Abstract
Tandem fluorescent protein timers (tFTs) report on protein age through time-dependent change in color, which can be exploited to study protein turnover and trafficking. Each tFT, composed of two fluorescent proteins (FPs) that differ in maturation kinetics, is suited to follow protein dynamics within a specific time range determined by the maturation rates of both FPs. So far tFTs were constructed by combining different slower-maturing red fluorescent proteins (redFPs) with the same faster-maturing superfolder green fluorescent protein (sfGFP). Towards a comprehensive characterization of tFTs, we compare here tFTs composed of different faster-maturing greenFPs, while keeping the slower-maturing redFP constant (mCherry). Our results indicate that the greenFP maturation kinetics influences the time range of a tFT. Moreover, we observe that commonly used greenFPs can partially withstand proteasomal degradation due to the stability of the FP fold, which results in accumulation of tFT fragments in the cell. Depending on the order of FPs in the timer, incomplete proteasomal degradation either shifts the time range of the tFT towards slower time scales or precludes its use for measurements of protein turnover. We identify greenFPs that are efficiently degraded by the proteasome and provide simple guidelines for design of new tFTs.
Abbreviations tFT – tandem fluorescent protein timer
FP – fluorescent protein
greenFP – green fluorescent protein
redFP – red fluorescent protein
Introduction
Fluorescent proteins (FPs) become fluorescent only upon correct folding of the polypeptide and formation of the fluorophore through a series of chemical reactions within the β-barrel fold. This maturation process takes place on time scales from minutes to hours, depending on the FP and the environment (Tsien, 1998; Shaner et al., 2005). The time delay between protein synthesis and completion of the maturation process is exploited in tandem fluorescent protein timers (tFTs) to measure the dynamics of various cellular processes (Khmelinskii et al., 2012). A tFT is a fusion of two FPs that differ in maturation kinetics and spectral properties (i.e. color), e.g. a slower-maturing red fluorescent protein (redFP) and a faster-maturing green fluorescent protein (greenFP) (Figure 1A). The greenFP is typically selected to be as fast-maturing as possible and thus reports on protein localization and abundance. Because of the difference in maturation kinetics between the greenFP and redFP moieties, the color of a tFT (defined as the redFP/greenFP ratio of fluorescence intensities) changes over time (Figure 1A). When a tFT is fused to a protein of interest, its color provides a measure of protein age that can be used to follow protein degradation, trafficking and segregation during cell division (Khmelinskii et al., 2012).
The range of protein ages that can be analyzed with a tFT, i.e. its time range, is strongly influenced by the maturation kinetics of the slower-maturing FP in the pair (Khmelinskii et al., 2012). Therefore, in the tFTs described thus far (Khmelinskii et al., 2012; Dona et al., 2013; Khmelinskii and Knop, 2014), different slower-maturing redFPs (mCherry, TagRFP, DsRed1) were combined with the same faster-maturing superfolder green fluorescent protein (sfGFP) (Pédelacq et al., 2006). Several properties make sfGFP a greenFP of choice for construction of tFTs: it is bright, it folds independently of the protein it is fused to, it is one of the faster-maturing greenFPs, with a maturation half-time of ~6 min in yeast (Pédelacq et al., 2006; Khmelinskii et al., 2012), and an additional V206R mutation ensures it is monomeric (Zacharias et al., 2002). Nevertheless, there is a wide range of greenFPs with other desirable properties such as high molecular brightness or high photostability (Dean and Palmer, 2014). However, their in vivo maturation rates, which determine their suitability for construction of new tFTs (Figure 1B), are not well characterized.
Here we sought to compare the performance of different greenFPs within tFTs. We constructed a series of mCherry-greenFP timers and examined their suitability for comparative measurements of protein age and protein degradation kinetics in yeast. We identified the stability of the greenFP fold as a new parameter that affects tFT behavior and directs the design and use of tFT as reporters of protein degradation.
Results
We constructed new putative tFTs by fusing the slower-maturing red fluorescent protein mCherry (Shaner et al., 2004) to different greenFPs: the commonly used monomeric yeast codon-optimized enhanced GFP (myeGFP) (GFPmut 3a from (Cormack et al., 1996) with the V206R dimerization-preventing mutation (Zacharias et al., 2002)), the fast-maturing GFPm (Yoo et al., 2007; Iizuka et al., 2011), and two greenFPs with superior brightness – Clover (Lam et al., 2012) and mNeonGreen (Shaner et al., 2013) (Figure S1, A and B). myeGFP, GFPm, Clover and mNeonGreen have red-shifted excitation peaks compared to sfGFP. Using such greenFPs is likely to improve signal-to-noise ratio in fluorescence imaging as autofluorescence of yeast cells and growth media is substantially reduced with excitation wavelengths longer than 488 nm (Figure S1C).
