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PEDS Advance Access originally published online on August 1, 2008
Protein Engineering Design and Selection 2008 21(10):613-622; doi:10.1093/protein/gzn040
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© The Author 2008. Published by Oxford University Press. All rights reserved. For Permissions, please e-mail: journals.permissions@oxfordjournals.org

Engineering of a monomeric fluorescent protein AsGFP499 and its applications in a dual translocation and transcription assay

Aynur Tasdemir1,4, Farid Khan2, Thomas A. Jowitt3, Lucia Iuzzolino1, Stefan Lohmer1, Sabrina Corazza1,5 and Thomas J. Schmidt4,5

1Axxam SpA, Via Olgettina 58, 20132 Milan, Italy 2Lumophore Limited, 20 Oakleigh Avenue, Timperley, Cheshire WA15 6QT, UK 3Faculty of Life Sciences, The University of Manchester, Michael Smith Building, Oxford Road, Manchester M13 9PT, UK 4Institute for Pharmaceutical Biology und Phytochemistry (IPBP), Westfälische Wilhelms-Universtität, Münster, Hittorfstraße 56, D-48149 Münster, Germany

5 To whom correspondence should be addressed. E-mail: thomschm{at}uni-muenster.de (T.J.S.)/sabrina.corazza.sc{at}axxam.com (S.C)


   Abstract
Top
 Abstract
Introduction
 Materials and methods
 Results
 Discussion
 Supplementary data
 Acknowledgements
 References
 
The tetrameric green fluorescent protein AsGFP499 from the sea anemone Anemonia sulcata was converted into a dimeric and monomeric protein by site-directed mutagenesis. The protein was engineered without prior knowledge of its crystal structure based on a sequence alignment of multiple proteins belonging to the GFP-family. Crucial residues for oligomerisation of AsGFP499 were predicted and selected for mutation. By introduction of a single site mutation (S103K) the A/B subunit was disrupted whereas two substitutions were necessary to separate the A/C subunit (T159K/F173E). This method can be applied as a general predictive method for designing monomeric proteins from multimeric fluorescent proteins. The maturation temperature was optimised to 37°C by a combination of Site-directed and random mutagenesis. In cell-based assays, the NFATc1A (nuclear factor of activated T-cells, subtype 1, isoform A)-AsGFP499 chimera formed massive cytoplasmic aggregates in HeLa cells, which prevented the shuttling of NFATc1A into the nucleus and consequentially its transcriptional activity. In contrast, the cells expressing the NFATc1A in fusion with our engineered dimeric and monomeric fluorescent mutants were homogeneously distributed throughout the cytoplasm, making these stable cell lines functional in both translocation and transcriptonal assays. This new dual cellular assay will allow the screening and discovery of new drugs that target NFAT cellular processes.

Keywords: AsGFP499/fluorescent/GFP/monomeric/NFATc1


   Introduction
Top
 Abstract
 Introduction
Materials and methods
 Results
 Discussion
 Supplementary data
 Acknowledgements
 References
 
In the early sixties, the first member of the green fluorescent protein (GFP) family, AvGFP, was discovered in the jellyfish, Aequorea victoria (Shimomura et al., 1962Go). It forms a β-can structure consisting of 11 anti-parallel β-sheets that is crossed by a buried {alpha}-helix (Ormo et al., 1996Go; F.Yang et al., 1996Go; T.T.Yang et al., 1996Go). The chromophore, an extended, conjugated {pi}-system located on the helix, is autocatalytically formed by the tripeptide sequence, S65/Y66/G67, which gives the protein its fluorescence (Heim et al., 1994Go; Cubitt et al., 1995Go). The unusual nature of its chromophore, unique among known chromoproteins, is the reason why GFP has become a very popular tool in cell biology as reporter gene, selection marker, fusion tag and biosensor (F.Yang et al., 1996Go; T.T.Yang et al., 1996Go; Tsien, 1998Go; Lippincott-Schwartz and Patterson, 2003Go).

Currently, the GFP family comprises >100 GFP-like proteins with different spectral properties, most of them coming from the Anthozoa class (Matz et al., 1999Go; Lukyanov et al., 2000Go; Wiedenmann et al., 2000Go). Contrary to monomeric AvGFP, all known Anthozoa GFP-like proteins reported to date form oligomers and exist mostly as tetramers (Baird et al., 2000Go; Vrzheshch et al., 2000Go; Mizuno et al., 2001Go; Wiedenmann et al., 2002Go; Shagin et al., 2004Go). The oligomerisation does not impair their application as reporter genes, selection markers or biosensors, but limits their use as fusion tags to study, for instance, protein localisation and dynamics in living cells (Baird et al., 2000Go; Lauf et al., 2001Go; Mizuno et al., 2001Go). A possible solution to this problem is the genetic engineering of the fluorescent protein (FP) of interest by mutagenesis, creating mutants that form functional monomeric variants. For instance in DsRed, isolated from Discosoma corals, 33 mutations were needed to disrupt the tetrameric structure and to maintain its fluorescence (Matz et al., 1999Go; Campbell et al., 2002Go). Other groups used this approach to generate dimeric and monomeric variants from different FPs by mutating similar key residues (Karasawa et al., 2003Go; Karasawa et al., 2004Go; Wiedenmann et al., 2004aGo,bGo).

