Quantum Dots in Bioanalysis: A Review of Applications Across

focal point reviewELEONORA PETRYAYEVA AND W. RUSS ALGAR*

DEPARTMENT OF CHEMISTRY, UNIVERSITY OF BRITISH COLUMBIA,

2036 MAIN MALL, VANCOUVER, BC V6T 1Z1, CANADA

IGOR L. MEDINTZ

CENTER FOR BIO/MOLECULAR SCIENCE AND ENGINEERING,

U.S. NAVAL RESEARCH LABORATORY CODE 6900,

4555 OVERLOOK AVENUE SW, WASHINGTON, DC 20375 USA

Quantum Dots in Bioanalysis:A Review of Applications Across

Various Platforms forFluorescence Spectroscopy

and Imaging

Semiconductor quantum dots (QDs) are bright-

ly luminescent nanoparticles that have found

numerous applications in bioanalysis and bio-

imaging. In this review, we highlight recent

developments in these areas in the context of

specific methods for fluorescence spectroscopy

and imaging. Following a primer on the

structure, properties, and biofunctionalization

of QDs, we describe select examples of how

QDs have been used in combination with

steady-state or time-resolved spectroscopic

techniques to develop a variety of assays,

bioprobes, and biosensors that function via

changes in QD photoluminescence intensity,

polarization, or lifetime. Some special attention

is paid to the use of Forster resonance energy

transfer–type methods in bioanalysis, including

those based on bioluminescence and chemilu-

minescence. Direct chemiluminescence, electro-

chemiluminescence, and charge transfer

quenching are similarly discussed. We further

describe the combination of QDs and flow

cytometry, including traditional cellular analy-

ses and spectrally encoded barcode-based assay

technologies, before turning our attention to

enhanced fluorescence techniques based on

photonic crystals or plasmon coupling. Finally,

we survey the use of QDs across different

platforms for biological fluorescence imaging,

including epifluorescence, confocal, and two-

photon excitation microscopy; single particle

tracking and fluorescence correlation spectros-

copy; super-resolution imaging; near-field scan-

ning optical microscopy; and fluorescence

lifetime imaging microscopy. In each of the

above-mentioned platforms, QDs provide the

brightness needed for highly sensitive detection,

the photostability needed for tracking dynamic

processes, or the multiplexing capacity needed

to elucidate complex systems. There is a clear

synergy between advances in QD materials and

spectroscopy and imaging techniques, as both

must be applied in concert to achieve their full

potential.

Index Headings: Quantum dot; Fluorescence;

Spectroscopy; Assay; Imaging; Microscopy;

Flow cytometry; Single molecule; Forster reso-

nance energy transfer (FRET); Multiplexing.

INTRODUCTION

In 2002, Applied Spectroscopy pub-lished its first review on quantumdots (QDs), ‘‘Quantum Dots: A

Primer,’’ by Murphy and Coffer.1 Theapplications of these luminescent nano-crystals have evolved tremendouslyover the last decade, particularly in theareas of bioimaging and bioanalysis.Since the seminal first demonstration ofQDs for biological imaging in 1998,2,3

thousands of new research articles onQDs have been published. Researchershave exploited the brightness, photosta-bility, size-dependent optoelectronicproperties, and superior multiplexingcapability of QDs for a myriad of

Received 3 December 2012; accepted 18December 2012.

* Author to whom correspondence should besent. E-mail: [email protected]: 10.1366/12-06948

APPLIED SPECTROSCOPY 215

applications.4–15 Some of the moreprominent applications include in vitrodiagnostics, energy transfer–based sens-ing, cellular and in vivo imaging, anddrug delivery and theranostics.6,16,17 Inparallel with these advances in bioimag-ing and bioanalysis, QD materials havealso evolved to provide greater flexibil-ity and capability. A wider range ofnanocrystal materials, functional coat-ings, and bioconjugate techniques areavailable to facilitate new applicationsof QDs. As we have noted previously,18

QDs have become bona fide multidis-ciplinary tools in much the same way asconventional fluorescent dyes, albeit notyet with the same extent of use. Therehas also been the realization that QDsshould not be viewed as wholesalereplacements for fluorescent dyes, butrather that QDs can be advantageous inmany applications, disadvantageous inothers, and even complementary to dyesin some cases.

The proliferation of QD materialsacross disciplines has been accompaniedby a similar proliferation of moreadvanced spectroscopic technologies,as well as diminishing costs and greatercommercial availability of importantoptical components (e.g., violet lasers,QD-specific filter sets). Indeed, use ofthe unique optical properties of QDs forbioanalysis and bioimaging is mootwithout a suitable measurement plat-form. Fortunately, new developments inapplied spectroscopy and the biologicalapplications of QDs are often synergis-tic. For example, the photoluminescence(PL) properties of QDs are ideal formaximizing the utility of spectral imag-ing and vice versa. Even the well-knownblinking of QD PL, which can compli-cate single molecule tracking, becomesvaluable for super-resolution imaging.19

In this focal point review, we provide anexpanded primer on QDs to complementthat written by Murphy and Coffer,1

briefly summarize the chemistry used tobiofunctionalize QDs, and highlightsome recent (2003–2012) biologicalapplications of QDs in the context ofspecific spectroscopic techniques. Thesetechniques include ensemble fluores-cence measurements based on intensity,polarization, or lifetime; energy andcharge transfer methods; flow cytometryand optical barcodes; enhanced fluores-

cence based on photonic crystals andplasmon coupling; epifluorescence, con-focal, and two-photon excitation (2PE)microscopy; single particle tracking andfluorescence correlation spectroscopy;and super-resolution imaging and near-field scanning optical microscopy. Eachexample highlights how QDs helpenable the full capability of a givenspectroscopic or imaging technique, andvice versa.

OPTICAL PROPERTIES OFQUANTUM DOTS

What is a Quantum Dot? QDs arecolloidal semiconductor nanocrystalswith dimensions between about 1 and10 nm. Excitons are generated in thenanocrystals upon the absorption oflight, and electron-hole recombinationleads to luminescence. Although depict-ed as spheres in most illustrations, QDsare crystalline materials with facets anda lattice structure analogous to the bulksemiconductor material. Depending onits size, each nanocrystal can comprisehundreds to thousands of atoms, a largefraction (.10%) of which are located atthe nanocrystal surface (i.e., a highsurface area-to-volume ratio). As de-scribed in more detail below, most of theQDs used in analytical applications aresynthesized as core/shell structures,where the core nanocrystal is overcoatedwith another semiconductor material toprotect and improve its optical proper-ties. The ‘‘flagship’’ QD material isundoubtedly core/shell CdSe/ZnS.

Absorption and Photolumines-cence. It was the unique photophysicalproperties of QDs that first generatedexcitement for biological imaging andanalysis. QDs have become renownedfor eye-catching photographs (Fig. 1A)of differentially sized QDs under ultra-violet (UV) illumination that show abright rainbow of PL. The bright PL isthe result of high quantum yields (U =0.1–0.9) combined with large molarextinction coefficients (105–107 M-1

cm-1). As shown in Figs. 1C and 1D,QDs have broad absorption spectra thatcontinuously increase in magnitudefrom their first exciton peak to shorterwavelengths in the near-UV. QD PLspectra are shifted to slightly longerwavelengths than the first exciton ab-sorption peak, such that an effective

Stokes shift .100 nm can be achieved.The PL is also spectrally narrow, with anapproximately Gaussian profile (fullwidth at half-maximum [FWHM] of25–35 nm). The stunning rainbow ofQD PL arises from the fact that the peakemission wavelength shifts as a functionof nanocrystal size and material. TheQD size and PL color can be selected bycontrolling the temperature and durationof crystal growth during synthesis.Photographs of the type in Fig. 1Aexemplify the utility of QDs for multi-plexed analyses and multicolor imaging:a single light source can excite manycolors of QD simultaneously (broadabsorption), and each PL contributioncan be readily resolved or deconvolved(narrow emission).

Other advantageous optical propertiesof QDs include excited state lifetimesthat tend to be longer than those offluorescent dyes (.10 ns), superiorresistance to photobleaching and chem-ical degradation (due to the inorganiccomposition and confinement of theexciton), and two-photon absorptioncross sections (103–104 GM) that areorders of magnitude larger than those offluorescent dyes.20 QDs are thus excel-lent probes for tracking dynamic pro-cesses over time, and for two-photonimaging of tissues or other complexbiological specimens where near-infra-red (NIR) excitation mitigates challeng-es associated with autofluorescence andattenuation of excitation light by strongprotein absorbance (e.g., hemoglobin) inthe visible region.21,22

Quantum Confinement and Core/Shell Structures. The size-dependenceof QD PL is the result of quantumconfinement. As a bulk material isreduced to nanoscale dimensions, thedensity of states decreases near theconduction band and valence bandedges, resulting in the emergence ofdiscrete excitonic states. The band gapenergy further increases with decreasingnanocrystal size as the exciton isconfined to smaller dimensions than itsBohr radius. The PL emission wave-length shifts since exciton recombina-tion occurs between the band edgestates. For example, bulk CdSe has aband gap energy of 1.76 eV and a Bohrexciton diameter of 9.6 nm,23 whereasthe band gap energy of 2–7 nm CdSe

216 Volume 67, Number 3, 2013

focal point review

nanocrystals decreases from 2.8 eV to1.9 eV, with PL shifting between 450and 650 nm. The range over which theband gap energy and PL wavelength canbe tuned by quantum confinementdepends on the material of the nano-crystal (vide infra) and its bulk band gapenergy. PL emission centered at wave-lengths between 380 and 2000 nm canbe obtained with appropriate selection ofthe semiconductor material and nano-crystal size.13

Although a QD is approximately aphysical representation of the particle-in-a-box concept, an important differ-ence is that the core nanocrystal does notprovide an infinite potential barrier forconfinement of the exciton. Further-more, the lattice structure of the nano-crystal abruptly terminates at its surfaceand can lead to localized ‘‘trap’’ stateswithin the quantum confined band gap.Trap states can sometimes be observedas band gap emission, showing up as abroad peak on the bathochromic side ofthe expected band edge emission. These

states, as well as leakage of the excitonicwavefunction outside the core nano-crystal, promote non-radiative pathwaysfor recombination of the exciton.23 Toimprove PL efficiency, the core nano-crystal can be coated with a few layersof a structurally similar semiconductorwith a higher band gap energy, as is thecase with widely used CdSe/ZnS andCdTe/ZnS QDs. Such an arrangement,where the core band edge states are bothintermediate in energy to those of theshell, is referred to as a Type Iheterostructure. This configuration isthe most common in bioanalytical ap-plications since it offers the best con-finement of the exciton (Fig. 2A) and thehighest rates of radiative recombination(i.e., brighter PL). Confinement is notcomplete, however, as shell growth istypically accompanied by a 5–10 nmbathochromic shift in the QD PLspectrum.

Other heterostructure configurationsare designed to localize the electron, theholes, or both outside of the core

nanocrystal. For example, in Type IIheterostructures (e.g., CdTe/CdSe,CdSe/ZnTe),24 the electron and holeare localized in the shell and core,respectively, or vice versa. This behav-ior arises from an offset between theband edge states of the core and shell(Fig. 2C). The exciton recombinesacross the core/shell interface and,consequently, the emission wavelengthcorresponds to an energy less than theband gap of either the core or shellmaterial. The decreased overlap betweenthe electron and hole wavefunctions alsoresults in lower absorption coefficientsand longer PL decay times. Type II QDsare potential NIR emitters and growth ofa second Type I shell (e.g., CdSe/CdTe/ZnSe)25 can enhance quantum yields;however, other Type I and alloyed NIR-emitting QDs (e.g., InAs/ZnSe, InAs/CdSe, InAs/InP, Cu:InP/ZnSe, InAsxP1–

x/InP/ZnSe) are also being activelydeveloped.26–28 Quasi Type II QDs haveonly a small offset between, for exam-ple, the conduction band edge states of

FIG. 1. (A) Size-tunable PL of CdSe QDs. The photograph was taken under UV illumination (365 nm). (B) Transmission electron microscopyimage of a CdSe/ZnS QD. [(A) and (B) reproduced with permission from Ref. 18. Copyright American Chemical Society 2011.] (C) Size-dependent absorption and fluorescence spectra of CdSe QDs. [Reproduced with permission from Ref. 23. Copyright American ChemicalSociety 2010.] (D) Absorption and PL spectra of ZnxCd1-xSe QDs with Zn mole fractions of (a) x = 0, (b) 0.28, (c) 0.44, (d) 0.55, and (e) 0.67.[Reproduced with permission from Ref. 56. Copyright American Chemical Society 2003.]


the core and shell, such that the electronis delocalized over the whole nano-crystal but the hole is confined to thecore (Fig. 2D).29 Inverse (or reverse)Type I QDs (e.g., CdS/CdSe, ZnSe/CdSe)30 are designed to localize both theelectron and hole into the shell. Theband edge states for the shell are bothintermediate to those of the core (Fig.2E). These configurations also require asecondary Type I shell (e.g., ZnSe/InP/ZnS)31 to enhance PL emission. Finally,lattice strain between the core and shellcan be used to tune the optical propertiesof certain QDs. For example, growth ofepitaxial shells of ZnS, ZnSe, CdS, orCdSe on small, soft CdTe cores can beused to shift band energies and thus PLemission. Compressive strain in the coreincreases the energy of its band edgestates, whereas synergistic tensile strainin the shell decreases the energy of itsband edge states.32 The effect of grow-ing thicker shells can be large enough toinduce Type II band alignment in a TypeI heterostructure such as CdTe/ZnSe.32

To date, Type II QDs have not found

widespread use in bioanalytical applica-tions.

Surface States and Effects. Theenergies of band edge states are not theonly determinants of QD PL. Even withgrowth of a Type I shell, surface statescan still affect the PL of real QDs (i.e.,imperfect structures). For example, the‘‘blinking’’ or fluorescence intermittencyof QDs, perhaps the second mostrenowned property after their size-tun-able emission, is associated with surfacestates. Blinking can be observed at thesingle particle level, has a power lawprobability distribution, and is a conse-quence of either (i) charging anddischarging of the core nanocrystal, or(ii) trapping of carriers at surface statesbefore they can relax to emissive bandedge core states.33 Auger recombinationis the predominant relaxation mecha-nism in charged QDs, resulting in veryefficient PL quenching until the QD coreis neutralized. Although detrimental insome applications of QDs, the observa-tion of blinking is useful to confirmtracking of a single QD,34,35 and it has

enabled super-resolution imaging,36 asdescribed later in this review.

In addition to blinking, QDs some-times exhibit other interesting opticalphenomena under high-intensity excita-tion. These phenomena include bluing,photobrightening, and photodarkening,all of which are observable in theensemble.37 Bluing corresponds to anirreversible hypsochromic shift in theband edge emission and is the manifes-tation of photooxidative etching of theaverage nanocrystal size.38 Brightening,or photoactivation, is an increase in theQD PL intensity under irradiation and isassociated with changes in the propertiesof the QD surface. These changes havebeen suggested to include the passiv-ation of defect states and danglingbonds,37 or displacement of trappedcharges,39,40 each leading to a decreasein a ‘‘dark fraction’’ of non-luminescentQDs in the ensemble. The extent ofphotobrightening, as well as the oppo-site effect, photodarkening, depends onthe duration and intensity of irradiation,although photodarkening seems to beinduced at higher irradiation intensities,

FIG. 2. Illustration of band gap engineering by selection of core and shell materials. The relative energy of conduction band and valenceband edge states between the core and shell determines the localization of the electron and hole, and the nature of the transition associatedwith exciton recombination, offering an additional means of tuning the optical properties of QDs. (A) Type I QD with localization of bothcarriers in the core. (B) Type II QD with localization of the electron in the shell. (C) Type II QD with localization of the hole in the shell. (D)Quasi-Type II QD with localization of the electron in both and the core and shell. (E) Inverse-Type I QD with localization of both carriers in theshell.


focal point review

above-gap excitation energies, and lon-ger irradiation times. The competitivekinetics of photobrightening and photo-darkening have been investigated andfound to yield different steady-state QDPL intensities for different irradiationintensities.40

The aforementioned dark fraction,which has been observed experimentallyvia fluorescence coincidence analysis, isinversely correlated to the ensemblequantum yield.41,42 It has been suggest-ed that the mechanism for formation ofthe dark fraction is analogous to that forblinking behavior,43 albeit that the darkfraction is not a by-product of blinkingover extended timescales.41,42 Interest-ingly, a decrease in the size of the darkfraction is responsible for the apparentincrease in the ensemble QD quantumyield that is frequently observed upon‘‘passivation’’ with adsorbed macromol-ecules such as proteins.44

The importance of the above-men-tioned effects in analytical applicationsof QDs is variable, depending on boththe characteristics of the batch of QDsused and the spectroscopic parameters ofthe experiment (e.g., laser power).Ensemble assay methodologies basedon one-time measurements at low powerexcitation tend to be relatively immune,whereas single molecule tracking exper-iments with high-intensity excitation arethe most susceptible to these effects. Ineither case, good or poor quality QDscan make a tremendous difference in theoutcome of an experiment. Maintainingcontinuity in the properties of QDmaterials is thus an ongoing challengein the field.

QUANTUM DOT MATERIALS

As alluded to above, QDs have beensynthesized from a broad range ofsemiconductor materials. The most pop-ular materials have been CdSe, CdTe,and their core/shell analogs, CdSe/ZnSand CdTe/ZnS. This popularity can beattributed to well-established syntheticprotocols, emission that can be size-tuned over the visible/NIR region, and,not least of all, commercial availability.Traditionally, emission has been tunedon the basis of core nanocrystal sizewith these materials, and the role of theType I shell has been to passivatedangling bonds on the surface of the

core, better confine the exciton (videsupra) and enhance the QD’s opticalproperties (e.g., the quantum yield canincrease by 20–35%).45,46 For thispurpose, the growth of a thin shell isimportant. For example, with CdSe/ZnSQDs, the 12% lattice mismatch betweenCdSe and ZnS necessitates that growthof the ZnS shell be limited to a fewatomic layers before lattice strain detri-mentally affects the PL properties.17

Thicker shells have been desirable torender QDs more robust or preventblinking.47 Effective approaches forgrowing thicker shells and relaxinglattice mismatch have included incorpo-rating a small amount of Cd into theshell material,48 and synthesis of gradi-ent or multi-shell structures (e.g., CdSe/CdS/ZnS).47,49,50 As an alternative tosize-tuning of PL, QD core materials canalso be alloyed. The PL emission ofternary alloyed QDs (e.g., CdSexTe1–x,CdSxSe1–x, CdxZn1–xS, Cd1–xZnxSe) canbe varied while maintaining a constantsize (Fig. 1D),51–56 and these materialsare also commercially available.

In addition to II–VI semiconductors,other materials used for QD synthesisinclude III–V (e.g., InP)57,58 or groupIV (e.g., Si)59,60 semiconductors. Tosome degree, the investigation of alter-native materials to CdSe and CdTe hasbeen driven by the perceived toxicity ofCd-based QDs (previous work61–63 pro-vides discussions on the complex issueof toxicity; QDs can be used in bothtoxic and nontoxic capacities). Althoughsynthesis protocols for alternative mate-rials are still being optimized to yieldoptical properties that match those ofCdSe/ZnS and CdTe/ZnS QDs, therehas been considerable progress. Forexample, InP/ZnS QDs64 (with emissionin the 480–750 nm range) and InP/ZnSe/ZnS QDs65 have been reportedwith U = 0.4–0.6 and a FWHM of 50–60 nm. In addition to the benefits ofNIR emission for in vivo applications,QD size plays an important role indetermining their fate in vivo. Renalclearance and minimal accumulation inorgans (e.g., spleen, kidney, liver) areobserved with nanoparticles ,5.5 nm inhydrodynamic diameter.66 Recently,Park et al. reported the synthesis ofhighly luminescent CuInxSey/ZnS core/shell QDs (U = 0.6), with emission

within the NIR biological window at741 nm, a FWHM of 175 nm, and anaverage diameter of 5 nm.67 With theexception of the large FWHM, theseQDs are almost ideal for prospective invivo applications. Some non–Cd QDmaterials (e.g., InP/ZnS, InGaP/ZnS) arecurrently available commercially.

Synthesis. Unfortunately, the labora-tory synthesis of high-quality colloidalQDs is still largely restricted to experi-enced chemists. Despite numerous at-tempts in the literature to synthesizeQDs in aqueous media by using conve-nient air-stable precursors, QDs withnarrow FHWM (a function of thedistribution of particle size, i.e., mono-dispersity) and high quantum yieldshave been almost exclusively obtainedthough solvothermal methods that useorganometallic precursors and nonpolarorganic solvents at high temperature andunder inert atmosphere (i.e., pyrolysis ofinorganic precursors).68–70 The possibleexception is the aqueous synthesis ofCdTe QDs, where quantum yields havebeen reported to reach 82% but aretypically closer to 40%.71–73 These QDscan also be relatively monodisperse,with FWHM values typically in therange of 30–60 nm.

Functionalization of QDs. Althoughthe optical properties of QD attract thelion’s share of excitement, experts havenow come to realize that the surface areaof the QD is almost as valuable: A QDcan serve as a nanoscale scaffold withphysicochemical properties and biologi-cal activity that can be tailored throughinterfacial chemistry and bioconjugation.Functionalization is done in multiplesteps, and the design and execution ateach step are critical to the efficacy of theQD in its intended application.18,74–78

Interfacial Chemistry. Since mostbiological applications use core/shellQDs, the inorganic shell is generallythe first site for modification. In partic-ular, high-quality QDs prepared bysolvothermal methods are coated withhydrophobic surfactants and requiremodification to render them water-solu-ble for biological applications. Asshown in Figs. 3i to 3v, there are twowell established routes to water-solubleQDs: (i) ligand exchange (i.e., replace-ment of the native surfactants), yieldingmore compact QDs; and (ii) encapsula-


tion with an amphiphilic polymer (i.e.,building around the native surfactants),typically yielding brighter QDs. Ideally,the core/shell QD PL properties areinsensitive to interfacial chemistry; how-ever, the typical few-atom-thick Type Ishells do not fully isolate the nanocrystalcore, and the optical properties of QDsare still somewhat affected by adsorbedmolecules, pH, temperature, and otherproperties of the local environment.79

This sensitivity is a consequence ofimperfect confinement of the exciton,nonuniform coverage of the shell mate-rial on the core, or both.48 Other

important considerations for the hydro-philic modification of QDs include thenet charge, colloidal stability (i.e.,resistance to aggregation), long-termcoating stability (i.e., stable associationbetween the organic coating and inor-ganic QD), compatibility with bioconju-gate chemistries (i.e., for attachingbiomolecules of interest), and resistanceto the nonspecific adsorption of proteinsand other biomolecules in a samplematrix (i.e., non-fouling). In the follow-ing paragraphs, we describe the chemis-try of coating QDs for aqueoussolubility in more detail, focusing first

on the interface exposed to bulk solutionand then discussing the interface be-tween the organic coating and theinorganic QD.

