The advantages of GFP-like proteins as fluorescent markers include their high stability and the ability to form for chromophore without auxiliary cofactors or substrate. Furthermore, FP fusion to target protein rarely affects the function of this protein. However, this is not always the case and sometimes fusion of GFP-like proteins to target protein may significantly affect its functions. For example, it has recently been shown that GFP strongly impairs actin-myosin interaction by binding to the actin-binding site of myosin [80
]. Another crucial advantage is that GFP-like proteins are non-toxic in most cases. GFP-like proteins were demonstrated to be highly resistant to many proteases [70
], detergents [37
], organic and chaotropic salts, alkaline pH and high temperature [71
]. The main trends of GFP-like protein applications in cell biology, disadvantages of fluorescent proteins, limiting their use, and methods to overcome these limitations are summarized in this section.
Reporters for gene activation and fusion tags. By using a fluorescent protein gene (reporter gene) that is under control of a promoter of interest and recording the FP fluorescence intensity, gene expression can be monitored. Gene of Aequorea
GFP was the first reporter gene [70
]. Henceforward, FPs have been successfully applied in a variety of cell cultures and organisms, such as bacteria [81
], yeast [82
], plants [83
], worms [84
], insects [60
] and vertebrates [84
]. Chimeric protein between FP and protein of interest (fusion protein) can be constructed by using standard cloning techniques. Fusion protein expression allows researchers to monitor target protein localization and to visualize dynamics of cell events.
The means of GFP-like protein applications in cell biology are widely diverse and include their use as partners for multicolor protein tagging [87
], as intracellular reporters of gene activation [89
], as markers of cell lineage during development and as markers of cell growth, including pathogenic bacteria and cancer cells [60
], as markers of protein, organelles and virus particles localization in living cells [92
], as population markers in symbiotic studies [95
], as reporters of bacterial phagocytosis [96
] and so on.
It is more preferable to use red FPs as fluorescent markers, because of higher tissue transparency and lower autofluorescence in this spectral region. Moreover, the use of red FPs in counterpart to Aequorea GFP increases the range of FPs applications, enabling multicolor labeling. The presence of cryptic introns in mRNA of wild-type DsRed1 and some others Anthozoa FPs can results in partial splicing in mammalian cells and cause premature translation termination with formation of less whole-length protein having low fluorescence intensity. This disadvantage can be easily overcome by removing putative cryptic introns as in the commercially available version of DsRed1 (Clontech, USA).
Many Anthozoa GFP-like proteins have a tendency to form high molecular weight aggregates both in vivo
and in vitro
]. Protein aggregation may hinder all possible applications due to considerable cellular toxicity and make impossible FRET-based applications (fluorescence resonance energy transfer), the study of protein-protein interactions and proper targeting to cell compartment. Aggregation generally takes place during heterologous over-expression of FPs in both bacterial and eukaryotic cells. FP aggregation can be observed as appearance of large fluorescent granules inside transfected cells, which results in ‘smearing’ of the fluorescent picture, so that nuclei and nucleoli are usually invisible.
The crystal structure of DsRed1 provided clues on the reasons for FP aggregation. Aggregates were suggested to form by properly folded native protein molecules, as they retain bright fluorescence. Two main reasons of protein aggregation were proposed. It can be driven by non-specific interactions of “sticky” hydrophobic patches on the molecular surface [98
]. Alternatively, it can be due to the electrostatic interactions between positively and negatively charged surfaces. However, DsRed1 was shown to contain no pronounced hydrophobic areas that may cause strong interactions between tetramers. A computer calculation of the electrostatic potential of tetrameric DsRed1 revealed that the protein surface is mostly negatively charged, except for a short N-terminal region of each monomer that contains a group of positively charged amino acid residues [41
]. Based on these observations, it was proposed that each tetramer is able form up to four salt bridges with adjacent tetramers, resulting in the net-like polymeric structure. Four valencies for electrostatic interactions make this structure very stable [41
Site-directed mutagenesis was used to solve the problem of FP aggregation, regarding positively charged patches at N-termini as a possible cause of aggregation. A variant of DsRed1, E57 (Val105Ala/Ile161Thr/Ser197Ala), was subjected to mutagenesis. Mutant E57 is characterized by fast maturation; its red fluorescence appears 2 times faster than of wild-type DsRed1 [84
]. As result, a mutant, denoted E57-NA with minimal level of aggregation both in vivo
and in vitro
, containing three substitutions (Arg2→Ala, Lys5→Glu, and Lys9→Thr) was created. E57-NA was very similar to E57 in terms of excitation–emission maxima, fluorescence brightness and maturation speed [41
], which made it promising for further use. A considerable increase in protein solubility by substitution of one to three positively charged amino acid residues at the N-termini to neutral or negatively charged residues represented a successful strategy for other Anthozoa GFP-like proteins (zFP538, zFP506, amFP486). These findings confirmed an electrostatic nature of interaction responsible for the formation of aggregates of GFP-like proteins.
