In vivo and in vitro protein solubility assays using split GFP

The rapid assessment of protein solubility is essential for evaluating expressed proteins and protein variants for use as reagents for downstream studies. Solubility screens based on antibody blots are complex and have limited screening capacity. Protein solubility screens using split beta-galactosidase in vivo and in vitro can perturb protein folding. Split GFP used for monitoring protein interactions folds poorly, and to overcome this limitation, we recently developed a protein-tagging system based on self-complementing split GFP derived from an exceptionally well folded variant of GFP termed 'superfolder GFP'. Here we present the step-by-step procedure of the solubility assay using split GFP. A 15-amino-acid GFP fragment, GFP 11, is fused to a test protein. The GFP 1-10 detector fragment is expressed separately. These fragments associate spontaneously to form fluorescent GFP. The fragments are soluble, and the GFP 11 tag has minimal effect on protein solubility and folding. We describe high-throughput protein solubility screens amenable both for in vivo and in vitro formats. The split-GFP system is composed of two vectors used in the same strain: pTET GFP 11 and pET GFP 1-10 (Fig. 1 and Supplementary Note online). The gene encoding the protein of interest is cloned into the pTET GFP 11 vector (resulting in an N-terminal fusion) and transformed into Escherichia coli BL21 (DE3) cells containing the pET GFP 1-10 plasmid. We also describe how this system can be used for selecting soluble proteins from a library of variants (Box 1). The large screening power of the in vivo assay combined with the high accuracy of the in vitro assay point to the efficiency of this two-step split-GFP tool for identifying soluble clones suitable for purification and downstream applications.     

Green fluorescent protein as a reporter for macromolecular localization in bacterial cells

Green fluorescent protein (GFP) is a highly useful fluorescent tag for studying the localization, structure, and dynamics of macromolecules in living cells, and has quickly become a primary tool for analysis of DNA and protein localization in prokaryotes. Several properties of GFP make it an attractive and versatile reporter. It is fluorescent and soluble in a wide variety of species, can be monitored noninvasively by external illumination, and needs no external substrates. Localization of GFP fusion proteins can be analyzed in live bacteria, therefore eliminating potential fixation artifacts and enabling real-time monitoring of dynamics in situ. Such real-time studies have been facilitated by brighter, more soluble GFP variants. In addition, red-shifted GFPs that can be excited by blue light have lessened the problem of UV-induced toxicity and photobleaching. The self-contained domain structure of GFP reduces the chance of major perturbations to GFP fluorescence by fused proteins and, conversely, to the activities of the proteins to which it is fused. As a result, many proteins fused to GFP retain their activities. The stability of GFP also allows detection of its fluorescence in vitro during protein purification and in cells fixed for indirect immunofluorescence and other staining protocols. Finally, the different properties of GFP variants have given rise to several technological innovations in the study of cellular physiology that should prove useful for studies in live bacteria. These include fluorescence resonance energy transfer (FRET) for studying protein-protein interactions and specially engineered GFP constructs for direct determination of cellular ion fluxes.     

Single-strand DNA aptamers as probes for protein localization in cells.

The accurate localization of proteins in fixed cells is important for many studies in cell biology, but good fixation is often antagonistic to good immunolabeling, given the density of well-preserved cells and the size of most labeled antibody probes. We therefore explored the use of single-stranded oligonucleotides (aptamers), which can bind to proteins with very high affinity and specificity but which are only approximately 10 kD. To evaluate these probes for general protein localization, we sought an aptamer that binds to a widely used protein tag, the green fluorescent protein (GFP). Although this quest was not successful, we were able to solve several practical problems that will confront any such labeling effort, e.g., the rates at which oligonucleotides enter fixed cells of different kinds and the extent of nonspecific oligonucleotide binding to both mammalian and yeast cell structures. Because such localization methods would be of particular value for electron microscopy of optimally fixed material, we also explored the solubility of aptamers under conditions suitable for freeze-substitution fixation. We found that aptamers are sufficiently soluble in cold organic solvents to encourage the view that this approach may be useful for the localization of specific proteins in context of cellular fine structure.     