One of the most widely used methodsfor dispersing QDs in aqueous solutionis to modify their outer surface withanionic carboxylate groups. At suffi-ciently basic pH and low ionic strength,electrostatic repulsion between QDsaffords a stable colloidal suspension;however, efficient charge screening athigh ionic strength, neutralization of thecarboxylates at acidic pH, or both yieldinsoluble aggregates of QDs.80,81 Car-

FIG. 3. Illustrative overview of the chemistry of core-shell QDs. Coatings for aqueous solubility are as follows: (i) amphiphilic polymercoating with carboxyl(ate) groups; (ii) amphiphilic polymer coating with PEG oligomers; (iii) dithiol ligand with a distal PEG oligomer; (iv)dithiol ligand with a distal zwitterionic functionality; and (v) dithiol ligand with a distal carboxyl(ate) group. Common R groups includecarboxyl, amine, and methoxy, although many others can be introduced (e.g., see vi, x, xi). Methods for conjugating biomolecules of interest(BOI) are as follows: (vi) biotin-streptavidin binding; (vii) polyhistidine self-assembly to the inorganic shell of the QD; (viii) amide couplingusing EDC/s-NHS activation; (ix) heterobifunctional crosslinking using succinimidyl-4-(N-maleimidomethyl)cyclohexane-1-carboxylate(SMCC; structure not shown); (x) aniline-catalyzed hydrazone ligation; and (xi) strain-promoted azide–alkyne cycloaddition. The doublearrows are intended to represent conjugation between the functional groups and, in principle, their interchangeability (not reactionmechanisms or reversibility). Not drawn to scale.


focal point review

boxylate coatings (Figs. 3i and 3v) alsotend to be prone to the nonspecificadsorption of proteins due to theircharges. Popular alternatives to carbox-ylate coatings are those featuring poly-(ethylene glycol) ([PEG]; Figs. 3ii and3iii) oligomers or zwitterionic moieties(Fig. 3iv). Both PEGylated and zwitter-ionic coatings offer colloidal stabilityover broad ranges of pH and ionicstrength, and minimal nonspecific ad-sorption for improved biocompatibili-ty.82,83 The advantage of zwitterioniccoatings over those based on PEG ismore compact size;84,85 however, PEGoligomers can be modified with a varietyof terminal functional groups (e.g.,carboxylic acids, amines, hydroxyl,PEG, biotin) with minimal impact onthe overall colloidal stability of theQDs.86

As noted above, there are two mainmethods for modifying QDs with func-tional groups such as carboxylic acidsand PEG oligomers. The first of thesemethods is ligand exchange that in-volves the replacement of hydrophobicsurfactants from QD synthesis withhigher affinity hydrophilic ligands viamass action. The most common ligandsare bifunctional molecules with thiolgroups that coordinate to Zn2þ sites onthe surface of the QD at one end, anddisplay carboxylate or PEG groups atthe other (Figs. 3iii to 3v). Althoughthiols will also coordinate to the Cd2þ atthe surface of a bare CdSe core, the ZnSshell is less prone to oxidation and Zn2þ

has higher binding affinity with basicligands, improving the coating stabilityof the final aqueous QDs.50 Coatingstability is also improved by usingbidentate ligands with two coordinatingthiol groups. For example, an extensivelibrary of bidentate ligands derived fromdihydrolipoic acid have been reported,including those appended with function-al group-terminated PEG oligomers81 orcompact zwitterionic moieties.85 Themajor challenge of ligand exchange withthiols is a reduction in the quantum yieldof the QD. Considerable efforts havebeen made to refine ligand exchangeprocedures to minimize such effects.87–

89 Commercially available QDs withhydrophobic surfactants are often madewater soluble by ligand exchange with

commercially available thiol ligands(e.g., 3-mercaptopropionic acid).3

Amphiphilic polymers are a secondtype of coating that can be applied toQDs, and that are designed to have amixture of hydrophilic groups andhydrophobic alkyl side chains. The alkylside chains interdigitate with alkyl-bearing surfactants from QD synthesis(e.g., trioctylphosphine oxide), leavingthe hydrophilic groups at the surface ofthe now water-soluble QDs (Figs. 3i and3ii). Common chemical strategies forpreparing amphiphilic polymers includepartial grafting of polyacrylic acid orpoly(maleic anhydride) backbones withalkyl amines, where the remaining siteson the backbone are left as carboxylicacids or appended with PEG chains.90–94

These polymer coatings better retain theoriginal brightness of synthesized QDssince they build an additional layer ontothe surface of the QD without alteringcoordination to the inorganic interface(i.e., less opportunity for forming sur-face traps). Polymer coatings also pro-vide good long-term coating stability,but typically larger hydrodynamic radiithan QDs coated with bifunctionalligands.13 Water-soluble QDs with am-phiphilic polymer coatings are availablecommercially, as are QDs coated withphospholipids that interact with the as-synthesized QDs in an analogous man-ner. Further details on the diversity ofpossible coatings for QDs, includingthose that are less widely used or stillemerging (e.g., coordinating poly-mers95,96) can be found in severalreviews.6,18,86,97

Bioconjugation of Quantum Dots.Bioconjugation strategies for QDs canbe broadly classified into (i) covalentcoupling and (ii) self-assembly/specificrecognition; both strategies have beenused to couple enzymes, proteins, pep-tides, antibodies, and oligonucleotides toQDs.6,75 It is critical to note that,without suitable bioconjugation, theutility of QDs in bioimaging andbioanalysis will be greatly hindered,regardless of their highly favorableoptical properties. Furthermore, irrepro-ducibility in bioconjugation tends totranslate into irreproducibility in exper-imental results. A key conceptual differ-ence between QDs and fluorescent dyesis that QDs are effectively surfaces that

can be modified with many biomole-cules at many different sites, whereasfluorescent dyes typically have onereactive group that labels one of manysites on a biomolecule. This differencecreates unique challenges for QDs andother nanoparticles, and these challengeshave been thoroughly reviewed else-where.75 Here, we briefly summarizesome of the most general and pragmaticstrategies for the bioconjugation of QDs,a few of which are illustrated in Figs. 3vito 3xi.

Covalent conjugation methods pro-vide a new chemical bond between abiomolecule of interest and the ligand orpolymer coating of a QD. The robust-ness of the linkage is a function of boththe bond stability and coating stability.The most common chemistry is tocouple amine-bearing biomolecules tocarboxylated QDs (or the oppositeconfiguration) by using amide-bondforming, water-soluble activating re-agents such as N-(3-dimethylaminoprop-yl)-N 0-ethylcarbodiimide (EDC) andsulfo-N-hydroxysuccinimide (s-NHS)(Fig. 3viii).10 This method is an effec-tive ‘‘shotgun’’ method that works wellin some applications and poorly inothers. With many proteins, this chem-istry neither provides good control overthe number of proteins conjugated perQD nor their orientation (potentiallyaffecting biological activity). Anothercommon outcome is a fraction of cross-linked aggregates that tend to result fromcoupling between the large number ofamine and carboxyl groups present onthe surface of a protein. EDC chemistryis often most effective with mono-reactive biomolecules, as is the casefor many synthetic oligonucleotides andpeptides. As an alternative to EDC,some commercial QD suppliers offerbioconjugation kits that target eitheramine or sulfhydryl groups on biomol-ecules, and that couple via hydrazoneligation (Fig. 3x) and heterobifunctionalcrosslinkers with maleimide groups(Fig. 3ix), respectively.98 These reac-tions tend to offer somewhat bettercontrol over the final bioconjugates.The liabilities of conventional covalentconjugation methods have generatedstrong interest in developing highlychemoselective ligation reactions thatprovide excellent control over nanopar-


ticle bioconjugation.75 The aforemen-tioned hydrazone ligation99 is one suchexample, as is copper-free strain-pro-moted azide-alkyne cycloaddition (Fig.3xi; often called ‘‘click chemis-try’’).100,101 Both of these chemistrieshave commercially available ‘‘chemicalhandles’’ that can be used to modifyamine-bearing QDs and biomoleculesfor subsequent ligation.102

Alternative bioconjugation strategiesbased on self-assembly and specificrecognition take advantage of high-affinity noncovalent interactions to as-semble biomolecules of interest to QDs.The best known example of specificrecognition is the tight-binding (femto-molar dissociation constants) betweenbiotin and the avidin family of tetrava-lent proteins (Fig. 3vi). Almost anybiomolecule can be biotinylated usingcommercially available kits and re-agents, assuming that it is not alreadysold with a biotin modification. Strepta-vidin-modified QDs are also availablecommercially, permitting widespreadaccess to a diverse array of QDbioconjugates. This strategy permits amoderate level of control over thenumber of biomolecules assembled perQD (conjugate valence) and their orien-tations; however, there are limitationsassociated with the heterogeneous at-tachment of the streptavidin to theunderlying QD coating.103 To date, thebioconjugate method that has providedthe best overall control is self-assemblybetween polyhistidine-appended bio-molecules and the ZnS shell of ligand-coated QDs (Fig. 3vii; nanomolar dis-sociation constants), thereby providingexcellent control over conjugate valenceand orientation.104 Both expressed pro-teins and commercially synthesizedpeptides can be readily obtained withpolyhistidine tags. Relatively facilemethods have also been developed forchemically ligating these tags to syn-thetic oligonucleotides.105 Polyhistidineassembly has also been extended tocommercial carboxylate polymer-coatedQDs.104 The primary advantage ofpolyhistidine self-assembly and biotin-streptavidin is that bioconjugation pro-ceeds almost quantitatively without needfor excess reagents and purification. Avariety of other self-assembly/recogni-tion methods have been developed, but

they do not yet enjoy the same wide-spread use and accessibility; thesemethods have been reviewed else-where.75

Comparison of Luminescent Nano-particles. The number of nanoparticlematerials with bioanalytical utility hasgreatly increased. In the first half of thedecade, review articles typically com-pared the optical properties of QDs withthose of conventional organic fluoro-phores; however, it is now important tocompare the properties of QDs withthose of other luminescent nanoparti-cles, including nanodiamonds (NDs),carbon nanodots (C-dots), grapheneoxide (GO), carbon nanotubes (CNTs),lanthanide-based upconversion nanopar-ticles (UCNPs), and fluorescent dye–doped silica nanoparticles (DSNPs). Asummary comparison of the physicaland optical properties of these lumines-cent NPs is given in Fig. 4. Note thateach type of nanoparticle has its ownbenefits and liabilities, and each appli-cation will have its own ‘‘best’’ material.We briefly elaborate on some of thecomparisons in Fig. 4 and note a recentreview article for each material below:

� Organic fluorophores: monoreactive;prone to photobleaching, less facilemultiplexing.14

� DSNPs: very bright, good resistanceto photobleaching; much larger insize, less facile multiplexing.106

� NDs: excellent resistance to photo-bleaching, high quantum yield; emis-sion is not easily tunable, lowextinction coefficient.107

� GO: resistant to photobleaching, in-trinsic aqueous solubility; broad PLemission is not easily tuned.108

� C-dots: non-blinking, excellent resis-tance to photobleaching; emissionwavelength depends on excitationwavelength, poorly understood mech-anism of PL.109

� CNTs: excellent resistance to photo-bleaching, NIR emission; challengingto obtain pure samples, weak PLintensity.110

� UCNPs: upconversion luminescence,more narrow emission lines; currentlyless developed coating and bioconju-gate chemistry, multiple emissionlines, larger size.111

Noble metal clusters (NCs) are anoth-

er type of luminescent nanomaterial thathas been gaining significant interest anddeserves some special attention here.These NCs are sometimes referred to as‘‘fluorescent noble metal QDs’’ due totheir optical properties and discrete size-evolved electronic states. NCs containbetween a few and a hundred atoms(e.g., Au, Ag, Pt), are smaller than 2 nm,exhibit no apparent plasmonic proper-ties, and have excitation and emissionbands similar to those of moleculardyes.112 Some very promising workhas been done toward the use of NCsfor biological imaging and analy-sis.112,113 For the purpose of this review,we treat NCs as a material distinct fromsemiconductor QDs. There are manypractical reasons for this distinction,even if both sets of optical properties(e.g., size-dependent fluorescence) arerooted in confinement phenomena. Forexample, NCs are more akin to mole-cules than nanoparticles,114 more sensi-tive to local environment,115 synthesizedquite differently,112,113 and their proper-ties cannot yet be rationally selected tothe degree of semiconductor QDs.113

Prospective biological applications ofNCs have been reviewed else-where,112,113,116 and they are not yet asdeveloped as those with QDs.

BIOANALYSIS ANDBIOIMAGING WITHQUANTUM DOTS

Spectrofluorometry. Most researchand clinical laboratories have access toeither a spectrofluorometer or fluores-cence plate reader. These instrumentsare stalwarts of assay development andhave been widely used with QDs. Here,we highlight in vitro bioanalyses basedon simple fluorescence intensity mea-surements, and, more prominently, For-ster-type energy transfer mechanisms,including fluorescence resonance energytransfer (FRET), bioluminescence ener-gy transfer (BRET), and chemilumines-cence energy transfer (CRET). Othertransduction mechanisms include chemi-luminescence (CL), electrochemilumi-nescence (ECL) and charge transfer(CT) quenching. In the following sec-tions, we denote the peak PL wave-length of QDs by using a subscriptnumber (e.g., QD525); if no material is


focal point review

mentioned explicitly, the reader shouldassume the QD material is CdSe/ZnS.

Fluorescence Intensity. Comparedwith conventional organic dyes, thebroad absorption and narrow emissionof QDs offer significant advantages inmultiplexed assays. Different colors ofQD label are generally associated withdifferent analytes of interest in a hetero-

geneous assay and are interrogatedsimultaneously. For example, Goldmanet al. demonstrated the simultaneousdetection of four toxins—cholera, ricin,shiga-like toxin 1, and staphylococcalenterotoxin B—in a sandwich immuno-assay with QD labels.117 The corre-sponding reporter antibodies wereconjugated with QD510, QD555, QD590,

and QD610, and offered limits of detec-tion (LODs) in the range of 3–300 ngmL-1. The assay was done in a singlemicrotiter plate well with excitation at330 nm. Many similar examples ofspectral multiplexing based on QD PLintensity can be found in the literature.

Fluorescence Resonance EnergyTransfer. A majority of FRET-based

FIG. 4. Comparison of the physical and optical properties of luminescent nanoparticles and organic fluorophores. Check marks ([)indicate the relative degree of favorability.


bioanalyses use QDs as donors fororganic dye acceptors, and these config-urations have several advantages overmore traditional dye–dye pairs. StrongQD absorption in the UV-blue region ofthe spectrum allows selection of anexcitation wavelength that minimizesdirect excitation of the acceptor dye.Furthermore, the narrow and size-tun-able QD PL permits optimization of thespectral overlap integral with onlylimited crosstalk between donor andacceptor emission. The surface area ofthe QD also supports modification withmultiple acceptor dyes, thus enhancingthe rate and efficiency of energy transfercompared with a discrete donor–accep-tor pair. A priori, the strong and broadlight absorption by QDs also suggeststhat they would be ideal acceptors;however, efficient and unavoidable di-rect excitation of the QDs, coupled withtheir relatively long excited state life-time, largely negates this advantagewhen paired with putative fluorescentdye donors (an excited state QD is not agood acceptor). The solution to thischallenge has been to pair QDs as FRETacceptors with luminescent lanthanidecomplexes as donors.12,118 Lanthanideions (e.g., Tb3þ, Eu3þ) typically haveexcited state lifetimes on the order of10-4–10-3 s (cf. 10-9–10-8 s for dyesand 10-8–10-7 s for QDs). As such,directly excited QDs return to theirground state and become good acceptorsfollowing a microsecond delay afterflash/pulsed excitation, whereas lantha-nide ions remain in their excited state asgood donors.12,105 Forster distances canreach ~10 nm with lanthanide-QDFRET pairs and .7 nm with QD-dyepairs,105 compared with ,6 nm withconventional dye–dye pairs.

In bioanalytical applications, the greatadvantage of FRET is the ability to turnQD PL ‘‘on’’ or ‘‘off’’ in response tobiorecognition events (e.g., ligand-re-ceptor binding, enzyme activity, DNAhybridization) or other physicochemicalstimuli (e.g., pH). Since measuredsignals are not strictly based on theaccumulation of QDs, FRET methodscan be applied in the ensemble anddown to the level of single particles.Numerous configurations using QDs andFRET have been reported for thedetection of metal ions,102,119 small

molecules,44,120,121 toxins,122 anddrugs;123 protease124,125 and nucle-ase126,127 activity; hybridization as-says;128,129 immunoassays;130 andpH.131,132 In each case, the underlyingidea is that a donor/acceptor is added orremoved from the vicinity of a FRET-paired QD, either physically (e.g., asso-ciation or dissociation) or through achange in its resonance (i.e., a largespectral shift). FRET-based sensing hasbeen thoroughly reviewed elsewhere,4,16

and we have limited ourselves to a fewmore recent examples here.

Lowe et al. devised a QD-basedmethod for the simultaneous detectionof protease and kinase activity.133 Thesetwo important classes of enzymes arefrequently causative agents of disease,points for therapeutic intervention, orboth. The method used a QD655-AlexaFluor 660 (A660) FRET-pair in combi-nation with a QD525-gold nanoparticle(Au NP) NSET-pair, as shown in Fig.5A. Nano-surface energy transfer(NSET) is a dipole–dipole mechanismthat is conceptually similar to FRET butoccurs over distances approaching 20nm.134,135 A peptide substrate for uro-kinase-type plasminogen activator([uPA], a serine protease) was labeledwith a 1.4 nm Au NP at one terminusand biotinylated at its other terminus forbinding with streptavidin-coated QD525.The QD PL intensity was inverselyproportional to the amount of proteolyticactivity that cleaved the peptide sub-strate to prevent association between theAu NP and QD525 (i.e., loss of NSET).Similarly, a peptide substrate for humanepidermal growth factor receptor 2(HER2) kinase incorporated a terminalpolyhistidine tag for self-assembly toligand-coated QD655, and a tyrosineresidue that was phosphorylated byHER2 kinase in the presence of ATPas a cofactor. A660-labeled anti-phos-photyrosine formed an immunocomplexwith the phosphorylated tyrosine andprovided the proximity for FRET. TheLOD was 50 ng mL-1 for uPA and 7.5nM for HER2, values that were belowthe 200 ng mL-1 and 15 nM, respec-tively, clinical cutoffs for positive/neg-ative breast cancer prognosis.

Although the current state-of-the-artfor QD multiplexing is to use N colors ofQD to detect N different analytes, Algar

et al. have recently shown that time-gatedFRET relays can be designed to detecttwo different analytes by using a singlecolor of QD vector.105,136 The time-gatedrelay comprised an approximately cen-trosymmetric array of luminescent Tb3þ

complexes and fluorescent dyes (AlexaFluor 647 [A647]) around a centralQD620. Due to the millisecond excitedstate lifetime of the Tb3þ complexes, twomodes of interrogation were possible:prompt (~0 ls delay after flash excita-tion; 20 ls integration time) and gated(~55 ls delay after flash excitation; 1 msintegration time). On the prompt time-scale, energy was transferred directlyfrom the QD to the A647 and providedone detection channel. On the gatedtimescale, energy was transferred fromthe Tb3þ to the QD and then to the A647,with analysis of the overall gated PL datayielding a second detection channelbased on Tb3þ-to-QD energy transfer.The time-gated FRET relay was appliedin a model two-plex DNA hybridizationassay (Fig. 5B),105 and in two-plexassays for protease activity.136 The latterassay included tracking the activation ofan inactive pro-protease by anotherprotease, where the activity of both theupstream activating protease and down-stream activated protease were quantita-tively measured. Although demonstratedwith trypsin and chymotrypsinogen, thistype of cascade occurs with many otherproteases, such as the caspases andmatrix metalloproteinases, which areinvolved in important biological signal-ing pathways and implicated in manydiseases. An additional non-multiplexedapplication of the time-gated FRET relayinvolved decoupling Tb3þ-to-QD energytransfer from any biorecognition process,so as to shift the QD-to-A647 energytransfer from the nanosecond time do-main to the microsecond time domain.105

Such a configuration is expected topermit rejection of background scatteringand autofluorescence (lifetime ,20 ns)from complex biological samples, andproof-of-concept was demonstrated forsensing DNA hybridization and proteaseactivity. Clearly, this configuration hasimportant ramifications for zero back-ground in vivo imaging.

Phase-sensitive detection and lock-inamplifiers are a well known tools of thespectroscopy trade and permit resolution


focal point review

of weak signals in very noisy environ-ments. These benefits of lock-in detectionhave been extended to fluorescenceimaging.137 Diaz et al. have suggestedthat photochromic FRET with QD donorsare a means by which to reversiblymodulate QD PL and reject backgroundsignals (Fig. 6).138 To this end, QD540

were coated with an amphiphilic polymerfunctionalized with pendant diheteroary-lethene photochromic dye molecules.The dyes were situated in the hydropho-bic microenvironment around the QDs tofacilitate optimal photoconversion uponmodulation of excitation light (green vs.UV). Outer functional groups on thepolymer were coupled with A647, whichhad constant fluorescence emission andwas used as an internal standard. UnderUV irradiation, the photochromic dyeswitched from an open to closed molec-ular form that was accompanied by theappearance of an absorption band inresonance with the QD PL. Consequent-ly, the QD PL was quenched by 52% dueto FRET. Under irradiation with greenlight, the closed form photochromic dyereverted to an open form with loss of theresonant absorption band and FRET. QDPL lifetimes decreased or increasedcommensurately with the FRET efficien-cy. Cyclic photomodulation of QD PL byalternating UV and visible excitation wasreproducible over 15 cycles. Beyondintensity-based lock-in detection, thesephotoswitchable QDs may have applica-tions in fluorescence lifetime imagingand super-resolution imaging.

FIG. 5. (A) FRET/NSET-based two-plex detection of enzyme activity via two orthogonalself-assembly strategies with QDs (biotin-streptavidin and polyhistidine coordination). (i)Enzyme activity of uPA and HER2 kinase was monitored via the degree of retention of QD

3

PL, which was quenched by the Au NP(NSET) or dye-labeled anti-phosphotyro-sine antibody (FRET) in the absence ofactivity. (ii) QD PL in the absence orpresence of each type of enzyme activity.[Reproduced with permission from Ref.133. Copyright American Chemical Society2012.] (B) Time-gated FRET relay for thetwo-plex detection of DNA hybridizationusing one QD vector. (i) Schematic of thetime-gated FRET relay concept, showingenergy transfer from a luminescent Tb3þ

complex (Tb) to the QD, and from the QD toa fluorescent dye (A647). (ii) QD-probeoligonucleotide conjugates and hybridiza-tion to assemble the FRET relay. (iii)Orthogonal calibration curves for detectionof the two DNA targets. [Reproduced withpermission from Ref. 105. Copyright Amer-ican Chemical Society 2012.]


Bioluminescence Resonance EnergyTransfer. Bioluminescence is the emis-sion of light from an excited state productof a biochemical reaction; for example,via the luciferase enzyme–catalyzed ox-idation of a luciferin substrate. Thistransient excited state can serve as adonor for Forster-type energy transferprovided that an acceptor is in sufficientproximity. The two most common BRETdonors include Renilla luciferase (Rluc),which catalyzes the oxidation of coelen-terazine with emission at ~480 nm; andFirefly luciferase, which catalyzes theoxidation of D-luciferin, with emission at~560–580 nm.139 Since these emissionprocesses do not require incident light, itis possible to use QDs as efficient BRETacceptors. The broad absorption of QDsprovides resonance with the blue-yellowemission of a BRET donor, and thetunable QD PL permits selection of a redor NIR acceptor emission that can becompletely resolved from the BRETdonor emission. This spectral separationis in contrast to organic fluorophores andfluorescent proteins that, due to their

small Stokes shifts, tend to have nontriv-ial emission overlap with BRET donors.

Rao’s group has done extensive workwith QD-BRET, including developingRluc-QD bioconjugates for multicolor,‘‘self-illuminating’’ imaging in vivo,140

and for multiplexed sensing of theproteolytic activity of matrix metallo-proteinase-7 (MMP-7) and uPA incomplex biological samples such asmouse serum and tumor secretions.141

In the latter, two mutant Rluc enzymeswere engineered with C-terminal aminoacid sequences that were both a sub-strate for either uPA or MMP-7 and alinker for conjugation to QDs.141 Con-jugation of the Rluc to the QD providedthe proximity necessary for efficientBRET (Fig. 7A). Hydrolysis of thesubstrate sequences by MMP-7 or uPAdisrupted this proximity, resulting in aloss of BRET-sensitized QD emission.The LODs for multiplexed detectionusing QD655-Rluc (MMP-7 substrate)and QD705-Rluc (uPA substrate) conju-gates were 1 and 500 ng mL-1,respectively. QD-BRET is generallyexpected to provide a multiplexing

capacity of two to four QD acceptorswhen paired with a given biolumines-cent donor system.

Chemiluminescence Resonance En-ergy Transfer. Chemiluminescence isanalogous to bioluminescence, except-ing that no enzyme is involved in thechemical reaction that produces anexcited state emitter. Also analogous toBRET, QDs are good CRET acceptorswhen in close proximity to a chemilu-minogenic reaction (e.g., luminol/hydro-gen peroxide [H2O2]) and canpotentially offer multiplexing capacityof two to four acceptors. Willner’s grouphas exploited catalytic hemin/G-quad-ruplex DNAzymes for the detection ofDNA, metal ions, aptamer–substratecomplexes, thrombin, glucose oxidase,and ATP by using QD-CRET.142–145

The hemin/G-quadruplex DNAzyme ex-hibits peroxidase-like activity that cancatalyze the chemiluminescent reactionbetween luminol and H2O2. For DNAdetection, three different colors of QDs(PL at 490, 560, and 620 nm) werefunctionalized with three different hair-pin oligonucleotide probes that were

FIG. 6. (A) A photoswitchable QD that transfers energy via FRET to a photochromic acceptor and that uses A647 as an internal standard.(B) Fluorescence emission upon excitation at 400 nm after photoconversion with either UV or visible light. (C) Dual-color excitation at 400nm and 600 nm to use A647 emission as an internal standard. [Reproduced with permission from Ref. 138. Copyright American ChemicalSociety 2012.]


focal point review

complementary to three target oligonu-cleotide sequences of interest.144 Thestem segment contained a horseradishperoxidase–mimicking DNAzyme se-quence, and the loop segment wascomplementary to the target. Hybridiza-tion caused opening of the hairpinstructure such that self-assembly of thehemin/G-quadruplex DNAzyme waspossible with evolution of chemilumi-nescence and CRET-sensitized QDemission. The three colors of QDprovided three resolvable signals forthe three targets of interest. Figure 7Billustrates this transduction strategy. Asimilar construct was developed for thedetection of vascular endothelial growthfactor (VEGF), a signaling proteinoverexpressed in cancer cells.146 TheVEGF induced assembly of the QD-hemin/G-quadruplex structure andCRET evolution. When compared withother transduction methods based onFRET (12 nM) and chemiluminescence(2.6 nM), detection of VEGF via CREToffered an improved LOD (875 pM).146

Chemiluminescence. In addition toserving as CRET acceptors, QDs candirectly participate in CL reactions as

the emitter. QD CL is generally obtainedusing H2O2 systems (e.g., H2O2,NaHCO3-H2O2, pyrogallol-H2O2) thatgenerate superoxide, �O2

–, and hydroxylradicals, �OH, that inject an electron intothe QD 1Se quantum confined state anda hole into the 1Sh state, respectively.Subsequent electron–hole recombinationgenerates QD luminescence. The QDCL intensity has been found to dependon the QD size and concentration, theoxidant and its concentration, surfactant,pH, and radical scavengers.147,148 De-spite the numerous advantages of CL-based biosensing (e.g., low background,low LOD, and a wide dynamic range),the use of QD CL remains limited withonly a few proof-of-principle studies forthe detection of metal ions,149 phenoliccompounds,150 and immunoglobulinG.150 Chen et al. demonstrated thedetection of L-ascorbic acid, a CLinhibitor, in human serum by usingCdSe/CdS QDs with a NaHCO3–H2O2

system that forms peroxymonocarbon-ate, HOOCO2

–, a reactive oxygen spe-cies.151 Despite these proof-of-conceptexamples, the origin of QD CL (e.g.,band gap states or band edge emission)

is not entirely clear, and further study isneeded to elucidate the mechanismbefore this transduction method can beused to its full potential.

