FRET - Förster Resonance Energy Transfer || Materials for FRET Analysis: Beyond Traditional Dye-Dye...

6Materials for FRET Analysis: Beyond Traditional Dye–DyeCombinationsKim E. Sapsford, Bridget Wildt, Angela Mariani, Andrew B. Yeatts, and Igor Medintz

6.1Introduction

The intrinsic sensitivity (r6 dependence) of F€orster (or fluorescence) resonanceenergy transfer (FRET) to nanoscale changes in the donor/acceptor separationdistance has made FRET an invaluable biophysical tool in a variety of applications,ranging from studying the structure and conformation of proteins and nucleic acidsto examining biomolecular interactions, including its use in in vitro and in vivobioassays [1–7]. While the myriad of FRET configurations and techniques currentlyin use are covered throughout this book, here we focus primarily on the materialsutilized as donor or acceptor probes in FRET rather than the process itself [3,5,8,9].Our 2006 review paper on this topic serves as the foundation for this updatedchapter [3]. The materials were divided into three main categories: organic materialsthat include “traditional” dye fluorophores, dark quenchers, polymers, and carbonnanomaterials (NMs); inorganic materials such as metal chelates, metal, andsemiconductor nanocrystals; and fluorophores of biological origin such as fluores-cent proteins (FPs), amino acids, and fluorescence generated from enzymaticbioluminescence (BL) and chemiluminescence (CL). These materials may functionas FRET donors and/or acceptors, depending upon experimental design. Many ofthe new materials developed and/or new donor–acceptor probe combinations usedaddress some of the inherent complications of more traditional FRET materials,including photobleaching, spectral cross talk, and direct excitation of the acceptorspecies, and examples of these will be highlighted and discussed throughout thechapter. Since the vast majority of FRET applications are biological in nature, theyroutinely involve some type of biomolecule labeling strategy, which ultimately playsa significant and fundamental role in the success and interpretation of the resultingFRET. Therefore, we begin the chapter with a brief discussion of the bioconjugationtechniques commonly utilized for FRETand points to consider, followed by sectionshighlighting the current and emerging materials that are used, or have the potentialto be used, in FRET applications.

FRET – Förster Resonance Energy Transfer: From Theory to Applications, First Edition.Edited by Igor Medintz and Niko Hildebrandt.� 2014 Wiley-VCH Verlag GmbH & Co. KGaA. Published 2014 by Wiley-VCH Verlag GmbH & Co. KGaA.

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6.2Bioconjugation

A large number of FRET-based applications involve the use and labeling of sometype of biomolecule (e.g., cell membrane, antibodies, nucleic acids, protein, andpeptides). Given the sensitivity of FRET to a number of parameters, the ability tocontrol the donor/acceptor labeling in the system under investigation is paramountto obtaining well-controlled and reproducible results that will aid in subsequentinterpretation of experimental data. The extent of such control is initially dictated bya combination of factors, including (i) the nature of the system under study (i.e., areinter- or intramolecular studies desired?), (ii) the availability/number (and reactivity,where applicable) of attachment/incorporation sites on the biomolecule(s) for thedonor/acceptor probes, (iii) the nature of the donor/acceptor probes (e.g., organicmolecule, fluorescent protein, and NM), (iv) the size of the donor/acceptor probes(especially protein-based and NMs) relative to the biomolecule(s), which caninfluence the system under study, and (v) the availability of the donor/acceptorprobes with the desired reactivity for bioconjugation and the nature of the linker(e.g., length and flexibility), connecting the donor/acceptor probes to the bio-molecule. For quantitative FRET the microenvironment dependency of the fluo-rophore’s photophysical properties and the uncertainty in probe position andorientation relative to the biomolecule should be considered when choosingdonor/acceptor probes and bioconjugation strategies [6]. Donor/acceptor probeattachment to a biomolecule can be achieved via a number of labeling techniquesthat can be chemically or biologically inspired in nature (Figure 6.1); for recentreviews see Refs [10–12].Bioconjugation based on forming interactions, typically covalent bonds between

the biomolecule and the probes, represents the most popular and traditional groupof chemistries used to date, and will likely remain the workhorse in the near futuredue to the commercial availability of a wide variety of donor/acceptor probesmodified with a number of reactive functionalities that facilitate bioconjugation.In the case of protein labeling, for example, the predominant chemically basedbioconjugation strategies target the naturally occurring amino acids lysine (Lys –

primary amine) and cysteine (Cys – thiol) with succinimidyl ester (NHS) ormaleimide reactive groups, respectively. Control of the donor/acceptor probelocations and stoichiometry is one of the many important considerations whendesigning FRET studies [5]. In the case of deoxyribonucleic acid (DNA)/ribonucleicacid (RNA) and short synthetic peptides, control of location and stoichiometry can beprogrammed into their structure via inclusion of the donor/acceptor moleculesthemselves or site-specific incorporation of thiol/amine groups for subsequentlabeling during synthesis [13,14]. However, proteins are generally more complexbecause they contain a number of primary amines that can cause difficulty incontrolling the location of labeling and usually results in variable dye-to-protein(D/P) ratios. Targeting thiols on Cys residues with maleimide chemistry is morespecific and can reduce the labeling variation, as these residues are far rarer.However, Cys residues are often critical to protein structural conformations as

166j 6 Materials for FRET Analysis: Beyond Traditional Dye–Dye Combinations

part of disulfide bonds and are typically buried below the protein surface, which canlimit access depending on the size of the chosen donor/acceptor probe. Thiols canbe chemically (typically via Lys interconversion) or recombinantly introduced intothe protein surface [15–17]. However, this too can be problematic as additional Cysresidues can “thiol-scramble” the protein structure during folding, and surface-exposed thiols can result in the formation of protein dimers, trimers, and so on,which, when purified, necessitate further reduction prior to labeling. There are acouple of excellent resources available for researchers interested in bioconjugationprotocols in general [18] and fluorescent labeling in particular [19].Bioorthogonal reactions (reactions that do not interfere with other biogroups

besides the target), which were born out of the desire to study biomolecules in theirnative environment, have the ability to address many of the controlled labelingconcerns outlined earlier [12,20]. Many of these reactions are based on organicchemistry-inspired covalent modifications and include the Staudinger ligation

Figure 6.1 Bioconjugation methods separated into chemistry- and biology-inspired techniques.

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reaction, ketone/aldehyde condensation reactions (bioorthogonal, depending on thesystem under study), and a variety of cycloaddition reactions, including thequintessential “click chemistry” azide/alkyne cycloaddition.There are a number of biologically inspired bioconjugation strategies that are

gaining popularity, which, although perhaps not bioorthogonal in the strictest sense,possess many of the stipulations and benefits required to be considered asbioorthogonal reactions [12,21]. Fluorescent proteins (FP) such as green fluorescentprotein (GFP) can be appended to existing proteins using recombinant techniquesgenerating protein chimeras, and although not considered a bioorthogonal chemis-try, this does allow the site-specific incorporation of the FP to the protein of interest(see Section 6.4.3) [22]. Likewise, nonnatural amino acid residues (Section 6.4.2),short peptide tags, and full proteins (such as enzymes), that are either fluorescent orallow specific labeling with fluorophores, can also be incorporated genetically intoprotein structures [10,12,23]. For example, the tetracysteine/biarsenical system,originally developed by Tsien and coworkers, demonstrated that proteins expressingan optimized Cys-Cys-X-X-Cys-Cys sequence [where X¼ could be any amino acid,but is traditionally proline-glycine (Pro-Gly)] would react with biarsenical-function-alized fluorophores (e.g., FlAsH and ReAsH) (Figure 6.2) [21,24,25]. Oligohistidine(His) peptides (e.g., His6) are peptide tags that have been used for conjugation. Theyare known to bind nickel-nitrilotriacetic acid (Ni2þ-NTA)-functionalized molecules,while the oligoaspartate (Asp) sequence (D4-tag, Asp4) has a strong binding affinityto multinuclear zinc(II) complexes [10,12,26,27].There are an expanding number of biologically inspired techniques that

harness genetically encoded peptide handles combined with enzymes to catalyzesmall-molecule (e.g., biotin and fluorescent dye) conjugation [10–12,16,23,28,29].

Figure 6.2 Bioconjugation using peptiderecognition. (a) Schematic showing the labelingof an engineered protein displaying a lineartetracysteine motif with bisarsenical containing

fluorophores. (b) Chemical structures of FlAsHand ReAsH. (Reprinted with permission fromRef. [25]. Copyright 2011, American ChemicalSociety.)


Jager et al., for example, modified a model protein, chymotrypsin inhibitor 2 (CI2),with a transglutaminase (TGase)-tag (Pro-Lys-Pro-Gln-Gln-Phe, where Gln is gluta-mine and Phe is phenylalanine) at its N-terminus [16]. TGase facilitated the couplingof the TGase-tag-modified CI2 with AlexaFluor 647 (Alexa647, A647) cadaverine(A647-(CH2)6-NH2), forming an isopeptide bond between the TGase-tag Gln residueand the primary amine on the A647 fluorophore. In another example, the Escherichiacoli enzyme biotin ligase has been shown to ligate biotin to proteins tagged with anacceptor peptide sequence Lys-Lys-Lys-Gly-Pro-Gly-Gly-Leu-Asn-Asp-Ile-Phe-Glu-Ala-Gln-Lys-Ile-Glu-Trp-His (where Leu is leucine, Asn is asparagine, Asp is asparticacid, Ile is isoleucine, Ala is alanine, and Trp is tryptophan) [28]. The biotin ligasealso accepted a ketone isostere of biotin as a cofactor resulting in ketone-function-alized proteins that could be subsequently modified with hydrazide- or hydroxyl-amine-functionalized molecules. The HaloTagTM [haloalkane dehalogenase (DhaA)]and SNAPTM-tag [O6-alkylguanine-DNA alkyltransferase (hAGT)] are labeling tech-niques that actually involve fusing the full enzymes (which self-label themselves) tothe protein of interest. The HaloTag utilizes fluorescently labeled haloalkanesubstrates, while the SNAP-tag uses fluorescent benzylguanine derivatives togenerate fluorescently labeled targets [10–12].Nanomaterials are increasingly being used as donor/acceptor probes in FRET

studies due to their many unique properties, and as such many of the bioconju-gation techniques described earlier are applicable here as well [8,9]. However, takinginto account their nanoscaffold nature, additional concerns should be consideredduring bioconjugation [5,30–32]. NM surfaces can be quite complex, comprising notonly the NM itself but also oftentimes additional stabilizing ligands that helpmaintain its aqueous solubility (e.g., colloidal stability) and prevent undesirableinteractions such as aggregation–agglomeration. In addition NMs are generallymuch larger than your typical fluorescent/quencher organicmolecules and are oftenon a similar size and scale or larger than most biomolecules, therefore the potentialinfluence of this size on the system under study should be carefully considered.Other factors to consider regarding bioconjugation include how the NM is stabilizedin solution and whether the reaction conditions might affect the stability or physicalproperties of the NM. A prime example that highlights this issue is the use ofcarbodiimide coupling chemistry to link carboxylic acid (COOH)-modified NMswith the primary amines on a protein, as discussed in a recent review by Algar et al.[30]. COOH-terminated ligands are commonly used to impart charge-based solu-bility and stability to inorganic NMs at basic pH. However, carbodiimide activation(often added in huge excess due to rapid hydrolysis of the reaction intermediate)converts the COOH to a less soluble o-acylisourea intermediate and hence can causereduced solubility and NM aggregation, severely impacting product quality andreproducibility. Another concern is whether the stabilizing surface ligands influencesubsequent interactions involving either the biomolecule attachment reaction or thesubsequent biorecognition event. Dennis et al. used FRET to investigate eightdistinct quantum dot (QD) coatings and their influence on the self-assembly of aHis-tag-labeled mCherry FP that interacts directly with the QD surface via metalaffinity coordination [33]. This coordination relies substantially on access of the

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His-tag to the QD surface, and the researchers found that even subtle changes in theorganic coating could significantly alter the accessibility and hence His-tag–mCherry coordination.In another example of reaction conditions that can impact the NM, the copper

(Cu) in the widely used Cu-catalyzed click chemistry has been found to be extremelydetrimental to the luminescent properties of QD materials [34]. As an alternativeBernardin et al. used a Cu-free strained click chemistry technique, coupling strainedcyclooctyne-functionalized QDs with azido biomolecules, and the QDs in this caseretained their strong luminescent properties [34].Surface ligands and the attachment chemistry used in bioconjugation can also

affect the performance of the NM bioconjugate in the desired application. Forexample, while developing a FRET-based QD–peptide sensor for monitoring botu-linum neurotoxin A (BoNT A) activity, Sapsford et al. found that the stabilizingligand sterically hindered the BoNT A from interacting with the peptide substrateassembled on the QD surface via metal affinity coordination, resulting in a non-responsive sensor [35]. The situation wasmitigated by conjugating the peptide to theterminal groups of the QD-stabilizing ligand, improving access for the BoNT A. Aswith any type of bioconjugation involving a surface, the conformation/orientation ofthe biomolecule upon immobilization is an essential component of its subsequentfunctionality and should be considered during study design. Loss of biomolecularactivity is to be expected if the recognition site of the biomolecule is positioned inclose proximity to the surface [30]. In a study of QD–DNA bioconjugates, Boenemanet al. found that the attachment chemistry strongly influenced the orientation ofDNA on a QD-poly(ethylene glycol) (PEG) surface [36]. His-tag-modified DNAattached directly to the QD surface resulted in a structure that, as predicted,extended out from the surface. Biotin-labeled DNA bound to streptavidin (SA)-modified QDs, however, did not follow predicted models, and the DNAwas found totake a number of random orientations on the QD surface, which was attributed tothe random attachment of SA to the QD. Random orientations are likely to result in adistribution of biomolecular activities, which can affect reproducibility of exper-imental results [30].Clearly, there is a wide range of bioconjugation techniques available to research-

ers, and choosing the most suitable method depends on the nature of the systemunder study. Some of these labeling techniques can be complex, requiring forexample genetic engineering expertise; however, the increase in commercialreagents and kits (from companies such as Life TechnologiesTM, Sigma-Aldrich,and Promega) for these types of bioconjugation reactions, especially with theintroduction of newer bioorthogonal chemistries, is encouraging more widespreadadoption. Regardless of the method chosen, having both the donor/acceptor probesat known and distinct locations on biomolecule(s) is most desirable in terms ofpostexperimental analysis. Care should be taken to ensure that donor/acceptorprobe modification does not influence the biomolecule functionality, especiallywhen trying to determine the functional characteristics of a biomolecule (wherelabeling may interfere with/alter structure, biomolecule conformational changes, orbiomolecule interactions).


6.3Organic Materials

As highlighted in this section, organic materials that possess the necessary photo-physical properties to be utilized as FRET donors or acceptors are a widely diversegroup and include molecules, macromolecules, polymers, and NMs.

6.3.1Ultraviolet, Visible, and Near-Infrared Emitting Dyes

The majority of donor/acceptor materials currently used in FRET applications areultraviolet (UV), visible (Vis), and near-infrared (IR) emitting organic dyes. These“traditional ” dyes are usually the first type of FRET material tested with other new ornon- “traditional” fluorescent materials in potential FRET systems. The most com-mon organic dye classes, shown in Figure 6.3, have several advantages, such ascommercial availability, cost-effectiveness, extensive characterization of FRET prop-erties, easy bioconjugation through NHS-ester, maleimide, hydrazide, or aminechemistries, availability in reactive form, and high quantum yields (QYs) andsolubilities [37,38]. Recent advances in click chemistry now allow companiessuch as Lumiprobe to provide organic dyes conjugated to azide and alkyne chemicalmoieties for further bioconjugation as well.Despite all the advantages of these traditional dyes, there are disadvantages too.

For instance, some have a high rate of photobleaching, may be sensitive to pH, andhave a propensity to self-quench when highly substituted on biomolecules. Some ofthe redder dyes have low solubility in aqueous solvents and for FRET in particular,the broad absorption/emission profiles and small Stokes shifts often lead to directexcitation of the acceptor, complicating subsequent analysis. The quest for neworganic dyes with the potential to overcome these limitations continues, mostrecently with materials such as the Chromeo [39] (currently sold by Active Motif ),CS1-6 near-IR (NIR) [40], and alkyne carbocation “cyanine-like ” dye families [41–43].Researchers have recently demonstrated a series of organic molecules that undergoexcited-state intramolecular protein transfer (ESIPT) for use in the development offluorescent chromophores with a large Stokes shift (LSS) [44– 46].Many resources are available to aid in choosing suitable donor–acceptor pairs,

including a number of FRET reviews [15,47– 49], as well as the Molecular ProbesHandbook [19] and a review by Wu and Brand [50] that offers an extensive list ofdonor–acceptor dye pairs and their respective R0 values. Life Technologies’ Fluo-rescence SpectraViewer (http://www.invitrogen.com/site/us/en/home/support/ResearchTools/Fluorescence-SpectraViewer.html) and Zeiss’ Fluorescence Dyeand Filter Database (https://www.micro-shop.zeiss.com/us/us_en/spektral.php)are Web-based programs that allow researchers to plot multiple dye absorption/emission profiles to optimize spectral overlap and choose appropriate filters. Anexcellent comparison of the physical and spectroscopic properties of a number ofred-absorbing dyes is provided by Buschmann [51]. See also the extended andupdated tables collated by van der Meer and provided later in this book.

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http://www.invitrogen.com/site/us/en/home/support/ResearchTools/Fluorescence-SpectraViewer.html

http://www.invitrogen.com/site/us/en/home/support/ResearchTools/Fluorescence-SpectraViewer.html

https://www.micro-shop.zeiss.com/us/us_en/spektral.php


Many current applications still rely on traditional dye–dye FRETcombinations dueto the many advantages of organic dyes as described earlier [3,47,52]. For instance,FRET-based biosensors have led to a deeper understanding of a number of biologicalphenomena such as integrin adhesiveness and signaling dynamics [53], plasmamembrane biophysical interactions [54], host–pathogen interactions [55], andprotein folding geometries and conformational states [56]. Similar dye combinationsare also useful for FRET-based biosensing, including glucose sensors [57] andbiological agent detection [58].Dye–dye FRET combinations have had tremendous impact on biomedical

research, specifically in the areas of nucleic acid analysis, DNA sequencing, andgenotyping [13,59–62]. In addition, molecular beacon probes used in nucleic acidanalysis and Scorpion real-time PCR assays are often FRET based [13,63]. Interest-ingly, use of DNA scaffolds incorporating donor/acceptor dyes has led to a morefundamental understanding of the orientational dependence of the dyes on FRETefficiency [64–66]. DNA microarrays, in which FRET-based DNA probes are immo-bilized on solid surfaces, are quickly becoming an exciting application with thepotential to increase sensitivity, specificity, and throughput of gene expression aswell as large-scale single-nucleotide polymorphism (SNP) discovery, detection, andgenotyping [67]. There is no doubt that new applications will continue to drive thedevelopment of novel donor–acceptor dye combinations that overcome currentdeficiencies in existing organic dyes.