To test whether these mCherry-greenFP fusions function as timers and to compare their ability to report on protein age, we analyzed cells expressing the proton pump Pma1 tagged with each putative tFT. Pma1 is an exceptionally stable protein of the plasma membrane (Thayer et al., 2014). During yeast bud formation, preexisting Pma1 molecules at the plasma membrane are retained in the mother cell, while the bud (future daughter cell) receives newly produced Pma1 (Takizawa et al., 2000; Malínská et al., 2003). Consequently, cells expressing Pma1 tagged with the mCherry-sfGFP timer exhibit lower mCherry/greenFP intensity ratios (R) at the plasma membrane in the bud (Rb) than in the mother cell (Rm) (Figure 1C) (Khmelinskii et al., 2012). We observed the same trend for all Pma1 fusions (Figure 1D), indicating that all tested mCherry-greenFP variants function as timers. The Rb/Rm ratio can be used for a qualitative comparison of greenFP maturation rates with an approximate model of tFT maturation (Figure 1B) (Khmelinskii et al., 2012). The Rb/Rm ratio was lowest for mCherry-sfGFP and mCherry-mNeonGreen timers (Figure 1D), indicating that sfGFP and mNeonGreen have the fastest maturation among all tested greenFPs. Consistent with slower maturation of Clover (Figure S1A), the Rb/Rm ratio was highest for the strain expressing Pma1-mCherry-Clover (Figure 1D). De novo fluorophore maturation of anaerobically synthesized GFPm is faster than maturation of sfGFP under the same conditions (Iizuka et al., 2011) (Figure S1A). However, in vivo maturation (which also includes protein folding) of GFPm appears to be slower than that of sfGFP, as the Rb/Rm ratio was higher for Pma1-mCherry-GFPm than for Pma1-mCherry-sfGFP (Figure 1D). The maturation kinetics of myeGFP is not known. The Rb/Rm ratio of Pma1-mCherry-myeGFP was lower than expected for a greenFP with a maturation half-time of 25 min (i.e. the maturation half-time of EGFP (Figure S1A) (Shaner et al., 2007)), indicating that myeGFP matures faster than EGFP but slower than sfGFP in yeast (Figure 1D).
We proceeded to examine the performance of all tFTs in comparative analysis of protein degradation kinetics. ln steady state, the mCherry/greenFP intensity ratio is determined by the degradation kinetics of the tFT fusion, such that strains expressing rapidly degraded fusions are expected to exhibit lower mCherry/greenFP intensity ratios (Figure 2A) (Khmelinskii et al., 2012). We generated two constructs for expression of N-terminal ubiquitin (Ubi) fusions with each tFT (Ubi-X-mCherry-greenFP): one with a methionine residue (M) and another with an arginine residue (R) at position X (Figure 2B). Co-translational cleavage of the ubiquitin moiety exposes a new N-terminus starting with residue X, which determines the stability of the fusion according to the N-end rule. Fusions with N-terminal arginine, but not with N-terminal methionine, are recognized by the E3 ubiquitin ligase Ubr1 and targeted for proteasomal degradation (Bachmair et al., 1986; Varshavsky, 2011) (Figure S2, A and B). Accordingly, for each tFT, the mCherry/greenFP intensity ratio was lower for the strain expressing Ubi-R-mCherry-greenFP (Figure 2C), indicating that all tested tFTs report on protein degradation kinetics.
The difference between the median mCherry/greenFP intensity ratios of strains expressing Ubi-M-mCherry-greenFP and Ubi-R-mCherry-greenFP fusions varied from ~ 5 fold for the mCherry-sfGFP timer to 1.4 – 2.1 fold for the other tFTs. Comparison of these differences to an approximate model of tFT turnover (Figure 2A, Supplemental Theory) revealed several inconsistencies between theory and experiment. For instance, the relative mCherry/greenFP intensity ratio of Ubi-R-mCherry-sfGFP was considerably lower than expected (Figure 2C). Moreover, tFTs with Clover and mNeonGreen exhibited similar relative mCherry/greenFP intensity ratios for Ubi-R-mCherry-greenFP (Figure 2C), despite the difference in maturation kinetics (Figure S1A) and distinct performance as reporters of protein age (Figure 1D). These observations suggest that a parameter other than greenFP maturation kinetics influences tFT behavior in the protein degradation assay. Immunoblotting of whole cell extracts with antibodies against GFP (or against an hemagglutinin (HA) epitope fused to the C-terminus of mNeonGreen) revealed several bands in addition to full length fusions. First, a band at ~45 kDa (band (h) in Figure 2D) that is explained by hydrolysis of the mature mCherry fluorophore during cell extract preparation, which leads to a break in the mCherry polypeptide (Gross et al., 2000; Shemiakina et al., 2012). Additionally, unexpected tFT fragments were detected around 33 kDa (~6 kDa larger than free greenFP) in all strains expressing Ubi-X-mCherry-greenFP fusions (band (c) in Figure 2D). These fragments were most prominently seen for the tFT with sfGFP, especially for the Ubi-R-mCherry-sfGFP fusion. We sought to determine the origin of these protein fragments (hereafter referred to as processed tFT fragments) and to examine whether their presence can account for the unexpected behavior of different tFTs described above.
