Proteolytic activity assayed by subcellular localization switching of a substrate

An approach to assay proteolytic activity in vivo by altering the subcellular localization of a labelled substrate was demonstrated. The assay included a protein shuttling between different cellular compartments and a site-specific recombinant protease. The shuttle protein used was the human immunodeficiency virus type 1 (HIV-1) Rev protein tandemly fused to the enhanced green fluorescent protein (EGFP) and the red fluorescent protein (RFP), while the protease was the site-specific protease VP24 from the herpes simplex virus type 1 (HSV-1). The fluorescent proteins in the Rev fusion protein were separated by a cleavage site specific for the VP24 protease. When co-expressed in COS-7 cells proteolysis was observed by fluorescence microscopy as a shift from a predominantly cytoplasmic localization of the fusion protein RevEGFP to a nuclear localization while the RFP part of the fusion protein remained in the cytoplasm. The cleavage of the fusion protein by VP24 was confirmed by Western blot analysis. The activity of VP24, when tagged N-terminally by the Myc-epitope, was found to be comparable to VP24. These results demonstrates that the activity and localization of a recombinantly expressed protease can be assessed by protease-mediated cleavage of fusion proteins containing a specific protease cleavage site.     

Visualizing and quantifying the differential cleavages of the eukaryotic translation initiation factors eIF4GI and eIF4GII in the enterovirus‐infected cell

Enterovirus (EV) infection has been shown to cause a marked shutoff of host protein synthesis, an event mainly achieved through the cleavages of eukaryotic translation initiation factors eIF4GI and eIF4GII that are mediated by viral 2A protease (2A^pro). Using fluorescence resonance energy transfer (FRET), we developed genetically encoded and FRET‐based biosensors to visualize and quantify the specific proteolytic process in intact cells. This was accomplished by stable expression of a fusion substrate construct composed of the green fluorescent protein 2 (GFP²) and red fluorescent protein 2 (DsRed2), with a cleavage motif on eIF4GI or eIF4GII connected in between. The FRET biosensor showed a real‐time and quantifiable impairment of FRET upon EV infection. Levels of the reduced FRET closely correlated with the cleavage kinetics of the endogenous eIF4Gs isoforms. The FRET impairments were solely attributed to 2A^pro catalytic activity, irrespective of other viral‐encoded protease, the activated caspases or general inhibition of protein synthesis in the EV‐infected cells. The FRET biosensors appeared to be a universal platform for several related EVs. The spatiotemporal and quantitative imaging enabled by FRET can shed light on the protease–substrate behaviors in their normal milieu, permitting investigation into the molecular mechanism underlying virus‐induced host translation inhibition. Biotechnol. Bioeng. 2009; 104: 1142–1152. © 2009 Wiley Periodicals, Inc.     

A Cell-based Fluorescence Resonance Energy Transfer (FRET) Sensor Reveals Inter- and Intragenogroup Variations in Norovirus Protease Activity and Polyprotein Cleavage

The viral protease represents a key drug target for the development of antiviral therapeutics. Because many protease inhibitors mimic protease substrates, differences in substrate recognition between proteases may affect their sensitivity to a given inhibitor. Here we use a cell-based FRET sensor to investigate the activity of different norovirus proteases upon cleavage of various norovirus cleavage sites inserted into a linker region separating cyan fluorescent protein and yellow fluorescent protein. Using this system, we demonstrate that differences in substrate processing exist between proteases from human noroviruses (genogroups I (GI) and II) and the commonly used murine norovirus (MNV, genogroup V) model. These altered the cleavage efficiency of specific cleavage sites both within and between genogroups. The differences observed between these proteases may affect sensitivity to protease inhibitors and the suitability of MNV as a model system for testing such molecules against the human norovirus protease. Finally, we demonstrate that replacement of MNV polyprotein cleavage sites with the GI or GII equivalents, with the exception of the NS6–7 junction, leads to the production of infectious virus when the MNV NS6 protease, but not the GI or GII proteases, are present.     