Electrochemiluminescence. In ECL,a chemiluminescent reaction is initiatedat the surface of an electrode. QDs cangenerate light under an alternatingapplied potential through an annihilationECL mechanism where cationic andanionic QDs neutralize one another toyield a luminescent excited state.152

However, QDs are more frequentlycombined with a coreactant to generateECL. Common coreactants include sul-fite (SO3

2–) or O2 for anodic ECL,153–155

and H2O2 or peroxydisulfate (S2O82–)

for cathodic ECL.156,157 Under theapplied potential, the coreactants areconverted into radicals that transfer anelectron or hole to the electrochemicallygenerated QDs to yield ECL. In mostapplications, QDs are cast onto orcomposited with other materials at thesurface of an electrode. The incorpora-tion with other nanomaterials (e.g.,carbon nanotubes, nanoflowers, gra-phene oxide, gold nanoparticles) isoften observed to enhance ECL intensi-

FIG. 7. (A) Example of QD-BRET: (i) construct with conjugated luciferase protein (Luc8) and (ii) BRET-sensitized PL from QD605, QD655,QD705, and QD800 that can be combined for multiplexed bioanalysis and imaging. The scissors in (i) indicate how a peptidyl linker on theluciferase can be used for sensing protease activity. [Reproduced with permission from Ref. 141. Copyright American Chemical Society2008 (i) and from Ref. 140. Copyright Macmillan Publishers Ltd: Nature Biotechnology 2006.] (B) Example of a QD-CRET construct formultiplexed detection of DNA: (i) QDs emitting at 620, 560, and 490 nm were functionalized with nucleic acid hairpin probe that, afterhybridization with target, formed a catalytic hemin–G-quadruplex that (ii) oxidized luminol to generate CRET; (iii) the luminescence spectrumof a mixture of the three colors of QD conjugates after hybridization with varying amounts of their DNA targets. [Reproduced with permissionfrom Ref. 144. Copyright American Chemical Society 2011.]


ties.158–161 The origin of ECL from QDsis strongly sensitive to surface chemistryand surface states. For example, in initialreports, ECL from CdSe QDs wasobserved from band gap states,162

whereas ECL from CdSe/ZnS QDscorresponded to band-edge emission.163

However, band-edge ECL has also beenobserved from CdSe and CdTe QDslacking shell structures.164–166 Furtherstudy is needed to gain better insightinto the charge transfer reactions at theQD interface and the resulting ECL.

Various ECL-based assays have beendeveloped for the detection of metalions, small molecules, drugs, enzymes,and DNA hybridization and to monitorcell surface carbohydrate expression, asthoroughly reviewed elsewhere.16,167,168

Transduction has primarily relied onanalytes exerting a quenching effect onECL, for example, via competitivecharge transfer or resonance energytransfer (ECL-RET).169,170 A criticallimitation in QD-ECL–based transduc-tion—and one that has persisted formore than a decade since the firstobservation of QD-ECL171—is the ab-sence of spectrally multiplexed assays.If QDs are to ever replace conventionalECL reagents (e.g., luminol), it isessential that they provide a multiplex-ing advantage. The absence of multi-plexed QD-ECL–based assays mayreflect the limited understanding of theQD-ECL mechanism compared with, forexample, well understood QD-FRET, anassay format increasingly used formultiplexed detection. Nonetheless,QD-ECL assays are making stridesforward, particularly in the area ofpotential point-of-care devices. Shi andcoworkers172 recently reported QD-modified carbon tape electrodes on alow cost, paper-based platform for the

"

FIG. 8. ECL detection of dopamine usingCdS QDs on carbon tape electrodes. (A)Instrument schematic. (B) ECL-potentialcurve and corresponding cyclic voltam-mogram of the CdS QD-modified carbontape electrode. (C) ECL intensity as afunction of dopamine concentration. Fivecycles of detection are shown for eachconcentration. The inset shows the corre-sponding calibration curve. [Reproducedwith permission from Ref. 172. CopyrightAmerican Chemical Society 2012.]


focal point review

detection of dopamine (Fig. 8). Thesimple approach of drop casting CdSQDs on adhesive carbon tape providedreproducible ECL over 31 cycles ofalternating cathodic and anodic potential(annihilation mechanism). Dopaminequenched the QD ECL and quantifica-tion was possible between 1 lM and 10mM. Another recent study by Wu et al.described a multiplexed immunoassayfor three cancer antigens—caricinoem-bryonic antigen, a-fetoproten, and pros-tate specific antigen—using ECL-RETon a microchip device with a 64-electrode array.173 Although simulta-neous detection of the three antigenswas demonstrated, multiplexing wasachieved on the basis of spatial registra-tion rather than the spectral multiplexingcapabilities of QDs. CdS nanorods andantigens were spotted on the electrodearray within the microfluidic platformand generated cathodic ECL in thepresence of S2O8

2– coreactant. Theelectrochemically generated CdS excitedstate was able to transfer its energy to atris(bipyridine)ruthenium(II) complex[Ru(bpy)3

2þ] acceptor that was a labelon an antibody bound to the co-immo-bilized antigen. Any antigen present in asample was detected via the increase inQD ECL intensity due to competitivebinding with the Ru(bpy)3

2þ-labeledantibodies. The method was successfulin identifying target cells from a com-plex cell mixture, and the amount ofantigen detected on the cell surface wasin good agreement with expectations.

Charge Transfer and QuantumDots. Many redox-active dyes, metalcomplexes, and other molecules have ademonstrated ability to engage in chargetransfer with QDs and quench their PL.These molecules have included Ru2þ-phenanthroline174,175 and Ru2þ–poly-pyridine complexes,176 ferrocene,177,178

bipyridinium dyes,179–182 and qui-nones,183–185 among others.186,187 Aswe discuss below, the modulation ofCT quenching, like the modulation ofFRET, can be used as an analyticalsignal in bioanalyses. Unlike FRET,however, the mechanism of PL quench-ing in these systems is not whollyunderstood. In general, quenching isthought to be due to photoinducedelectron transfer (PET) between anexcited state QD and a proximal mole-

cule with an occupied or unoccupiedstate intermediate in energy to thevalence and conduction band edge statesof the QD, but many mechanistic detailsremain unclear. A significant challengeis reconciling analytically applied QD-CT systems that generally compriseaqueous core/shell QDs and redox activemoieties as labels on biomolecules, withphysical studies of QD-CT dynamicsthat use hydrophobic core-only QDswith adsorbed redox active moieties inorganic solvent. Although the latter areimportant for applications of QDs in, forexample, solar energy conversion,188 thestudies bypass much of the complexityinherent to bioanalytical systems.

In general, spectroscopic measure-ments (e.g., transient absorption) ofnonaqueous, abiotic QD-CT systemssupport a PET mechanism, but thereare many interesting observations thatsuggest more complex determinants ofCT rates and PL quenching efficiencies.For example, electron transfer is typi-cally faster with smaller QDs,174,182,187

and this result has been attributed to adecrease in the free energy change forthe CT reaction (i.e., energy gap be-tween the QD conduction band state andproximal LUMO of the quencher) as thesize of the QD increases.182 However,other studies suggest a role for surfacestates in the CT mechanism: blinkingdynamics and electron transfer rateshave been correlated;189 so-called‘‘gray’’ states (quenched, but not non-emissive) have been observed with holetransfer to proximal dye molecules189

and with hole-trapping during blink-ing;190 and some of the core only QDsused in CT studies exhibit band gapPL181 (for example). Stationary absorp-tion spectra of a biotic QD-CT system,CdSe/ZnS QDs assembled with Ru2þ-phenanthroline (Ru-phen)–labeled pep-tides, have suggested hole transfer tosurface states with one type of water-soluble ligand coating (negativelycharged) and transfer to both surfaceand core states with another type ofcoating (neutral).174 Here, PL quenchingwas attributed to charging-induced non-radiative relaxation pathways (e.g., Au-ger recombination or hole-trapping)191

that became more efficient with decreas-ing nanocrystal size due to greaterspatial overlap between the transferred

charge and the exciton. Studies ofabiotic QD-CT systems have begun toaddress the effect of ligands (e.g.,length)192 and the binding mode193 ofadsorbed redox active dyes to the QDsurface; however, it is unclear howmuch insight abiotic studies will providegiven the significantly different condi-tions of the experiments. Indeed, onestudy with hydrophobic CdSe/ZnS QDsfound an electrochemical band gap (viacyclic voltammetry) that correspondedto the measured optical band gap, andobserved efficient CT quenching of theQD PL by ferrocene;194 another studywith aqueous CdSe/ZnS QDs foundonly electrochemically active oxidationlevels within the band gap and observedno apparent quenching by ferrocene.174

There is clearly a need for morefundamental studies of the CT dynamicsassociated with QD bioconjugates.

The limited understanding of QD-CTsystems notwithstanding, there is strongpotential for the use of redox activemolecules as dark quenchers for QDs.Compared with FRET, CT quenching isbeneficial in that is does not have therequirement of spectral overlap and mayoffer greater ability to probe redoxactive biological processes. Anotherexpected advantage of charge transferis an exponential dependence of thequenching efficiency on the distancebetween the QD and redox-active moi-ety.175 This sensitivity has been borneout in several unimolecular sensingconstructs developed by Benson’s groupfor the detection of maltose,195,196

palmitate,197 lead,198 and thrombin.175

These constructs have been reviewedelsewhere16 and were based on confor-mational changes associated with recep-tor proteins or oligonucleotides/aptamers upon binding with their cog-nate target. The proteins and oligonu-cleotides were labeled with Ru-phen andconjugated to CdSe/ZnS QDs such thattheir conformational changes altered theseparation between the QD and Ru-phen, resulting in changes in QD PLintensity. Unimolecular sensing config-urations of this type cannot be so readilydesigned with FRET-based transduction,which is less sensitive to small changesin donor-acceptor separation (inversesixth power-dependence). CT quenchingcan also be used in more conventional


on-off formats common with FRET-based detection; for example, Medintzet al. self-assembled Ru-phen–labeledpeptide substrates to CdSe/ZnS QDs asprobes for the detection of the proteaseschymotrypsin and thrombin.174 Initiallyquenched due to the proximal Ru-phen,QD PL was recovered with proteolyticactivity that cleaved the Ru-phen fromthe QD. Enzyme kinetic parameterswere obtained by fitting the assay datawith the Michaelis–Menten equation.Medintz et al. further demonstratedproof-of-concept for highly multiplexedCT quenching with eight colors of QDs(PL maxima at 510, 537, 555, 565, 581,590, 610, and 635 nm), including aGaussian deconvolution algorithm forresolution of each PL signal and theirmodulation by Ru-phen (Fig. 9A).11

In addition to sensing configurationsthat use hole accepting metal complexesto quench QD PL, there are examplesthat use electron accepting molecules,such as quinones, to monitor enzymeactivity199,200 and intracellular pH,183 orbipyridinium, to monitor receptor–sub-strate interactions.180 Initially, Yildiz etal. electrostatically adsorbed a bipyridi-nium dye to the surface of QDs andfound that a macrocyclic receptor,cucurbit[7]uril, could disrupt the CTquenching interaction through competi-tive host–guest interactions.180 Cui et al.reversed this approach by modifyingCdTe QDs with thiolated cucurbit[6]uril(CB[6]) via self-assembly.201 The CB[6]improved the colloidal stability of theQDs and, more pertinently, bound anitrobenzene amine electron acceptorthrough a host–guest interaction, pro-viding the proximity needed for CT

FIG. 9. (A) Resolution of six QD PL signals in a mixture where each QD was quenched to adifferent degree by Ru-phen. The measured composite spectrum is shown in green, and thebest fit to the data is shown in black. Up to eight QD PL signals can be resolved. The inset is

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a cartoon representation of a Ru-phen–labeled peptide undergoing CT with a QD.[Reproduced with permission from Ref. 11.Copyright American Chemical Society2009.] (B) A QD-CT pH sensor: (i) mecha-nism of CT quenching via electron transferbetween the QD an a dopamine modifiedpeptide; (ii) intracellular pH sensing overtime via the microinjection of QD550-dopa-mine conjugates (progressive quenching)and red fluorescent nanospheres (FLX;invariant) into COS-1 cells. [Adapted withpermission from Ref. 183. Copyright Mac-millan Publishers Ltd: Nature Materials2010.]


focal point review

quenching. Both of the above-men-tioned approaches are expected to beuseful for monitoring molecular recog-nition processes or the dynamics ofprotein–ligand interactions.

Quinones are another potent electronacceptor for QDs.183,199,200,202 Freemanet al. functionalized QD620 with peptidescontaining a phosphotyrosine residue (86 2 per QD) to monitor the activity ofalkaline phosphatase (ALP).200 Initially,QDs were unquenched and retained theirPL upon hydrolysis of the phosophotyr-osine residue by ALP. However, in thepresence of a reporter enzyme, tyrosi-nase, the resultant tyrosine residue isoxidized to a dopaquinone residue thatquenched QD PL via CT. Quantificationof ALP was demonstrated over the range0.05–0.5 units in the presence of 25units of tyrosinase. Similarly, directmodification of the QD surface withphosphotyrosine afforded detection ofALP over a range of 0.01–0.13 units.Medintz et al. developed an intracellularpH sensor by taking advantage of thepH-dependent conversion of dopaminefrom its hydroquinone form (acidic pH)to its quinone form (basic pH).183 Thequinone form efficiently quenches QDsvia electron transfer. PEG-coated QD550

were functionalized with peptides mod-ified at their N terminus with dopamine,resulting in progressive quenching ofQD PL as the number of labeledpeptides increased and the pH increased(pH 6–12). For more robust pH mea-surements, the dopamine-peptide–QDconjugates were mixed with pH-insen-sitive red fluorescent nanospheres toprovide a reference signal. The mixturewas microinjected into cells and bymeasuring the QD PL intensity relativeto the red fluorescence nanospheres, itwas possible to quantitatively track thegradual (60 min) increase in cytosolicpH after acidic extracellular medium(pH 6.5) was exchanged for basicextracellular medium (pH 11.5) in thepresence of a cell-permeabilizing drug(Fig. 9B). Naturally, this QD-CT assem-bly also functions as an in vitro pHsensor, and can be interrogated on thebasis of pH dependent changes in QDPL lifetime (measured relative to a pHinsensitive dye such as Cy5).183

Fluorescence Polarization (FP). FPprovides information about the Brow-

nian motion-driven rotational dynamicsof emitting fluorophores. These dynam-ics are a function of the size of afluorophore bioconjugate, such that po-larization increases as the molecularweight of the labeled biomolecule in-creases and rotation slows. Increases inpolarization can thus signal biomolecu-lar interactions (e.g., ligand-receptorbinding) when one of the componentsis fluorescently labeled. The narrowsize-tunable QD PL is also potentiallyideal for developing multiplexed FPassays. For example, Tian et al. demon-strated an FP-based competitive immu-noassay for the simultaneous detectionof two tumor markers in human serum,carcinoembryonic antigen (CEA) and a-fetoprotein (AFP), by using CdTe/CdSQD520 and QD620 conjugated with theseproteins.203 Binding of the correspond-ing antibodies increased the effectivesize of the QD conjugates and increasedthe emission polarization. Notably, thereare only a few examples of QD-basedFP assays reported in the literature. Thisscarcity may be due to the intrinsicallylow polarization of QD emission: QDshave a so-called ‘‘bright plane’’ (emis-sion orientated in any direction alongthis plane) and a ‘‘dark axis’’ (the c-axis)rather than a linear transition dipole likeorganic dyes.204,205 Another potentialchallenge is the large geometric size(comparable to most proteins) and highmolecular weight of QDs, which reducethe relative effect of individual bindingevents on rotational dynamics. The workby Tian et al. suggests that this short-coming can be addressed by takingadvantage of the multivalent nature ofQD conjugates. Since multiple CEA orAFP proteins were conjugated per QD,multiple antibodies were bound per QDto provide a much larger change inrotational dynamics. Such formats mayhold promise for further FP assays basedon QDs.

Flow Cytometry and Optical Barc-odes. In flow cytometry, single cells (orparticles) are optically interrogated in anordered flow stream, including side-scatter intensity; forward-scatter intensi-ty; and, most pertinently, fluorescenceintensity. The parallel detection ofmultiple fluorescent tags associated withcellular biomarkers (i.e., multiplexing) iscritical to the utility of flow cytometry in

high-throughput biomedical researchand clinical diagnosis. In particular, thiscapability allows for complex studiesthat correlate the presence or absence ofmultiple cellular markers with specificimmunophenotypes, responses to certainstimuli, or other investigations wheredistinct subpopulations of cells need tobe resolved. By virtue of their brightnessand multiplexing capability, QDs are anatural fit with flow cytometry. Indeed,the addition of QDs to the repertoire offluorescent labels has increased themultiplexing capacity of flow cytometryto 17, as demonstrated by Roederer’slaboratory.206 Using a combination ofnine organic fluorophores and eightcolors of QD, antigen-specific T cellsfrom a human immunodeficiency virus(HIV)-positive individual were immu-nophenotyped. The fluorescent probesconsisted of QDs functionalized witheither peptide-major histocompatibilitycomplexes or antibodies, and organicfluorophores conjugated to antibodies.QD PL was detected using an octagonalarray of photomultiplier tubes anddichroic/bandpass filters with laser exci-tation at 408 nm (Fig. 10A). The studyfound that T cells specific for a givenpathogen, or even a particular epitope onthat pathogen, can have different phe-notypes, suggesting that the propertiesof individual T-cell clones could play arole in cell differentiation.206 Kovtun etal. conjugated QD655 with agonists fordopamine transporter (DAT) to measureDAT activity and regulation in live cellswith a flow-cytometric method.207 Cellscultured in a microtiter plate wereexposed to agonist candidates, incubatedwith a biotinylated reference agonist(competitive assay), washed, and incu-bated with QD655-streptavidin conju-gates. The cells were then collectedfrom each well in the plate and analyzedvia flow cytometry. The assay was ableto determine half maximal inhibitoryconcentration values for agonists andindicate downregulation of DAT expres-sion demonstrating direct pharmaceuti-cal relevance.207

In addition to traditional flow cytom-etry applications, high-throughput mul-tiplexed screening assays have beendeveloped around flow cytometry in-strumentation by using optically encod-ed bead technology.208 Although


possible (and commercialized) usingfluorescent dyes,209 QDs are a superiorplatform for optical encoding and mul-tiplexed analysis by virtue of theirspectrally narrow PL and the ability toexcite multiple colors of QDs by using asingle laser line.210,211 QD-based barc-odes are read using a combination ofcolor and PL intensity; a barcode systemwith N resolvable levels of PL intensityand m different colors can theoreticallyprovide (Nm – 1) unique codes. Forexample, QD-barcodes designed withsix colors and six intensity levels have acoding capacity .45 000, and thesecodes can be read with 99.7% accuracyby using data processing algorithms.212

Chan’s group recently characterized aQD-barcode–based DNA hybridizationassay, identifying optimal probe oligo-nucleotide and target sequence lengthsfor fast hybridization kinetics and goodhybridization efficiency.213 A modelthree-plex hybridization assay was de-signed using microbeads encoded withQD500 and QD600 at different ratios toobtain three unique codes, and thetargets were non-purified, non-amplifiedDNA sequences from a restriction digestof plasmid DNA. The total samplepreparation and analysis time was ,1h and the dynamic range was 0.02–100fmol.213 Chan’s group also reported asimilar flow cytometric method for rapid(,10 min), sensitive (femtomole level),and parallel detection of genetic markersfor the infectious diseases HIV, hepatitisB, hepatitis C, syphilis, and malaria(Fig. 10B).214 In another related study,

FIG. 10. (A) Schematic of an octagonal photomultiplier tube (PMT) array for eight-colorflow cytometry using QD labels. (i) QD PL spectra and selection of optical components. Bluediamonds indicate the cut-on wavelengths of long-pass dichroic filters; gray bars indicate

3

the transmission range of band pass filters.The colored bars above the spectra indi-cate the spectral detection range for eachQD. (ii) Geometry of the detector array,illustrating the dichroic mirrors (gray) andbandpass filters (colored). [Adapted withpermission from Ref. 206. Copyright Mac-millan Publishers Ltd: Nature Medicine2006.] (B) Example of a spectrally encodedbarcode-based assay using QDs and a flowcytometer: (i) sandwich assay format withbarcoded probe and universal reporter(A647); (ii) assay methodology; (iii) map-ping of three different spectral barcodesbased on the relative PL of QD500 andQD600. The insets show color images of thethree different barcodes. [Reproduced withpermission from Ref. 214. Copyright Amer-ican Chemical Society 2011.]


focal point review

Xu et al. demonstrated a flow cytometricQD-barcode method for the simulta-neous analysis of 10 single-nucleotidepolymorphisims.215 Continued develop-ment of QD-barcode assays is expectedto result in improved methods forsensitive, high-throughput assays forbroad panels of pathogen and diseasemarkers.

Enhanced Fluorescence Tech-niques. Photonic Crystals. Photoniccrystals (PCs) are structures that havesub-wavelength periodicity between twomaterials with different dielectric con-stants, resulting in a ‘‘photonic bandgap’’ that does not allow propagation ofcertain wavelengths of light in some (orall) directions within the PC.216 Cun-ningham’s group has extensively devel-

oped PC slabs as substrates for a varietyof assays with enhanced fluorescentdetection capabilities.217 The slabs com-prise one- or two-dimensional (2D)grating patterns of titanium dioxide(TiO2) on a quartz (SiO2) substrate,and they can be engineered to have twooptical resonances. For a given fluoro-phore, one of these resonances can betuned to its excitation wavelength, thuspermitting amplification of the electricnear-field intensity at the surface of thePC (enhanced excitation); the secondresonance can be tuned to the emissionwavelength and redirect emitted fluores-cence along the PC toward the opticaldetector via Bragg scattering (enhancedextraction). These two enhancementmechanisms are multiplicative and canincrease fluorescence signals by 2–3orders of magnitude.217 For example,2D PC slabs with resonances close to488 nm (excitation) and 616 nm (emis-sion) have been used to enhance the PLof red-emitting CdSe/ZnS QDs by afactor of 108 (Fig. 11).218 The use ofQDs in PC-based biosensing applica-tions is underdeveloped. The potentialchallenge for spectral multiplexing isengineering multiple PC resonances forenhanced extraction of several colors ofQD PL, although the broad QD absorp-tion will permit effective use of theexcitation enhancement across multiplecolors. However, considering the well-established protocols for QD-based im-munoassays,219 and the ability to fabri-cate microtiter plates with embeddedphotonic crystals,220 the future develop-ment of high-sensitivity, high-through-put screening methods with PCs andQDs can be anticipated.

Plasmon-Coupled Fluorescence.The immobilization of QDs on metallicheterostructures or thin films with plas-mon resonances can provide significantenhancements in PL intensity.221 Theseenhancements are a consequence ofmore intense local electric fields forexcitation, increased radiative rates, andcoupling to plasmon modes to yielddirectional, rather than isotropic, emis-sion.222,223 The magnitude of the en-hancement depends on the size, shape,and type of metal substrate, as well asthe distance between the emitter and themetal surface: short distances (e.g., ,5nm) induce fluorescence quenching

through new non-radiative pathways,whereas intermediate distances (e.g.,10–20 nm) permit enhancement via theabove-mentioned processes. The studyand optimization of metal nanostructuresto enhance fluorescence and other scat-tering processes are very active areas ofresearch and have included investiga-tions of QD PL enhancement. In one ofthe earliest reports, Kulakovich et al.used layer-by-layer assembly (LBL) tocontrollably place CdSe/ZnS QDs atfixed distances from Au NPs (12–15nm diameter).224 When the QDs werelocated 11 nm from the Au NPs, a PLenhancement factor of 5 was ob-served.224 In a separate study, Chen etal. found an ~two-fold PL enhancementwhen two colors of QD were LBL-assembled onto ~100 nm Ag NPs at theoptimum separation distance (Fig.12A).225 Subsequently, Song et al.fabricated a periodic silver nano-island(~100 nm) array that produced a 50-foldPL enhancement when the plasmonicfeatures of the substrate were resonantwith QD655 PL emission.226 Pompa etal. observed a similar 30-fold enhance-ment in QD550,598,625 PL when coupledto a periodic nano-pattern of goldtriangles,227 and Leong et al. observeda 15-fold enhancement when QD weresandwiched between a 2D array of goldnanodisks and colloidal Au NPs withcontrolled spacing.228 The current un-derstanding of plasmon enhanced fluo-rescence and the best methods for itsimplementation are far from complete;however, very sensitive assay methodswith QD labels will undoubtedly be builtaround such plasmonic nanostructuresand arrays. For example, a fluorescentdye–based immunoassay on a plasmonicnanostructure array has recently beenreported to provide an ~106-fold im-provement in the achievable LOD com-pared with a planar glass substrate.229

Ultrasensitive multiplexed assays will bepossible if similar enhancements can beobtained with QDs.

Traditional planar metal films, wellknown for surface plasmon resonance(SPR) assays and imaging, can alsosupport plasmon enhanced QD PL. Oneof the first such examples was reportedby Robelek et al., who demonstrated amultiplexed DNA hybridization assayby using SPR-enhanced fluorescence

FIG. 11. (A) Scanning electron microsco-py image of PC slab. The scale bars are 500nm. (B) Enhancement of QD PL intensity: (i)108-fold when the PC is resonant with theexcitation light; (ii) 13-fold enhancementwhen the PC is not resonant with theexcitation light. The circle on the left ofeach image shows the QD PL without a PCstructure. [Adapted with permission fromRef. 218. Copyright Macmillan PublishersLtd: Nature Nanotechnology 2007.]