Although the tetramerization complicates FP applications as partners for FRET and as fusion-partner for a target protein, oligomerization does not limit the use of FPs as markers of gene expression [10
]. Tetrameric nature of many GFP-like proteins can result in abnormal localization of a protein tagged by FPs. Furthermore, as the oligomerization of many proteins involved in signal transduction leads to their activation, the fusion of a signal protein to FPs can cause constitutive signaling. Intensive mutagenesis of DsRed1 resulted in monomeric variant mRFP1, containing 33 substitutions: 3 in the hydrophobic interface and 10 in the hydrophilic interface, 3 in short the N-terminal region, 13 internal to the β-barrel and 4 surface mutation with unknown effect on protein structure and function [12
]. Mutant mRFP1 possesses 1.3-times lower extinction coefficient and 3.2-times less quantum yield than DsRed1, but it has 10-times higher maturation speed. mRFP1 still contains fraction of “green” chromophore, which can impede with FRET and multicolorlabeling applications. Novel monomeric RFPs, such as mOrange, mStrawberry, mCherry, mRaspberry and mPlum have substantially enhanced maturation rate, brightness and photostability [13
]. Recently developed monomeric variant of eqFP578, TagRFP, has even higher brightness than mCherry, which would make TagRFP a protein of choice as a monomeric fluorescent tag in red region of the spectrum. The other monomeric mutant of eqFP578, mKate, has compared to mCherry brightness, but within spectral range of 650–800 nm mKate is essentially brighter than any known monomeric red and far-red fluorescent proteins. Thus mKate is preferable fluorescent marker in far-red region of the spectrum.
Several other approaches have been developed to overcome oligomerization of Anthozoa FPs. One of them is covalent head-to-tail linkage of double copies of the same FPs; formed tandem dimeric structure may prevent oligomerization. This approach was successfully applied for HcRed1 [10
] and dimeric variants of DsRed1 [12
]. It was shown that tandem proteins, containing 4- and 12-residue linker between monomers for HcRed1 and dimeric variants of DsRed1 respectively, had the best properties in terms of chromophore maturation rate and final fluorescence intensity. Both tandem proteins demonstrated the same spectral properties as their parent proteins. Expression of tandem HcRed1 with β-actin fusion resulted in labeled patterns indistinguishable from those produced by widely used EGFP-fusion constructs, thus indicating the potency of tandem HcRed1-fusion proteins using in in vivo
labeling of cytoskeleton structures. Fusion constructions of tandem dimeric DsRed1 with connexin43 were correctly transported to the membrane and successfully formed functional connexin channels, but were unable to assemble into a large gap junction.
The other approach to overcome oligomerization is to use pseudo-monomeric form of tetrameric fluorescent proteins. The strategy is based on the co-expression of FP-fused protein of interest with a large excess of free non-fluorescent variant of this FP (FP-helper). This leads to the formation of FP heterooligomers containing only a single target protein linked to tetrameric FP-tag, which therefore can be considered as pseudo-monomers. The feasibility of this method has been demonstrated with red fluorescent mutant of asulCP, M355NA, fused to human cytoplasmic β-actin and asulCP-Ala148Cys as FP-helper [100
]. L929 fibroblasts were transfected with IRES-containing vector for bicystronic expression of the helper asulCP-Ala148Cys and M355NA–actin, or with M355NA–actin under IRES control, but without asulCP-Ala148Cys. As Cap-dependent translation is several times more effective than IRES- dependent one, asulCP-Ala148Cys is produced in higher concentration with regard to M355NA–actin. M355NA–actin expression alone resulted in a high level of cytoplasmic aggregation, in contrast, the templates, obtained with co-expression of M355NA–actin with excess of asulCP-Ala148Cys, were indistinguishable from those produced by EGFP-actin constructs.
