Fluorescence assays to identify protein concentration without adding a large peptide sequence?

Fluorescence assays to identify protein concentration without adding a large peptide sequence?

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I'm trying to find a way of tagging a protein with something visually quantifiable to track protein concentration through potential purification steps and screen for the most efficient such steps. However, GFP and its cousins are very large proteins that would change the properties I'm targeting for purification, making them non-starters.

Is there a tag that through fluorescence, colorimetrics, etc. can be used to quantify concentration of a single protein while minimally altering chemical/physical properties of the protein for purification?


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Lodish H, Berk A, Zipursky SL, et al. Molecular Cell Biology. 4th edition. New York: W. H. Freeman 2000.

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AMP-activated protein kinase (AMPK) plays a central role in regulating metabolic activity by downregulating ATP-consuming processes and upregulating catabolism in response to environmental stresses that decrease the cellular [ATP]:[AMP] ratio. Upon activation, AMPK phosphorylates and inactivates acetyl CoA carboxylase (ACC, involved in fatty acid biosynthesis), HMG-CoA reductase (a key regulator of cholesterol synthesis and the target of the statin class of drugs) [1], as well as glycogen synthase [2]. AMPK is a heterotrimeric complex consisting of α, β, and γ subunits [3, 4], and is directly regulated by at least three processes: activation through phosphorylation of Thr-172 in the activation loop of the catalytic α subunit by LKB1 [5-9] or CaMKKα or β [10, 11], deactivation via dephosphorylation at this site by PP2 [12], and allosteric activation via AMP [13].

AMPK is broadly expressed in a variety of tissues, with different isoforms being dominantly expressed in different tissues. The trimeric complex consists of α, β, and γ subunits, with two isoforms identified for each of the α and β subunits, and three identified for the γ subunit, giving rise to a possibility of twelve isoforms of AMPK. In liver, the α1 and α2 subunits are present in approximately equal amounts, with the β1 and γ1 being the predominant forms of their respective subunits [14]. In contrast, in heart and in skeletal muscle the α2 form predominates over the α1 form by a greater than a 2:1 ratio, and the β2 subunit is predominant over the β1 form in these tissues as well [15].

Therapeutic targeting of AMPK activation has been shown to have positive results in models of diabetes. The small molecule drug metformin has been used for approximately 50 years in the treatment of type 2 diabetes, and functions by indirectly promoting AMPK phosphorylation and activation [16]. Another small molecule, AICAR (5-aminoimidazole-4-carbozamide riboside), has proven to be a useful tool for pharmacologically activating AMPK without perturbing the cellular [ATP]:[AMP] ratio. AICAR is taken up into cells via the adenosine transport system and then phosphorylated to form AICAR monophosphate, or ZMP [17], which activates AMPK by binding to the allosteric AMP-binding site. Recently, the results of a highly miniaturized radiometric assay were reported, in which of a library of over 700,000 compounds was screened for AMPK activators or inhibitors [18], and optimization of hits led to a thienopyridone-based small molecule (A-769662) that activated rat-liver or recombinant AMPK (α1β1γ1) with an EC50 of approximately 800 nM, and showed positive effects in the treatment of diabetic ob/ob mice [19]. Interestingly, this compound was able to increase AMPK activity in vitro in the presence of saturating amounts of AMP, suggesting the possibility of multiple allosteric sites that can be exploited to develop compounds that activate AMPK. Additionally, AMPK activation has emerged as a therapeutic target for atherosclerosis and cancer [20].

In addition to therapeutic interest in AMPK activation, there is evidence that either direct inhibition [21] or a leptin-induced decrease [22, 23] in AMPK activity in the hypothalamus can reduce food intake and body weight. Compound C, a pyrazo[1,5-a]pyrimidine compound that inhibits AMPK was discovered in a high-throughput screen [16] and subsequently shown to decrease food intake in mice [21]. Expression of a dominant negative form of AMPK in mouse hypothalamus was shown to reduce both food intake and weight gain, with opposite results seen when a constitutively active form of AMPK was expressed [22].

Because of the key role that AMPK plays in maintaining metabolic homeostasis and the diversity of isoforms that exist in different tissues, there is a need for simple, homogenous, non-radiometric methods to identify and characterize small-molecule modulators of AMPK activity. To address this need, we have developed a suite of assays (Fig. ​ 1 1 ) for different stages of the discovery process. We have developed a time resolved Förster resonance energy transfer (TR-FRET) assay that is well suited to HTS applications, due to its inherent resistance to common forms of assay interference such as colored, fluorescent, or precipitated compounds [24]. Further, we have developed secondary assays that provide a response that is directly proportional to the amount of product formed, and are therefore well suited to detailed mechanistic studies (or hit-confirmation) of small-molecule modulators of AMPK activity. The first of these formats is a FRET-based format in which a peptide substrate for AMPK is labeled on its termini with a coumarin / fluorescein FRET pair. After the kinase reaction, a site-specific protease is added that preferentially cleaves non-phosphorylated substrate, thereby decreasing the FRET signal that remains intact in the non-cleaved, phosphorylated product [25]. In this manner, changes in FRET can be directly correlated to substrate phosphorylation. The second format is based upon the principal of chelate-enhanced fluorescence (CHEF), in which phosphorylation of a serine, threonine, or tyrosine residue causes phosphate-dependent chelation of magnesium between the phosphate group and a non-natural, fluorogenic amino acid (Sox), that has been incorporated into the peptide sequence and that becomes fluorescent upon this chelating event [26]. The FRET-based assay format provides a convenient method for automated compound profiling, and the CHEF-based assay provides the ability to directly monitor purified AMPK activity in real-time.

