Fluorescent Probes and Quenchers in Studies of Protein Folding and Protein-Lipid Interactions

Abstract

Fluorescence spectroscopy provides numerous methodological tools for structural and functional studies of biological macromolecules and their complexes. All fluorescence-based approaches require either existence of an intrinsic probe or an introduction of an extrinsic one. Moreover, studies of complex systems often require an additional introduction of a specific quencher molecule acting in combination with a fluorophore to provide structural or thermodynamic information. Here, we review the fundamentals and summarize the latest progress in applications of different classes of fluorescent probes and their specific quenchers, aimed at studies of protein folding and protein-membrane interactions. Specifically, we discuss various environment-sensitive dyes, FRET probes, probes for short-distance measurements, and several probe-quencher pairs for studies of membrane penetration of proteins and peptides. The goals of this review are: (a) to familiarize the readership with the general concept that complex biological systems often require both a probe and a quencher to decipher mechanistic details of functioning and (b) to provide example of the immediate applications of the described methods.

Graphical Abstract

Fluorescence spectroscopy provides numerous methodological tools for structural and functional studies of biological macromolecules and their complexes. However, studies of complex systems often require an additional introduction of a specific quencher molecule acting in combination with a fluorophore to provide structural or thermodynamic information. In this review, we outline the fundamentals and summarize the latest progress in applications of different classes of fluorescent probes and their specific quenchers, aimed at studies of protein folding and protein-membrane interactions

1. Introduction

Fluorescence spectroscopy is a powerful tool to study the structure and dynamics of proteins and other biological macromolecules.^[1-2] Superb sensitivity of fluorescence measurements, ensured by the high signal-to-noise ratio due to the spectral Stokes shift, provides important advantages over other techniques of optical spectroscopy.^[3-5] Rapid development of the high-resolution fluorescence spectroscopy methods is accompanied by the continued progress in a design of novel classes of fluorophores, including near infrared (NIR)^[6-7] and two-photon absorbing organic dyes, as well as genetically encoded fluorescent proteins.^[8] In recent years, several excellent reviews have been published on this topic,^[9-10] focusing on the development and applications of fluorescent probes and labels.^[11-16] In this review, we summarize progress in applying different classes of fluorescent probes and quenchers, resulting in methodological advances in studies of protein structure and folding, and protein-membrane interactions.

2. Applications of Intrinsic and Extrinsic Environment-Sensitive Dyes as a Tool for Monitoring Protein Binding, Refolding, and Membrane Interactions

Intrinsic protein fluorescence, usually originating from aromatic amino acids such as tryptophan, tyrosine, and phenylalanine, has long been used by biophysics for studying protein stability, misfolding and aggregation.^[3,17] Given latest developments in site-selective labeling, we felt it would be prudent to expand the term “intrinsic” fluorescent probe to include other fluorophores, integrated into the protein during its biosynthesis, such as natural and non-natural amino acids (Scheme 1), and chimeric fluorescent proteins. Here, we also classify synthetic probes covalently attached to biomacromolecules post-translationally (Schemes 2-3) as intrinsic fluorescent labels. Thus, the main feature (and advantage) of intrinsic fluorescent probes is their ability to be introduced selectively at certain sites at proteins.^[18-21]

Scheme 1.

Fluorescent amino acids. Native amino acids: phenylalanine (Phe), tyrosine (Tyr), tryptophan (Trp). Unnatural fluorescent amino acids; 5-fluorotryptophan (5F-Trp), 7-azatryptophan (7ATrp), 2-amino-3-(5-(dimethyl-amino)-naphthalene-1-sulfonamide)propanoic acid (dansylalanine), (7-hydroxy-coumarin-4-yl)ethyl-L-glycine (7-HCaa), Anap (3-(6-acetylnaphthalen-2-yl-amino)-2-aminopropanoic acid),^[28] N-[4-(N,N-dimethylamino)-phthalimidyl]-L-alanine (4-DMAP).^[29]

Scheme 2.

Thiol-reactive probes for site-selective labelling: monobromobiname, BODIPY FL N-(2-aminoethyl) maleimide, IANBD ester (N-((2-(iodoacetoxy) ethyl)-N-methyl)-amino-7-nitrobenz-2-oxa-1,3-diazole), 7-diethylamino-3-[N-(2-maleimido-ethyl)-carbamoyl] coumarin (MDCC) and 5-(N-iodoacetyl-aminoethyl)-naphthylamine-1-sulfonic acid (IAEDANS).

Scheme 3.

Structure of some thiol-reactive AlexaFluor dyes.

In contrast, we define “extrinsic” probes as those not attached to biological macromolecules covalently, but associate with the target non-selectively via hydrophobic or electrostatic interactions.^[22] These dyes can be applied for measuring surface hydrophobicity, probing active sites of enzymes, monitoring protein-ligand interactions, and detecting protein aggregation or fibrillation. Such extrinsic fluorescence probes often operate by an “on/off” principle, so that they are hardly fluorescent in an aqueous environment, however, they become highly fluorescent upon adsorption into hydrophobic binding sites within proteins.^[17] To strengthen the binding interactions of extrinsic probes with protein or lipid membranes, they can further be functionalized by long alkyl tails or polar functional moieties (see Scheme 5 for more detail).

Scheme 5.

Fluorescent probes, fatty acids and amines: dibutylamino stilbazolium butylsulfonate (di-4-ASPBS), 1-[6-(dimethylamino)naphthalen-2-yl]propan-1-one (Prodan), 6-lauroyl-2-(dimethylamino)-naphtalene (Laurdan), tryptophan octyl ester (TOE), 4-(hexadecylamino)-7-nitrobenz-2-oxa-1,3-diazole (NBD-C₁₆),1,1′-dioctadecyl-3,3,3,3′-tetramethyl-indocarbocyanine (DiI-C₁₈).

2.1. Fluorescent Amino Acids as Intrinsic Probe

Native amino acids, such as phenylalanine, tyrosine and tryptophan, are often part of peptides and proteins, so that they are used as an intrinsic fluorescent probe for studying protein structure (Scheme 1).^[23-24] Their unique photophysical properties are utilized for studying the structure and folding of proteins without disrupting their native folds.^[3,25] Intrinsic protein fluorescence can be used to look at the impact of mutations or other protein modifications on conformational changes, measure enzyme activities, or investigate the binding of ligands.^[26-27]

An emission spectrum and intensity of a Trp fluorophore change depending on the environment polarity, which makes its fluorescence a valuable indicator of the conformational state of a protein and its folding process.^[5,30] For example, if Trp residues are deeply buried in the hydrophobic pocket of protein, they exhibit the blue-shifted fluorescence compared to those exposed to the aqueous environment^[3,26,31]

Using a tryptophan as an intrinsic fluorescence reporter has been instrumental in studying membrane interactions of proteins and peptides.^[5,31-32] Numerous examples of utilizing Trp fluorescence to study structure, kinetics and thermodynamics of peptide-lipid interactions exist.^[32-45]