6.3.2Quencher Molecules

The use of quenching molecules in FRET-based applications continues to bepopular. The primary advantage of using these molecular acceptors over theirfluorescent counterparts is the elimination of background fluorescence due todirect acceptor excitation or reemission. Typically, quenchers take the form oforganic molecules or metallic materials such as gold (Au) (Section 6.5.3). Figure 6.4offers a visual representation of a variety of organic quencher families that arecommercially available. Two of the most common quenching acceptor molecules,Dabcyl (4,-(40-dimethylaminophenylazo)benzoic acid) and Dabsyl (4-dimethylami-noazobenzene-40-sulfonyl), have absorption maxima centered at 485 and 466 nm,respectively. Another recent addition to nonfluorescent quencher dyes is IRDyeQC-1, which is characterized by a broad absorption peak between 550 and 950 nm,

Figure 6.3 Organic UV and visible fluorescentdyes. (a) Structures of the common organic UVand visible fluorescent dyes. Typicalsubstituents at the R position include CO2

�,SO3

�, OH, OCH3, CH3, and NO2; Rx markstypical location of the bioconjugation linker.(Reprinted with permission from Ref. [3].) (b)Plot of fluorophore brightness versus the

wavelength of maximum absorption (max) forthe major classes of fluorophores. The colorof the structure indicates its wavelength ofmaximum emission (em). For clarity, onlythe fluorophoric moiety of some moleculesis shown. (Reprinted with permission fromRef. [37]. Copyright 2008, AmericanChemical Society.)

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effectively allowing the quenching of both near-infrared and the more commonlyused visible donor dyes [68]. Other quencher families that tend to have wide-rangeabsorption spectra include the trademarkedQSY, QXL, ATTO, BlackBerry, and BlackHole Quenchers. These broad absorption spectra decrease design constraints andallow quencher molecules to function as acceptors for many dyes. One applicationwhere quenchers are often applied is DNA analysis, specifically molecular beaconscoupled with organic dye donors [13,48,49]. The primary advantage of this donor–

Figure 6.4 Organic quencher molecules. (a)Example of structures of the common quenchermolecules. Substituents R are listed. Rx markstypical location of the bioconjugation linker.(Reprinted with permission from Ref. [3].)

(b) Some common, commercially available,quencher families along with absorbancemaxima and spectral regions covered by aparticular quencher family. (Adapted withpermission from Ref. [3].)


quencher configuration is that singular or individual donor channels can bemonitored and, if sufficient spectral separation is achieved, utilized for “multi-plexing” with a variety of other donor–quencher pairs. Typical applications of thesemethods have measured DNA permeability of polyelectrolyte thin films [69] andutilized catalytic DNA biosensors to detect lead (Pb) ions [70]. Another techniqueemploys quencher-labeled substrate analogues used in conjunction with dye-labeledproteins for FRET-based biosensing of nutrients through displacement [71].Reduced size of donor/quencher pairs can also improve the accuracy in determiningprotein dynamics. For example, the use of thioamide as a quencher fabricated fromthe protein’s own backbone, through a single-atom substitution, minimized per-turbation of the system [72]. Current techniques also look into increasing theefficiency of quencher molecules through the use of binding molecules withadvantageous three-dimensional conformations and charge density allowingincreased local dye concentrations [73]. One of the few examples of FRET, whereorganic quencher molecules are coupled to nonorganic fluorophores, involves QDdonors (see Section 6.5.5).

6.3.3Environmentally Sensitive Fluorophores

While the vast majority of fluorophores will respond to a certain extent to aperturbation in their microenvironment, some exhibit much higher sensitivitythan others and as such are classified as environmentally sensitive. These fluo-rophores exhibit some change in their spectral characteristics (absorption/emissionprofiles) in response to a change in their microenvironment, such as pH, ioninteractions, or another moiety such as oxygen (O2), solvation, polarity (solvato-chromic fluorophores), or rigidity, and these dyes are usually defined by the analyteor condition that they respond to most favorably [19,47]. A large number ofenvironmentally sensitive fluorophores are found in the small organic moleculesclass of fluorescent probes, however, other classes of fluorophores such as FPs havealso been found to be sensitive to specific changes in their environment [37,74–79].Life Technologies offers a wide range of environmentally sensitive fluorophores,

including dyes sensitive to reactive oxygen species (ROS), pH, calcium (Ca2þ),magnesium (Mg2þ), Zn2þ, sodium (Naþ), chloride (Cl�), potassium (Kþ), andmembrane potential [19]. The dyes are offered in a variety of forms depending on theapplication, including cell-permeant and cell-impermeant, or can be modified withfunctional groups that aid in subsequent conjugation if desired. Many of the targetions are important signaling molecules in molecular cell biology, and intracellularmeasurements are essential to understanding the processes that govern cellularfunction [76,77]. In the case of intracellular pH, for example, two functionalmicroenvironments should be considered, the cytosol (pH 6.8–7.4) and the acidicorganelles (pH 4.5–6.0), with the exact choice of fluorescent probe dependent on itspKa [19,76]. Oregon Green and LysoSensors are appropriate for the more acidicorganelle environment, while fluorescein derivatives, the polar 20,70-bis-(2-carbox-yethyl)-5-(and-6)-carboxy-fluorescein (BCECF) derivative (developed by Tsien) and


the proprietary seminaphthorhodafluors (SNARF) dyes, function optimally in thepH �6.0–8.0 range and are popular choices for cytosol pH measurements [19,80].Nakata et al. recently developed two new SNARF derivatives, SNARF-F and SNARF-Cl, which, while maintaining the characteristic spectral changes of the originalSNARF dye, have improved cell permeability and in the case of SNARF-F animproved pKa (7.38) for cytosol measurements (Figure 6.5) [81]. Tweaking ofthe fluorophore chemical structure can improve the fluorescent properties, andthe target interactions of many of these fluorescent probes and combinatorialapproaches offer an interesting alternative to more traditional and rational designmethods for identifying new improved fluorescent candidates, recently reviewed inRef. [82].While environmentally sensitive fluorophores function alone adequately as

qualitative intensity-based fluorescent probes, the combination of FRET usingenvironmentally sensitive dyes has been demonstrated for detection of pH, ammo-nia (NH3), and carbon dioxide (CO2) [83,84]. When target quantification is desired,especially in studies involving complex cellular environments, FRETrepresents onemechanism in which to achieve ratiometric measurements that are independent offluorescent signal fluctuations [77,85]. Signal fluctuations can result from variationsin local probe concentration, sample thickness, pH, or temperature, which make

Figure 6.5 Environmentally pH-sensitiveSNARF dyes. (a–c) Absorbance (solid line)and fluorescence (dashed line) spectra of10 mM of (a) SNARF, (b) SNARF-F, and (c)SNARF-Cl at pH 5.0 (red) and pH 10.0(blue), excited at the isosbestic point,

respectively. (d and e) Photos of (d) colorand (e) fluorescence of SNARF (10 mM),SNARF-F (10 mM), and SNARF-Cl (10 mM) atpH 5.0 and pH 10.0. (Reprinted withpermission from Ref. [81]. Copyright 2011,Elsevier.)


subsequent interpretation of single-intensity data complicated. Ratiometric FRET-based sensors for the analysis of nucleoside polyphosphates, pH, temperature,hydrogen peroxide (H2O2), mercury (Hg), and chromium (Cr) ions have all beendemonstrated [79,86–90].Environmentally sensitive fluorophores are commonly incorporated into nano-

particles (NPs) for sensing applications. For example, Childress et al. developed dye-doped polymer NPs for ratiometric fluorescence detection of Hg(II) ions in aqueoussolution [89]. NPs of the conjugated polymer (CP) poly[(9,9-dioctylfluorenyl-2,7-diyl)-co-(1,4-benzo-{2,10-3}-thiadiazole)] (PFBT), which fluoresced green-yellow,were doped with a nonfluorescent Hg(II)-responsive rhodamine dye. The rho-damine dye, upon interacting with Hg(II) ions, converts to an orange-red fluores-cent form, resulting in FRET between the donor PFBT NPs and the activatedacceptor rhodamine dye. The sensing NPs couldmeasure low levels of Hg(II) ions inthe 0.7–10 ppb range, and the ratiometric nature of the FRETsensor eliminated anyissues related to environmental or instrumental fluctuations. Using a more con-ventional platform, Kurishita et al. developed FRET-based ratiometric chemosensorsfor detection of nucleoside polyphosphates such as adenosine-50-triphosphate (ATP)[87]. The sensor was based upon a large off–on fluorescence enhancement thatoccurred when a xanthene-based Zn(II) complex bound ATP (Figure 6.6a). Thechemosensor combined a coumarin (blue fluorescence) donor with the xanthene-based Zn(II) complex acceptor, which developed green fluorescence upon ATPbinding. Ultimately the chemosensor was tested in live Henrietta Lacks (HeLa) cells,where initial staining resulted in green fluorescence due to the presence of ATP inthe cells (Figure 6.6c). Introduction of 2-deoxyglucose (2-DG) and/or potassiumcyanide (KCN), which inhibited ATP synthesis, resulted in a significant decrease inFRET, causing an increase in the blue donor emission compared to untreated cells(Figure 6.6d and e).Nucleic acid binding dyes represent a specific class of environmentally

sensitive organic dyes that warrant special mention [91,92]. These dyes formcomplexes with nucleic acids, resulting in significant off–on fluorescenceenhancements, which also incorporate intermolecular homo-FRET to a largeextent. Dyes include 40,6-diamidino-2-phenylindole (DAPI), the bisbenzimide-based dyes (collectively called the Hoechst dyes), OliGreen, ethidium bromide(EtBr), propidium iodide, and the cyanine dyes (PicoGreen, YOYO, and TOTOfamilies of dyes; SYBR Green I and SYBR Gold) [91,92]. Cosa et al. performed acomprehensive study comparing the photophysical properties of a number ofthese dyes alone and upon binding single-stranded DNA (ssDNA) or double-stranded DNA (dsDNA) [91]. All these dyes interact with dsDNA, with somesuch as DAPI and the Hoechst dyes, binding specifically with the minor grooveof adenine (A)–thymine (T)-rich sequences [91,92]. Most of the dyes also seemto interact with ssDNA and some, such as EtBr, have also been found to interactwith RNA [91,92]. The combination of nucleic acid sensitive dyes and FRET hasbeen used in a number of studies dealing with understanding how the dyesinteract with DNA [92–94] and in the detection of single-nucleotide polymor-phisms (SNPs) and short tandem repeats (STRs) [95–97].


6.3.4Dye-Modified Microspheres/Nanomaterials

There are a wide range of materials that can and have been modified with organicdyes to generate fluorescent microspheres or NMs, although the most commonplatforms are either polymer or silica based. Modification typically involves eitherencapsulation of the fluorophores within the material core or interaction with theparticle surface. Recent advances in NM synthesis and improved surface function-alization techniques have allowed the increased use of dye-labeled NMs in particular,which, from a size perspective, is a benefit in FRET-based applications. Dye-labelednano- and micro-sized particles have been prepared through ionic interaction [98],miniemulsion [99,100], covalent conjugation [101], or encapsulation of fluorophoremolecules during synthesis [102–105], and the benefits of these formats result inincreased signal, decreased photobleaching, increased surface functionalizability,and lower limits of detection than small-molecule fluorophores [106]. Due to thepopularity of diagnostic and research techniques utilizing fluorescence detectionsystems, commercial fluorescent particles are available from a wide variety ofsources such as Molecular Probes, Phosphorex, Spherotech, Bangs Laboratories,Sigma-Aldrich, and Polysciences. Besides standard fluorophores, fluorescent micro-spheres loaded with europium (Eu) chelates are available, extending the utility ofsuch labels to time-resolved energy transfer configurations (Section 6.5.1) [107].A current trend in microspheres and NM synthesis involves using FRET to tune

the spectral properties of the resulting fluorescent microspheres/NMs, generatingfluorescent FRET-based tags that have improved Stokes shifts and unique fluores-cent signatures that can be excited by a single wavelength (Figure 6.7)[99,100,102,108–111]. These FRET systems can also be designed to contain

Figure 6.7 Polymeric FRET-based NPs for invivo imaging. The particles were assembledfrom diblock copolymers of poly(D,L-lactic-co-glycolic acid) and maleimide-activated PEG,which were also encapsulated in both thedonor (1,10-dioctadecyl-3,3,30,30-tetramethylindodicarbocyanine) and acceptor(1,10- dioctadecyl-3,3,30,30-tetramethylindotricarbocyanine) fluorophores.FRET resulted in a large Stokes shift (>100 nm)

of the emission maxima, and the transferefficiency could be fine-tuned by furtheradjusting the doping ratio of the donor andacceptor fluorophores. The optimizedformulation was less than 100 nm in size,brighter than quantum dots, stable inbiological media, and demonstrated similarbiodistribution to most NMs. (Reprinted withpermission from Ref. [102]. Copyright 2012,American Chemical Society.)


photochromic dyes (discussed further in Section 6.3.6) that increase signal detectionby decreasing background emission [99].A variety of hybrid fluorescent core–shell silica NPs with interesting optical

properties have been developed, such as, dye-labeled silica-coated silver (Ag) NPswith enhanced FRET properties [112,113] and Cornell dots (CU dots) [114,115]developed by Wiesner at Cornell University. CU dots are synthesized by covalentlyconjugating dye molecules to a silica precursor before being condensed to form adye-rich core. Silica sol–gel monomers are then added to form a denser outersilica network.The use of dye-labeled microparticles and NPs in FRET-based analytical and

research assays continues to grow as these materials are exploited in nanoscopicruler measurements [116], biosensing of infectious agents [105,111], in vivo biol-abeling [102], flow cytometry [117,118], SNP genotyping [119,120], and diseasediagnosis using protein nanoarrays [121].

6.3.5Dendrimers and Polymer Macromolecules

Dendritic and polymer macromolecules are increasingly being used in fluores-cence-based applications. Dendritic architectures represent a class of repeatedlybranched or tree-like polymeric structures of which the subclasses of dendrimersand dendrons have found particular application in molecular imaging, sensing,photovoltaics, and energy harvesting [122–124]. Dendrimers, in particular, arehighly ordered macrostructures comprised of a distinct core, branched mid, andbranched surface/periphery regions. The ability to tailor dendrimer structureswith inclusion of multiple functional groups during synthesis allows precisecontrol over the position and orientation of fluorophores and attachment ofbiomolecules leading to well-defined macromolecules [122–124]. This precisestructural control has made dendrimers interesting synthetic platforms in whichto mimic Nature’s light-harvesting structures, reviewed in Ref. [124], and in thedevelopment of new photovoltaic materials [125]. Of the many types of den-drimers reported, the poly(amidoamine) (PAMAM)-based materials are the mostutilized because the many primary amines enable facile functionalization[122,123]. The ability to label these dendritic structures with multiple fluoro-phores results in increased absorption cross sections and higher fluorescenceintensities, which is ideal for bioassay and imaging applications [122,123]. Thisstrategy is employed in a class of dendron-based fluorogenic dyes known asdyedrons, which comprise multiple donor Cy3 dyes coupled to a single malachitegreen (MG) acceptor [126]. Free in solution, the MG acceptor acts as a quencherdue to unconstrained internal structural rotations, however, upon binding tofluorogen-activating proteins (FAPs) such as single-chain variable fragment anti-bodies, the MG acceptor becomes activated, making it highly fluorescent whenexcited via FRET from the Cy3 donors. Such dyes have potential application astargeted probes in sensitive homogeneous (no wash) imaging. Inherently fluo-rescent cationic dendritic structures comprising primarily phenylene-ethynylene


have been used as donors for the detection of peptide nucleic acid (PNA)/DNAhybridization, where the neutral PNA probe was labeled with a fluoresceinacceptor [127]. Davis et al. used inherently fluorescent cationic and anionicdiphenylacetylene dendrimers doped into cellulose acetate to generate solid-stateelectrospun nanofiber sensor arrays for protein detection [128]. These nanofiberfilms selectively interacted with proteins (including metal- and nonmetal-contain-ing proteins), which resulted in fluorescent quenching in distinct patterns, due tovarying interactions with the different protein structures; this allows specificprotein identification even in complex mixtures. Dendrimers have also been usedas carriers for multiple materials such as combinations of drugs, nucleic acids,antibodies, fluorescent tags, and/or contrasting agents that can be used forimaging, drug delivery, or bioassay applications [123]. For example, Myc et al.used folic acid modified dendrimers to target delivery of a FRET-based apoptoticsensor to cancer cells that could be used to determine the efficacy of a chemo-therapeutic or other targeted treatment by monitoring cell death in real time [129].Although synthetic procedures for dendrimer synthesis are available in theliterature, dendrimers are also commercially sold with functionalities that canbe further modified by the end user, which may aid in more widespread use(Dendritech, Polymer Factory Sweden AB, and Dendritic Nanotechnologies Inc.).Qiagen offers dendrimers specifically functionalized to bind both DNA and cellsfor cellular transfection.Fluorescent polymers are a related class of fluorophores that can be either

intrinsically fluorescent such as CPs or functionalized with multiple fluorophores[3,130]. Similar to dendrimers, fluorescent polymers are characterized by largemolar absorption coefficients and therefore high fluorescence. However, due to theirinherent polydispersity the emission from fluorescent polymers is typically notlocalized, resulting from energy transfer processes along the whole chain, with a netresult of diffuse emission [131]. Thus fluorescent polymers cannot be consideredpoint donors for FRET. Nonetheless, fluorescent polymers, especially CPs, havefound application in a number of fluorescence-based studies, including FRET [130].Fluorescent CPs, cationic conjugated polymers (CCPs), and conjugated polyelec-trolytes (CPEs) are particularly popular for developing FRET-based DNA biosensors[132], with applications ranging from detection of DNA [133,134], DNA hybridiza-tion [135], DNAmethylation [136], and SNPs [137]. They have also been incorporatedinto sensing schemes for proteins [138] and Hg ions [89,139]. The group of McNeilland coworkers recently developed a series of multicolor conjugated polymer dots(CPdots) (Figure 6.8) [140–142]. These polymer dots, referred to as CPdots and laterPdots, are made from semiconducting polymer materials such as PPE, PFPV, orPFBT (see Figure 6.8 for full chemical name), are �4 nm in diameter with highfluorescent intensities, and could be readily functionalized [140–142]. Others havestudied these Pdotmaterials, including Chan et al. who developed FRET-based Pdotswith photoswitching capabilities [143,144]. These were created through the incor-poration of photochromic spiropyran molecules into a PFBT polymer and wereintended for use in bioimaging applications. In another example, a pH-sensing Pdotwas created by functionalizing PPE Pdots with pH-sensitive fluorescein [145].


