Dynamics of protein synthesis and degradation through the cell cycle

ABSTRACT

Protein expression levels depend on the balance between their synthesis and degradation rates. Even quiescent (G₀) cells display a continuous turnover of proteins, despite protein levels remaining largely constant over time. In cycling cells, global protein levels need to be precisely doubled at each cell division in order to maintain cellular homeostasis, but we still lack a quantitative understanding of how this is achieved. Recent studies have shed light on cell cycle-dependent changes in protein synthesis and degradation rates. Here we discuss current population-based and single cell approaches used to assess protein synthesis and degradation, and review the insights they have provided into the dynamics of protein turnover in different cell cycle phases.

Introduction

More than 50% of cellular dry weight consists of proteins [1], which act as building blocks as well as main executors of most cellular functions. Their continuous synthesis and degradation allow to replace old proteins with new ones and to alter the composition of the proteome during cell differentiation or in response to intra- or extracellular cues. The complex interplay of protein synthesis, folding, trafficking and degradation is referred to as protein homeostasis (or proteostasis). The precise execution of each step involved in the regulation of protein levels is crucial to the function of the cell and thus to the health of the whole organism, as their mis-regulation can lead to a variety of diseases [2].

The birth of a new protein is the end-result of several consecutive steps and can be regulated at various levels. It is initiated by the onset of transcription, followed by pre-mRNA maturation, nuclear export of the mRNA, translation and protein folding. Newly synthesized proteins can also be subject to a myriad of posttranslational modifications, such as phosphorylation, glycosylation, ubiquitination and many others [3]. Importantly, while often used as a synonym for protein translation, in this review we will employ “protein synthesis” as the net result of all gene expression steps leading to the synthesis of a protein, including transcription, pre-mRNA maturation, mRNA export, and translation.

Protein degradation generally follows first order decay kinetics, with half-lives varying widely across different proteins depending on their function and the cell type/organism in which they are expressed. Regulatory proteins such as transcription factors are often short-lived, which allows for rapid adjustment of their levels in response to internal or external stimuli. In contrast, housekeeping proteins are not subject to short-term fluctuations in their levels and are thus generally long-lived [4]. In addition, protein lifetimes in fast-cycling cell types, such as prokaryotes or yeast, are usually shorter than in cell types with a longer cell cycle [5,6].

The major route for selective protein degradation is the ubiquitin-proteasome pathway [4]. To target a protein for degradation, a chain of ubiquitin molecules is added to the side chain of one of its lysine residues. Polyubiquitination is a multistep process, mediated by 3 main classes of enzymes: ubiquitin-activating enzymes (E1), ubiquitin-conjugating enzymes (E2) and ubiquitin ligases (E3). E3 enzymes mediate substrate specificity [7] and are consequently the most heterogeneous class of enzymes in the ubiquitination pathway [8]. Polyubiquitinated proteins are then recognized and degraded by the proteasome, a large multisubunit protease complex consisting of two major subcomplexes: the core particle (CP), also referred to as the 20S subunit, which executes the degradation reaction, and the regulatory particle (RP), also referred to as the 19S subunit, which mediates target recognition [9].

Each individual step involved in protein synthesis and degradation needs to be tightly regulated to control protein expression levels. How this is achieved is not fully understood. In non-dividing (G₀) cells, the levels of the vast majority of proteins are kept constant, even though proteins are still continuously turned over. In contrast, cycling cells such as embryonic stem cells [10,11], intestinal stem cells, transit amplifying cells [12] and hematopoietic stem cells [13] continuously need to increase their biomass, thus more proteins need to be synthesized than degraded. Importantly, cycling cells need to be able to precisely double their global protein levels at each cell cycle, as imbalances in protein homeostasis might broadly compromise cellular functions.

Cycling mammalian cells transit through four consecutive phases: G1, S, G2 and M, each of which consists of a specific series of events that lead to the doubling of protein content to generate two new cells [4]. The transition from one cell cycle phase to the next is regulated by complex molecular programs whose main executors are specific pairs of cyclins and cyclin-dependent kinases (CDKs) [14]. In addition, the cell cycle contains several checkpoints, which ensure that the cell is fully prepared to enter the next phase [15]. The cell cycle poses several major challenges to gene expression and thus global protein turnover, such as DNA duplication during S phase and chromosome condensation during M phase. How cells control the rates of protein synthesis and degradation throughout the cell cycle to allow protein level doubling is not understood.

