Removal of the Initial Methionine in Venus for FRET

I'm working on building some FRET reporters. In addition to a cleavage site (of varying composition from 15-18AA), a 1 AA linker, I'm using Venus and Cerulean.

Initially I was worried that 18AA would be too long, but it does in fact give a fair signal. For reference, I believe the Förster radius (Ro) is 54Å.

I am now working on further optimization. Does the removal of the intial methionine cause any problems for Venus if it is the second protein in the FRET pair? The basic layout is:

Cerulean - Link - Cleavage - Link - Venus

And I'm wondering if cutting the Met from Venus might bring me 1 AA closer and therefore a slightly better signal. I have seen it published both ways, and I was wondering if anyone had tried both or just knew theoretically that it wouldn't matter.

Activated Ras Protein Accelerates Cell Cycle Progression to Perturb Madin-Darby Canine Kidney Cystogenesis*

In a number of human cancer cells, K-RAS is frequently mutated and activated constitutively, culminating in the induction of continuous cell growth, a hallmark of cancer cells. It is still unclear, however, how the mutated K-RAS induces morphological abnormalities in cancerous tissues. To investigate the mechanism underlying the K-RAS-induced morphological changes, we utilized an auxin-dependent protein expression system, which enabled us to rapidly induce and evaluate constitutively active K-Ras in MDCK (Madin-Darby canine kidney) cysts, a model for polarized epithelial structure. Cells carrying the constitutively active KRasV12 gene were morphologically indistinguishable from normal cells in two-dimensional culture. However, in a gel of extracellular matrix, KRasV12-expressing cells failed to form a spherical cyst. When KRasV12 induction was delayed until after cyst formation, some cells in the cyst wall lost polarity and were extruded into and accumulated in the luminal space. With effector-specific mutants of KRasV12 and inhibitors for MEK and PI3-kinase, we found that both the Raf-MEK-ERK and PI3-kinase axes are necessary and sufficient for this phenotype. Live cell imaging with cell cycle indicators showed that KRasV12 expression promoted cell cycle progression, which was prevented by either MEK or PI3-kinase inhibitors. From these results, we provide a model wherein active-Ras induces cell cycle progression leading to apical cell extrusion through Raf and PI3-kinase in a cooperative manner. The system developed here can be applied to drug screening for various cancers originating from epithelial cells.

This work was supported by grants from the Ministry of Education, Culture, Sports, Science, and Technology of Japan and the Mochida Memorial Foundation for Medical and Pharmaceutical Research.

Supported by research fellowships from the Japan Society for the Promotion of Science for Young Scientists.

Single Molecule Tools, Part B:Super-Resolution, Particle Tracking, Multiparameter, and Force Based Methods

Michael A. Thompson , . W.E. Moerner , in Methods in Enzymology , 2010

2.4 EYFP as a photoswitchable emitter

The use of EYFP as a photoswitchable emitter vastly expands the number of biological specimens immediately available for super-resolution imaging. In most reported PALM imaging, the photoactivatable fluorescent protein has been selected from various sophisticated constructs such as PA-GFP, Dronpa, Kaede, tdEosFP, Dendra2, rsFastLime, PA-mCherry, and rsCherryRev ( Betzig et al., 2006 Geisler et al., 2007 Niu and Yu, 2008 Stiel et al., 2008 Subach et al., 2009 ). However, single immobilized and apparently bleached yellow FPs (S65G, S72A, T203Y or S65G, S72A, T203F) were shown to reactivate with violet light more than 10 years ago ( Dickson et al., 1997 ), and the closely related enhanced yellow fluorescent protein EYFP (S65G, V68L, S72A, T203Y) was recently used for live-cell PALM imaging of the C. crescentus structural protein MreB ( Biteen et al., 2008 ). EYFP is widely used for routine fluorescent protein fusions due to the absence of physiological perturbations such as agglomeration and mislocalization, and N-terminal EYFP-MreB fusions have been previously shown to be functional in C. crescentus and other bacteria, making this a physiologically relevant system ( Carballido-López and Errington, 2003 Figge et al., 2004 Gitai et al., 2004 ). Furthermore, single-molecule imaging of EYFP-MreB with 514-nm excitation has previously shown that this fluorescent protein is a bright single-molecule emitter for live-cell imaging ( Kim et al., 2006 ).

Figure 2.5 shows EYFP photoreactivation in live cells ( Biteen et al., 2008 ). Figure 2.5A shows a single imaging frame where the fluorescence image is superimposed over a negative-contrast white-light transmission image of the C. crescentus cell. Two single EYFP-MreB molecules can be identified in panel A, which was acquired after the initial bleaching step. After further imaging with 514-nm light, all fluorophores had bleached, as observed in panel B. EYFP reactivation was achieved after this initial bleaching step by administering a 2-s dose of 407-nm laser illumination. This pulse length, as well as the reactivation intensity of 10 3 –10 4 W cm − 2 , was chosen such that, at most, one EYFP molecule was reactivated in each diffraction-limited region and a different subensemble of EYFP molecules was activated in each cycle of this process.