6.3.6Photochromic Dyes

Materials that can reversibly switch between two forms/states upon exposure toelectromagnetic radiation, in which each form/state has different photophysicalproperties, are known as photochromic dyes [146,147]. While there are bothinorganic and organic photochromic materials, organic photochromic dyes arethe most popular, having a wide range of uses from decoration and eyeglass lens

Figure 6.8 Conjugated polymer dots (CPdots).(a) Chemical structures of fluorescent CPdot:the polyfluorene derivative poly(9,9-dioctylfluorenyl-2,7-diyl) (PFO), the copolymerpoly[{9,9-dioctyl-2,7-divinylene-fluorenylene}-alt-co-{2-methoxy-5-(2-ethylhexyloxy)-1,4-phenylene}] (PFPV), the poly[(9,9-dioctylfluorenyl-2,7-diyl)-co-(1,4-benzo-{2,10,3}-thiadiazole)] (PFBT), and the poly(phenylene

vinylene) derivative poly[2-methoxy-5-(2-ethylhexyloxy)-1,4-phenylenevinylene] (MEH-PPV). (b) Photographs of aqueous CPdotsuspensions under room light (left) and UVlight (right) illumination. (c) Absorption spectraand (d) fluorescence spectra of the conjugatedpolymer dots. (Reprinted with permission fromRef. [140]. Copyright 2008, American ChemicalSociety.)


coatings to optical switches and data storage. There are a number of photochromicdyes and various mechanisms that cause their photochromic transformations,and these have been extensively reviewed [146–150]. Of these, spiro-based (e.g.,spiropyran) and increasingly diarylethene-based photoswitchable compoundshave found particular application in FRET, so-called photochromic FRET(pcFRET) [147,149,151–153]. The pcFRET technique is particularly useful inFRET imaging applications, especially on a single-protein level, where pcFRETcan be used to turn the FRET process “off” or “on” thereby creating an internalcontrol and eliminating false-positive or false-negative signals due to highintrinsic autofluorescence, interactions with other endogenous proteins, and/or low FRET efficiencies [152,154–157]. FRET imaging sensitivity can be furtherenhanced when pcFRET is used in conjunction with techniques such as opticallock-in detection (OLID) [156,158].Spiro-based materials represent one of the more prominent types of photochro-

mic dyes, and exist in a closed spiro (colorless) form featuring an absorbance at<400 nm, which undergoes a ring-opening rearrangement upon UV exposure to anopen merocyanine (colored) form with an absorbance from 500 to 700 nm(Figure 6.9a) [151]. Photoswitchable spironaphthoxazine (NISO) has been conju-gated to tetramethylrhodamine (TMR) in a FRET-based strategy to enhance intra-cellular imaging [159]. When NISO is present in its merocyanine (colored) form, itacts as a FRET acceptor for TMR diminishing its emission. However, when it isswitched to its spiro (colorless) state, TMR donor emission increases due todiminished FRET. Spiropyran-based photochromic materials have been incorpo-rated into a number of FRET formulations, including hyperbranched polymermicelles containing a hydrophobic fluorescent dye nitrobenzoxadiazolyl derivative[160], gadolinium-complexed materials for use in magnetic resonance imaging(MRI)-based deep tissue gene expression mapping [161], and QDs [162]. Using aspiropyran-based nitrospirobenzopyran (Nitro-BIPS)-conjugated fluorescent pro-tein acceptor, FRET could be modulated with cycles of 365 and 546 nm light.This technique has been used to measure FRET efficiencies below 1% within acell [157]. Also utilizing photochromic BIPS, a photoswitchable QD (psQD) has beendeveloped for pcFRET [162]. By exposing the QD to white or UV light, the BIPS istransferred from colored merocyanine that acts as a FRET acceptor to colorlessspiropyran that will not act as a FRET acceptor to the QD donor. In this way theemission of the QD can be modulated. Spiropyran-based dyes have also beenincorporated into a number of luminescent NM-based probes for bioimaging[99,154,163,164]. Chen et al., for example, prepared FRET-based multicolor fluores-cent and photoswitchable polymer NPs by incorporating two fluorescent dyes (EANIand NBDAA) (Figure 6.9b) and photoswitchable spiropyran into methyl meth-acrylate-based NPs via copolymerization (Figure 6.9b) [99]. By varying the ratios ofthe dyes, the emission signatures could be tuned such that the NPs exhibit multiplecolors under a single excitation.Another increasingly common class of organic photochromic dyes includes the

diarylethene-based photoswitchable compounds, which, like the spiro-based mate-rials, undergo a structural open–closed photochromic transition [152,153,165–167].


Diarylethenematerials have been used to create photoswitchable dendrimers, wherethe diarylethene acts as a FRET acceptor that quenches the attached Cy3 donoremission via FRET, when switched off in the closed form [167]. Photochromic FRETimaging using this dendrimer was demonstrated within both HeLa cells and zebra

Figure 6.9 Spiro-based photochromic dyes. (a)The closed spiro form (SP – colorless)undergoes a ring-opening rearrangement uponUV exposure to the open merocyanine (MC –

colored) form. (b) Generation of FRET-basedmulticolor photoswitchable fluorescent NPs bycovalently incorporating two (EANI and SPMA)or three fluorescent dyes (EANI, NBDAA, andSPMA) under excitation at 385 nm [4-ethoxy-9-allyl-1,8-naphthalimide (EANI) and allyl-(7-nitro-benzo[1,2,5]oxadiazol-4-yl)-amine (NBDAA)].

(c) Fluorescence emission spectra of three NPsamples with different NBDAA feed (forsamples NP-N1, NP-N3, and NP-N5, theNBDAA feed increased at a certain value) aftervisible light irradiation and UV irradiation. (d)Photograph of three NP dispersions (NP-N1,NP-N3, and NP-N5) after visible lightirradiation and UV irradiation in the darkenvironment. (Reprinted with permission fromRef. [99]. Copyright 2012, American ChemicalSociety.)


fish, with the potential for use in detailed imaging of cellular processes within cellsor organisms. Photoswitchable QDs were developed by Diaz et al. by coating QDdonors with an amphiphilic polymer containing diheteroarylethene acceptors and aspectrally separate Alexa647 dye to act as an internal standard by facilitatingratiometric measurements (Figure 6.10) [165,166]. The properties of this NP couldmake it potentially useful for intracellular imaging applications.

Figure 6.10 Diarylethene-based photochromicdyes, and generation of photoswitchable QDs(psQDs). (a) The open diarylethene form(oPC – colorless) undergoes a ring-closingrearrangement upon UV exposure to the closeddiarylethene form (cPC – colored). (b) Spectralsignatures of the dual-color psQD components.Superposition of absorbance (solid lines) andemission (filled areas) spectra of PC, QD, andAlexa647, demonstrating the PC spectraloverlap with the QD but not with the Alexa647.The spectra are normalized by their peak values.

(c) Schematic of the dual-color psQD. Thefluorescence of the QD is modulated by thephotoconversion of the PC, while the Alexa647fluorescence is constant. The PC in the openform (oPC) is photoconverted with UVirradiation to the closed form (cPC), whichcan then be photoreversed by direct excitationwith visible light, or via FRET from the QDacting as a donor. (Reprinted withpermission from Ref. [165]. Copyright 2012,American Chemical Society.)


In addition to organic dyes for pcFRET, photoswitchable FPs that can be encodedin the genome have also been demonstrated, and are discussed in more detail inSection 6.4.3 [155,168,169]. The use of pcFRET shows great promise for sensitive,high-resolution (diffraction unlimited) imaging of cellular processes, including rareprotein interactions and individual protein movement within a cell. As a greatertoolbox of photoswitchable materials for pcFRETcontinues to develop and research-ers work on improving their inherent properties, pcFRET applications are sure tocontinue to grow.

6.3.7Carbon Nanomaterials

Carbon NMs represent a diverse class of materials with a variety of differing physicaland chemical properties. This diversity is a direct result of a range of allotropiccarbon material forms, including diamond, fullerene spheres and nanotubes,graphite and graphene, along with amorphous carbon. Of these many types,graphene-based sheets, nanodiamonds (NDs), luminescent carbon nanodots (C-dots), carbon NPs (CNPs), and carbon nanotubes (CNTs) possess relevant opticalproperties of interest for FRET applications.Of the graphene family of materials, graphene oxide (GO) sheets are reportedly

fluorescence “superquenchers” that possess long-range energy transfer propertiesthat make them ideal in FRET studies [170]. Most of the GO-based FRET sensors todate use DNA-based molecular recognition, in the form of molecular beacons,aptamers, or DNAzymes, for the specific detection of a range of target analytes fromsmall molecules, such as heavy metals and mycotoxins, to proteins includingthrombin, DNA, and even whole cancer cells [170–175]. Donor species rangefrom traditional organic fluorophores to QDs and upconverting NPs (UCNPs);the diversity reflects the superquenching abilities of the GOmaterials. Wu et al., forexample, demonstrated the simultaneous detection of the mycotoxins ochratoxin A(OTA) and fumonisin B (FB1) using two types of UCNPs modified with specificaptamers [173]. GO was used as the universal acceptor in the sensing scheme(Figure 6.11). In the absence of the target mycotoxins, the aptamer-modifed UCNPsinteracted with the GO surface resulting in FRET and effective quenching of theUCNP luminescence. Addition of the mycotoxins, which bind to the aptamer-modifed UCNP, altered the GO–aptamer-modifed UCNP interaction, resulting inan off–on sensor whose resulting luminescence spectra was mycotoxin specific.Dependent on the synthetic approach, GO sheets, especially the reduced form, in

suspension or solid thin films can exhibit luminescent properties and have beenused as donors in combination with Au NP acceptors for FRET detection of DNAhybridization and microcystins [176–178]. Graphene sheets smaller than 10 nmhave also been found to possess photoluminescent (PL) properties [includingupconversion (UC) and downconversion PL], and are referred to as graphenequantum dots (GQDs) [179,180]. Fan et al. found that 2,4,6-trinitrotoluene (TNT)effectively quenched GQDs luminescence via FRETupon the p–p stacking interac-tion that occurs between the two species [181].


CNTs comprise graphene tubes and, like the sheets, can either possess lumines-cent or superquenching properties depending upon morphology, synthesis, andpurity. In general single-walled CNTs (SWCNTs) and sometimes double-walledCNTs (DWCNTs) are found to have luminescent properties [182], while multiwalledCNTs (MWCNTs) are considered superquenchers [183]. CNTs have successfullybeen used as donors and acceptors in FRET applications where they have beencoupled with traditional organic dyes, QDs, and even lanthanide ions [184–187]. Forexample, QD-labeled ssDNA is found to undergo a strong interaction with CNTs,resulting in significant quenching of the QD luminescence [187]. Binding of thetarget influenza A virus DNA resulted in a significant decrease in the DNA–CNTinteraction and an increase in QD emission, with a limit of detection (LOD) of9.4 nM and excellent single-base mismatch discrimination. Synthesis and purity ofthe CNTs appear to be a key requirement for their successful application in FRET.While studying the fluorescence quenching of the dyes dansyl hydrazine andpanacyl bromide covalently attached to SWCNTs, Chiu et al. found that the panacylbromide quenching, unlike dansyl hydrazine, was very sensitive to the CNTpurification method, specifically the metal impurities left over from CNTmanufac-ture, suggesting care should be taken when interpreting data [185].As an alternative to CNTs and GO sheets, which can be quite large as discrete

labels, CNPs, NDs, and C-dots are relatively small and compact labels that are

Figure 6.11 GO sheets as universalquenchers in FRET-based assays. (a)Preparation of aptamers–UCNPs formycotoxin detection. (b) Schematicillustration of the multiplexed upconversion

FRET bioassay using aptamers–UCNPs(donors) and GO (universal acceptor) forFB1 and OTA detection. (Reprinted withpermission from Ref. [173]. Copyright 2012,American Chemical Society.)


starting to find application in FRET [188–191]. CNP acceptors (quenchers) coupledwith UCNPs have been applied to the detection of thrombin, using aptamers, andmatrix metalloproteinase-2 (MMP-2), using a polypeptide substrate [192,193]. NDs,whose intense fluorescence properties arising from nitrogen-vacancy (NV) pointdefects in their nanocrystalline structure, have been investigated as donors in FRETstudies with a number of near-IR dyes [194–196]. C-dots that are sub-10 nm particlesthat become fluorescent upon surface passivation have yet to be applied to FRETapplications, but like NDs have great potential as donors.Many of these carbon NMs are still relatively new and studying their inherent

physical properties and understanding themechanisms that govern them is still verymuch a work in progress. This is hampered somewhat by the fairly complex andnormally poorly controlled methods typically used to generate carbon NMs, such aschemical vapor deposition, electric arc discharge, or laser ablation. These methodstypically produce a range of products containing a variety of impurities that have toundergo some type of purification in order to obtain the desired end product. Thatsaid, there are an increasing number of manufacturers who offer various carbonNMs (e.g., Sigma-Aldrich, Carbon Solutions Inc., Nano-C1, Microdiamant, andNanoAmor: Nanostructured and Amorphous Materials Inc.), oftentimes premodi-fied with functional groups that aid in solubility and bioconjugation, which mayencourage more widespread application.

6.4Biological Materials

Biological materials and biologically inspired materials (i.e., nonnatural aminoacids), similar to the organic materials, are a diverse group including, molecules,proteins, and protein complexes. In addition, biological reactions creating bio- orchemiluminescence are an interesting alternative to the requirement for an externalexcitation source.

6.4.1Natural Fluorophores

Of the various naturally occurring fluorophores, including certain amino acidresidues, reduced nicotinamide cofactors (NADH and NADPH), flavins (FADand FMN), porphyrins, and pyridoxal derivatives, it is the aromatic amino acids,Trp, tyrosine (Tyr), and Phe shown in Figure 6.12 that dominate FRET applications[47]. The primary advantage to using these naturally occurring amino acids is thatthey have an endogenous presence in proteins, and when combined with FRETanalysis can be used to study protein structure and dynamics [197]. Imaging studiesof proteins can thus be completed with limited to no modification and even if theseresidues are not present in a protein, they generally can be incorporated into thepeptide sequence with a minimal effect on its size, structure, and subsequentinteractions. The strong UV absorbance of proteins at 280 nm (commonly used for


quantitation) as well as an emission at �340–360 nm, originate mostly from theindole ring of Trp; Tyr and Phe contribute to amuch lesser extent [47]. The negligibleQY (�0.02) of Phe makes it less amenable to FRET, except perhaps in intraproteinconfigurations. Tyr is prone to quenching and energy transfer to Trp, leaving Trp asthe most reliable residue for FRET (detailed in Refs [47,50,198]). A potential liabilityin the use of Trp for FRET is that the excitation lines and any donor/acceptor dyeswill be confined to the UV region. The fluorescence from these residues is alsoenvironmentally sensitive and so their placement deep within a protein structurewill produce results that differ from those at the terminus of a small peptide.The primary application of endogenous fluorophores, in particular Trp, is to study

the structure and function of proteins and peptides. FRET exchanges often occurbetween Trp and other fluorophores [such as nonnatural amino acids (see Section6.4.2), or organic dyes], but FRET can also occur between two Trp residues, termedhomo-FRET. Because of the low QYof Trp and its random distribution in moleculesof interest, this homo-FRET rarely occurs. However, Kayser et al. found it to be aparticularly useful tool for studying structure–function relationships in monoclonalantibodies, which were found to have an unusually high Trp content [199]. Anotheruse of Trp as a natural fluorophore for FRET is to probe protein folding/unfolding,an important yet poorly understood biological process [200–202]. For example, Jhaet al. used FRET to study the unfolding of a small protein, monellin [202]. Thetransition of this small protein, from an unfolded to a folded state, is not completelyunderstood but by using a naturally occurring Trp and additionally labeling theprotein with the FRET acceptor thionitrobenzoate, researchers were able to deter-mine that monellin unfolds gradually rather than all in onemotion [202]. Visser et al.utilized Trp homo-FRET to characterize protein folding of apoflavodoxin, demon-strating the potential of this method as a powerful means to understand some of thestill unknown mechanisms of protein folding [201]. Another salient exampleleveraging Trp’s fluorescent properties to determine protein structure and functionis its use in understanding the membrane transport protein LacY [203]. LacY is asugar/Hþ symporter of E. coli bacteria and is widely studied. Trp fluorescence orquenching was used to determine the conformation of LacY and the function of the

Figure 6.12 Naturally fluorescent amino acid residues. Structures of the aromatic amino acids,tryptophan (Trp), tyrosine (Tyr), and phenylalanine (Phe).

6.4 Biological Materials j189

symporter. Understanding protein structural variations could also aid in determin-ing the pathogenic process of certain diseases. Lee et al., studying the humana-synuclein protein associated with Parkinson’s disease, used FRET between a fixedTrp donor and a modified 3-nitrotyrosine acceptor to demonstrate an elongatedstructure for the mutant protein associated with the disease [204].While amino acids tend to dominate endogenous fluorophore FRET research,

porphyrins, which physiologically form transition metal complexes [e.g., iron (seeSection 6.5.2)], are found to have strong luminescent properties and have beendemonstrated as FRET acceptors and donors in various formats. Lovell et al., forexample, employed a caspase-3-specific peptide sequence modified with a rho-damine donor and porphyrin acceptor to monitor caspase activation in single cellsfollowing induction of cell death [205]. In the pursuit of artificial light-harvestingsystems, porphyrin donors immobilized onto a clay surface (acceptor) were found toreach energy transfer efficiencies approaching the ideal 100% [206].

