Attenuation of green fluorescent protein half-life in mammalian cells

Pete Corish and Chris Tyler-Smith1

CRC Chromosome Molecular Biology Group, Department of Biochemistry, University of Oxford, South Parks Road, Oxford OX1 3QU, UK


    Abstract
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 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
The half-life of the green fluorescent protein (GFP) was determined biochemically in cultured mouse LA-9 cells. The wild-type protein was found to be stable with a half-life of ~26 h, but could be destabilized by the addition of putative proteolytic signal sequences derived from proteins with shorter half-lives. A C-terminal fusion of a PEST sequence from the mouse ornithine decarboxylase gene reduced the half-life to 9.8 h, resulting in a GFP variant suitable for the study of dynamic cellular processes. In an N-terminal fusion containing the mouse cyclin B1 destruction box, it was reduced to 5.8 h, with most degradation taking place at metaphase. The combination of both sequences produced a similar GFP half-life of 5.5 h. Thus, the stability of this marker protein can be controlled in predetermined ways by addition of the appropriate proteolytic signals.

Keywords: cyclin destruction box/green fluorescent protein/half-life/PEST sequence


    Introduction
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
The green fluorescent protein (GFP) from the jellyfish Aequoria victoria provides a visual marker that can be used in a wide variety of organisms (Chalfie et al., 1994Go; Tsien, 1998Go). The wild-type protein has been significantly improved for use in mammalian cells by the incorporation of mutations such as S65T and F64L, which change the absorption spectrum and increase the rate of fluorophore formation (Heim et al., 1995Go) and by `humanizing' the codon usage (Zolotukhin et al., 1996Go). GFP folds into a unique, compact structure, the ß-can (Ormö et al., 1996Go; Yang et al., 1996Go), that is strongly resistant to chemical denaturation (Ward and Bokman, 1982Go) and is observed to be stable in most cells (Cubitt et al., 1995Go). This is well suited to purposes such as marking cell lineages, but for monitoring dynamic processes such as gene expression or chromosome transmission, a less stable protein with a more rapid turnover would be more suitable.

The role of controlled protein degradation as a method of regulating many cellular biochemical processes has only recently been fully appreciated and the pathways involved in the accurate targetting of proteins for proteolysis are an increasingly important area of study. Failure of the key components in these pathways can be causative in human disease (Kishino et al., 1997Go; Matsuura et al., 1997Go) and responsible for incomplete cell-cycle progression (Ghislain et al., 1993Go; Finley et al., 1994Go; Pu and Osmani, 1995Go). Central to this process of degradation is the 26S proteosome, a 2000 kDa multi-protein complex which is distinct from the compartmentalized lysosomal proteolysis machinery and is responsible for the ATP-dependent turnover of 80–90% of the cell's protein content (Lee and Goldberg, 1998Go). Proteins are generally, but not exclusively, identified for systematic degradation in the proteosome by ubiquitination. The specificity of this process is in part defined by the action of a family of E2-ubiquitin conjugating enzymes, which catalyse the final ubiquitin addition, but is also conferred by sequence motifs in the target protein which act as proteolytic signals.

One such motif, the PEST sequence, is found extensively in short-lived proteins including metabolic enzymes, transcription factors and their regulators, signalling pathway components and certain cyclins (Rechsteiner and Rogers, 1996). PEST sequences are enriched for proline, glutamate, serine and threonine in a negatively or neutrally charged background and removal of this region from short-lived proteins results in more stable derivatives (Tyers et al., 1992Go; Pu and Osmani, 1995Go; Tsurumi et al., 1995Go). The metabolic enzyme ornithine decarboxylase (ODC) is a key regulatory control point in polyamine biosynthesis and is regulated by 26S proteosome degradation, albeit without prior ubiquitination, which is mediated by two PEST regions. Transfer of the carboxy-terminal ODC PEST region to a reporter enzyme, dihydrofolate reductase, resulted in an increased turnover rate of the DHFR protein (Loetscher et al., 1991Go).

An alternative mechanism of targetting proteins for rapid degradation is the `cyclin destruction box' (CDB), which leads to ubiquitination and degradation of A- and B-type cyclins by the 26S proteosome at the end of mitosis. This process is promoted by the anaphase promoting complex (APC), which possesses E3-ubiquitin ligase activity, as part of its role as regulator of the metaphase–anaphase transition checkpoint. Transfer of the CDB to protein A led to destabilization of this protein in Xenopus extracts (Glotzer et al., 1991Go).