(Fig. 12B).230 Two target oligonucleo-

tide sequences, complementary to two

different capture probes immobilized on

the gold film, were labeled with QD565

and QD655. Maximum PL intensity from

the QDs was observed at the SPR angle

(618, 543 nm laser excitation), where

excitation of surface plasmons was most

efficient. Jin et al. reported an SPR-

based immunoassay for prostate specific

membrane antigens.231 Plasmon-cou-

pled PL provided a 7-fold enhancement

in the measured PL compared with far-

field excitation of PL, and this enhance-

ment was attributed to directional emis-

sion at the SPR angle (478, 405 nm laser

excitation). Malic et al. also demonstrat-

ed SPR-based DNA hybridization as-

says and immunoassays by using NIR-

emitting QDs as reporters.232 However,

in this work, the reflectivity was used as

the analytical parameter (as in traditional

SPR) rather than PL. The QDs provided

a 25-fold signal amplification due to a

putative combination of its optical mass

(large size and high dielectric constant)

and coupling of its emission into

propagating surface plasmons. The min-

imum detectable DNA concentration

was 100 fM. This type of experiment

FIG. 12. (A) Plasmonic enhancement of QD PL. (i) Change in QD PL as a function of distance (controlled by LBL assembly of polymerlayers) from a cross-hatched patterned array of Au (black squares) and Ag (blue triangles) NPs. (ii) False-colored QD PL image collected with488 nm excitation, showing enhancement of QD PL on a cross-hatched pattern of Ag NPs. (iii) Image of the Ag NP pattern via reflection of the488 nm excitation light, showing strong absorption by the Ag NPs. The white dot is a region with Ag NPs; the green dot is a region without.[Reproduced with permission from Ref. 225. Copyright American Chemical Society 2009.] (B) Surface plasmon–enhanced fluorescenceimaging of a DNA array: (i) instrument schematic; (ii) images obtained from sequential hybridization of DNA targets labeled with green (left)and red (right) QDs; (iii) correlation between the SPR reflectivity (solid line) and QD PL intensity (dotted line) as a function of angle ofincidence. [Reproduced with permission from Ref. 230. Copyright American Chemical Society 2004.]


focal point review

must rely on spatial registration (i.e.,discrete spots) and SPR imaging formultiplexing and thus does not take fulladvantage of the optical properties ofQDs, but it is interesting from theperspective of multimodal detection(SPR and PL).

Microscopy and Imaging. Since thetwo seminal breakthroughs in 1998,when Bruchez et al. demonstrated mul-ticolor imaging of fixed cells by usingQDs (staining F-actin filaments and thenucleus),2 and Chan and Nie demon-strated live cell imaging with receptor-mediated endocytosis of transferrin-QDconjugates,3 QDs have steadily becomea more prominent player in the micros-copist’s toolbox. In this section, wedescribe the utility of QDs in severalmicroscopy techniques.

Epifluorescence Microscopy. Immu-nocytochemical (ICC) and immunohis-tochemical (IHC) labeling have beentwo of the most widespread applicationsof QDs in microscopy. Primary orsecondary antibody-QD conjugates canbe used for direct and indirect staining,respectively, albeit that the latter is oftenpreferred due to the high cost of primaryantibodies and the potential loss ofactivity upon conjugation. Severalgroups have reported labeling of fixedcells and tissue with QD-antibody con-jugates. For example, Wu et al. demon-strated the use of QD probes for ICClabeling of different subcellular targets,including cell surface receptors (HER2),cytoskeletal components (actin and mi-crotubules), and nuclear antigens asso-ciated with SK-BR-3 (human breastcancer) or 3T3 cells (mouse fibro-blast).10 QD-secondary antibody conju-gates, or a combination of QD-streptavidin conjugates with biotinylatedsecondary antibody (or phalloidin foractin), were used for labeling. Turningto IHC, Chen et al. detected caveolin-1and proliferating cell nuclear agent(PCNA) in a lung cancer tissue micro-array.233 Compared with conventionalIHC, QD-based labeling improved de-tection rates from 47% to 57% forcaveolin-1 and from 77% to 86% forPCNA. Similarly, QD-IHC has beenshown to improve the sensitivity ofquantitative detection of the HER2breast cancer marker in formalin-fixedparaffin-embedded tissue specimens.234

Ruan et al. found that QDs also providedexcellent long-term stability (PL re-mained after 2–4 weeks) in addition tobetter sensitivity for IHC labeling ofprostate stem cell antigen in specimensfrom prostate resections or prostatecto-mies.235 QDs are also well suited tomultiplexed staining of tissue biopsyspecimens. Five-color molecular profil-ing of human prostate cancer cells byusing QD-IHC has been described byXing et al.,236 who used one color of QDas an internal standard to target ahousekeeping gene product that wasexpressed at a constant level. Fourbiomarkers—vimentin, N-cadherin, re-ceptor activator of nuclear factor jBligand, and E-cadherin—were labeledwith secondary antibody conjugates ofQD525, QD565, QD605, and QD655,respectively. An internal standard, elon-gation factor-1a, was labeled withQD705 (Fig. 13A). Here, epifluorescencewas used in combination with spectralimaging (vide infra). In general, moreinsight can be obtained from the detec-tion of more biomarkers in parallel,suggesting a key role for the multiplex-ing advantages of QDs in diagnosticpathology.

In addition to labeling fixed cells andtissues, epifluorescence microscopy andQDs have been widely applied for livecell labeling and imaging.237 A report byDelehanty et al. demonstrated spatio-temporal multicolor labeling of liveA594 cells (human alveolar adenocarci-noma) by using mixed delivery tech-niques over several days (Fig. 13B).238

Initially, QD520 were delivered to cellsby using a cationic polymer, and thecells were cultured for 3 days so that theQD520 were largely within late endo-somes. QD635-cell–penetrating peptideconjugates were then delivered to cellsso as to label early endosomes, followedby cytosolic microinjection of QD550-cyanine 3 (Cy3) conjugates and subse-quent incubation with Cy5-QD635-Arg-Gly-Asp (RGD) peptide conjugates tolabel aVb3 integrins on the cell mem-brane. When combined with four narrowbandpass filters, these QD probes pro-vided four distinct spectral windows forimaging each of the aforementionedcellular components with excitation at457 nm. The QD550-Cy3 and Cy5-QD635-RDG conjugates were FRET

pairs where the QDs were used asantennae to enable observation of Cy3/Cy5 PL with excitation at 457 nm(nonresonant with the dye absorption)and a reduction of dye photobleachingrates (indirect excitation via FRET).238

QDs with emission wavelengths analo-gous to those of Cy3/Cy5 can also beused directly, as demonstrated withQD580 instead of Cy3. Additional ex-amples of live cell imaging are discussedbelow in the context of other microsco-py techniques.

Confocal Microscopy. Confocal mi-croscopy is a high-resolution imagingtechnique that is capable of axialsectioning. It particularly benefits fromthe brightness of QDs due to its reducedlight throughput compared with epi-fluorescence microscopy (the trade-offfor higher resolution). Although lessimportant for the spinning disk confocalformat, the enhanced photostability ofQDs is another significant advantage inthe laser scanning format. Matsuno et al.used QDs and confocal laser scanningmicroscopy to generate three-dimen-sional (3D) images of the intracellularlocalization of either growth hormone orprolactin, along with its correspondingmRNA, by using combined immunohis-tochemical labeling (QD655-antibodyconjugates) and in situ hybridization(QD605-oligonucleotide conjugates).239

The spatial distributions of mRNA andits encoded protein provide insight intocellular protein synthesis. Chan et al.used similar methodology to simulta-neously visualize vesicular monoaminetransporter 2 mRNA and tyrosine hy-droxylase protein within the cytoplasmof dopaminergic neurons.240 Lee andcoworkers used confocal imaging withQDs to observe agonist-induced endo-cytosis of two types of tagged G protein-coupled receptors (GPCRs): (i) influenzahemagglutinin (HA) peptide-tagged k-OR and (ii) green fluorescent protein(GFP)-tagged A3AR (adenosine recep-tors).241 These GPCRs are overex-pressed on the membrane of humanosteosarcoma cells. The k-OR receptorwas labeled with QD-anti-HA (anti-body) conjugates, and the QD and GFPPL signals allowed real-time parallelvisualization (Fig. 13C) of the internal-ization of both GPCR types whenstimulated with agonists. This format is


potentially useful for high-throughputscreening of agonists for drug discovery(GPCRs are involved in several majordiseases).241

Spectral Imaging. Spectral imagingincludes both the hyperspectral andmultispectral varieties, and offers supe-rior informing power compared withconventional optical filter–based imag-ing. Image cubes (x, y, k) are collectedand comprise a stack of planar imagesacquired across a series of wavelengths,k, or bands thereof (e.g., a PL spectrumat each image pixel). Even with emis-sion overlap between different fluoro-phores, the use of spectral unmixing(i.e., deconvolution) and a library ofreference spectra permits quantitativeresolution of the unique fluorescencecontribution from each emitter anddiscrimination of background autofluo-rescence. This technical capability isideal for pairing with the multiplexingadvantages of QDs to facilitate detectionand visualization of multiple biomarkersin complex biological specimens. Re-cent studies have highlighted the utilityof QD-based spectral imaging for study-ing human cancers. Li’s group usedmultispectral imaging and IHC labelingwith QD605-streptavidin conjugates (viabiotinylated secondary antibodies) toidentify three molecular markers inbreast cancer tissue: HER2, estrogenreceptor, and progesterone recep-tor.242,243 Five different molecular sub-types of breast cancer cell heterogeneitywere identified and corresponded todifferent 5-year patient prognoses.243

Spectral imaging was largely used toseparate out tissue autofluorescence, andthis separation was further aided by thebrightness of QDs. In contrast, Nie’s

FIG. 13. (A) Four-plex profiling of tumor biomarkers. Fluorescence image (left) of highlymetastatic prostate cancer cells acquired with a color charge-coupled device camera. Thecells were labeled with QD565-, QD605-, QD655-, and QD705-antibody conjugates bound to N-cadherin, elongation factor (EF)–1a, E-cadherin, and vimentin, respectively. Single-cell PLdata are shown on the right (acquired by spectral imaging). [Adapted with permission fromRef. 236. Copyright Macmillan Publishers Ltd: Nature Protocols 2007.] (B) Multicolorspatiotemporal strategy for labeling various subcellular compartments and structures with

3

QDs and an example of a correspondingepifluorescence image. [Reproduced withpermission from Ref. 238. Copyright Amer-ican Chemical Society 2011.] (C) QD-basedscreening of GPCR agonists using QD-anti-HA conjugates with specific binding to HA-k-OR overexpressing cells (U2OS). Ago-nist-induced translocation of k-OR is clear-ly tracked by migration of membrane-bound QD conjugates to the cytoplasmicregion. The cells also express a GFP-labeled adenosine receptor (A3AR). [Adapt-ed with permission from Ref. 241. Copy-right John Wiley and Sons 2012.]


focal point review

group has used multispectral imagingwith multiple colors of QD to reliablydetect and characterize tumor cells inc o m p l e x t i s s u e m i c r o e n v i r o n -ments.244,245 In one study, four proteinbiomarkers—CD15, CD30, CD45, andPax5—were simultaneously mapped inlymph node biopsy specimens by usingfour colors of QD-secondary antibodyconjugates (QD525, QD565, QD605, andQD655) as shown in Fig. 14A.244 This

method permitted reliable identificationof low-abundance (~1%) Hodgkins’sand Reed–Sternberg cells; identifyingsuch cells is essential for differentiatingHodgkin’s lymphoma from non-Hodg-kin’s lymphoma and benign lymphoidhyperplasia.244 In another study, fourprotein markers associated with prostatecancer—E-cadhedrin, high-molecular-weight cytoketarin (CK HMW), p63,and a-methylacyl CoA racemase

(AMACR)—were quant i t a t ive lymapped in biopsy specimens (Fig.14B).245 As in the previous study, thebiomarkers were detected using fourdifferent colors of QD-secondary anti-body conjugate (QD565, QD605, QD655,QD705), revealing molecular and mor-phological details that are unseen withtraditional staining methods. One of themost important observations was thatprogressive changes in benign prostate

FIG. 14. (A) Multispectral imaging of QDs for detection of rare Hodgkin’s and Reed–Sternberg (HRS) tumor cells in Hodgkin’s lymphoma.The images show HRS malignant cells and infiltrating immune cells on lymph node tissue specimens. The HRS cells (arrows) exhibitedcharacteristic staining pattern: membrane staining (CD30 positive, red), Golgi staining (CD15 positive, white), and nuclear staining (Pax5positive, blue). Staining patterns were clearly distinct from infiltrating B cells (blue nuclear staining) and T cells (green membrane staining).The scale bar in the top image is 100 lm; the scale bar in the bottom image is 10 lm. [Reproduced with permission from Ref. 244. CopyrightAmerican Chemical Society 2010.] (B) Composite spectral image of a single malignant cell (arrow) in the basal layer of a largely benignprostate gland, surrounded by malignant cells (blue staining surrounded by dotted lines). The scale bar is 20 lm. Four different proteinbiomarkers, labeled with different colors of QD-antibody conjugates, are highlighted in green (E-cadherin), white (CK HMW), blue (AMACR),and red (p63). [Reproduced with permission from Ref. 245. Copyright American Chemical Society 2010.] (C) Five-color immunohisto-chemical labeling of a fixed mouse spleen tissue section with QD-antibody conjugates, imaged via multispectral imaging. A mergedfluorescence-false color image is shown, highlighting the following cellular biomarkers: CD45/blue, CD31/yellow, CD11b/aqua, CD4/green,and CD11c/red. [Reproduced with permission from Ref. 98. Copyright American Chemical Society 2011.]


glands start with a single malignant celland ultimately lead to a malignantgland.245 Overall, spectral imaging withQDs is an ideal methodology fortackling the challenge of tumor hetero-geneity that exists at the molecular,cellular, and tissue-architecture levels,as well as between individuals. A greaterunderstanding of tumor growth mecha-nisms and more rigorous classificationschemes are expected to lead to moreeffective stage-specific and personalizedtreatments of cancer. Jennings et al.demonstrated spectral imaging of fixedmouse spleen tissue sections IHC la-beled with five colors of QD-antibodyconjugate (QD525, QD565, QD605,QD625, QD650).98 B cells, T cells,leukocytes, thymocytes, and macro-phages were labeled with the QDs viaantibodies targeting CD45 (type Cprotein tyrosine phosphatase receptor),CD11c (integrin aX), CD31 (plateletendothelial cell adhesion molecule),CD4 (a glycoprotein), and CD11b(integrin aM) antigens, respectively(Fig. 14C).98

Another prospective application ofspectral imaging with QDs is intracellu-lar thermometry. QDs exhibit a bath-ochromic shif t wi th increasingtemperature due to alteration of electronlattice interactions.246 Yang et al. usedthis effect to measure intracellular heatgeneration in NIH/3T3 murine fibroblastcells after Ca2þ stress and cold shock.247

QD PL collected from the specimenthrough an inverted microscope wasdirected to a spectrograph to measuretemperature-dependent spectral shifts,revealing an inhomogeneous intracellu-lar temperature response. The impor-tance of intracellular temperaturemeasurements is described further inthe section on fluorescence lifetimeimaging microscopy (FLIM).

Two-Photon Fluorescence Micros-copy. Another technique widely used forbiological imaging is two-photon fluo-rescence microscopy (2PE). It offershigh resolution, comparable to that ofconfocal microscopy, by limiting exci-tation of fluorescence to a small focalvolume where there is sufficient flux ofNIR photons for 2PE. The addedadvantage of 2PE is that NIR excitationpenetrates more deeply into tissues andgenerates less autofluorescence than the

visible excitation used in conventional(one-photon excitation [1PE]) confocalmicroscopy. Multiple fluorescent dyescan also be simultaneously interrogateddue to frequent overlap in their 2PEspectra, even when their 1PE spectrahave minimal overlap.248 2PE fluores-cence microscopy, however, is stillprone to photobleaching of dyes,248

and such dyes already have limitedbrightness in 2PE due to their smalltwo-photon absorption cross sections(typically,300 GM).249 It is here thatQDs are distinctly advantageous due totheir resistance to photobleaching andremarkably large two-photon absorptioncross sections (103–104 GM).14,20

One application of QDs in 2PEmicroscopy is high resolution cellularimaging with minimal autofluorescencebackground and photodamage. For ex-ample, Wang et al. imaged CdTe QDs inhighly autofluorescent BY-2-T (tobacco)cells and found that the signal-to-noise(S/N) ratio, or QD PL-to-autofluores-cence ratio, increased from ~3 to 11when the switch was made from 1PE at405 nm to 2PE at 800 nm.250 This studyalso found that, unlike 1PE at 405 or488 nm (2 mW), 2PE at 800 nm (20mW) caused no discernible photodam-age to QGY human hepatocellularcarcinoma cells. Bharali et al. deliveredfolate-conjugated InP/ZnS QDs to KBcells (human epithelial) via receptor-mediated uptake and imaged their accu-mulation in multivesicular bodies with2PE at 800 nm.251 Geszke et al.similarly used folate conjugation and2PE to deliver and image ZnS:Mn/ZnSQDs in T47D and MCF-7 breast cancercells.252 Tu et al. demonstrated thatparamagnetic Mn-doped Si QDs couldfunction as a non-cytotoxic, multimodalcontrast agent for magnetic resonanceimaging and 2PE fluorescence imagingof macrophage cells.253 Selvin’s groupwas able to extend their 2D, 1PEfluorescence-imaging-with-one-nanome-ter accuracy (FIONA) technique to 3Dimaging by using 2PE in combinationwith QDs (Fig. 15).254 2PE FIONA wasfirst validated by tracking the steps ofindividual QD655-labeled myosin Vmotors on F-actin with nanometer accu-racy (step size, 35.8 6 6.3 nm for 2PEand 35.4 6 7.0 nm for 1PE). Impres-sively, widefield illumination at 785 nm

was used for the 2PE tracking and wasuniquely enabled by the efficient 2PE ofQDs. To achieve 3D imaging, widefieldillumination was replaced with multi-point scanning (x-, y-, z-axes) of a 9 3 9array of diffraction-limited focal spotsby using a holographic beam-splitter.LamB receptors on live Escherichia colicells were labeled with QD605 and foundto occur as spatial helices or bands onthe cell membrane. In a second exper-iment, basal breast cancer cells withmembranous epidermal growth factorreceptors (EGFR) were incubated withQD605-epidermal growth factor conju-gates to induce internalization. Afterfixing the cells, 3D-2PE FIONA wasable to visualize individual QD-labeledEGFR endosomes with an (x, y, z)accuracy of ~ 2–3 nm. The 2PE-FIONAtechnique provided a five-fold enhance-ment in S/N compared with imagingwith 1PE total internal reflection mi-croscopy.254

Another useful application for QDs in2PE microscopy is in vivo imaging.Larson et al. demonstrated in vivo 2PEimaging of mouse vasculature (i.e.,angiography) in skin and adipose tissueby injection of amphiphilic polymer–coated CdSe/ZnS QD550 and 2PE at 880nm.20 Conventional fluorescein–dextran(70 kDa) angiography (2PE at 780 nm)revealed less detail than the QD-basedangiography, even when using five-foldmore excitation power at half the tissuedepth. Collagen was imaged in parallelwith the QDs via its second harmonicgeneration. Stroh et al. similarly dem-onstrated two-color 2PE (at 800 nm)imaging for tracking cells in vivo.255

QD590 were conjugated with the HIVTAT protein to label bone marrowlineage–negative cells. The cells wereco-injected with phospholipid-coatedQD470 into a mouse with an MCaIVtumor, where the QD470 PL permittedimaging of blood flow and the QD590 PLpermitted tracking of the recruitment ofbone marrow–derived cells to the tumor.Although the NIR excitation used in2PE microscopy addresses the penetra-tion of tissue by excitation light, thedepth of imaging remains limited (,500lm)21 by scattering and absorption ofshorter wavelength fluorescence (e.g.,visible wavelengths) emitted from with-in the tissue. Although the brightness of


focal point review

QDs with NIR 2PE can help to addressthis challenge, another significant ad-vantage of QDs in this regard is theability to select certain sizes and mate-rials to emit NIR PL. For example, CdTeQD800 can be imaged using 2PE at 900nm, such that both the excitation andemission wavelengths fall within thetissue optical window between ~650and 950 nm.256 Measurements on tissuephantoms suggested imaging could bedone at a depth of 1.6 mm, approxi-mately twice the depth of imaging with1PE of NIR CdTexSe1–x/CdS ternaryQDs (albeit that the latter was measuredwith real heart and femur tissue257).

Single Molecule/Particle Spectros-copy. Single Molecule/Particle Imag-ing. Single molecule spectroscopyprovides access to physicochemicaldetails that are lost when averaging overthe ensemble; for example, revealingheterogeneous subpopulations or asyn-chronous dynamics.258,259 Due to theirbrightness and resistance to photo-bleaching, QDs are ideal for singlemolecule or, more accurately, particle

measurements. The propensity of QDsto blink is both a benefit and a liabilityas it confirms tracking of a single QDbut also complicates the analysis of PLtrajectories.260 Since size polydispersitydoes not exist at the level of single QDs,the FWHM of their PL tends to decreaseto ~15 nm.260

Single QD detection has been used tomeasure biomolecular interactions (e.g.,DNA hybridization, DNA-protein bind-ing, ligand-receptor binding), and todetect small molecules via two-colorcolocalization, FRET, and, more recent-ly, charge transfer.261–263 In the latter,CdSe/ZnS QD560 were functionalizedwith ferrocene-maltose binding protein(MBP) conjugates, resulting in CTquenching (Fig. 16A).263 MBP under-goes a conformational change (scissor-ing) upon binding maltose, resulting inan increase in the distance between theferrocene and the QD with a corre-sponding increase in the QD PL. Bymonitoring individual QDs, it waspossible to detect maltose with a dy-namic range spanning a remarkable five

orders of magnitude (100 pM–10 lM).Interestingly, unlike single-pair FRETsystems that show on-off switching ofQD PL, the single QD-CT systemexhibited constant emission with anincrease in intensity upon binding malt-ose.263 This behavior was attributed to a‘‘gray’’ state associated with CT. Solu-tion-phase single-pair FRET (spFRET)with QDs has also been used as aplatform for sensitive assays. For exam-ple, Zhang et al. conjugated streptavi-din-QD605 with biotinylated captureprobe oligonucleotides for a sandwichDNA hybridization assay, where thereporter oligonucleotides were labeledwith Cy5 acceptors.264 spFRET wasmonitored by flowing the sample solu-tion (mixed with QDs, capture andreporter oligonucleotides) through aglass microcapillary with laser excita-tion at 488 nm in a small observationvolume. A fluorescence burst coinci-dence analysis between the donor andacceptor detection channels was used toobserve FRET and measure DNA hy-bridization. The LOD was 4.8 fM target,

FIG. 15. 2PE-FIONA 3D imaging with QDs. (A) Instrument schematic. The holographic 9 3 9 excitation matrix used for 3D imaging is shownin the inset. (B) Images of live E. coli cells labeled with QD605, illustrating the superior resolution of 2PE widefield imaging compared with1PE widefield imaging. (C) 3D localization of single QD655-EGFR receptor conjugates on the membrane of a breast cancer cell. [Reproducedwith permission from Ref. 254. Copyright American Chemical Society 2011.]


FIG. 16. (A) Single molecule biosensing using QD-MBP-ferrocene conjugates: (i) schematic of the experiment illustrating the decrease inCT quenching efficiency upon binding maltose; (ii) schematic of the microfluidic channel used for sample delivery and the optical setup; (iii)titration curve for maltose detection spanning 100 pM–10 mM. [Reproduced with permission from Ref. 263. Copyright American ChemicalSociety 2012.] (B) Illustration of QD-DNA conjugates and representative 3C burst coincidence analysis results for the two-plex detection ofDNA hybridization using single molecule fluorescence detection and spFRET: (i) HIV-1 gene detection, signaled by the coincidence of Alexa


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a value 100-fold better than the LODassociated with molecular beacons (0.48pM). The assay was applied to thedetection of Kras point mutations inclinical samples from patients withovarian cancer.264 Subsequently, Zhangand Hu extended this spectroscopicmethod to the multiplexed detection ofthe HIV-1 and HIV-2 genes, relying ona three-color (3C) burst coincidenceanalysis (Fig. 16B).265 Here, two sand-wich DNA hybridization assays weredone in parallel using streptavidin-QD605, two biotinylated capture probestargeting HIV-1 and HIV-2, and corre-sponding reporter oligonucleotides thatwere labeled with either Alexa Fluort488 (A488) or A647, respectively.Directly excited (488 nm laser line)fluorescence bursts from A488, whencoincident with QD605 PL bursts, wereindicative of presence of HIV-1 se-quence. FRET-sensitized fluorescencebursts from A647, when coincident withQD PL bursts, were indicative of HIV-2.265 This study is another example oftaking advantage of the surface areaoffered by QDs to assemble multiplebiomolecular probes.

In addition to bioanalytical applica-tions, Pons et al. used spFRET betweenCdSe/ZnS QDs and dye-labeled poly-histidine-tagged MBP to confirm thatbiomolecules self-assemble to QDs ac-cording to a Poisson distribution.266

That is, an ensemble with an averageof N biomolecules (e.g., proteins, pep-tides, oligonucleotides) per QD is actu-ally heterogeneous and comprisesdiscrete subpopulations with . . . N – 2,N – 1, N, Nþ 1, Nþ 2 . . . biomoleculesper QD, where the abundance of eachsubpopulation is weighted according toPoisson statistics. With an average offour or more biomolecules per QD, theunconjugated subpopulation decreasesto 2% of the ensemble.