6.4.2Nonnatural Amino Acids

Intrinsic natural probes such as Trp, Tyr, and Phe are highly useful for visualizationof protein structure, movements, and interactions, and as mentioned are eithernaturally occurring or can be introduced without significantly impacting the proteinstructure. However, due to low QYs of both Phe and Tyr, Trp is the only widely usednatural fluorescent probe. To overcome this limitation nonnatural (also calledunnatural or noncanonical) amino acids have been fabricated [207–213]. Thesenonnatural amino acids can provide larger QYs and new FRET pairs for proteinstructure function analysis. One recently developed fluorescent nonnatural aminoacid p-cyanophenylalanine (PheCN) can be incorporated into a protein, minimallydisturbing the native protein structure [214], and can act as a FRET donor to Trp(Figure 6.13) [207]. The photophysics of this useful nonnatural amino acid hasrecently been characterized [211,215]. The Trp–PheCN FRET pair has been used todetermine detailed protein folding and unfolding in two small proteins, the villinheadpiece subdomain (HP35) and the lysin motif (LysM) domain [200]. Two othernonnatural amino acids, 7-azatryptophan (7AW) and 5-hydroxytryptophan (5HW)(Figure 6.13), were recently found to be FRET acceptors to PheCN [210]. The7AW–PheCN FRET pair had a greater separation of fluorescent spectrums thanPheCN–Trp. Moreover PheCN, Trp, and 7AW can be used in a multistep FRETsystem to investigate interactions of three points on a protein (see Section 6.6) [210].Even newer nonnatural FRET pairs are continuously being developed. Recently,L-4-cyanophenylalanine (pCNPhe) and 4-ethynylphenylalanine (pENPhe) were usedas FRET donors to Trp in order to probe the hydrophobic core of the proteinT4 lysozyme [216]. Another nonnatural amino acid was created by combiningp-aminophenylalanine derivatives with BODIPY fluorophores, generating amaterialwith an emission wavelength greater than 500 nm [208]. These amino acids wereincorporated into a calmodulin-binding peptide and FRET used to probe proteinbinding and resulting conformational changes.


Figure 6.13 Nonnatural amino acids. (a)Structures of some selected nonnatural aminoacids, p-cyanophenylalanine (PheCN), 7-azatryptophan (7AW), and 5-hydroxytryptophan(5HW), derived from the naturally fluorescentamino acids with the chemical modificationsshown in red. (b) Improving Stokes shift ineCFP using FRET with a nonnatural amino acid.Three-dimensional model of eCFP, whichcarries the fluorescent amino acid (1) at thesurface-exposed position 39, is based on thecrystal structure of eCFP (PDB entry 2WSN).Chemical structures of (bottom) the eCFPfluorophore 4-[(1H-indol-3-yl)methylidene]

imidazolin-5-one (5; lex¼ 434 nm,lem¼ 476 nm) and the fluorescent nonnaturalamino acid L-(7-hydroxycoumarin- 4-yl)ethylglycine (1; lex¼ 360 nm, lem¼ 450 nm)that together form a FRET pair. Normalizedabsorption and fluorescence spectra of 1 [Abs(1) and Flu(1)] and eCFP [Abs(eCFP) and Flu(eCFP)]. The major absorption band of eCFPshows considerable overlap with thefluorescence spectrum of 1, thus fulfilling aprerequisite for efficient FRET. (Reprinted withpermission from Ref. [217]. Copyright 2011,American Chemical Society.)


Nonnatural amino acids can also be used to modify FPs to enhance theirfluorescent properties via intramolecular FRET [209]. The nonnatural amino acidL-(7-hydroxycoumarin-4-yl)ethylglycine, for example, was incorporated into recom-binant cyan FP (CFP) [217]. The modified FP underwent FRET between thenonnatural amino acid (donor) and the FP’s natural chromophore (acceptor),resulting in emission at 476 nm with a 365 nm excitation wavelength (Figure 6.13).This large apparent Stokes shift of 110 nm is much greater than the natural 40 nmStokes shift of CFP alone.Nonnatural amino acids have also been applied to time-resolved FRET (TR-FRET),

which can be used to evaluate protein folding dynamics. Historically, Trp has beenused as a donor for TR-FRETmeasurements; however, a significant drawback is thatit exhibits a high degree of variation in its fluorescent lifetime, depending on proteinconformations. In an attempt to overcome this, an analogue of Trp, 5-fluorotrypto-phan (5F-Trp) has been proposed as a better candidate for TR-FRET [212]. The 5F-Trp has more homogenous decay kinetics than Trp and is less environmentallysensitive, making it an ideal donor for TR-FRET for the determination of molecularstructure in proteins.As new nonnatural fluorophores are designed, and the study of these and

natural fluorophores progress, these materials could provide an evengreater utility for understanding proteins at a molecular level utilizing FRETtechniques.

6.4.3Green Fluorescent Protein and Derivatives

FPs represent an increasingly diverse class of fluorophores that have shown greatpotential as genetically encoded fluorescent tags for assessing protein location andfunction (monitoring protein–protein interactions) in cell studies and the devel-opment of in vivo (signaling dynamics such as calcium ions) and in vitrobiosensors [4,22,168,218–221]. While GFP derived from the jellyfish Aequoreavictoria represents the prototypical fluorophore of this protein family, various GFPmutations and Anthozoa (coral) homologues provide an increasingly diverserange of photophysical properties (Table 6.1 and Figure 6.14), which stemfrom their internal chromophores [155,168,169,218,222–224]. Newman et al.give an excellent monograph of the FP basics [168]. To briefly summarize, FPsself-generate their intrinsic chromophore from key internal amino acid residues,which nestle deep in the core of the characteristic 11-stranded b-barrel FPstructure. The final photophysical properties of the mature FP are governed bythe extent of p-conjugation, subsequent chromophore transitions, and interac-tions with the surrounding amino acid microenvironment with the chromophore(Figure 6.14) [155,168,169].Key to the success of FPs has been the ability to genetically encode them via

commercial plasmids that can be expressed in a variety of cells/organisms(available from ClonTech Laboratories, Inc., Life Technologies, Evrogen, andMBL Intl. Corp.). The genetically encoded FPs are commonly attached to


Table 6.1 Properties of some representative FPs summarized from www.microscopyu.com andRef. [168].

Protein Source In vivostructure

Absorbancemax. nm

Emissionmax. nm

ExtinctioncoefficientM – 1 cm – 1

Quantumyield

Blue

EBFP A. victoria Monomera) 383 445 29 900 0.31TagBFP E. quadricolor Monomer 399 456 52 000 0.63mBlueberry2 D. striata Monomer 402 467 51 000 0.48ECFP A. victoria Monomera) 439 476 32 500 0.4CyPet A. victoria Monomera) 435 477 35 000 0.51TagCFP E. quadricolor Monomer 458 480 37 000 0.57

GreenGFP (wt) A. victoria Monomera) 395/475 509 21 000 0.77Turbo GFP Copepod sp. Dimer 482 502 70 000 0.53EGFP A. victoria Monomera) 484 507 56 000 0.6TagGFP2 E. quadricolor Monomer 483 506 56 500 0.6mWasabi Clavularia

coralMonomer 493 509 70 000 0.8

Yellow

TagYFP E. quadricolor Monomer 508 524 64 000 0.6EYFP A. victoria Monomera) 514 527 83 400 0.61mCritrine A. victoria Monomer 516 529 77 000 0.76Ypet A. victoria Monomera) 517 530 104 000 0.77

Orange

mBanana D. striata Monomer 540 553 6000 0.7mOrange D. striata Monomer 548 562 71 000 0.69OFP Cerianthus sp. Tetramer 548 573 60 000 0.64TurboRFP E. quadricolor Dimer 553 574 92 000 0.67

Red

DsRed D. striata Tetramer 558 583 75 000 0.79TagRFP E. quadricolor Monomer 555 584 100 000 0.48mTangerine D. striata Monomer 568 585 38 000 0.3AsRed2 A. sulcata Tetramer 576 592 56 200 0.05mStrawberry D. striata Monomer 574 596 90 000 0.29mCherry D. striata Monomer 587 610 72 000 0.22

Near-IR

mRaspberry D. striata Monomer 598 625 86 000 0.15mPlum D. striata Monomer 590 649 41 000 0.1mNeptune E. quadricolor Monomer 600 650 67 000 0.2eqFP670 E. quadricolor Dimer 605 670 70 000 0.06

a) Weak Dimer formation.b) E ¼ Entacmaea genus.c) D ¼ Discosoma genus.


http://www.microscopyu.com

proteins of interest through the creation of FP chimeras that can subsequently beused in cellular studies of protein location, protein–protein interactions, and thedevelopment of biosensors to monitor cell signaling processes [4,168,219,220]. Inaddition to the benefit of genetic manipulation, there are a number of advantagesand disadvantages to the use of FPs as fluorescent tags that should be factoredinto their use as FRET donors/acceptors. FPs have a wide range of QYs (seeTable 6.1), ranging from 0.04 for AQ143 to 0.91 for ZsGreen, which depend on

Figure 6.14 Fluorescent proteins. (a)Structure of A. victoria GFP showing thedimensions of the protein, the intrinsicallyderived p-HBI chromophore, and severalkey residues surrounding thechromophore (image generated usingPyMOL open access and PDB ID 1w7s).

(b) Chromophore structures of representativeFP color variants within each spectral class.The conjugated ring structure of eachchromophore is colored according to itsemission profile. (Reprinted with permissionfrom Ref. [168]. Copyright 2011, AmericanChemical Society.)


the mutations present and final chromophore structure, although the majorityare generally good with QYs > 0.5 [168]. Certain FPs have been found to possesstwo-photon absorption properties that can be very advantageous in deep tissueimaging applications [225,226].Photophysically, FPs can take several hours to fully mature as the final

chromophore is formed through the protein folding process and any subsequentchemical transitions [155,169,168,221]. The relative brightness of their fluores-cence intensity is found to be intimately linked to the efficiency of the FP foldingprocess and mutation time [155,168,221]. FPs generally have relatively broadabsorption and emission profiles that may preclude “multiplex” analysis. FPs arealso prone to photobleaching and have known susceptibilities to pH, tempera-ture, O2 concentration, and other environmental conditions. FPs are of a fairlylarge size from a fluorescent tag point of view, Mw �25–30 kD and upwards,which can be problematic in terms of maintaining the desired function of thelabeled target protein. In addition, certain FPs have a tendency toward theformation of oligomers (dimers and tetramers), which can further confoundthe size issue [168]. Also, location of the FP-tag in the target protein must becarefully chosen so as not to significantly impact FP maturation and thereforebrightness [168]. Researchers continue to develop FP mutations that attempt toaddress a number of these issues, including improved photostability [223],decreased oligomerization [222], emission in the near–far IR region [169,227],improved Stokes shifts [155], or photoactivatable (including photochromic)properties [155,168,169,228].Use of FP pairs in FRET and bioluminescence resonance energy transfer (BRET)

(see Section 6.4.6) applications is ever expanding, with dramatic implications for invivo imaging, biosensors, and cellular studies in particular [22,155,168,219,229–235].FPshave revolutionized thedetection and study of cellular events, and themore recentcombination of FPs and FRET imaging has taken this detection to new levels ofprecision, allowing the study of protein–protein interactions and tracking biochemicaland protein signaling dynamics, reviewed in a number of excellent publications[22,155,168,219,220,229,232,236]. There are twomain FRET–FP strategies employedwhen studying cellular processes, the nature of which is dependent on the processbeing investigated. In the first strategy, typically employed for measuring inter-molecular protein–protein interactions, it is common to tag each protein with eitherthe donor or acceptor FP (Figure 6.15a), as FRETemission will only take place whenthe two (or more) proteins of interest interact. In the second strategy a protein orbiosensor construct is labeled with both the donor and acceptor FPs, and interactionwith the target species of interest results in some measurable change in the FRETsignal. There are a number of different biological processes that can be monitoredusing the second strategy, including protease activity and activation, Ca2þ ionfluctuations,measuring secondmessengers such as cyclic adenosinemonophosphate(cAMP) and cyclic guanosine monophosphate (cGMP), studying phosphoinositidedynamics, andGprotein-coupled receptor (GPCR) activation, andhence awider rangeof formats are/can be employed, some of which are highlighted in Figure 6.15b, andrecently reviewed in Refs [22,168,236].


Figure 6.15 Representative FRET-basedsensor formats that incorporate FPs. (a)Monitoring protein–protein interactions. Here,each protein is labeled with a FP that uponinteraction results in FRET. (b) Biosensors inwhich binding of a small molecule induces theassociation of two distinct moieties within thesingle polypeptide chain. (c) Biosensors forposttranslational modification. (d) Biosensors

in which a protein undergoes a conformationalchange upon binding its small-moleculeligand. (e) Biosensors for protease activity.The donor is CFP and the acceptor is YFP inthese representations, however, a variety ofother FP FRET combinations could besubstituted. (Reprinted with permissionfrom Ref. [22]. Copyright 2009, AmericanChemical Society.)


While the use of FP pairs in FRET has been constantly expanding, advances needto be made continually to enhance its effectiveness. Shortcomings in FP–FRETinclude a large range in brightness in FPs, cross talk due to large emission spectra ofFPs, slow development of intramolecular sensors, and limited ability to create stablytransfected cells lines with FRET FPs. Though recent advances have achieved somesuccess in expressing FP–FRET pairs in a cell line [237,238], difficulty still exists inachieving this goal [236]. With advances in developing cell lines to express FP–FRETpairs, more research can be conducted using these systems, as researchers will beable to more readily conduct experiments without the difficulty of transientlytransfecting cells. Brightness and cross talk issues can be mitigated by furtherimproving FPs for FRET. Development of additional FPs for FRET has also led tomultiparameter imaging using dual FRETpairs. Development of this technology hasimportant research implications as entire signaling cascades could potentially beimaged in the same cell. The most commonly used FRET pair is CFP–yellow FP(YFP), but even with this pair significant cross talk exists [229]. New, recentlydeveloped FPs include enhanced GFP (EGFP) and mCherry, which have similarbrightness and reduced cross talk compared to CFP–YFP [239]. Lam et al. demon-strated a Clover–mRuby2 FP–FRET pair combination for monitoring kinase aci-tivity, guanosine triphosphate hydrolase (GTPase) activity, and transmembranevoltage with significantly improved photostability, FRET dynamic range, and emis-sion ratio changes versus CFP–YFP [224]. An orange fluorescent protein with a largeStokes shift (LSSmOrange) has been developed, which can be used for intracellularimaging, potentially allowing two FRET pairs in combination with CFP–YFP [240].In a salient example of multiparameter imaging, four different cellular events wererecorded simultaneously using FP–FRET imaging [241]. Here, two CFP–YFP FRETsensors that could be spatially resolved were combined with a spectrally distinctmCherry–mORange FRET pair and a fourth sensor Fura Red. The development ofnew FRET protein pairs for multiple parameter imaging could greatly expand theuse of FP–FRET and will accelerate discovery of cellular processes [242], cancerresearch [236], and toxin detection [243].Photoswitchable FPs, a subset of the larger FP community, for pcFRET have also

been demonstrated and have great potential for high-resolution imaging applica-tions [155,168,169]. Photochromic/photoswitchable FPs are under continuousdevelopment, and the various mechanisms/factors that govern photoswitchinghave recently been reviewed [155,168,244]. Switching is thought to occur primarilythrough a cis–trans isomerization of the FP chromophore, as demonstrated withisolated synthetic chromophore analogues [245,246], however the chromophoreenvironment within the FP structure also plays a key role in determining a FPsswitchability [78]. Recently, it has been discovered that substitution of certain keyamino acids in the chromophore environment within the FP structure can improveand/or restore the photochromic behavior of the FP chromophore, leading toimproved photoswitchable FP mutations for pcFRET studies [78,244,247]. Grotjo-hann et al., for example, generated a reversibly switchable EGFP (rsEGFP) mutantthat could be cycled �1200 times before experiencing a 50% reduction in


fluorescence, in stark contrast to FP Dronpa that experiences a 50% reduction afteronly 10 cycles [247]. The red FP (rsTagRFP) can be photoswitched from “on” and“off” fluorescent states using 445 and 570 nm lights, respectively [248]. ThersTagRFP was subsequently used as an acceptor with an enhanced YFP (EYFP)donor to monitor protein interactions in pcFRET studies of living cells, where theon–off switching provided confirmation of the FRET signal and the proteininteraction. Other photoswitchable FPs include Dronpa [249] (and variantsbsDronpa and Padron [228]), rsCherry [250], and rsCherryRev [250].While pairing two FPs for FRET is the most common combination when

utilizing FPs, there are increasing examples of FP donors or acceptors beingpaired with other fluorescent materials for FRETapplications. There are a numberof BRETexamples discussed in Section 6.4.6 and also multi-FRETexamples wherethe FP is coupled with light-harvesting complexes or proteins (see Section 6.6).Rice created a kinesin C-terminal GFP fusion and labeled the kinesin withtetramethylrhodamine, allowing FRET monitoring of protein conformationalchanges upon binding nucleotides [251]. Hoffman dual labeled a GPCR systemwith CFP and the FlAsH system to monitor receptor activation demonstratingthat, unlike the equivalent CFP–YFP-labeled system, downstream signaling wasnot disrupted by the FlAsH acceptor [252]. FP acceptors combined with QDdonors are becoming an increasingly common combination, especially in thedevelopment of biosensors for measuring protease activity [230] and intracellularpH (Figure 6.16) [79].Clearly, FP-based FRET has already made a significant impact on our under-

standing of cellular processes, and as the materials themselves continue to evolveand improve, more sophisticated applications can be expected.

Figure 6.16 QD–FP FRET-based pH sensor.(a) Schematic demonstration of the pH-dependent energy transfer between the QD andthe FP. In an acidic environment, energytransfer to the FP FRET acceptor is minimal,yielding a high QD signal; at neutral or basicpH, energy transfer is more efficient, producingan enhanced FRET signal. (b) A pH titration ofQD–FP probes containing the FP acceptormOrange M163K showing increased energytransfer at alkaline pHs with a clear isosbesticpoint. Cellular imaging of QD–mOrange pHsensor. (c) Schematic of probe color changesduring progression through the endocyticpathway. FRET efficiency is high in the neutralpH of the extracellular environment and earlyendosome. FRET efficiency decreases as the

endosome matures and the endosomal pHdrops, resulting in diminished emission frommOrange and recovery of some QD signal. Anyprobe that escapes the endosome regains itselevated FRET efficiency in the pH neutralcytoplasm. (d) Fluorescence microscopyimages immediately after delivery of the probeand 2 h post delivery. The QD images (left)demonstrate consolidation of the probe inthe endosomes over time; images of thedirect excitation of mOrange (center) and FRETemission (right) indicate a clear decrease inthe mOrange emission and the FRETefficiency of the probe with maturation of theendosome. (Reprinted with permissionfrom Ref. [79]. Copyright 2012, AmericanChemical Society.)