We wish to use destabilized GFPs to assay chromosome transmission in mammalian cells. We therefore engineered fusion proteins that link GFP to an ODC PEST sequence and a cyclin B1 destruction box region, to determine if these signals can reduce the resistance of wild-type GFP to proteolysis. The stabilities of these fusion proteins in mouse cells were compared with that of unmodified GFP.


    Materials and methods
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 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Plasmid construction

All constructs were produced in a modified version of the high-level mammalian constitutive expression vector pCAGG-Zeo (Niwa et al. 1991Go), allowing coordinated expression of GFP and Zeocin (Invitrogen) drug resistance from a single transcript incorporating an internal ribosome entry site. The `humanized' S65T variant of GFP (Heim et al. 1995Go) was cloned as a 720 bp Ecl136II/XbaI fragment into EcoRI-cut pCAGG-Zeo, after end-filling of recessed 3' ends. In the resulting plasmid, the GFP gene was flanked by reconstituted EcoRI sites, the downstream 3' site was inactivated to facilitate subsequent cloning steps. The plasmid was further modified by removal of a 760 bp region containing a polyoma virus origin of replication resulting in the 7.8 kb plasmid pCAGG–GFP.

The carboxy-terminal PEST sequence corresponding to codons 423–449 of the mouse ornithine decarboxylase gene (mODC) was amplified by PCR from LA-9 genomic DNA using the primers ODC-1 (5'-GAG CTG TAC AAG CAT GGC TTC CCG CCG GAG-3') and ODC-2 (5'-GAG CTG TAC ATT AAC GGT CCA TCC CGC TCT C-3') which contain BsrGI sites (underlined). A 100 bp product was isolated and cloned into the unique BsrGI site at the 3' end of the GFP gene in pCAGG–GFP to produce an in-frame fusion of the PEST sequence to the carboxy terminal of the GFP protein. Sequence analysis of this plasmid, pGFP–PEST revealed a single base deletion in the PCR-derived fragment at the intended TAA termination codon (in bold). This frameshift mutation resulted in the addition of a sequence of nine amino acids (NVQVKLPRL), not found in either mODC or GFP, before a TAG termination codon was reached (Figure 1AGo).



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Fig. 1. (A) Structures of the variant GFP constructs expressed from the pCAGG-Zeo vector and their predicted protein product sizes. Additional amino acid sequences, resulting from cloning procedures, not found in either GFP, murine ODC PEST or cyclin destruction box (CDB), are shown shaded. Amino acid numbering refers to residue positions in the original mouse ODC and cyclin B1 proteins and not to the GFP fusions. (B) SDS–PAGE analysis of immunoprecipitated 35S-radiolabelled GFP variants from stably transformed LA-9 cell lines.

 
Cell-cycle specific degradation of GFP was achieved using an amino-terminal fusion of the mouse cyclin B1 destruction box (CDB) motif to GFP. A 348 bp fragment from the 5' end of mouse cyclin B1 was amplified by PCR from embryonic stem cell genomic DNA using primers CYC-1 (5'-CGG AAT TCT CTG ATT TTG GAG GAG CCA TG-3') and CYC-2 (5'-ATG AAG CTT TTC AAG TTC AGG TTC AGG C-3') which included EcoRI and HindIII restriction sites (underlined), respectively. The resulting amplified product was cloned, after digestion, into partial HindIII/EcoRI-cut pCAGG–GFP producing an in-frame fusion to the N-terminal end of GFP, resulting in plasmid pCDB–GFP. Five amino acids (KLAAT), derived from vector sequences, separate the 3' end of the cyclin fragment from the original GFP initiation codon. Sequencing confirmed the cyclin sequence was free of PCR-induced mutations and corresponded to a published variant murine B1 cyclin (Hanley-Hyde et al., 1992Go).

Sub-cloning of the mODC PEST sequence from pGFP–PEST into the BsrGI site of pCDB–GFP resulted in plasmid pCDB–GFP–PEST in which both putative proteolytic targetting signals were incorporated in the same GFP fusion protein.