Schematic of fluorescent assay formats used to characterize AMPK activators of inhibitors. (A) The TR-FRET format detects association between fluorescein labeled, phosphorylated peptide and a terbium-labeled phosphospecific antibody. (B) The FRET-based format uses a peptide substrate terminally labeled with a coumarin-fluorescein FRET pair and measures the amount of phosphorylated product due to a decrease in sensitivity of the phosphorylated peptide to proteolysis. (C) The CHEF-based format uses a peptide substrate that incorporates the non-natural, fluorogenic Sox residue. Upon phosphorylation of a proximal serine, threonine, or tyrosine residue, the Sox moiety forms a Mg 2+ mediated bridge between the Sox residue and the phosphate group and becomes fluorescent.

Chemically-induced protein dimerization is involved in many key regulatory pathways throughout the biological world. [ 1 ] A number of natural products exert their downstream effect by the stabilization of naturally occurring interactions. This can occur via an allosteric effect, whereby an initial interaction between the dimerizer and the target protein results in the formation or uncovering of a new binding interface on the protein, enabling a protein-protein interaction (PPI) that is not otherwise observed. Alternatively, the dimerizer can act by the direct binding of two or more target proteins. This can either involve the binding of one target protein preferentially to create a new, high affinity protein-ligand binding surface for the second protein, or binding to a pre-existing interface between the two proteins can increase the lifetime or affinity of a given PPI. [ 2 ] The former is best characterized by the rapamycin-mTOR interaction. [ 3 ] The latter is typically induced by large macrolide natural products these include the 14-3-3 adapter proteins, whereby stabilization of these chaperone proteins with a range of targets has been demonstrated with natural products such as fusicoccin A and cotylenin A. [ 2, 4 ]

Within the same family as rapamycin, other notable examples include the immunosuppressants FK506 and cyclosporin, both of which are now licensed for therapeutic use. [ 5-7 ] Rapamycin is one of the most widely studied examples of this family. By binding to a small hydrophobic pocket on FKBP12, a new FKBP12-rapamycin binding surface is generated that displays a strong affinity for mTOR (0.2 nM Kd). [ 8 ] Subsequent binding to mTOR results in the inhibition of wide reaching mTOR signaling pathways, [ 9 ] which acts to regulate critical pathways in cell growth and metabolism, as well as proliferation and survival. [ 10 ]

Whilst a variety of techniques have been developed and deployed for the identification of PPI inhibitors, [ 11 ] there are very few examples for the de novo discovery of compounds that upregulate the direct association of two proteins. [ 12 ] Increasingly, chemically-induced protein dimerization is being used for the investigation of complex signaling networks, with sequestering of target proteins providing spatiotemporal control of intracellular proteins. [ 13 ] This typically involves the fusion of protein domains that have prior characterized ligands to the proteins of interest. [ 14-16 ] Taking advantage of chemically-induced protein dimerization for therapeutic or research use first requires a known natural protein binder or dimerizer, However, not all desirable protein targets have an identified natural product ligand that can be used as a starting point for these studies. The stabilization of existing interactions, or the artificial association of two naturally unrelated proteins, represents both a significant research and therapeutic potential in the PPI field.

We have previously reported a platform for the identification of PPI inhibitors that combines a bacterial reverse two-hybrid system (RTHS), a well-established means of investigating PPIs, [ 17-19 ] with an in vivo library generation technique called split-intein circular ligation of peptides and proteins (SICLOPPS). This combined approach allows for the intracellular generation and screening of cyclic peptide libraries of up to a hundred million members for inhibitors of a given PPI. [ 20-24 ] Further developments of this approach include the construction of a bidirectional fluorescent two-hybrid system (FTHS). [ 25 ] Here, we report the construction and validation of a three-hybrid system (3HS) incorporating both a life/death selection and a fluorescent marker for combination with SICLOPPS libraries, allowing for the high-throughput identification of cyclic peptides that induce or upregulate the dimerization of two target proteins. Such a system could enable the identification of new starting points for small-molecule stabilizers or inducers of dimerization.