I



6.4.4Light-Harvesting Proteins

Phycob iliproteins (PBPs) are the colored component prot eins found in natu rallyoccurring ph yc obil isome -b as ed li ght-ha rves ting c omplexes (se e Se ction 6. 6)[253,254]. T heir intense c olor originate s from the combination of ch romophoresthe y contain, te rmed phycobilins. Examples of phycobilins include ph yc ocyano-bi lin ( PCB – blue ), phycoerythrobilin (PEB – red), p hycourobilin (PUB – yellow ),and phyc obiliviolin (PXB – purple). In phycobilisome-based s ystems, the re arefour ma in PBP s: allop hycocyanin (APC), phycocyanin (PC), phycoerythrin (P E),and phycoerythrocyani n (PEC); s ome s pectrosc opic details a re summarized i nTa ble 6.2 [254,255]. Pe ridinin– chlorophyll– protein (Per CP or PCP) is a light-h a r v e s t i ng c o m p l e x f o u nd i n d i no fl agellates and has a smaller M W (35 kDa) thanmost of t he 100 – 240 kDa PBPs mentioned, which can b e bene fi cial whe n t he sizeof the fl u ores cent lab el is a concern. Sin ce t heir i nception as fl uorescent prob es,the pu ri fi ed PBP s themselves (e .g., phyco fl uor prob es), compl exes c ont aini ngmultiple P BPs (e.g., PBXL), or the smaller Per CP materials h ave b ecomecommon fl uores cent lab els in bioas says , espe cially in fl ow cytometry[254,256– 259]. The ir use a s fl uorophores in F RET has b een more limited todate, a nd may b e due in part to t heir relatively large size – although some of th esmaller materials being develop ed (e .g., Per CP and CryptoFluor TM ) may addressthis c onc ern. That said, th ey have be en demonst rated i n sandwic h-based p eptid eand a ntibody FRET a ssays for target analyte detect ion and f or de mons tratingFRET in combination with Au a nd QD NMs [121,257,260– 264]. The combinationof APC (acceptor) and Eu-cryp t ate (donor ) h as b een use d for th e time-res olve dFRET-based detection of telomerase activity [262], inhibitors of hepatitis C viruscore dimerization [264], and small-molecule inhibitors of HIV-1 fusion [263]. Anumber of companies sell PBP- and PerCP-based materials, including Life

Table 6.2 Properties of some representative phycobiliproteins summarized fromwww.columbiabiosciences.com and Ref. [254].

Protein Approx.MWkDa

Types ofphycobilinspresent

Approximatenumber ofphycobilins

Absorbancemax. nm

Emissionmax. nm

ExtinctioncoefficientM – 1 cm – 1

Quantumyield

Allophycocyanin(APC)

100 PCB 6 652 657.5 2.4� 105 0.68625a)

R-Phycoerythrin(RPE)

240 PEB andPUB

34 565 573 1.96� 106 0.84498a)

B-Phycoerythrin(RPE)

240 PEB andPUB

34 545 572 2.41� 106 0.98563.5a)

Note: Exact properties are dependent on the origin of the protein PCB: phyocyanobilin, PEB:phycoerythrobilin, and PUB: phycourobilin.a) Additional absorbance peak.


http://www.columbiabiosciences.com

Technologies, Jackson ImmunoResearch (mainly streptavidin antibody-labeledand secondary antibody-labeled materials), the SureLight1 series from ColumbiaBiosciences, and the CryptoFluor materials from Martek Biosciences Corp. (soldby Sigma-Aldrich).

6.4.5DNA-Based Macrostructures/Nanotechnology

DNA is an incredibly complex polymeric material comprising four monomers –adenine (A), thymine (T), guanine (G), and cytosine (C) – that can be singlestranded (ss) or doubled stranded (ds), which results when two complementarystrands hybridize [265]. Researchers are increasingly interested in using thepolymeric nature of DNA to synthesize unique 2D and 3D DNA-based macro-molecular/nano structures [265–268]. The use of functionalized DNA nano-structures for light harvesting and charge separation has recently beenreviewed [269]. Researchers have already demonstrated the potential of theseDNA structures as fluorescent labels using some relatively simple DNAconstructs. Accumulation of perylene- and pyrene-based fluorophores inDNA duplexes has been shown and produces fluorescent excimer structuresvery strongly, which possess large Stokes shifts, making them ideal donorsin FRET applications [270,271]. Kumar and Duff engineered some uniqueDNA–protein complexes that demonstrated potential as light-harvesting com-plexes [272]. The Armitage group developed DNA tetrahedron and duplexfluorescent nanotags using FRET to shift the emission of the DNA nanotagfurther into the red region relative to the donor dye alone (Figure 6.17)[273–275]. Although the majority of these DNA-based structures are designedin-house, the DNA sequences themselves are often synthesized and purchasedcommercially [ from companies such as Integrated DNA Technologies (IDT)].Genisphere1 sells a 3DNA-based dendrimer that is marketed for signalamplification in a number of bioassays. Given the ease with which fluorescentdyes can be incorporated into these unique structures, through either interca-lation or covalent attachment, it seems likely that the utility of these materialsas fluorescent labels and their subsequent use in FRET-based applications willincrease in the future.

6.4.6Enzyme-Generated Bioluminescence

Enzyme-generated bioluminescence (BL) is used in a particularly advantageousvariant of FRET known as BRET. BL is a naturally occurring phenomenon foundin certain beetles and bacterial or marine species, where various substrates(luciferins) react with enzymes (luciferases) in the presence of O2 (and sometimesother cofactors) to produce light emission (Table 6.3) [276–281]. The exact wave-length of the light emission is found to be dependent on a number of factors,


Figure 6.17 DNA NMs for FRET. (a) Assemblyof DNA tetrahedron nanotags. Four strandswith partially complementary sequences formthe DNA tetrahedron nanostructure templatefor the self-assembly of intercalator dyes. Blacksections represent two-nucleotide long, single-stranded hinges. (b) Schematic description ofET (energy transfer) in a tetrahedron nanotagloaded with YOYO-1 intercalated dyes andcovalently attached Cy3 acceptor dyes.

(c) Fluorescence emission of tetrahedronnanotags with 0–4 covalently attached Cy3molecules. Spectra acquired by excitation at440 nm. Samples contained 50 nM DNAtetrahedron and 1.28M YOYO-1. ET efficienciesgiven in legend were determined by thepercentage of decrease in the YOYO-1 emissionat 509 nm. (Reprinted with permission fromRef. [274]. Copyright 2009, American ChemicalSociety.)


including the structure of the luciferin, the nature of the luciferase, and thepresence of accessory proteins, physiologically, GFP or its derivatives [277]. BLitself has found application as a reporter in many bioassays, both in vitro andin vivo [278–282].The use of the BL reaction as a donor within energy transfer mechanisms is, in

fact, an intrinsic process observed in many sea creatures such as A. victoria(jellyfish) and Renilla reniformis (sea pansy), where accessory proteins (e.g., GFP)modify the color of the emission through BRET [277]. Researchers have usedBRET in a number of applications, most notability for studying protein–proteininteractions, leading to drug discovery, and increasingly for biosensing and in vivoimaging [283–288]. As with FRET, BRET depends upon spectral overlap betweenthe donor emission and the acceptor absorption, and it is similarly efficient overdistances up to 10 nm [278,284,287]. The principal advantage of BRET is removalof the excitation source, negating problems such as light scattering, high back-ground noise, direct acceptor excitation, photodamage to cells, and photobleach-ing effects [287,289].The most commonly exploited luciferases for BL are the eukaryotic firefly and

Renilla luciferases (Rluc), with the wild-type enzymes generating blue-green emis-sion. DNA vectors with the desired luciferase gene or plasmids can be purchasedthrough a variety of sources such as Promega, New England Biolabs, and TargetingSystems. These vector complexes are then internalized by the cell of choice, wherethe luciferase gene can be transcribed by ribosomes to produce the desired enzyme.Luciferases can also be fused to other proteins or fragmented to monitor proteininteractions of interest [276,290].Improving the BL properties of these systems is an active field, and research

ranges from generating a wider variety of BL colors, spanning the visible to near-infrared wavelengths, to improving the emission kinetics of the BL, by increasingthe intensity and/or decay of half-lives (reviewed in Ref. [278]). Research in thisarea is two pronged, focusing on the luciferases themselves and the luciferinsubstrates, with a wide range of protein mutants and substrate analogues nowproduced/utilized (reviewed in Ref. [278]). Sun et al. recently reviewed progress inD-luciferin amino analogues that produced emissions ranging from 460 to 609 nmwith wild-type luciferase [291]. Other D-luciferin analogues, reviewed by theMeroni group, give insight into a variety of chemical manipulations that mayresult in altered emitted light [292]. Caged luciferin, for example, is an alternativefirefly luciferase substrate designed for intracellular delivery available fromMolecular Probes [293]. Caged luciferin has also recently been synthesized forthe real-time in vivo imaging of H2O2 production in living mice [294]. Coelenter-azine substrates have similarly been altered to provide both enhanced bright-ness and enhanced duration of photon emission, including ViviRenTM andEnduRenTM from Promega, and several analogues with different emissionsare available from Molecular Probes and Biotium [295,296]. In an effort tocontinuously improve BRET, recent studies by Zhang and coworkers have shownimprovements in both sensitivity and limit of detection by 10-fold and 7-fold,respectively, through the use of an enhanced buffer environment [297].


As illustrated in Table 6.3, a variety of luciferase types can be combined withessentially three different chemical substrates to produce BL for BRET applica-tions. Bacterial luciferases, for example, catalyze the oxidation of reduced flavinmononucleotide (FMNH2) and a long-chain aliphatic aldehyde in the presence ofO2, yielding blue light [298,299]. Bacterial luciferase, however, is not optimized formost mammalian cell lines and the substrate, FMNH2, is rapidly oxidized in air,generating short bursts of light and not a steady BL emission [300]. Fireflyluciferases differ from bacterial luciferases in that they require an additionalcofactor, ATP, in order to catalyze the oxidation of the substrate luciferin, leadingto the emission of green-yellow light. Initial BL in this case is intense and thendecays to a low-sustained luminescence, which can be aided by an additionalcofactor, coenzyme A, to yield more stable, high-intensity luminescence [301].Rluc is one of the most commonly utilized luciferase systems and uses thesubstrate coelenterazine in the presence of O2. It has been widely adapted tomammalian cell lines, and hRluc is available from a variety of sources, includingPromega and PerkinElmer Life Sciences; cofactors are not needed. Improvementshave been made upon the wild-type enzyme, producing the often used mutants,namely Rluc2 and Rluc8, which have improved BL properties [287,302,303].Another coelenterazine-based luciferase that has begun finding wide applicationis derived from Gaussia princeps (Gluc or hGluc) and marine copepods (BLcrustacean), and has been optimized for expression in both bacterial and mam-malian cells [304,305]. The low molecular mass of Gluc (20 kDa) compared to Rluc(36 kDa) addresses problems associated with steric constraints in chimeric proteinfusions. Gaussia luciferase expressed in mammalian cells reportedly generateslight up to 1000-fold brighter than that of native Renilla [306,307]. BL photo-proteins from jellyfish and hydroid species, namely aequorin (from A. victoria)and obelin (from Obelia longissima), respectively, are also coelenterazine-basedenzymes that differ from luciferases in that the enzyme is complexed to itscoelenterazine substrate and is Ca2þ sensitive [308]. The principal application ofthese photoproteins has been as Ca2þ reporters [309,310]. In BRET applicationsaequorin has been employed to monitor the protein–protein interactions betweenSA and a biotin carboxyl carrier protein [311].In terms of BRET these BL enzymes are most commonly coupled with FPs, a

combination that has been fueled by the desire to push the BRET emission furtherinto the near-IR for optimal in vivo imaging. This has been facilitated by theincreasing number of mutant BL enzymes, substrate analogues, and mutant FPsthat allow good spectral overlap of the generated BL with the FP absorption[155,218,278]. As a result there are a variety of BL enzyme–FP BRETconfigurations,termed BRET x [288,302,312,313]. In BRET1 the BL enzyme variant is Rluc or Rluc8and the accepting protein is a GFP variant, YFP. Oxidation of the substratecoelenterazine-h by Rluc results in BL with a 480 nm peak, which, through energytransfer to YFP, generates a fluorescent emission peak at 530 nm. Typical uses ofBRET1 are ligand screening applied to real-time detection of protein–protein andprotein–ligand interactions, such as agonist-induced interactions of the GPCRsfamily of receptors [287,288,314]. BRET has been used to study a number of the


GPCRs, irrespective of their G protein-coupling selectivity [287,288], and is particu-larly amenable to high-throughput formats [287]. BRET2 utilizes the BL enzymevariants Rluc, Rluc2, or Rluc8 and acceptor GFP2 (a GFP variant with Ex 400 nm andEm 511nm). Reaction of the coelenterazine analogue substrate, DeepBlueC (coe-lenterazine 400A, sold commercially by Biotium, Inc., PerkinElmer, and NanoLightTechnology), with the Rluc enzyme causes a BL emission at �400 nm, resulting inexcitation of GFP2 and an endpoint emission at 511 nm [287,288,314]. Thisconfiguration works like a standard BRET assay, but has a larger apparent Stokesshift resulting in more spectral resolution between the donor–acceptor pair. Thistechnique was successfully applied to visualize protein–protein interactions in mice[312] and RNA detection and quantification (Figure 6.18a) [315]. BRET3 again usesRluc, Rluc2, or Rluc8 and the fluorescent protein mOrange with a coelenterazinesubstrate [313]. Advantages of the BRET3 combination include several-fold improve-ment in light intensity, as well as improved spatial and temporal resolution formeasuring intracellular events in a single cell. This improved BRET strategy allowsthe visualization of protein–protein interactions within small living animals[287,288,313,314]. The Gambhir group has subsequently demonstrated additionalBRETx systems with various combinations of Rluc variants (Rluc8 and Rluc8.6)combined with two red FPs, TagRFP, and TurboFP635, using the substratescoelenterazine and its analogue coelenterazine-v to generate a red light-emitting600–650 nm reporter system for in vivo protein–protein association studies [302].BRETx systems are initially characterized/optimized by generating a fusion

protein of the BL enzyme and the FP, and these fusion proteins representinteresting tags in their own right and may be useful labels for a variety of bioassays[316]. For example, they have been used in sequential BRET–FRET assays, termedSRET, for detection of heteromerization in plasma membranes and ratiometricprotease assays [286,314,317,318]. Extended BRET (eBRET) typically uses theenzyme variant Rluc or Rluc8 with YFP (Ex 480 nm and Em 530 nm). Apresubstrate,a protected form of coelenterazine (EnduRen from Promega), is metabolized byendogenous esterases into coelenterazine-h similar to that used in BRET1. Thisprovides a steady supply of substrate for luciferase oxidation within cells forextended periods of time up to 24 h [285,287,314].Although less common, BL enzymes have also been coupled with traditional

organic dyes and increasingly NMs, especially QDs as acceptors [283,286,319–321].Currently QD–BRET has been used for sensing protease activity, protein–proteininteractions and in vivo imaging, as reviewed in Ref. [283]. Enhanced BRET betweenthe firefly Photinus pyralis luciferase variant PpyGRTS quantum rods (QRs) as theenergy acceptor has also been described recently, with BRET ratios dependent uponenzyme loading, rod aspect ratio, and donor–acceptor distances (Figure 6.18b) [320].Protease activity sensing in a QD–BRETsystem relies on a peptide substrate linkagebetween the luciferase and the QD that is cleaved in the presence of the protease ofinterest, reducing the BRET-based QD emissions [321]. QD–BRET has also beenused to study protein–protein and receptor–ligand interactions. For example, Raoand coworkers fused luciferase (Luc8) to HaloTag protein and functionalized QDswith HaloTag ligands to study the resulting BRET that occurred when the HaloTag


Figure 6.18 (a) General strategy for BRET-based RNA detection. Probe1 consists of a 20-mer oligonucleotide conjugated at the 50 end tothe thermostable bioluminescent protein (RL8).Probe2 consists of a 20-mer oligonucleotideconjugated at the 30 end to the fluorescentprotein GFP2. Both the probes arecomplementary to different portions of thesame mRNA target gene. Thus, the targetmRNA serves as a scaffold upon which theprobes can bind, bringing the proteins intoproximity to one another. When RL8 oxidizes itssubstrate, the energy produced is nonradiativelytransferred to GFP2, which then emits photonsat a characteristic wavelength as itschromophore returns to the ground state. Adual probe assay testing sensitivity with mixedpopulations of in vitro-transcribed cRNA astargets. RL8 and GFP2 were combined withvarious amounts of Fluc cRNA, while keeping

the level of total cRNA constant bysupplementing with nontarget cRNA.Statistically significant BRET signal was seen foras little as 1 mg Fluc cRNA. Inset: raw imageobtained in IVIS-200. Rows from top to bottommatch columns left to right of figure. Left image:shows GFP2 filter; Right image: shows RL8 filter.(Reprinted with permission from Ref. [315].Copyright 2008, American Chemical Society.)(b) BRET between QRs and firefly luciferaseenzymes. (i) BRET efficiency plots for PpyGRTSdonors and QR acceptors. The summary of theBR measured with respect to aspect ratio atL¼ 5 and 10 for (ii) CdSe/CdS and (iii) CdSe/CdS/ZnS QRs. (iv) Illustration of themicrostructure of the particular QRs studied,including dot-in-dot, rod-in-rod, and dot-in-rodtypes. (Reprinted with permission fromRef. [320]. Copyright 2012, American ChemicalSociety.)


protein irreversibly bound the HaloTag ligands [286]. Interestingly, in vivo imagingwith a QD–BRET system has the potential to overcome the natural tissue scatteringand autofluorescence, which affects a large number of BRET acceptors with shortwavelength emissions. QDs are available with emission in the red and near-IRranges, which coupled with their broad absorption profiles makes them an excellentacceptor for BRET. This was first shown in a mouse model where superficial anddeep tissues were imaged using QDs with a 655 nm emission, which were coupledto Luc8 [319]. As highlighted later in the QD section (Section 6.5.5), the relativelynarrow emission profiles open up the possibility of multiplexed sensing of protein–protein interactions using a number of QD–BRET pairs [286,321].The BRET platform has been used for sensing a variety of analytes, including

Ca2þ, through the use of the described photoproteins and ATP. BRET is possiblymost often used for monitoring protein–protein interactions such as the earlierdescribed GPCR activation as well as the QD–BRET system for determiningreaction kinetics of HaloTag protein binding to HaloTag ligands [286]. Similarly,BRET has been utilized for nucleic acid hybridization assays and immunoassays,with the latter available for purchase from a variety of companies. As an exampleof a nucleic acid hybridization assay, Kumar et al. utilized Rluc bound to anoligonucleotide probe and a QD on the nucleic acid target [322]. Hybridization ofthe probe to target strands increased resonance energy transfer and QD emission.Cell-based BRET assays as well as in vivo BRET imaging are also becoming morepopular as more BRET pairs become available for the selected applications asdescribed, although care should be taken if quantitative BRET data is desired[285,287,314].