Fluorescent labeling of antibody fragments using split GFP

Antibody fragments are easily isolated from in vitro selection systems, such as phage and yeast display. Lacking the Fc portion of the antibody, they are usually labeled using small peptide tags recognized by antibodies. In this paper we present an efficient method to fluorescently label single chain Fvs (scFvs) using the split green fluorescent protein (GFP) system. A 13 amino acid tag, derived from the last beta strand of GFP (termed GFP11), is fused to the C terminus of the scFv. This tag has been engineered to be non-perturbing, and we were able to show that it exerted no effect on scFv expression or functionality when compared to a scFv without the GFP11 tag. Effective functional fluorescent labeling is demonstrated in a number of different assays, including fluorescence linked immunosorbant assays, flow cytometry and yeast display. Furthermore, we were able to show that this split GFP system can be used to determine the concentration of scFv in crude samples, as well an estimate of antibody affinity, without the need for antibody purification. We anticipate this system will be of widespread interest in antibody engineering and in vitro display systems.     

Enhanced protein expression in the baculovirus/insect cell system using engineered SUMO fusions

Recombinant protein expression in insect cells varies greatly from protein to protein. A fusion tag that is not only a tool for detection and purification, but also enhances expression and/or solubility would greatly facilitate both structure/function studies and therapeutic protein production. We have shown that fusion of SUMO (small ubiquitin-related modifier) to several test proteins leads to enhanced expression levels in Escherichia coli. In eukaryotic expression systems, however, the SUMO tag could be cleaved by endogenous desumoylase. In order to adapt SUMO-fusion technology to these systems, we have developed an alternative SUMO-derived tag, designated SUMOstar, which is not processed by native SUMO proteases. In the present study, we tested the SUMOstar tag in a baculovirus/insect cell system with several proteins, i.e. mouse UBP43, human tryptase beta II, USP4, USP15, and GFP. Our results demonstrate that fusion to SUMOstar enhanced protein expression levels at least 4-fold compared to either the native or His(6)-tagged proteins. We isolated active SUMOstar tagged UBP43, USP4, USP15, and GFP. Tryptase was active following cleavage with a SUMOstar specific protease. The SUMOstar system will make significant impact in difficult-to-express proteins and especially to those proteins that require the native N-terminal residue for function.     

Assessing the binding and endocytosis activity of cellular receptors using GFP-ligand fusions

We have developed a simple scheme for characterizing ligand-receptor binding and post-binding activity on living cells. Our approach makes use of green fluorescent protein (GFP) as an auto-fluorescent tag to label protein ligands. We have constructed GFP-tagged ligands that can be expressed in bacteria as soluble fusion proteins. A cell-binding assay using fluorescence-activated cell sorting (FACS) demonstrates that GFP-tagged proteins retain their wild-type receptor-binding specificity. Using this assay, we measure ligand binding on unfixed cells and demonstrate receptor specificity using specific competitors. To determine the ability of receptor targets to internalize, we developed a second FACS-based assay to detect the rate and percentage of internalized ligand in living cells. Noninternalizing control ligands and fluorescence microscopy of treated cells confirm that our assay is reliable for determining receptor internalization activity.     

From fixed to FRAP: measuring protein mobility and activity in living cells

Experiments with fluorescence recovery after photobleaching (FRAP) started 30 years ago to visualize the lateral mobility and dynamics of fluorescent proteins in living cells. Its popularity increased when non-invasive fluorescent tagging became possible with the green fluorescent protein (GFP). Many researchers use GFP to study the localization of fusion proteins in fixed or living cells, but the same fluorescent proteins can also be used to study protein mobility in living cells. Here we review the potential of FRAP to study protein dynamics and activity within a single living cell. These measurements can be made with most standard confocal laser-scanning microscopes equipped with photobleaching protocols.     