Selection of fluorescent proteins (FP) for the multicolor protein labeling. In the best currently available proteins for each of blue, cyan, green, yellow, orange, red and far-red wavelength ranges are summarized. We would like to briefly review here some criteria to choose the right fluorescent proteins for experiment.
Current Mostly Used Fluorescent Proteins
- Higher intrinsic brightness: Recommended brightness is at least 30% of that of common EGFP.
- Monomeric state: even a weak FP dimerization may cause incorrect cellular labeling pattern. The best recommended approach is comparing the pattern with the immunostaining using antibodies to the labeled protein. Less good is comparing the cellular labeling pattern with that of the EGFP labeling. At last run a native SDS-PAGE gel with the purified FP.
- Faster chromophore formation: the shorter time for fluorescence maturation will result in detecting the faster intracellular events. Recommended maturation time is less than 30 minutes for the time to achieve a half-maximum of the FP fluorescence.
- Higher pH-stability: the higher pH stability of FP is the brighter labeling in acidic organelles such as endosomes, lysosomes, etc. Recommended pKa value is less than 6.0 (here, pKa is pH value at which FP looses a half of its brightness).
- Higher photostability: the larger stability of the chromophore in the high-power light irradiation is the longer you are able imaging the cell. The most photostable FP for each respective wavelength range is recommended.
- Larger spectral separation of two FPs to be used simultaneously: with common filters and standard microscopic techniques 50 nm or more difference between both the FP excitation and FP emission peaks is recommended.
Fluorescent timers. Red fluorescent proteins from Anthozoa species (such as DsRed1) are characterized by much slower maturation than Aequorea
GFP. This property could be used in developmental timing studies. When expressed under the same promoter, red fluorescence of DsRed1 appears 18–20 hours later than green fluorescence of Aequorea
GFP. If the activation of expression occurs soon after cell division or differentiation, then color of fluorescence is indicative of the cell differentiation order [60
Some of Anthozoa RFPs, such as zoan2RFP, mcavRFP, rfloRFP [1
] and mutant DsRed-E5 [84
], change their spectral properties during maturation. DsRed-E5 and zoan2RFP can be considered as absolute fluorescent timers, since at early stage of maturation they fluoresce in green spectral region and after complete maturation they exhibit red fluorescence. At the same time, mature mcavRFP and rfloRFP have fluorescence spectra with both green and red peaks.
DsRed-E5 and zoan2RFP can be used to obtain precise information about activation and downregulation of target promoters. Theoretically, the appearance of green fluorescence indicate recent promoter activation, yellow-orange fluorescence means continuous activation and red one shows cessation of promoter activity.
DsRed-E5 was used to trace the activity of the Otx-2
promoter, which is involved in Xenopus laevis
development. A Xenopus laevis
embryo was microinjected with a plasmid containing DsRed-E5 gene under control of the Otx-2
promoter. Expression of Otx-2
was primary observed in the telencephalic and ventral diencephalic brain regions. At the tadpole stage, Otx-2
expression in these regions was almost completely suppressed, but high level of Otx-2
expression was found in the mesencephalic area [101
Studies of diffusion and transport of proteins, organelles and cells. Aequorea
GFP, as well as the majority of Anthozoa FPs, can be irreversibly photobleached being irradiated with the intensive light at their absorption maxima for a prolonged time (about 10–103
c). Presumably because Anthozoa RFPs have more complex chromophore structure and can possess several chromophore forms they show more complex photobleaching kinetic than Aequorea
EGFP does. DsRed1 and its mutants, including monomeric mRFP1, are characterized by two-exponential photobleaching kinetic, as EGFP displays mono-exponential behavior. The spatially restricted chromophore photobleaching can be used to visualize protein and organelle motility within the cell [102
] by methods of fluorescence recovery after photobleaching (FRAP) and fluorescence loss in photobleaching (FLIP).