Methods of Protein Purification: 4 Methods

The methods used in protein purification, can roughly be divided into analytical and preparative methods.

The distinction is not exact, but the deciding factor is the amount of protein, that can practically be purified with that method. Analytical methods aim to detect and identify a protein in a mixture, whereas preparative methods aim to produce large quantities of the protein for other purposes, such as structural biology or industrial use. In general, the preparative methods can be used in analytical applications, but not the other way around.

Method # 1. Extraction:

Depending on the source, the protein has to be brought into solution by breaking the tissue or cells containing it. There are several methods to achieve this Repeated freezing and thawing, sonication, homogenization by high pressure or permeabilization by organic solvents. The method of choice depends on how fragile the protein is and how sturdy the cells are.

After this extraction process soluble protein will be in the solvent, and can be separated from cell membranes, DNA, etc. by centrifugation. The extraction process also extracts proteases, which will start digesting the proteins in the solution. If the protein is sensitive to proteolysis, it is usually desirable to proceed quickly, and keep the extract cooled, to slow down proteolysis.

Method # 2. Precipitation and Differential Solubilisation:

In bulk protein purification, a common first step to isolate proteins is precipitation with ammonium sulphate (NH4)2SO4. This is performed by adding increasing amounts of ammonium sulphate and collecting the different fractions of precipitate protein. One advantage of this method is that it can be performed inexpensively with very large volumes.

The first proteins to be purified are water-soluble proteins. Purification of integral membrane proteins requires disruption of the cell membrane in order to isolate any one particular protein from others that are in the same membrane compartment. Sometimes a particular membrane traction can be isolated first, such as isolating mitochondria from cells before purifying a protein located in a mitochondrial membrane.

A detergent such as sodium dodecyl sulphate (SDS) can be used to dissolve cell membranes and keep membrane proteins in solution during purification however, because SDS causes denaturation, milder detergents such as Triton X-100 or CHAPS can be used to retain the protein’s native conformation during purification.

Method # 3. Ultracentrifugation:

Centrifugation is a process that uses centrifugal force to separate mixtures of particles of varying masses or densities suspended in a liquid. When a vessel (typically a tube or bottle) containing a mixture of proteins or other particulate matter, such as bacterial cells, is rotated at high speeds, the angular momentum yields an outward force to each particle that is proportional to its mass.

The tendency of a given particle to move through the liquid because of this force is offset by the resistance the liquid exerts on the particle. The net effect of “spinning” the sample in a centrifuge is that massive, small, and dense particles move outward faster than less massive particles or particles with more “drag” in the liquid.

When suspensions of particles are “spun” in a centrifuge, a “pellet” may form at the bottom of the vessel that is enriched for the most massive particles with low drag in the liquid. The remaining, non-compacted particles still remaining mostly in the liquid are called the “supernatant” and can be removed from the vessel to separate the supernatant from the pellet.

The rate of centrifugation is specified by the angular acceleration applied to the sample, typically measured in comparison to the g. If samples are centrifuged long enough, the particles in the vessel will reach equilibrium wherein the particles accumulate specifically at a point in the vessel where their buoyant density is balanced with centrifugal force. Such an “equilibrium” centrifugation can allow extensive purification of a given particle.

Sucrose gradient centrifugation:

A linear concentration gradient of sugar (typically sucrose glycerol, or Percoll) is generated in a tube such that the highest concentration is on the bottom and lowest on top. A protein sample is then layered on top of the gradient and spun at high speeds in an ultracentrifuge. This causes heavy macromolecules to migrate towards the bottom of the tube faster than lighter material.

During centrifugation in the absence of sucrose, as particles move farther and farther from the centre of rotation, they experience more and more centrifugal force (the further they move, the faster they move). The problem with this is that the useful separation range within the vessel is restricted to a small observable window.

Spinning a sample twice as long does not mean the particle of interest will go twice as far in fact, it will go significantly farther. However when the proteins are moving through a sucrose gradient, they encounter liquid of increasing density and viscosity.

A properly designed sucrose gradient will counteract the increasing centrifugal force, so the particles move in close proportion to the time they have been in the centrifugal field. Samples separated by these gradients are referred to as “rate zonal” centrifugations. After separating the protein/particles, the gradient is then fractionated and collected.

Method # 4. Chromatographic Methods:

Usually a protein purification protocol contains one or more chromatographic steps. The basic procedure in chromatography is to flow the solution containing the protein through a column packed with various materials. Different proteins interact differently with the column material, and can thus be separated by the time required to pass the column, or the conditions required to elute the protein from the column. Usually proteins are detected as they are coming off the column by their absorbance at 280 nm.

Many different chromatographic methods exist:

1. Size Exclusion Chromatography:

Chromatography can be used to separate protein in solution or denaturing conditions by using porous gels. This technique is known as size exclusion chromatography. The principle is that smaller molecules have to traverse a larger volume in a porous matrix. Consequentially, proteins of a certain range in size will require a variable volume of eluant (solvent) before being collected at the other end of the column of gel.