Figure 2.5 . Reactivation of EYFP-MreB fusions in live C. crescentus cells. (A) Single 100-ms acquisition frames showing isolated EYFP-MreB molecules (a, c, and e) upon photoactivation and no single-molecule emission (b, d, f) after photobleaching. The spot in the bottom left of each image is an imaging artifact. (B) Bulk reactivation of a sample of 22 cells. The grey lines indicate 2-s pulses of 407 nm light. (Adapted and reproduced with permission from Biteen et al., 2008 . Copyright 2008 Nature Publishing Group.)

In Fig. 2.5B , the total emission intensity from 22 live C. crescentus cells expressing EYFP-MreB is displayed as a function of time. After an initial bleaching period, flashes of 407-nm activation energy were applied between imaging cycles. These reactivation cycles were used to calculate the photoreactivation quantum yield of the EYFP fluorophore. The measured relationship between activation time and percent reactivation can be plotted and is quasi-linear. In measurements of EYFP-MreB in live C. crescentus cells, 370 photons are absorbed by each EYFP molecule per second of the activation pulse. From the slope of the plot, a reactivation quantum yield of 1.6 × 10 –6 was found for EYFP ( Biteen et al., 2009 ). This number is on the same order of magnitude as the activation quantum yield for PA-GFP, and only 1 order of magnitude smaller than the photoswitching quantum yield of the highly engineered protein, tdEos (see Table 2.1 ). EYFP can therefore be viewed as a useful photoswitchable fluorophore for super-resolution imaging.


Centralspindlin is an evolutionarily conserved, constitutive 2:2 heterotetrameric complex of a kinesin-6 subunit MKLP1 and a non-motor subunit CYK4. In this article, we shall use the term “MKLP1” to refer collectively to the orthologs of the mammalian KIF23/MKLP1 [1], such as Caenorhabditis elegans ZEN-4, and the term “CYK4” to denote the orthologs of C. elegans CYK-4 [2], such as mammalian RACGAP1/MgcRacGAP [3]. Centralspindlin plays essential roles in cytokinesis by forming central spindle and midbody microtubule bundle structures, by recruiting and regulating various factors at the site of division, and by anchoring the plasma membrane in the intercellular bridge while the daughter cells are waiting for abscission [4–11]. A point mutation in KIF23/MKLP1 is the cause of congenital dyserythropoietic anemia type III, which is characterized by large multinucleated erythroblasts in bone marrow [12]. Both the MKLP1 and CYK4 subunits are essential for microtubule bundling by centralspindlin [3,13]. In vitro, neither MKLP1 alone nor CYK4 alone can efficiently bundle microtubules. In vivo, depletion of either component or point mutations that affect the formation of the centralspindlin heterotetramer cause the central spindle defects [2,3,14–19]. Strikingly, although genetic screens in C. elegans for suppressors of such complex-disrupting mutations (S15L in CYK-4 or D520N in ZEN-4) so far identified 15 independent point mutations, all reside within limited regions CYK-4 12–39 and ZEN-4 477–515. It is likely that these findings define the binding interfaces between these subunits [3,13] (summarized in Fig 1A). These are included in the minimal domains of CYK-4 and ZEN-4 sufficient for in vitro reconstitution of the stable complex between them (CYK-4 1–120 and ZEN-4 435–555). These data emphasize the importance of heterotetramer formation for microtubule bundling and suggest that the tetramer assembly is achieved through compact domains without extensive contact, such as a long four-helix bundle. However, it remains unclear how CYK4 contributes to microtubule bundling.

(A) A schematic of the domain structures of CYK-4 and ZEN-4 and their fragments expressed as fusion proteins with purification tags, which were removed during purification. Magenta and cyan vertical segments denote the residues whose mutation disrupts the CYK-4–ZEN-4 interaction and whose mutation suppresses the interaction defect, respectively. Expected behaviors of the ZEN-4 constructs in dimerization, microtubule-binding (MT-binding) and clustering are indicated (+ or –) based on previous results [13,34], except for microtubule-binding of Z555ΔN (*), which is based on results described below. CC1, CC2: coiled coil predicted with high and low propensity, respectively C1: C1 domain RhoGAP: Rho-family GTPase-activating protein domain Kinesin: kinesin motor domain ARFB: ARF6-binding domain [9,10], BIO: 20 aa biotinylation tag. (B–E) Protein preparations used in this study were run on SDS-PAGE gels and visualized with Coomassie stain. Note that the version of Z601 used in Z601/C120, which lacks the biotinylation tag, showed a slightly larger mobility than those used alone or in a complex with C40G.