Single QD tracking has also beenused to study the diffusion dynamics ofmembrane proteins associated with neu-

ronal cells, including the distributionand surface mobility of neuroreceptorsfor glycine,267 c-aminobutyric acid,268

a7 neuronal nicotinic acetylcholine,269

and 2-amino-3-(3-hydroxy-5-methyl-isoxazol-4-yl)propanoic acid,270 at bothsynaptic and extrasynaptic sites. Inanother example, Tsien’s group studiedthe vesicular secretion of neurotransmit-ters by loading individual synapticvesicles with single QD605.271,272 Track-ing of these QDs, in combination with asmall pH-dependent change (15%) inQD PL intensity between vesicles (pH5.5) and the extracellular matrix (pH7.3), was used to distinguish betweenfull collapse fusion (complete vesicleintegration with membrane) and kiss-and-run ([K&R], transient vesicularfusion with membrane, retrieval, andrelease) mechanisms. It was found thatK&R was more prominent under high-activity demand conditions. Lowe et al.investigated the selectivity mechanismof the nuclear pore complex in HeLacells by tracking single QDs decoratedwith multiple copies of importin-bbinding domain of snurportin-1, theimport receptor for small moleculenucleoriboproteins.273 The import orrejection of the QD cargos was moni-tored, and the authors were able toderive a model for cargo translocationthat consisted of cargo capture, filtering,translocation, and release into the nucle-us. Wells et al. demonstrated that QDscould be used to track the 3D molecularmotions inside live cells and on theirsurface (Fig. 16C).274 The imagingsystem used four diffraction-limitedoverlapping confocal volume elements,implemented with four optical fibers,and provided a localization accuracy of50 nm along the x- and y-axes, and 80nm along the z-axis, with 5 ms temporalresolution. Experimentally, QD-immu-noglobulin E (IgE) conjugates werebound to high-affinity IgE receptors(FceRI) on rat tumor mast cells. Themotion of single QD-IgE-FceRI com-

plexes was tracked, revealing 3D surfacetopology of the cell membrane andpermitting extraction of diffusion rates(0.21 lm2 s-1). When the complexeswere crosslinked with the addition ofdinitrophenyl-bovine serum albumin an-tigen, it was possible to observe ligand-mediated endocytosis and measure therate of vesicular transit (950 nm s-1).274

Fluorescence Correlation Spectros-copy. Fluorescence (cross) correlationspectroscopy [F(C)CS] measures time-based intensity fluctuations in one ormore fluorescence signals as singleemitters diffuse in and out of a verysmall observation volume. Analogous tothe single particle methods discussedunder Single Molecule/Particle Imaging,the brightness of QDs and their photo-stability are significant advantages inF(C)CS, although their blinking behav-ior can again introduce extraneousfluctuations.275,276 Nonetheless, FCShas become an important tool forphysically characterizing QDs. It hasbeen widely used to determine thehydrodynamic radii of QDs functional-ized with different organic coatings orunder different conditions, and it hasbeen proposed as a method for accu-rately measuring QD concentrations andextinction coefficients.275,277,278 In ad-dition, Cramb’s group has publishedseveral studies where QD-FCCS hasbeen used to monitor ligand–receptorinteractions. A pair of early studiesinvestigated the streptavidin–biotin in-teraction in association with QDs (i.e.,QD-streptavidin and QD-biotin conju-gates).279,280 It was found that thestreptavidin–biotin dissociation constant(Kd = koff/kon) increased from 10-15 Mwith the native system to 10-10 M inassociation with QDs, where the asso-ciation rate (kon) decreased by 6 ordersof magnitude and the dissociation rateincreased by 1 order of magnitude.280

2PE-FCCS was also used to interrogatethe binding kinetics between the agonistLeu-enkephalin (BLEK) and human l

3

Fluor 488 (A488) and QD PL; (ii) HIV-2 gene detection signaled by the coincidence of A647 and QD PL; and (iii) simultaneous detection of HIV-1 and HIV-2 genes via the coincidence of A488, QD, and A647 PL. [Adapted with permission from Ref. 265. Copyright American ChemicalSociety 2010.] (C) 3D tracking of single QDs using an optical fiber–based confocal imaging system: (i) instrument schematic; (ii) 3Dtrajectory for a QD–IgE-FceRI conjugate over 250 s, where early times are indicated in red and late times are indicated in blue; (iii) image ofthe cell for which the trajectory was generated. [Reproduced with permission from Ref. 274. Copyright American Chemical Society 2010.]


opioid receptor (hMOR) in cell mem-brane nanopatches.281 QD605 were con-jugated with BLEK to bind the hMORreceptors, and the FCCS data wereconverted into fractional receptor occu-pancy for Hill plot analysis and extrac-tion of Kd. Interestingly, the conjugationof BLEK to comparatively large QDsdid not affect its binding properties.Cramb’s group has developed 3C-FCCSthat is greatly facilitated by the multi-plexing advantages of QDs (Fig.17).281–283 Polymer spheres barcodedwith QD525, QD605, and QD655 could besuccessfully identified and quantitatedin the presence of more than an 800-foldexcess of free QDs, and the size of thetriply labeled spheres could be deter-

mined.281 3C-FCCS was also able toidentify and determine the size of DNAtrimers (~80 nm) labeled with the threecolors of QD.282 The technique does notrequire long acquisition times (,1 min)and is expected to be useful formeasuring molecular exchanges in sig-nal transduction pathways, or the as-sembly of tripartite biomolecularcomplexes.

Wiseman’s group has used QDs aslabels in image correlation spectroscopy(ICS) for measuring biomolecular diffu-sion constants.284–286 Although theblinking of QDs affects diffusion mea-surements made with temporal ICS,reciprocal space (k-space) ICS (kICS)can separate PL fluctuation contributions

due to blinking and transport.284,285

kICS has been used to measure thediffusion coefficient of QD-labeled gly-cosyl phosphatidylinositol–anchoredprotein CD73 in the membrane of livefibroblast cells.284 Furthermore, by tak-ing advantage of changes in blinkingautocorrelation as a function of QDclustering, it was possible to observeT-cell receptor clustering upon activa-tion with antigen.286

Super-Resolution Imaging. Opticalmicroscopy is an unrivaled tool forobtaining structural and molecular in-formation at the micrometer scale.However, the opportunity for insight atthe nanometer scale, where complex andimportant biochemistry remains to be

FIG. 17. Three-color FCCS using QDs. (A) Instrument diagram. (B) Schematic of QD nanobarcode. (C) Diagonal triple cross-correlationfunction decays for QD655 at different concentrations. (D) Count rate trajectories collected for QD525 (green), QD605 (orange), and increasingamounts of QD655 (red): (i) 0 lL, (ii) 6 lL, and (iii) 10 lL. [Reproduced with permission from Ref. 283. Copyright American Chemical Society2012.]


focal point review

elucidated, is limited by the Abbediffraction limit (~k/2 or typically200–300 nm). To address this shortcom-ing, a variety of super-resolution imag-ing techniques have emerged. Onegroup of these techniques, referred toas single molecule localization methods,includes stochastic optical reconstruc-tion microscopy (STORM), photoacti-vated localization microscopy, andsuper-resolution optical fluctuation im-aging (SOFI). These methods rely on theswitching of individual fluorophoresbetween bright and dark states, whichhas a natural parallel with the blinkingbehavior of QDs. Indeed, Lidke et al.used an independent component analysismethod to resolve two closely spaced(23 nm) QD emitters, based on theirblinking, with a standard wide-fieldmicroscope.19 Chien et al. similarly usedthe blinking of QDs to image microtu-bules in fixed CHO (Chinese hamsterovary) cells with 30 nm resolution andan acquisition time that was less than 1

min.287 QDs were also used to developthe SOFI imaging technique that recordsa movie to capture fluorescence inter-mittency (i.e., blinking) over time andstatistically analyzes the signal fluctua-tions.36 Since the blinking of QDsfollows power law statistics and occursover all time scales, arbitrary frame ratescan be used. The advantages of thetechnique include a five-fold improve-ment in resolution and reduced back-ground, as demonstrated by labeling thea-tubulin network of fixed 3T3 fibro-blast cells with QD625-secondary anti-body conjugates (Fig. 18).36 There is anongoing effort to combine QDs andSTORM for in vivo super-resolutionimaging.288,289

Near-Field Scanning Optical Mi-croscopy. Near-field scanning opticalmicroscopy (NSOM) is a scanningprobe imaging technique that providesoptical resolution below the diffractionlimit, with simultaneous topographicinformation. Cai’s group has combined

QDs and NSOM for high-resolution(~50–100 nm) imaging of cell surfacereceptors in several studies.290–292 Forexample, one study focused on imagingantigen-specific T-cell receptor (TCR)response to activation and expansion.290

Vc2Vd2 T-cell surface receptors specif-ically bind phosphoantigens, resulting inT-cell expansion (i.e., rapid proliferationof clonal T cells) as part of an immuneresponse. QD655 were conjugated withanti-TCR antibodies and used to labelthe TCRs on T cells obtained frommacaques. The NSOM-measured distri-bution of QD PL across individual cellswas correlated to the cell surfacedistribution of TCRs. Before activationand expansion, a larger number ofsmaller TCR sites were associated withVc2Vd2 T cells (~5 6 1 3 103,primarily 50 nm in size) and moreuniformly distributed compared with anonengaging ab T-cell phenotype (~26 1 3 103, .90 nm in size). Afteractivation (i.e., infection of the ma-

FIG. 18. 3D SOFI of the a-tubulin network in 3T3 cells labeled with QD625: (A) Time-averaged image from all frames and (B) SOFI image. Thescale bar is 2 lm. [Reproduced with permission from Ref. 36. Copyright the authors 2009.]


caques), the TCRs aggregated on theVc2Vd2 T-cell membrane, and thispattern was sustained in daughter cells.The aggregated receptors recognizedphosphoantigen and imbued theVc2Vd2 T cells with much more potenteffector function, suggesting an impor-tant role for TCR aggregation in im-mune response.290 In these experiments,the QD655 were found to be a muchmore effective label than several com-mon fluorescent dyes, permitting repeat-ed imaging of cells by virtue of theirresistance to both photobleaching andchemical degradation. Building on thisapproach, these authors subsequentlyused two-color NSOM to visualize theclustering and interactions of membrane

receptors during TCR/CD3-mediatedsignaling in T cells.293 Images ofQD605 and QD650 PL were combinedwith topographical information (Fig. 19)and indicated co-clustering of CD3 withCD4 or CD8 co-receptors during T-cellactivation. The authors also noted thatco-stimulation of CD28 receptors sig-nificantly enhanced the clustering ofTCR/CD3 with CD4 co-receptors, butnot CD8 co-receptors. Cai’s group alsoused QD-antibody conjugates andNSOM to investigate the distributionof CD4 receptor proteins on T-helpercell surfaces291 and hyaluronan receptorCD44 on the surface of mesenchymalstem cells.292 In the latter, blinking wassuggested to be useful for identifying

single QDs, and preferential enrichmentof QD-antibody conjugates was ob-served on cell filopodia (a sensoryorganelle important in migration andadhesion).292 The enrichment corre-sponded to clusters of CD44 domains(200–600 nm in size) that were primar-ily observed at the peaks and ridges ofcell surface topography plots. Suchdetail would not have been discernedwithout the dual topographical andoptical capability of the NSOM tech-nique, facilitated by the use of QDs.

Fluorescence Lifetime and Fluores-cence Lifetime Imaging Microscopy.Although QD PL intensity measure-ments are straightforward and versatile,reliable quantitative measurements arevery challenging when the concentrationof QDs is not controlled. Such is almostalways the case when, for example, QDsare delivered to cells, QDs are immobi-lized at an interface, or QDs are subjectto dilution in flow systems. Althoughratiometric PL intensity methods addressthis challenge, ratiometric QD probesare not available for every application.Fluorescence (or PL) lifetime analysis isindependent of QD concentration and isalso very well suited to quantitativemeasurements when concentration isuncertain. In addition to being a measureof FRET and CT quenching efficiency,changes in QD PL properties, such aslifetime, can reflect physicochemicalchanges in the local microenvironment(e.g., temperature, pH) by virtue ofchanges in radiative and non-radiativerates.294,295 Jaque’s group has investi-gated this phenomenon for thermometryapplications and found that optimumtemperature sensitivity can be obtainedwith smaller sized QDs and the use ofCdTe over CdSe.296 Spectral shifts areaccompanied by changes in PL lifetimewhere, for example, CdTe QD515 (1 nmin diameter) exhibited a thermal sensi-tivity of -0.017 8C-1 (measured as therelative change in lifetime per degreeCelsius) between 27 and 50 8C. Thissensitivity was comparable to rhodamineB–doped microspheres (0.0016 8C-1)and Kiton red dye (0.011 8C-1),296

suggesting that QDs might be usefulprobes for FLIM-based measurement ofintracellular temperature. Such measure-ments are important for identifying‘‘hot’’ malignant cells (higher metabolic

FIG. 19. NSOM images of T cell receptors (TCR/CD3) labeled with QD-antibodyconjugates: (A) CD4 T cells co-stimulated with anti-CD3 and anti-CD28 antibodies; and (B)CD4 T cells stimulated with anti-CD28 only. Scale bars are 1 lm. The images on the leftshow topographic data, the images in the right show the merged topographic-QD PL data.PL from QD-labeled CD3 is shown in blue; PL from QD-labeled CD4 is shown in red;colocalization of both PL signals appears violet. The co-stimulation of cells (A) showsincreased co-clustering of CD3-CD4 nanodomains on the membrane. Figure reprinted withpermission under the Creative Commons Attribution License from ref. 293. Copyright 2009the authors.


focal point review

activity compared with healthy cells)and monitoring hyperthermic treatmentof those cells.

FLIM can also be used for visualiza-tion and quantitative sensing with QDprobes based on FRET or CT quench-ing. For example, Pai and Cotlet mod-ified vaterite (CaCO3) microparticleswith CdSe/ZnS QD525-TDTomato (fluo-rescent protein) conjugates and mea-sured the QD-TDTomato FRETefficiency (~67%) via two-colorFLIM.297 Although only a proof-of-concept study, this format could bereadily extended to sensing proteolyticactivity by expressing the fluorescentprotein with a peptide linker that was asubstrate for a protease of interest.125,141

Ruedas-Rama et al. developed a FLIM-based chloride ion (Cl–) sensor based onthe CT quenching interaction betweenCdSe/ZnS QD610 and lucigenin, a chlo-ride-sensitive indicator dye.298 The luci-genin was conjugated to the QDs toprovide the proximity for CT quenching,resulting in a marked decrease in the QDPL intensity and its lifetime (from 20 to9 ns). However, Cl– dynamicallyquenches lucigenin, and the collisionallucigenin–Cl– interaction competed withthe QD–lucigenin CT interaction. In-creasing concentrations of Cl– led to aprogressive recovery of the QD PLlifetime, including a linear response for0.5–50 mM Cl– in a complex solutionmimicking an intracellular matrix.298

FLIM-based sensing of Cl– was demon-strated, but once again only as a proof-of-concept. Nevertheless, as FLIM withfluorescent proteins and organic dyescontinues to grow in importance forcellular imaging and sensing,299–301 theutility of combining QDs with FLIMwill also grow.

SUMMARY AND OUTLOOK

The role for QDs in bioanalysis andbioimaging has grown considerably overthe last decade, and this growth can beexpected to continue. QDs haveemerged as much more than an alterna-tive to fluorescent dyes, providing themeans to address new challenges inbiochemical and biophysical research.As the applications of QDs have ex-panded, there has been a progressiveevolution of QD materials and biocon-jugates, permitting even greater tailoring

of the physical and optical properties ofQDs. This review has summarized theimportant properties of QDs and high-lighted several QD-based methods inbioanalysis and bioimaging. Thesemethods rely on QD PL intensity,polarization, or lifetime for detection,as well as modulation of these propertiesvia Forster-type energy transfer (FRET,BRET, CRET), CL and ECL, CTquenching, and coupling with plasmonicstructures or photonic crystals. Fluorom-eters, plate readers, flow cytometers, andother steady-state or time-resolved spec-troscopic platforms have been used todevelop assays around QDs, whereasimaging methods have included the useof epifluorescence, confocal and 2PEmicroscopy, spectral imaging, singleparticle tracking, FCS, super-resolutionimaging, NSOM, and FLIM. In each ofthe above-mentioned methods, QDsoffer nontrivial advantages: the bright-ness needed for sensitive detection, thephotostability needed for tracking dy-namic processes, the multiplexing capa-bility needed to elucidate complexsystems, or the nanoscale interfaceneeded for biomolecular engineering ofnovel probes and biosensors. AlthoughQD materials have been commercializedfor research and development purposes,one of the challenges moving forwardwill be to translate the fruits of theselabors into commercialized, QD-enabledtechnologies; for example, assay kitsand diagnostic platforms. In the interim,QDs will continue their rise to promi-nence as powerful and versatile tools forbioanalysis and bioimaging.

ACKNOWLEDGMENTS

Eleonora Petryayeva is grateful to the NaturalSciences and Engineering Research Council ofCanada for support through a postgraduate fellow-ship. W. Russ Algar and Eleonora Petryayevaacknowledge the University of British Columbiafor financial support of this research. Igor L.Medintz acknowledges the Office of Naval Re-search, the Naval Research Laboratory (NRL), theNRL Nanosciences Institute, the Defense Ad-vanced Research Projects Agency, and the DefenseThreat Reduction Agency Joint Science andTechnological Office (DTRA-JSTO) Military In-terdepartmental Purchase Requisition B112582M.

1. C.J. Murphy, J.L. Coffer. ‘‘Quantum Dots: APrimer’’. Appl. Spectrosc. 2002. 56(1):16A-27A.

2. M. Bruchez, M. Moronne, P. Gin, S. Weiss,A.P. Alivisatos. ‘‘Semiconductor Nanocrys-

tals as Fluorescent Biological Labels’’. Sci-ence. 1998. 281(5385): 2013-2016.

3. W.C.W. Chan, S.M. Nie. ‘‘Quantum DotBioconjugates for Ultrasensitive NonisotopicDetection’’. Science. 1998. 281(5385):2016-2018.

4. I.L. Medintz, H. Mattoussi. ‘‘Quantum Dot-Based Resonance Energy Transfer and ItsGrowing Application in Biology’’. Phys.Chem. Chem. Phys. 2009. 11(1): 17-45.

5. M.K. Wagner, F. Li, J.J. Li, X.F. Li, X.C. Le.‘‘Use of Quantum Dots in the Developmentof Assays for Cancer Biomarkers’’. Anal.Bioanal. Chem. 2010. 397(8): 3213-3224.

6. H. Mattoussi, G. Palui, H.B. Na. ‘‘Lumines-cent Quantum Dots as Platforms for Probingin Vitro and in Vivo Biological Processes’’.Adv. Drug Delivery Rev. 2012. 64(2):138-166.

7. C. Frigerio, D.S.M. Ribeiro, S.S.M. Ro-drigues, V.L.R.G. Abreu, J.A.C. Barbosa,J.A.V. Prior, K.L. Marques, J.L.M. Santos.‘‘Application of Quantum Dots as AnalyticalTools in Automated Chemical Analysis: AReview’’. Anal. Chim. Acta. 2012. 735: 9-22.

8. M.F. Frasco, N. Chaniotakis. ‘‘BioconjugatedQuantum Dots as Fluorescent Probes forBioanalytical Applications’’. Anal. Bioanal.Chem. 2010. 396(1): 229-240.

9. X. Michalet, F.F. Pinaud, L.A. Bentolila,J.M. Tsay, S. Doose, J.J. Li, G. Sundaresan,A.M. Wu, S.S. Gambhir, S. Weiss. ‘‘Quan-tum Dots for Live Cells, in Vivo Imaging,and Diagnostics’’. Science. 2005. 307(5709):538-544.

10. X.Y. Wu, H.J. Liu, J.Q. Liu, K.N. Haley,J.A. Treadway, J.P. Larson, N.F. Ge, F.Peale, M.P. Bruchez. ‘‘ImmunofluorescentLabeling of Cancer Marker Her2 and OtherCellular Targets with Semiconductor Quan-tum Dots’’. Nat. Biotechnol. 2003. 21(1):41-46.

11. I.L. Medintz, D. Farrell, K. Susumu, S.A.Trammell, J.R. Deschamps, F.M. Brunel,P.E. Dawson, H. Mattoussi. ‘‘MultiplexCharge-Transfer Interactions between Quan-tum Dots and Peptide-Bridged RutheniumComplexes’’. Anal. Chem. 2009. 81(12):4831-4839.

12. D. Geissler, L.J. Charbonniere, R.F. Ziessel,N.G. Butlin, H.G. Lohmannsroben, N. Hil-debrandt. ‘‘Quantum Dot Biosensors forUltrasensitive Multiplexed Diagnostics’’. An-gew. Chem. Int. Ed. 2010. 49(8): 1396-1401.

13. I.L. Medintz, H.T. Uyeda, E.R. Goldman, H.Mattoussi. ‘‘Quantum Dot Bioconjugates forImaging, Labelling and Sensing’’. Nat. Ma-ter. 2005. 4(6): 435-446.

14. U. Resch-Genger, M. Grabolle, S. Cavaliere-Jaricot, R. Nitschke, T. Nann. ‘‘QuantumDots versus Organic Dyes as FluorescentLabels’’. Nat. Methods. 2008. 5(9): 763-775.

15. W.C.W. Chan, D.J. Maxwell, X.H. Gao, R.E.Bailey, M.Y. Han, S.M. Nie. ‘‘LuminescentQuantum Dots for Multiplexed BiologicalDetection and Imaging’’. Curr. Opin. Bio-technol. 2002. 13(1): 40-46.

16. W.R. Algar, A.J. Tavares, U.J. Krull. ‘‘Be-yond Labels: A Review of the Application ofQuantum Dots as Integrated Components ofAssays, Bioprobes, and Biosensors Utilizing


Optical Transduction’’. Anal. Chim. Acta.2010. 673(1): 1-25.

17. S.J. Rosenthal, J.C. Chang, O. Kovtun, J.R.McBride, I.D. Tomlinson. ‘‘BiocompatibleQuantum Dots for Biological Applications’’.Chem. Biol. 2011. 18(1): 10-24.

18. W.R. Algar, K. Susumu, J.B. Delehanty, I.L.Medintz. ‘‘Semiconductor Quantum Dots inBioanalysis: Crossing the Valley of Death’’.Anal. Chem. 2011. 83(23): 8826-8837.

19. K. Lidke, B. Rieger, T. Jovin, R. Heintz-mann. ‘‘Superresolution by Localization ofQuantum Dots Using Blinking Statistics’’.Opt. Express. 2005. 13(18): 7052-7062.

20. D.R. Larson, W.R. Zipfel, R.M. Williams,S.W. Clark, M.P. Bruchez, F.W. Wise, W.W.Webb. ‘‘Water-Soluble Quantum Dots forMultiphoton Fluorescence Imaging in Vivo’’.Science. 2003. 300(5624): 1434-1436.

21. F. Helmchen, W. Denk. ‘‘Deep Tissue Two-Photon Microscopy’’. Nat. Methods. 2005.2(12): 932-940.

22. R.H. Na, I.M. Stender, L.X. Ma, H.C. Wulf.‘‘Autofluorescence Spectrum of Skin: Com-ponent Bands and Body Site Variations’’.Skin Res. Technol. 2000. 6(3): 112-117.

23. A.M. Smith, S. Nie. ‘‘Semiconductor Nano-crystals: Structure, Properties, and Band GapEngineering. Acc’’. Chem. Res. 2009. 43(2):190-200.

24. S. Kim, B. Fisher, H.J. Eisler, M. Bawendi.‘‘Type-II Quantum Dots: CdTe/CdSe(Core/Shell) and CdSe/ZinTe(Core/Shell) Hetero-structures’’. J. Am. Chem. Soc. 2003.125(38): 11466-11467.

25. B. Blackman, D. Battaglia, X. Peng. ‘‘Brightand Water-Soluble Near IR-Emitting CdSe/CdTe/ZnSe Type-II/Type-I Nanocrystals,Tuning the Efficiency and Stability byGrowth’’. Chem. Mater. 2008. 20(15):4847-4853.

26. R. Xie, X. Peng. ‘‘Synthetic Scheme forHigh-Quality InAs Nanocrystals Based onSelf-Focusing and One-Pot Synthesis ofInAs-Based Core–Shell Nanocrystals’’. An-gew. Chem. Int. Ed. 2008. 47(40):7677-7680.

27. R. Xie, X. Peng. ‘‘Synthesis of Cu-DopedInP Nanocrystals (d-dots) with ZnSe Diffu-sion Barrier as Efficient and Color-TunableNIR Emitters.’’. J. Am. Chem. Soc. 2009.131(30): 10645-10651.

28. S.W. Kim, J.P. Zimmer, S. Ohnishi, J.B.Tracy, J.V. Frangioni, M.G. Bawendi. ‘‘En-gineering InAsxP1-x/InP/ZnSe III-V AlloyedCore/Shell Quantum Dots for the Near-Infrared’’. J. Am. Chem. Soc. 2005.127(30): 10526-10532.

29. R.G. Xie, K. Chen, X.Y. Chen, X.G. Peng.‘‘InAs/InP/ZnSe Core/Shell/Shell QuantumDots as Near-Infrared Emitters: Bright,Narrow-Band, Non-Cadmium Containing,and Biocompatible’’. Nano Res. 2008. 1(6):457-464.