One of the advantages of using Trp as a probe is that the conformational heterogeneity of the environment can be studies by simultaneously monitoring the position of the emission spectrum and its widths.^[31] An illustrative example of the use of the intrinsic Trp fluorescence for examining the protein-membrane interactions is the case study of the membrane-proximal external region (MPER) of the membrane-interacting C-terminal domain of the HIV-1 gp41 fusion protein.^[46] MPER is a Trp-rich peptide containing five tryptophan residues (Figure 1a). The shape and position of the Trp fluorescence spectrum can be used to interpret its location within the lipid membrane. Figure 1b (blue) shows the red-shifted Trp fluorescence of MPER, which is indicative of the interfacial (IF) location of the peptide, when the peptide is added externally to preformed LUVs. However, the blue-shifted emission indicating a much more hydrophobic environment of the tryptophan residues is characteristic of the transmembrane (TM) conformation of the MPER peptide, which is populated when vesicles are formed in the presence of the peptide (Figure 1b, red).^[46]

Figure 1.

(a) Amino acid sequence of the membrane-proximal external region (MPER) of membrane-interacting C-terminal domain of the HIV-1 gp41 fusion protein. (b-c) Tryptophan fluorescence of the MPER peptide in various membranous environment: the MPER peptide in solution (black), added to preformed POPC vesicles (MPER + LUV, blue) and coextruded with POPC vesicles (MPER/LUV, red). (d-g) Fluorescence titration of the unlabeled MPER peptide (d-e) and the NBD-labeled W666C mutant of the MPER peptide (f-g) with POPC LUV in 50 mM phosphate buffer solution at pH 8.^[46]

Spectral heterogeneity of the MPER peptide can be further analyzed by fitting the fluorescence spectra with a log-normal distribution and plotting the spectral width as a function of the spectral maximum (Figure 1c). For the MPER in solution and MPER + LUV samples, some spectral broadening was observed, which is accompanied by an upward deviation from the linear baseline that can be drawn through position-width points corresponding to spectral classes I, II, and III identified by Burstein and coworkers.^[47-48] The relatively large spectral width of MPER/LUV indicate substantial heterogeneity in the environments of the five Trp residues of MPER, which allowed us to suggest a TM rather than an IF membrane orientation.^[46] The fluorescence titration experiment shown in Figures 1d and 1e allowed us to estimate the free energy of partitioning (ΔG) of MPER with POPC LUV, which was found to be −8.1 kcal/mol. Notably, the replacement of W666 with the Cys-attached NBD probe did not perturb the overall strength of the membrane binding of MPER with the POPC bilayer, as seen by the good agreement with the lipid titration experiment shown in Figure 1f and 1 g, respectively.^[46]

Although natural fluorescent amino acids have become a convenient tool in protein biophysics, their shortcoming is short-wavelength emission and low-intense fluorescence. Therefore, some unnatural fluorescent amino acids with red-shifted absorption and bright emission have recently been designed (Scheme 1).^[10,49-55] Over the past decade, several approaches based on the direct covalent incorporation of fluorescent unnatural amino acids into proteins have been suggested.^[28,56-60] An example is dansylalanine (Scheme 1), which was genetically encoded in Saccharomyces cerevisiae by using an amber nonsense codon and corresponding orthogonal tRNA/aminoacyl-tRNA synthetase pair.^[56] Noncanonical amino acid Anap has been used as an energy donor in FRET-based experiments to probe short-range intra-protein distances in maltose-binding proteins.^[61] 7-Hydroxycoumarin moiety in 7-HCaa forms a FRET pair with native Trp residues, enabling monitoring of substrate-induced conformational changes in hexokinases.^[62] Environment-sensitive fluorescence of Anap has also been utilized to monitor the dynamics of internal pore opening in K_v channels^[63] and voltage-dependent motion of the catalytic region of voltage-sensing phosphatase.^[64] Dansylalanine has an environmentally sensitive fluorophore, which after selective incorporation into human superoxide dismutase, was used to monitor protein unfolding.^[56]

2.2. Thiol-Reactive Probes for Site-Selective Protein Labeling

Intrinsic tryptophan fluorescence has some disadvantages because proteins usually have multiple tryptophan residues, which complicate the interpretation of the complex fluorescence from multi-Trp proteins due to the environment-sensitive nature of individual Trp residues. Site-directed fluorescence labeling of protein by organic dyes appears to be a promising alternative to intrinsic protein fluorescence.^{[10,19,21,53,65]} Selective protein labeling with thiol-reactive fluorescent probes, such as Bimane, BODIPY, NBD and AEDANS (Scheme 2), revealed increasing promise and is widely used.^{[10,20,50,66-69]}

The common site-specific labeling protocols utilize commercially available fluorophores with a succinimide ester or maleimide reactive group that targets the primary amino or thiol groups. Like Trp, most labeling fluorescent probes display high sensitivity to environmental polarity. The small-sized fluorophores, such as bimane and NBD with a short linker, are small enough and do not perturb the native structure of the protein. Synthetic fluorescence probes have a high molar absorption coefficient (ε > 25,000 M⁻¹cm⁻¹) and bright emission with tunable peak position.^[50] Today, organic fluorophores can be easily synthesized, and hundreds of fluorescent probes are available commercially.^[11,70]

Applications associated with single-molecule fluorescence-correlation spectroscopy (FCS) require that fluorescent dyes have reasonably high solubility in water, low environment-sensitivity (including pH-independent fluorescence) and the absence of unwanted interactions between a dye and peptide after conjugation.^[71] For these purposes, a series of AlexaFluor dyes (Scheme 3) have been designed to accomplish most these requirements.^[72-74]

Advantages of FCS applications are focused on molecular interactions resulting in large changes in hydrodynamic volume, occurring upon binding between ligand and receptor, peptide and liposome, yielding large changes in diffusion correlation times of a fluorescent probe.^[75-77] For example, membrane interactions of AlexaFluor-labeled peptide and proteins were measured using FCS for WALP23 and WALP27 peptides.^[78]

Popular donor-acceptor pairs for a single-molecule FRET are AlexaFluor-488/AlexaFluor-594,^[79-80] AlexaFluor-488/AlexaFluor-647,^[81-82] AlexaFluor-555/AlexaFluor-647,^[83-86] respectively. AlexaFluor dyes are also used in FRET experiments in combination with other fluorophores.^[87-88]

2.3. Biosynthetically Incorporated Fluorescent Proteins and Lanthanide Binding Sites as an Advanced Tool for Biophysical and Cellular Studies

Biosynthetically incorporated fluorescent proteins (FPs) have revolutionized the molecular biology by enabling protein labeling in the cells.^[89-92] FPs can be appended onto existing proteins by using recombinant techniques, thus allowing the endogenous expression of fluorescent protein chimeras.^[93] Genetically encoded FPs with widely diverse and tunable colors have profoundly renewed and quickly revolutionized biological cell imaging (Scheme 4).^[94-97]

Scheme 4.