Although MKLP1, a kinesin-6 [20], has the kinesin motor domain at its N-terminus, it is distinct from other N-terminal kinesins (N-kinesins). In kinesins-1, -2, -3, -4, -5, and -7, the catalytic core, which contains an ATP-binding pocket and a microtubule-binding surface, is connected to the coiled coil via a short conserved sequence motif of about 15 aa, so-called the "neck linker" [21–23]. The neck linker docks to the catalytic core in a manner bidirectionally coupled to the nucleotide and microtubule-binding states of the catalytic core. Thus, it plays crucial roles in force-generation by individual motor domains and in the mechanochemical coordination between the two heads in walking kinesin dimers. Interestingly, MKLP1 does not have a recognizable neck linker. Instead, the catalytic core of MKLP1 is connected to the coiled coil via a longer "neck" sequence of 60–70 aa, which contains dispersed helix/coil-breaking proline residues (“MKLP1” in S1 Fig) [3]. Importantly, this neck region of MKLP1 contains the CYK4 binding site. Because of these unusual mechanistic features, the molecular structure of the heterotetrameric centralspindlin complex of CYK4 and MKLP1 is of great interest. Although a recent study using electron paramagnetic resonance spectroscopy reported a change in the mobility of ZEN-4 neck residues upon CYK-4 binding [24], it remains unclear how this influences the configuration of the two motor domains and how this modulates microtubule bundling by the complex.

Here we performed direct visualization of the dynamic structure of centralspindlin by atomic force microscopy. Our data indicate that CYK4 binding to the neck domain of the MKLP1 dimer remodels the configuration between the two motor domains. Furthermore, by using in vitro functional assays, we demonstrate that this reconfiguration optimizes the centralspindlin complex for the formation of and accumulation to the antiparallel microtubule bundles crucial for cytokinesis.


Since MetQ has a high affinity for l -methionine, preparation of the apo-MetQ requires cycles of unfolding and refolding to remove the bound ligand. 11 As an alternative approach, mutation of Asn229 to Ala (N229A) in the binding pocket of the E. coli MetQ was found to substantially decrease the affinity toward methionine, thereby greatly facilitating preparation of the substrate-free form of MetQ. Despite extensive efforts with both approaches, we could not crystallize the substrate-free form of E. coli MetQ. While screening structurally characterized MetQs from the RCSB Protein Data Bank, we were able to prepare and crystallize a substrate-free form of the N. meningitides MetQ (PDB 3IR1 14 ), containing the Asn to Ala mutation at residue 238 (N238A corresponding to N229A of the E. coli MetQ) to disrupt substrate binding. As this asparagine residue interacts with both the α-amino and α-carboxyl groups of l -methionine, mutation to alanine would be expected to significantly impair ligand binding. Indeed, isothermal titration calorimetry (ITC) studies of ligand binding to the wild type and N238A forms of N. meningitides MetQ quantitate the impact of this mutation on the binding of l -methionine, with the dissociation constant (Kd) changing by a factor of over 10 5 , from 0.2 nM to 78 μM the binding of d -methionine is also impaired, but to a lesser extent as the Kd changes from 3.5 to 240 μM (Figure 1 and Table 1).

Proteins Ligands Kd (μM) ΔH (kJ mol −1 ) Incompetent fractiona a Incompetent fraction is the fraction of MetQ that is apparently not able to bind to titrant.
Wild-type l -methionine 0.00020 [0.00017, 0.00034]b b About 68.3% confidence intervals determined by error-surface projection 18 are shown in square brackets.
−83 [−84, −81] 0.06 [0.049–0.067]b b About 68.3% confidence intervals determined by error-surface projection 18 are shown in square brackets.
Wild-type d -methionine 3.5 [2.5, 4.6] −55 [−57, −53] 0.06 [0.049–0.067]b b About 68.3% confidence intervals determined by error-surface projection 18 are shown in square brackets.
N238A l -methionine 78 [69, 89] −42 [−50, −37] 0.04 [0, 0.067]b b About 68.3% confidence intervals determined by error-surface projection 18 are shown in square brackets.
N238A d -methionine 240 [190, 310] −31 [−34, −29] 0.31 [0.28–0.34]b b About 68.3% confidence intervals determined by error-surface projection 18 are shown in square brackets.
  • Abbreviation: ITC, isothermal titration calorimetry.
  • a Incompetent fraction is the fraction of MetQ that is apparently not able to bind to titrant.
  • b About 68.3% confidence intervals determined by error-surface projection 18 are shown in square brackets.

The crystal structure of the substrate-free N238A MetQ was determined at 1.56 å resolution (Figure 2a, PDB 6CVA) and reveals an open conformation with an accessible substrate cavity. Superposition of one domain of the l -methionine bound MetQ (PDB: 3IR1) on the corresponding domain of substrate-free N238A MetQ reveals a 42° hinge-type rotation that separates the two SBP domains and opens up the substrate-binding cavity (Figure 2b). This “Venus-fly trap” hinge movement contrasts with that observed for the relationship between liganded MetQ and substrate-free MetQ in the MetNIQ complex, that are related by a 24° twist around an axis perpendicular to the interface between the two lobes. 7 The root mean square deviation (rmsd) in Cα positions between the substrate-free N238A and the l -methionine bound form of N. meningitidis MetQ is 3.7 å, while the corresponding rmsd between the two substrate-free forms of MetQ (N238A and the conformation of MetQ in complex with MetNI, PDB 6CVL) is 4.4 å.