In this review we start by describing the most commonly used methods to monitor protein synthesis and degradation. We distinguish between population-based bulk biochemical approaches and fluorescence-based single cell methods, which both have their specific advantages and disadvantages (Table 1). In addition, some approaches allow to study synthesis or degradation of specific proteins, whereas other methods measure synthesis and degradation rates on a global level. Furthermore, we discuss what studies based on those methods have revealed about cell cycle-dependent changes and the interplay between protein synthesis and degradation.

Table 1.

Overview of the most commonly used methods to measure protein synthesis and degradation.

Method	Measures	Specificity	Advantages	Disadvantages
Bulk biochemical methods
Metabolically labeled amino acids or nucleotides	Synthesis OR degradation	Global OR protein-specific	No genetic manipulation Genome-wide studies possible Synthesis and degradation can be studied in the same sample	Low temporal resolution No single cell resolution No in vivo studies Cell cycle studies require synchronization drugs
Incorporation of puromycin (PUNCH-P)	Synthesis	Global AND protein-specific
Ribosome profiling	Synthesis	Global AND protein-specific
Cycloheximide chase & Western Blot	Degradation	Global OR protein-specific
SILAC in combination with MS	Synthesis AND degradation	Global AND protein-specific
Fluorescence-based approaches
Incorporation of puromycin (OP-Puro)	Synthesis	Global	Single cell resolution In silico synchronization (no drugs required) Studies of short-term fluctuations and dynamic changes	Often require tedious genetic engineering Fusion with tag or fluorescent proteins might alter half-lives Risk of phototoxicity or bleaching
Controllable destabilization domain (e.g. ecDHFR)	Synthesis	Protein- or regulatory domain- specific
SunTag system	Synthesis	Protein-specific
Cycloheximide & fluorescence microscopy	Degradation	Protein-specific
Bleach chase	Degradation	Protein-specific
Fluorescently labeled protein tags (SNAP/Halo/Clip-tags)	Degradation	Protein-specific
Fluorescent protein timers	Synthesis AND degradation	Protein-specific

Methods to monitor protein turnover

Bulk biochemical approaches

Protein synthesis rates

Protein levels are often indirectly inferred from quantifications of mRNA levels, which are relatively straightforward using well-established methods such as qPCR, RNA-seq or microarrays. However, the relatively poor correlations between mRNA and protein levels suggest that mRNA quantification alone may not be a reliable proxy for protein synthesis rates [16–18]. Thus, measuring translation rates directly or in combination with transcription rates is required to obtain a more accurate assessment of protein synthesis rates. Traditionally, global protein synthesis has been measured using the incorporation of radioactively or isotopically labeled amino acids into nascent polypeptide chains. More recent studies have also used the translation inhibitor puromycin, which is incorporated into the C-terminus of a nascent polypeptide and induces premature chain termination. The use of biotin-conjugated puromycin allows the puromycylated polypeptides to be captured, purified and further analyzed by mass spectrometry [19]. Another approach to study protein synthesis rates is to directly quantify mRNAs undergoing translation by ribosome profiling, which relies on the isolation and deep sequencing of ribosome-protected mRNA fragments [20,21]. This method enables a transcriptome-wide assessment of ribosome occupancy, thus providing a snapshot of the global translational landscape at a sub-codon resolution. It has the additional advantage to allow for mapping translation start sites and thereby identifying novel open reading frames (ORFs).

Protein degradation rates

Similar to approaches allowing to monitor protein synthesis, early studies quantifying protein degradation mainly relied on metabolic labeling with radioactively or isotopically labeled elements [22]. This approach allows for determining either global or protein-specific degradation rates, the latter including a pull-down step with a specific antibody. Another widely used method to monitor protein degradation is cycloheximide chase, in which protein synthesis is blocked by cycloheximide and the decay of the remaining protein pool is monitored by western blotting at different time-points [23]. In contrast to radioactive or isotopic labeling, this method does not require immunoprecipitation of a specific protein of interest. However, cycloheximide induces considerable cellular stress and ultimately cell death, which limits its application to short-lived proteins.