6.4.7Enzyme-Generated Chemiluminescence

Conceptually, there is little difference between the mechanism of BL (Section6.4.6) and enzyme-generated chemiluminescence (CL), other than in CL theluminophore is a synthetic substrate [278]. CL substrates include luminol and itsderivatives, 1,2-dioxetanes and acridinium esters, which are brought to an excitedstate through an enzymatically catalyzed reaction [282]. In direct CL, enzymaticactivity upon the substrate leads to electromagnetic radiation and an electronicallyexcited intermediate, which luminesces. Indirect or sensitized CL occurs whenthe energy is instead donated to another molecule, which in turn luminesces[323]. Table 6.4 describes some common CL substrates, processing enzymes, andchemical reactions. Applications of CL include immunoassays, protein blotting,DNA probe assays [324], detection systems in separative and flow-assistedanalytical techniques [325], measurement of target analytes with biospecificprobes, measurements of substrates, cofactors, or quenchers, and in vitro andin vivo imaging [282].CL resonance energy transfer (CRET) is a concept and laboratory technique that is

widely underutilized in current research; however, enhanced substrates, altered


enzymes, and NMamplification are slowly widening the range of useful applications[282,323,326,327]. The most widely used enzymes for CRET systems includehorseradish peroxidase (HRP) and the HRPmimic hemin/G-quadruplex DNAzyme[327–329]. Although a variety of CL substrates exist for the HRP enzyme, luminoland its luminogenic derivatives remain the most popular (Table 6.4) [282,330,331].As illustrated in the table, HRP oxidizes luminol to a luminescent species in thepresence of hydrogen peroxide, yielding blue emission at 425 nm. Luminol isusually employed in conjunction with an enhancer such as luciferin, fluorescein,or a phenolic compound (e.g., para-iodophenol) [332–334], which is thought toincrease the sensitivity of the assay through intermolecular energy transfer. HRPenzymatic conversion of acridan substrates generates higher luminescent intensitythan luminol, when the luminescent acridinium esters intermediates decay, emit-ting yellow light (530 nm) [335–337]. Alkaline phosphatase is another enzyme thathas shown promise in CRET applications. This enzyme catalyzes the oxidation of1,2-dioxetane luminogenic substrates as shown in Table 6.4 [338,339].CRET has shown promise in microchip electrophoresis [340], measurement of

target molecules and proteins such as ATP [341], human immunoglobulin G (IgG)[342], alpha fetoprotein (a cancer marker) [343], microRNA [344], DNA, metal ions,and aptamers [328], with biospecific probes that are often single-stranded DNA.Analogous to BRET, there is no outside excitation, though generally the QY in CL islower than BL. Much work has been done using NMs, including QDs [326], Au NPs[343], and graphene to enhance the QY, overall CRET brightness, and usability. Agreat review of many of these techniques can be found in Ref. [327]. In addition toNPs, magnetic beads have also been used to create a sensing platform that aids in aseparation protocol when working with complex protein samples such as serum[342]. Recently, work by the Willner group has focused on using CRET to generatephotocurrents. Through the use of a QD acceptor associated with electrical leads,CRET occurring through the enzymatic action of hemin/G-quadruplex HRP onluminol with triethanolamine as an electron donor, results in a detectable photo-current [345].CL and CRET research continues to evolve at a relatively slow pace compared to

FRETand BRET. Recent advances with the use of QDs and other NMs, coupled withthe large number of recombinant enzymes available, the low-cost commercialsubstrates, and control over emission wavelength, will drive further explorationof CRET for sensors and other applications.

6.5Inorganic Materials

Inorganic materials typically take the form of chelates, doped nano- or micro-particles or NMs. They have a range of unique properties including bright lumi-nescence, strong quenching abilities, large Stokes shifts, and long luminescentlifetimes that make them highly desirable for energy transfer applications asdiscussed in more detail later.

6.5 Inorganic Materials j211

6.5.1Luminescent Lanthanide Complexes and Doped Nano-/Microparticles

Luminescent lanthanides are a prominent class of long-lifetime fluorophores usedby the energy transfer research community for a wide range of bioapplications,mainly focused on in vitro and in vivo sensing and imaging [9,346–349]. Themajorityof the trivalent lanthanide ions are luminescent, with terbium (Tb), Eu, samarium(Sm), thulium (Tm), and dysprosium (Dy) emitting in the visible spectrum, whileytterbium (Yb), neodymium (Nd), and erbium (Er) emitting in the near-IR. Theiremission spectra are comprised of several well-separated narrow lines coupled withlong excited-state lifetimes on the order of several milliseconds. Long lifetime dyes(fluorescent lifetime t> 100 ns–ms) have a number of technical advantages overconventional fluorescence dyes (t¼ 1–5 ns). The principal benefit arises from theability to gate out (through time-resolved measurements) background fluorescencefrom direct excitation of acceptor dyes, scattering, and autofluorescence from cellsand biomolecules, which can dramatically improve sensitivity. Use of time-basedmeasurements may also necessitate more complex equipment than steady-statefluorimeters. However, because these are long-lifetime dyes (microsecond–milli-second) many standardmicrotiter well plate readers are available withmeasurementcapabilities in this timescale.For bioapplications, lanthanide ions are either complexed within an organic

chelate/cryptate ligand producing classical coordination metal complexes [lumines-cent lanthanide complexes (LLCs)] or doped into ceramic-type materials andformulated as nano-/microparticles. These complexes help to improve the opticalproperties of the lanthanides as well as their photostability and chemical stability,discussed in more detail later [9,347–349].The LLC chelate ligands vary in form, but include derivatives of polyamino-

carboxylates, cyclen, hydroxyquinoline, salicylamide, and phenylporphyrin, whichhave been recently reviewed [348]. The chelate ligands fulfill a number offunctional roles in the development of successful LLC bioprobes, with ongoingresearch to further improve/match their properties to target applications[3,347–351]. First, the lanthanide ion must be tightly bound within these chelatecomplexes, resulting in higher thermodynamic and photochemical stability,which shields the lanthanide ion from the quenching effects of the surroundingsolution. Second, compared to common dyes, lanthanide ions have very lowextinction coefficients (�1M�1 cm�1), making them difficult to excite directly.Thus, the chelate label contains an organic chromophore, referred to as the light-harvesting antenna molecule or sensitizer, which is placed in close proximity tothe ion. The sensitizing molecule absorbs incident light and due to closeproximity transfers this energy to the lanthanide ion, presumably by a Dextermechanism. Finally, the chelate label should possess a reactive group allowingbioconjugation. Commercial sources of lanthanide probes include CIS-Bio Inter-national (cryptate-based probes), PerkinElmer (LANCE1), Life Technologies(LanthaScreenTM), GE Healthcare (europium–TMT chelates), Lumiphore, andSigma-Aldrich.


Other than their long luminescent lifetime, LLCs have a number of otherproperties that make them excellent donors in luminescent resonance energytransfer (LRET) studies. For example, LLCs generally have a large effective Stokesshift, due to the separation of the absorption (by the organic chelate) and theemission spectra of the complex (from the lanthanide ion following energy transferfrom the organic chelate), which reduces the potential for direct excitation of theacceptor. In addition, their emission spectrum manifest in the form of multiple,distinct, and sharp emission bands, allowing LLCs to be coupled tomultiple acceptordyes [352,353]. Tb, for example, has good spectral overlap with fluorescein, rho-damine, Cy3, and a number of AlexaFluor dye acceptors (Figure 6.19a)[346,352,353]. LLCs have also been found to undergo a phenomenon termednonoverlapping FRET (nFRET), a mechanism not fully understood, which occursbetween a LLC donor and a spectrally nonoverlapping acceptor [354,355]. Using aDNA hybridization assay, Vuojola et al. investigated nFRET between Eu(III) chelateand various AlexaFluor dyes (with varying degrees of spectral overlap with the Eudonor). They found nFRET to be very efficient over short distances, more efficientthan predicted using conventional FRET theory, and unlike FRET, nFRETwas foundto be temperature dependent, leading the authors to conclude that a thermalexcitation process was involved as part of the nFRET mechanism [354].LRET using LLCs has been applied in a number of applications, including

monitoring protein–protein interactions in cells [356], monitoring orthogonal lig-and-dependent protein–peptide binding events [352], high-throughput screening ofpotential drug candidates [357], and numerous in vitro bioassays [347,353,358–361].Kupstat et al., for example, developed a homogeneous time-resolved immunoassay forprostate-specific antigen (PSA) that was sensitive and quantitative, and could beincorporated into a point-of-care testing (POCT) device [360]. A sandwich immuno-assay formatwas used inwhich the two antibody species that recognized and bound todifferent epitopes on the PSA were labeled with either the donor (Eu trisbipyr-idine) or the acceptor (APC protein). The presence of PSA brought the twoantibodies and hence the donor/acceptor species into close proximity, resulting inLRET, with LODs two orders of magnitude below the clinical PSA cutoff of 4 ng/ml. A similar format, using a Tb donor and five different acceptor dyes, wasrecently used to detect five different lung cancer tumor markers simultaneously ina 50 ml human serum sample [362]. Li et al. developed an adenosine sensor usingan aptamer-based sensor design, which functioned by inducing a conformationalchange that disrupted the LRET (Figure 6.19b) [361]. The sensor was able to detectselectively 60 mM of adenosine in undiluted serum samples.Themarriage of LLC donors and QD acceptors is a powerful combination in LRET

studies and takes full advantage of the many unique properties each brings to thetable, such as bright fluorescence (QDs), large Stokes shifts (both the QDs andthe LLCs), and time-gated measurements (LLCs) [363]. This donor/acceptor combi-nation has found application in luminescent microscopy, time-resolved immuno-assays, measuring protease activity, and detecting nucleic acid hybridization[364–367]. Algar and coworkers in particular have developed a series of time-gatedFRETrelays, demonstrating the use of QDs as simultaneous acceptors and donors in



bioassays for monitoring protease activity and nucleic acid hybridization [365,366].In the case of the protease activity bioassays, the authors were able to demonstratemultiplex protease activity detection using a single QD color (Figure 6.20) [368].Here, the QD (which acts as both a donor and an acceptor) was functionalized withpeptide substrates for trypsin (labeled with a luminescent Tb complex donor) andchymotrypsin (labeled with an AlexaFluor dye acceptor). Activity of chymotrypsinresulted in a decrease in prompt FRET (between the QD and the AlexaFluor dye),while trypsin activity resulted in loss of the time-gated FRET (between the Tbcomplex and the QD).LLCs have also been incorporated into thin-film layers or NPs, either within the

core–shell structure or bound to ligands on the NP surface, for a range ofapplications, including LRET-based bioassays, molecular imaging, and multiplexsignal labels [369–373]. Song et al. developed core–shell nanostructures aimed atimproving the LRET efficiency of the NMs, which comprised a rhodamine-functionalized silicon dioxide (SiO2) core surrounded by a Tb chelate-modifiedSiO2 shell [372]. The resulting core–shell nanostructures had an energy transferefficiency of 80%, a large F€orster distance range of 5.7–11.3 nm, and an emissionlifetime of �0.25ms.Besides LLCs, lanthanide ions are also doped into host ceramic materials, such as

oxides and fluorides, to generate phosphor/luminescent materials with uniqueupconverting properties [upconverting phosphors (UCPs)]. Upconversion is anonlinear phenomenon where a material sequentially absorbs long-wavelengthphotons and subsequently emits shorter wavelength emission, that is, convertsred to visible light, a different mechanism to multiphoton absorption, where thephotons are absorbed simultaneously [348,374]. The UCPs are routinely formulatedas NPs (UCNPs), where they have all the benefits of the LLC materials (i.e., time-resolved measurements, sharp emission profiles, etc.), but in addition UC of theexcitation light makes them excellent biolabels for in vivo imaging and in vitrobioassays. UCNPs allow the use of cheaper red excitation sources, avoid backgroundautofluorescence from complex biological samples (improving sensitivity), and usenear-IR excitation (typically �980 nm) allowing greater tissue penetration depths,while minimizing photodamage to biological samples [375–379].The most common crystalline host material used for generating UCNPs is the

fluoride NaYF4 that is either doped with one type of lanthanide species or, as is morecommon, codoped with two lanthanide species [374,377]. Codoping improvesthe UC efficiency with Yb3þ and Er3þ, representing a popular combination.

Figure 6.19 LLC materials for FRET. (a)Excitation (Ex) and emission (Em) spectra ofTb3þ chelate (black) versus fluorescein (green),and Alexa633 (red). (Reprinted with permissionfrom Ref. [352]. Copyright 2007, AmericanChemical Society.) (b) Scheme of the adenosinesensor design based on a Tb complexconjugated to a DNA aptamer. (c) Steady-stateemission spectra of the aptamer sensor upon

the addition of increased concentrations ofadenosine in the HEPES buffer solution(lex¼ 344 nm). (d) Emission intensity of thesensor at 545 nm as a function of adenosineconcentration. Inset: Shows the selectivity of thesensor toward adenosine over othernucleosides at 5mM concentration. (Reprintedwith permission from Ref. [361]. Copyright2012, American Chemical Society.)

3


Here, the Yb3þ dopant acts as the near-IR absorbing ion (sensitizer), while the Er3þ

acts as the emitter/activator ion [374]. UCNPs can be prepared using a number ofsynthetic procedures, including precipitation/coprecipitation, hydrothermal/solvo-thermal thermolysis-based techniques, and laser annealing; for more detail refer torecent publications [377–384]. Ultimately, the goal is to prepare highly crystallinestructures (which improves the overall UC efficiency) that have a small particle sizecombined with low size distribution, uniform dissemination of the doped lantha-nide ions, good aqueous solubility, and the ability to bioconjugate, if required

Figure 6.20 LLC–QD combinations. (a)Principle of the time-gated Tb!QD!A647FRET relay. Optical excitation of theconjugates yields excited-state Tb and anexcited-state QD (�), and FRET2 is observedon a prompt timescale (emission <100 nsand integration time 20 ms). The extent ofFRET2 is measured via the ratio of promptA647 and QD PL, rp. Following a suitabletime delay (60 ms) during which the QDreturns to its ground state, FRET1 andsubsequent FRET2 can be observed. Theextent of FRET1 is measured in a time-gatedobservation window via the ratio of gatedQD and Tb PL, rg. (b) Schematic of a time-

gated FRET relay for multiplexed proteasesensing. A central CdSe/ZnS QD is coatedwith compact zwitterionic ligands (CL4) andassembled with polyhistidine (His6)-appended peptide substrates. The peptides,labeled with either Tb or A647, serve assubstrates, SubTRP and SubChT, for TRPand ChT, respectively. The cleavage sites arehighlighted in the peptide sequences.Proteolytic activity disengages FRET andalters the prompt and gated PL ratios, qpand qg, which are used as analyticalsignals. (Reprinted with permission from Ref.[365]. Copyright 2012, AmericanChemical Society.)


[374,377,380,385,386]. Common techniques to improve the aqueous solubility ofthese inherently hydrophobic materials include polymer surface coatings (such assilica shells), layer-by-layer techniques, ligand exchange, and surfactant addition(including phospholipids), reviewed in Refs [376,377,380].Lanthanide materials doped into oxides and fluorides, which take advantage of

time-resolved measurements (but not the UC potential), have been used in LRETstudies for a range of applications including labels in bioassays, imaging, and NP/bioconjugation characterization [387–390]. However, fully exploiting the UC prop-erties is more common and UCNPs have a wide range of applications, includingdisplay devices, solar cells, optoelectronics, lasers, catalysis, and biolabeling, such asbioassays, imaging, and therapy, which have been the topic of recent reviews[374,377,380,385,386,391,392]. In most cases the UCNPs donors are coupledwith the traditional organic fluorophore acceptors, although there have been reportsof UCNPs coupling with QD [393] or Au NP [394] acceptor NMs. LRET-based in vitrosensors using UCNPs have been successfully developed for small molecules, suchas ammonia [395] and glucose [394], nucleic acid hybridization [396], and proteindetection, such as caspase-3 [397] and avidin [392]. UCNPs combined with energytransfer assays have also been used in various imaging studies, for example, to lookat the intracellular fate of small interference RNA (siRNA) upon uptake into cells[398]. Cheng et al. developedmulticolor UCNPs for multiplex in vivomouse imagingby tuning their emissions via LRET [399]. NaYF4 NPs doped with Er3þ/Yb3þ (greenemission – donor) were functionalized with an amphiphilic polymer before beingloaded with the fluorophores rhodamine B, rhodamine 6 G, or the quencher TideQuencher 1 (acceptor) to produce three different colored UCNPs (Figure 6.21). Todemonstrate the imaging utility of these materials, five UCNP materials – the threedescribed plus unmodified NaYF4: Yb, Er (green) and NaYF4: Yb, Tm (red) – wereinjected subcutaneously into the back of nude mice, excited using a 980 nm laserand imaged using a MaestroTM imaging system (PerkinElmer), which capturedmultispectral fluorescence images (Figure 6.21). Spectral unmixing by the Maestroimaging system clearly distinguished where each population of the UCNPs waslocated in the mouse model, demonstrating the multiplex capability of the LRET-color tuned UCNPs, when combined with the spectral imaging technology.

6.5.2Luminescent Transition Metal Complexes

Transition metals integrated into organic complexes, either as classical coordinationmetal complexes or as organometallic compounds (metal complexes containing atleast one metal–carbon bond), are found to have unique luminescent properties thattypically arise from a triplet metal–ligand charge transfer process and are reviewedin Refs [9,400–404]. These complexes possess a number of favorable characteristicsthat make them suitable for luminescent applications, including long excited-statelifetimes (100 ns–ms), high photostability, and often a large Stokes shift. Thesecomplexes have found particular application in cell imaging [9,400–404]. Of thetransition metals, ruthenium (Ru) complexes remain the most popular, however


increasingly iridium (Ir), rhenium (Re), and occasionally osmium (Os) and plati-num (Pt) have been used in cell imaging applications [9,400–405]. The current factorlimiting widespread adoption of these types of materials is probably commercialavailability, asmost of thematerials are synthesized in-house. Sigma-Aldrich offers aseries of reactive Ru complexes, originally developed by Lakowicz as anisotropylabels [406,407]. These Ru complexes have lifetimes of t� 500 ns, small extinctioncoefficients (14 500M�1 cm�1), relatively low QYs (0.05), high photostability, fairlylarge Stokes shift, and absorption close to the visible spectrum, but, most impor-tantly, they are functionalized with moieties that facilitate bioconjugation.While cell imaging dominates the biological applications of these luminescent

transition metal complexes, there have been some examples of their use in LRET-based studies. Ru complexes have been used as donors in immunoassays for humanserum albumin (HSA) [407,408] and for CO2 when coupled with the environ-mentally sensitive Sudan III disazo acceptor dye [409]. There are also examples of Ru

Figure 6.21 Multicolor imaging of UCNPs.(a) Multicolor UCL images of three UCNP–dye complexes and a mixture of the three.The images were obtained by the Maestro invivo imaging system after spectral unmixing.(b) UCL emission spectra of solutions ofUCNP1, UCNP2, UCNP1/RhB, UCNP1/R6 G,and UCNP1/TQ1 recorded by the Maestro invivo imaging system under the 980 nm NIRlaser excitation. Inset: Fluorescence spectra

of RhB and R6 G under green lightexcitation. (c) Multicolor in vivo UCL imagingof LRET-tuned UCNPs in mice. Left image: Invivo multicolor UCL images of a nude mousesubcutaneously injected with five colors ofUCNPs solutions after spectral unmixing.Right image: A white light image of theimaged mouse. (Reprinted with permissionfrom Ref. [399]. Copyright 2011, AmericanChemical Society.)


complex acceptors, inmetal complex – protein binding studies and thrombin activityassays [410,411]. More recently, Ru complexes have played the role of both acceptorsand donors in a three-color FRETrelay for studying DNA and polypeptide dynamicsand interactions [412,413]. Os(II) complexes have been coupled as acceptors inFRET studies to QD donors [414,415], and Ir(III) FRETprobes have been developedfor cysteine and homocysteine detection [416]. A Zn porphyrin cofactor in Zn(II)-substituted horse heart cytochrome c was shown to serve as a donor to a Alex660acceptor in cytochrome c unfolding studies [417]. CPEs (see Section 6.3.5), modifiedwith transition metal complexes, have also been developed for protein sensingapplications [138,418]. The CPEs are composed of main donor segments andtransition metal complex-modified acceptor units, which in aqueous solutionundergo polymer aggregation, resulting in efficient FRET. Addition of certainproteins disrupts the polymer aggregate structure, resulting in a measurabledecrease in FRET efficiency. Demonstrations include HSA detection using Pt(II)-modified CPEs [418] and histone detection using Ir(III)-modified CPEs [138].