MetQ can bind to other methionine derivatives, including d -methionine, but preparing these forms has been complicated by the high affinity for l -methionine. For example, a previous effort to crystallize d -methionine-bound MetQ was made by Yang et al 14 by growing a methionine auxotroph E. coli B384 strain in the media containing 50 mg L −1 of d -methionine. Due to the presence of amino acid racemases in E. coli, however, they obtained the l -methionine bound structure. 14 Hence, to crystallize the d -methionine bound N. meningitides MetQ, we developed an alternative approach by unfolding/refolding MetQ during the MetQ purification to remove the endogenous bound l -methionine. A sample of unfolded/refolded MetQ was then mixed with 10 mM d -methionine prior to the crystallization. The crystal structure of the d -methionine bound MetQ was solved at 1.68 å resolution with a clear density of d -methionine in the binding cavity (Table 2, PDB 6DZX). Six molecules are present in the asymmetric unit of the d -methionine bound MetQ crystal. As a control, we also determined the structure of the l -methionine bound N. meningitides MetQ crystallized under similar conditions (Table 2, PDB 6OJA). Superposition of the l - and d -methionine bound MetQ structures reveal their similar conformation, with rmsd ∼0.1 å (Figure 3a). While these structures are similar to the previously determined 3IR1 structure, 14 rmsd ∼0.4 å, the space groups and packing interactions are distinct in these two studies.

Not surprisingly, l - and d -methionine bind to the same site, interacting with the same set of conserved residues. This observation is consistent at all six MetQ molecules present in the asymmetric unit. The α-amino and α-carboxyl groups of both l - and d -methionine interact with residues R156, N213, and N238 located on one domain of MetQ, while from the other lobe, residues Y81, F98, H100, and Y103 pack around the methionine thioether group (Figure 3b). These binding residues are highly conserved among different bacterial MetQ homologs. 11-14 The hydrogen bonds between the N238 side chain and the methionine α-amino and α-carboxyl groups appear to contribute significantly to the binding affinity, since mutation of this residue to alanine weakens the binding of l -methionine by over 10 5 . Despite binding to the same site on MetQ, the detailed interactions for d - and l -methionine differ as reflected in the ∼10 4 -fold difference in Kd values. The origin of this difference in binding affinity is difficult to identify since the hydrogen-bonding network involving the α-amino and α-carboxyl groups is largely unchanged between the two structures, including the positioning of water molecules (Figure 3c). Short contacts to the methionine CB group are observed in both structures, with ∼3.3 å distances observed to the Y98 CO in the l -methionine structure, and to the Y81 OH groups in the d -methionine structure. The side chains of l - and d -methionine adopt distinct rotameric conformations for the N-Cα-Cβ-Cγ Cα-Cβ-Cγ-Sδ and Cβ-Cγ-Sδ-Cε torsion angles, which are approximately −175°, −175°, and −70° for l -methionine and +170°, −80°, and −60° for d -methionine the latter values correspond to −170°, +80°, and + 60°, respectively, for l -methionine. Hence, while l - and d -methionine exhibit distinct “ttm” and “tpp” rotamers, these rotamers are found in comparable abundances in proteins (∼6% each 19 ), suggesting that the side chain torsion angle conformation does not contribute significantly to the differences in binding affinity (for reference, the most common methionine rotamer is “mmm” with a 20% frequency). Consequently, while there are differences in the details of the binding interactions between l - and d -methionine to NmMetQ, a qualitative evaluation does not identify any obvious feature as dominating the differences in binding affinities for these two ligands.

The higher affinity of ATP-bound MetNI for ligand-free MetQ is not surprising given the high affinity of MetQ for l -methionine (they co-purify) and the necessity to dissociate the ligand from MetQ for transport to occur.

Three distinct conformational states have been characterized to date for MetQ: one liganded and two ligand-free forms. The findings of a recent single molecule FRET study illustrating the conformational richness of SBPs, and the role of conformational dynamics in substrate transport, 20 suggests that additional conformations of MetQ may be identified, perhaps for larger methionine derivatives with modified amino or carboxyl groups, 1-4 or in complexes with different states of the MetNI transporter. This study provides a foundation for addressing the coupling between the conformation of MetQ, the affinities of MetQ for MetNI and methionine derivatives that is at the heart of the specificity of ligand transport by the methionine transporter an important next step is to define the kinetics of these interactions and deciphering how MgATP hydrolysis is coupled to ligand translocation.