30. X. Zhong, R. Xie, Y. Zhang, T. Basche, W.Knoll. ‘‘High-Quality Violet- to Red-Emit-ting ZnSe/CdSe Core/Shell Nanocrystals’’.Chem. Mater. 2005. 17(16): 4038-4042.

31. S. Kim, J. Park, T. Kim, E. Jang, S. Jun, H.Jang, B. Kim, S.-W. Kim. ‘‘Reverse Type-IZnSe/InP/ZnS Core/Shell/Shell Nanocrys-tals: Cadmium-Free Quantum Dots for Vis-

ible Luminescence’’. Small. 2011. 7(1):70-73.

32. A.M. Smith, A.M. Mohs, S. Nie. ‘‘Tuning theOptical and Electronic Properties of Colloi-dal Nanocrystals by Lattice Strain’’. Nat.Nanotechnol. 2009. 4(1): 56-63.

33. C. Galland, Y. Ghosh, A. Steinbruck, M.Sykora, J.A. Hollingsworth, V.I. Klimov, H.Htoon. ‘‘Two Types of Luminescence Blink-ing Revealed by Spectroelectrochemistry ofSingle Quantum Dots’’. Nature. 2011.479(7372): 203-U275.

34. S. Clarke, F. Pinaud, O. Beutel, C.J. You, J.Piehler, M. Dahan. ‘‘Covalent Monofunction-alization of Peptide-Coated Quantum Dotsfor Single-Molecule Assays’’. Nano Lett.2010. 10(6): 2147-2154.

35. M.V. Ehrensperger, C. Hanus, C. Vannier,A. Triller, M. Dahan. ‘‘Multiple AssociationStates between Glycine Receptors and Ge-phyrin Identified by SPT Analysis’’. Biophys.J. 2007. 92(10): 3706-3718.

36. T. Dertinger, R. Colyer, G. Iyer, S. Weiss, J.Enderlein. ‘‘Fast, Background-Free, 3D Su-per-Resolution Optical Fluctuation Imaging(SOFI)’’. P. Natl. Acad. S. USA. 2009.106(52): 22287-22292.

37. S.F. Lee, M.A. Osborne. ‘‘Brightening,Blinking, Bluing and Bleaching in the Lifeof a Quantum Dot: Friend or Foe?’’.ChemPhysChem. 2009. 10(13): 2174-2191.

38. K.X. Hay, V.Y. Waisundara, Y. Zong, M.-Y.Han, D. Huang. ‘‘CdSe Nanocrystals asHydroperoxide Scavengers: A New Ap-proach to Highly Sensitive Quantification ofLipid Hydroperoxides’’. Small. 2007. 3(2):290-293.

39. A. Javier, G.F. Strouse. ‘‘Activated andIntermittent Photoluminescence in ThinCdSe Quantum Dot Films’’. Chem. Phys.Lett. 2004. 391(1-3): 60-63.

40. J.U. Sutter, D.J.S. Birch, O.J. Rolinski. ‘‘TheEffect of Intensity of Excitation on CdSe/ZnSQuantum Dots: Opportunities in Lumines-cence Sensing’’. Appl. Phys. Lett. 2011.98(2): 021108.

41. J. Yao, D.R. Larson, H.D. Vishwasrao, W.R.Zipfel, W.W. Webb. ‘‘Blinking and Non-radiant Dark Fraction of Water-SolubleQuantum Dots in Aqueous Solution’’. P.Natl. Acad. S. USA. 2005. 102(40):14284-14289.

42. T. Pons, I.L. Medintz, D. Farrell, X. Wang,A.F. Grimes, D.S. English, L. Berti, H.Mattoussi. ‘‘Single-Molecule ColocalizationStudies Shed Light on the Idea of FullyEmitting versus Dark Single Quantum Dots’’.Small. 2011. 7(14): 2101-2108.

43. N. Durisic, P.W. Wiseman, P. Grutter, C.D.Heyes. ‘‘A Common Mechanism Underliesthe Dark Fraction Formation and Fluores-cence Blinking of Quantum Dots’’. ACSNano. 2009. 3(5): 1167-1175.

44. I.L. Medintz, A.R. Clapp, H. Mattoussi, E.R.Goldman, B. Fisher, J.M. Mauro. ‘‘Self-Assembled Nanoscale Biosensors Based onQuantum Dot FRET Donors’’. Nat. Mater.2003. 2(9): 630-638.

45. M.A. Hines, P. Guyot-Sionnest. ‘‘Synthesisand Characterization of Strongly Luminesc-ing ZnS-Capped CdSe Nanocrystals’’. J.Phys. Chem. 1996. 100(2): 468-471.

46. B.O. Dabbousi, J. RodriguezViejo, F.V.Mikulec, J.R. Heine, H. Mattoussi, R. Ober,K.F. Jensen, M.G. Bawendi. ‘‘(CdSe)ZnSCore-Shell Quantum Dots: Synthesis andCharacterization of a Size Series of HighlyLuminescent Nanocrystallites’’. J. Phys.Chem. B. 1997. 101(46): 9463-9475.

47. X.Y. Wang, X.F. Ren, K. Kahen, M.A.Hahn, M. Rajeswaran, S. Maccagnano-Zach-er, J. Silcox, G.E. Cragg, A.L. Efros, T.D.Krauss. ‘‘Non-Blinking SemiconductorNanocrystals’’. Nature. 2009. 459(7247):686-689.

48. J. McBride, J. Treadway, L.C. Feldman, S.J.Pennycook, S.J. Rosenthal. ‘‘Structural Basisfor Near Unity Quantum Yield Core/ShellNanostructures’’. Nano Lett. 2006. 6(7):1496-1501.

49. D.V. Talapin, I. Mekis, S. Gotzinger, A.Kornowski, O. Benson, H. Weller. ‘‘CdSe/CdS/ZnS and CdSe/ZnSe/ZnS Core-Shell-Shell Nanocrystals’’. J. Phys. Chem. B. 2004.108(49): 18826-18831.

50. R.G. Xie, U. Kolb, J.X. Li, T. Basche, A.Mews. ‘‘Synthesis and Characterization ofHighly Luminescent CdSe-Core CdS/Zn0.5Cd0.5S/ZnS Multishell Nanocrystals’’.J. Am. Chem. Soc. 2005. 127(20):7480-7488.

51. M.D. Regulacio, M.Y. Han. ‘‘Composition-Tunable Alloyed Semiconductor Nanocrys-tals. Acc’’. Chem. Res. 2010. 43(5): 621-630.

52. N. Al-Salim, A.G. Young, R.D. Tilley, A.J.McQuillan, J. Xia. ‘‘Synthesis of CdSeSNanocrystals in Coordinating and Noncoor-dinating Solvents: Solvent’s Role in Evolu-tion of the Optical and Structural Properties’’.Chem. Mater. 2007. 19(21): 5185-5193.

53. L.A. Swafford, L.A. Weigand, M.J. Bowers,J.R. McBride, J.L. Rapaport, T.L. Watt, S.K.Dixit, L.C. Feldman, S.J. Rosenthal. ‘‘Ho-mogeneously Alloyed CdSxSe1-x Nanocrys-tals: Synthesis, Characterization, andComposition/Size-Dependent Band Gap’’. J.A m . C h e m . S o c . 2 0 0 6 . 1 2 8 ( 3 7 ) :12299-12306.

54. R.E. Bailey, S. Nie. ‘‘Alloyed SemiconductorQuantum Dots: Tuning the Optical Proper-ties Without Changing the Particle Size’’. J.Am. Chem. Soc. 2003. 125(23): 7100-7106.

55. X. Zhong, Y. Feng, W. Knoll, M. Han.‘‘Alloyed ZnxCd1-xS Nanocrystals withHighly Narrow Luminescence SpectralWidth’’. J. Am. Chem. Soc. 2003. 125(44):13559-13563.

56. X. Zhong, M. Han, Z. Dong, T.J. White, W.Knoll. ‘‘Composition-Tunable ZnxCd1-xSeNanocrystals with High Luminescence andStability’’. J. Am. Chem. Soc. 2003. 125(28):8589-8594.

57. O.I. Micic, S.P. Ahrenkiel, A.J. Nozik.‘‘Synthesis of Extremely Small InP QuantumDots and Electronic Coupling in TheirDisordered Solid Films’’. Appl. Phys. Lett.2001. 78(25): 4022-4024.

58. D. Battaglia, X. Peng. ‘‘Formation of HighQuality InP and InAs Nanocrystals in aNoncoordinating Solvent’’. Nano Lett. 2002.2(9): 1027-1030.

59. A. Shiohara, S. Prabakar, A. Faramus, C.-Y.Hsu, P.-S. Lai, P.T. Northcote, R.D. Tilley.‘‘Sized Controlled Synthesis, Purification,


focal point review

and Cell Studies with Silicon QuantumDots’’. Nanoscale. 2011. 3(8): 3364-3370.

60. F. Erogbogbo, K.T. Yong, I. Roy, R. Hu,W.C. Law, W.W. Zhao, H. Ding, F. Wu, R.Kumar, M.T. Swihart, P.N. Prasad. ‘‘In VivoTargeted Cancer Imaging, Sentinel LymphNode Mapping and Multi-Channel Imagingwith Biocompatible Silicon Nanocrystals’’.ACS Nano. 2011. 5(1): 413-423.

61. L.Y.T. Chou, W.C.W. Chan. ‘‘Nanotoxicol-ogy No Signs of Illness’’. Nat. Nanotechnol.2012. 7(7): 416-417.

62. S. Ghaderi, B. Ramesh, A.M. Seifalian.‘‘Fluorescence Nanoparticles ‘‘Quantum Dotsas Drug Delivery System and Their Toxicity:A Review’’. J. Drug Targeting. 2011. 19(7):475-486.

63. F.M. Winnik, D. Maysinger. ‘‘Quantum DotCytotoxicity and Ways To Reduce It’’. Acc.Chem. Res. 2012. DOI: 10.1021/ar3000585.

64. S. Xu, J. Ziegler, T. Nann. ‘‘Rapid Synthesisof Highly Luminescent InP and InP/ZnSNanocrystals’’. J. Mater. Chem. 2008.18(23): 2653-2656.

65. T. Greco, C. Ippen, A. Wedel. ‘‘InP/ZnSe/ZnS Core-Multishell Quantum Dots forImproved Luminescence Efficiency’’. Proc.SPIE. 2012. 8424: 842439.

66. H.S. Choi, W. Liu, P. Misra, E. Tanaka, J.P.Zimmer, B.I. Ipe, M.G. Bawendi, J.V.Frangioni. ‘‘Renal Clearance of QuantumDots’’. Nat. Biotechnol. 2007. 25(10):1165-1170.

67. J. Park, C. Dvoracek, K.H. Lee, J.F.Galloway, H.-e. C. Bhang, M.G. Pomper,P.C. Searson. ‘‘CuInSe/ZnS Core/Shell NIRQuantum Dots for Biomedical Imaging’’.Small. 2011. 7(22): 3148-3152.

68. Z.A. Peng, X.G. Peng. ‘‘Formation of High-Quality CdTe, CdSe, and CdS NanocrystalsUsing CdO as Precursor’’. J. Am. Chem. Soc.2001. 123(1): 183-184.

69. L.H. Qu, Z.A. Peng, X.G. Peng. ‘‘AlternativeRoutes Toward High Quality CdSe Nano-crystals’’. Nano Lett. 2001. 1(6): 333-337.

70. W.W. Yu, X.G. Peng. ‘‘Formation of High-Quality CdS and UII-VI SemiconductorNanocrystals in Noncoordinating Solvents:Tunable Reactivity of Monomers’’. Angew.Chem. Int. Ed. 2002. 41(13): 2368-2371.

71. W.J. Zhang, G.J. Chen, J. Wang, B.C. Ye,X.H. Zhong. ‘‘Design and Synthesis ofHighly Luminescent Near-Infrared-EmittingWater-Soluble CdTe/CdSe/ZnS Core/Shell/Shell Quantum Dots’’. Inorg. Chem. 2009.48(20): 9723-9731.

72. Z.T. Deng, O. Schulz, S. Lin, B.Q. Ding,X.W. Liu, X.X. Wei, R. Ros, H. Yan, Y. Liu.‘‘Aqueous Synthesis of Zinc Blende CdTe/CdS Magic-Core/Thick-Shell Tetrahedral-Shaped Nanocrystals with Emission Tunableto Near-Infrared’’. J. Am. Chem. Soc. 2010.132(16): 5592.

73. Z.M. Yuan, Q. Ma, A.Y. Zhang, Y.Q. Cao, J.Yang, P. Yang. ‘‘Synthesis of Highly Lumi-nescent CdTe/ZnO Core/Shell Quantum Dotsin Aqueous Solution’’. J. Mater. Sci. 2012.47(8): 3770-3776.

74. W.J. Parak, T. Pellegrino, C. Plank. ‘‘Label-ling of Cells with Quantum Dots’’. Nano-technology. 2005. 16(2): R9-R25.

75. W.R. Algar, D.E. Prasuhn, M.H. Stewart,

T.L. Jennings, J.B. Blanco-Canosa, P.E.Dawson, I.L. Medintz. ‘‘The ControlledDisplay of Biomolecules on Nanoparticles:A Challenge Suited to Bioorthogonal Chem-istry’’. Bioconjugate Chem. 2011. 22(5):825-858.

76. K.E. Sapsford, W.R. Algar, L. Berti, K.B.Gemmill, B. Casey, E. Oh, M.H. Stewart,I.L. Medintz. ‘‘Functionalizing Nanoparticleswith Biological Molecules: DevelopingChemistries that Facilitate Nanotechnology’’.Chem. Rev., paper in press, 2012.

77. K.E. Sapsford, K.M. Tyner, B.J. Dair, J.R.Deschamps, I.L. Medintz. ‘‘Analyzing Nano-material Bioconjugates: A Review of Currentand Emerging Purification and Characteriza-tion Techniques’’. Anal. Chem. 2011. 83(12):4453-4488.

78. I. Medintz. ‘‘Universal Tools for Biomolec-ular Attachment to Surfaces’’. Nat. Mater.2006. 5(11): 842-842.

79. P.A.S. Jorge, M.A. Martins, T. Trindade, J.L.Santos, F. Farahi. ‘‘Optical Fiber SensingUsing Quantum Dots’’. Sensors. 2007. 7(12):3489-3534.

80. K. Susumu, H.T. Uyeda, I.L. Medintz, T.Pons, J.B. Delehanty, H. Mattoussi. ‘‘En-hancing the Stability and Biological Func-tionalities of Quantum Dots via CompactMultifunctional Ligands’’. J. Am. Chem.Soc. 2007. 129(45): 13987-13996.

81. K. Susumu, B.C. Mei, H. Mattoussi. ‘‘Mul-tifunctional Ligands based on DihydrolipoicAcid and Polyethylene Glycol to PromoteBiocompatibility of Quantum Dots’’. Nat.Protoc. 2009. 4(3): 424-436.

82. B.C. Mei, K. Susumu, I.L. Medintz, J.B.Delehanty, T.J. Mountziaris, H. Mattoussi.‘‘Modular Poly(ethylene glycol) Ligands forBiocompatible Semiconductor and GoldNanocrystals with Extended pH and IonicStability’’. J. Mater. Chem. 2008. 18(41):4949-4958.

83. W. Liu, M. Howarth, A.B. Greytak, Y.Zheng, D.G. Nocera, A.Y. Ting, M.G.Bawendi. ‘‘Compact Biocompatible Quan-tum Dots Functionalized for Cellular Imag-ing’’. J. Am. Chem. Soc. 2008. 130(4):1274-1284.

84. E. Muro, T. Pons, N. Lequeux, A. Fragola,N. Sanson, Z. Lenkei, B. Dubertret. ‘‘Smalland Stable Sulfobetaine Zwitterionic Quan-tum Dots for Functional Live-Cell Imaging’’.J. Am. Chem. Soc. 2010. 132(13): 4556.

85. K. Susumu, E. Oh, J.B. Delehanty, J.B.Blanco-Canosa, B.J. Johnson, V. Jain, W.J.Hervey, W.R. Algar, K. Boeneman, P.E.Dawson, I.L. Medintz. ‘‘MultifunctionalCompact Zwitterionic Ligands for PreparingRobust Biocompatible Semiconductor Quan-tum Dots and Gold Nanoparticles’’. J. Am.Chem. Soc. 2011. 133(24): 9480-9496.

86. Y.J. Zhang, A. Clapp. ‘‘Overview of Stabi-lizing Ligands for Biocompatible QuantumDot Nanocrystals’’. Sensors. 2011. 11(12):11036-11055.

87. D. Liu, P.T. Snee. ‘‘Water-Soluble Semicon-ductor Nanocrystals Cap Exchanged withMetalated Ligands’’. ACS Nano. 2011. 5(1):546-550.

88. Y.J. Zhang, A.M. Schnoes, A.R. Clapp.‘‘Dithiocarbamates as Capping Ligands for

Water-Soluble Quantum Dots’’. ACS Appl.Mater. Interfaces. 2010. 2(11): 3384-3395.

89. D.J. Zhou, Y. Li, E.A.H. Hall, C. Abell, D.Klenerman. ‘‘A Chelating Dendritic LigandCapped Quantum Dot: Preparation, SurfacePassivation, Bioconjugation and SpecificDNA detection’’. Nanoscale. 2011. 3(1):201-211.

90. E.E. Lees, T.L. Nguyen, A.H.A. Clayton, P.Mulvaney, B.W. Muir. ‘‘The Preparation ofColloidally Stable, Water-Soluble, Biocom-patible, Semiconductor Nanocrystals with aSmall Hydrodynamic Diameter’’. ACS Nano.2009. 3(5): 1121-1128.

91. W.W. Yu, E. Chang, J.C. Falkner, J.Y.Zhang, A.M. Al-Somali, C.M. Sayes, J.Johns, R. Drezek, V.L. Colvin. ‘‘FormingBiocompatible and Nonaggregated Nano-crystals in Water Using Amphiphilic Poly-mers’’. J. Am. Chem. Soc. 2007. 129(10):2871-2879.

92. T. Pellegrino, L. Manna, S. Kudera, T. Liedl,D. Koktysh, A.L. Rogach, S. Keller, J.Radler, G. Natile, W.J. Parak. ‘‘HydrophobicNanocrystals Coated with an AmphiphilicPolymer Shell: A General Route to WaterSoluble Nanocrystals’’. Nano Lett. 2004.4(4): 703-707.

93. C. Luccardini, C. Tribet, F. Vial, V. Marchi-Artzner, M. Dahan. ‘‘Size, Charge, andInteractions with Giant Lipid Vesicles ofQuantum Dots Coated with an AmphiphilicMacromolecule’’. Langmuir. 2006. 22(5):2304-2310.

94. D. Janczewski, N. Tomczak, M.Y. Han, G.J.Vancso. ‘‘Synthesis of Functionalized Am-phiphilic Polymers for Coating QuantumDots’’. Nat. Protoc. 2011. 6(10): 1546-1553.

95. I. Yildiz, B. McCaughan, S.F. Cruickshank,J.F. Callan, F.M. Raymo. ‘‘BiocompatibleCdSe-ZnS Core-Shell Quantum Dots Coat-ed with Hydrophilic Polythiols’’. Langmuir.2009. 25(12): 7090-7096.

96. W.H. Liu, A.B. Greytak, J. Lee, C.R. Wong,J. Park, L.F. Marshall, W. Jiang, P.N. Curtin,A.Y. Ting, D.G. Nocera, D. Fukumura, R.K.Jain, M.G. Bawendi. ‘‘Compact Biocompat-ible Quantum Dots via RAFT-MediatedSynthesis of Imidazole-Based Random Co-polymer Ligand’’. J. Am. Chem. Soc. 2010.132(2): 472-483.

97. J.K. Oh. ‘‘Surface Modification of ColloidalCdX-Based Quantum Dots for BiomedicalApplications’’. J. Mater. Chem. 2010. 20(39):8433-8445.

98. T.L. Jennings, S.G. Becker-Catania, R.C.Triulzi, G. Tao, B. Scott, K.E. Sapsford, S.Spindel, E. Oh, V. Jain, J.B. Delehanty, D.E.Prasuhn, K. Boeneman, W.R. Algar, I.L.Medintz. ‘‘Reactive Semiconductor Nano-crystals for Chemoselective Biolabeling andMultiplexed Analysis’’. ACS Nano. 2011.5(7): 5579-5593.

99. D.E. Prasuhn, J.B. Blanco-Canosa, G.J.Vora, J.B. Delehanty, K. Susumu, B.C.Mei, P.E. Dawson, I.L. Medintz. ‘‘Combin-ing Chemoselective Ligation with Polyhisti-dine-Driven Self-Assembly for the ModularDisplay of Biomolecules on Quantum Dots’’.ACS Nano. 2010. 4(1): 267-278.

100. H.S. Han, N.K. Devaraj, J. Lee, S.A.Hilderbrand, R. Weissleder, M.G. Bawendi.


‘‘Development of a Bioorthogonal and High-ly Efficient Conjugation Method for Quan-tum Dots Using Tetrazine-NorborneneCycloaddition’’. J. Am. Chem. Soc. 2010.132(23): 7838.

101. A. Bernardin, A. Cazet, L. Guyon, P.Delannoy, F. Vinet, D. Bonnaffe, I. Texier.‘‘Copper-Free Click Chemistry for HighlyLuminescent Quantum Dot Conjugates: Ap-plication to in Vivo Metabolic Imaging’’.Bioconjugate Chem. 2010. 21(4): 583-588.

102. D.E. Prasuhn, A. Feltz, J.B. Blanco-Canosa,K. Susumu, M.H. Stewart, B.C. Mei, A.V.Yakovlev, C. Loukov, J.M. Mallet, M.Oheim, P.E. Dawson, I.L. Medintz. ‘‘Quan-tum Dot Peptide Biosensors for MonitoringCaspase 3 Proteolysis and Calcium Ions’’.ACS Nano. 2010. 4(9): 5487-5497.

103. K. Boeneman, J.R. Deschamps, S. Buckhout-White, D.E. Prasuhn, J.B. Blanco-Canosa,P.E. Dawson, M.H. Stewart, K. Susumu,E.R. Goldman, M. Ancona, I.L. Medintz.‘‘Quantum Dot DNA Bioconjugates: Attach-ment Chemistry Strongly Influences theResulting Composite Architecture’’. ACSNano. 2010. 4(12): 7253-7266.

104. A.M. Dennis, D.C. Sotto, B.C. Mei, I.L.Medintz, H. Mattoussi, G. Bao. ‘‘SurfaceLigand Effects on Metal-Affinity Coordina-tion to Quantum Dots: Implications forNanoprobe Self-Assembly’’. BioconjugateChem. 2010. 21(7): 1160-1170.

105. W.R. Algar, D. Wegner, A.L. Huston, J.B.Blanco-Canosa, M.H. Stewart, A. Arm-strong, P.E. Dawson, N. Hildebrandt, I.L.Medintz. ‘‘Quantum Dots as SimultaneousAcceptors and Donors in Time-Gated ForsterResonance Energy Transfer Relays: Charac-terization and Biosensing’’. J. Am. Chem.Soc. 2012. 134(3): 1876-1891.

106. S.W. Bae, W.H. Tan, J.I. Hong. ‘‘FluorescentDye-Doped Silica Nanoparticles: New Toolsfor Bioapplications’’. Chem. Commun. 2012.48(17): 2270-2282.

107. V.N. Mochalin, O. Shenderova, D. Ho, Y.Gogotsi. ‘‘The Properties and Applications ofNanodiamonds’’. Nat. Nanotechnol. 2012.7(1): 11-23.

108. K.P. Loh, Q.L. Bao, G. Eda, M. Chhowalla.‘‘Graphene Oxide as a Chemically TunablePlatform for Optical Applications’’. Nat.Chem. 2010. 2(12): 1015-1024.

109. S.N. Baker, G.A. Baker. ‘‘LuminescentCarbon Nanodots: Emergent Nanolights’’.Angew. Chem. Int. Ed. 2010. 49(38):6726-6744.

110. H.C. Wu, X.L. Chang, L. Liu, F. Zhao, Y.L.Zhao. ‘‘Chemistry of Carbon Nanotubes inBiomedical Applications’’. J. Mater. Chem.2010. 20(6): 1036-1052.

111. F. Wang, D. Banerjee, Y.S. Liu, X.Y. Chen,X.G. Liu. ‘‘Upconversion Nanoparticles inBiological Labeling, Imaging, and Therapy’’.Analyst. 2010. 135(8): 1839-1854.

112. Y.C. Shiang, C.C. Huang, W.Y. Chen, P.C.Chen, H.T. Chang. ‘‘Fluorescent Gold andSilver Nanoclusters for the Analysis ofBiopolymers and Cell Imaging’’. J. Mater.Chem. 2012. 22(26): 12972-12982.

113. L. Shang, S.J. Dong, G.U. Nienhaus. ‘‘Ultra-Small Fluorescent Metal Nanoclusters: Syn-

thesis and Biological Applications’’. NanoToday. 2011. 6(4): 401-418.

114. J. Zheng, P.R. Nicovich, R.M. Dickson.‘‘Highly Fluorescent Noble-Metal QuantumDots’’. Annu. Rev. Phys. Chem. 2007. 58:409-431.

115. I. Diez, R.H.A. Ras. ‘‘Fluorescent SilverNanoclusters’’. Nanoscale. 2011. 3(5):1963-1970.

116. Y.Z. Lu, W. Chen. ‘‘Sub-Nanometre SizedMetal Clusters: from Synthetic Challenges tothe Unique Property Discoveries’’. Chem.Soc. Rev. 2012. 41(9): 3594-3623.