Photobleaching of Anthozoa RFPs and YFPs, which contain at least two different types of chromophore (green and red/yellow) [36
], can lead to the change of spectral characteristics of these proteins. In the tetramer of fluorescent protein composed of monomers with both types of chromophore, green fluorescence is suppressed by energy transfer from green chromophore to red/yellow chromophore. Selective photobleaching of red/yellow chromophore results in an increased green fluorescence. When DsRed1 was used for cellular and subcellular labeling, resultant green-to-red color change was 1.2–2.0×102
in HEK293 and CHO-K1 cells, and this spectral changes persisted for >30 h [103
]. This type of photoconversion can be applied for selective optical labeling of whole cell, organelles and proteins.
Irradiation of some Anthozoa RFPs with very intense red light can induce the photoconversion of the red chromophore into blue- and far-red spectral species. For instance, irradiation of free or immobilized DsRed1 with light of high intensity at the wavelength 532 nm for 14 h resulted in a decrease of absorption peak at 558 nm and its shift up to 574 nm, besides new absorption peak at 386 nm appeared with 25 % of initial peak at 558 nm magnitude [104
]. At the same time, green chromophore absorption peak at 475 nm changed neither amplitude nor position. When exited at the wavelength 390 nm, fluorescence spectrum of DsRed1 after photoconversion had maximum at 500 nm (due to an energy transfer from blue to green chromophore) and pronounced shoulder at 450 nm (50 % of the maximum amplitude); when exited at the wavelength 570 nm, photoconverted DsRed1 possess fluorescence spectrum peaked at about 595 nm. Keeping in the dark of modified protein for 24 h did not lead to any changes in spectral properties. This type of photoconversion is hardly to be used in cell biology, but it can be applied in biotechnology to label DsRed1-containing immobilized structures by spectrally resolved blue fluorescence [104
KFPs and PAFPs give an opportunity of spatio-temporal labeling and monitoring of living cells, organelles and molecules within the cell. One of the main PAFPs application is protein labeling. By using PAFP-tagged proteins and selectively irradiating the part of the cell, one can induce photochemical chromophore transitions (photoconversion of PAFP from non-fluorescent to fluorescent state or change the color of fluorescence) and observe protein localization, turnover and trafficking by means of confocal microscopy [105
]. For example, PA-GFP–α-tubulin fusion protein has been used to visualize peripheral microtubules motion and their inclusion into a mitotic spindle [106
]. DNA or RNA molecules can be labeled with PAFPs by fusing of DNA/RNA-binding domain fused with PAFP and introducing the corresponding nucleotide sequence into nucleic acid of interest [107
]. In contrast to irreversibly photoconverted PAFPs, KFP1 and Dronpa permit repeated and successive photoactivation of different region of the cell. By using Dronpa–ERK1 fusion protein and repeatedly photoactivating and quenching the fluorescence signal in the nucleus and cytoplasm of a single cell the import and export of ERK1 kinase in the response to the cell stimulation have been studied [108
Cellular organelles can be labeled by PAFPs via fusing of PAFP to polypeptide targeting signals or to proteins that have specific localization. PAFPs have been used to study endocytosis [26
], exocytosis, phagocytosis [109
], the assembly and disassembly of cellular organelles [26
], and to follow the exchange of organelle content [26
KFP1 has been used in mRNA microinjection assays to monitor Xenopus laevis
embryo development [33
]. In vivo
applications for PAFPs also include tracking cells in cancer and metastasis, tracking unicellular organisms, free living and in a host, tracking viruses or protein particles in a host [113
CALI applications. The method CALI (Chromophore Assisted Laser Inactivation) is successfully used for acute inactivation of proteins in living cells [114
]. This method is based on the ability of some fluorescent dye (fluorescein, malachite green, ReAsH and F1AsH) to produce reactive oxygen species (ROS) upon laser irradiation at a wavelength of light absorbed by the dye [117
]. Such dyes are termed photosensitizers. At the cellular level, subjection of photosensitizer-tagged protein to mild illumination for a limited time interval results in precise inactivation of the target while neighboring molecules remain intact. CALI has been successfully applied in functional studies of various proteins [118
]. As applied to the living tissues, high level of ROS production can lead to the target damaging by necrosis or apoptosis [121
]. Some photosensitizers tend to accumulate in tumors, and thus found their use in photodynamic therapy of cancer [124
]. While known photosensitizers are chemical compounds that must be added into living systems exogenously.