The FP under study, AsGFP499 cloned from Anemonia sulcata, is a protein composed of 228 amino acids (Wiedenmann et al., 2000Go). Here we present the generation and characterisation of dimeric and monomeric variants of the tetrameric AsGFP499 created by site-directed mutagenesis (SDM), which was guided by a multi-sequence alignment. Since most mammalian cells are cultured at 37°C, the maturation temperature of the FP should be also at 37°C to ensure optimal function in cell-based applications. Therefore, in addition to monomer design, we took a similar approach as Crameri et al. (1996Go) who applied random mutagenesis to select mutants with chromophore maturation at 37°C .

Furthermore we demonstrate here the suitability of the AsGFP499 mutants as fusion tags for NFATc1A (nuclear factor of activated T-cells, subtype 1, isoform A) in translocation and functional transcription assays. NFAT transcription factors are intimately involved in the regulation of multiple important biological functions including immune response, development and bone homeostasis (Ranger et al., 1998aGo,bGo; Graef et al., 1999Go; Graef et al., 2001Go; Crabtree and Olson, 2002Go; Graef et al., 2003Go; Horsley et al., 2003Go; Chang et al., 2004Go; Winslow et al., 2006Go).

Finally, HeLa cells, expressing the NFATc1A-FP chimeras and the luciferase (Luc) reporter gene under control of the NFAT response element were developed, providing a dual assay system which allows the detection of both, the NFATc1A translocation and NFATc1A-controlled transcriptional activity. This dual cell-based assay format will enable the discovery of new drugs targeting the NFATc1A signal transduction pathway.


   Materials and methods
Top
 Abstract
 Introduction
 Materials and methods
Results
 Discussion
 Supplementary data
 Acknowledgements
 References
 
Cloning and mutagenesis of bacterial expression vectors

The coding sequences for EGFP, DsRed2 and humanised AsGFP499 [hAsGFP499 (Supplementary data, Table S2)] were amplified from the vectors pEGFP-N1, pDsRed2-N1 (Clontech) and pcDNA3-hAsGFP499, respectively. The fragments were subcloned into pET28a(+)-vector (Novagen) and transformed into electro-competent BL21(DE3)Gold bacteria (Stratagene). Positive clones were confirmed by DNA sequencing.

Site-directed mutagenesis

SDM was carried out on the construct pET28a(+)-hAsGFP499 using the QuikChange SDM kit (Stratagene) according to the manufacturer’s instructions. The resulting vectors were transformed into electro-competent BL21(DE3)Gold cells. Mutations were confirmed by DNA sequencing.

Random mutagenesis

The coding sequences of hAsGFP499(S103K/R150E) and hAsGFP499(S103K/T159K/F173E) were cloned into pRSET-B vector (Invitrogen). Random mutagenesis was performed using the Genemorph random mutagenesis kit II (Stratagene) (Miyazaki, 2003Go). The resulting vectors were transformed into electro-competent BL21(DE3)Gold cells and grown at 37°C. Mutants of interest were sequenced and recloned into pET28a(+) vector.

Cloning of mammalian expression vectors

For mammalian cell expression, EGFP in pEGFP-N1 fusion vector (Clontech) was substituted with the coding sequences of hAsGFP499 and its mutants (pFP-N1). Further, the coding sequence of NFATc1A was amplified without its stop-codon from pcDNA3-NFATc1A by PCR. The fragment was inserted in frame to the FPs to generate the fusion constructs pNFATc1A-FP-N1. Positive clones were confirmed by DNA sequencing.

All primer pairs used for cloning are listed in the Supplementary data, Table S3.

Protein expression and purification

Fluorescent colonies were picked and grown with antibiotic, overnight on a shaker at 37°C in 2 ml liquid LB medium. Five millilitre of fresh medium was inoculated with 0.1 ml of the overnight bacterial cultures and grown to A600 of 0.6–0.8. Depending on the FP, expression was induced either at RT, 30 or 37°C by addition of IPTG to 0.5 mM. After overnight incubation, 4 ml of the cultures were centrifuged and pellets were lysed in 200 µl BPER-II (Pierce) solution added with 30 mM Tris–HCl pH 8.0, 300 mM NaCl, 10 µg/ml DNAse and EDTA-free protease-inhibitor-cocktail (Boehringer Mannheim). Finally, the bacterial lysates were centrifuged and the supernatants were analysed on pseudo-native SDS–PAGE (sodium dodecyl sulphate–polyacrylamide gel electrophoresis).

For large-scale protein purification, 100 ml LB medium containing kanamycin were inoculated with an overnight culture of fluorescent bacteria and grown at 37°C until A600 of 0.6–0.8 was reached. Protein production was induced with 0.5 mM IPTG. Bacterial pellets were lysed and centrifuged as described earlier, and subsequent protein purifications of the soluble fractions were performed on 1 ml His-Trap columns (GE Healthcare) using ÄKTAexplorer 100 (GE Healthcare). Bound proteins were washed with 10 mM Tris–HCl + 150 mM NaCl pH 8.0 buffer containing a gradient of imidazole (10–400 mM). Following protein elution by 500 mM imidazole, samples were exchanged into 10 mM Tris–HCl pH 7.4, 150 mM NaCl, using Amicon 10 filter units (Millipore). The protein concentrations were determined using the BCA Assay Kit (Pierce) and further used in biochemical characterisation.

Sodium dodecyl sulphate–polyacrylamide gel electrophoresis

Thermally denatured (for SDS–PAGE) and non-denatured [for pseudo-native SDS–PAGE (Baird et al., 2000Go)] protein samples prepared with x5 SDS sample buffer containing 200 mM DTT were loaded on 16% Tris-Glycine gels (Invitrogen) and separated in Tris-Glycine-SDS running buffer at pH 8.0. SeaBlue plus2 (Invitrogen) and (His)6-tagged DsRed2 (tetramer) and EGFP (monomer) were used as protein standards. Following separation, gels were illuminated with UV light to detect fluorescent bands and then stained with Coomassie blue.