Two mechanisms could generate processed tFT fragments: proteolytic cleavage of the tFT by an endopeptidase (model 1) and incomplete degradation of the tFT by the proteasome (model 2) (Figure 3A). In model 1, the fraction of processed tFT in steady state is expected to be independent of the degradation kinetics of the tFT fusion. In contrast, is expected to be proportional to the degradation kinetics of the tFT fusion in model 2 (further details in Supplemental Theory). In both models processed tFT fragments accumulate only if their degradation rate constant (k2 in Figure 3A) is small. Immunoblotting analysis of strains expressing a series of Ubi-X-mCherry-sfGFP fusions with different stabilities showed that the fraction of processed tFT increases as a function of protein degradation kinetics, in agreement with model 2 (Figure 3B). Moreover, accumulation of processed tFT fragments depended on proteasomal activity (Figure S2C). These observations were not specific to the Ubi-X-mCherry-greenFP fusions. Processed tFT fragments could also be detected in strains expressing misfolded cytoplasmic proteins tagged with mCherry-sfGFP and the fraction of processed tFT decreased upon deletion of E3 ubiquitin ligases involved in degradation of these misfolded proteins (Figure S2, D and E). Finally, similar ~33 kDa fragments have been observed when GFP fusions are degraded by proteasomes with impaired processivity, e.g. in mutants of the proteasome-associated E3 ubiquitin ligase Hul5 (or its mammalian ortholog UBE3C) (Zhang and Coffino, 2004; Aviram and Kornitzer, 2010; Martínez-Noël et al., 2012; Chu et al., 2013). We conclude that accumulation of processed tFT fragments is caused by incomplete proteasomal degradation of tFT fusions and that these fragments appear to be relatively stable in the cell.
What features of the mCherry-sfGFP timer are responsible for its incomplete degradation? We considered two possibilities: the linker connecting mCherry to sfGFP and the robust fold of sfGFP. The linker between mCherry and greenFP in the tFTs is composed of a leucine (L) and an aspartate (D) residues followed by five glycine-alanine (GA) repeats (LD(GA)5). GA rich sequences can prevent complete protein degradation by impairing the ability of the proteasome to unfold its substrate (Hoyt et al., 2006; Daskalogianni et al., 2008). To determine the role of the linker in the accumulation of processed tFT fragments, we tested shorter LDGAG or LDGS linker sequences between mCherry and sfGFP. Although shorter linkers changed the relative amounts of different processed tFT fragments, they did not reduce the total amount of processed fragments (Figure 4A) nor did they alter the ability of the mCherry-sfGFP timer to report on protein degradation kinetics (Figure 4B). In contrast, exchanging sfGFP for other greenFPs, while keeping the LD(GA)5 linker between mCherry and greenFP constant, reduced the amount of processed tFT fragments (Figure 2D). Moreover, fragments resulting from incomplete proteasomal degradation could be detected when using sfGFP but not mCherry alone as a tag (Figure S2F). This suggests that the stability of the greenFP fold, not the linker, is responsible for incomplete degradation of tFT fusions, although we cannot exclude the possibility that processed tFT fragments from timers other than mCherry-sfGFP are rapidly degraded in the cell.
To test the possibility that incomplete degradation is caused by the robust fold of sfGFP, we attempted to destabilize the sfGFP fold by reverting five mutations (S30R, Y39N, N105T, Y145F and I171V) responsible for the superfolder nature of sfGFP (Pédelacq et al., 2006) (sfGFP5M) or by reverting the F64L mutation known to improve GFP folding at 37°C (Cormack et al., 1996; Tsien, 1998) (sfGFPL64F). However, tFTs with these sfGFP mutants behaved similarly to the mCherry-sfGFP timer (Figure 4, C and D). Nevertheless, in a reverse experiment, introducing the F64L mutation into Clover increased the accumulation of processed tFT fragments in the strain expressing Ubi-R-mCherry-CloverF64L (Figure 4C). This accumulation correlated with a reduced mCherry/greenFP intensity ratio of Ubi-R-mCherry-CloverF64L relative to the Ubi-M-mCherry-CloverF64L fusion, bringing it close to Ubi-R-mCherry-sfGFP (Figure 4D). The F64L mutation did not affect the accumulation of processed tFT fragments for the Ubi-M-mCherry-CloverF64L fusion, possibly because the longer half-life of Ubi-M-tFT fusions gives enough time for complete folding and maturation of wild type Clover. These observations support the idea that the stability of the greenFP fold is responsible incomplete degradation of tFT fusions.
Next we sought to destabilize the sfGFP fold through circular permutations. Circular permutation can drastically impair folding of classical GFP (Baird et al., 1999; Topell et al., 1999). Although sfGFP is largely tolerant of circular permutations (Pédelacq et al., 2006), some circularly permutated sfGFP variants can be efficiently unfolded and more rapidly degraded by the AAA+ ClpXP protease in vitro (Nager et al., 2011). We constructed a series of tFTs with different circular permutations (cp) of sfGFP (Figure 5A) and examined their behavior in the degradation assay with N-terminal ubiquitin fusions. Accumulation of processed tFT fragments was reduced to below the detection limit with all circular permutations except cp3 (Figure 5B). This is consistent with higher mobility of the β7-β11 sheets in the β-barrel fold and the tendency of GFP to start unfolding from β7-β11 (Huang et al., 2007; Zimmer et al., 2014). As seen in the case of Clover (Figure 4, C and D), reduced accumulation of processed tFT fragments correlated with a reduced relative difference in mCherry/sfGFP ratios between the stable and unstable Ubi-X-mCherry-greenFP fusions (Figure 5C). Taken together, these experiments indicate that the stability of the greenFP fold is responsible for proteasome-dependent processing of tFTs and concomitant accumulation of tFT fragments.