Structure of some popular FPs: Enhanced cyan fluorescent protein (ECFP, PDB 1CV7), wild-type green fluorescent protein (GFP, PDB 4KW4), enhanced yellow fluorescent protein (EYFP, PDB 1YFP), mCherry red fluorescent protein (mCherry, PDB 6YLM). The secondary structure of a FP carrier is shown as β-barrel and a chromophore is shown in vdW representation. For clarity, the structure of the chromophore alone is given by a licorice model. Excitation (Ex), emission (Em) maxima and fluorescent quantum yield (Qy) are given for comparison.

Green Fluorescent Protein (GFP) from the Aequorea victoria jellyfish contains the prototypical p-hydroxy-benzylidene-imidazolinone chromophore, located in the central helix and surrounded by 11 β-strands. Early experimental and theoretical studies of GFP have revealed a complex interplay between a relatively simple chromophore formed after specific biosynthesis, and the spatial and dynamic organization of its protein carrier (Scheme 4).^[94,98-99]

Today, through mutation and selection, many GFP variants were designed that differ in protein and chromophore structure, intending to tune their absorbance and emission characteristics. Popular variants of more enhanced and stable GFP were produced, such as cyan, and yellow fluorescent proteins (ECFP, EYFP, etc.). While FPs are primarily being used for in vivo labeling of cells, some advanced FRET-based FP sensors and indicators were also developed.^[100]

Genetically encoded FPs have the disadvantage of their large size, so they may alter the structure and dynamics of tagged protein, which outweighs their advantageous fluorescence features. Therefore, using much smaller genetically encoded tags—the lanthanide binding tag (LBT) is a promising alternative. This approach pioneered a novel modification of FRET, as a technique to measure distances at a molecular level, referred to as lanthanide-based resonance energy transfer (LRET).^[101] LRET requires chelating molecules that bind lanthanide ions and also protect them from collisional quenching from the aqueous media^[102]. In this setup, the lanthanide ions are used as an energy donor, transferring excitation to an acceptor.^[101]

Application of LRET to Shaker potassium channels expressed in live Xenopus oocytes has been reported.^[103] Two small genetically encoded tags, the LBT and the hexa-histidine (6-His) tag, were introduced to the potassium channel without distorting the protein’s function. The 6-His tag is capable of binding transition metal ions, such as Ni²⁺, Co²⁺, and Cu²⁺ (Figure 2a). Quenching of luminescence decay of LBT-bound Tb³⁺ donor was observed in the presence of the acceptor 6-His tag complexed with either Ni²⁺ or Cu²⁺, which allows probing distance changes in Shaker potassium channels (Figure 2b-c).^[103]

Figure 2.

(a) Schematic representation of the Shaker K1 channel monomer illustrating the positions of the LBT (between transmembrane, TM, segments S3 and S4) and of the four 6-His tags tested. Displayed is the voltage sensing domain, comprised of TM segments S1-S4, and the pore region, segments S5-pore loop-S6, of the adjacent subunit. Inset shows the LBT site with a bound Tb³⁺ ion. (b-c) Tb³⁺ luminescence decay of Shaker channel constructs with the LBT in the S3-S4 loop with and without a 6-His tag close to S1 and S3 for Ni²⁺ and Cu²⁺, respectively. Adapted from^[103]. Copyright © 2007. The Biophysical Society. Published by Elsevier Inc. All rights reserved.

2.4. Extrinsic Environment-Sensitive Organic Fluorophores

Properties of the solvation shell modulate the spectroscopic behavior of environment-sensitive organic fluorophores. This phenomenon is referred to as solvatochromism and solvato-fluorochromism and describe the pronounced change in absorption or emission spectra following a change in the solvent polarity.^[4] Formation–disruption of hydrogen bonds with solvent molecules and solvent-dependent changes of dye geometry can strongly alter fluorescence intensity and lifetime.^[104-106] Dramatic fluorescence quenching may occur in water due to the formation by solute-solvent H-bonds.^[106] In addition, the solvent can influence the dye energetics, particularly the inversion of n* (non-fluorescent) and π* (fluorescent) energy levels. Comprehensive reviews of solvato-chromic and solvatofluorochromic dyes are given elsewhere.^{[3,15,107-108]}

Fluorescent probes functionalized with fatty acids are frequently used to study the structure, dynamics, and local polarity of biological membranes (Scheme 5).^[109] For instance, Prodan and Laurdan were used to monitor time-dependent solvent relaxations, allowing monitoring of local viscosity and polarity in a lipid bilayer,^[110-115] and discriminating cholesterol content in living cell membranes.^[116] Membrane-bound TOE is a good model for studying the intrinsic fluorescence of protein-membrane interactions.^[117-119] Fluorescent fatty amines NBD-C_n (n = 8–16) have been utilized to characterize phase transitions, lipid desorption and flip-flop in lipid membranes.^[120-121]

Hydrocarbon-tailored fluorophores are better alternative to noncovalent aromatic dyes,^[122-123] such as pyrene, DPH, ANS, as extrinsic probes for studying protein organization because their binding affinity are governed and tuned by the fatty acid chain. For instance, the association of a homologues series of fluorescent NBD-C_n probes with BSA was studied in an aqueous solution as a function of alkyl chain length n = 4,8,12,14,16. It was found that the binding free energy of NBD-C_n probes towards BSA was linearly dependent on the length of the alkyl chain.^[124]

2.5. Fluorescent Lipids

Fluorescent analogues of natural lipids have allowed to monitor membrane dynamics in different regions of a bilayer and various cellular activities^[125-129] as well as assist in studies of thermodynamics and kinetics of protein partitioning into the bilayers.^[38,130-132] Depending on a labeling position of a fluorescent moiety, they are divided into two main classes: (i) acyl chain labeled and (ii) head-group labeled (Scheme 6).^[125]

Scheme 6.

Fluorescent lipids:1-palmitoyl-2-(2-pyrenedecanoyl)-sn-glycero-3-phosphocholine (Pyr₁₀-PC), 1-palmitoyl-2-(12-[(7-nitro-2-1,3-benzoxadiazol-4-yl)amino]-dodecanoyl)-sn-glycero-3-phosphocholine (C₁₂-NBD-PC), 1-palmitoyl-2-{6-[(7-nitro-2-1,3-benzoxadiazol-4-yl)amino]hexanoyl)-sn-glycero-3-phosphocholine (C₆-NBD-PC), 2-(4,4-difluoro-5-butyl-4-bora-3a,4a-diaza-s-indacene-3-octanoyl)-1-hexadecanoyl-sn-glycero-3-phosphocholine (BODIPY-PC), 1,2-dipalmitoyl-sn-glycero-3-phosphoethanolamine-N-(7-nitro-2-1,3-ben-zoxa-diazol-4-yl) (NBD-PE), 1,2-dipalmitoyl-sn-glycero-3-phosphoethanol-amine-N-(lissamine rhodamine B sulfonyl) (ammonium salt) (rhodamine-PE), Texas Red 1,2-dihexadecanoyl-sn-glycero-3-phosphoethanolamine (Texas Red PE).