An easy-to-use FRET protein substrate to detect calpain cleavage in vitro and in vivo

Calpain-1 and -2 are Ca^2 +-activated intracellular cysteine proteases that regulate a wide range of cellular functions through the cleavage of their protein substrates. Unlike degradative proteases, calpains make limited, transformative cleavages, typically in accessible sequences linking discrete subdomains, to irreversibly alter substrate functions. The biological roles of calpain and their interplay with calcium signaling are of significant biomedical interest as biomarkers and potential therapeutic targets in a growing number of diseases including Alzheimer's, cancer and fibrosis. Unfortunately, many of the colorimetric and fluorimetric assays that have been developed to study calpain activity suffer from low sensitivity and/or poor calpain specificity. To address the need for a highly sensitive and calpain-specific substrate suitable for in vitro and in vivo calpain activity analysis, we have developed a protein FRET probe. We inserted the optimized calpain cleavage sequence PLFAAR between cyan fluorescent protein (CFP) and yellow fluorescent protein (YFP) and modulated its flanking sequences for optimal calpain cleavage. We demonstrate greater sensitivity and calpain-specificity of an optimal 16-residue PLFAAR-based FRET substrate compared to a standard α-spectrin-based probe. The 16-residue PLFAAR protein FRET substrate is not significantly cleaved by trypsin, chymotrypsin, cathepsin-L or caspase-3, and is highly sensitive to both calpain-1 and -2. After transfection of the substrate gene into breast cancer cells the PLFAAR protein FRET product was cut in lysed wild-type cells but not in those with a calpain knock-out phenotype. Blockage of substrate cleavage in the lysates by endogenous and exogenous calpastatin was observed, and was overcome by adding extra calpain.     

Imaging FRET between spectrally similar GFP molecules in single cells

Fluorescence resonance energy transfer (FRET) detection in fusion constructs consisting of green fluorescent protein (GFP) variants linked by a sequence that changes conformation upon modification by enzymes or binding of ligands has enabled detection of physiological processes such as Ca(2+) ion release, and protease and kinase activity. Current FRET microscopy techniques are limited to the use of spectrally distinct GFPs such as blue or cyan donors in combination with green or yellow acceptors. The blue or cyan GFPs have the disadvantages of less brightness and of autofluorescence. Here a FRET imaging method is presented that circumvents the need for spectral separation of the GFPs by determination of the fluorescence lifetime of the combined donor/acceptor emission by fluorescence lifetime imaging microscopy (FLIM). This technique gives a sensitive, reproducible, and intrinsically calibrated FRET measurement that can be used with the spectrally similar and bright yellow and green fluorescent proteins (EYFP/EGFP), a pair previously unusable for FRET applications. We demonstrate the benefits of this approach in the analysis of single-cell signaling by monitoring caspase activity in individual cells during apoptosis.     

Sequential bioluminescence resonance energy transfer-fluorescence resonance energy transfer-based ratiometric protease assays with fusion proteins of firefly luciferase and red fluorescent protein

We report here the preparation of ratiometric luminescent probes that contain two well-separated emission peaks produced by a sequential bioluminescence resonance energy transfer (BRET)–fluorescence resonance energy transfer (FRET) process. The probes are single soluble fusion proteins consisting of a thermostable firefly luciferase variant that catalyze yellow-green (560 nm maximum) bioluminescence and a red fluorescent protein covalently labeled with a near-infrared fluorescent dye. The two proteins are connected by a decapeptide containing a protease recognition site specific for factor Xa, thrombin, or caspase 3. The rates of protease cleavage of the fusion protein substrates were monitored by recording emission spectra and plotting the change in peak ratios over time. Detection limits of 0.41 nM for caspase 3, 1.0 nM for thrombin, and 58 nM for factor Xa were realized with a scanning fluorometer. Our results demonstrate for the first time that an efficient sequential BRET–FRET energy transfer process based on firefly luciferase bioluminescence can be employed to assay physiologically important protease activities.     

Visualization of human immunodeficiency virus protease inhibition using a novel Förster resonance energy transfer molecular probe