In the context of protein purification, the eluant is usually pooled in different test tubes. All test tubes containing no measurable trace of the protein to purify are discarded. The remaining solution is thus made of the protein to purify and any other similarly-sized proteins.

2. Ion Exchange Chromatography:

Ion exchange chromatography separates compounds according to the nature and degree of their ionic charge. The column to be used is selected according to its type and strength of charge. Anion exchange resins have a positive charge and are used to retain and separate negatively charged com pounds, while cation exchange resins have a negative charge and are used to separate positively charged molecules.

Before the separation begins a buffer is pumped through the column to equilibrate the opposing charged ions. Upon injection of the sample, solute molecules will exchange with the buffer ions as each competes for the binding sites on the resin. The length of retention for each solute depends upon the strength of its charge.

The most weakly charged compounds will elute first, followed by those with successively stronger charges. Because of the nature of the separating mechanism, pH, buffer type, buffer concentration, and temperature all play important roles in controlling the separation. Ion exchange chromatography is a very powerful tool for use in protein purification and is frequently used in both analytical and preparative separations.

3. Affinity Chromatography:

Affinity Chromatography is a separation technique based upon molecular conformation, which frequently utilizes application specific resins. These resins have ligands attached to their surfaces which are specific for the compounds to be separated. Most frequently, these ligands function in a fashion similar to that of antibody-antigen interactions. This “lock and key” fit between the ligand and its target compound makes it highly specific, frequently generating a single peak, while all else in the sample is un-retained.

Many membrane proteins are glycoproteins and can be purified by lectin affinity chromatography. Detergent solubilized proteins can be allowed to bind to a chromatography resin that has been modified to have a covalently attached lectin.

Proteins that do not bind to the lectin are washed away and then specifically bound glycoproteins can be eluted by adding a high concentration of a sugar that competes with the bound glycoproteins at the lectin binding site. Some lectins have high affinity binding to oligosaccharides of glycoproteins that is hard to compete with sugars, and bound glycoproteins need to be released by denaturing the lectin.

4. Metal Binding:

A common technique involves engineering a sequence of 6 to 8 histidines into the C-terminal of the protein. The polyhistidine binds strongly to divalent metal ions such as nickel and cobalt. The protein can be passed through a column containing immobilized nickel ions, which binds the polyhistidine tag. All untagged proteins pass through the column.

The protein can be eluted with imidazole, which competes with the polyhistidine tag for binding to the column, or by a decrease in pH (typically to 4.5), which decreases the affinity of the tag for the resin. While this procedure is generally used for the purification of recombinant proteins with an engineered affinity tag (such as a 6xHis-tag or Clontech’s HAT tag), it can also be used for natural proteins with an inherent affinity tor divalent cations.

5. Immunoaffinity Chromatography:

Immunoaffinity chromatography uses the specific binding of an antibody to the target protein to selectively purify the protein. The procedure involves immobilizing an antibody to a column material, which then selectively binds the protein, while everything else flows through. The protein can Ix eluted by changing the pH or the salinity. Because this method does not involve engineering in a tag, it can be used for proteins from natural sources.

6. HPLC:

High performance liquid chromatography or high pressure liquid chromatography is a form of chromatography applying high pressure to drive the solutes through the column faster. This means that the diffusion is limited and the resolution is improved. The most common form is “reversed phase” HPLC, where the column material is hydrophobic.

The proteins are eluted by a gradient of increasing amounts of an organic solvent, such as acetonitrile. The proteins elute according to their hydrophobicity. After purification by HPLC the protein is in a solution that only contains volatile compounds, and can easily be lyophilized. HPLC purification frequently results in denaturation of the purified proteins and is thus not applicable to proteins that do not spontaneously refold.


Immunoprecipitation (IP) procedures are applied to a large variety of molecular biology assays, including protein purification, concentration, co-immunoprecipitation and chromatin immunoprecipitation [2–8]. The classical IP procedure is often coupled to immunoblotting analysis, a technique suited to low throughput analysis of only a few samples [10, 20, 21]. Here we describe and characterize FLIP (Fluorescence IP), which couples conventional IP to the direct observation and/or quantification of a fluorescently tagged target protein on the surface of beads. This approach is faster and cheaper than using IB to measure the success of an IP, and is thus amenable to high-throughput screens. A major limitation of the IP assay is often the availability of antibodies able to efficiently immunoprecipitate the target protein [22]. Here we applied FLIP to the screening of mouse monoclonal antibodies in order to quickly screen and identify those able to immuneprecipitate their corresponding target protein.