Cell culture and transfections

Mouse cell line LA-9 was maintained in DMEM supplemented with 10% fetal calf serum (Globepharm) and selection for drug-resistant lines made with 600 µg/ml Zeocin. Cell lines with stable integrations of the various GFP constructs were produced by electroporation of LA-9 cells with 10 µg of linearized plasmid DNA. Colonies were visualized and selected for GFP expression using an inverted fluorescence microscope (Leica DMIRB) with an FITC filter set (Leica I3) and several independent colonies chosen for expansion and subsequent analysis. For analysis of the CDB–GFP dynamics, cells were synchronized in S phase by treatment with 2 µg/ml aphidicolin (Sigma) for 16 h. FACS analysis of GFP fluorescence was performed on a Becton-Dickinson Facscalibur machine using the supplied FITC filter set.

Metabolic labelling of proteins and immunoprecipitation

GFP half-life was determined by immunoprecipitation of pulse-labelled proteins. Approximately 1x106 cells (30–40% confluent in 10 cm plates) were incubated with 50 µCi of 35S-labelled methionine (TranSLabel, ICN Pharmaceuticals) in 4 ml of methionine-deficient DMEM (Sigma) for 4 h at 37°C to pulse label all proteins including GFP. After this time, the medium was replaced with fully supplemented, unlabelled DMEM and an initial sample taken to determine the incorporation of the 35S-labelled methionine at the start of the experiment. Between five and seven further samples were taken during the next 48 h chase period at approximately 6 h time points.

For each time point, cells were lysed in 1 ml of TETN250 buffer (25 mM Tris–Cl pH 7.5, 5 mM EDTA, 1% Triton X-100, 250 mM NaCl, 1 mM PMSF), the soluble fraction pre-cleared with rabbit serum and GFP selectively immunoprecipitated using 1 µl of undiluted rabbit anti-GFP polyclonal antibody (IgG fraction; Clontech). GFP–antibody complexes were collected using a 10% formalin-fixed Staphylococcus aureus cell suspension (Immunoprecipitin; Life Technologies) and washed with lysis buffer before separation on 12% SDS–PAGE gels by standard methods (Harlow and Lane, 1988Go). Gels were dried under vacuum after fixing in methanol–acetic acid–glycerol and electronic autoradiography was carried out using an Instant Imager (Packard Bell Instruments). The total radioactivity in bands corresponding to GFP or its tagged variants was calculated by volume integration using the built-in software after background correction. The GFP half-life was calculated by linear regression analysis of log(total radioactivity per band) against time.


    Results
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 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Stably transformed cell lines containing each of the three fusion protein constructs or the S65T unmodified CAGG– GFP (Figure 1AGo) were produced in mouse LA-9 cells. The integrity of the fusion proteins produced was determined both by fluorescence microscopy and by immunoprecipitation. Visually, all cell lines displayed green fluorescence, which indicated that they were expressing the GFP fusions although with intensity levels and distribution patterns specific for each construct (described below). Immunoprecipitation of 35S-labelled proteins using a polyclonal anti-GFP antibody produced bands at the predicted size for each of the fusion proteins, from GFP at 27 kDa (Tsien, 1998Go) to the combination-tagged CDB–GFP–PEST variant at 43 kDa (Figure 1BGo). Several additional bands, 39 kDa or larger, were also observed even in the parental untransfected LA-9 control lane and represent non-GFP proteins with affinity for the polyclonal antibody. While one of these bands co-migrates with the CDB–GFP fusion protein, the relative intensity of the bands indicates that this was an unrelated band and did not result from contamination of the parental cell line, which was consistently non-fluorescent.

Immunoprecipitated samples of pulse-chase labelled protein extracts were taken over a 48 h period to determine the stabilities of each of the fusion proteins (Figure 2Go). As predicted from the structure, CAGG–GFP was a stable protein whose destruction followed first-order kinetics with a half-life of approximately 26 h. The fusion protein containing the C-terminal mouse ODC PEST sequence was also degraded with first-order kinetics, but at an increased rate, resulting in a reduced half-life of 9.8 h. This represents a significant destabilization of the protein of 2.6-fold compared to CAGG–GFP.



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Fig. 2. GFP stability is reduced in variants carrying proteolytic signal sequences. Linear regression lines for each of the four tested GFP variants are shown (r2 = 0.937, 0.93, 0.919 and 0.964, respectively). Horizontal dashed lines indicate 50% decreases in activity on a log scale, representing one to four half-lives. No 48 h time point is displayed for CDB–GFP–PEST as band activity levels were indistinguishable from background.