We thank S. R. Adams and J. Babendure for their comments on the manuscript. This work was supported by a National Institutes of Health grant to R.Y.T. and postdoctoral fellowship to A.Y.T, a grant from the Alliance for Cellular Signaling (to J.Z. and R.Y.T.) and the Howard Hughes Medical Institute. J.Z. is supported in part by a postdoctoral fellowship from La Jolla Interfaces in Science and Burroughs Wellcome Fund, and R.E.C. is supported in part by a postdoctoral fellowship from the Canadian Institutes of Health Research.

Fig. 3

(a) Excitation and (b) emission steady-state anisotropies of Cerulean. The high value of emission anisotropy, > 0.3 , suggests limited flexibility of the fluorophore within the protein structure. The rapid decrease of the excitation anisotropy is consistent with FRET between aromatic amino acids and the fluorescent protein chromophore.

Chapter 6 - Genetically Encoded Probes for Measurement of Intracellular Calcium

Small, fluorescent, calcium-sensing molecules have been enormously useful in mapping intracellular calcium signals in time and space, as chapters in this volume attest. Despite their widespread adoption and utility, they suffer some disadvantages. Genetically encoded calcium sensors that can be expressed inside cells by transfection or transgenesis are desirable. The last 10 years have been marked by a rapid evolution in the laboratory of genetically encoded calcium sensors both figuratively and literally, resulting in 11 distinct configurations of fluorescent proteins and their attendant calcium sensor modules. Here, the design logic and performance of this abundant collection of sensors and their in vitro and in vivo use and performance are described. Genetically encoded calcium sensors have proved valuable in the measurement of calcium concentration in cellular organelles, for the most part in single cells in vitro. Their success as quantitative calcium sensors in tissues in vitro and in vivo is qualified, but they have proved valuable in imaging the pattern of calcium signals within tissues in whole animals. Some branches of the calcium sensor evolutionary tree continue to evolve rapidly and the steady progress in optimizing sensor parameters leads to the certain hope that these drawbacks will eventually be overcome by further genetic engineering.


Deubiquitinating enzymes (DUBs) are proteases that fulfill crucial roles in the ubiquitin (Ub) system, by deconjugation of Ub from its targets and disassembly of polyUb chains. The specificity of a DUB towards one of the polyUb chain linkages largely determines the ultimate signaling function. We present a novel set of diubiquitin FRET probes, comprising all seven isopeptide linkages, for the absolute quantification of chain cleavage specificity of DUBs by means of Michaelis–Menten kinetics. Each probe is equipped with a FRET pair consisting of Rhodamine110 and tetramethylrhodamine to allow the fully synthetic preparation of the probes by SPPS and NCL. Our synthetic strategy includes the introduction of N,N′𠄋oc‐protected 5�rboxyrhodamine as a convenient building block in peptide chemistry. We demonstrate the value of our probes by quantifying the linkage specificities of a panel of nine DUBs in a high‐throughput manner.

Ubiquitin (Ub), a 76 amino acid protein, is a post‐translational modifier that is crucial for a wide range of cellular processes, including protein degradation, trafficking, and signaling.1 Ub is generally attached via its C‐terminal carboxylate to the side𠄌hain amine of a lysine residue in the target protein, thereby forming an isopeptide bond. Target proteins are frequently modified with a polyUb chain, in which multiple Ub modules are successively linked at the N terminus (linear polyUb) or any of the seven internal lysine residues (isopeptide‐linked polyUb: K6, K11, K27, K29, K33, K48, and K63). The type of polyUb chain largely determines its signaling function.1

Ubiquitination is mediated by the concerted action of three enzymes, E1 (activating), E2 (conjugating), and E3 (ligase), the particular combination of which provides specificity for the protein target or polyUb chain topology. Removal of Ub from its targets and disassembly of polyUb chains are catalyzed by deubiquitinating enzymes (DUBs). About 100 human DUBs have been identified2 some exhibit Ub linkage specificity. DUB action can rescue proteins from proteasomal degradation and alter Ub signaling functions through chain remodeling in a linkage‐specific manner.1 The synthesis of diubiquitin (diUb) has made it possible to study processing by DUBs.3 In order to determine specificity, a DUB can be incubated with either a native diUb molecule4 or with a diUb activity�sed probe5 of a given linkage. However current methods do not allow fast and absolute quantification of DUB linkage specificity, and furthermore cannot separate this specificity into binding affinity and catalytic turnover rate (K m and k cat, respectively, in Michaelis–Menten kinetics).

The application of FRET pairs has proved useful in the study of DUB activity, Ub chain conformation, and Ub‐interacting proteins.6 In order to investigate chain cleavage specificity across all isopeptide linkages, we developed a full chemical synthesis of all seven isopeptide‐linked diUb FRET pairs. These pairs carry a novel dye‐pair suitable for FRET and compatible with solid phase peptide synthesis (SPPS). We determined K m and k cat values of linkage‐specific DUBs that are used in Ub chain restriction analysis,7 in order to obtain insight into their catalytic action.