Here we show that FLIP has a similar sensitivity to the IP/IB assay and that FLIP can also be miniaturized to increase throughput by minimizing both volume of cell lysate and amount of antibody used for IP. Another advantage of FLIP is the use of fluorescently labeled target protein, which ensures a degree of specificity for the antibodies that pass FLIP. Indeed, given that the target protein is the only fluorescently tagged protein in the cell lysate used for IP, the presence of fluorescence coating the agarose beads after immunoprecipitation indicates selective binding of the tested antibody to the target protein. On the other hand, the FLIP assay is “blind” to any possible non-specific binding of the tested antibody to proteins different from the fluorescently labeled target protein. Another possible limitation of FLIP is that it does not currently give any information about the ability of the tested antibody to recognize the target protein in its denatured state and therefore FLIP cannot predict the behavior of antibodies in immunoblotting assays. To this end, FLIP could theoretically be used using protein solutions/lysates pre-treated with denaturing conditions such as high SDS concentrations and/or reducing agents [5]. This approach may be possible and somehow informative because of the high stability of GFP and GFP derived fluorophores that can withstand denaturing conditions undergoing partial denaturation and still remaining fluorescent [23].

Also, FLIP cannot assay the performance of an antibody in IP of endogenous proteins because it relies on the overexpression of the target protein. Nevertheless, we show here that FLIP can easily be integrated into classical IP/IB procedures with no disruption and no diminished sensitivity of the latter. This allows for further characterization of the antibodies after FLIP screening for the IB analysis of bands of the endogenous protein. Moreover, FLIP relies on the overexpression of the target protein as a fluorophore-tagged fusion and therefore requires the cloning of the gene of interest into a suitable expression vector. To this end we developed a flexible mammalian expression vector named HuEV-A that combines the simplicity of Gateway cloning with the flexibility of our 3xFLAG-V5-YFP tag [18, 19, 24]. We used the Life Technologies Ultimate ORF collection, compatible with Gateway cloning (Gateway entry clone format) to construct a HuEV-A library of proteins. Also, the HuEV-A tag can be dramatically reduced in size with simple Cre recombination to a 3xFLAG-V5 tag, or the entire tag can be eliminated by employing FLP recombination.

The FLIP assay is very similar in basic concept to the previously described LUMIER (luminescence-based mammalian interactome mapping) assay [12, 13] which has been applied to a wide range of applications, including identification of new protein-protein interactions [12, 25–27], validation of yeast two hybrid screening hits [28, 29] and identification of particular antibodies in patients’ blood (Luciferase Immunoprecipitation Systems or LIP) [30]. LUMIER exploits the overexpression of a target protein fused to Renilla luciferase and utilizes quantification of luminescence signal after immunoprecipitation. As for FLIP, the success of the immunoprecipitation of the target fusion protein can be efficiently and easily determined by high-throughput measurements: measurement of luminescence after incubation with a luciferase substrate in the case of LUMIER and LIP, and measurement of the fluorescence coating the immunoprecipitated beads in the case of FLIP. Despite the great success and the many applications of the LUMIER system no alternative assays with high-throughput capability has been developed for IP analysis until now.

Our FLIP assay is not meant to substitute for the LUMIER assay, which is probably more sensitive compared to IP/WB and FLIP analysis itself. Our FLIP assay is meant to complement the previously known techniques (IP/WB and LUMIER assays) in high-throughput procedures challenging for luciferase reactions or in contexts in which microscopy observations are more amenable than luciferase signal quantification or in which fluorophore tagging is preferable to Renilla tagging. The FLIP assay is an additional tool for the fast screening of IP reactions. FLIP can also be envisioned as an orthogonal assay to the LUMIER system because “preys” tagged with FLAG and a fluorophore of choice can be used with Renilla tagged “baits” (see [12]) to verify and quantify efficiency of IP before luminescence quantification. In this context our HuEV-A expression vector that includes a FLAG, V5 and YFP tag would be optimal.

Compared to IP-Western blotting, some information such as the nonspecific reactivity of the antibody towards non-target proteins, and/or the preferential reactivity towards specific isoforms or splice variants of the target protein, are lost. Nevertheless, we foresee an extremely useful application of FLIP in screening for antibodies that function efficiently in IP assays. For instance, many unpurified antibodies can be tested by FLIP directly using hybridoma supernatants. After selection of the FLIP-positive clones, a reduced number of the antibodies would need to be purified and characterized further by IP/IB or other assays. Moreover, because of the minimal material used in the FLIP analysis, a new IP does not need to be performed again for the FLIP-positive antibodies, as the left-over beads from FLIP analysis can be used for the follow-up characterizations.

Here we show that FLIP is a fast and reliable method that can partially substitute and easily complement the conventional IP/IB procedure. This new technique can be directly applicable to the high-throughput screening for the identification of IP-grade antibodies.

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Development of a Whole-Cell Assay for Peptidase Activity.