Size-exclusion chromatography

The apparent molecular weights (MW) of the purified (His)6-tagged FPs were determined using size-exclusion chromatography (SEC) on a pre-calibrated Superdex 200HR10/30 column (GE Healthcare) connected to ÄKTAexplorer 100 (GE Healthcare) at a flow rate of 0.5 ml/min. Hundred microlitre purified protein samples at 2 mg/ml were separated using 10 mM Tris–HCl buffer containing 150 mM NaCl at pH 7.4, and the absorbance at 280, 400 and 480 nm was measured in real-time.

SEC coupled to multi-angle laser light scattering

Proteins were separated according to size on a 24/30 gel filtration column (GE, Amersham, UK) run in 20 mM Tris–HCl pH 7.4, 200 mM NaCl at 0.71 ml/min using a Dionex BioLC HPLC (Camberley, UK). Proteins resolved as one peak with little evidence of aggregation or other species being present. Proteins were passed through a Wyatt EOS 18-angle laser photometer (Wyatt Technology, Santa Barbara, CA, USA) with the 13th detector replaced with a QELS detector (Wyatt Technology) for the simultaneous measurement of hydrodynamic radius (Rh). This was coupled to an Optilab rEX refractive index detector (Wyatt Technology), and the Rh, MW moments and concentration of peaks were analysed using Astra 4.98 (Wyatt Technology). Analysis was performed using a differential refractive increment of 0.18 ml/g and mass was extrapolated using the Zimm plot.

Absorbance spectroscopy

A Cary 400 Scan UV-visible absorbance spectrophotometer (Varian) with a 1 cm path length microcuvette was used for absorbance spectra measurements in PBS (phosphate-buffered saline) at 25°C. The protein concentration of each AsGFP sample was determined using the BCA protein assay kit (Pierce) using GFPuv (Crameri et al., 1996Go) as a standard and Beer–Lambert law was applied to calculate the molar extinction coefficient (M–1 cm–1).

Fluorescence spectroscopy

The Cary Eclipse Fluorescence spectrophotometer (Varian) was used to obtain the fluorescence emission profiles of all proteins. For quantum yield (QY) measurements, EGFP was used as a standard, such that the AsGFP samples matched the absorbances at 490 nm in PBS. After excitation at 490 nm, the fluorescence emission spectra peaks for EGFP and for AsGFP samples were integrated using the Cary analysis software. The QY of the AsGFP samples were calculated by direct comparison of the integrated emission intensity with the known QY of 0.6 for EGFP (Tsien, 1998Go).

Cell line, culture conditions and transfections

HeLa cells were cultured in Alpha Minimum Essential Medium ({alpha}-MEM; Gibco-BRL), supplemented with 100 UI/ml penicillin, 100 µg/ml streptomycin and 10% heat-inactivated FBS at 37°C in 95% humidified air and 5% CO2. Cell transfections were performed using Fugene 6 (Roche).

HeLa/NFAT-Luc reporter cell line generation

The HeLa wt cell line was stably co-transfected with 0.8 µg NFAT response element, pNFAT-TA-Luc and 0.2 µg pPUR (Clontech). Antibiotic resistant clones were selected (2.5 µg/ml) and maintained (0.5 µg/ml) in puromycine containing medium. NFAT-TA-Luc carrying clones were isolated by limiting dilutions at 1 cell/well (c/w) in 96-well plates. Confluent clones were replicated using an automatic dispensing/diluting device (IGEL; OpalJena GmbH, Jena/Germany). The following day, cells were treated with 1 µM Trichostatin A (TSA; Sigma) overnight at 37°C and day after loaded with Tyrode-buffer. After injection of triton/luciferin mix the Luc activity in relative light units (RLU) was immediately recorded using a Lumibox CCD camera-based luminescence reader designed and built by Bayer Technologies GmbH (Wuppertal, Germany).

NFAT-Luc/NFATc1A-FP cell line generation

Stable HeLa/NFAT-Luc cells (MOCK) were transfected with pNFATc1A-FPs-N1 fusion vectors and antibiotic resistant clones were selected (2 mg/ml) and maintained (0.8 mg/ml) gentamicin (G418) containing medium. NFATc1A-FP positive clones were isolated by limiting dilutions at 1 c/w in 96-well plates. Confluent clones were replicated and following day incubated with Tyrode-buffer plus 100 µM adenosine 5'-triphosphate (ATP; Sigma) for 5 h at 37°C before measurement of Luc activity.

NFAT-Luc assay

MOCK and MOCK/NFATc1A-FP cells were seeded at 10 000 c/w density overnight in 96-well plates. The following day the growth medium was removed and cells were either incubated in Tyrode-buffer with different concentrations of ATP (1 mM to 100 nM) for 5 h at 37°C or pre-treated 1 h before stimulation with 50 µM ATP, with various concentrations of Cyclosporin A [CsA 5 µM–320 pM (Sigma)] or Tacrolimus [FK506 5 µM–10 pM (Sigma)] at 37°C. Each single concentration was tested in quadruplicates. The Luc activity was then measured. To determine the mean EC50 for ATP and IC50 for both CaN (calcineurin) inhibitors, CsA and FK506, RLU values were corrected against the background and normalised. The normalised RLU values were plotted versus the molarity of the substances using the PRISM analysis software (GraphPad, San Diego, CA, USA).