Lipophilic fluorescent lipids are effective probes for cell membrane confocal fluorescence imaging.^[133] Therefore, head-group-labeled lipids are commonly used as fluorescent dopants to lipid bilayers.^[134] A charged phosphate linker between a fluorescent moiety and the phospholipid anchor favors the precise location of a fluorophore in the plasma membrane of living cells.^[125,135] The typical protein-lipid FRET-binding assay consists of a donor dye-labeled protein (Trp, NBD, AlexaFluor-488) and a lipid membrane doped with an acceptor dye (Rhodamine-PE, Dansyl-PE), so that the partitioning of the protein causes changes in a FRET signal between the probes.^[38,130,136] For example, Loura and coworkers used C₁₂-NBD-PC as a FRET donor to examine the ability of a model peptide (Lys6Trp or K6 W) to reorganize DPPC:DPPS bilayers.^[137]

An example of applying of probe-labeled lipids as acceptors in the FRET-based protein-membrane binding assay is shown in Figure 3. In this example, binding of the pH-low insertion peptide (pHLIP), containing two native Trp residues, to Dansyl-labeled LUV was monitored by FRET signal changes.^[38] pHLIP is a protein designed from the sequence of the C-helix of bacteriorhodopsin and its pH-dependent membrane interactions are associated with the protonation of Asp residues.^[138] pHLIP–P20G is a mutant (Figure 3a), lacking the helix-breaking proline in the middle of the hydrophobic stretch. In the absence of acceptor-labeled LUVs, pHLIP–P20G shows Trp emission only. Upon titrating the pHLIP solution with Dansyl-PE-LUVs, the peptide partitioning to membranes is monitored by the decrease in the intensity of the Trp donor peak accompanied by the corresponding increase in Dansyl emission at 520 nm (Figure 3b).

Figure 3.

Application of lipid-attached probes as FRET acceptors to the protein donor fluorescence to study membrane binding. (a) Scheme of FRET-binding assay, in which peptide binding to a bilayer is monitored by FRET occurring between a donor-dye-labeled pHLIP and Dansyl-PE acceptor-doped bilayer. (b) FRET measurements between Trp (donor) present in pHLIP–P20G and POPC-LUV containing 2% Dansyl-PE (acceptor) by gradual increasing LUV concentration.^[38]

2.6. Dual-Fluorescence ESIPT Probes

A promising class of environmentally sensitive dyes is based on a 3-hydroxyflavone (3HF) scaffold.^[15,139-144] The key feature of these probes is the excited-state intramolecular proton transfer (ESIPT) reaction,^[145-146] which occurs through an intramolecular hydrogen bond bridge between the 3-OH group and the flavonoid carbonyl oxygen (Figure 4a). ESIPT results in an extremely fast (k_ESIPT > 10¹²s⁻¹) phototautomerization from the normal state (N*) to the tautomer state (T*),^{[144,147-148]} observed experimentally as dual-band emission (Figure 4b).^[139,147] Importantly, the intensity ratio of normal-to-tautomer emission bands depends on the probe environment, which makes possible ratiometric fluorescence detection of local changes in the microenvironment. The environment-sensitive ESIPT phenomenon of 3HF probes has been utilized for monitoring solvent polarity,^{[139,149 150]} metal ion sensing^[144,151] pH.^[152-153]

Figure 4.

(a) Schematic presentation of the excited state intramolecular proton transfer (ESIPT) reaction in 3-hydroxyflavones (3HF). (b) Dual-band fluorescence spectra of 3-hydroxyflavone dye. (c) Dual-fluorescence fluorescent conjugates, nonnatural amino acids and alkyl-functionalized membrane probes based on 3HF-dyes: 6-acetamido-4′-(dimethylamino)-3-hydroxyflavone (MFL), 3-[2-(4-methoxyphenyl)-3-hydroxy-4-oxo-4H-chromen-6-yl]-$L$-alanine (M3HFaa), 3-[2-(2-furyl)-3-hydroxy-4-oxo-4H-chromen-6-yl]-L-alanine (3HCaa).

Numerous dual-fluorescence probes based on 3HF-scaffold have been designed as covalent labeling agents for proteins and peptides, non-natural fluorescent $L$-amino acids, and alkyltailored membrane probes (Figure 4c).^{[15,49,154-155]} Noncovalent 3HF probes have revealed good selectivity and sensitivity toward various biomolecules.^[50,156] 3HFs show a significant “turn-on” fluorescence effect upon selective binding to proteins, such as lysozymes^[157] and albumins.^[156,158] For instance, 3HCaa was incorporated through solid phase synthesis into the zinc finger domain of the HIV-1 nucleocapsid protein for studying the interaction with oligonucleotides.^[159] Membrane interactions of melittin and magainin 2 labeled by an MFL probe at the N-terminus were studied using changes in the dual fluorescence signal.^[160-161]

3HF-based probes are also promising fluorescent agent with a low perturbation impact on the lipid membrane physical state.^[162-165] Due to the low molecular mass (<500 Dalton) and low toxicity, some fluorescent 3HFs have been utilized for fluorescent cell imaging.^[166,168] In addition, they could be used as a fluorescent indicator of β-glucosidase activity.^[169-170] Comprehensive reviews of 3HF probe design, ESIPT-sensing principles and applications are available elsewhere.^[15,171-173]

3. Distance Mapping by a Long-Range Energy Transfer and Short-Range Fluorescence Quenching Techniques

Electronic excitation energy can be non-radiatively transferred between a pair of two different or identical dyes, one of which acts as a donor and another as an energy acceptor.^[1,3,174] Because energy transfer occurs from the excited donor, its fluorescence signal is reduced or quenched in the presence of an acceptor. Excitation energy transfer can occur through either dynamic or static mechanisms. Quantitative estimation of dynamic fluorescence quenching is well described by Forster^[175] and Dexter^[176] theories.