Yield, solubility and conformational quality of soluble proteins are not simultaneously favored in recombinant Escherichia coli

Many enzymes or fluorescent proteins produced in Escherichia coli are enzymatically active or fluorescent respectively when deposited as inclusion bodies. The occurrence of insoluble but functional protein species with native-like secondary structure indicates that solubility and conformational quality of recombinant proteins are not coincident parameters, and suggests that both properties can be engineered independently. We have here proven this principle by producing elevated yields of a highly fluorescent but insoluble green fluorescent protein (GFP) in a DnaK- background, and further enhancing its solubility through adjusting the growth temperature and GFP gene expression rate. The success of such a two-step approach confirms the independent control of solubility and conformational quality, advocates for new routes towards high quality protein production and intriguingly, proves that high protein yields dramatically compromise the conformational quality of soluble versions.     

Use of Green Fluorescent Protein in mouse embryos

Green Fluorescent Protein (GFP) has rapidly been established as a versatile and powerful cell marker in many organisms. Initial problems in using it in mammalian cells were solved by introducing mutations to increase its solubility at higher temperatures, such that GFP has now been used as a reporter in both gene expression and cell lineage studies, and to localize proteins within mammalian cells. GFP has two unique advantages: (i) the protein becomes fluorescent in an autocatalytic reaction, so that it can be introduced into any cell type simply as a cDNA or mRNA, or as protein; (ii) it is "bright" enough to be visualized in living cells under conditions that do not cause photodamage to the cells. In this article we outline the ways in which we have used GFP mRNA and cDNA in our studies of mouse cell lineages, and to characterize the behavior of proteins within the embryos.     

Identifying protein domains by global analysis of soluble fragment data

The production and analysis of individual structural domains is a common strategy for studying large or complex proteins, which may be experimentally intractable in their full-length form. However, identifying domain boundaries is challenging if there is little structural information concerning the protein target. One experimental procedure for mapping domains is to screen a library of random protein fragments for solubility, since truncation of a domain will typically expose hydrophobic groups, leading to poor fragment solubility. We have coupled fragment solubility screening with global data analysis to develop an effective method for identifying structural domains within a protein. A gene fragment library is generated using mechanical shearing, or by uracil doping of the gene and a uracil-specific enzymatic digest. A split green fluorescent protein (GFP) assay is used to screen the corresponding protein fragments for solubility when expressed in Escherichia coli. The soluble fragment data are then analyzed using two complementary approaches. Fragmentation “hotspots” indicate possible interdomain regions. Clustering algorithms are used to group related fragments, and concomitantly predict domain location. The effectiveness of this Domain Seeking procedure is demonstrated by application to the well-characterized human protein p85α.     

Quantitative Analysis of α-Synuclein Solubility in Living Cells Using Split GFP Complementation

Presently incurable, Parkinson's disease (PD) is the most common neurodegenerative movement disorder and affects 1% of the population over 60 years of age. The hallmarks of PD pathogenesis are the loss of dopaminergic neurons in the substantia nigra pars compacta, and the occurrence of proteinaceous cytoplasmic inclusions (Lewy bodies) in surviving neurons. Lewy bodies are mainly composed of the pre-synaptic protein alpha-synuclein (αsyn), an intrinsically unstructured, misfolding-prone protein with high propensity to aggregate. Quantifying the pool of soluble αsyn and monitoring αsyn aggregation in living cells is fundamental to study the molecular mechanisms of αsyn-induced cytotoxicity and develop therapeutic strategies to prevent αsyn aggregation. In this study, we report the use of a split GFP complementation assay to quantify αsyn solubility. Particularly, we investigated a series of naturally occurring and rationally designed αsyn variants and showed that this method can be used to study how αsyn sequence specificity affects its solubility. Furthermore, we demonstrated the utility of this assay to explore the influence of the cellular folding network on αsyn solubility. The results presented underscore the utility of the split GFP assay to quantify αsyn solubility in living cells.     