6.5.3Noble Metal Nanomaterials (Gold, Silver, and Copper)

Au, Ag, and other noble metal, such as Cu, NMs exhibit unique size- and shape-dependent optical properties, due to surface plasmon resonances in the visible range(see Figure 6.22 for Au example) [419,420]. These particles typically have largerextinction coefficients (�105 cm�1M�1), more stable/nonfluctuating signal inten-sities, and greater resistance to photobleaching when compared to small-moleculefluorophores [421]. Due to their strong absorbance, they are often used as quenchers

Figure 6.22 Au NPs. (a) Normalized UV–Visabsorption spectra of Au NPs with differentsizes. (b) Photographs of the colloidal Au NPswith different diameters (2.4–89 nm).Concentrations of Au NPs are 670 nM (2.4 nm),56 nM (5.5 nm), 17 nM (8.2 nm), 2.3 nM(16 nm), 0.17 nM (38 nm), and 0.013 nM

(89 nm). (c) TEM images of Au NPs. Averagesize and standard deviation are reported foreach sample. Scale bars are 20 nm (top) and40 nm (bottom). (Reprinted with permissionfrom Ref. [419]. Copyright 2011, AmericanChemical Society.)


(acceptors) in FRET applications; however, highly luminescent Au and Ag QDs ornanoclusters have been synthesized, leading to their potential role as fluorescentFRET donors/acceptors [422–424]. Excellent reviews on the properties of Au[425,426] and Ag [427–429] NMs commonly used in optical applications can befound in the literature.Au NPs can be produced in various sizes using either the citrate reduction (16–

147 nm diameter) or the Brust–Schiffrin method (1.5–5.2 nm diameter) [425,430].These NPs can also be produced in various shapes such as spheres, rods, shells,cages, and plates [431–433]. Similarly, Ag NPs are often produced in various sizesthrough the reduction of Ag salts, typically a stronger reducing agent leads to smallersized particles (<40 nm), which are often less stable over time than the larger NPs[434]. Biogenically produced nanoAg provides an environmentally friendly synthesisroute [435], and various oligonucleotide sequences have been used to template Agnanoclusters [422]. Commercial sources of Au and Ag NMs are available in a widevariety of geometries such as spheres, rods, and shells from relatively newermanufacturers such as nanoComposix, NANOCS, and NanoPartz. More specializedcompanies such as NanoRod, LLC or Microspheres-Nanospheres offer a widevariety of Au nanorods or metallic nanospheres, respectively. Commercially availa-ble copper NPs are less common but can be purchased in organic solution fromSkySpring Nanomaterials, Inc.One of the intrinsic benefits of using Au and Ag NPs is that they are readily

functionalized with ligands containing specific terminal chemical moieties (e.g.,carboxyl or amine) or biomolecules through reactions with exposed thiol groups thatdirectly attach to the NM surface via formation of an Au–sulfur (S) [436] or Ag��Sbond [437]. Companies such as Structure Probe Inc., Nanoprobes, EB Sciences, andResearch Diagnostics Inc., offer an extensive array of colloidal Au in many sizes,which are available functionalized with a variety of bioconjugates. British BiocellInternational offers a variety of colloidal Au and Ag also prefunctionalized aschemical or biological conjugates.Au is by far the most commonly used material in optical applications, and while

Ag materials are mostly utilized for their antimicrobial properties, the increased useof Ag NMs in optical applications stems from their high QYs, photostability, andstrong fluorescence intensities [427,438–443]. As a result of their native oxide layer,copper NMs have been observed to enhance fluorescence signal [444], however,recent syntheses that decrease the native oxide layer may increase the use of Cu NPsas fluorescence quenchers in future optical applications [445].In terms of energy transfer studies, the interaction of noble metal NMs and

fluorophores can be quite complex, resulting in either quenching or plasmonicenhancement of the proximal fluorophores fluorescent signal [446]. Plasmonicenhancement, observed in metal NPs coated with fluorescent dyes, is an energytransfer-type phenomenon between the excited-state fluorophore and the plasmonresonance of the proximal metal surface/particle [428,429,447–449]. Successfulplasmon enhancement requires careful spacing between the fluorophore and themetal structure and factors such as metal type, NP size, and fluorophore can allinfluence this complex process [450–456]. Plasmon enhancement has been


exploited to increase FRET efficiency between DNA bound fluorophores [453–456],and more viable configurations are expected in the near future.Au and Ag (to a lesser extent) NMs are more commonly used for their “super-

quencher” abilities, that is, the ability to quench fluorescence over a broad range ofwavelengths. The fluorescence quenching ability is dependent on a number offactors, including the NP size and shape, fluorophore distance from the NP surface,fluorophore dipole orientation, and the amount of overlap between fluorophoreemission and NP absorption spectra, all of which influence the radiative andnonradiative decay rates [432,445,457–459]. A number of detailed studies havecharacterized the fluorescence quenching, via dipole–metal interactions, of variousfluorophores attached near the surface of Au NPs, and both FRET and nanometalsurface energy transfer (NSET) mechanisms have been proposed for emissionquenching, which have a d�6 and d�4 distance dependency on efficiency, respec-tively [445,460–465]. The NSETmodel extends the efficient nonradiative quenchingdistance between the fluorophore and the proximal Au NP, effectively extending themolecular ruler capabilities, and this has been put to good use in a number of energytransfer studies [445,460–465].The quenching abilities of Au NMs in FRET/NSET configurations are com-

monly exploited in a number of bioassay formats. Molecular beacon-based assaysfor DNA sensing measurements, for example, produce 100-fold sensitivityenhancements using Au NMs compared to previous dye–dye combinations[466–469]. Au NP–fluorophore complexes show promise for in vitro diagnosticapplications as “noses,” with the ability to discern between various bacteriaspecies and strains, proteins in complex solutions, and between cancerous andhealthy cell lines [470]. These complexes have also been used for the detection ofmalaria antigens [471], DNA analysis [472,473], and for in vivo probes for reactiveoxygen species, hyaluronidase, and protease detection systems [458]. Aptamersmodified with fluorophores and subsequently combined with Au NPs have beenused for the multiplex detection of adenosine, Kþ ions and cocaine [474], andnanorulers for measuring binding-site distances on live cell surfaces [475]. Inaddition, Au NPs have been tested as quenchers for semiconductor QDs (seeSection 6.5.5). QD–Au NP systems have been used as probes to monitor real-timeintracellular gene expression [476], to detect DNA hybridization events [477–480],and for TNT detection [481]. Polystyrene microspheres surface modified with AuNPs and QDs have been proposed as suitable FRET-probes for bioassays,including measuring protease activity [482]. Results from these Au NP–QDquenching demonstrations suggest that this FRET configuration has tremendouspotential. Besides the lower background and improved sensitivity, the abilityto label both the Au NP and QD with multiple biological moieties mayimprove avidity.While Au and Ag NPs are typically used for their quenching abilities, other

applications utilizing their ability to scatter or fluoresce light are increasing. Highlyfluorescent Au and Ag QDs consisting of only a few clusters of noble metal atomshave been synthesized [423,424], and they show potential as labels in in vitro andin vivo imaging [483]. Larger clusters of a few nanometer thicknesses show promise


as low photobleaching alternatives in cancer cell imaging [484]. Much like theirsemiconductor counterparts, these Au and Ag QDs (or nanodots, NDs) have size-tunable emission maxima, shifting to longer wavelengths with increasing nano-cluster size, although emission can also be influenced by other factors such assurface ligands and crystallinity of the NM [424].Although not strictly energy transfer based, plasmonic rulers consisting of two

plasmonic particles in close proximity, capable of detecting molecular binding andfolding events, offer several advantages over traditional dye-based FRETapplicationsin that they prevent photobleaching as well as allow for a 10-fold increase inmeasurement range [485,486]. Plasmonic rulers may also be fabricated in threedimensions allowing a more spatially complete understanding of complex molecu-lar events [487].The wide variety of materials and applications incorporating the light-altering

properties of noble metal NPs suggests the increasing importance of these particlesin optical applications.

6.5.4Silicon-Based Materials

A number of Si-based materials, considered metalloid in nature, have been foundto have intrinsic fluorescent properties that are worth mentioning. Silole mole-cules and polymers, for example, are Si-containing five-membered cyclic dienesstructures that are found to become highly fluorescent upon aggregation [488].The silole molecule 1,1,2,3,4,5-hexaphenylsilole (HPS) generates a strong aggre-gation-induced luminescence at 495 nm. Amorphous silica (SiO2) NPs have alsobeen found to exhibit inherent luminescence due to oxygen-stabilized defects inthe SiO2 lattice [489], although it is more common to dope silica NM structures(core, core–shell, and shell structures) with organic fluorophores in the pursuit offluorescent NMs (see Section 6.3.4). Si NPs are a much more commonly utilizedluminescent form and are increasingly used as bioimaging agents due to their lowtoxicity, resistance to photobleaching as well as their bright size-dependentphotoluminescence and broad excitation spectra [490–495]. Si NPs have beeninvestigated as fluorescent tags for DNA [496], photonic barcode devices [497],potentially nontoxic materials for in vivo and in vitro imaging [498] includingbiodegradable imaging systems [499], theranostic systems in which particles canimage as well as potentially treat cancer cells photodynamically [500,501], andbiomodal imaging systems in which iron-doped Si particles exhibit magnetic aswell as fluorescent properties [502]. The synthesis of Si NPs remains tricky, butnew methods for synthesizing and stabilizing them have been reported [503–505],including those that provide a variety of particle geometries such as “flowerlike”polyhedron [506], nanowires, and clusters [507]. An extensive review of Si NPsynthesis and physical properties can be found in Refs [495,508]. Future appli-cations using Si-based NPs as FRET donors can be expected because of theincredible photostability, tunability, and facile surface modification of thesematerials.


6.5.5Semiconductor Nanocrystals

Colloidal luminescent semiconductor nanocrystals, or QDs, have a range of poten-tial applications, including optoelectronics (lighting and advanced displays), optics(lasers), solar energy, biotechnology, and medicine [509]. However, since theirinception as biological labels, applications in the biological arena have developedexponentially [283,510–515]. QDs can function either as a passive fluorescent labelor in a more integrated/active role, where the QD both acts as a scaffold forbiorecognition and is intimately involved in signal transduction, through mecha-nisms such as FRET, BRET, charge transfer (CT), or CRET [283,512,513,516–518].QDs possess a number of unique electro-optical properties that make them idealenergy transfer labels [283,512,513,515,518,519]. Benefits include a size and chem-ical composition dependent emission that is narrow and symmetric in profile, highquantum yields and extinction coefficients (enabling single-molecule detection),broad absorption profile, large Stokes shift, large two-photon absorption properties,and excellent resistance to photobleaching and chemical degradation[283,512,513,515,518–521]. QDs have been synthesized from a range of binaryand ternary alloys such as ZnS, CdSe, CdTe, InP, GaN, PbS, ZnO, InGaAs, andCdZnS, and their exact emission, which can span from UV-Vis to infrared, is foundto be dependent on both the chemical composition and NP size, resulting in atunable emission profile (Figure 6.23) [515,522,523]. For FRET, in particular, thismeans that QD donors can be size-tuned or “dialed in” to have better spectral overlapwith a particular acceptor dye, improving FRET efficiency. The broad absorptionspectra and large Stokes shift found in QDs is also of benefit for FRET studies, as itallows excitation of mixed QD donor populations at one wavelength far removedfrom their emissions and also facilitates selection of an excitation wavelength thatcorresponds to the acceptors absorption minima, thus reducing direct excitationbackground signals. The ability to excite multiple QD donors using a singleexcitation wavelength, combined with their narrow and symmetric emission (whichmakes deconvolution of multiple fluorescent signals simpler), makes them attract-ive labels for multiplex applications [32,321,513,524–529].As with any of the potential FRET labels discussed in this chapter, there are issues

to consider before using QDs. QDs have been found to blink under continuousexcitation, which may be problematic for single-molecule studies. QDs have a finitesize that can be both a benefit and a liability for FRET. In addition, the “assynthesized” semiconducting nanocrystals are inherently hydrophobic, requiringsome type of modification, that is, surface coating, to facilitate water solubility whilemaintaining their optical properties. Aqueous solubility can be achieved using threemain approaches: direct aqueous synthesis using hydrophilic stabilizing agents, capexchange of hydrophobic surfactants with hydrophilic ligands, and encapsulation/coating with amphiphilic species, polymers, or silica coatings, reviewed in Refs[512,513,515,519,530]. Encapsulation, in particular, can significantly influence thehydrodynamic radius of the final QD and hence impact the distance-dependentFRET efficiency [512,515,518,519]. A number of researchers have focused on


developing ligands that can facilitate both aqueous solubility and subsequentbioconjugation of CdSe/ZnS QDs, while keeping the overall hydrodynamic radiuscompact [512,513,515,519,531]. These ligands have evolved from simple designsthat facilitate aqueous solubility to multifunctional modular designs that comprisean anchor group that interacts with the QD surface (commonly a monodentate or

Figure 6.23 Dependence of fluorescenceemission wavelengths of QDs on theirchemical composition. The CdSe group isexpanded to demonstrate the size-dependent absorption and emission

profiles of the CdSe QDs, obtained by Pengsynthesis and different heating times: 3, 5,7, 10, 14, 20, 25, and 30min. (Reprintedwith permission from Ref. [515]. Copyright2013, Elsevier.)


bidentate thiol), a hydrophilic segment that imparts solubility (e.g., PEG, zwitter-ionic nature), and a terminal functional group that can provide solubility and be usedfor bioconjugation (e.g., ��OH, ��NH2, ��COOH, ��Biotin, and ��azide or��alkyne for click chemistry) [518,531,532]. Susumu et al., for example, designedcompact zwitterionic inspired ligands for preparing QDs and Au NPs [533]. Thezwitterionic nature of the ligands greatly improved the pH stability of the QDs,compared to dihydrolipoic acid (DHLA) only preparations, without impacting thehydrodynamic radius of the functionalized QDs (Figure 6.24) [533].In situations where FRETefficiency is found to be low, the finite QD size can be an

advantage allowing multiple acceptors to be assembled around the central QDscaffold, improving the FRET efficiency and potentially enhancing subsequentbiomolecular interactions (as recently demonstrated in proteolytic digestion studies[368]), but this may not be desirable for all applications [3,518]. In addition,dimensionality of the NM may be a factor influencing the FRET efficiency, asfound in the case of a QD donor (spheres versus rods) coupled to multiple acceptors[534–536].The broad absorption profile of QDs serves them well as FRETdonors, however it

can be problematic for their application as FRETacceptors, where direct excitation ofthe QD by the excitation source is an issue [517]. With careful choice of the donorspecies, there are a number of instances where QDs are excellent energy transferacceptors, including BRETand CRET assays (Sections 6.4.6 and 6.4.7, respectively),which do not require an excitation source, or the use of long-lifetime donors,therefore allowing the performance of time-gated/resolved measurements, whichcan factor out any direct QD excitation [517,524].There are a number of commercial suppliers of QD materials, recently reviewed

in Ref. [515], that are either functionalized with chemical handles that facilitatebioconjugation (e.g., ��OH, ��NH2, ��COOH, and -biotin) or are bioconjugatedwith SA or various secondary antibodies (e.g., goat antihuman, goat antimouse,or goat antirabbit). While there are increasing varieties of commercially availableQD materials, the most popular ones still remain the CdSe and CdTe corematerials and the CdSe/ZnS core–shell QDs. There are reviews and detailedmonographs describing QDs synthesis using various materials, generally involvingwet chemistry techniques such as high-temperature organometallic synthesis,microwave or gamma irradiation, and aqueous colloidal and sol–gel methods[515,523,537,538]. Various biotemplated fabrication approaches have also beenproposed for QD production, ranging from whole organism synthesis (bacteria,yeast, and viruses) to biomolecule-based ligands that facilitate nucleation andcapping of the QDs during synthesis (nucleic acids and peptide sequences)[539–543]. While greener in terms of reagents and reaction conditions, thesebiofabrication methods tend to produce lower quality QDs, in terms of QY andsize polydispersity, compared to high-temperature chemical synthetic techniques,but this may improve as our understanding of the underlying mechanisms thatgovern these syntheses grows [539–543].As discussed in Section 6.2, there are a number of diverse strategies that exist for

attaching biomolecules to QDs, including covalent coupling, electrostatic/metal


Figure 6.24 Improving aqueous solubility ofQDs through DHLA surface ligands. (a)Chemical structures of DHLA and the DHLA-based ligands used to stabilize QDs. (b–d)Physical characterization of a series of compactligand (CL)-coated QDs. (b) Gel electrophoreticseparation of 550 nm emitting QDs cap-exchanged with the indicated ligands. Gels wererun on 1% agarose gel in 1 TBE buffer (pH 8.3)at �7 V/cm for�10min. (c) Hydrodynamic sizedistribution of 550 nm emitting QDs cap-exchanged with DHLA: 10.8 (2.7 nm), CL1: 8.6(1.8 nm), CL2: 9.3 (1.7 nm), CL3: 9.5 (2.1 nm),CL4: 9.8 (2.2 nm), and DHLA-PEG750-OCH3:11.5 (2.5 nm) measured by dynamic light

scattering. Data is plotted in arbitrary units ofscattering intensity. (d) PL images (left) for a setof 0.5mMQDs capped with the CL1 compactligands in different buffers at pH 2–13. The550 nm emitting CdSe/ZnS QDs were used andexcited with a UV lamp at 365 nm. Images weretaken <20min and �4 weeks after samplepreparation. PL images (right) for a set of0.5mM–550 nm emitting QDs coated withDHLA or the indicated compact ligands in 3MNaCl solution. Images were taken 1 day aftersample preparation for DHLA and after 90 daysfor CL1�CL4. (Reprinted with permission fromRef. [533]. Copyright 2011, American ChemicalSociety.)