 
There was no significant difference in fluorescence intensities between GFP and GFP–PEST lines when examined visually despite the increased turnover of the fusion protein. This observation was quantitatively confirmed by subsequent FACS analysis and indicates that the steady-state levels of the protein are the same. This was not the case for CDB fusion proteins that showed highly variable levels of expression between individual cells from unsynchronized cultures of the same clonal line. In particular, cells undergoing mitosis (as determined by a `rounding off' phenotype) or cells which had recently completed cytokinesis appeared non-fluorescent and revealed a bimodal distribution in FACS analysis (Figure 3DGo). The left-hand peak in this profile corresponded to cells undergoing mitosis which, nevertheless, showed minimal levels of fluorescence 2–3 times brighter than the non-fluorescent parental LA-9 cells (Figure 3BGo). During this period of the cell cycle, many proteins are being degraded by the 26S proteosome and so complete removal of the excessive amounts of GFP produced may be hindered by saturation of the degradation machinery.



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Fig. 3. (A) Cyclical changes in fluorescence intensity in aphidicolin S-phase-synchronized cultures of LA-9 cells expressing the CDB–GFP fusion protein compared with wild-type GFP. Fluorescence was measured as the portion of cells 200x brighter or greater than the non-fluorescent LA-9 parental line. This results in a percentage of cells (15–20%) from the left-most tail of the positive control being classed as non-fluorescent. (B) FACS profile of unsynchronized untransfected LA-9 cells used as the negative control (mean fluorescence = 2.9). (C) FACS profile of S-phase-synchronized cells expressing CAGG–GFP (mean fluorescence = 227). (D) and (E) FACS profiles of unsynchronized and S-phase-synchronized cells expressing CDB–GFP (mean fluorescence = 124, with geometric mean of 53.7 and 170, respectively). (F) Phase contrast (upper panels) and fluorescent (lower panels) images of a pair of cells, expressing the CDB–GFP fusion protein, photographed over a 25 h period, illustrating the dynamics of GFP fluorescence throughout the cell cycle.

 
The cyclin destruction box produced the most significant effect on protein stability, as the CDB–GFP fusion protein had a half-life of 5.8 h averaged over the 48 h sample period (Figure 2Go). However, this rate was not constant during this time, as demonstrated when synchronized, rather than unsynchronized, cultures were examined (Figure 3AGo). In this case, the majority of GFP turnover was seen to occur as cells entered mitosis, during which the GFP half-life was 4.9 h. Outside the mitotic phase, the protein remained predominantly stable. Thus, the figure of 5.8 h reflects the fact that not all cells in the unsynchronized culture are actively degrading GFP to the same extent.

The addition of the PEST motif to this CDB–GFP protein marginally reduced the average half-life further to 5.5 h. The outstanding feature of this latter `combination' fusion is that overall levels of fluorescence were the lowest for any of the fusions, such that no labelled protein could be detected on the final time point sample. Also, FACS analysis of unsynchronized cells expressing CDB–GFP–PEST showed a bimodal distribution of fluorescence where the lower peak coincided with the non-fluorescent negative control (data not shown). Thus, addition of the PEST region does not appear to make a significant difference to fluorescence when protein levels are non-limiting; at lower concentrations, its effect is more evident.

The most prominent consequence of fusing the GFP sequence to the cyclin destruction box was highlighted through a time-dependent analysis of fluorescence development and decay in cells from synchronized culture (Figure 3A and FGo). Cells from lines containing either the CAGG–GFP or CDB–GFP variants were blocked in S phase using the DNA synthesis inhibitor aphidicolin (Sourlingas and Sekeri-Pataryas, 1996Go). The FACS profiles of these cells immediately after removal of the synthesis block showed similar distributions of fluorescence intensities, with wild-type GFP being slightly higher (Figure 3C and EGo). However, over a period of 26 h, corresponding to approximately one and a half cell cycles for LA-9 cells, the population profiles differed dramatically. In order to measure the early stages of fluorescence loss, observed as an initial moderate shift in the FACS profile peak to the left in Figure 3DGo, cells were defined as fluorescent if they exhibited a relative fluorescence intensity of >=200 units. Inevitably, below this threshold some cells were included as non-fluorescent despite expressing significant levels of GFP. By this criterion, the percentage of fluorescent cells remained constant (80–85%) in the line expressing CAGG–GFP, whereas the CDB–GFP line showed a loss of fluorescence down to 30% as the synchronized cells entered mitosis. Fluorescence levels increased subsequently as cells progressed through G1 and S phase before a second decrease occurred as cells entered their second mitotic division. The reduced peak in fluorescence levels during this second cycle (62% compared with 78%) results from the loss of synchronization in the population after this length of time. These dynamic changes in CDB–GFP fluorescence with cell cycle progression are illustrated in Figure 3FGo, in which each cell of a synchronized pair undergoes mitosis resulting in reductions in fluorescence levels. At the four-cell stage, once the cells are in interphase, fluorescence levels have returned to a maximum and are comparable between all cells. Although it is not possible from these data to establish the exact timing of these changes with respect to the cell cycle, the pattern is consistent with that reported for normal cyclin B1 accumulation and degradation (Glotzer et al., 1991Go).