The Forster resonance energy transfer (FRET) mechanism, also known as the Coulomb mechanism, considers classical dipole-dipole interactions between the transition dipoles of the donor and acceptor. Fluorescence quenching by the Förster theory is extremely dependent on the donor-acceptor distance ($r$) and their relative orientation ($k^2$). FRET can operate at distances $r$ up to 100 Å, decreasing at a rate of 1/R⁶ (Equation 1-2).^[175]

$$k_{FRET}(r)=\frac{1}{{\tau }_D}{\left(\frac{R_0}{r}\right)}^6$$ (1)

$$E_{FRET}=1∕\left[1+{(r∕R_0)}^6\right]$$ (2)

The Dexter mechanism, known also as exchange or collisional energy transfer, is based on short-range interactions that depend on the spatial overlap of donor and quencher molecular orbitals.^{[174,176,179]} Therefore, the efficiency of Dexter-type energy transfer decreases with $e^{-r}$, where $r$ is quencher distance (Equation 3).

$$k_{DEXTER}(r)=KJ\text{exp}\left(\frac{-2r}{L}\right)$$ (3)

where $K$ is a specific orbital interaction constant, $J$ is the spectral overlap integral, $\text{exp}(2r∕L)$ is the orbital overlap, and $L$ is the van der Waals radii of the molecules.

Static fluorescence quenching occurs through the formation of a ground state complex. The donor and acceptor (quencher) species bind together to form a complex or an intramolecular dimer already in the electronic ground state.

In this Section, applications of FRET-binding assays, long-range FRET, and short-distance residue-to-residue FRET quenching are illustrated for studies of protein-membrane interactions, protein misfolding, and local rearrangement of protein structure.

3.1. Long-Range Forster Resonance Energy Transfer for Monitoring Protein-Lipid Interactions and Large-Scale Protein Refolding

The application of FRET is a common strategy to detect binding between two fluorescence-labeled particles.^{[93,174,180-181]} In this approach, a FRET ratio is changed because a donor probe transfers energy to an acceptor probe when they are within the Förster’s distance.^{[38,130-131,182]} This approach can be applied for protein-protein and protein-ligand binding.^[10,183] For instance, the effective Förster radius for Trp-Dansyl FRET is 21 Å, which allows for estimating protein-protein binding. The thermodynamics of helix-helix association in bacteriorhodopsin was studied using native and Dansyl-labeled-bacteriorhodopsin.^[34] The binding affinity of small molecules for Trp-containing proteins was determined using FRET between endogenous Trp residues in Streptavidin and the coumarin-derived fluorophore Pacific Blue (PB).^[184]

FRET is increasingly being used to determine distances, structures, and dynamics of proteins at the ensemble and the single-molecule level.^{[50 87 185-186]} Intramolecular FRET usually occurs within inter-dye distances from 30 up to 100 Å that is comparable to the dimensions of most biological macromolecules.^{[174-175,187-188]}

Figure 5 shows an example of FRET-based measurements of pH-dependent refolding of the anti-apoptotic protein Bcl-xL.^[189] The Bcl-xL construct was designed, so that the AlexaFluor-488 donor was attached at D189C site (yellow) and an N-terminally conjugated mCherry fluorescence protein served a FRET acceptor (magenta). The membrane insertion of Bcl-xL leads to its refolding and the release of its N-terminal BH4 helix.^[132] The release of the BH4 helix could be measured in the presence of 1TOCL:2POPC LUV by loss of FRET between an N-terminally conjugated mCherry fluorescent protein and the fluorophore AlexaFluor-488 (A488) (Figures 5a-c). These spectral changes indicate a loss of FRET between both fluorophores and suggest the increase in distance between donor and acceptor, attributed to the release of the N-terminal BH4 helix.^[132,189] The FRET efficiency, measured using single-molecule FRET (smFRET) as a function of pH, was characterized by a progressive shift of the FRET distributions to lower efficiencies (Figure 5d). These data suggest the presence of multiple stable intermediate conformations during the refolding/membrane insertion of Bcl-xL, each with characteristic FRET distance.^[189]

Figure 5.

Application of FRET measurements to study conformational changes in a protein selectively labeled with an Alexa probe and a conjugated Fluorescent Protein. (a) Bcl-xL construct labeled by AlexaFluor-488-mCherry FRET dyes. (b) Experimental FRET set-up is based on the presence of FRET when the mCherry acceptor is close to the AlexaFluor-488 donor and the lack of FRET in the refolded/inserted form of Bcl-xL due to the increase in distance between the donor-acceptor pair. (c) Steady state measurements show a progressive increase in donor A488 intensity at 518 nm accompanied by a decrease in the acceptor mCherry intensity at 605 nm as a function of pH. (d) The smFRET efficiency is characterized by a progressive shift of the distributions to lower FRET efficiencies as a function of pH.^[132,189]

3.2. Short-Range Fluorescence Quenching as a Ruler for Distance Mapping

Tryptophan-induced fluorescence quenching can map interactions within a protein and trace protein refolding in real time by monitoring changes in the distance-dependent quenching of specific fluorophores caused by the presence of a nearby Trp residue.^[21,190-191] Bimane fluorescence (Scheme 2) is quenched by Trp over relatively short distances (< 10 Å), offering a near-instant snapshot of a local protein conformation.^[18,68] This short-distance quenching technique has been used to study the structure and dynamics of close-contact interactions in an ion channel measured by the quenching of a covalently attached bimane fluorophore by a nearby Trp residue.^[192] The correlation between the enzymatic activity of TL lipase and its polarity-induced conformational changes has also been examined by Trp-bimane quenching.^[193-194] Recently, it has been shown that Tyrosine could quench fluorescence of nearby fluorophores in a similar manner as a Trp-bimane pair does.^[195] Moreover, Trp was able to quench the fluorescence of other fluorophores, such as some oxazine dyes, ATTO 655 and Rhodamine 6G in solution as well as bound to proteins.^[196-197] Using steady-state and time-resolved fluorescence spectroscopy, together with single-molecule FCS measurements, the formation of rapidly quenched complexes was suggested, in which nonradiative deactivation occurs by photoinduced electron transfer (PET) mechanisms.^[196]

An example of probing pH-triggered conformational rearrangements of the T-domain of diphtheria toxin by short-range Trp-bimane fluorescence quenching and long-range Trp-AEDANS FRET measurements is shown in Figure 6.^[198-199]

Figure 6.

Example of the distance measurements during conformational switching in a protein by using a combination of long-range and short-range fluorescence. (a) pH-triggered histidine protonation of the T-domain result in structural changes in proximity of residues Q369 (C-terminal helix TH9) and W206 (N-terminal helix TH1) in membrane-incompetent W-state (cyan) and membrane-competent W⁺ -state (yellow), as estimated by MD simulations. (b) Trp-to-bimane proximity changes (black and red arrows in panel a) are monitored by fluorescence decay, which is strongly quenched in W-state (pH 8, black), but not in W⁺ -state (pH 5.5, red). (c) Long-range FRET measurements between two energy donors, W206 and W281, and an Q369C-AEDANS energy acceptor, demonstrating the retention of the overall compact conformation of the T-domain.