Split‐Cre recombinase effectively monitors protein–protein interactions in living bacteria

The ability of Cre recombinase to excise genetic material has been used extensively for genome engineering in prokaryotic and eukaryotic cells. Recently, split‐Cre fragments have been described that advance control of recombinase activity in mammalian cells. However, whether these fragments can be utilized for monitoring protein‐protein interactions has not been reported. In this work, we developed a protein‐fragment complementation assay (PCA) based on split‐Cre for monitoring and engineering pairwise protein interactions in living Escherichia coli cells. This required creation of a dual‐fluorescent reporter plasmid that permits visualization of reconstituted Cre recombinase activity by switching from red to green in the presence of an interacting protein pair. The resulting split‐Cre PCA faithfully links cell fluorescence with differences in binding affinity, thereby allowing the facile isolation of high‐affinity binders based on phenotype. Given the resolution of its activity and sensitivity to interactions, our system may prove a viable option for poorly expressed or weakly interacting protein pairs that evade detection in other PCA formats. Based on these findings, we anticipate that our split‐Cre PCA will become a highly complementary and useful new addition to the protein‐protein interaction toolbox.     

Determining Membrane Protein Topology Using Fluorescence Protease Protection (FPP)

The correct topology and orientation of integral membrane proteins are essential for their proper function, yet such information has not been established for many membrane proteins. A simple technique called fluorescence protease protection (FPP) is presented, which permits the determination of membrane protein topology in living cells. This technique has numerous advantages over other methods for determining protein topology, in that it does not require the availability of multiple antibodies against various domains of the membrane protein, does not require large amounts of protein, and can be performed on living cells. The FPP method employs the spatially confined actions of proteases on the degradation of green fluorescent protein (GFP) tagged membrane proteins to determine their membrane topology and orientation. This simple approach is applicable to a wide variety of cell types, and can be used to determine membrane protein orientation in various subcellular organelles such as the mitochondria, Golgi, endoplasmic reticulum and components of the endosomal/recycling system. Membrane proteins, tagged on either the N-termini or C-termini with a GFP fusion, are expressed in a cell of interest, which is subject to selective permeabilization using the detergent digitonin. Digitonin has the ability to permeabilize the plasma membrane, while leaving intracellular organelles intact. GFP moieties exposed to the cytosol can be selectively degraded through the application of protease, whereas GFP moieties present in the lumen of organelles are protected from the protease and remain intact. The FPP assay is straightforward, and results can be obtained rapidly.     

Expression of green or red fluorescent protein (GFP or DsRed) linked proteins in nonmuscle and muscle cells

The introduction of the green fluorescent protein (GFP) plasmids that allow proteins and peptides to be expressed with a fluorescent tag has had a major impact on the field of cell biology. It has enabled the dynamics of a wide variety of proteins to be analyzed that could not otherwise be detected in live cells. Transient transfections of muscle and nonmuscle cells with plasmids encoding various cytoskeletal proteins ligated to green fluorescent protein or Ds red protein allow changes in the cytoskeletal network to be studied in the same cell for time periods up to several days. With this approach, proteins that could not be purified and directly labeled with fluorescent dyes and microinjected into cells can now be expressed and visualized in a wide variety of cells. Procedures are presented for transfection of the nonmuscle cell, PtK2, and primary cultures of embryonic chick myocytes, and for studying the live transfected cells.     

New molecular reporters for rapid protein folding assays

The GFP folding reporter assay uses a C-terminal GFP fusion to report on the folding success of upstream fused polypeptides. The GFP folding assay is widely-used for screening protein variants with improved folding and solubility, but truncation artifacts may arise during evolution, i.e. from de novo internal ribosome entry sites. One way to reduce such artifacts would be to insert target genes within the scaffolding of GFP circular permuted variants. Circular permutants of fluorescent proteins often misfold and are non-fluorescent, and do not readily tolerate fused polypeptides within the fluorescent protein scaffolding. To overcome these limitations, and to increase the dynamic range for reporting on protein misfolding, we have created eight GFP insertion reporters with different sensitivities to protein misfolding using chimeras of two previously described GFP variants, the GFP folding reporter and the robustly-folding "superfolder" GFP. We applied this technology to engineer soluble variants of Rv0113, a protein from Mycobacterium tuberculosis initially expressed as inclusion bodies in Escherichia coli. Using GFP insertion reporters with increasing stringency for each cycle of mutagenesis and selection led to a variant that produced large amounts of soluble protein at 37 degrees C in Escherichia coli. The new reporter constructs discriminate against truncation artifacts previously isolated during directed evolution of Rv0113 using the original C-terminal GFP folding reporter. Using GFP insertion reporters with variable stringency should prove useful for engineering protein variants with improved folding and solubility, while reducing the number of artifacts arising from internal cryptic ribosome initiation sites.     