In the FRET�sed assay (Figure  1 ) the reagents consist of two Ub modules, one equipped with a donor fluorophore and the other with an acceptor these are specifically linked by a native isopeptide bond to each of the seven lysine residues. We reasoned that the best position for fluorophore attachment would be the N termini of both Ub modules, because the distance between the N termini ranges from 30 to 50 Å, based on available crystallographic data (Table S1 in the Supporting Information), an ideal distance for FRET. Because the fluorophores need to be compatible with all synthetic steps (see below), we developed a new FRET pair by using 5�rboxyrhodamine110 (Rho) as the donor and 5�rboxytetramethylrhodamine (TAMRA) as the acceptor. Fluorescein, the more commonly used FRET donor, was initially tried but proved incompatible with the desulfurization step in the final synthesis step (see below) and was therefore replaced by Rho. Upon addition of a DUB, the diUb FRET pair is cleaved, thereby resulting in loss of the FRET signal and hence an increase in donor emission.

Principle of the FRET�sed diUb cleavage assay. Upon cleavage of the diUb FRET pair by a DUB, the FRET signal is lost.

A major problem in the use of Rho (but also TAMRA) in SPPS (Scheme  1 𠂚) is that when Rho is attached to an amine in a globally side𠄌hain‐protected peptide, the 1�rboxylate moiety of Rho is in an open conformation and can react upon further extension of the peptide chain (Scheme  1 𠂚, 12). In addition, the coupling of Rho is generally difficult because of the poor solubility and intrinsic reactivity of the aniline moieties. We therefore prepared N,N′𠄋oc‐protected Rho. When this molecule is coupled to a peptide, the dual Boc protection locks the 1�rboxylate in the closed lactone form, thus making it unreactive (Scheme  1 𠂚, 34). We modified the method reported by Grimm and Lavis8 to prepare N,N′𠄋oc‐protected Rho 10 (Scheme  1 ). 5�rboxyfluorescein (5) was converted in four steps into ditriflate 8. Buchwald–Hartwig coupling with BocNH2 resulted in the formation of N,N′𠄋oc‐protected Rho (9). Use of ethyl ester protection of the 5�rboxylate allowed selective liberation of the 5�rboxylate without affecting the Boc groups, thereby resulting in 10. In contrast to unprotected Rho, 10 is very soluble in organic solvents, can easily be coupled under standard peptide coupling conditions, and can be prepared on a multi‐gram scale.

The seven diUb FRET pairs 17𠂚g were constructed by native chemical ligation (NCL) between Rho‐Ub‐thioester 14 and TAMRA‐Ub containing a γ‐thioLys building block3, 9 16 (Scheme  2 ). The individual Ub modules where synthesized by linear SPPS on hyper�id‐labile trityl resin.3 DiBoc‐protected Rho (10) was coupled to Ub1� (11) on resin to result in 12, which was subsequently cleaved from the resin under mild acidic conditions without affecting the global protection scheme. Methyl𠄃‐(glycylthio)‐propionate was coupled to the liberated C‐terminal carboxylate to give 13. Global deprotection under strong acidic conditions followed by cation exchange and RP‐HPLC purification gave Rho‐Ub‐thioester 14. It is of note that the Boc groups on Rho are concomitantly removed during the global deprotection, thereby restoring its fluorescent properties. TAMRA‐Ub modules containing γ‐thioLys on each of the respective lysine positions (16𠂚g) were prepared by coupling the 5�rboxy isomer of TAMRA to the Ub1� polypeptides 15𠂚g, followed by global deprotection and purification (Figure S1 in the Supporting Information). Methionine𠄁 was replaced by the isostere norleucine to prevent oxidation of the thioether moiety. NCL reactions between 14 and 16𠂚g, followed by desulfurization under radical conditions,10 purification by RP‐HPLC, and gel filtration gave the final seven diUb FRET pairs 17𠂚g in good yield and purity.

The purities of 17𠂚g were confirmed by LCMS analysis (Figure  2 𠂚, B, and Supporting Information) and gel analysis (Figure  2 𠂜). Upon excitation at 466 nm, the emission spectra of the diUb FRET pairs and Rho‐Ub and TAMRA‐Ub revealed that all seven molecules show a clear FRET signal (Figure  2 𠂝). We performed fluorescence lifetime imaging microscopy (FLIM) to determine FRET efficiencies of all the FRET pairs (Figure  2 𠂞, Table S3) these were found to be 0.45𠄰.60, depending on the linkage, thus demonstrating efficient FRET in all these molecules.

Characterization of diUb FRET pairs 17𠂚g. A)𠁚nalytical HPLC and B) MS of Lys6‐linked diUb FRET pair 17𠂚. C) SDS‐PAGE analysis. D)𠁞mission spectra recorded at λ ex=466 nm. E)𠁟RET efficiencies (E) determined by FLIM.

