performed mitochon-drial enrichment followed by blue native PAGE (BN PAGE)separation of protein complexes, finding different Kd valuesfor some subunits of the respective sub-complexes and theholo-complexes [89, 90].

New strategies to assess protein interactions might alsoprove useful in turnover studies. In a recent tobacco cell cul-ture study, researchers combined BN PAGE and LC-MS/MSand then applied multivariate data analysis to the result-ing protein identifications from different molecular weightfractions. Using this approach, investigators identified hun-dreds of protein interactors with dozens of these known aslong-lived protein interactions but in addition the authorscharacterised more ephemeral protein interactions and sug-gested several novel protein complex combinations [120].A combination of these techniques would be expected toyield the greatest information. Plant tissues collected overa stable-isotope labelling time course could be fractionated,perhaps using differential centrifugation, and the resulting


Proteomics 2014, 14, 579–592 587

sub-cellular fractions further separated by native separa-tions to retain protein complex information [120]. Large-scaleturnover information on proteins, complexes and even sub-complexes in combination with other protein [121] and com-plex level assessments [122] could prove invaluable.

Besides complex level information, another daunting chal-lenge will be determining the role that PTMs play in proteinstability and turnover [75]. Myriad of protein modifications ex-ist that have been characterised to varying degrees in terms oftheir prevalence and physiological effects within organisms.For some of the better understood modifications, large-scaleintegrative studies are just beginning to assess this complex-ity, with one study addressing the role of phosphorylation,acetylation and ubiquitination in biological networks, empha-sising the need for integration of large data sets [123]. Whileplant studies typically lag behind those of animal systems,there has been an impressive collection and cataloguing ofphosphorylation and ubiqutination sites in different plant sys-tems as well as studies that have explored the physiologicalimplications for some of these modifications [30, 124–127].N-terminal processing of proteins is another class of mod-ifications that is fairly common that has been shown toaffect protein stability as well as localisation. Frequently,the N-terminal methionine is removed and an N-terminal�-acetylation or myristolation moiety is attached. In one sur-vey from Arabidopsis, a large fraction of proteins were mod-ified and interestingly nuclear-encoded plastid proteins wereshown to possess N-terminal acetylation following signal pep-tide removal, suggesting organelle-specific machinery whichis not present in animals [128]. There are also mitochondrial-specific peptidases in rice and Arabidopsis responsible forN-terminal modification that are believed to play a role inprotein stability [129].

Besides enzymatic modification, proteins can also becomemodified or damaged through day-to-day operations withinthe cell with many of these modifications being irreversible.However, two common modifications, cysteine and methio-nine oxidation, are reversible by the actions of glutaredox-ins and thioredoxins in the case of cysteine and methioninesulphoxide reductases in the case of methionine. These en-zymes are known to play protective roles in developmentand stress [130, 131]. Another common modification to pro-teins, resulting from daily wear and tear, is the formation ofisoaspartatyl residues from deamidation of asparagine andisomerisation of aspartic acid. A highly conserved class ofenzymes, protein isoaspartyl methyltransferases, repair thisdamage and are known to play important roles in seed viabil-ity and germination [132,133]. Future studies that investigatethe roles of these repair systems and their relation to proteinturnover in plants should provide insight into our under-standing of plant physiology and the role of repair in slowingthe need for protein degradation.

Beyond the direct impact of a PTM on the turnover of a pro-tein, modifications to the synthetic and catabolic machinerycan affect protein dynamics. There are established examplesof PTMs, particularly phosphorylation, impacting protein

synthesis indirectly by their effects on the ribosomal appara-tus [29], some of which are unique to plants [134]. Indirect ef-fects on a protein’s stability can also be accomplished by mod-ifications to the proteasomal machinery [50]. Interestingly,one proteomic report suggested that the small ubiquitin-like modifier PTM participates in a post-transcriptional re-sponse to various stresses, potentially impacting proteinsynthesis [135].

Application of turnover studies in combination with othersystems level approaches will also leverage the informationavailable from a given experiment. By integrating transcrip-tomic, proteomic and metabolomic methodologies one wouldanticipate synergistic gains in knowledge. In one impressiveexample, investigators assessed synthesis and degradation ofproteins and their associated transcripts for 5000 proteins inmouse cells, finding that protein synthesis played the largestrole in determining protein abundance under the steady-state experimental conditions employed [19]. In another re-port, protein dynamics were determined for Chlamydomonasreinhardtii and samples were also subjected to metabolomicanalysis for control and salt-stressed cells [79]. The authorsfound noticeable metabolomic changes with minimal pro-teomic perturbations and assigned metabolite shifts to theidea of metabolic flexibility within the existing proteome andPTMs, thus avoiding expensive protein synthesis when pos-sible [79]. These are just a couple of recent examples of howturnover studies are being combined with other systems biol-ogy techniques, but they set the scene for the combination ofsystems level technologies and methodologies in the future.

The authors have declared no conflict of interest.

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