Given the utility of FACS as a quantitative library screening tool, we sought to assay for peptidase activity by displaying reporter substrates on the surface of Esherichia coli (Fig. 1A). Reporter substrates were designed consisting of a peptide ligand that binds the fluorescent probe streptavidin-conjugated phycoerythrin (SA-PE) and a peptide substrate oriented such that cleavage removes the SA-PE-binding ligand from the cell surface. In this way, protease activity toward a given substrate would be detectable by monitoring whole-cell fluorescence by using FACS. Reporter substrates were displayed on E. coli by using circularly permutated outer membrane protein X (CPX), which presents both N- and C termini on the cell surface, enabling presentation of passenger peptides as nonconstrained, terminal fusions (18). As a control, a substrate-reporter display vector was constructed incorporating a known enteropeptidase cleavage site (DDDDK) flanked by flexible peptide linker sequences, as “spacers,” allowing protease access to the substrate, and a SA-PE-binding peptide ligand. Cells displaying the substrate were fluorescently labeled with SA-PE, resulting in a >20-fold increase in mean fluorescence intensity over background autofluorescence, as measured by flow cytometry (Fig. 2). Incubation of this cell population with enteropeptidase before labeling with SA-PE resulted in a ≈20-fold decrease in mean fluorescence intensity (Fig. 2B), whereas a negative control cell population displaying the sequence GGQSGQ exhibited minimal change in fluorescence (Fig. 2A). These results demonstrated that enzymatic cleavage of reporter substrates could be detected as a decrease in fluorescence intensity of cells by using FACS and hydrolysis is not due to cleavage outside of the designated substrate region.

CLiPS. (A) Display of reporter substrates, consisting of a substrate peptide and fluorescent-probe peptide ligand, on the surface of E. coli as fusions to the N terminus of circularly permuted outer membrane protein OmpX (CPX). Substrate cleavage results in a reduction of cellular fluorescence as detected by flow cytometry. (B) Substrate libraries are screened by depleting the library pool of clones that do not display a peptide and then enriching clones with hydrolyzed substrates.

Measurement of substrate conversion by FACS. Flow cytometry analysis of bacterial cell populations displaying either linker (GGSGGS) (A) or canonical substrate (DDDDK) (B) before (gray line) and after (black line) treatment with enteropeptidase. During library screening for enteropeptidase substrates, cell populations collected from screen 1A (C) and screen 3B (D) were analyzed by flow cytometry before (gray line) and after (black line) enteropeptidase treatment. The loss of fluorescence due to treatment, shown by the shift in the black line, demonstrates enrichment of enteropeptidase substrates after screen 3B (D).

To extend this single-cell substrate cleavage assay to identify optimal substrates for a given protease, a substrate library was constructed in E. coli by combinatorial randomization of six sequential amino acid positions within the substrate region (Fig. 1A). This CLiPS had a theoretical diversity of 6.4 × 10 7 unique amino acid sequences. The constructed library contained 1.5 × 10 8 independent transformants. Thus, this library is expected to include all possible 5-mer and 4-mer substrate sequences with >95% and 99% confidence limits, respectively, assuming a random distribution (19). Using the whole-cell activity assay, a screening methodology was devised to isolate library members displaying substrates cleaved by a given protease and, thereby, identify optimal substrates.

Determination of Enteropeptidase and Caspase-3 Specificity by Using CLiPS.

To demonstrate the general utility of CLiPS, the 6-mer substrate library was screened to identify optimal substrates for two unrelated proteases: caspase-3 and enteropeptidase. These proteases recognize the canonical substrates DEVD↓ (20) and DDDDK↓ (4), respectively. Caspase-3 was chosen to validate CLiPS, because specificity has been investigated extensively by using both substrate phage and fluorogenic substrates (21, 22). In contrast, enteropeptidase specificity is less well characterized and has been investigated primarily by using individually synthesized, fluorogenic substrate variants (23). For each protease, optimal substrates were identified by performing a two-step screen for hydrolysis (Fig. 1B). First, library members that display the affinity epitope were purified by sorting (Fig. 1B), thereby removing library members that do not display substrates (i.e., members with stop codons and frameshift mutations). The resulting library population was amplified by growth, treated with protease, labeled with SA-PE, and cells with reduced fluorescence resulting from substrate hydrolysis were collected (Fig. 1B). After three cycles of screening for enteropeptidase substrates, >95% of the enriched library displayed reporter substrates and exhibited cleavage similar to the canonical substrate (Fig. 2 C and D). Therefore, a final sort was performed to identify enteropeptidase substrates that hydrolyzed more rapidly than the canonical substrate.