Conformational reorganization of the protein occurring upon histidine protonation (W⁺ -state) results in an increase in the distance between Q369 in the C-terminus of helix TH9 and W206 in the N-terminus of helix TH1. Short-range FRET quenching was used to monitor changes along this distance by replacing Q369 with a cysteine and labeling it with a bimane dye. Fluorescence lifetime measurements of the bimane-labeled T-domain at pH 8 showed highly quenched kinetics with a pronounced subnanosecond component (Figure 6b, black). The quenching is, however, decreased at pH 5.5, suggesting some increase in Trp-to-bimane distance. At intermediate pH 6, the decay could be represented by the mixture of quenched and unquenched fluorescence kinetics.^[199]

Despite pH-induced local rearrangement of the T-domain probed by short-range sensitive Trp-to-bimane quenching, the overall compact conformation of whole protein was retained, as confirmed by another FRET experiments between native Trps and an AEDANS dye selectively attached at Q369C residue (Figure 6c).^[199] At pH 8.0 (black) and pH 5.5 (red), fluorescence excitation spectra measured at the AEDANS acceptor emission band at 470 nm showed no variation in the FRET ratio, which suggests the retention of the folded structure. However, GndHCL-induced denaturation of the T-domain protein resulted in the intensity loss of the excitation band of the Trp donor at 280–290 nm (blue), indicative of global unfolding accompanied by losing long-range FRET.^[199]

Other examples of Trp-AEDANS FRET quenching were focused to determine membrane topology of Colicin E1,^[200] Ca²⁺-dependent structural coupling of globular domains of Calmodulin,^[201] and the structure of N-terminal domain of M13 major coat protein.^[202]

To use FRET measurements as a molecular-size ruler for distance mapping, it is crucial to discriminate between static and dynamic quenching mechanism, operating by different distance dependences.^[203]In complex systems, probe-quencher double-labeled systems, actual quenching mechanism is often unknown; therefore, it cannot be automatically assumed, but instead, it should be validated by steady-state and time-resolved fluorescence measurements.^{[21,93,180,203-204]}

An example of such validation is given in Figure 7. The transition metal ion Forster resonance energy transfer (tm-FRET) approach utilized short-range fluorescence quenching of peptide-attached fluorophores by transition metal ions, such as Ni²⁺ or Cu²⁺.^[205-208]

Figure 7.

Fluorescence quenching of bimane-labeled peptide by transition metal ions (Cu²⁺ and Ni²⁺). (a) Molecular models of a Bi–C2H6H10 peptide composed of a bimane dye attached to an ideal a-helical by the cysteine residue is shown in blue, whereas the two histidines comprising a metal binding site and Cu²⁺ ion are shown in red and green, respectively. The three panels correspond to the three representative dihedral angles of the bimane linker, estimated by molecular modelling. The arrows show the center-to-center distances between the centers of a bimane dye and a metal ion. Steady-state (b) and time-resolved (c) fluorescence quenching of bimane-labeled peptide C2H6H10 using transition by metal ions.^[209]

In recent studies using this approach, a series of helical peptides was designed in which a thiol-reactive bimane dye was attached to a cysteine residue, whereas transition metal ions (Cu²⁺ and Ni²⁺) were chelated by a pair of histidines placed at different positions along the sequence.^[210] Figure 7b shows steady-state fluorescence titration of Bi–C2H6H10 peptide^[205,210] with Cu²⁺ and Ni²⁺ ions resulting in a substantial intensity reduction. The measurements were performed for the following samples: (i) containing no quencher (B1) and at saturating concentrations of Ni²⁺ (B2) and Cu²⁺ (B3). Despite the substantial reduction in intensity in the presence of metal ions (F_M/F), corresponding fluorescence decays were superimposable. These quenching results provide strong evidence of the static nature of quenching, which is likely caused by shorter-range orbital coupling between a dye and a transition metal ion quencher.^[209] This example demonstrates that the FRET-like sixth power of distance dependence of quenching cannot be a priori assumed for short-range quenching, so that time-resolved measurements are required for distinguishing among various quenching mechanisms.

While interpreting short-range quenching, it is important to keep in mind that the quenching efficiency has different distance dependence for Forster and Dexter mechanisms.^[211] For example, ignoring the contribution of Dexter mechanism can lead to the errors in the interpretation of the scale of the conformational changes in proteins. The correct interpretation can be ensured by the comparison of the static and dynamic quenching (see Figure 2 and Figure 7 for examples)^[209].

It has been shown that thioamides quench Trp and Tyr fluorescence in a distance-dependent manner, which can be used to monitor the binding of thioamide-containing peptides to proteins.^[211] Thioamide-containing amino acids have been shown to quench a wide range of fluorophores through distinct mechanisms. In a series of polyproline peptides with terminally-attached thioamide and a fluorophore, it was found that thioamide quenched Tyr fluorescence by FRET, while Trp quenching occurs through an alternate Dexter mechanism.^[212]

4. Fluorescence Quenching in Membranes and Quantitative Analysis of Membrane Protein Insertion

4.1. Dye-Labeled Lyso-Phospholipid FRET Quenchers for Studies of Membrane Protein Topology

Dye-labeled Lyso-glycerolipids, such as LysoMC and LysoUB (Scheme 7), are rapidly and completely partitioned into LUVs. Their lipid tail anchors a headgroup-attached dye at an interface of a lipid bilayer. The significant advantage of Lyso-lipids is that they can be added from aqueous stocks to preformed LUVs externally as micelles, and will reside only in the outer leaflet of the lipid bilayer. This asymmetric labeling of membranes with Lyso-lipids can be used for determining membrane topology (e. g., cis- or trans-leaflet location) of proteins and peptides appropriately labeled with Trp (using LysoMC) and NBD probes (using LysoUB).^{[5,136,213-215]}

Scheme 7.

Dye-labeled Lyso-phospholipids, designed for measurements of membrane topology of proteins and peptides. LysoMC (N-(7-hydroxyl-4-methylcoumarin-3-acetyl)-1-palmitoyl-2-hydroxy-sn-glycero-3-phosphoethanol-amine) and LysoUB (UniBlue-A-1-palmitoyl-2-hydroxy-sn-glycero-3-phos-phoethanolamine) are FRET quenchers (acceptors) for Trp and NBD, respectively.

It has been shown that 4-methyl-coumarine-labeled Lyso-phospholipid (LysoMC), after the incorporation into LUV, quenches the fluorescence of membrane-bound Trp that reside in the same monolayer as the probe.^[213] The Trp quenching occurs by the Förster energy transfer mechanism with an apparent ${\mathrm{R}}_0=25$ Å, which is of comparable size to the thickness of the hydrocarbon core of a lipid bilayer.^[213,216] This quenching method has also been used to estimate the rate of trans-bilayer diffusion (flip-flop) in POPC and POPG bilayers.^[213]

The further modification of the Trp-LysoMC FRET method has been suggested by replacing a weakly-emitting Trp probe by the brightly fluorescent NBD and using a non-fluorescent LysoUB quencher to decrease the background due to direct acceptor excitation (Scheme 7 and Figure 8).^{[214,217-218]}

Figure 8.