NFAT-translocation assay in living cells

Stable MOCK/NFATc1A-FP cells were grown on 35 mm glass bottom dishes at 37°C. For maximum gene expression, cells were treated overnight with 1 µM TSA. The following day, cells were either incubated in Tyrode-buffer alone or Tyrode-buffer plus 5 µM CsA or FK506 for 1 h at 37°C. Translocation was then initiated by adding 100 µM ATP and fluorescence images were captured every 60 s for 20 min using a confocal microscope (BioRad Leica).

Supporting online information

Multi-sequence alignment, list of primers, DNA sequence of hAsGFP499.


   Results
Top
 Abstract
 Introduction
 Materials and methods
 Results
Discussion
 Supplementary data
 Acknowledgements
 References
 
Multi-sequence alignment

At the time of this study, the crystal structure of AsGFP499 was unknown. Therefore, a multi-sequence alignment strategy was taken to identify key residues that may be involved in the subunit interfaces. These residues would be promising targets for SDM for the disruption of intermolecular contacts responsible for oligomerisation. Crystal structures of GFP-like proteins were retrieved from the protein data bank [PDB-codes (Berman et al., 2000Go): 1GGX (Wall et al., 2000Go); 1MOU (Prescott et al., 2003Go); 1UIS (Petersen et al., 2003Go); 1XQM (Wilmann et al., 2005Go); 1XA9 (Remington et al., 2005Go); 1B9C (Battistutta et al., 2000Go) and 1EMM (Palm et al., 1997Go)] and based on secondary structure elements these proteins were aligned manually with each other and with the protein sequences of 122 further GFP-like proteins of unknown crystal structures (Bairoch, 2000Go) (Supplementary data, Table S1). Using this alignment, the secondary structure elements of AsGFP499 were predicted. The predicted subunit interfaces of AsGFP499 were compared with the corresponding residues in the crystal structure of the native tetrameric DsRed (Wall et al., 2000Go; Yarbrough et al., 2001Go), which is known to occur at two chemically distinct protein interfaces. The hydrophobic interface, termed the A/B-interface, is formed between the chains A and B as well as C and D, whereas the hydrophilic A/C-interface is a result of the interaction of chains A and C as well as B and D.

Engineering of a dimeric variant of AsGFP499 by SDM

In the predicted A/B-interface of AsGFP499, the residue S103 is in a position homologous to T106 in DsRed, which has been previously described to form a hydrogen bond with its counterpart on chain B (Yarbrough et al., 2001Go). T106 in DsRed as well as S103 in AsGFP499, according to our alignment, are centrally located in the A/B-interface and completely surrounded by a number of hydrophobic amino acids (in DsRed: V96; V104; I225; V227, in AsGFP499: V93; V101; L122). The substitution of S103 with lysine could thus lead to a disruption of the A/B-interface, since two positively charged residues in close proximity to each other would significantly weaken the contact. Indeed, the mutation S103K in AsGFP499 led to a fully functional dimeric FP. We termed this mutant As-Dimer1. Interestingly, we found that the substitution of T106 with lysine in DsRed caused a complete loss of its fluorescence activity (data not shown).

Engineering of a monomeric variant of AsGFP499 by SDM

Based on our alignment, the analysis of the predicted A/C-interface of AsGFP499 revealed a more complex interaction network than in case of the A/B-interface. A number of promising mutation sites from our sequence alignment were targeted by SDM: (i) a salt-bridge between the residues E97 (chain A) and R150 (chain C), also occurring in DsRed (E100, chain A - R153, chain C), where a randomly inserted mutation R153E was included in the mutations leading to a monomeric variant (Campbell et al., 2002Go); (ii) hydrogen bond(s) between R157 (chain A) and T159 and/or S171, (both chain C), also observed in the corresponding residues of the eqFP611 from the sea anemone Entacmaea quadricolor, where Y157 forms a hydrogen bond to S171 (Petersen et al., 2003Go); (iii) a {pi}-stacking between F173 (chain A) and its counterpart (chain C).

In order to disrupt the salt-bridge, a variety of mutants of our dimeric variant As-Dimer1 was generated in which either of the two residues forming the salt bridge was altered. Many mutants lost their fluorescence activity or still formed stable dimeric proteins. However, we generated the mutant As-pMono1 [AsGFP499(S103K/R150E)], which showed partial monomeric character, but became fluorescent only at RT when expressed in Escherichia coli (E.coli) cultures.

Secondly, we addressed the mentioned hydrogen bond(s) between R157 (chain A) and T159 and/or S171 (both chain C). All three residues were altered by mutagenesis to perturb these potential hydrogen bond interaction(s), which, however, did not result in any functional monomeric variant.

Attempts to disrupt potential {pi}-stacking interaction between F173 (chain A) with its counterpart on chain C by replacing F173 with the positively charged residue arginine did not alter the dimeric structure (data not shown), but exchange by the negatively charged amino acid glutamate led to a variant with partial monomeric character (data not shown). However, the mutant AsGFP499(S103K/F173E) was fluorescent when expressed at RT in E.coli cultures, but not at higher temperatures.

A single site mutation on the A/C-interface did not produce a monomeric FP, however, several As-Dimer1 variants were generated by combining mutations targeting two or all of the residues predicted in the A/C-interface. Finally, the combination of two mutations (T159K/F173E) within the As-Dimer1 yielded a monomeric variant (As-Mono1) that still exhibited fluorescence, but only when expressed at RT in bacteria.