Combined measurements of protein synthesis and degradation rates

Technological advances in the field of mass spectrometry have considerably increased the throughput and resolution of pulse-chase experiments and thereby provided a more quantitative and global picture of gene expression. In 2011, Schwanhäusser and colleagues published the first genome-wide study of both RNA and protein turnover [6]. They used stable isotope labeling by amino acids in cell culture (SILAC) followed by liquid chromatography and online tandem mass spectrometry (LC-MS/MS) to monitor protein synthesis and degradation, as well as pulse labeling with the nucleoside 4-thiouridine (4sU) to simultaneously measure RNA turnover. This allowed to quantitatively dissect the contribution of different steps of gene expression in determining protein levels on a genome-wide scale. They found that cellular protein abundance is mostly controlled at the level of protein translation and that functionally related proteins often show similar dynamics of RNA and protein turnover. Complementing these findings, Jovanovic et al. have studied the changes in gene expression upon stimulation of mouse immune bone marrow-derived dendritic cells (DC) with lipopolysaccharide (LPS), using a similar genome-wide approach allowing to monitor both RNA and protein turnover simultaneously [24]. They observed that the acquisition of novel cellular functions related to the stimulation response is mainly controlled at the level of transcription. In contrast, they showed that proteins performing basic cellular functions such as ribosomal proteins or components of the cytoskeleton are rather regulated by changes in translation or protein degradation.

One major drawback of bulk biochemical approaches is their low temporal resolution due to the limited number of time-points that can be acquired. Thus, short-term fluctuations in protein turnover cannot be assessed using those methods. More importantly, they only provide population averages, thus masking cell-to-cell variability in proteostasis and unsynchronized, single cell fluctuations in protein synthesis and degradation rates. Finally, they require the use of cell cycle synchronizing drugs to investigate cell cycle-related processes, which often results in incomplete synchronization and can compromise certain steps of gene expression, such as interference with translation by the microtubule-destabilizing agent nocodazole [25].

Single cell imaging approaches

Protein synthesis rates

In contrast to population-based biochemical approaches, fluorescence or luminescence microscopy-based methods provide single cell resolution. Several approaches have been established to study transcription, both in fixed cells, e.g. by single molecule fluorescence in situ hybridization (smFISH) [26], and in live cells, e.g. by using the MS2 system [27] or luminescence imaging of a destabilized luciferase reporter [28]. In contrast, measuring translation rates in single cells has been more challenging. Similarly to the PUNCH-P approach described above, puromycin can also be used as a fluorescence-based approach, since puromycilated polypeptides can be visualized and quantified by conjugating them to a fluorescent dye (OP-Puro) [29]. However, this only allows for studying global translation rates and does not provide protein specificity. In contrast, an approach based on the E. coli destabilization domain from dihydrofolate reductase (ecDHFR) that can be stabilized by its ligand trimethoprim (TMP) [30] allows to monitor translation rates of specific mRNAs in single live cells. The ecDHFR can be conjugated to a translation regulatory domain and fused to a fluorescent reporter, which leads to the immediate degradation of the fusion protein and a reduction of the reporter fluorescence to a basal level in the absence of TMP. Upon addition of TMP, the accumulation of fluorescence can be used to monitor real-time translation rates [30].

Finally, the visualization of translation in vivo with single mRNA resolution has been rendered possible by the recently developed SunTag system. Briefly, this approach is based on the fusion of a protein of interest to an array of SunTag peptides, which can be recognized specifically by a GFP-tagged single-chain intracellular antibody (scFv-GFP) [31,32]. As soon as the newly translated SunTag peptides exit the ribosome, they are labeled by the scFv-GFP, which allows for in vivo monitoring of translation. This approach has been applied to study various aspects of translation, such as measurements of ribosome numbers, initiation and elongation rates, and changes in translation rates in response to various stimuli [31,32].