Two permutations, cp7 and cp9, reduced the molecular brightness of sfGFP (Figure 5D). Another two, cp12 and cp13, appeared to affect protein folding and expression levels (Figure 5D), and the maturation kinetics of sfGFP, as suggested by the lack of difference in mCherry/sfGFP ratios between the two Ubi-X-mCherry-greenFP fusions (Figure 5C). However, the cp8 permutation did not impair protein expression and had only a minor effect on the molecular brightness of sfGFP (Figure 5D). sfGFP(cp8) exhibited the fastest maturation kinetics among all tested circular permutation, as evidenced by the performance of the mCherry-sfGFP(cp8) timer in the analysis of Pma1 protein age (Figure 1D). Moreover, accumulation of processed tFT fragments was strongly reduced in human embryonic kidney HEK293T cells when using the mCherry-sfGFP(cp8) timer instead of mCherry-sfGFP (Figure S3). Thus, sfGFP(cp8) represents an alternative to sfGFP without potential artifacts caused by incomplete proteasomal degradation.
We sought to determine how incomplete proteasomal degradation affects the properties of tFTs and their use in studies of protein dynamics. We incorporated proteasome-dependent processing into a model of tFT maturation and turnover for two tFTs, mCherry-sfGFP and sfGFP-mCherry. Proteasomal degradation typically requires an unstructured region in the substrate to initiate degradation (Prakash et al., 2004). We assumed that no such initiation region is present in the mCherry-sfGFP and sfGFP-mCherry timers. It is implicit in the model that degradation is initiated within the tagged protein of interest and proceeds in a processive manner until the sfGFP fold is reached, at which point the remaining polypeptide can be either released from the proteasome with a defined probability or completely degraded. Incomplete degradation of proteins tagged at the C-terminus with mCherry-sfGFP or sfGFP-mCherry produces respectively free sfGFP or free sfGFP-mCherry in the model (further details in Supplemental Theory). Our experiments indicate that the mCherry-sfGFP timer functions as a degradation reporter with C-terminally tagged proteins (Figure 2) (Khmelinskii et al., 2012). However, the time range of this timer shifts towards more stable proteins when the probability of incomplete tFT degradation is increased in the model (Figures 6A). This shift does not depend on whether Förster resonance energy transfer (FRET) from sfGFP to mCherry is considered in the model (Supplemental Theory and Figure S4). The difference in mCherry/sfGFP intensity ratios between Ubi-M-mCherry-sfGFP and Ubi-R-mCherry-sfGFP fusions is expected to increase with increasing probability of incomplete tFT degradation (Figure 6A). The experimentally observed behavior of Clover and sfGFP-based tFTs with different levels of incomplete tFT degradation is consistent with this prediction (Figures 4D and 5C).
In contrast to the mCherry-sfGFP timer, incomplete degradation of the sfGFP-mCherry timer as a C-terminal tag is predicted to abolish the monotonic relationship between mCherry/sfGFP intensity ratio and protein half-life (Figure 6B). We tested this prediction in the degradation assay with N-terminal ubiquitin fusions. To ensure initiation of degradation N-terminally to sfGFP, we separated the N-terminal degradation signal (N-degron) from the sfGFP-mCherry timer with a spacer protein Don1 and an HA tag (Ubi-X-spacer-sfGFP-mCherry). Immunoblotting of whole cell extracts confirmed the accumulation of processed tFT fragments with the expected size, ~6 kDa larger than free sfGFP-mCherry, in these strains (Figure S5A). In contrast to strains expressing Ubi-X-mCherry-sfGFP fusions, the mCherry/sfGFP intensity ratio was largely independent of the degradation rate for Ubi-X-spacer-sfGFP-mCherry fusions (Figures 6C and S5B). This result is consistent with the model predictions and suggests that the probability of incomplete sfGFP degradation is substantial (~0.5 or higher) (Figure 6B). Moreover, it shows that the sfGFP-mCherry timer should not be used as a degradation reporter with C-terminally tagged proteins. However, it is important to stress that the sfGFP-mCherry fusion is a timer, i.e. it changes color with time, and thus should report on the age of C-terminally tagged proteins in the absence of protein degradation. Indeed cells expressing Pma1-sfGFP-mCherry exhibited lower mCherry/sfGFP intensity ratios at the plasma membrane in the bud than in the mother cell, similarly to Pma1-mCherry-sfGFP (Figure 1D). Moreover, the sfGFP-mCherry timer, but not the mCherry-sfGFP timer, faithfully reported on the degradation kinetics of N-terminally tagged proteins (Figures 6D and S5C). Together, these results indicate that different tFT variants should be used for N- and C-terminal protein tagging, such that the sfGFP moiety is placed distally to the region in the protein of interest where proteasomal degradation is initiated. Importantly, this theoretical analysis was performed under the assumption that processed tFT fragments are stable in the cell (Figure 6, A and B), and thus establishes an upper limit for the influence of proteasome-dependent processing of tFTs on their ability to report on protein degradation kinetics. Increasing the degradation rate of processed tFT fragments reduces their impact and, at the limit of infinitely fast degradation, mCherry-sfGFP and sfGFP-mCherry timers are both reporters of protein degradation kinetics when used for C-terminal tagging, with behavior indistinguishable from a hypothetical mCherry-sfGFP timer that is completely degraded by the proteasome (Supplemental Theory and Figure S6).