The in vivo high‐throughput screening (HTS) of human immunodeficiency virus (HIV) protease inhibitors is a significant challenge because of the lack of reliable assays that allow the visualization of HIV targets within living cells. In this study, we developed a new molecular probe that utilizes the principles of Förster resonance energy transfer (FRET) to visualize HIV‐1 protease inhibition within living cells. The probe is constructed by linking two fluorescent proteins: AcGFP1 (a mutant green fluorescent protein) and mCherry (a red fluorescent protein) with an HIV‐1 protease cleavable p2/p7 peptide. The cleavage of the linker peptide by HIV‐1 protease leads to separation of AcGFP1 from mCherry, quenching FRET between AcGFP1 and mCherry. Conversely, the addition of a protease inhibitor prevents the cleavage of the linker peptide by the protease, allowing FRET from AcGFP1 to mCherry. Thus, HIV‐1 protease inhibition can be determined by measuring the FRET signal's change generated from the probe. Both in vitro and in vivo studies demonstrated the feasibility of applying the probe for quantitative analyses of HIV‐1 protease inhibition. By cotransfecting HIV‐1 protease and the probe expression plasmids into 293T cells, we showed that the inhibition of HIV‐1 protease by inhibitors can be visualized or quantitatively determined within living cells through ratiometric FRET microscopy imaging measurement. It is expected that this new probe will allow high‐content screening (HCS) of new anti‐HIV drugs. © 2011 American Institute of Chemical Engineers Biotechnol. Prog., 2011     

Function-based isolation of novel enzymes from a large library

Here we describe a high-throughput, quantitative method for the isolation of enzymes with novel substrate specificities from large libraries of protein variants. Protein variants are displayed on the surface of microorganisms and incubated with a synthetic substrate consisting of (1) a fluorescent dye (2) a positively charged moiety (3) the target scissile bond, and (4) a fluorescence resonance energy transfer (FRET) quenching partner. Enzymatic cleavage of the scissile bond results in release of the FRET quenching partner while the fluorescent product is retained on the cell surface, allowing isolation of catalytically active clones by fluorescence-activated cell sorting (FACS). Using a synthetic substrate with these characteristics, we enriched Escherichia coli expressing the serine protease OmpT from cells expressing an inactive OmpT variant by over 5,000-fold in a single round. Screening a library of 6 x 10(5) random OmpT variants by FACS using a FRET peptide substrate with a nonpreferred Arg-Val cleavage sequence resulted in the isolation of variant proteases with catalytic activities enhanced by as much as 60-fold. This approach represents a potentially widely applicable method for high-throughput screening of large libraries on the basis of catalytic turnover.     

The eIF4G-eIF4E complex is the target for direct cleavage by the rhinovirus 2A proteinase.

The 2A proteinases (2Apro) of certain picornaviruses induce the cleavage of the eIF4G subunit of the cap-binding protein complex, eIF4F. Several reports have demonstrated that 2Apro of rhinovirus and coxsackievirus B4 cleave eIF4G directly. However, it was suggested that in poliovirus infection, the 2Apro induces the activation of a cellular proteinase which in turn cleaves eIF4G. Furthermore, it is not clear whether eIF4G is cleaved as part of the eIF4F complex or as an individual polypeptide. To address these issues, recombinant eIF4G was purified from Sf9 insect cells and tested for cleavage by purified rhinovirus 2Apro. Here we report that eIF4G alone is a relatively poor substrate for cleavage by the rhinovirus 2Apro. However, an eIF4G-eIF4E complex is cleaved efficiently by the 2Apro, suggesting that eIF4F is a preferred substrate for cleavage by rhinovirus 2Apro. Furthermore, 2Apr drastically reduced the translation of a capped mRNA. An eIF4G-eIF4E complex, but not eIF4G alone, was required to restore translation.     

Fluorescence resonance energy transfer between blue-emitting and red-shifted excitation derivatives of the green fluorescent protein

We report fluorescent resonance energy transfer (FRET) between two linked variants of the green fluorescent protein (GFP). The C terminus of a red-shifted variant of GFP (RSGFP4) is fused to a flexible polypeptide linker containing a Factor X_a protease cleavage site. The C terminus of this linker is in turn fused to the N terminus of a blue variant of GFP (BFP5). The gene product has spectral properties that suggest energy transfer is occurring from BFP5 to RSGFP4. Upon incubation with Factor X_a, the protein is cleaved, and the two fluorescent proteins dissociate. This is accompanied by a marked decrease in energy transfer. The RSGFP4::BFP5 fusion protein demonstrates the feasibility of using FRET between two GFP derivatives as a tool to monitor protein-protein interactions; in addition, this construct may find applications as an intracellular screen for protease inhibitors.     