Increasing the maturation temperature of As-Mono1

An higher maturation temperature was achieved by subjecting As-pMono1 and As-Mono1 to three cycles of random mutagenesis and screening for fluorescent bacterial colonies that grew at 37°C. Although none of the As-Mono1 descendants revealed strict monomeric character on pseudo-native gels, one dimeric mutant, As-Dimer2 [AsGFP499(S94T/S103K/R150K/S171 N/K206R)], a mutant of As-pMono1, was selected for further characterisation, because it efficiently developed fluorescence at 37°C. Based on the mutations leading to improved maturation temperature of As-Dimer2, a number of variants of As-Mono1 were created, which led to the monomeric mutant As-Mono2 [AsGFP499(S94T/S103K/T159K/F173E)]. This mutant could be functionally expressed in cells at 37°C.

Pseudo-native SDS–PAGE

Pseudo-native SDS–PAGE of DsRed2 and wt AsGFP499 showed both proteins migrating as tetramers with an apparent MW between 64 and 98 kDa. In contrast, As-Dimer1 migrated between DsRed2 and EGFP as an apparent dimer at ~50 kDa with a faint band at lower MW. The mutants, As-Mono1 and As-Mono2, ran with similar mobilities as EGFP of ~28 kDa for a monomeric protein. In addition, As-pMono1 had both a monomer band and a faint band at the position of a dimer (shown by an arrow in Fig. 1A, Lane 8), indicating partial monomeric/dimeric character. The randomly generated mutant As-Dimer2 did not show a sharp band. Its oligomeric nature could not be deduced using this method (Fig. 1A, Lane 9). However, all bands were clearly fluorescent under UV excitation, indicating the formation of the mature chromophore (data not shown). Thermally denatured SDS–PAGE gel runs produced non-fluorescent bands, with all of the samples appearing as monomeric proteins with mobilities between 22 and 36 kDa. The Coomassie stained gel was also used to estimate the purity of the samples, which was >90%, except for DsRed2 (Fig. 1B, Lane 2).


Figure 1
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  		Fig. 1. SDS–PAGE of purified His-tagged recombinant proteins. Lane 1: Protein MW standard marker (labels express relative MW in kDa); Lane 2: DsRed2; Lane 3: EGFP; Lane 4: AsGFP499; Lane 5: As-Dimer1; Lane 6: As-Mono1; Lane 7: As-Mono2; Lane 8: As-pMono1 and Lane 9: As-Dimer2. (A) Coomassie blue stained pseudo-native SDS–PAGE. (B) Coomassie blue stained SDS–PAGE.

 
Size-exclusion chromatography

The SEC experiments showed that the determined apparent MW of As-Dimer1 (~39 kDa), As-Mono2 (~23 kDa) and EGFP (~27 kDa) are smaller in size than wt AsGFP499 (~78 kDa) and DsRed2 (~79 kDa) (Fig. 2A and B and Table I). Regarding the absorption profiles obtained in SEC-chromatograms for wt AsGFP499, As-Dimer1 and As-Mono2, As-Mono2 exhibited a decreased absorption at 480 nm together with an increased absorption at 400 nm relatively to the protein absorption at 280 nm (Fig. 2A).


Figure 2
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  		Fig. 2. SEC chromatograms of purified proteins. Absorbance at 280 nm (solid lines), 400 nm (dashed lines) and 480 nm (dotted lines). The chromatograms are normalised to the peaks at 280 nm. Overlaid plots of AsGFP499 (1); As-Dimer1 (2), As-Mono2 (3) (A); and As-pMono1 with dimeric and monomeric fraction (B).

 

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  		Table I. Biophysical properties of wt AsGFP499 and its mutants including GFPuv, EGFP and DsRed2

 
In agreement with the SDS–PAGE results, As-pMono1 displayed here two separated peaks at ~39 and 23 kDa. Interestingly, the peak corresponding to a dimer showed absorptions at 400 and 480 nm similar to wt AsGFP499 and As-Dimer1, whereas the monomeric peak was comparable to As-Mono2 (Fig. 2B and Table I).

SEC coupled to multi-angle laser light scattering

The absolute MW and the Rh for wt AsGFP499 and its mutants were determined by SEC-MALLS (SEC coupled to multi-angle laser light scattering). Wt AsGFP499 was confirmed to exist as a tetramer (~102 kDa). The dimeric nature of As-Dimer1 (~54 kDa) and As-Dimer2 (~51 kDa), as well as the monomeric nature of As-Mono2 (~30 kDa), was confirmed.

Absorbance spectroscopy

The absorption spectra of all AsGFP samples displayed two distinct maxima at 482 nm (major peak) and 406 nm (minor peak). Compared with wt AsGFP499, both As-Dimer1 and As-Dimer2 showed an increased absorption at 482 nm with respect to the minor peak. The most notable spectral change occurred within As-Mono2, where the 482 nm peak is reduced such that its intensity equals the 406 nm peak (Fig. 3A).


Figure 3
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  		Fig. 3. Absorbance spectra of AsGFP499 and its mutants at 1 mg/ml (A) and fluorescence emission spectra at 1 µM protein concentration in PBS (B).

 
Fluorescence spectroscopy

The fluorescence emission spectra of As-Dimer1 and As-Dimer2 protein samples were very similar to wt AsGFP499 with peak maxima at 498 nm, whereas As-Mono2 was slightly shifted towards longer wavelengths and emitted maximally light at 500 nm (Fig. 3B). Moreover, the QY of wt AsGFP499, As-Dimer1 and As-Dimer2 were also very similar with a QY ~0.5, while As-Mono2 had a QY of 0.4. Nevertheless, all proteins were extremely bright, having marginally less QY compared with EGFP (Tsien, 1998Go). Table I gives a summary of all spectroscopic properties.