Protein degradation rates

Inhibition of protein synthesis with cycloheximide can also be used in combination with time-lapse fluorescence microscopy to monitor the decay of fluorescently labeled proteins, but this is only suitable to study short-lived proteins because of cytotoxicity. In contrast, the “bleach-chase” approach does not require translational inhibitors [33]. This method requires cells expressing an endogenous protein fused to a fluorescent reporter. These are then partially bleached by a short pulse of light, which irreversibly renders a fraction of the fluorophores non-fluorescent. After the pulse, the bleached protein population decays and is replaced by newly synthesized fluorescent proteins, which can be observed by time-lapse microscopy. At the same time, a population of unbleached cells is imaged. The protein half-life can then be inferred by subtracting the fluorescence intensities of bleached and unbleached cells. However, since this approach relies on the comparison between the averages of two different populations of cells (bleached vs. unbleached) it does not allow to determine protein degradation rates of individual cells.

Time-lapse imaging of pulse-labeled fluorescent tags, such as SNAP-tags [34], Halo-tags [35] or Clip-tags [36] allow measuring protein degradation rates in single cells. The tags are fused to a protein of interest and can be covalently bound by a fluorescent ligand (Figure 1(a)). Upon washing of the residual ligand or blocking with a non-fluorescent version of the ligand, the decay of the labeled protein population can be monitored, which allows for assessing protein half-lives (Figure 1(a,b)). This approach has been successfully established to measure protein decay of cell populations in vivo [37] and single cell protein half-lives in in vitro cultured adherent cells [38–40]. Importantly, fluorescently labeled protein tags can not only be used to assess cell-to-cell variability in protein half-lives, but also to monitor transient changes in protein degradation [39] (Figure 1(c)), as they can provide a higher temporal resolution than bulk biochemical approaches.

Figure 1.

Fluorescently labeled protein tags to monitor protein degradation. (a): A protein of interest is fused to a SNAP/Halo/Clip-tag, which can be pulse-labeled with a fluorescent ligand. Upon washing of the residual dye, the decay of the labeled protein population can be observed. (b): Protein degradation follows an exponential decay. A₀: Initial fluorescence intensity, b: decay rate, T_1/2: half-life. (c): Schematic representation of how SNAP/Halo/Clip-tags can be used to identify changes in protein degradation.

Tandem fluorescent timers: simultaneous measurements of protein synthesis and degradation rates in single live cells

The fluorescence-based approaches described above do not allow for simultaneous measurements of protein synthesis and degradation rates. Thus, until very recently it has not been possible to determine how synthesis and degradation interplay to control protein expression levels in individual cells. However, a new study reported the development of mammalian cell-optimized fluorescent timers, allowing to disentangle changes in protein synthesis and degradation in single live cells [39]. Fluorescent timers are fluorescent proteins that change their emission spectrum over time, which provides a measure of the age of a protein pool. Traditional timers consist of only one fluorophore that switches its emission spectrum due to a series of chemical reactions, but their application has been impeded by their low brightness and tendency to form oligomers [41,42]. These limitations were overcome by the engineering of tandem fluorescent protein timers (tFTs), which consist of a fusion of the fast maturing green fluorescent protein superfolder GFP (sfGFP) and a slower maturing red fluorescent protein such as mCherry, TagRFP, Td-Tomato, mKate2 or mOrange2 [39,43,44] (Figure 2(a)). Tandem fluorescent timers have mainly been used to assess changes in protein stability or age-dependent protein localization [43–45], as the lag in maturation time between the two fluorophores allows for distinguishing old from newly made proteins and thereby provides an estimate for the average age of a protein pool. However, it is in fact possible to use fluorescent timers to gain quantitative insights into protein synthesis and degradation rates. Alber et al. recently leveraged on this idea by combining single cell time-lapse imaging of timer-tagged proteins with computational modeling, using a set of differential equations to infer both protein synthesis and degradation in single cells [39]. Figure 2(b) illustrates how tandem fluorescent timers can be used to distinguish changes in protein synthesis from changes in protein degradation. An increase in protein synthesis leads to a fast increase in green fluorescence, followed by a slower increase in red fluorescence due to the slower maturation of the red fluorescent protein. In contrast, a decrease in protein degradation leads to an increase in red fluorescence that is more pronounced than the increase in green fluorescence, and thus the green/red ratio decreases. In combination with computational modeling, this allows to extract temporal changes in protein synthesis and degradation rates in single live cells.

Figure 2.