Recently it was reported that the mutant developed from Hydrozoa jellyfish chromoprotein anm2CP, KillerRed, is capable of ROS generation upon green light irradiation. KillerRed is the first fully genetically encoded photosensitizer. The mechanism of ROS generation by KillerRed remains unclear but it was shown that amino acid residues Asn145 and Ala161, spatially close to the chromophore, are necessary for the effect, indicating a key role of chromophore surrounding for its capability to generate ROS. Development of KillerRed should give a second breath to the CALI of molecules in living cells. Potentially, KillerRed can be fused to entire collections of open reading frames providing an instrument for high-throughput analysis of protein function in living cells. KillerRed provides much lower 2 % non-specific inactivation, while ReAsH produce up to 21 % nonspecific protein inactivation [125
Another possible field of KillerRed application is CALI of nucleic acids. KillerRed, being fused to an RNA- or DNA-binding domain, can be targeted to a specific RNA or a gene [126
]. If KillerRed induces breaks or damage in adjacent nucleic acid strand then it should be possible to inactivate expression of target genes temporally or even permanently by a pulse of light.
As KillerRed is a fully genetically encoded photosensitizer and don’t require any exogenous compounds it can be used in stable cell lines and transgenic animals. Expression of KillerRed under a specific promoter provides a unique opportunity to investigate cell fate in developing and adult organisms by spatially and temporally controlled cell killing.
Finally, KillerRed opens new perspectives for the photodynamic therapy too. Recent studies on nude mice demonstrated accumulation of GFP-expressed bacteria or virus within various tumors [127
]. Possibly, bacteria or viruses expressing KillerRed can be used not only for visualization but also for light-induced killing of tumor cells.
Protein aggregation studies. A broad range of human diseases known as protein conformational or protein misfolding diseases arises from the failure of a specific peptide or protein to adopt its native functional conformational state. The obvious consequences of misfolding are protein aggregation (and/or fibril formation), loss of function, and gain of toxic function. Some proteins have an intrinsic propensity to assume a pathologic conformation, which becomes evident with aging or at persistently high concentrations. Interactions (or impaired interactions) with some endogenous factors (e.g., chaperones, intracellular or extracellular matrixes, other proteins, small molecules) can change conformation of a pathogenic protein and increase its propensity to misfold. Misfolding can originate from point mutation(s) or result from an exposure to internal or external toxins, impaired posttranslational modifications (phosphorylation, advanced glycation, deamidation, racemization, etc.), an increased probability of degradation, impaired trafficking, lost binding partners or oxidative damage. All these factors can act independently or in association with one another.
Misfolding diseases can affect a single organ or be spread through multiple tissues. The largest group of misfolding diseases, including numerous neurodegenerative disorders and the amyloidoses, originates from the conversion of specific proteins from their soluble functional states into stable, highly ordered, filamentous protein aggregates, known as amyloid fibrils, and from the deposition of these aggregated material in the variety of organs and tissues. In each of these pathological states, a specific protein or protein fragment changes from its natural soluble form into insoluble fibrils, which accumulate in a variety of organs and tissues [128
]. Amyloid-like fibrils display many common properties including a core cross-β-sheet structure in which continuous β-sheets are formed with β-strands running perpendicular to the long axis of the fibrils [135
]. Morphologically, they typically consist of 2–6 unbranched protofilaments 2–5 nm in diameter associated laterally or twisted together to form fibrils with 4–13 nm diameter (e.g., see [136
]). A current set of known proteins involved in protein deposition diseases associated with the formation of extracellular amyloid fibrils or intracellular inclusions with amyloid-like characteristics includes about 70 proteins which are responsible for more than 160 diseases and syndromes [139
]. It has been pointed out that these disorders can be broadly grouped into neurodegenerative conditions, where aggregation occurs in the brain, nonneuropathic localized amyloidoses, where aggregation takes place in a single type of tissue other than the brain, and nonneuropathic systemic amyloidoses, where aggregation affects multiple tissues [140
]. Protein deposition diseases can be sporadic (85%), hereditary (10%) or even transmissible, as in the case of prion diseases (5%) [140