Application of wt AsGFP and its mutants in cell-based assays

It has been frequently reported that oligomeric FPs are not ideal fusion partners for labelling target proteins (Baird et al., 2000Go; Vrzheshch et al., 2000Go; Lauf et al., 2001Go; Mizuno et al., 2001Go; Shagin et al., 2004Go). In the present work the suitability of AsGFP499 mutants as fusion markers was tested by investigating the translocation and transcriptional behaviour of NFATc1A when fused to hAsGFP499 or a mutant of interest in HeLa cells. Therefore, HeLa cells were stably transfected with the NFATc1A-FP fusion constructs and an NFAT-Luc reporter vector carrying three copies of the NFAT-binding site derived from the IL-2 promoter and fused to the Luc gene. Cells were stimulated with ATP, which increases intracellular Ca2+ concentration by acting on P2 purinoreceptors, inducing inositol 1,4,5-triphosphate (IP3)-mediated release of Ca2+ from intracellular stores (Badminton et al., 1996Go; Ferrari et al., 1999Go). Elevated Ca2+ activates the Ca2+/calmodulin-dependent protein phosphatase CaN, which in turn dephosphorylates cytoplasmic NFAT and thus promotes its nuclear translocation. In the nucleus dephosphorylated NFAT binds to promoter regions, containing NFAT response elements and modulates the transcription of targeted genes (Jain et al., 1993Go; Luo et al., 1996Go; Beals et al., 1997Go; Rao et al., 1997Go; Klee et al., 1998Go; Crabtree, 1999Go; Okamura et al., 2000Go). The immunosuppressive drugs CsA and Tacrolimus (FK506) were used to inhibit NFATc1A. Both drugs interfere with the NFAT signalling pathway at the level of CaN activation and block the nuclear shuttling of NFAT proteins, thereby preventing their action on target genes (Emmel et al., 1989Go; Mattila et al., 1990Go; Randak et al., 1990Go; Brabletz et al., 1991Go; Liu et al., 1991Go; Liu, 1993Go).

NFAT translocation assay

Live-cell images of NFATc1A-hAsGFP499 showed formation of large aggregates in the cytoplasm before stimulation (0 min). After addition of 100 µM ATP the aggregation presumably prevented an efficient translocation of the fusion protein into the nucleus (20 min) (Fig. 4). In a control experiment transiently transfected cells with NFATc1A-DsRed2 showed similar aggregation events (data not shown). In contrast, fusions of NFATc1A to As-Dimer1, As-Mono2 and EGFP as a positive control (Kehlenbach et al., 1998Go, Scott et al., 2001Go) were homogenously distributed in the cytoplasm prior to stimulation and translocated significantly into the nucleus within 20 min after addition of ATP. The translocation of NFATc1A-FPs was also investigated in presence of 5 µM CsA or FK506. Regardless of the FP used, the translocation of NFATc1A-FPs was blocked by both inhibitors and fluorescence was observed mainly in the cytoplasm after stimulation (Fig. 4).


Figure 4
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  		Fig. 4. Translocation of NFATc1A from the cytoplasm into the nucleus in living cells: HeLa/NFAT-Luc cells stably expressing NFATc1A in fusion with FPs were treated overnight with TSA and translocation was initiated by stimulation with ATP in presence or absence of CsA or FK506. Images were taken at time 0 and 20 min after stimulation with 100 µM ATP.

 
NFAT-Luc transcriptional assay

To assess whether the function as a transcription factor of NFATc1A is affected by fusion to a FP, its transcriptional activity was measured using a Luc reporter system. Positive control cells were either stably expressing NFATc1A only or as a fusion with EGFP. The results demonstrated that the cell lines harbouring the AsGFP constructs were comparable to the positive control cells. All cell lines were sensitive to activators and inhibitors of the NFAT signalling pathway (Fig. 5).


Figure 5
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  		Fig. 5. Concentration response curves for ATP, CsA and FK506 in HeLa/NFAT-Luc cells stably expressing the NFATc1A alone or in fusion with FPs. Cell lines are indicated as follows: MOCK (open diamond), NFATc1A-hAsGFP499 (multiplication sign), NFATc1A-As-Dimer1 (plus sign), NFATc1A-As-Mono2 (open square), NFATc1A-EGFP (inverted open triangle) and NFATc1A (open triangle). Cells were treated with various concentrations of ATP (A), CsA (B) and FK506 (C). The NFAT mediated Luc activity is given in RLU in (A), (B) and (C). Each data point represents the mean ± SE of four replicates. Normalised RLU values are shown for ATP (D), CsA (E) and FK506 (F).

 
Extracellular ATP induced Luc activity resulted in NFAT transcription activity in a dose-dependent manner. In all cell lines tested, including MOCK cells, the calculated mean EC50 values for ATP was ~125 µM, whereas the maximal RLU values obtained by the untagged or differently tagged NFATc1A varied significantly in the order: NFATc1A (~424 000) > NFATc1-EGFP (~250 000) > NFATc1A-As-Dimer1 (~77 100)=NFATc1A-As-Mono2 (~57 800) >> NFATc1A-hAsGFP499 (~11 650) >> MOCK (~1400). This indicates that the increased Luc activity was dependent on the recombinant expression of NFATc1A either alone or as a fusion with FPs (Fig. 5, Table II).