Discussion
We report here on the use of different greenFPs for analysis of protein dynamics with tFTs. All tested greenFPs can be combined with mCherry to obtain tFTs that report on protein age and degradation kinetics, indicating that greenFPs mature faster than mCherry. Our analysis shows that the greenFP maturation kinetics influences the time range of mCherry-greenFP timers (Figure 1). In addition, we observe that greenFPs (especially sfGFP), but not mCherry, can withstand proteasomal degradation in yeast (Figures 2 and S2F), consistent with previous reports of incomplete proteasomal degradation of GFP (Zhang and Coffino, 2004; Aviram and Kornitzer, 2010; Martínez-Noël et al., 2012; Chu et al., 2013). The efficiency of FP degradation by the proteasome could differ in other organisms and cell types, as proteasome processivity varies between species (Kraut et al., 2012).
Two features are necessary for a protein to become a proteasome substrate: a degradation signal (degron) and a degradation initiation site (Schrader et al., 2009). These features are provided by the model substrates used to investigate tFT behavior in the degradation assay (Figure 2). Therefore, proteasomal degradation of a tFT starts from the site where it is linked to the model substrate and proceeds by unfolding and channeling of the polypeptide into the catalytic core of the 19S particle of the proteasome. Protein sequences that cannot be efficiently channeled into the catalytic core can be release from the proteasome. Several factors can contribute to this release: the protein sequence adjacent to the released domain, the stability of the fold of the released domain, and an intrinsic probability of the proteasome to release substrates that is independent of the power stroke underlying substrate unfolding (Kraut et al., 2012; Fishbain et al., 2015). Our analysis indicates that incomplete degradation of sfGFP fusions depends on the stability of the sfGFP fold (Figures 4 and 5), although the molecular details, i.e. the determinants in sfGFP, are unclear.
The outcome of proteasomal processing of tFT fusions depends on the order of FPs in the timer relative to the region in the protein from which proteasomal degradation begins. If mCherry is placed proximal to the degradation initiation site (e.g. in mCherry-sfGFP timer as a C-terminal tag), proteasomal processing produces tFT fragments of ~33 kDa (Figure 2). These contain the greenFP moiety and a short ~6 kDa tail that presumably corresponds to the length of the unfolded polypeptide channeled into the proteasome catalytic core before release of the fragments. In this situation, the timer functions as a reporter of protein age and degradation kinetics but its time range is shifted towards more stable proteins compared to a hypothetical timer that is completely degraded by the proteasome (Figure 6A). This shift should increase the difference in mCherry/greenFP intensity ratios between the stable Ubi-M-tFT and unstable Ubi-R-tFT fusions (inset in Figure 6A), as experimentally observed for the mCherry-sfGFP timer compared to mCherry-sfGFP(cp8) or mCherry-mNeonGreen (Figures 2C and 5C). Moreover, the increased cytoplasmic greenFP fluorescence resulting from incomplete tFT degradation should be taken into account when measuring protein age at subcellular level. We note that the ~33 kDa tFT fragments are distinct from the ~26 kDa fragments observed upon incomplete lysosomal/vacuolar degradation of GFP fusions, which have been exploited to follow the cytoplasm to vacuole targeting (Cvt) pathway and autophagy (Shintani and Klionsky, 2004; Klionsky et al., 2008). If in turn greenFP is located proximal to the degradation initiation site, proteasomal processing results in the release of ~60 kDa tFT fragments containing both FPs and a short ~6 kDa tail (Figure S5A). In this case the timer can be still used to measure local protein age but it no longer reports on protein degradation kinetics (Figure 6B). From these observations we can derive a short list of recommendations for design and use of tFTs as reporters of protein age and degradation:
The time range of a tFT is mostly determined by the maturation kinetics of the slower-maturing FP (Khmelinskii et al., 2012), although the maturation kinetics of the faster-maturing FP, the extent of proteasomal processing and potential FRET between the two FPs also contribute (Figures 2A, 6A and S4A). Thus, to construct a tFT for a given time range, a slower-maturing FP with maturation kinetics matching the time scale of interest should be paired with a faster-maturing FP that is as bright and fast-maturing as possible to lower the detection limit of the timer (Khmelinskii and Knop, 2014).
Ideally, characterization of a new tFT should involve testing both arrangements of FPs as reporters of protein age and protein degradation in the experimental system of interest to obtain qualitative system-specific estimates of FP maturation rates, efficiency of proteasomal degradation and time range of the tFT. In yeast, this can be done using the Pma1 protein age and Ubi-X-tFT degradation assays (Figures 1D and 2C).