(a) Structure of soluble diphtheria toxin T-domain with highlighted NBD-labeled residues L350C-NBD and P378C-NBD. FRET quenching of NBD by LysoUB within quenching radius R, as illustrated by a shaded circle. (b) Application of NBD/LysoUB FRET quenching method for distinguishing topologies of the interfacial (IF) intermediate and final transmembrane (TM) conformations of helixes TH8-TH9 of the membrane-inserted T-domain.

An illustration of the NBD-LysoUB FRET quenching method is given in Figure 8 for the case of the membrane topology of helices TH8 and TH9 of the translocation domain (T-domain) of diphtheria toxin.^[130] To test whether the insertion topology of the T-domain corresponds to either interfacial or transmembrane configuration, an NBD probe was attached to the very tip of helix TH9 using the L350C mutant (Figure 8a). After mixing the T-domain with LUVs at low pH and incubation for 1 hour, the T-domain insertion was probed quantitatively by measuring the fluorescence lifetime τ of the L350C-NBD label for two samples, one in the absence of the quencher (τ₀) and another one after adding 2% of LysoUB quencher (τ_q). If helices TH8-TH9 adopt the TM conformation upon insertion, the L350C-NBD probe will be translocated across the bilayer to its trans-side, after which the probe becomes inaccessible to the externally added LysoUB quencher. On the contrary, if helix TH9 remains interfacial at the cis-side, the probe quenching will be essential (Figure 8b). It was found that when the T-domain was incubated with LUV containing high fraction of anionic lipids, the calculated lifetime ratio of τ₀/τ_q was 1.12, suggesting translocation of the probe to the trans-side of the bilayer. In a control experiment using the NBD-labeled P378C mutant, for which no translocation is expected during TH8-TH9 insertion, the lifetime ratio τ₀/τ_q was equal to 1.72, providing further evidence for the TM topology.^[130] In contrast, in the absence of anionic lipids, no difference in quenching of NBD attached to either side of TH9 was observed, suggesting that a non-productive interfacial intermediate state is stabilized in bilayers formed of zwitterionic lipids (the latter was confirmed by the independent functional studies^[219]).

Other examples of application of the NBD-LysoUB FRET quenching include estimating the membrane topology of Annexin B12^[220] and cytoplasmic ATPase motor protein SecA.^[221]

4.2. Bromine-Labeled and Spin-Labeled Phospholipids for Measurements of Membrane Penetration via Depth-Dependent Quenching

One of the important structural methods that utilizes fluorescence probes on proteins and lipid-localized short-range quenchers is the depth-dependent fluorescence quenching.^{[215 222-225]} The probes for such studies are usually Trp residues or NBD dyes, selectively attached to Cys residues. The quenchers are either heavy atoms like bromine or paramagnetic nitroxide spin labels, covalently attached to the polar/hydrophilic head-group or specific regions of acyl chains of phospholipids (Scheme 8). Using a series of quencher-labeled lipids, in which a quencher is placed at a defined depth within a bilayer, makes it possible to estimate a precise membrane penetration depth of a given fluorophore.^[225] Depth-dependent fluorescence quenching of protein-labeled fluorophore occurs predominantly via collisional quenching mechanism.^{[117,226,227]}

Scheme 8.

Structures and transverse depths of lipid-attached fluorescence quenchers. (Top) Dibrominated phospholipids: 1-palmitoyl-2-(n,n-dibromo)-stearoyl-sn-glycero-3-phosphocholine (n,n-Br-PC). (Bottom) Paramagnetic spin-labeled phospholipids: Tempo-PC (1-palmitoyl-2-oleoyl-sn-glycero-3-phospho-(TEMPO)-choline), n-Doxyl-PC (1-palmitoyl-2-stearoyl-(n-Doxyl)-sn-glycero-3-phospho-choline) spin-labeled lipids. The numbers below the structures correspond to the average immersion depth (e. g., distance from the bilayer center) for each of the quenchers.

Dibrominated phospholipids have been utilized as quenchers for tryptophan and pyrene fluorescence (Scheme 8).^[228] Incorporation of bromo-lipids into LUVs cause only minor perturbations of the membrane. Transverse position of bromine atoms in bromolipids has been determined by X-ray diffraction.^[229]

The lipid-attached paramagnetic spin moieties, such as Tempo or Doxyl (Scheme 8), can quench fluorescence from other organic dyes, including NBD, rhodamine, bimane and anthracene. The membrane-embedded spin-quenchers, such as Tempo-PC and Doxyl-PCs, are characterized by a broad distribution across a bilayer, as shown by MD simulations.^[230] To account for the trans-leaflet quenching of highly penetrated proteins or peptides, the spin-probes are distributed in both leaflets of the lipid bilayer. To extract quantitative information about membrane-penetration depth, two practical methods exist, such as the parallax approach^[226,231] and distribution analysis.^{[223,225,227]}

The distribution analysis (DA) method does not require any assumption about the immersion depth of a fluorophore; instead, it uses a series of 4–6 bromo- or spin-quenchers positioned at the known depths from the center of a bilayer up to its interface.^{[215,225,232-233]} Measuring steady-state and time-resolved fluorescence for a series of quenchers gives a “quenching profile” (QP), which can be fitted by a single or double Gaussian function.^[136,223]

An illustration of the DA method is given in Figure 9 for the case of the depth-dependent quenching of NBD-PE by a series of six quenchers with the different immersion depth of its paramagnetic moiety, such as Tempo-PC and n-Doxyl-PCs with n = 5, 7, 10, 12, and 14 (Scheme 8), respectively. The NBD-PE probe and the spin-labeled quenchers were membrane-embedded by co-extrusion in POPC-LUV. In addition, the quencher concentration (Q) was gradually varied from 7.5 to 30 mol%. As seen in Figure 9a-b, the addition of the Tempo-PC quencher leads to significant quenching of both steady-state fluorescence intensity and shortening of NBD fluorescence decays.^[234]

Figure 9.

Depth-dependent fluorescence quenching of NBD–PE in a lipid bilayer. (a–b) The steady-state and lifetime fluorescence quenching of NBD–PE by Tempo–PC in POPC-LUVs, as shown for the samples containing different quencher content. (c-d) Distribution analysis (DA) of depth-dependent fluorescence quenching profiles of NBD–PE obtained with a series of the six spin-labeled lipids. (c) Quenching profiles of steady-state fluorescence quenching of NBD–PE in LUVs containing the different quencher concentrations. (d) Application of DA methodology to lifetime quenching (squares) and “differential” quenching (circles). See the text and refs^{[223,234-235]} for more detail.