Green fluorescent protein as a novel tool to measure apoptosis and necrosis

BACKGROUND: Several apoptosis-detecting methods are currently available. Many of them are work intensive and require the additional use of antibodies, dyes, specific substrates, or enzymatic reactions. A simple, fast, and reliable method was developed to test for apoptosis or necrosis using mouse and human cell lines (e.g., Jurkat, A20.2J, and PB3c cells) stably transfected with a vector coding for green fluorescent protein (GFP) as indicator cells. METHODS: Apoptosis in GFP-transfected cell lines was induced either by soluble Fas-Ligand (sFasL), recombinant human TRAIL (rhTRAIL), or interleukin-3 (IL-3) deprivation. Necrosis was induced by polyclonal anti-A20 and complement treatment of GFP-transfected A20. Cells were analyzed by flow cytometry for GFP fluorescence. Propidium iodide and Annexin V staining were used to confirm the results obtained with the GFP-method. RESULTS: Live GFP-transfected cells show a strong fluorescence intensity, which is significantly diminished upon induction of apoptosis, whereas necrotic GFP-transfected cells almost completely lose their GFP-associated fluorescence. Apoptosis but not necrosis of GFP-transfected cells was blocked by the use of a caspase inhibitor. The results are highly comparable to conventional apoptosis-detecting methods. CONCLUSIONS: The advantage of our GFP-based assay compared with other methods is the analysis of apoptosis or necrosis without the necessity for additional staining or washing steps, making it an ideal tool for screening apoptotic or necrotic stimuli.     

基于GFP的FRET应用

绿色荧光蛋白（GFP）是一种活性荧光标记,已被用来研究基因表达、分子定位,蛋白质折叠和转运;荧光共振能量转移（FRET）是一种无损伤的光学检测方法,能检测到小于纳米的距离变化。将GFP的活性定位标记功能与FRET的高分辨率相结合。为活体研究生物分子的功能和命运开创了新的篇章。作者在介绍GFP和FRET原理的基础上,综述了基于GFP的FRET在蛋白酶活性,蛋白质间相互作用 构象改变研究中的应用。     

Simultaneous visualization of two protein complexes in a single plant cell using multicolor fluorescence complementation analysis

Bimolecular fluorescence complementation (BiFC) is an approach used to analyze protein–protein interaction in vivo, in which non-fluorescent N-terminal and C-terminal fragments of a fluorescent protein are reconstituted to emit fluorescence only when they are brought together by interaction of two proteins to fuse both fragments. A method for simultaneous visualization of two protein complexes by multicolor BiFC with fragments from green fluorescent protein (GFP) and its variants such as cyan and yellow fluorescent proteins (CFP and YFP) was recently reported in animal cells. In this paper we describe a new strategy for simultaneous visualization of two protein complexes in plant cells using the multicolor BiFC with fragments from CFP, GFP, YFP and a red fluorescent protein variant (DsRed-Monomer). We identified nine different BiFC complexes using fragments of CFP, GFP and YFP, and one BiFC complex using fragments of DsRed-Monomer. Fluorescence complementation did not occur by combinations between fragments of GFP variants and DsRed-Monomer. Based on these findings, we achieved simultaneous visualization of two protein complexes in a single plant cell using two colored fluorescent complementation pairs (cyan/red, green/red or yellow/red).