    Discussion
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 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
The most notable impact of green fluorescent protein technology to date in cell biology has been its use in fusion proteins for monitoring a specific protein's localization, trafficking and processing (Corbett et al., 1995Go; Rizzuto et al., 1996Go; Oatey et al., 1997Go; Tsien, 1998Go). For these purposes, GFP already has been engineered to be brighter and to exhibit a wider range of excitation and extinction frequencies, which are amenable to double labelling studies (Tsien, 1998Go). However, further development of GFP will undoubtedly take the direction of utilizing GFP per se as an indicator of cellular state, as opposed merely to conferring fluorescence on previously non-fluorescent proteins of interest. It is to this class of modification that the GFP variants described in this work belong.

The reduction in GFP stability seen in the mODC PEST-tagged variant, while significant, still places GFP at the higher end of stability compared with more traditional reporters of gene activity such as luciferase which has a half-life of approximately 3 h (Thompson et al., 1991Go). In addition, the lack of enzymatic amplification of a signal with GFP means that its sensitivity is limited, as cytoplasmic concentrations of approximately 1 µM are required to distinguish signal from auto-fluorescent background (Niswender et al., 1995Go). However, as most enzymatic methods involve cell disruption, this improvement in using GFP non-invasively with a shorter half-life will be suitable for many applications. In particular, for monitoring chromosome transmission, where expression of GFP to readily detectable levels is not problematic, the shorter half-life means that parentally derived cytoplasmic GFP is removed quickly, allowing the measurement of de novo GFP synthesis from the daughter cells.

GFP may yet be a viable alternative for in vivo gene expression studies if the destabilization can be improved to reduce the half-life further. A destabilized GFP variant, d2GFP, has recently been described (Li et al., 1998Go) which utilizes a PEST region very similar to that used in this work. However, the reported half-life of this variant is 2 h and further reports of a 1 h variant (d1GFP) are based on fluorescence measurements, not biochemical purification. The discrepancy between those claims and the results presented here has been investigated to distinguish between cell differences, experimental protocols and intrinsic properties of the GFP variants used. We have confirmed the increased rate of degradation of the d1GFP variant, compared with the GFP–PEST described here (data not shown), and using the more accurate method of pulse-chase labelling we have determined its half-life to be 50 min in human HT1080 cells.

This result is remarkable considering the overall homology between the two mODC PEST sequences used. The d2GFP variant is extended compared with GFP–PEST by only 13 amino acids, but the core PEST region is maintained between both. This clearly implicates `non-PEST motif' residues as having a major contributory role to the rapid turnover of the mouse ornithine decarboxlase protein ordinarily. What is more significant, however, is that in the most unstable d1GFP variant the conserved Glu residues in the PEST region are mutated to Ala. While these changes would theoretically predict a weaker PEST motif, in practice the alterations result in a 50% reduction in protein stability. Consequently, the presence of a PEST motif within a protein merely implies a propensity to instability and cannot indicate the magnitude of the degradation rate even when close to or identical with the consensus PEST sequence.

While the patterns of protein degradation for the cyclin–GFP fusion proteins are more complex than the simple linear profiles obtained with the PEST fusions, these variants are potentially useful as non-invasive indicators of cell-cycle status. The FACS profile of unsynchronized cells expressing these proteins (Figure 3DGo) is an indirect measurement of the proportion of cells in defined stages of the cell cycle. Although more detailed investigation of the correlation between fluorescence levels and cellular status is necessary, the lower peak of the fluorescence distribution corresponded broadly to mitotic and early G1 cells. Therefore, this form of GFP could be used as an alternative to procedures, such as propidium iodide staining, which require fixation with concomitant cell death. Indeed, in the cell synchronization experiments described (Figure 3A–EGo), the fluorescence levels were a direct indication of the efficiency of the aphidicolin treatment.