In applications that involve complex protease-containing mixtures, such as cellular lysates or tissue extracts, we anticipated that specificity could be identified by removing substrates that are cleaved by an appropriate background mixture. For this reason, we investigated whether substrates of a target protease can be determined in the presence of cell lysates. Nonspecifically cleaved substrates were depleted from the library first by incubation with E. coli lysate protein that does not contain the target protease, caspase-3 (Fig. 1B). Subsequently, cells displaying specifically cleaved substrates were isolated after incubating the library with lysate from E. coli-expressing caspase-3 (Fig. 1B). This process ensured that cleavage during screening was due to caspase-3 activity and not endogenous E. coli proteolytic activity. Two cycles of screening resulted in the enrichment of library members exhibiting caspase-3 dependent cleavage. Incubation of the enriched library with caspase-3 containing lysates, but not caspase-3-free lysates, resulted in a reduction of the mean fluorescence intensity of the population, as measured by flow cytometry (data not shown). Single clones from the enriched library were isolated from the remaining population by plating. Thus, CLiPS was capable of identifying caspase-3-specific substrates in the presence of a complex mixture.

Characterization of Substrate Cleavage Kinetics.

The use of multicopy substrate display on whole cells enabled simple and direct quantitative characterization of cleavage kinetics. Consequently, flow cytometry was used to rank individual isolated clones on the basis of substrate conversion, and those clones exhibiting >50% conversion were identified by DNA sequencing (Tables 1 and 2). Substrates efficiently cleaved by caspase-3 revealed a strong substrate consensus of DxVDG (Table 1), in agreement with the known specificity of caspase-3. The substrates identified for enteropeptidase shared a consensus sequence of D /ERM, indicating a substrate preference at the P1′ position (Table 2). Interestingly, enteropeptidase substrates identified by CLiPS were cleaved more rapidly than the canonical sequence, DDDDK (Table 2). Four isolated clones with high conversion were investigated further to quantify cleavage kinetics. Clones exhibiting multiple arginine residues were excluded to avoid substrates that may have multiple cleavage sites. Individual substrate displaying clones (e.g., EP4.1 EP, enteropeptidase) exhibited uniform substrate turnover (Fig. 3A), as determined by flow cytometry. In this way, the extent of conversion for each clone could be determined at several different time points and fit to a Michaelis–Menton model (Fig. 3B). The observed second-order rate constant (kcat/KM) for the most rapidly cleaved substrate (EP4.3 SGDRMW) was 13-fold greater than that for the canonical substrate DDDDK (Table 3).

Caspase-3 substrates identified by using CLiPS with a 6-mer library

Enteropeptidase substrates identified by using CLiPS with a 6-mer library

Enteropeptidase substrate cleavage kinetics. (A) Time-dependent substrate conversion for clone EP 4.1 (VDYRFL) measured by FACS. (B) Average conversion for cell surface displayed enteropeptidase substrates identified by using CLiPS: VDYRFL (○), SGDRMW (▵), and SGERMM (×) with canonical DDDDK (♢). Data were fit to Michaelis–Menton model, which is shown as a line for each substrate.

Comparison of kinetic constants determined by using CLiPS, fluorescent protein FRET substrates, or synthetic peptides

To determine how cleavage kinetics (kcat/KM) measured by using surface displayed reporter substrates relate to those measured in solution, two independent approaches were applied to measure kcat/KM for soluble substrates. Because enteropeptidase is often used to remove peptide affinity tags, substrates were assayed in the context of a fusion protein. Specifically, fluorogenic substrates were constructed by using fluorescent proteins that exhibit FRET (CyPet and YPet) (24) and were used to determine protease cleavage kinetics as described in ref. 25. CyPet-YPet substrates for enteropeptidase having recognition sequences of DDDDKG, GGSGGS, or four sequences identified by CLiPS (EP4.1, EP4.2, EP4.3, or EP4.6) were constructed, expressed in E. coli, and purified. Substrate conversion by enteropeptidase was measured in real-time by fluorimetry and fit to Michaelis–Menton kinetics (Table 3). In relative agreement with whole-cell assays, the CLiPS substrate, SGDRMW, cleaved at a rate 17-fold faster than DDDDK. Absolute values of kcat/KM for cell-surface-tethered and soluble substrates differed systematically, but importantly, the relative ranking of the cleavage rates of individual substrates was identical in either context. To further confirm the improved hydrolysis rate for the SGDRMW substrate, relative to DDDDK, fluorogenic peptide substrates were synthesized and cleavage was measured by using fluorimetry (Table 3). The kcat/KM of the CLiPS-identified substrate SGDRMW was >5-fold higher than that of DDDDK. Collectively, these results demonstrate that whole-cell fluorescence assays provide a reliable means to quantitatively measure and rank cleavage kinetics of individual substrate sequences and that CLiPS enables identification of substrates with improved cleavage kinetics.


For many experiments, the first consideration will be what color (spectral properties) of fluorescent tag to use. The spectral properties of a fluorophore are given by its excitation and emission spectra. The excitation spectrum describes the wavelengths of light that, when absorbed, will result in the fluorophore reemitting light the spectrum of that emitted light is the emission spectrum. These are often reduced to numbers describing the peak excitation and emission wavelengths, but the spectra are often broad, and this simplification can obscure important information. To image two fluorescent tags in different channels requires that the excitation and emission spectra of one tag be sufficiently separate from those of the other tag (typically 60–100 nm) so that filters can be chosen that selectively detect each protein.