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  		Table II. In cell-based assays calculated mean EC50 and IC50 values for ATP, CsA and FK506

 
In presence of CsA and FK506, the Luc activity was reduced in a dose dependent manner. The mean IC50 values for these CaN-blockers were independent from the cell line used. The calculated mean IC50‘s for CsA and FK506 were ~286 and 6 nM, respectively (Fig. 5, Table II). The measured IC50‘s for FK506 and CsA were consistent with the reported inhibitory potency of FK506 (Siekierka et al., 1989Go; Schwaninger et al., 1993Go). Noteworthy, all NFATc1A over-expressing cells showed relatively high basal Luc activities compared with MOCK cells when incubated without ATP (Fig. 7). The higher the observed basal values, the higher maximal RLU values were observed by treatment with ATP.


Figure 7
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  		Fig. 7. The histogram shows the basal values of NFAT-driven Luc response measured in various HeLa cell lines stably co-expressing the NFAT response-element either with NFATc1A or NFATc1A in fusion with FPs.

 

   Discussion
Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
Supplementary data
 Acknowledgements
 References
 
When engineering an oligomeric protein to generate a monomeric version, knowledge of the structure allows the application of rational approaches to design mutants with the desired properties. Since the structure of AsGFP499 was unknown at the time of this study, secondary structure elements had to be predicted by multiple sequence alignment and compared with FPs with known 3D(three-dimensional)-structures. SDM was then used to modify amino acids that were predicted to contribute to the intermolecular contacts responsible for oligomerisation. A dimeric mutant (As-Dimer1) was obtained by mutation of a single residue at the hydrophobic A/B-interface, namely the polar residue (S103). The replacement of only hydrophobic residues in the A/B-interface had been applied in other oligomeric FPs, including DsRed monomerisation (Campbell et al., 2002Go; Karasawa et al., 2003Go; Wiedenmann et al., 2004aGo,bGo).

The disruption of the remaining A/C-interface was more challenging. It has been proposed that manipulations at this interface may cause a decrease of the folding temperature, because the interactions here are likely involved in the folding process of the β-can structure (Wiedenmann et al., 2005Go). Indeed, all of our A/C-mutants showed fluorescence at temperatures <37°C. In addition, we generated mutants that lost completely fluorescence or led to higher oligomerisation states. The optimisation of the maturation temperature of As-Mono1 was finally achieved by combining Site-directed and random mutagenesis.

The majority of FPs are known to form a compact, rigid globular β-can structure, presumably to shield the chromophore from solvent (Reid and Flynn, 1997Go; Zimmer, 2002Go). SEC-MALLS analysis provided not only data on absolute molecular mass, but also on apparent hydrodynamic size (Rh), which depends on shape (conformation) as well as mass (Table I). As expected from crystal structures of other FPs, wt AsGFP499 had a compact and tetrameric arrangement similar to DsRed2, with Rh values of ~3.2 nm. In the case of As-Dimer1 and As-Dimer2, the Rh values of 2.6 nm were an indication of its more tightly folded state and its expected structural arrangements of side-to side β-barrel cylindrical contacts (rather than an end to end arrangement). The Rh for As-Mono2 values could not be fitted due to concentration restraints. Nevertheless, As-Mono2 is also expected to be a tightly folded protein with a buried chromophore that allows it to exhibit fluorescence. The unique feature of As-Mono2 is that it can be excited equally at 406 nm and 482 nm using a UV or blue laser, respectively. The QY of the novel dimers were ~0.5 with As-Mono2 having marginally less QY. However, all proteins were extremely fluorescent and QY were comparable to EGFP (Table I) (Tsien, 1998Go).

In retrospect, the recently published X-ray structure of AsGFP499 (Nienhaus et al., 2006Go) gave us the opportunity to examine these targeted interface residues. The crystal structure confirmed that all of the predicted and mutated residues were important for oligomerisation to maintain interfacial interactions. Consistent with our multi-sequence alignment, analysis of the A/B-interface revealed that S103 in chain A forms a hydrogen bond with its counterpart in chain B. Furthermore, the A/C-interface displays all three predicted interactions: the salt-bridge between R150 and E97, the {pi}-stack between two phenylalanines, F173 from the A- and C-chain, respectively, and the hydrogen-bond between the amino acids T159 and R157 (Fig. 6A and B).


Figure 6
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  		Fig. 6. Molecular representation of the network of interactions at the interfaces of AsGFP499, (nomenclature of monomers according to the DsRed crystal structure that is used throughout as reference). (A) A/B (equivalent to A/D in crystal structure of AsGFP499)-interface, showing the interaction of serine 103 with its counterpart. (B) A/C (equivalent to A/B in crystal structure AsGFP499)-interface, illustrating important interactions for oligomerisation: the salt-bridge formed by glutamate 97 and arginine 150, {pi}-stacking between F173 and its counterpart, {pi}-cation interaction between F173 and R157 and a hydrogen bond between R157 and T159. (C) A water bridge connects the chromophore of chain C with H227 of chain A.