For tFTs based on mCherry and greenFPs (or any timer in which one of the FPs is resistant to proteasomal degradation), the FPs should be arranged such that the degradation-resistant greenFP ends up distal to the degradation initiation site in the tagged protein of interest. Thus, the mCherry-sfGFP timer should be used for C-terminal tagging, whereas the sfGFP-mCherry timer – for N-terminal tagging. For internal tagging, both timers should be tested to first identify the degradation initiation site based on the size of processed tFT fragments.
Generally, a tFT in which both FPs are efficiently degraded by the proteasome is preferable, assuming its time range is suitable for the intended analysis. However, tFTs with incomplete proteasomal degradation can be advantageous in some situations. For instance, although the time range of mCherry-sfGFP(cp8) and mCherry-mNeonGreen timers is in principle more suitable to study degradation of unstable proteins than mCherry-sfGFP (Figure 6A), the mCherry-sfGFP timer could facilitate detection of extremely unstable and low abundance proteins due to increased absolute greenFP fluorescence that comes from accumulation of processed tFT fragments. Moreover, processed tFT fragments can be exploited as a marker of proteasomal degradation. For example, for a protein tagged at the C-terminus with mCherry-sfGFP, accumulation of ~33 kDa fragments is indicative of proteasomal degradation (Figure S2C), whereas lysosomal/vacuolar degradation should produce ~26 kDa fragments (Shintani and Klionsky, 2004; Klionsky et al., 2008).
tFTs with incomplete proteasomal degradation such as mCherry-sfGFP can be used for systematic analysis of protein degradation as the mCherry/sfGFP intensity ratio in steady state is a monotonic function of protein half-life (Figure 6A). However, care should be exercised when comparing proteins degraded by the proteasome and lysosome/vacuole pathways, which produce distinct tFT degradation fragments and differ in local environment that can affect FP maturation and brightness (Khmelinskii and Knop, 2014).
To follow protein dynamics, fast-maturing FP tags are needed to detect proteins as early as possible after synthesis. Our analysis indicates that mNeonGreen (Lam et al., 2012) is a good alternative to sfGFP: it is similarly fast-maturing but significantly less resistant to proteasomal degradation in yeast than sfGFP (Figures 1 and 2). The molecular brightness of mNeonGreen is similar to sfGFP when using excitation/emission wavelengths optimal for sfGFP and ~5 times higher with wavelengths optimal for mNeonGreen (Figure S1). The only limitation is that filter sets matching the excitation/emission peaks of mNeonGreen or lasers around its 506 nm excitation peak are not commonly used in fluorescence microscopy. The cp8 circular permutation of sfGFP is another alternative. When used as a C-terminal tag, sfGFP(cp8) is completely degraded by the proteasome both in yeast and human cells, and its maturation kinetics and brightness are similar to sfGFP (Figures 5 and S3).
In conclusion, our study provides a detailed characterization of tFTs as reporters of protein age and degradation. This should facilitate their application in different areas of cellular and organismal research. Our work also emphasizes the notion that fluorescent proteins are not neutral tags. Careful characterization of their properties is required for correct interpretation of protein dynamics measurements across spatial and temporal dimensions.
Materials and Methods
Yeast methods and plasmids
Yeast genome manipulations (gene deletions and tagging) were performed using conventional PCR targeting, as described (Janke et al., 2004). Yeast strains and plasmids used in this study are listed in Supplemental Tables 1 and 2, respectively. Yeast codon-optimized sequences of all fluorescent proteins were obtained using full gene synthesis. All Ubi-X-tFT constructs are based on previously described Ubi-X-β-galactosidase fusions, where X is followed by a 40-residue sequence that starts with histidine (Bachmair et al., 1986). The sequence of the ubiquitin-independent ODC23 degron (MSCAQESITSLYKKAGSENLYFQ) was obtained from plasmid pCT334 (Renicke et al., 2013). The Don1 coding sequence was amplified from genomic DNA of strain ESM356-1 (Supplemental Table 1). Standard site-directed mutagenesis was used to introduce point mutations into sfGFP or Clover and change the linker between mCherry and sfGFP. Circular permutations of sfGFP were amplified from a plasmid carrying two copies of sfGFP fused with a short linker (GSGAG). The N-termini of circular permutations 3, 7, 8, 9, 12 and 13 are the amino acids at position 51, 129, 140, 145, 189 and 189 of sfGFP, respectively. All plasmid sequences are available upon request.
Models of tFT maturation and turnover
Fluorescence intensity curves depicting maturation of a pool of mCherry-sfGFP molecules initialized in the non-mature state in the absence of protein production and degradation (Figure 1A) were calculated using a two-step maturation model for mCherry (maturation half-times of 16.91 and 30.3 min) and one-step maturation model for sfGFP (maturation half-time of 5.63 min), as described (Khmelinskii et al., 2012). To examine the influence of greenFP on tFT maturation (Figure 1B), the maturation half-time of greenFP was varied between 5 and 45 min in 5 min steps. All curves were normalized to the point of complete maturation.
To examine the influence of greenFP maturation on the relationship between protein half-life and mCherry/greenFP intensity ratio in steady state (Figure 2A), mCherry/greenFP intensity ratios were calculated according to equation E33 (Supplemental Theory), using a probability of incomplete tFT degradation z = 0, mCherry maturation half-times of 16.91 and 30.3 min, greenFP maturation half-time between 5 and 45 min varied in 5 min steps and a population doubling time of 90 min.