In many cases, the choice of colors is dictated by the instrumentation to which one has access and the fluorescent proteins it is designed to detect. For example, nearly all of the microscopes in the imaging center I direct can image blue, green, red, and infrared fluorescent proteins, but only a few microscopes are equipped with filters for cyan and yellow fluorescent proteins (CFP and YFP, respectively). I advise users of our center to avoid CFP and YFP if possible to maximize their imaging options. This is particularly true for instruments using laser illumination such as confocal, total internal reflection fluorescence, and light-sheet microscopes and flow cytometers, for which changing excitation wavelengths is expensive and difficult.

In general, I recommend starting with green and red fluorescent proteins, as these tend to be the brightest and best studied and have been found to work well for many applications. Somewhat surprisingly, EGFP still performs well in many systems, although newer proteins such as mClover3 and mNeonGreen outperform it in mammalian cells (Shaner et al., 2013 Bajar et al., 2016b). For red fluorescent proteins, mCherry was for many years the protein of choice, but it is now being supplanted by brighter and more photostable proteins. It appears likely that mScarlet (Bindels et al., 2017) will be the new red fluorescent protein of choice, but other proteins, such as mRuby3, TagRFP-T, and mKate2, may be worth considering (Shaner et al., 2008 Shcherbo et al., 2009 Bajar et al., 2016b). If additional colors are needed, mTagBFP2 (Subach et al., 2011) can be used with these with minimal cross-talk, although it is less bright than EGFP and phototoxicity is a concern with the near-ultraviolet (UV) excitation required. For a fourth color, a near-infrared fluorescent protein can be used, although these require a ligand. tdsmURFP is the brightest near-infrared fluorescent protein, although it requires supplementation with a biliverdin methyl ester to achieve maximum brightness (Rodriguez et al., 2016b). The next brightest monomeric option is mIRFP670 (Shcherbakova et al., 2016). However, both of these proteins are very new and have not been studied extensively. An alternative option is to use a self-labeling tag like Halo tag and an infrared dye these dyes, such as Cy5 or Alexa 647, are substantially brighter than the infrared fluorescent proteins. Four-color imaging with this combination is relatively straightforward, as both the fluorescent proteins and relevant filter sets are readily available.

CFP, YFP, and a red fluorescent protein can be used for three-color imaging (Livet et al., 2007), and an alternate system for four-color imaging has recently been demonstrated with mTurquoise2, Clover, mKO2, and mMaroon1 (Bajar et al., 2016a). In principle, both of these combinations should be extensible with the incorporation of a blue and infrared fluorescent protein to five and six colors. However, these combinations are less well tested, and filter sets for them are not widely available.

Other possibilities for imaging more than four fluorescent proteins at once include the use of long–Stokes shift proteins, which have a large separation between their excitation and emission wavelengths, enabling multiplexing with short–Stokes shift proteins. For example, T-Sapphire is a UV-excited GFP variant that can be multiplexed with the mWasabi green fluorescent protein, which is not UV excited (Ai et al., 2008). This pair can be further combined with mTagBFP2, allowing three proteins to be imaged in the spectral space previously used for two. The near-infrared proteins iRFP670 and iRFP720 are sufficiently spectrally separate to allow two-color imaging (Shcherbakova and Verkhusha, 2013), suggesting that iRFP720 could be multiplexed with other near-infrared fluorescent proteins to access an additional channel. However, iRFP720 is dimeric, and filter sets for this protein are not common, complicating use of this option. Another option for multiplexing larger numbers of colors is to acquire fluorescence at many wavelengths and use computational tools to separate overlapping fluorophore spectra (Zimmermann, 2005 Cutrale et al., 2017), although this requires specialized hardware and software.


Imaging confluently stained DNA molecules is complicated by issues of photodamage and phototoxicity mediated by organic dyes. Although fluorescent proteins do not generally suffer such issues and are well-targeted by many fusion techniques to cellular and molecular structures, they do not offer the ubiquitous binding patterns inherent to simple dyes. As such, we combined the virtues of organic and protein fluors by creating fluorescent fusion proteins (FP-DBP) featuring tightly binding peptides that promiscuously cover DNA molecules and when expressed within E. coli cells enabled the localization of nucleoid structures. Controllable expression of FP-DBPs will likely foster other new routes to localization of nucleic acids within living cells and offers yet another important advantage over organic dyes. Aside from reversible staining via pH shifts, we have also demonstrated FP-DBP confluent staining of naked DNA molecules in ways comparable to organic dyes, but featuring minimal photodamage during irradiation, and minimal perturbation of the DNA polymer contour length.

We thank Prof. Jungho Kim and Prof. Kwanghwan Jung for providing the mCherry and eGFP genes.

Watch the video: gA based fluorescence assay (August 2022).