DUB‐mediated cleavage of our new diUb FRET pairs was first assessed by incubation with USP7 and OTUD2, two well‐studied DUBs from the two largest DUB families and for which cleavage of unlabeled diUbs has been reported. Reactions were analyzed by SDS‐PAGE (Figures S2 and S3) which showed that the diUb selectivity of both DUBs was in good agreement with reported data.4a,4b We then incubated OTUD2 with Lys11‐ and Lys27‐linked diUb FRET pairs (17𠂛 and 17𠂜, respectively) and the corresponding unlabeled diUbs. We also included a 1:1 mixture of the FRET pair and the unlabeled diUb for both linkages. SDS‐PAGE analysis (Figures  3 𠂚, S4 and S5) showed that both the FRET pair and the unlabeled diUb substrates were equally processed, thus we concluded that the attached fluorophores do not affect DUB activity.

A) SDS‐PAGE analysis of Lys11‐linked diUb cleavage. OTUD2 was incubated with unlabeled diUb, FRET pair 17𠂛, or a 1:1 mixture samples were taken after 10, 30, 60, and 180 min. B) Michaelis–Menten kinetics of TRABID for Lys29‐, Lys33‐ and Lys63‐linked diUb, as determined by the FRET assay.

We next applied our FRET reagents for the quantification of diUb linkage‐specificity for nine DUBs derived from three different DUB families each was shown to display a distinct specificity (Table  1 ).4b We incubated the DUBs with all diUb FRET pairs at a fixed concentration (0.5 μ m, to keep initial fluorescence constant for all samples) with an increasing concentration of the unlabeled diUb (0� μ m ). The enzyme concentration was chosen such that the reaction proceeded linearly for at least 20 min (Supporting Information). The total amount of processed diUb was then determined by monitoring the increase in donor fluorescence over time from this the rates of initial velocity were calculated and fitted to the Michaelis–Menten equation (Figure  3 𠂛 and Supporting Information), from which K m and, k cat were determined.

Table 1

Kinetic characterization of DUBs that are used in Ub chain restriction analysis7 for the diUb FRET pairs 17𠂚g. [a]

DUBLinkage K m [μ m ] k cat [s 𢄡 ] k cat/K m [ m 𢄡  s 𢄡 ]
AMSHLys63 45.40.002759
AMSH* [b] Lys632.40.1770�
OTUB1* [b] Lys48 38.60.6617�
Lys11 52.40.0085162
Lys63 57.90.0061105
TRABIDLys29 40.10.0531317
Lys63 54.00.034627
OTUD2Lys11 87.94.450�

[a] Values in italics were obtained by extrapolation beyond the highest substrate concentration. [b]�tivated versions of AMSH and OTUB1 were created by fusing them to their activators (STAM and UBE2D2, respectively).11

Table  1 shows the data for all combinations of DUB and diUb substrate for which activity could be measured. Overall, the individual diUb linkage types cleaved by each DUB were consistent with published qualitative data.4 As expected from earlier findings, the unspecific DUB USP21 showed similar activity towards most linkages.4a The virus�rived DUB vOTU, which is also considered to be unspecific, showed some interesting results: the catalytic efficiency (k cat/K m ) for Lys6 was two times higher than for Lys48, and four times higher than for Lys11 and Lys63 diUbs this can largely be attributed to differences in k cat rather than K m . Another interesting result was for AMSH. In agreement with earlier findings, this DUB had absolute specificity for Lys63 diUb,4c although the overall efficiency was rather low. Remarkably, the recently reported fusion of AMSH with its natural activator STAM211 resulted in a more than 1000𠄏old increase in catalytic efficiency, which can be attributed to increases in both affinity and catalytic turnover. Taken together, these data show that our quantitative assessment of the DUB linkage specificity is in accordance with reported data and that new insights can be obtained from the kinetic parameters.

In summary, the set of all seven isopeptide‐linked diUb FRET pairs allows absolute quantification of DUB linkage specificity. Our synthetic strategy, which includes a convenient N,N′𠄋oc‐protected Rho building block, allows efficient preparation of these reagents in large quantities. The assay requires low amounts of material, can easily be automated, and can be used in high‐throughput small‐molecule screening or for the assessment of (di)Ub binding domains.6c Overall, we believe that our diUb FRET probes will be of great value in ongoing efforts to crack the ubiquitin code and that the FRET pair presented here will facilitate FRET‐pair synthesis in general.