]. All these diseases, being very different clinically, share similar molecular mechanisms where a specific protein or protein fragment changes from its natural soluble form into insoluble fibrils [128
]. Although approximately 70 different proteins are known to be involved in protein deposition diseases, they are mostly unrelated in terms of sequence or structure and prior to fibrillation the amyloidogenic polypeptides causing diseases may be rich in β-sheet, α-helix, β-helix, or be natively unfolded [134
]. Furthermore, the proteinaceous deposits found in patients with any of the protein misfolding diseases beside a major protein component that comprises from a causative protein and forms the core, posses several additional associated species, including metal ions, glycosaminoglycans, the serum amyloid P component, apolipoprotein E, collagen, and many others [144
For many years it has been generally assumed that the ability to form amyloid fibrils is limited to a relatively small number of proteins, essentially those found in the diseases, and that these proteins posses specific sequence motifs encoding the unique structure of the amyloid core. However, recent studies have established that many diseases unrelated proteins were shown to form fibrils [129
]. It is even believed that virtually any protein can be forced to fibrillate if the appropriate conditions are found [129
]. The structural diversity of amyloidogenic proteins and close similarity of the resultant fibrils imply that considerable structural rearrangements have to occur in order for fibril formation to happen. A general hypothesis of fibrillogenesis states: structural transformation of a polypeptide chain into a partially folded conformation represents an important prerequisite for protein fibrillation [134
]. These aggregation-prone intermediates would be structurally different for different proteins. Furthermore, intermediate might contain different amount of ordered structure even for the same protein undergoing different aggregation processes. It is believed that the precursor of soluble aggregates is the most structured, whereas amyloid fibrils are formed from the least ordered conformation (cf. [147
]). It has been also pointed out that the variations in the amount of the ordered structure in the amyloidogenic precursor might be responsible for the formation of fibrils with distinct morphologies [148
The formation of amyloid fibrils does not represent the only pathological hallmark of conformational or protein deposition diseases. In several neurodegenerative disorders (as well as in numerous in vitro experiments) the protein depositions are composed of the amorphous aggregates, cloud-like inclusions without defined structure. Similarly, soluble oligomers represent another alternative final product of the aggregation process. The choice between three aggregation pathways, fibrillation, amorphous aggregate formation or oligomerization, is determined by the amino acid sequence (which could be modified by mutation) and by the peculiarities of the protein environment.
Obviously, the progress in understanding the pathology of protein misfolding diseases and in rational design of drugs to inhibit or reverse protein aggregation depends on our ability to study the details of the misfolding process, to follow the aggregation process, and to see and analyze the structure of the aggregated particles. GFP-like proteins were shown to represent a set of unique tools that allow visualization and analysis of aggregated structures and aggregation process both in vitro and in vivo.
When production of misfolded proteins exceeds the cellular capacity to degrade them, the proteins are deposited in large aggregates surrounding the microtubule (MT)-organizing center (MTOC) and ensheathed in a cage of vimentin. This subcellular structure was termed the aggresome and it was proposed that the formation of an aggresome is a general cellular response to the presence of aggregated, nondegraded proteins [149
]. The process of aggresome formation by a GFP-250 chimera composed of the GFP fused to a 250–amino acid fragment of the cytosolic protein p115 was investigated in great details [150
]. p115 is a protein involved in the membrane transport. It is peripherally associated with membranes, and has been localized to the Golgi [151
] and to the transport intermediates carrying cargo from the ER to the Golgi [153
]. Time-lapse image analysis in living cells was used to characterize the dynamics of aggresome formation. This analysis revealed that small aggregates were formed peripherally and traveled on microtubules in a minus-end direction to MTOC region where they remained as distinct but closely apposed particulate structures. Furthermore, aggresome formation interfered with correct Golgi localization and disrupted the normal astral distribution of microtubules [153
Novel GFP-based technique called ‘split GFP complementation assay’ has been introduced to detect the aggregated state of proteins in vitro
or in prokaryotes [154