Evaluation of the CFP-substrate-YFP system for protease studies: advantages and limitations

A protease can be defined as an enzyme capable of hydrolyzing peptide bonds. Thus, characterization of a protease involves identification of target peptide sequences, measurement of activities toward these sequences, and determination of kinetic parameters. Biological protease substrates based on fluorescent protein pairs, which allow for use of fluorescence resonance energy transfer (FRET), have been recently developed for in vivo protease activity detection and represent a very interesting alternative to chemical substrates for in vitro protease characterization. Here, we analyze a FRET system consisting of cyan and yellow fluorescent proteins (CFP and YFP, respectively), which are fused by a peptide linker serving as protease substrate. Conditions for CFP-YFP fusion protein production in Escherichia coli and purification of proteins were optimized. FRET between CFP and YFP was found to be optimum at a pH between 5.5 and 10.0, at low concentrations of salt and a temperature superior to 25 degrees C. For efficient FRET to occur, the peptide linker between CFP and YFP can measure up to 25 amino acids. The CFP-substrate-YFP system demonstrated a high degree of resistance to nonspecific proteolysis, making it suitable for enzyme kinetic analysis. As with chemical substrates, substrate specificity of CFP-substrate-YFP proteins was tested towards different proteases and kcat/Km values were calculated.     

Determinants of substrate recognition by poliovirus 2A proteinase.

Poliovirus proteinase 2A (2Apro) is autocatalytically released from the viral polyprotein by cleavage in cis of a Tyr-Gly dipeptide at its own amino terminus, resulting in separation of the P1 structural and P2-P3 nonstructural protein precursors. A second Ty-Gly dipeptide within 3D polymerase is cleaved by 2Apro in trans, but this is not essential for viral proliferation. The mechanism which limits cleavage to only 2 of the 10 Tyr-Gly dipeptides within the poliovirus polyprotein has not been characterized. We have therefore undertaken a systematic mutational analysis of the VP1-2A site to elucidate determinants of substrate recognition by 2Apro. The P2 and P1' positions are important determinants for cis cleavage of this site, whereas a variety of substituents could be tolerated at the P2', P1, and P3 positions. The requirements for trans cleavage of this site were more stringent. We found that the 2Apro of coxsackievirus type A21 and rhinoviruses 2 and 14 have stringent requirements similar to those of poliovirus 2Apro for cleavage in trans.     

Covalent labeling of cell-surface proteins for in-vivo FRET studies

Fluorescence resonance energy transfer (FRET) is a powerful technique to reveal interactions between membrane proteins in live cells. Fluorescence labeling for FRET is typically performed by fusion with fluorescent proteins (FP) with the drawbacks of a limited choice of fluorophores, an arduous control of donor-acceptor ratio and high background fluorescence arising from intracellular FPs. Here we show that these shortcomings can be overcome by using the acyl carrier protein labeling technique. FRET revealed interactions between cell-surface neurokinin-1 receptors simultaneously labeled with a controlled ratio of donors and acceptors. Moreover, using FRET the specific binding of fluorescent agonists could be monitored.     

A cleavage enzyme-cytometric bead array provides biochemical profiling of resistance mutations in HIV-1 Gag and protease

Most protease-substrate assays rely on short, synthetic peptide substrates consisting of native or modified cleavage sequences. These assays are inadequate for interrogating the contribution of native substrate structure distal to a cleavage site that influences enzymatic cleavage or for inhibitor screening of native substrates. Recent evidence from HIV-1 isolates obtained from individuals resistant to protease inhibitors has demonstrated that mutations distal to or surrounding the protease cleavage sites in the Gag substrate contribute to inhibitor resistance. We have developed a protease-substrate cleavage assay, termed the cleavage enzyme- cytometric bead array (CE-CBA), which relies on native domains of the Gag substrate containing embedded cleavage sites. The Gag substrate is expressed as a fluorescent reporter fusion protein, and substrate cleavage can be followed through the loss of fluorescence utilizing flow cytometry. The CE-CBA allows precise determination of alterations in protease catalytic efficiency (k(cat)/K(M)) imparted by protease inhibitor resistance mutations in protease and/or gag in cleavage or noncleavage site locations in the Gag substrate. We show that the CE-CBA platform can identify HIV-1 protease present in cellular extractions and facilitates the identification of small molecule inhibitors of protease or its substrate Gag. Moreover, the CE-CBA can be readily adapted to any enzyme-substrate pair and can be utilized to rapidly provide assessment of catalytic efficiency as well as systematically screen for inhibitors of enzymatic processing of substrate.     