Tandem fluorescent timers allow to simultaneously monitor protein synthesis and degradation. (a): Upon production of the fusion protein, superfolder GFP (sfGFP), matures fast, whereas the red fluorescent protein (FP) matures more slowly. (b): Simulations showing how tandem fluorescent timers can be used to distinguish changes in protein synthesis from changes in protein degradation. Figure reused from [38], Figure 1(d), with permission from Molecular Cell (License number: 4472970734996).

In summary, while population-based bulk biochemical approaches enable proteome-wide studies, mostly without genetic manipulation, fluorescence-based approaches allow for single cell studies and provide a high temporal resolution. However, fluorescence-based approaches can be affected by phototoxicity or bleaching. Fluorescent tags and fusion proteins might also alter endogenous protein turnover, especially when fused to the N-terminus of the protein of interest (N-end rule) [46]. In addition, expression of fluorescent fusion proteins from strong exogenous promoters may lead to changes in turnover, for example due to altered stoichiometry relative to interaction partners [47]. Thus, it is advisable to use endogenous, C-terminal fusions and to carefully test for potential alterations in protein degradation, e.g. by using bulk biochemical methods to directly compare the half-life of a fluorescent fusion protein and its endogenous version. Despite these potential limitations, single cell fluorescence-based approaches are of particular interest for studying cell cycle related processes, as cells can be in silico synchronized and cell cycle-associated processes can therefore be studied without the use of cell cycle-synchronizing drugs.

Dynamics of protein turnover through the cell cycle

Cell cycle-dependent changes in protein synthesis rates

While cell cycle-related changes in the expression of key regulators of cell cycle progression have been known for a long time [19,48–50], how global transcriptional and translational rates are altered in different cell cycle phases has only started to be addressed recently. The most pronounced changes in global transcriptional activity take place during mitosis. Due to the condensation of chromosomes, the RNA polymerase II machinery was suggested to be physically excluded from chromatin, causing a complete and global shutdown of transcriptional activity [51,52]. However, a more recent study showed that transcription is retained at low levels during mitosis [53]. During early G1 phase, 50% of transcribed genes display a hyperactive transcriptional state that leads to a spike in transcription [54]. A further perturbation to transcription occurs when each gene copy is duplicated during S phase. While this should in principle lead to an increase in mRNA synthesis, there is evidence for buffering mechanisms involving H3K56 methylation on newly deposited histones [55].

Translational activity has also been shown to globally decrease during mitosis, which was suggested to facilitate the selective translation of certain specific mRNAs by internal ribosomal entry sites [56–60]. However, these studies mostly relied on mitotic synchronization of cells by microtubule-destabilizing drugs such as nocodazole, which can interfere with translation [25] and thus complicate the interpretation of experimental results. More recently, a study re-addressed global regulation of translation during mitosis using the potentially less perturbing CDK1 inhibitor RO-3306 for synchronization in G2 phase [61]. Metabolic labeling revealed a global decrease in translation by 35%, in line with previous findings. Furthermore, mitosis-specific ribosome profiling revealed a subset of 200 mRNAs that are strongly up- or down- regulated at the translational level during M-phase [61]. In contrast to mitosis, population-based studies revealed no major changes in global protein synthesis rates during G1, S and G2 phase.

Alber et al. have recently monitored changes in protein synthesis of several proteins in single cells throughout the cell cycle. While synthesis rates increased over the cell cycle for several proteins they studied, degradation rates essentially remained constant [39]. This indicates that between the beginning of G1 phase and the end of G2 phase, synthesis rates can change in a protein-specific manner and result in different protein accumulation profiles. They then further focused on the time interval around cell division, and in agreement with previous studies found a decrease in protein synthesis for a large number of proteins [39].

In summary, both population-based and single cell studies suggest that the most pronounced global changes in protein synthesis occur during cell division, which can be attributed to large changes in both transcription and translation rates.

Cell cycle dependent changes in protein degradation rates

While the regulation of transcriptional and translational efficiency throughout the cell cycle is relatively well studied on a global and transcript-specific level, our knowledge about cell cycle dependent changes in protein degradation has long been limited to very specific cases such as cyclins and other key cell cycle regulatory proteins [62] .