 
Spectroscopic and oligomeric studies of the mutants revealed some interesting features of AsGFP499 and perhaps FPs in general. In SEC experiments, the absorption profile of the dimeric fraction (480 > 400 nm) of As-pMono1 [AsGFP499(S103/R150E)] turned out to be different from the monomeric fraction (480 ≤ 400 nm) (Fig. 2A). This suggests an altered balance between the neutral and anionic protonation state of the chromophore depending on the oligomerisation state of the protein. Analysis of the crystal structure of AsGFP499 supports this hypothesis. Closer inspection of the A/C-interface interactions between the chromophore in chain A and the protein matrix of chain C, showed a network of hydrogen bonds, linking the phenolic oxygen of the chromophore in chain A with histidine 227 in chain C via four intervening water molecules (Fig. 6C). Thus, it is possible that the positively charged protonated histidine residue stabilises the negatively charged deprotonated form of the phenolic chromophore through this ‘water route’. This mechanism of stabilisation would be consistent with the increased absorption at 480 nm of the A/C-dimer. A chromophoric interaction has also been reported for DsRed within the A/C-interface (Wall et al., 2000Go).

Application of almost all cellular assay types for high throughput screening and high content screening are carried out by using stable cell lines, which guarantees more data consistency and less variability compared with cellular assay types based on transient gene expression methods (Wilson et al., 2002Go). Therefore a further goal of this study was to develop a cell line stably co-expressing the NFATc1A-FP chimeras and the NFAT-Luc reporter gene to evaluate its use in translocation and functional assays testing compounds that interact with the NFAT-signalling pathway. To our knowledge, this is the first dual-assay system in one stable cell line specifically designed for a single member of the NFAT-family.

Wt AsGFP499 in fusion with NFATc1A cannot be used for translocation assays, due to formation of aggregates that prevented any visible translocation into the nucleus (Fig. 4). In contrast, EGFP (Kehlenbach et al., 1998Go; Scott et al., 2001Go) and the mutants As-Dimer1 and As-Mono2 fusions translocated visibly into the nucleus. In the presence of CaN inhibitors, CsA and FK506, the translocation of all fusion products was efficiently blocked (Fig. 4). This demonstrates that the translocation event of the chimeras is dependent on NFATc1A, which is a known substrate for CaN.

Wt AsGFP499 and its derivatives do not interfere with the transcriptional activity of NFATc1A fusions. The transcriptional activity of NFATc1A-As-Dimer1 and NFATc1A-As-Mono2 was significantly increased compared with MOCK cells (55-fold and 40-fold, respectively). It is noteworthy that an 8-fold Luc activity of NFATc1A-hAsGFP499 with respect to the MOCK cells was detected, although we did not observe significant nuclear import in our translocation assays. A plausible explanation would be that very low concentrations of mono- and dimeric NFATc1A-AsGFP499 might exist at equilibrium with the predominant tetrameric form. These non-tetrameric fusion proteins would be capable of translocating into the nucleus and could hence be responsible for the observed increase of transcription activity. In presence of CaN inhibitors the transcriptional activity of untagged and fusion products of NFATc1A was blocked. This is consistent with our results from the translocation assay and confirms that the transcription of Luc relies on NFATc1A nuclear import. The EC50 value of ATP and IC50 values of CsA and FK506 determined demonstrate that the signal transduction mechanism of NFATc1A is not altered by the chimeras (Fig. 5, Table II).

All NFATc1A over-expressing cell lines showed increased basal Luc activity in our functional assay (Fig. 7). This phenomenon might be a result of a shifted equilibrium between phosphorylated and unphosphorylated NFATc1A in the cytoplasm. Since NFATc1A needs to be phosphorylated by specific cytoplasmic kinases to prevent its translocation in non-activated cells (Scott et al., 1997Go), the endogenous amount of kinase activity in NFATc1A over-expressing cells might not be sufficient to maintain the entire population of NFATc1A in the phosphorylated state. The increased basal values would thus be an artefact of over-expression.

In summary, the present investigations led to a successful conversion of tetrameric AsGFP499 into dimeric and monomeric variants guided by a rational strategy of sequence alignments. The exchange of only three amino acids led to a monomeric variant of AsGFP499. A fourth mutation had to be introduced in order to increase maturation temperature of the protein. This new FP, As-Mono2, shows different spectral properties and matures in the temperature range compatible for its use as fluorescence marker in cellular assay systems. The findings of this study have thus revealed interesting new details on the A/B and A/C subunit interactions responsible for oligomer formation of FPs. Furthermore, analysis of the absorbance characteristics of mono- and dimeric variants of AsGFP499 yielded new insights in protein-chromophore interactions mediated by intervening water molecules, which may be a general principle of interaction in some FPs with a higher oligomerisation state. More detailed theoretical investigations of this phenomenon will be an interesting subject for further studies.

Finally, using simultaneous stable transfections of HeLa cells with the NFAT-Luc reporter element and NFATc1A-As-Dimer1 or NFATc1A-As-Mono2 fusion constructs, we generated for the first time a cell-based assay system allowing the investigation of NFATc1A translocation and transcriptional activity within the same cell line. This system, and the perspective to develop further analogous assays by the fusion of As-Dimer1 or As-Mono2 with other transcription factors, represents a promising tool in the search for new drugs intervening with the function of transcription factors.


   Supplementary data
Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 Supplementary data
Acknowledgements
 References
 
Supplementary data are available at PEDS online.


   Footnotes
 
Edited by Andreas Kungl


   Acknowledgements
Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 Supplementary data
 Acknowledgements
References
 
We thank Daniele Carettoni for many fruitful discussions and help for the biochemical analysis, Paolo Guarnieri for the bioinformatic support, Johannes C. Hermann and Tod Flak for reading the manuscript.


   References
Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 Supplementary data
 Acknowledgements
 References
 
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Received April 21, 2008; revised June 27, 2008; accepted July 2, 2008.


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