Author summary

There is abundant evidence that insufficient energy, or energy failure, contributes to the pathophysiology of many inherited and degenerative diseases, cancer, and aging. However, we understand little about how cellular energy levels are regulated. To begin to address this gap, we developed a screening approach that uses a fluorescent biosensor to measure relative levels of ATP in individual cells and used this approach to identify those genes that are most essential to maintaining energy levels. We screened approximately 2,200 genes and found that mitochondrial ribosomal proteins are particularly critical in maintaining energy levels and enabling cell growth. We also used this approach to identify genetic targets that could be amenable to an energy-based therapy via supplementation with the mitochondrial cofactor coenzyme Q10 (CoQ10). These included CoQ10 biosynthetic genes associated with human disease as well as a subset of genes not previously linked to CoQ10 biosynthesis. Our screening paradigm thus reveals mechanisms by which cellular energy levels are controlled. It can also be used to identify genetic defects that will be responsive to energy-based therapies. Application of this approach may additionally enable powerful screens for other key metabolites to further advance a genome-scale understanding of the regulation of metabolism.

Citation: Mendelsohn BA, Bennett NK, Darch MA, Yu K, Nguyen MK, Pucciarelli D, et al. (2018) A high-throughput screen of real-time ATP levels in individual cells reveals mechanisms of energy failure. PLoS Biol 16(8): e2004624.

Academic Editor: Rong Tian, University of Washington, United States of America

Received: October 30, 2017 Accepted: July 26, 2018 Published: August 27, 2018

Copyright: © 2018 Mendelsohn et al. This is an open access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.

Data Availability: All relevant data are within the paper and its Supporting Information files. Specifically, S1 Table contains screen data for all genes in all conditions.

Funding: Joan and David Traitel Family Trust. KN, BAM, NKB, MAD, KY, MKN, and MN. The funder had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript. Betty Brown’s Family. KN, BAM, NKB, MAD, KY, MKN, and MN. The funder had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript. Hagar Family Foundation. JLN and DP. The funder had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript. Burroughs Wellcome Fund Medical Scientist Career Award. KN, BAM, MAD, and KY. The funder had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript. Pediatric Scientist Development Program (grant number K12-HD000850). BAM. The funder had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript. NIH (grant number R01 NS091902, 5P30 NS069496, U54CA196519, R01 DA036858, U01 CA168370, DP2 GM119139, RR018928, S10-RR028962, K99/R00 CA204602). R01 NS091902: KN, 5P30 NS069496: KN, BAM, NKB, MAD, KY, MKN, and MN, U54CA196519: JLN, R01 DA036858: LG, MAH, and MK, U01 CA168370: LG, MAH, and MK, DP2 GM119139: MK, S10-RR028962: Gladstone flow cytometry core: LG, K99/R00 CA204602. The funder had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript. St. Baldrick’s Scholar Award. JLN and DP. The funder had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript. Howard Hughes Medical Institute. LG, MH, and MK. The funder had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript. CFAR grant (grant number P30-AI-027763). Gladstone flow cytometry core. The funder had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript. James B. Pendelton Charitable Trust. Gladstone flow cytometry core. The funder had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.

Competing interests: The authors have declared that no competing interests exist.

Abbreviations: 2DG, 2-deoxyglucose ATPbasal, ATP levels under basal conditions ATPgly, ATP levels under glycolytic conditions ATPresp, ATP levels under respiratory conditons CFP, cyan fluorescent protein CoQ10, coenzyme Q10 COS, CV-1 (simian) in origin, and carrying the SV40 genetic material CRISPRi, clustered regularly interspaced short palindromic repeats interference dCas9, dead CRISPR-associated protein 9 ECAR, extracellular acidification rate FACS, fluorescence-activated cell sorting FBS, fetal bovine serum FCCP, carbonyl cyanide-4-(trifluoromethoxy)phenylhydrazone FRET, fluorescence resonance energy transfer GROWTHbasal, growth under basal conditions GROWTHglyc, growth under glycolytic conditions GROWTHresp, growth under respiratory conditions HGNC, HUGO Gene Nomenclature Committee HPLC, high-performance liquid chromatography KRAB, Kruppel-associated box MELAS, mitochondrial encephalopathy, lactic acidosis, and stroke-like episodes MICOS, mitochondrial contact site and cristae organizing system mVenus, monomeric Venus NDUFS4, NADH:ubiquinone oxidoreductase subunit S4 OCR, oxygen consumption rate PVDF, polyvinylidene fluoride qRT-PCR, quantitative real-time reverse transcription PCR RNAi, RNA interference ROS, reactive oxygen species sgRNA, single guide RNA shRNA, short hairpin RNA TCA, tricarboxylic acid Tom20, translocase of outer membrane 20 kDa subunit

Author information

Matthew Y. Tang and Marta Vranas: These authors contributed equally to this work


Department of Neurology and Neurosurgery, McGill Parkinson Program, Neurodegenerative Diseases Group, Montreal Neurological Institute, McGill University, Montréal, Québec, Canada H3A 2B4

Matthew Y. Tang, Andrea I. Krahn, Shayal Pundlik & Edward A. Fon

Department of Pharmacology and Therapeutics, Groupe de Recherche Axé sur la Structure des Protéines, McGill University, Montréal, Québec, Canada H3G 1Y6

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