117. E.R. Goldman, A.R. Clapp, G.P. Anderson,H.T. Uyeda, J.M. Mauro, I.L. Medintz, H.Mattoussi. ‘‘Multiplexed Toxin AnalysisUsing Four Colors of Quantum Dot Fluorore-agents’’. Anal. Chem. 2004. 76(3): 684-688.

118. F. Morgner, S. Stufler, D. Geissler, I.L.Medintz, W.R. Algar, K. Susumu, M.H.Stewart, J.B. Blanco-Canosa, P.E. Dawson,N. Hildebrandt. ‘‘Terbium to Quantum DotFRET Bioconjugates for Clinical Diagnos-tics: Influence of Human Plasma on Opticaland Assembly Properties’’. Sensors. 2011.11(10): 9667-9684.

119. C.S. Wu, M.K.K. Oo, X.D. Fan. ‘‘HighlySensitive Multiplexed Heavy Metal Detec-tion Using Quantum-Dot-Labeled DNA-zymes’’. ACS Nano. 2010. 4(10): 5897-5904.

120. R. Freeman, Y. Li, R. Tel-Vered, E. Sharon,J. Elbaz, I. Willner. ‘‘Self-Assembly ofSupramolecular Aptamer Structures for Op-tical or Electrochemical Sensing’’. Analyst.2009. 134(4): 653-656.

121. E.R. Goldman, I.L. Medintz, J.L. Whitley, A.Hayhurst, A.R. Clapp, H.T. Uyeda, J.R.Deschamps, M.E. Lassman, H. Mattoussi.‘‘A Hybrid Quantum Dot-Antibody Frag-ment Fluorescence Resonance Energy Trans-fer-Based TNT Sensor’’. J. Am. Chem. Soc.2005. 127(18): 6744-6751.

122. K.E. Sapsford, J. Granek, J.R. Deschamps,K. Boeneman, J.B. Blanco-Canosa, P.E.Dawson, K. Susumu, M.H. Stewart, I.L.Medintz. ‘‘Monitoring Botulinum NeurotoxinA Activity with Peptide-FunctionalizedQuantum Dot Resonance Energy TransferSensors ’’ . ACS Nano. 2011. 5(4):2687-2699.

123. K.R. Lee, I.-J. Kang. ‘‘Effects of DopamineConcentration on Energy Transfer BetweenDendrimer–QD and Dye-Labeled Antibody’’.Ultramicroscopy. 2009. 109(8): 894-898.

124. I.L. Medintz, A.R. Clapp, F.M. Brunel, T.Tiefenbrunn, H.T. Uyeda, E.L. Chang, J.R.Deschamps, P.E. Dawson, H. Mattoussi.‘‘Proteolytic Activity Monitored by Fluores-cence Resonance Energy Transfer ThroughQuantum-Dot-Peptide Conjugates’’. Nat. Ma-ter. 2006. 5(7): 581-589.

125. K. Boeneman, B.C. Mei, A.M. Dennis, G.Bao, J.R. Deschamps, H. Mattoussi, I.L.Medintz. ‘‘Sensing Caspase 3 Activity withQuantum Dot-Fluorescent Protein Assem-blies’’. J. Am. Chem. Soc. 2009. 131(11):3828.

126. S. Huang, Q. Xiao, Z.K. He, Y. Liu, P.Tinnefeld, X.R. Su, X.N. Peng. ‘‘A HighSensitive and Specific QDs FRET Bioprobefor MNase’’. Chem. Commun. 2008. (45):5990-5992.

127. M. Suzuki, Y. Husimi, H. Komatsu, K.Suzuki, K.T. Douglas. ‘‘Quantum Dot FRETBiosensors That Respond to pH, to Proteo-lytic or Nucleolytic Cleavage, to DNASynthesis, or to a Multiplexing Combina-tion’’. J. Am. Chem. Soc. 2008. 130(17):5720-5725.

128. V.J. Bailey, H. Easwaran, Y. Zhang, E.Griffiths, S.A. Belinsky, J.G. Herman, S.B.Baylin, H.E. Carraway, T.H. Wang. ‘‘MS-qFRET: A Quantum Dot-Based Method forAnalysis of DNA Methylation’’. GenomeRes. 2009. 19(8): 1455-1461.

129. W.R. Algar, U.J. Krull. ‘‘Towards Multi-Colour Strategies for the Detection ofOligonucleotide Hybridization Using Quan-tum Dots as Energy Donors in FluorescenceResonance Energy Transfer (FRET)’’. Anal.Chim. Acta. 2007. 581(2): 193-201.

130. M.D. Kattke, E.J. Gao, K.E. Sapsford, L.D.Stephenson, A. Kumar. ‘‘FRET-Based Quan-tum Dot Immunoassay for Rapid and Sensi-tive Detection of Aspergillus amstelodami’’.Sensors. 2011. 11(6): 6396-6410.

131. A.M. Dennis, W.J. Rhee, D. Sotto, S.N.Dublin, G. Bao. ‘‘Quantum Dot-FluorescentProtein FRET Probes for Sensing Intracellu-lar pH’’. ACS Nano. 2012. 6(4): 2917-2924.

132. R.C. Somers, R.M. Lanning, P.T. Snee, A.B.Greytak, R.K. Jain, M.G. Bawendi, D.G.Nocera. ‘‘A Nanocrystal-Based RatiometricpH Sensor for Natural pH Ranges’’. Chem.Sci. 2012. 3(10): 2980-2985.

133. S.B. Lowe, J.A.G. Dick, B.E. Cohen, M.M.Stevens. ‘‘Multiplex Sensing of Protease andKinase Enzyme Activity via OrthogonalCoupling of Quantum Dot–Peptide Conju-gates’’. ACS Nano. 2011. 6(1): 851-857.

134. T.L. Jennings, M.P. Singh, G.F. Strouse.‘‘Fluorescent Lifetime Quenching near d =1.5 nm Gold Nanoparticles: Probing NSETValidity’’. J. Am. Chem. Soc. 2006. 128(16):5462-5467.

135. T. Pons, I.L. Medintz, K.E. Sapsford, S.Higashiya, A.F. Grimes, D.S. English, H.Mattoussi. ‘‘On the Quenching of Semicon-ductor Quantum Dot Photoluminescence byProximal Gold Nanoparticles’’. Nano Lett.2007. 7(10): 3157-3164.

136. W.R. Algar, A.P. Malanoski, K. Susumu,M.H. Stewart, N. Hildebrandt, I.L. Medintz.‘‘Multiplexed Tracking of Protease ActivityUsing a Single Color of Quantum Dot Vectorand a Time-Gated Forster Resonance EnergyTransfer Relay’’. Anal. Chem. 2012. 84(22):10136-10146.

137. G. Marriott, S. Mao, T. Sakata, J. Ran, D.K.Jackson, C. Petchprayoon, T.J. Gomez, E.Warp, O. Tulyathan, H.L. Aaron, E.Y.Isacoff, Y.L. Yan. ‘‘Optical Lock-in Detec-tion Imaging Microscopy for Contrast-En-hanced Imaging in Living Cells’’. P. Natl.Acad. S. USA. 2008. 105(46): 17789-17794.

138. S.A. Diaz, L. Giordano, T.M. Jovin, E.A.Jares-Erijman. ‘‘Modulation of a Photo-switchable Dual-Color Quantum Dot Con-taining a Photochromic FRET Acceptor andan Internal Standard’’. Nano Lett. 2012.12(7): 3537-3544.

139. Z.Y. Xia, J.H. Rao. ‘‘Biosensing and ImagingBased on Bioluminescence Resonance Ener-


focal point review

gy Transfer’’. Curr. Opin. Biotechnol. 2009.20(1): 37-44.

140. M.K. So, C.J. Xu, A.M. Loening, S.S.Gambhir, J.H. Rao. ‘‘Self-Illuminating Quan-tum Dot Conjugates for in Vivo Imaging’’.Nat. Biotechnol. 2006. 24(3): 339-343.

141. Z.Y. Xia, Y. Xing, M.K. So, A.L. Koh, R.Sinclair, J.H. Rao. ‘‘Multiplex Detection ofProtease Activity with Quantum Dot Nano-sensors Prepared by Intein-Mediated SpecificBioconjugation’’. Anal. Chem. 2008. 80(22):8649-8655.

142. X.Q. Liu, R. Freeman, E. Golub, I. Willner.‘‘Chemiluminescence and Chemilumines-cence Resonance Energy Transfer (CRET)Aptamer Sensors Using Catalytic Hemin/G-Quadruplexes’’. ACS Nano. 2011. 5(9):7648-7655.

143. A. Niazov, R. Freeman, J. Girsh, I. Willner.‘‘Following Glucose Oxidase Activity byChemiluminescence and ChemiluminescenceResonance Energy Transfer (CRET) Process-es Involving Enzyme-DNAzyme Conju-gates’’. Sensors. 2011. 11(11): 10388-10397.

144. R. Freeman, X.Q. Liu, I. Winner. ‘‘Chemilu-minescent and Chemiluminescence Reso-nance Energy Transfer (CRET) Detection ofDNA, Metal Ions, and Aptamer-SubstrateComplexes Using Hemin/G-Quadruplexesand CdSe/ZnS Quantum Dots’’. J. Am.Chem. Soc. 2011. 133(30): 11597-11604.

145. E. Golub, A. Niazov, R. Freeman, M.Zatsepin, I. Willner. ‘‘PhotoelectrochemicalBiosensors Without External Irradiation:Probing Enzyme Activities and DNA Sens-ing Using Hemin/G-Quadruplex-StimulatedChemiluminescence Resonance EnergyTransfer (CRET) Generation of Photocur-rents’’. J. Phys. Chem. C. 2012. 116(25):13827-13834.

146. R. Freeman, J. Girsh, A.F.J. Jou, J.A.A. Ho,T. Hug, J. Dernedde, I. Willner. ‘‘OpticalAptasensors for the Analysis of the VascularEndothelial Growth Factor (VEGF)’’. Anal.Chem. 2012. 84(14): 6192-6198.

147. H. Chen, L. Lin, Z. Lin, G.S. Guo, J.M. Lin.‘‘Chemiluminescence Arising from the De-composition of Peroxymonocarbonate andEnhanced by CdTe Quantum Dots’’. J. Phys.Chem. A. 2010. 114(37): 10049-10058.

148. Z.P. Wang, J. Li, B. Liu, J.Q. Hu, X. Yao,J.H. Li. ‘‘Chemiluminescence of CdTe Nano-crystals Induced by Direct Chemical Oxida-tion and Its Size-Dependent and Surfactant-Sensitized Effect’’. J. Phys. Chem. B. 2005.109(49): 23304-23311.

149. X.Z. Li, J. Li, J.L. Tang, J. Kang, Y.H.Zhang. ‘‘Study of Influence of Metal Ions onCdTe/H2O2 Chemiluminescence’’. J. Lumin.2008. 128(7): 1229-1234.

150. Z.P. Wang, J. Li, B. Liu, J.H. Li. ‘‘CdTeNanocrystals Sensitized Chemiluminescenceand the Analytical Application’’. Talanta.2009. 77(3): 1050-1056.

151. H.I. Chen, R.B. Li, L. Lin, G.S. Guo, J.M.Lin. ‘‘Determination of l-Ascorbic Acid inHuman Serum by Chemiluminescence Basedon Hydrogen Peroxide-Sodium HydrogenCarbonate-CdSe/CdS Quantum Dots Sys-tem’’. Talanta. 2010. 81(4-5): 1688-1696.

152. J.P. Lei, H.X. Ju. ‘‘Fundamentals and Bio-analytical Applications of Functional Quan-

tum Dots as Electrogenerated Emitters ofChemiluminescence’’. TrAC, Trends Anal.Chem. 2011. 30(8): 1351-1359.

153. X. Liu, L. Guo, L. Cheng, H. Ju. ‘‘Determi-nation of Nitrite Based on Its QuenchingEffect on Anodic Electrochemiluminescenceof CdSe Quantum Dots’’. Talanta. 2009.78(3): 691-694.

154. L. Zhang, L. Shang, S. Dong. ‘‘Sensitive andSelective Determination of Cu2þ by Electro-chemiluminescence of CdTe QuantumDots’’. Electrochem. Commun. 2008.10(10): 1452-1454.

155. X. Liu, H. Ju. ‘‘Coreactant Enhanced AnodicElectrochemiluminescence of CdTe Quan-tum Dots at Low Potential for SensitiveBiosensing Amplified by Enzymatic Cycle’’.Anal. Chem. 2008. 80(14): 5377-5382.

156. H. Han, Z. Sheng, J. Liang. ‘‘Electrogener-ated Chemiluminescence from Thiol-CappedCdTe Quantum Dots and Its Sensing Appli-cation in Aqueous Solution’’. Anal. Chim.Acta. 2007. 596(1): 73-78.

157. H. Huang, Y. Tan, J. Shi, G. Liang, J.-J. Zhu.‘‘DNA Aptasensor for the Detection of ATPBased on Quantum Dots Electrochemilumi-nescence’’. Nanoscale. 2010. 2(4): 606-612.

158. G.F. Jie, P. Liu, L. Wang, S.S. Zhang.‘‘Electrochemiluminescence ImmunosensorBased on Nanocomposite Film of CdSQuantum Dots-Carbon Nanotubes Combinedwith Gold Nanoparticles-Chitosan’’. Electro-chem. Commun. 2010. 12(1): 22-26.

159. R. Zhang, L. Fan, Y. Fang, S. Yang.‘‘Electrochemical Route to the Preparationof Highly Dispersed Composites of ZnO/Carbon Nanotubes with Significantly En-hanced Electrochemiluminescence fromZnO’’. J. Mater. Chem. 2008. 18(41):4964-4970.

160. T. Wang, S.Y. Zhang, C.J. Mao, J.M. Song,H.L. Niu, B.K. Jin, Y.P. Tian. ‘‘EnhancedElectrochemiluminescence of CdSe QuantumDots Composited with Graphene Oxide andChitosan for Sensitive Sensor. Biosens’’.Bioelectron. 2012. 31(1): 369-375.

161. J. Wang, Y. Shan, W.-W. Zhao, J.-J. Xu, H.-Y. Chen. ‘‘Gold Nanoparticle EnhancedElectrochemiluminescence of CdS ThinFilms for Ultrasensitive Thrombin Detec-tion’’. Anal. Chem. 2011. 83(11): 4004-4011.

162. N. Myung, Z.F. Ding, A.J. Bard. ‘‘Electro-generated Chemiluminescence of CdSeNanocrystals’’. Nano Lett. 2002. 2(11):1315-1319.

163. N. Myung, Y. Bae, A.J. Bard. ‘‘Effect ofSurface Passivation on the ElectrogeneratedChemiluminescence of CdSe/ZnSe Nano-crystals’’. Nano Lett. 2003. 3(8): 1053-1055.

164. S.K. Poznyak, D.V. Talapin, E.V. Shevchen-ko, H. Weller. ‘‘Quantum Dot Chemilumi-nescence’’. Nano Lett. 2004. 4(4): 693-698.

165. Y. Bae, N. Myung, A.J. Bard. ‘‘Electrochem-istry and Electrogenerated Chemilumines-cence of CdTe Nanoparticles’’. Nano Lett.2004. 4(6): 1153-1161.

166. H. Jiang, H. Ju. ‘‘ElectrochemiluminescenceSensors for Scavengers of Hydroxyl RadicalBased on Its Annihilation in CdSe QuantumDots Film/Peroxide System’’. Anal. Chem.2007. 79(17): 6690-6696.

167. H.P. Huang, J.J. Li, J.J. Zhu. ‘‘Electro-

chemiluminescence Based on Quantum Dotsand Their Analytical Application’’. Anal.Methods. 2011. 3(1): 33-42.

168. J. Li, S.J. Guo, E.K. Wang. ‘‘RecentAdvances in New Luminescent Nanomate-rials for Electrochemiluminescence Sen-sors ’’ . RSC Advances. 2012. 2(9):3579-3586.

169. X. Liu, H. Jiang, J. Lei, H. Ju. ‘‘AnodicElectrochemiluminescence of CdTe Quan-tum Dots and Its Energy Transfer forDetection of Catechol Derivatives’’. Anal.Chem. 2007. 79(21): 8055-8060.

170. X. Liu, L. Cheng, J. Lei, H. Ju. ‘‘DopamineDetection Based on Its Quenching Effect onthe Anodic Electrochemiluminescence ofCdSe Quantum Dots’’. Analyst. 2008.133(9): 1161-1163.

171. Z.F. Ding, B.M. Quinn, S.K. Haram, L.E.Pell, B.A. Korgel, A.J. Bard. ‘‘Electrochem-istry and Electrogenerated Chemilumines-cence from Silicon Nanocrystal QuantumDots ’’ . Sc ience . 2002. 296(5571) :1293-1297.

172. C.G. Shi, X. Shan, Z.Q. Pan, J.J. Xu, C. Lu,N. Bao, H.Y. Gu. ‘‘Quantum Dot (QD)-Modified Carbon Tape Electrodes for Repro-ducible Electrochemiluminescence (ECL)Emission on a Paper-Based Platform’’. Anal.Chem. 2012. 84(6): 3033-3038.

173. M.S. Wu, H.W. Shi, L.J. He, J.J. Xu, H.Y.Chen. ‘‘Microchip Device with 64-SiteElectrode Array for Multiplexed Immunoas-say of Cell Surface Antigens Based onElectrochemiluminescence Resonance Ener-gy Transfer’’. Anal. Chem. 2012. 84(9):4207-4213.

174. I.L. Medintz, T. Pons, S.A. Trammell, A.F.Grimes, D.S. English, J.B. Blanco-Canosa,P.E. Dawson, H. Mattoussi. ‘‘InteractionsBetween Redox Complexes and Semicon-ductor Quantum Dots Coupled via a PeptideBridge’’. J. Am. Chem. Soc. 2008. 130(49):16745-16756.

175. M.D. Swain, J. Octain, D.E. Benson. ‘‘Un-imolecular, Soluble Semiconductor Nanopar-ticle-Based Biosensors for Thrombin UsingCharge/Electron Transfer’’. BioconjugateChem. 2008. 19(12): 2520-2526.

176. M. Sykora, M.A. Petruska, J. Alstrum-Acevedo, I. Bezel, T.J. Meyer, V.I. Klimov.‘‘Photoinduced Charge Transfer BetweenCdSe Nanocrystal Quantum Dots and Ru-Polypyridine Complexes’’. J. Am. Chem.Soc. 2006. 128(31): 9984-9985.

177. M. Malicki, K.E. Knowles, E.A. Weiss.‘‘Gating of Hole Transfer from PhotoexcitedPbS Quantum Dots to Aminoferrocene by theLigand Shell of the Dots’’. Chem. Commun.2013. DOI: 10.1039/C2CC32895J.

178. D. Dorokhin, N. Tomczak, A.H. Velders,D.N. Reinhoudt, G.J. Vancso. ‘‘Photolumi-nescence Quenching of CdSe/ZnS QuantumDots by Molecular Ferrocene and FerrocenylThiol Ligands’’. J. Phys. Chem. C. 2009.113(43): 18676-18680.

179. M.J. Ruedas-Rama, E.A.H. Hall. ‘‘A Quan-tum Dot-Lucigenin Probe for Cl’’. Analyst.2008. 133(11): 1556-1566.

180. I. Yildiz, M. Tomasulo, F.M. Raymo. ‘‘AMechanism to Signal Receptor-SubstrateInteractions with Luminescent Quantum


Dots’’. P. Natl. Acad. S. USA. 2006.103(31): 11457-11460.

181. A.J. Morris-Cohen, M.T. Frederick, L.C.Cass, E.A. Weiss. ‘‘Simultaneous Determi-nation of the Adsorption Constant and thePhotoinduced Electron Transfer Rate for aCds Quantum Dot-Viologen Complex’’. J.A m . C h e m . S o c . 2 0 1 1 . 1 3 3 ( 2 6 ) :10146-10154.

182. F. Scholz, L. Dworak, V.V. Matylitsky, J.Wachtveitl. ‘‘Ultrafast Electron Transfer fromPhotoexcited CdSe Quantum Dots to Meth-ylviologen’’. ChemPhysChem. 2011. 12(12):2255-2259.

183. I.L. Medintz, M.H. Stewart, S.A. Trammell,K. Susumu, J.B. Delehanty, B.C. Mei, J.S.Melinger, J.B. Blanco-Canosa, P.E. Dawson,H. Mattoussi. ‘‘Quantum-Dot/Dopamine Bio-conjugates Function as Redox CoupledAssemblies for in Vitro and Intracellular pHSensing’’. Nat. Mater. 2010. 9(8): 676-684.

184. X. Ji, G. Palui, T. Avellini, H.B. Na, C. Yi,K.L. Knappenberger, H. Mattoussi. ‘‘On thepH-Dependent Quenching of Quantum DotPhotoluminescence by Redox Active Dopa-mine’’. J. Am. Chem. Soc. 2012. 134(13):6006-6017.

185. R. Freeman, T. Finder, R. Gill, I. Willner.‘‘Probing Protein Kinase (CK2) and AlkalinePhosphatase with CdSe/ZnS Quantum Dots’’.Nano Lett. 2010. 10(6): 2192-2196.

186. J. Callan, R. Mulrooney, S. Kamila, B.McCaughan. ‘‘Anion Sensing with Lumines-cent Quantum Dots–A Modular ApproachBased on the Photoinduced Electron Transfer(PET) Mechanism’’. J. Fluoresc. 2008. 18(2):527-532.

187. J. Huang, D. Stockwell, Z. Huang, D.L.Mohler, T. Lian. ‘‘Photoinduced UltrafastElectron Transfer from CdSe Quantum Dotsto Re-Bipyridyl Complexes’’. J. Am. Chem.Soc. 2008. 130(17): 5632-5633.

188. S. Ruhle, M. Shalom, A. Zaban. ‘‘Quantum-Dot-Sensitized Solar Cells’’. Chem-PhysChem. 2010. 11(11): 2290-2304.

189. S. Jin, J.-C. Hsiang, H. Zhu, N. Song, R.M.Dickson, T. Lian. ‘‘Correlated Single Quan-tum Dot Blinking and Interfacial ElectronTransfer Dynamics’’. Chem. Sci. 2010. 1(4):519-526.

190. W. Qin, P. Guyot-Sionnest. ‘‘Evidence forthe Role of Holes in Blinking: Negative andOxidized CdSe/CdS Dots’’. ACS Nano.2012. 6(10): 9125-9132.

191. M. Shim, C.J. Wang, P. Guyot-Sionnest.‘‘Charge-Tunable Optical Properties in Col-loidal Semiconductor Nanocrystals’’. J. Phys.Chem. B. 2001. 105(12): 2369-2373.

192. M. Tagliazucchi, D.B. Tice, C.M. Sweeney,A.J. Morris-Cohen, E.A. Weiss. ‘‘Ligand-Controlled Rates of Photoinduced ElectronTransfer in Hybrid CdSe Nanocrystal/Poly(-viologen) Films’’. ACS Nano. 2011. 5(12):9907-9917.

193. A.J. Morris-Cohen, M.D. Peterson, M.T.Frederick, J.M. Kamm, E.A. Weiss. ‘‘Evi-dence for a Through-Space Pathway forElectron Transfer from Quantum Dots toCarboxylate-Functionalized Viologens’’. J.Phys. Chem. Lett. 2012. 3(19): 2840-2844.

194. D. Dorokhin, N. Tomczak, D.N. Reinhoudt,A.H. Velders, G.J. Vancso. ‘‘Ferrocene-

Coated CdSe/ZnS Quantum Dots as Electro-active Nanoparticles Hybrids’’. Nanotechnol-ogy. 2010. 21: 285703.

195. M.G. Sandros, D. Gao, D.E. Benson. ‘‘AModular Nanoparticle-Based System forReagentless Small Molecule Biosensing’’. J.A m . Ch em . S o c . 2 0 0 5 . 1 2 7 ( 3 5 ) :12198-12199.

196. M.G. Sandros, V. Shete, D.E. Benson.‘‘Selective, Reversible, Reagentless MaltoseBiosensing with Core-Shell SemiconductingNanoparticles’’. Analyst. 2006. 131(2):229-235.

197. B.P. Aryal, D.E. Benson. ‘‘Electron DonorSolvent Effects Provide Biosensing withQuantum Dots’’. J. Am. Chem. Soc. 2006.128(50): 15986-15987.

198. V.S. Shete, D.E. Benson. ‘‘Protein DesignProvides Lead(II) Ion Biosensors for ImagingMolecular Fluxes Around Red Blood Cells’’.Biochemistry. 2009. 48(2): 462-470.

199. R. Gill, R. Freeman, J.-P. Xu, I. Willner, S.Winograd, I. Shweky, U. Banin. ‘‘ProbingBioca ta ly t i c Trans format ions wi thCdSe-ZnS QDs’’. J. Am. Chem. Soc.2006. 128(48): 15376-15377.

200. R. Freeman, T. Finder, R. Gill, I. Willner.‘‘Probing Protein Kinase (CK2) and AlkalinePhosphatase with CdSe/ZnS Quantum Dots’’.Nano Lett. 2010. 10(6): 2192-2196.

201. S.C. Cui, T. Tachikawa, M. Fujitsuka, T.Majima. ‘‘Photoinduced Electron Transfer ina Quantum Dot-Cucurbituril SupramolecularComplex’’. J. Phys. Chem. C. 2011. 115(5):1824-1830.

202. C. Burda, T.C. Green, S. Link, M.A. El-Sayed. ‘‘Electron Shuttling Across the Inter-face of CdSe Nanoparticles Monitored byFemtosecond Laser Spectroscopy’’. J. Phys.Chem. B. 1999. 103(11): 1783-1788.