To examine the influence of incomplete tFT degradation on the relationship between protein half-life and mCherry/sfGFP intensity ratio in steady state (Figure 6, A and B), mCherry/sfGFP intensity ratios were calculated for the mCherry-sfGFP and sfGFP-mCherry timers according to equations E33 and E34, respectively (Supplemental Theory), using a population doubling time of 90 min, mCherry maturation halftimes of 16.91 and 30.3 min for both full-length fusion and processed form, sfGFP maturation half-time of 5.63 min for both full-length fusion and processed form, a degradation rate constant of processed tFT fragments k2 = 0 and varying the probability of incomplete tFT degradation z between 0 and 1 in steps of 0.1.
To examine the influence of FRET within a tFT on the relationship between protein half-life and mCherry/sfGFP intensity ratio in steady state (Figure S4A), mCherry/sfGFP intensity ratios were calculated for the mCherry-sfGFP timer according to equation E39 (Supplemental Theory), using a population doubling time of 90 min, a probability of incomplete tFT degradation z = 0, mCherry maturation half-times of 16.91 and 30.3 min, sfGFP maturation half-time of 5.63 min and varying the FRET efficiency E between 0 and 1 in steps of 0.1. For a fixed FRET efficiency E (0, 0.5 or 1) (Figures S4, B-D), the calculations were extended by varying the probability of incomplete tFT degradation z between 0 and 1 in steps of 0.1, using a degradation rate constant of processed tFT fragments k2 = 0 and sfGFP maturation half-time of 5.63 min for both full-length fusion and processed form.
To examine how degradation of processed tFT fragments influences the relationship between protein half-life and mCherry/sfGFP intensity ratio in steady state (Figure S6), mCherry/greenFP intensity ratios were calculated for the mCherry-sfGFP and sfGFP-mCherry timers according to equations E33 and E34, respectively (Supplemental Theory), using a population doubling time of 90 min, mCherry maturation halftimes of 16.91 and 30.3 min for both full-length fusion and processed form, sfGFP maturation half-time of 5.63 min for both full-length fusion and processed form, a probability of incomplete tFT degradation z = 1 and varying the half-life of processed tFT fragments H2 between 10 and 1000 min, in addition to 0 and infinity. The half-life of processed tFT fragments H2 is related to the degradation rate constant of processed tFT fragments k2 as k2 = ln(2)/H2.
Fluorescence microscopy
Strains were grown at 30°C in low fluorescence medium (synthetic complete medium prepared with yeast nitrogen base lacking folic acid and riboflavin (CYN6501, ForMedium)) to 0.4−1.2×107 cells ml−1 and attached to glass-bottom 96-well plates (MGB096-1-2-LG-L, Matrical) using Concanavalin A (C7275, Sigma) as described (Khmelinskii and Knop, 2014). Images from the middle of the cell were acquired on a DeltaVision Elite system (Applied Precision), consisting of an inverted epifluorescence microscope (IX71; Olympus) equipped with an LED light engine (SpectraX, Lumencor), 475/28 and 575/25 excitation, and 525/50 and 624/40 emission filters (Semrock), a dual-band beam splitter 89021 (Chroma Technology), a 100x NA 1.4 UPlanSApo oil immersion objective (Olympus), an sCMOS camera (pco.edge 4.2, PCO) and a motorized stage contained in a temperature-controlled chamber. Image correction and quantification were performed using ImageJ (Schneider et al., 2012). Dark signal and flat field corrections were applied to all images as described (Khmelinskii and Knop, 2014). Outlines of mother and bud compartments were manually defined in the sfGFP channel using a 5 pixel wide segmented line with a spline fit and applied to the mCherry channel. Fluorescence measurements at the plasma membrane were corrected for background using autofluorescence of the growth medium measured in close proximity to each individual cell.
Flow cytometry
Strains were grown at 30°C in synthetic medium lacking leucine (to select for plasmids) to a density of 0. 4−1.2×107 cells ml−1. Single cell fluorescence intensities, forward and side scatter were measured for at least 6000 cells per sample on a BD FACSCanto RUO Special Order System (BD Biosciences) equipped with 488 nm and 561 nm lasers, 505 nm and 600 nm long pass filters, 530/30 nm and 610/20 nm band pass filters. Multi-spectral beads (3.0-3.4 μm Sphero Rainbow Calibration Particles (6 peaks), #556286, BD Biosciences) were used to control for fluctuations in excitation laser power. Data analysis was performed with Flowing Software 2 www.flowingsoftware.com): measurements were gated for cells, followed by gating for cells with fluorescence above background to exclude cells that lost the expression plasmids. Sample measurements were corrected for background using autofluorescence levels of a control strain carrying an empty plasmid. mCherry/sfGFP intensity ratios were calculated for each individual cell.