However, it is still debatable as to how accurately GFP fluorescence intensity is a reliable indicator of GFP levels in a cell. Two factors that significantly affect the fluorescent properties of GFP, namely the requirement for post-translational oxidative fluorophore formation and the sensitivity to cellular pH, could result in fluorescence intensities being lower than actual GFP concentration. Western analysis indicates that a good correlation exists between fluorescence intensity and protein concentration (Li et al., 1998Go), but does not distinguish between immature, non-fluorescent and mature, fluorescent forms of GFP.

Considering this correlation between fluorescence levels and protein content, it is paradoxical that cell lines containing the unstable GFP–PEST construct (half-life 9.8 h) have fluorescence intensities not significantly different to those of lines containing its more stable GFP counterpart (half-life 26 h). Differences in steady-state protein levels between these lines would predict that the more unstable lines are also the least intense for GFP fluorescence, contrary to the actual observation. A similar observation was reported for the d2GFP variant (Li et al., 1998Go), although in our experimental system lines containing this construct evidently displayed weaker fluorescence both in FACS analysis and by microscopy. Although this could indicate that non-fluorescent immature GFP molecules are preferentially degraded, we believe this to be unlikely. Instead, it is probable that proteolytic products from 26S proteosome activity maintain a degree of fluorescence (Tsien, 1998Go) but are not detected by the immunoblot technique used to measure half-life, thereby increasing the apparent GFP concentration. Thus, reduction in fluorescence intensity is proportional to a decrease in GFP half-life, but not to the same magnitude. The d1GFP variant that is 30 times more unstable than the wild-type GFP is, nonetheless, only ~70–80% less intense by FACS analysis.

The results also shed some light on the biology of proteolytic signalling itself. The GFP–PEST fusion construct has been tested in human HT1080 and HeLa-S3 cell lines where the fusion protein localized to give a punctate pattern. The distribution of GFP foci appeared limited to the cytoplasm, partially concentrated around the nuclear envelope (data not shown). This phenotype was only seen in LA-9 cells when the brighter EGFP variant was fused to mouse ODC PEST, although the phenotype was less pronounced than in human cells. This mottled pattern is unlikely to be due to sub-cellular localization of the 26S proteosome, which is known to be found in both the nucleus and cytoplasm (Coux et al., 1996Go), but may represent natural sequestering of proteins marked for degradation away from the active cellular protein conterparts. Equally, the phenotype may be artefactual, with the very high levels of expression from the pCAGG vector resulting in protein aggregation complexes.

The distribution of the CDB–GFP protein was also consistent with known cyclin B1 characteristics (Hagting et al., 1998Go). A pronounced nuclear localization was observed in mouse and human HT1080 cells (Figure 3FGo, 0 h, lower panel). Nuclear-cytoplasmic translocation is a normal feature of cyclin B1 expression and is mediated by a putative cytoplasmic retention or nuclear export signal between residues 129 and 157 (Hagting et al., 1998Go; Toyoshima et al., 1998Go) which is absent from the CDB–GFP construct (Figure 1AGo). In this respect, CDB–GFP is a useful starting point for the in vivo analysis of the 5' region of the gene which is responsible for the dynamic movement of the cyclin B1 protein throughout the cell cycle.

In conclusion, it is evident from the results using the PEST and CDB fusions that the barrel-like structure of GFP is not refractory to major modifications that alter the basic properties of GFP biology. In these examples, GFP stability can be permanently compromised by the addition of the appropriate proteolytic signal sequences. Engineering opportunities are, however, limited to the carboxy- and amino-terminal ends for GFP, as internal modifications destroy the protective barrel structure and expose the fluorophore rendering it non-fluorescent (Dopf and Horiagon, 1996Go). However, the ease by which new derivatives can be produced and tested directly in living cells will allow GFP to be tailored to a variety of individual uses.


    Acknowledgments
 
We thank Elizabeth Burns for plasmid donations and Steve Kain and Jason Li of Clontech for sharing unpublished results. P.C. was supported by the MRC and C.T.-S. by the Cancer Research Campaign.


    Notes
 
1 To whom correspondence should be addressed. E-mail: chris{at}bioch.ox.ac.uk Back


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 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
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Received December 11, 1998; revised August 26, 1999; accepted September 1, 1999.