Quenching profiles (QP) of steady-state fluorescence quenching of NBD–PE plotted as F₀/F-1 versus the immersion depth of all six quenchers could be well fitted by the double Gaussian function, resulting in the NBD immersion depth h_m= 13.9 Å and its distribution width σ = 7.2 Å (Figure 9c). The same procedure for the lifetime quenching provided the “dynamics” QP, which accounts for the thermal “broadening” of the probe location within a bilayer. Finally, the application of DA methodology can provide the “differential” QP (circles in Figure 9d), calculated as the difference between the total F₀/F-1 quenching (circles in Figure 9c) and “dynamic” quenching.^[234] Thus, a combination of steady-state and time-resolved measurements could generate a “differential” QP, reducing the contribution from the transverse diffusional quenching occurring during the excited-state lifetime. Such a procedure results in narrower, better-defined QPs, compared to those obtained by traditional fluorescence intensity measurements.

Using a series of 6,7-Br-PC, 9,10-Br-PC, and 11,12-Br-PC, membrane interactions and penetration depth of the amino-terminal domain of huntingtin^[37] and arginine-rich cell-penetrating peptide RW16^[236] were examined using the DA approach. The DA method has been used to measure the depths of penetration of NBD-labeled mutants of the diphtheria toxin T-domain^[235,237] and Bcl-xL,^[132] the membrane topology of NBD-labeled MPER peptide^[46] and BAX protein.^[238]

The applications of the DA method to distinguish between the interfacial membrane topology of the central hairpin (helix α6) of Bcl-xL and the transmembrane one of the translocation domain (T-domain, helix α9) of diphtheria toxin are summarized in Figure 10. Single-cysteine mutants V161C and W169C of Bcl-xL were site-selectively labeled by an NBD probe at the ends of helix α6. The differential quenching profiles shown in Figures 10a-b suggest that Bcl-xL helix α6 is deeply penetrated into the bilayer, as seen from the membrane depth of V161C-NBD (11.3 Å) and W169C-NBD (10.6 Å) probe; so that the helix α6 does not insert in a transmembrane orientation (Figure 10e).^[132] Comparison of the differential quenching profiles for P378C-NBD and I364C-NBD placed at helix α9 of the diphtheria toxin T-domain is shown for the two limiting cases of shallow (P378C–NBD) and deep penetration of the probe (I364C-NBD) in Figure 10c-d, confirming the transmembrane topology (Figure 10e).^[235]

Figure 10.

Illustration of the difference in membrane penetration of helices of the central hairpin of Bcl-xL (left panels, helix α6^[132]) and the translocation domain of diphtheria toxin (right panels, helix α9^[235]). Differential depth-dependent quenching profiles of NBD probe selectively attached to a single cysteine residue at position 161 (a) or 169 of Bcl-xL (b), and at position 378 (c) or 364 (d) of the diphtheria toxin T-domain (see the text for more detail), (e) Schematic representation of the membrane topology of a-helix 6 of Bcl-xL and a-helix 9 of the T-domain (corresponding $h_m$ parameters are given in parenthesis). Reproduced with permission from^[240]. © 2023 The Authors. Published by Elsevier B.V.

In addition, a method of a “dual fluorescence quenching” has been developed to estimate membrane penetration depths of Trp residues in membranes.^[239] This method utilizes the combination of two quenchers, lipophilic 10-doxylnonadecane (10-DN) and aqueous acrylamide. The resulting ratio of the quenching efficiencies between the two quenchers (Q-ratio) have been suggested to have a linear dependence on a Trp depth in membranes.^[239]

5. Summary and Perspective

Applications of different classes of intrinsic and extrinsic fluorescent probes in combination with their specific quenchers are overviewed in context of studies of protein folding and protein-membrane interactions. We discussed various environment-sensitive dyes, FRET probes, probe-quencher pairs for short distance measurements as well as paramagnetic quenchers for quantitative estimation of depth-dependent quenching of membrane proteins and peptides. In the not-so-distant future, we expect the development of more labeling techniques to selectively introduce various fluorescent probes into proteins in the cellular or native-like environments. In addition, we foresee an increasing use of the fluorescence spectroscopy in conjunction with various computer simulations, aiming at reconstructing atomistic or near-atomistic processes of physiological and biomedical importance. The first examples of such joined refinements of fluorescence experiments and results of the all-atom MD simulations have been particularly useful for studies of membrane-interacting proteins,^{[42,46, 119, 235, 241-243]} which are notoriously difficult to study by regular structural methods.

Glossary

AlexaFluor

AlexaFluor^® dye family

ECFP

enhanced cyan fluorescent protein

EYFP

enhanced yellow fluorescent protein

GFP

green fluorescent protein

LUV

large unillamelar lipid vesicle

LysoMC

4Me-coumarine-labeled Lyso-PE, FRET quencher for Trp

LysoUB

UniBlue-A-labeled Lyso-PE, FRET quencher for NBD

mCherry

red fluorescent protein

FRET

Förster resonance energy transfer

smFRET

single-molecule Förster resonance energy transfer

DA

distribution analysis

FCS

fluorescence correlation spectroscopy

FP

fluorescent protein

QP

quenching profile

MD simulation

molecular dynamics simulation

pHLIP

pH-low insertion peptide

NBD, AEDANS

synthetic fluorescence probes

Biographies

Dr. Alexander Kyrychenko obtained his Ph.D. in Physical Chemistry in 1998 and Doctor of Science in Chemistry in 2017 from Kharkiv National University, Kharkiv, Ukraine. He worked as a Postdoctoral Fellow in Institute of Physical Chemistry, Warsaw, Poland (1998-1999) and Chalmers University of Technology, Gothenburg, Sweden (1999-2002). During 2008 to 2013 he worked as a Visiting Researcher in the University of Kansas Medical Center, Kansas City, U.S.A. He is working as a Senior Research Associate (2014-2021), a Head of Department of Physical-Organic Chemistry in Research Institute of Chemistry (since 2021) and Professor of School of Chemistry (since 2016) of V.N. Karazin Kharkiv National University, Kharkiv, Ukraine. His current research interests focus on the fluorescence spectroscopy and computational chemistry of fluorescent probes and sensors for biomedical applications.

After receiving his PhD from the Institute of Biochemistry in Kyiv, Ukraine, Alexey Ladokhin has moved to the US, where he worked as a research associate at the University of Virginia, Johns Hopkins University, and the University of California-Irvine. He has started his independent lab in 2004 at the University of Kansas Medical Center, where he is currently holding a position of Professor of Biochemistry and Molecular Biology. Over the years, Alexey Ladokhin served as a Guest Editor in various journals, including the most recent BBA Advances special issue entitled “Highlights of Ukrainian Molecular Biosciences”. He is currently serving as the Editor-in-Chief for the Journal of Membrane Biology of the Nature/Springer family. Dr. Ladokhin’s NIH-funded research is dedicated to deciphering structural, functional, and thermodynamic aspects of protein-membrane interactions.