More recently, two studies have provided evidence for more global cell cycle-dependent changes in protein degradation rates. In several cultured cell lines, the periodic activation of the Wnt-signaling cascade leads to inhibition of glycogen synthase kinase 3 (GSK3) during mitosis. Since GSK3 kinase-mediated phosphorylation is often leading to target protein degradation, this causes a stabilization of GSK3 target proteins specifically during M phase, which is referred to as Wnt-dependent stabilization of proteins (Wnt/STOP) [63]. Wnt/STOP might promote cell growth, as its inhibition was shown to decrease in the size of G1 cells [63]. Since 20% of all cellular proteins are predicted GSK3 target proteins, Wnt/STOP might affect a substantial fraction of the proteome [64]. Using their fluorescent timer approach, Alber and colleagues recently made the surprising finding that about 40% of proteins are stabilized during cell division [39]. These results were further confirmed by fluorescent pulse-labeling of SNAP-tagged proteins. Importantly, the culture conditions that were used maintain the Wnt signaling pathway constantly on, bypassing a potential Wnt/STOP mechanism and thus suggesting that the observed stabilization of proteins is caused by a different mechanism.

Protein homeostasis at the single cell level

The combination of tandem fluorescent timers with computational modeling and the use of fluorescently labeled protein tags has also provided the unique opportunity to simultaneously quantify protein synthesis and degradation in single live cells [39]. One outstanding question in the field was to what extent protein synthesis and degradation vary between cells, and whether their rates correlate in single cells. Alber et al. discovered that protein degradation rates vary substantially between individual cells, and that these differences are accompanied by corresponding changes in protein synthesis rates [39]. Simulations suggested that cell-to-cell variability in protein expression levels is higher in cells where protein synthesis and degradation are uncorrelated, indicating that a positive correlation between protein synthesis and degradation allows to buffer protein expression variability. Furthermore, degradation rates in single cells were correlated for different proteins, suggesting that the observed cell-to-cell variability is global rather than protein-specific. Taken together, these findings indicate that cells with similar protein levels might actually have quite different underlying global turnover rates, which may affect a variety of cellular processes, such as stress response or differentiation.

Perspectives

Elucidating how protein synthesis and degradation are regulated and how they interplay is of fundamental interest to understand how cells control protein levels. Recent technological advances have allowed for studying protein synthesis and degradation simultaneously, both at the population and single cell level [6,24,39]. The development of live cell approaches allowing time-resolved measurements of several steps of gene expression will be key to gain a quantitative understanding of the information flow from DNA to protein. The use of fluorescent timers to disentangle changes in protein synthesis and degradation represents a first step towards this goal. While CD-tagging allows for rapid generation of cell libraries by non-targeted protein fusion with a fluorescent timer or other tags [65–67], the rapid development of higher throughput CRISPR-Cas9 knock-in approaches will allow for more precise targeting of a large numbers of proteins. The development of brighter fluorescent timers will also hopefully enable studying homeostasis of lowly expressed proteins.

Recent advances have greatly increased our understanding of global cell cycle-dependent changes in protein synthesis and degradation rates (Figure 3). While a reduction in transcriptional activity can be explained by the condensation of chromatin during mitosis, the global decreases in translation and protein degradation rates are more intriguing. In particular, the mechanisms causing a decrease in protein degradation rates independently from Wnt/STOP are not known. Finally, what controls and coordinates protein synthesis and degradation to allow doubling of protein levels at each cell cycle also remains to be investigated. Fortunately, these fascinating new questions should now be within experimental reach thanks to methodological advances that raised them in the first place, and will hopefully be addressed in the near future.

Figure 3.

Summary of currently known global changes in protein synthesis (blue) and degradation (red) over the cell cycle.

Funding Statement

This work was supported by the Teofilo Rossi di Montelera e di Premuda Foundation and an anonymous donor, both advised by CARIGEST SA [NA];Fondation Pierre Mercier [NA].

Acknowledgments

This work was supported by the Pierre Mercier Foundation (to D.M.S.), and the Teofilo Rossi di Montelera e di Premuda Foundation and an anonymous donor, both advised by CARIGEST SA (to D.M.S).

Disclosure statement

No potential conflict of interest was reported by the authors.