]. This method is based on separating the GFP into two soluble and spontaneously associating fragments, that when mixed together spontaneously complement, resulting in GFP folding and formation of the fluorophore. Here, a protein of interest is fused to a small GFP fragment via a flexible linker. The complementary GFP fragment is expressed separately. Neither fragment alone is fluorescent. When mixed, the small and large GFP fragments spontaneously associate, resulting in GFP folding and formation of the fluorophore [154
]. The largest challenge in this endeavor was a poor foldability of the GFP fragments. To overcome this problem, several pairs of fragments were tested from either the folding reporter GFP, which contains the mutations F99S, M153T, V163A [77
], F64L and S65 Thttp://www.nature.com/nbt/journal/v23/n1/full/nbt1044.html - B16#B16
], or the exceptionally stable 'superfolder' GFP, containing the folding reporter GFP mutations and S30R, Y39N, N105T, Y145F, I171V and A206V [154
]. Although coexpression of the superfolder GFP fragments containing amino acids 1–214 (GFP 1–10) and 214–230 (GFP 11) gave fluorescent Escherichia coli
colonies, the complication with this pair was insolubility of superfolder GFP 1–10. Next, superfolder GFP 1–10 was evolved by DNA shuffling [156
] to improve its solubility and increase its complementation. This procedure gave a variant, termed GFP 1−10 OPT, which was moderately soluble and in addition to the folding reporter GFP mutations, contained S30R, Y145F, I171V and A206V substitutions from superfolder GFP and seven new mutations: N39I, T105K, E111V, I128T, K166T, I167V and S205T [154
]. In the original study, split GFP complementation provided a robust quantitative monitoring of the aggregation process of test proteins in vitro
and in Escherichia coli
]. Recently, this technique was successfully applied to quantitatively measure tau protein aggregation in situ
allowing determination of the key mediators of tau aggregation and early aggregation processes in living mammalian cells [157
Among the various neurodegenerative diseases, the poly(Q) diseases, including Huntington disease and various types of spinocerebellar ataxia, are of special interest. Poly(Q) diseases (also known as the CAG repeat diseases) are a group of at least nine inherited neurodegenerative disorders caused by abnormal expansions of the poly(Q) stretch within disease-causing proteins. These expansions of the poly(Q) stretch to above 40 glutamines trigger the disease-causing proteins to aggregate into insoluble β-sheet-rich amyloid fibrils [158
]. To provide a model system for investigating common pathogenic features, the behavior of poly(Q) expansions fused to GFP and expressed in Caenorhabditis elegans
has been examined [160
]. In this model, the effect of polyglutamine expansions in C. elegans
was examined by expressing GFP fusion proteins with 19 or 82 glutamine residues (Q19-GFP or Q82-GFP) in body wall muscle cells. Although Q19-GFP was distributed evenly throughout the body wall muscle cells, Q82-GFP formed discrete intracellular aggregates. These aggregates appeared early in embryogenesis, increased in number and size during development from the larval to the adult stage and correlated with a delay in larval to adult development. It has been also shown that the toxic effect of poly(Q) expression and the formation of aggregates can be reversed by coexpression of the yeast chaperone Hsp104 [160
Recently, fluorescence correlation spectroscopy (FCS) based on GFP-like protein fused to a target protein was employed to explore oligomerization of the poly(Q) proteins in cells [161
]. A time-dependent increase in the diffusion time and particle size of expanded poly(Q)-GFP fusion proteins expressed in cultured cells was detected by FCS, indicating oligomer formation. Intriguingly, the poly(Q)-binding peptide QBP1 was shown to suppress poly(Q)-GFP oligomer formation [161
]. Based on these observations it has been concluded that FCS of GFP-fused proteins is a useful technique to monitor the oligomerization of disease-causing proteins in cells as well as its inhibition in the conformational diseases.
Large-scale search for compounds that inhibit protein aggregation and which therefore potentially could be used as therapeutic agents for the prevention or treatment of protein misfolding diseases poses an enormous challenge. A novel high-throughput GFP-based screen capable of isolating inhibitors of Aβ aggregation from large libraries of inactive candidates was recently developed. This technology used a fusion of Aβ42
to GFP and was based on the observation that in the absence of aggregation inhibition, the rapid misfolding and aggregation of Aβ42
caused the entire fusion protein to misfold, thereby preventing green fluorescence. Compounds that inhibited Aβ42
aggregation enable GFP to fold into its native structure and were identified by the increased fluorescent signal. This GFP-based method is rapid and inexpensive and can be used to screen large combinatorial libraries for inhibitors of protein misfolding and aggregation [162