The Use of a Stably Expressed FRET Biosensor for Determining the Potency of Cancer Drugs

Many cancer drugs are intended to kill cancer cells by inducing apoptosis. However, the potency assays used for measuring the bioactivity of these products are generally cell viability assays which do not distinguish between cell death and growth inhibition. Here we describe a cell-based fluorescence resonance energy transfer (FRET) biosensor designed to measure the bioactivity of apoptosis inducing cancer drugs. The biosensor contains cyan fluorescent protein (CFP) linked via caspase 3 and caspase 8 specific cleavage recognition sequences to yellow fluorescent protein (YFP). Upon caspase activation, as in the case of apoptosis induction, the linker is cleaved abolishing the cellular FRET signal. This assay closely reflects the mechanism of action of cancer drugs, in killing cancer cells and therefore can function as a potency test for different cancer drugs. We rigorously demonstrate this through characterization of a class of proteins targeting the death receptors. The one-step assay appears to be superior to other apoptosis-based assays because of its simplicity, convenience, and robustness.     

Single-molecule FRET study of SNARE-mediated membrane fusion

Membrane fusion is one of the most important cellular processes by which two initially distinct lipid bilayers merge their hydrophobic cores, resulting in one interconnected structure. Proteins, called SNARE (soluble N-ethylmaleimide-sensitive factor-attachment protein receptor), play a central role in the fusion process that is also regulated by several accessory proteins. In order to study the SNARE-mediated membrane fusion, the in vitro protein reconstitution assay involving ensemble FRET (fluorescence resonance energy transfer) has been used over a decade. In this mini-review, we describe several single-molecule-based FRET approaches that have been applied to this field to overcome the shortage of the bulk assay in terms of protein and fusion dynamics.     

Visualization of ternary complexes in living cells by using a BiFC-based FRET assay

Studies of protein interactions have increased our understanding and knowledge of biological processes. Assays that utilize fluorescent proteins, such as fluorescence resonance energy transfer (FRET) and bimolecular fluorescence complementation (BiFC), have enabled direct visualization of protein interactions in living cells. However, these assays are primarily suitable for a pair of interacting proteins, and methods to visualize and identify multiple protein complexes in vivo are very limited. This protocol describes the recently developed BiFC-FRET assay, which allows visualization of ternary complexes in living cells. We discuss how to design the BiFC-FRET assay on the basis of the validation of BiFC and FRET assays and how to perform transfection experiments for acquisition of fluorescent images for net FRET calculation. We also provide three methods for normalization of the FRET efficiency. The assay employs a two-chromophore and three-filter FRET setup and is applicable to epifluorescence microscopes. The entire protocol takes about 2-3 weeks to complete.     

Design of FRET-based GFP probes for detection of protease inhibitors

In this study, tandem Green fluorescent protein (GFP) fusion proteins were designed to detect proteolytic activity of thrombin based on the principle of fluorescence resonance energy transfer (FRET). The thrombin-specific recognition sequence, LVPR, was strategically placed in between a cyan-emitting mutant of the green fluorescent protein and an enhanced yellow-emitting fluorescent protein to allow thrombin-specific cleavage with detectable changes of FRET signal. A 4.6-fold increase of fluorescence emission ratio was observed upon addition of thrombin. This FRET-based probe was further tested for dose-dependent effects of thrombin specific inhibitor, hirudin. Our result showed a nice correlation between fluorescence emission ratios and concentrations of hirudin with subnanomolar sensitivity. We propose that FRET-based GFP probes can be used for high-throughput screening of protease inhibitors.     

Luminometric method for screening retroviral protease inhibitors

We have developed a sensitive luminometric assay for determining the activity of retroviral proteases that uses proteolytic cleavage of polypeptide substrate immobilized on Ni-NTA HisSorb Strips microplates. The protease substrate derived from the Gag precursor protein of Mason-Pfizer monkey virus (M-PMV) was conjugated with horseradish peroxidase (HRP), which catalyzes oxidation of luminol in the assay. The cleavage of the substrate was monitored as a decrease in luminescent signal caused by the release of the cleavage product conjugated to HRP. Testing of a set of M-PMV protease inhibitors confirmed that this method is sufficiently sensitive and specific for high-throughput screening of retroviral protease inhibitors.