affinity-driven self-assembly, and biotin–avidin chemistry (Figure 6.25) [283]. How-ever, given their NMnature, additional issues (also discussed in Section 6.2), such asthe influence of the surface ligand on bioconjugation reactions and subsequentbiomolecular interactions and how the attachment chemistry influences the NMstability and biomolecule orientation, should be considered for QDs [30,33–36,515].QDs as donors in FRET applications have been coupled to a range of

acceptor materials for energy transfer studies, including fluorescent dyes, NMs(such as Au and carbon), fluorescent proteins, and polymers (Figure 6.26a)[79,184,283,462,482,513,516,517,520,524,544–548]. As mentioned earlier, theyare also adept FRET acceptors when coupled with the appropriate donors, such

Figure 6.25 QD bioconjugation – anillustration of some selected surfacechemistries and conjugation strategies that areapplied to QDs. The gray periphery around theQD represents a general coating. This coatingcan be associated with the surface of the QD via(e) hydrophobic interactions, or ligandcoordination. Examples of the latter include (a)monodentate or bidentate thiols, (b) imidazole,polyimidazole (e.g., polyhistidine), ordithiocarbamate (not shown) groups. Theexterior of the coating mediates aqueoussolubility by the display of (c) amine or carboxyl

groups, or (d) functionalized PEG. Commonstrategies for bioconjugation include (a) thiolmodifications or (b) polyhistidine ormetallothionein (not shown) tags that penetratethe coating and interact with the surface of theQD, (f) electrostatic association with thecoating, (g) nickel-mediated assembly ofpolyhistidine to carboxyl coatings, (h)maleimide activation and coupling, (i) activeester formation and coupling, and (j) biotinlabeling and SA–QD conjugates. The figure isnot to scale. (Reprinted with permission fromRef. [283]. Copyright 2010, Elsevier.)



as long-lifetime lanthanide materials (Figure 6.26b) [513,517,549]. QD donors/acceptors have also been applied in a number of studies looking at multi-FRETprocesses, discussed in more detail in Section 6.6. Algar et al., in addition, recentlydemonstrated QDs as simultaneous acceptors and donors in time-gated FRETrelays bioassays for monitoring protease activity and nucleic acid hybridization(Figure 6.20) [366]. QDs have also been shown to be sensitive to CTprocesses andcoupling with redox active species, such as dopamine, Ru, rhodamine B, and Os,which often leads to a competition between energy and charge transfer processesin the resulting complex (Figure 6.26c) [415,513,550–553].Since our previous review [3], the use of QDs in energy transfer-based applications

has truly blossomed and has been reviewed in a number of excellent monographs[283,513,515–517,524,554,555]. In vitro applications dominate the QD FRET litera-ture, including assays for detection of specific targets ranging from small moleculesto proteins and whole organisms (e.g., bacteria), enzyme activity monitoring,tracking intracellular gene delivery, solid-phase assays, and QD-enabled single-molecule detection. There are hosts of biomolecular interactions/processes that arestudied using QD energy transfer formats and these include nucleic acid inter-actions, binding protein conformation changes, antibody binding, aptamer inter-actions, and protease cleavage [283,513,515–518,524,554]. Assay formats found inconjunction with QD FRETare quite varied and include (i) cleavage-based assays (e.g., proteases, kinases, and DNAzymes) (Figure 6.27) [321,365,368,526–529,556–558], (ii) conformational change-based assays (e.g., aptamers, binding proteins, andDNA molecular beacons) [518,559–561], (iii) displacement assays (e.g., antibodiesand binding proteins) (Figure 6.28) [554,562], (iv) various immunoassays (includingdirect, displacement, and sandwich) [367,562–564], (v) nucleic acid hybridization[518,555], and (vi) assays based on acceptor spectral changes (mainly pH or ionsensitive dyes) (Figure 6.16) [79,527,556]. Given the unique photophysical propertiesof QDs, our increasing fundamental understanding of these unique materials whenused in energy transfer configurations, and the availability of improved synthesisand bioconjugation methods, we can expect continued utilization in many FRET-based biological assays, with increasing emphasis on multiplexed and in vivodetection [321,524,526–529,549,565].

Figure 6.26 QDs as FRET acceptors anddonors. (a) QDs are good FRET donors forfluorescent proteins (FPs), dye, and Au NPacceptors. The dashed circle represents anarbitrary F€orster distance (R0) measured fromthe QD center. The scale on the right indicateshow R0 proportionally increases as the numberof proximal acceptors (a) increases. Conversely,QDs can function as acceptors for Tbcomplexes and BL luciferase donors. (b)Qualitative spectral overlap (shaded) for a625 nm emitting CdSe/ZnS QD as (i) donor tofluorescent dye acceptor (Alexa647, A647) and(ii) acceptor to a Tb chelate donor. (c) CT

quenching is an alternative method ofmodulating QD PL: (i) an electron acceptor (e.g., quinone) has an unoccupied energy levelintermediate in energy to the 1Sh and 1Se band-edge states to which the excited QD transfersan electron, and (ii) an electron donor (e.g., Ruphenanthroline) has an occupied intermediateenergy level and transfers an electron to theQD. Charge transfer inhibits radiativerecombination of the exciton. Both the redoxactive species are illustrated as peptidesconjugates. (Reprinted with permission fromRef. [513]. Copyright 2011, American ChemicalSociety.)

3


Figure 6.27 Cleavage-based FRET assays. (a) Aprotease assay where (i) an acceptor dye-labeled peptide is assembled on a QD donor viaa polyhistidine tag. The QD-dye proximity in thebioconjugate is sufficient for FRET. (ii) Proteaseactivity (scissors) cleaves the peptide and

disrupts FRET, restoring the QD PL. (Reprintedwith permission from Ref. [283]. Copyright2010, Elsevier.) (b) Proteolytic assay data fromexposing a constant concentration of 550 nmemitting QDs conjugated to four Texas Redsubstrate peptides to a constant concentration


6.6Multi-FRET Systems

Multistep FRET is a naturally occurring process exemplified by the light-harvestingsystems found in many biological species, which allows them to harness light notreadily absorbed by chlorophyll for photosynthesis [254,566]. Phycobilisomes aresupramolecular complexes found in blue-green cyanobacteria and various algaesuch as glaucophytes, red, and cryptomonad [253,254]. Phycobilisomes have avariable composition that is organism dependent, but typically consists of multiplePBP subunits (see Section 6.4.4). As mentioned in Section 6.4.4, within thephycobilisome system there are four main types of PBP – PC, PE, PEC, andAPC – that bind the phycobilin chromophores (e.g., PUB, PEB, and PCB) thatgive these proteins their intense colors [254]. The multi-FRET process and flow ofenergy in phycobilisomes is PE–PC–APC–photosynthetic reaction center (chloro-phyll) [254]. Energy transfer efficiency in this system approaches nearly 100%, andresearchers have yet to match experimentally the complexity or efficiency of thisnaturally selected energy harvesting system.An increasing number of biologically inspired and artificial synthetic multi-FRET

systems have been developed to precisely space or orient multiple fluorophores withthe goal of characterizing and mimicking the natural light-harvesting process. Suchsystems have potential use in solar cells, nanoscale photonic devices, and otheroptoelectronic applications [269,567,568]. More commonly, these multi-FRET con-figurations are developed as a means to extend the optical ruler and are used toelucidate biological configurations, study protein and DNA interactions, and forbiosensing applications [135,210,365,366,412,413,569].While artificial synthetic building block structures have been developed, such as

perylene bisimide-calix[4]arene arrays by Hippius et al., to control the position andorientation of chromophores, biologically inspired platforms are more common[570]. DNA is perhaps the most attractive biologically inspired platform for multi-FRET configurations due to (i) its predictable structure/chemistry, (ii) the inherentability to introduce fluorophores at specific sites, (iii) the ability to hybridizemultipledye-labeled oligos to a complimentary strand, and (iv) the ability to control theorientation of the attached fluorophores [3]. DNA can be synthesized with multiplefluorophores at specific terminal or internal sites or with thiol/amine/biotin or other

of caspase-3 enzyme. Derived Km and Vmax

values are given. An R2 of 0.98 was obtained forthe fitting of the curve. (Reprinted withpermission from Ref. [556]. Copyright 2010,American Chemical Society.) (c) Multiplexedassay of proteases by using QDs with differentcolors on a glass slide. SA-QD525, SA-QD605,and SA-QD655 were used (from left to right).Biotinylated peptide substrates for MMP-7,caspase-3, and thrombin were conjugated tothe AuNPs, and then the resulting Pep-AuNPs

were associated with SA-QD525, SA-QD605,and SA-QD655, respectively: (i) SA-QDs only,(ii) SA-QDsþ respective Pep-AuNPs, (iii) SA-QDsþ Pep-AuNPsþMMP-7, (iv) SA-QDsþ Pep-AuNPsþ caspase-3, (v) SA-QDsþ Pep-AuNPsþ thrombin, and (vi)QDsþ Pep-AuNPsþmixture of the respectiveprotease and its inhibitor. (Reprinted withpermission from Ref. [528]. Copyright 2008,American Chemical Society.)

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6.6 Multi-FRET Systems j231

modifications allowing custom labeling. Altering donor/acceptor spacing is facile inthis configuration and allows fine-tuning of FRET efficiency [569]. Such fine-tuningand control of multi-FRETusing DNA constructs has been used for light harvestingand charge separation using DNA nanostructures [269,567], development of com-binatorial FRET-tags for SNP detection, [571] and DNA-based photonic wires [568].Proteins have also been used in the development of multi-FRET configurations.

Maltose binding protein (MBP), for example, has been either triple labeled withFAM, tetramethylrhodamine, and Cy5, [572] or dual labeled with QD and Cy3 beforebinding Cy3.5-labeled b-cyclodextrin [71] in the development of maltose biosensors.Rogers et al. incorporated a multi-FRET PheCN, Trp, and 7AW system intotwo model protein systems: HP35 and a designed bba motif (BBA5) to study

Figure 6.28 Displacement format usingantibody-functionalized QDs for FRET-basedTNT detection. (a) Schematic of the assay.When TNB-BHQ-10 is bound to the QD-TNB2-45 conjugate, QD fluorescence isquenched via FRET. As TNT is added to theassay, it competes for binding to theantibody fragment and the QD fluorescenceincreases following TNB-BHQ-10 release

from the conjugate. (b) Results from titrationof the QD-TNB2-45-TNB-BHQ-10 assemblywith TNT and the indicated TNT analogues.These assemblies were constructed using530 nm emitting QDs. Each data point is anaverage of three measurements, and errorbars represent the standard deviation.(Reprinted with permission from Ref. [562].Copyright 2005, American Chemical Society.)


urea-induced conformational changes [210]. The nonnatural amino acid fluoro-phores, PheCN and 7AW (Section 6.4.2), were incorporated by mutation of the C-and N-terminus. Such a three-color FRET system extends the working range of themolecular ruler and yields information about the relative positions between thethree fluorophores.In addition to the use of the more traditional organic fluorophores for multi-FRET

applications, substantial development in the areas of NMs, time-resolved fluorescentreagents, and BRET has led to material combinations, with unique advantages inmulti-FRET applications. QDs NMs can act as donors and acceptors in FRETconfigurations as well as provide a scaffold in which to immobilize the multiplecomponents of a multi-FRETsystem. Medintz et al. developed a multi-FRETmaltosebiosensor using QDs (initial donor) functionalized with Cy3-labeled MBP thatfurther bound Cy3.5-labeled b-cyclodextrin, a sugar analogue that was displacedfrom the QD–MBP complex in the presence of maltose [71]. Boeneman et al.developed DNA photonic wires using a QD modified with an ssDNA backbonetemplate that bound four separate short complementary sequences (labeled withdifferent dyes) (Figure 6.29) [568]. The system was used to study the sequentialFRET from the initial QD donor to Cy3/Cy5/Cy5.5/Cy7, and it was found that whilethe initial QD-quenching efficiency was high (80–90%), the amount of energysubsequently emitted by Cy5 and Cy5.5 was relatively low at �2.2 and 1%,respectively, with the Cy7 acting as an IR quencher. QDs can also function assimultaneous donors and acceptors in multi-FRET systems, as demonstrated inDNA hybridization and protease activity detection assays [135,365,366].The use of long-lifetime dyes, such as Ru complexes and LLCs, can provide

additional dynamic information about a system or enhance the biosensing capabili-ties of a platform [365,366,412,413]. The groups of Kumke and Bannwarth, forexample, created a three-color FRET system for protein and DNA analysis using acarbostyril donor–Ru complex (acceptor/relay)–anthraquinone (quencher) combi-nation [412,413]. In the case of DNA analysis, they found that the short luminescentlifetimes gave information about the rotation of the dyemolecules themselves, whilethe long lifetimes yielded information regarding the overall dynamics within theDNA macromolecule itself [412]. Algar et al. developed and characterized multi-FRET systems for monitoring DNA hybridization and protease activity [365,366].Here, a Tb chelate–QD–Alexa647 combination was used in which the Tb chelateassumed the role of initial donor and facilitated time-gated measurements, ulti-mately allowing multiplexed biosensing based on a single-color QD scaffold(discussed in more detail in Sections 6.5.1 and 6.5.5, and also see Figure 6.20).SRET, the sequential combination of BRET and FRET, is a fairly recent develop-

ment in the multiresonance energy transfer arena [317,318,573]. Carriba et al., forexample, labeled three different membrane receptor-interacting proteins with Rluc,GFP, YFP, or DsRed, using SRET to study the heteromers that formed uponexposure to agonists [317]. Protease activity has also been monitored via SRET(Figure 6.30) [318]. Here, the peptide-based probe comprised a peptide sequence(containing the protease-specific cleavage site) flanked by a thermostable fireflyluciferase that produced yellow-green BL (BRET), and a red FP labeled with a near-


infrared fluorescent dye (Alexa680) (FRET). The intact peptide probe undergoesefficient SRET, resulting in acceptor emission of the near-infrared fluorescent dye (at705 nm). Addition of the protease results in a decrease in acceptor emission, due todisruption of the SRETprocess. Xiong et al. developed SRET-based NPs for near-IRin vivo imaging of the lymphatic networks and vasculature of xenografted tumorsin mice [573]. The NPs were composed of a fluorescent polymer, poly[2-methoxy-5-((2-ethylhexyl)oxy)-p-phenylenevinylene] (MEH-PPV), which was subsequentlydoped with the near-IR dye (NIR775) and its surface modified with a COOH-

Figure 6.29 Multi-FRET systems using DNAtemplates. (a) Schematic of the configurationconsisting of 530 nm QDs self-assembledwith four (His)6-peptide-DNA hybridized withCy3 in 1/Cy5 in 2/Cy5.5 in 3/Cy7 in 4. (b)Composite PL spectra from 530 nm QD

donors self-assembled with unlabeled DNA,DNA with Cy3 in position-1, Cy3 in 1/Cy5 in2, Cy3 in 1/Cy5 in 2/Cy5.5 in 3, Cy3 in 1/Cy5in 2/Cy5.5 in 3/Cy7 in 4. (Reprinted withpermission from Ref. [568]. Copyright 2010,American Chemical Society.)


terminated PEG polymer that facilitated Luc8 and tumor-targeting ligand (RGDpeptide) bioconjugation. BRET occurred between the Luc8 and MEH-PPV in thepresence of substrate coelenterazine, with sequential FRET occurring between theMEH-PPV and NIR775. The NPs demonstrated good blood circulation and tumortargeting in mice models. Although there have been fairly limited SRET demon-strations to date, given the improved sensitivity afforded by the self-illuminatingnature of these SRET-based systems utility is bound to increase.

Figure 6.30 Multi-FRET systems – sequentialBRET then FRET – for protease sensing. (a–c)Factor Xa detection using a unique sequentialBRET–FRET combination. (a) The peptide-based probe comprised a peptide sequence(containing the protease-specific cleavage site)flanked by a thermostable firefly luciferase thatproduces yellow-green BL, and a red FP labeledwith a near-infrared fluorescent dye (Alexa680).

(b) When intact the peptide probe undergoesefficient BRET/FRET resulting in acceptoremission of the near-infrared fluorescent dye (at705 nm). (c) Addition of the protease factor Xaresults in a decrease in acceptor emission, dueto disruption of the BRET/FRET process, asillustrated in the time course spectra.(Reprinted with permission from Ref. [318].Copyright 2011, Elsevier.)


6.7Summary and Outlook

FRET is clearly an invaluable biophysical tool for a variety of applications, rangingfrom fundamental studies of the structure and conformation of biological materialsand the in vivo examination of biomolecular interactions, to more applied useswhere FRET signal transduction has been utilized for in vitro and in vivo bioassays,for healthcare diagnostics/screening, defense, environment, and food safety. Therange of new and improved donor/acceptor probe materials continues to expand toaddress some of the inherent complications of more traditional FRET materials,including photobleaching, spectral cross talk and direct excitation of the acceptorspecies. NMs, in particular, are increasingly being used as donor/acceptor probes inFRETstudies due to their many unique photophysical properties and their inherentnanoscaffolding capabilities that can be used to improve FRET efficiency. Hand inhand with donor/acceptor materials development that has expanded the use ofFRET has been the evolving bioconjugation techniques, especially bioorthogonalmethods that facilitate greater site-specific control of the donor/acceptor labeling.With further advances in the areas of NMs and bioconjugation techniques, weanticipate FRET to become an increasingly appreciated tool in a wide range offundamental and applied applications.

Acknowledgments

K.E.S acknowledges Division of Biology, FDA andMCMi, FDA for financial support.K.E.S would also like to thank Ms. S. Spindel and Dr. K. Butler for their commentsand review of this chapter. The mention of commercial products, their sources, ortheir use in connection withmaterial reported herein is not to be construed as actualand/or implied endorsement of such products by the Department of Health andHuman Services.

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