Colony fluorescence measurements
Strains were grown to saturation at 30°C in synthetic medium lacking leucine and pinned onto agar plates with synthetic medium lacking leucine in 384-colony format, with 4 technical replicates for each strain. For pinning a RoToR pinning robot (Singer Instruments) was used. Fluorescence intensities of colonies were typically measured after 24 h of growth at 30°C using an Infinite M1000 Pro plate reader (Tecan). Measurements in mCherry (587/10 nm excitation, 610/10 nm emission, optimal detector gain) and sfGFP (488/10 nm excitation, 510/10 nm emission, optimal detector gain) channels were performed from the top at 400 Hz frequency of the flash lamp, with 20 flashes averaged for each measurement. For relative measurements of molecular brightness of different greenFPs (Figure S1B), fluorescence intensities were additionally measured in a third channel (505/5 nm excitation, 516/5 nm emission, optimal detector gain). Measurements of control colonies without fluorescent protein expression were used to correct all measurements for background.
To acquire excitation spectra (Figure S1C), exponentially growing cultures of strains yMaM26 and yMaM32 (Supplemental Table 1) were transferred to glass-bottom 96-well plates (MGB096-1-2-LG-L, Matrical) and allowed to settle down. Fluorescence emission at 510/5 nm was measured from the bottom with excitation wavelengths between 350 and 600 nm, in 2 nm steps, at 400 Hz frequency of the flash lamp, with 1 flash per step.
Immunoblotting and time-course experiments
Strains were typically grown at 30°C in synthetic medium lacking leucine to 107 cells ml−1. One-milliliter samples were mixed with 150 μl of 1.85 M NaOH and 10 μl β-mercaptoethanol, and flash frozen in liquid nitrogen. Whole cell extracts were prepared as previously described (Knop et al., 1999), separated by SDSPAGE (NuPAGE Novex 4-12% Bis-Tris protein gels (Life Technologies)) followed by blotting and probed with mouse anti-HA (12CA5), mouse anti-myc (9E10), rabbit anti-GFP (ab6556, abcam), rabbit anti-YFP (Miller et al., 2015), mouse anti-Pgk1 (22C5D8, Molecular Probes) and rabbit anti-Zwf1 (Miller et al., 2015) antibodies. Secondary goat anti-mouse (IgG (H+L)-HRP, Dianova) and goat anti-rabbit (IgG (H+L)-HRP, Dianova) antibodies were used for detection. Imaging was performed on a LAS-4000 system (Fuji) using the increment exposure routine to avoid overexposure. Image quantification was performed in ImageJ (Schneider et al., 2012). All measurements were corrected for local background.
For cycloheximide chases, strains were grown at 30°C in synthetic complete medium to ~0.8×107 cells ml−1 before addition of cycloheximide to 100 μg/ml final concentration. One-milliliter samples taken at each time point were immediately mixed with 150 μl of 1.85 M NaOH and 10 μl β-mercaptoethanol, flash frozen in liquid nitrogen and processed for immunoblotting as detailed above.
For the proteasome inhibition experiment (Figure S2C), strains yMaM67 and yMaM957 (Supplemental Table 1) were grown at 30°C to ~0.8×107 cells ml−1 in synthetic medium lacking leucine and with raffinose (2% w/v) as carbon source. Expression of Ubi-R-mCherry-sfGFP was induced by addition of galactose (2% w/v) alongside inhibition of proteasome activity by addition of MG132 (40 mM stock in DMSO) to 80 μg/ml final concentration or DMSO as control. Whole cell extracts of samples collected before and after induction were prepared and analyzed by SDS-PAGE followed by immunoblotting, as detailed above.
Human cell culture and immunoblotting
The sequence encoding PARK6 was amplified from the plasmid pcDNA3.1/Pink1 (Meissner et al., 2011) and cloned into the pcDNA5/mCherry-sfGFP vector (Khmelinskii et al., 2012). pcDNA5/PARK6-mCherry-sfGFP(cp8) was cloned by overlap assembly PCR of a human codon-optimized sfGFP(cp8), which was obtained by full gene synthesis, into the pcDNA5/PARK6-mCherry-sfGFP background.
HEK293T cells were grown in Dulbecco’s modified Eagle’s medium supplemented with 10% (v/v) fetal bovine serum at 37°C in 5% (v/v) CO2. Transient transfections were performed using 25 kDa linear polyethylenimine (Polysciences) (Durocher et al., 2002) and harvested 24 h thereafter. To block mitochondrial protein import, cells were incubated for 3 h with 10 μM carbonyl cyanide m-chlorophenyl hydrazone (Sigma) before cell lysis.
Whole cell extracts were separated by SDS-PAGE followed by blotting onto a PVDF membrane and probed with a monoclonal anti-GFP antibody (11814460001, Roche). A secondary goat anti-mouse IgG-HRP antibody (sc-2005, Santa Cruz) was used for detection. Imaging was performed on a LAS-4000 system (Fuji).
Acknowledgements
We thank Christof Taxis for reagents and Marc Zimmer for comments on the manuscript. This work was supported by the Deutsche Forschungsgemeinschaft (DFG) through the Sonderforschungsbereich 1036 (SFB1036) (M.K., A.M., B.B. and M.K.L.) and the Hartmut Hoffmann-Berling International Graduate School of Molecular and Cellular Biology (HBIGS) (C.H.).Figure 1. Analysis of protein age with different tFTs.