203. J. Tian, L. Zhou, Y. Zhao, Y. Wang, Y.Peng, S. Zhao. ‘‘Multiplexed Detection ofTumor Markers with Multicolor QuantumDots based on Fluorescence PolarizationImmunoassay’’. Talanta. 2012. 92(0): 72-77.

204. S.A. Empedocles, R. Neuhauser, M.G. Ba-wendi. ‘‘Three-Dimensional OrientationMeasurements of Symmetric Single Chro-mophores Using Polarization Microscopy’’.Nature. 1999. 399(6732): 126-130.

205. I.H. Chung, K.T. Shimizu, M.G. Bawendi.‘‘Room Temperature Measurements of the3D Orientation of Single CdSe QuantumDots Using Polarization Microscopy’’. P.Natl. Acad. S. USA. 2003. 100(2): 405-408.

206. P.K. Chattopadhyay, D.A. Price, T.F. Harp-er, M.R. Betts, J. Yu, E. Gostick, S.P.Perfetto, P. Goepfert, R.A. Koup, S.C. DeRosa, M.P. Bruchez, M. Roederer. ‘‘Quan-tum Dot Semiconductor Nanocrystals forImmunophenotyping by Polychromatic FlowCytometry’’. Nat. Med. 2006. 12(8):972-977.

207. O. Kovtun, E.J. Ross, I.D. Tomlinson, S.J.Rosenthal. ‘‘A Flow Cytometry-Based Do-pamine Transporter Binding Assay UsingAntagonist-Conjugated Quantum Dots’’.Chem. Commun. 2012. 48(44): 5428-5430.

208. P.O. Krutzik, G.P. Nolan. ‘‘Fluorescent CellBarcoding in Flow Cytometry Allows High-Throughput Drug Screening and Signaling

Profiling’’. Nat. Methods. 2006. 3(5):361-368.

209. Luminex Corporation. ‘‘Luminex’’. 2012.www.luminex.com [accessed Nov 12 2012].

210. M.Y. Han, X.H. Gao, J.Z. Su, S. Nie.‘‘Quantum-Dot-Tagged Microbeads for Mul-tiplexed Optical Coding of Biomolecules’’.Nat. Biotechnol. 2001. 19(7): 631-635.

211. A. Agrawal, C.Y. Zhang, T. Byassee, R.A.Tripp, S.M. Nie. ‘‘Counting Single NativeBiomolecules and Intact Viruses with Color-Coded Nanoparticles’’. Anal. Chem. 2006.78(4): 1061-1070.

212. K.C. Goss, G.G. Messier, M.E. Potter. ‘‘DataDetection Algorithms for Multiplexed Quan-tum Dot Encoding’’. Opt. Express. 2012.20(5): 5762-5774.

213. Y.L. Gao, W.L. Stanford, W.C.W. Chan.‘‘Quantum-Dot-Encoded Microbeads forMultiplexed Genetic Detection of Non-Amplified DNA Samples’’. Small. 2011.7(1): 137-146.

214. S. Giri, E.A. Sykes, T.L. Jennings, W.C.W.Chan. ‘‘Rapid Screening of Genetic Bio-markers of Infectious Agents Using QuantumDot Barcodes’’. ACS Nano. 2011. 5(3):1580-1587.

215. H.X. Xu, M.Y. Sha, E.Y. Wong, J. Uphoff,Y.H. Xu, J.A. Treadway, A. Truong, E.O’Brien, S. Asquith, M. Stubbins, N.K.Spurr, E.H. Lai, W. Mahoney. ‘‘MultiplexedSNP Genotyping Using the Qbead (TM)System: A Quantum Dot-Encoded Micro-sphere-Based Assay’’. Nucleic Acids Res.2003. 31(8): e43.

216. J.D. Joannopoulos, S.G. Johnson, J.N. Winn,R.D. Meade. Photonic Crystals: Molding theFlow of Light. Princeton, NJ: PrincetonUniversity Press, 2008. 2nd ed.

217. B.T. Cunningham, R.C. Zangar. ‘‘PhotonicCrystal Enhanced Fluorescence for EarlyBreast Cancer Biomarker Detection’’. J.Biophotonics. 2012. 5(8-9): 617-628.

218. N. Ganesh, W. Zhang, P.C. Mathias, E.Chow, J.A.N.T. Soares, V. Malyarchuk,A.D. Smith, B.T. Cunningham. ‘‘EnhancedFluorescence Emission from Quantum Dotson a Photonic Crystal Surface’’. Nat. Nano-technol. 2007. 2(8): 515-520.

219. K.E. Sapsford, S. Spindel, T. Jennings, G.L.Tao, R.C. Triulzi, W.R. Algar, I.L. Medintz.‘‘Optimizing Two-Color SemiconductorNanocrystal Immunoassays in Single WellMicrotiter Plate Formats’’. Sensors. 2011.11(8): 7879-7891.

220. B.T. Cunningham, P. Li, S. Schulz, B. Lin,C. Baird, J. Gerstenmaier, C. Genick, F.Wang, E. Fine, L. Laing. ‘‘Label-Free Assayson the BIND System’’. J. Biomol. Screen.2004. 9(6): 481-490.

221. S.H. Cao, W.P. Cai, Q. Liu, Y.Q. Li.‘‘Surface Plasmon-Coupled Emission: WhatCan Directional Fluorescence Bring to theAnalytical Sciences?’’. Annu. Rev. Anal.Chem. 2012. 5: 317-336.

222. J.R. Lakowicz, J. Malicka, I. Gryczynski, Z.Gryczynski, C.D. Geddes. ‘‘Radiative DecayEngineering: The Role of Photonic ModeDensity in Biotechnology’’. J. Phys. D: Appl.Phys. 2003. 36(14): R240-R249.

223. C.D. Geddes, J.R. Lakowicz. ‘‘Editorial:


focal point review

Metal-Enhanced Fluorescence’’. J. Fluoresc.2002. 12(2): 121-129.

224. O. Kulakovich, N. Strekal, A. Yaroshevich,S. Maskevich, S. Gaponenko, I. Nabiev, U.Woggon, M. Artemyev. ‘‘Enhanced Lumi-nescence of CdSe Quantum Dots on GoldColloids’’. Nano Lett. 2002. 2(12):1449-1452.

225. Y.-H. Chan, J. Chen, S.E. Wark, S.L. Skiles,D.H. Son, J.D. Batteas. ‘‘Using PatternedArrays of Metal Nanoparticles to ProbePlasmon Enhanced Luminescence of CdSeQuantum Dots’’. ACS Nano. 2009. 3(7):1735-1744.

226. J.H. Song, T. Atay, S.F. Shi, H. Urabe, A.V.Nurmikko. ‘‘Large Enhancement of Fluores-cence Efficiency from CdSe/ZnS QuantumDots Induced by Resonant Coupling toSpatially Controlled Surface Plasmons’’.Nano Lett. 2005. 5(8): 1557-1561.

227. P.P. Pompa, L. Martiradonna, A. Della Torre,L. Carbone, L.L. del Mercato, L. Manna, M.De Vittorio, F. Calabi, R. Cingolani, R.Rinaldi. ‘‘Fluorescence Enhancement in Col-loidal Semiconductor Nanocrystals by Me-tallic Nanopatterns. Sensors. Actuat. B-Chem. 2007. 126(1): 187-192.

228. K. Leong, Y. Chen, D.J. Masiello, M.T. Zin,M. Hnilova, H. Ma, C. Tamerler, M.Sarikaya, D.S. Ginger, A.K.-Y. Jen. ‘‘Coop-erative Near-Field Surface Plasmon En-hanced Quantum Dot Nanoarrays’’. Adv.Funct. Mater. 2010. 20(16): 2675-2682.

229. L. Zhou, F. Ding, H. Chen, W. Ding, W.Zhang, S.Y. Chou. ‘‘Enhancement of Immu-noassay’s Fluorescence and Detection Sensi-tivity Using Three-Dimensional PlasmonicNano-Antenna-Dots Array’’. Anal. Chem.2012. 84(10): 4489-4495.

230. R. Robelek, L. Niu, E.L. Schmid, W. Knoll.‘‘Multiplexed Hybridization Detection ofQuantum Dot-Conjugated DNA SequencesUsing Surface Plasmon Enhanced Fluores-cence Microscopy and Spectrometry’’. Anal.Chem. 2004. 76(20): 6160-6165.

231. L.-H. Jin, S.-M. Li, Y.-H. Cho. ‘‘EnhancedDetection Sensitivity of Pegylated CdSe/ZnSQuantum Dots-Based Prostate Cancer Bio-markers by Surface Plasmon-Coupled Emis-sion. Biosens’’. Bioelectron. 2012. 33(1):284-287.

232. L. Malic, M.G. Sandros, M. Tabrizian.‘‘Designed Biointerface Using Near-InfraredQuantum Dots for Ultrasensitive SurfacePlasmon Resonance Imaging Biosensors’’.Anal. Chem. 2011. 83(13): 5222-5229.

233. H. Chen, J. Xue, Y. Zhang, X. Zhu, J. Gao,B. Yu. ‘‘Comparison of Quantum DotsImmunofluorescence Histochemistry andConventional Immunohistochemistry for theDetection of Caveolin-1 and PCNA in theLung Cancer Tissue Microarray’’. J. Mol.Histol. 2009. 40(4): 261-268.

234. C. Chen, J. Peng, H.S. Xia, G.F. Yang, Q.S.Wu, L.D. Chen, L.B. Zeng, Z.L. Zhang,D.W. Pang, Y. Li. ‘‘Quantum Dots-BasedImmunofluorescence Technology for theQuantitative Determination of HER2 Expres-sion in Breast Cancer’’. Biomaterials. 2009.30(15): 2912-2918.

235. Y. Ruan, W.M. Yu, F. Cheng, X.B. Zhang,S. Larre. ‘‘Detection of Prostate Stem Cell

Antigen Expression in Human ProstateCancer Using Quantum-Dot-Based Technol-ogy’’. Sensors. 2012. 12(5): 5461-5470.

236. Y. Xing, Q. Chaudry, C. Shen, K.Y. Kong,H.E. Zhau, L. WChung, J.A. Petros, R.M.O’Regan, M.V. Yezhelyev, J.W. Simons,M.D. Wang, S. Nie. ‘‘Bioconjugated Quan-tum Dots for Multiplexed and QuantitativeImmunohistochemistry’’. Nat. Protoc. 2007.2(5): 1152-1165.

237. J.K. Jaiswal, H. Mattoussi, J.M. Mauro, S.M.Simon. ‘‘Long-Term Multiple Color Imagingof Live Cells using Quantum Dot Bioconju-gates’’. Nat. Biotechnol. 2003. 21(1): 47-51.

238. J.B. Delehanty, C.E. Bradburne, K. Susumu,K. Boeneman, B.C. Mei, D. Farrell, J.B.Blanco-Canosa, P.E. Dawson, H. Mattoussi,I.L. Medintz. ‘‘Spatiotemporal MulticolorLabeling of Individual Cells Using Peptide-Functionalized Quantum Dots and MixedDelivery Techniques’’. J. Am. Chem. Soc.2011. 133(27): 10482-10489.

239. A. Matsuno, J. Itoh, S. Takekoshi, T.Nagashima, R.Y. Osamura. ‘‘Three-Dimen-sional Imaging of the Intracellular Localiza-tion of Growth Hormone and Prolactin andTheir mRNA Using Nanocrystal (Quantumdot) and Confocal Laser Scanning Micros-copy Techniques’’. J. Histochem. Cytochem.2005. 53(7): 833-838.

240. P.M. Chan, T. Yuen, F. Ruf, J. Gonzalez-Maeso, S.C. Sealfon. ‘‘Method for MultiplexCellular Detection of mRNAs Using Quan-tum Dot Fluorescent in Situ Hybridization’’.Nucleic Acids Res. 2005. 33(18): e161.

241. J. Lee, Y.-J. Kwon, Y. Choi, H.C. Kim, K.Kim, J. Kim, S. Park, R. Song. ‘‘QuantumDot-Based Screening System for Discoveryof G Protein-Coupled Receptor Agonists’’.ChemBioChem. 2012. 13(10): 1503-1508.

242. C. Chen, S.R. Sun, Y.P. Gong, C.B. Qi,C.W. Peng, X.Q. Yang, S.P. Liu, J. Peng, S.Zhu, M.B. Hu, D.W. Pang, Y. Li. ‘‘QuantumDots-Based Molecular Classification ofBreast Cancer by Quantitative Spectroanal-ysis of Hormone Receptors and HER2’’.Biomaterials. 2011. 32(30): 7592-7599.

243. X.-Q. Yang, C. Chen, C.-W. Peng, J.-X.Hou, S.-P. Liu, C.-B. Qi, Y.-P. Gong, X.-B.Zhu, D.-W. Pang, Y. Li. ‘‘Quantum Dot-Based Quantitative Immunofluorescence De-tection and Spectrum Analysis of EpidermalGrowth Factor Receptor in Breast CancerTissue Arrays’’. Int. J. Nanomedicine. 2011.6: 2265-2273.

244. J.A. Liu, S.K. Lau, V.A. Varma, B.A.Kairdolf, S.M. Nie. ‘‘Multiplexed Detectionand Characterization of Rare Tumor Cells inHodgkin’s Lymphoma with MulticolorQuantum Dots’’. Anal. Chem. 2010. 82(14):6237-6243.

245. J. Liu, S.K. Lau, V.A. Varma, R.A. Moffitt,M. Caldwell, T. Liu, A.N. Young, J.A.Petros, A.O. Osunkoya, T. Krogstad, B.Leyland-Jones, M.D. Wang, S.M. Nie. ‘‘Mo-lecular Mapping of Tumor Heterogeneity onClinical Tissue Specimens with MultiplexedQuantum Dots’’. ACS Nano. 2010. 4(5):2755-2765.

246. Y.P. Varshni. ‘‘Temperature Dependence ofthe Energy Gap in Semiconductors’’. Phys-ica. 1967. 34(1): 149-154.

247. J.-M. Yang, H. Yang, L. Lin. ‘‘Quantum DotNano Thermometers Reveal HeterogeneousLocal Thermogenesis in Living Cells’’. ACSNano. 2011. 5(6): 5067-5071.

248. A. Diaspro, G. Chirico, M. Collini. ‘‘Two-Photon Fluorescence Excitation and RelatedTechniques in Biological Microscopy’’. Q.Rev. Biophys. 2005. 38(2): 97-166.

249. G.S. He, L.S. Tan, Q. Zheng, P.N. Prasad.‘‘Multiphoton Absorbing Materials: Molecu-lar Designs, Characterizations, and Applica-t ions ’’ . Chem. Rev. 2008. 108(4):1245-1330.

250. T. Wang, J.Y. Chen, S. Zhen, P.N. Wang,C.C. Wang, W.L. Yang, Q. Peng. ‘‘Thiol-Capped CdTe Quantum Dots with Two-Photon Excitation for Imaging High Auto-fluorescence Background Living Cells’’. J.Fluoresc. 2009. 19(4): 615-621.

251. D.J. Bharali, D.W. Lucey, H. Jayakumar,H.E. Pudavar, P.N. Prasad. ‘‘Folate-Recep-tor-Mediated Delivery of InP Quantum Dotsfor Bioimaging Using Confocal and Two-Photon Microscopy’’. J. Am. Chem. Soc.2005. 127(32): 11364-11371.

252. M. Geszke, M. Murias, L. Balan, G.Medjahdi, J. Korczynski, M. Moritz, J.Lulek, R. Schneider. ‘‘Folic Acid-Conjugat-ed Core/Shell ZnS: Mn/ZnS Quantum Dotsas Targeted Probes for Two Photon Fluores-cence Imaging of Cancer Cells’’. ActaBiomater. 2011. 7(3): 1327-1338.

253. C. Tu, X. Ma, P. Pantazis, S.M. Kauzlarich,A.Y. Louie. ‘‘Paramagnetic, Silicon Quan-tum Dots for Magnetic Resonance and Two-Photon Imaging of Macrophages’’. J. Am.Chem. Soc. 2010. 132(6): 2016-2023.

254. R. Zhang, E. Rothenberg, G. Fruhwirth, P.D.Simonson, F. Ye, I. Golding, T. Ng, W.Lopes, P.R. Selvin. ‘‘Two-Photon 3D FIO-NA of Individual Quantum Dots in anAqueous Environment’’. Nano Lett. 2011.11(10): 4074-4078.

255. M. Stroh, J.P. Zimmer, D.G. Duda, T.S.Levchenko, K.S. Cohen, E.B. Brown, D.T.Scadden, V.P. Torchilin, M.G. Bawendi, D.Fukumura, R.K. Jain. ‘‘Quantum Dots Spec-trally Distinguish Multiple Species within theTumor Milieu in Vivo’’. Nat. Med. 2005.11(6): 678-682.

256. L.M. Maestro, J.E. Ramirez-Hernandez, N.Bogdan, J.A. Capobianco, F. Vetrone, J.G.Sole, D. Jaque. ‘‘Deep Tissue Bio-ImagingUsing Two-Photon Excited CdTe Fluores-cent Quantum Dots Working within theBiological Window’’. Nanoscale. 2012.4(1): 298-302.

257. W. Jiang, A. Singhal, B.Y.S. Kim, J. Zheng,J.T. Rutka, C. Wang, W.C.W. Chan. ‘‘As-sessing Near-Infrared Quantum Dots forDeep Tissue, Organ, and Animal ImagingApplications’’. J. Lab. Autom. 2008. 13(1):6-12.

258. S. Weiss. ‘‘Measuring Conformational Dy-namics of Biomolecules by Single MoleculeFluorescence Spectroscopy’’. Nat. Struct.Biol. 2000. 7(9): 724-729.

259. S. Weiss. ‘‘Fluorescence Spectroscopy ofSingle Biomolecules’’. Science. 1999.283(5408): 1676-1683.

260. T. Pons, H. Mattoussi. ‘‘Investigating Bio-logical Processes at the Single Molecule


Level Using Luminescent Quantum Dots’’.Ann. Biomed. Eng. 2009. 37(10):1934-1959.

261. M.P. Bruchez. ‘‘Quantum Dots Find TheirStride in Single Molecule Tracking’’. Curr.Opin. Chem. Biol. 2011. 15(6): 775-780.

262. F. Pinaud, S. Clarke, A. Sittner, M. Dahan.‘‘Probing Cellular Events, One Quantum Dotat a Time’’. Nat. Methods. 2010. 7(4):275-285.

263. S.R. Opperwall, A. Divakaran, E.G. Porter,J.A. Christians, A.J. DenHartigh, D.E. Ben-son. ‘‘Wide Dynamic Range Sensing withSingle Quantum Dot Biosensors’’. ACSNano. 2012. 6(9): 8078-8086.

264. C.Y. Zhang, H.C. Yeh, M.T. Kuroki, T.H.Wang. ‘‘Single-Quantum-Dot-Based DNANanosensor’’. Nat. Mater. 2005. 4(11):826-831.

265. C.Y. Zhang, J. Hu. ‘‘Single Quantum Dot-Based Nanosensor for Multiple DNA Detec-tion’’. Anal. Chem. 2010. 82(5): 1921-1927.

266. T. Pons, I.L. Medintz, X. Wang, D.S.English, H. Mattoussi. ‘‘Solution-Phase Sin-gle Quantum Dot Fluorescence ResonanceEnergy Transfer’’. J. Am. Chem. Soc. 2006.128(47): 15324-15331.

267. M. Dahan, S. Levi, C. Luccardini, P.Rostaing, B. Riveau, A. Triller. ‘‘DiffusionDynamics of Glycine Receptors Revealed bySingle-Quantum Dot Tracking’’. Science.2003. 302(5644): 442-445.

268. C. Bouzigues, M. Morel, A. Triller, M.Dahan. ‘‘Asymmetric Redistribution ofGABA Receptors During GABA GradientSensing by Nerve Growth Cones Analyzedby Single Quantum Dot Imaging’’. P. Natl.Acad. S. USA. 2007. 104(27): 11251-11256.

269. T. Buerli, K. Baer, H. Ewers, C. Sidler, C.Fuhrer, J.-M. Fritschy. ‘‘Single ParticleTracking of alpha 7 Nicotinic AChR inHippocampal Neurons Reveals RegulatedConfinement at Glutamatergic and GABAer-gic Perisynaptic Sites’’. PLoS ONE. 2010.5(7): e11507.

270. M. Heine, L. Groc, R. Frischknecht, J.C.Beique, B. Lounis, G. Rumbaugh, R.L.Huganir, L. Cognet, D. Choquet. ‘‘SurfaceMobility of Postsynaptic AMPARs TunesSynaptic Transmission’’. Science. 2008.320(5873): 201-205.

271. Q. Zhang, Y.Q. Cao, R.W. Tsien. ‘‘QuantumDots Provide an Optical Signal Specific toFull Collapse Fusion of Synaptic Vesicles’’.P. Natl. Acad. S. USA. 2007. 104(45):17843-17848.

272. Q. Zhang, Y. Li, R.W. Tsien. ‘‘The DynamicControl of Kiss-and-Run and VesicularReuse Probed with Single Nanoparticles’’.Science. 2009. 323(5920): 1448-1453.

273. A.R. Lowe, J.J. Siegel, P. Kalab, M. Siu, K.Weis, J.T. Liphardt. ‘‘Selectivity Mechanismof the Nuclear Pore Complex Characterizedby Single Cargo Tracking’’. Nature. 2010.467(7315): 600-U126.

274. N.P. Wells, G.A. Lessard, P.M. Goodwin,M.E. Phipps, P.J. Cutler, D.S. Lidke, B.S.Wilson, J.H. Werner. ‘‘Time-ResolvedThree-Dimensional Molecular Tracking inLive Cells’’. Nano Lett. 2010. 10(11):4732-4737.

275. S. Doose, J.M. Tsay, F. Pinaud, S. Weiss.

‘‘Comparison of Photophysical and ColloidalProperties of Biocompatible SemiconductorNanocrystals Using Fluorescence CorrelationSpectroscopy’’. Anal. Chem. 2005. 77(7):2235-2242.

276. T. Liedl, S. Keller, F.C. Simmel, J.O. Radler,W.J. Parak. ‘‘Fluorescent Nanocrystals asColloidal Probes in Complex Fluids Mea-sured by Fluorescence Correlation Spectros-copy’’. Small. 2005. 1(10): 997-1003.

277. C. Dong, J. Ren. ‘‘Measurements for MolarExtinction Coefficients of Aqueous QuantumDots’’. Analyst. 2010. 135(6): 1395-1399.

278. R.F. Heuff, J.L. Swift, D.T. Cramb. ‘‘Fluo-rescence Correlation Spectroscopy UsingQuantum Dots: Advances, Challenges andOpportunities’’. Phys. Chem. Chem. Phys.2007. 9(16): 1870-1880.

279. J.L. Swift, R. Heuff, D.T. Cramb. ‘‘A Two-Photon Excitation Fluorescence Cross-Cor-relation Assay for a Model Ligand-ReceptorBinding System Using Quantum Dots’’.Biophys. J. 2006. 90(4): 1396-1410.

280. J.L. Swift, D.T. Cramb. ‘‘Nanoparticles asFluorescence Labels: Is Size All ThatMatters?’’. Biophys. J. 2008. 95(2): 865-876.

281. J.L. Swift, M.C. Burger, D. Massotte, T.E.S.Dahms, D.T. Cramb. ‘‘Two-Photon Excita-tion Fluorescence Cross-Correlation Assayfor Ligand-Receptor Binding: Cell Mem-brane Nanopatches Containing the Humanmu-Opioid Receptor’’. Anal. Chem. 2007.79(17): 6783-6791.

282. H.M. Wobma, M.L. Blades, E. Grekova,D.L. McGuire, K. Chen, W.C.W. Chan, D.T.Cramb. ‘‘The Development of Direct Multi-colour Fluorescence Cross-Correlation Spec-troscopy: Towards a New Tool for TrackingComplex Biomolecular Events in Real-Time’’. Phys. Chem. Chem. Phys. 2012.14(10): 3290-3294.

283. M.L. Blades, E. Grekova, H.M. Wobma, K.Chen, W.C.W. Chan, D.T. Cramb. ‘‘Three-Color Fluorescence Cross-Correlation Spec-troscopy for Analyzing Complex Nanoparti-cle Mixtures’’. Anal. Chem. 2012. 84(21):9623-9631.

284. N. Durisic, A.I. Bachir, D.L. Kolin, B.Hebert, B.C. Lagerholm, P. Grutter, P.W.Wiseman. ‘‘Detection and Correction ofBlinking Bias in Image Correlation TransportMeasurements of Quantum Dot TaggedMacromolecules’’. Biophys. J. 2007. 93(4):1338-1346.

285. A.I. Bachir, D.L. Kolin, K.G. Heinze, B.Hebert, P.W. Wiseman. ‘‘A Guide to Accu-rate Measurement of Diffusion Using Fluo-rescence Correlation Techniques withBlinking Quantum Dot Nanoparticle La-bels’’. J. Chem. Phys. 2008. 128(22).

286. S. Boyle, D.L. Kolin, J.G. Bieler, J.P.Schneck, P.W. Wiseman, M. Edidin. ‘‘Quan-tum Dot Fluorescence Characterizes theNanoscale Organization of T Cell Receptorsfor Antigen’’. Biophys. J. 2011. 101(11):L57-L59.

287. F.-C. Chien, C.W. Kuo, P. Chen. ‘‘Localiza-tion Imaging Using Blinking QuantumDots’’. Analyst. 2011. 136(8): 1608-1613.

288. National Science Foundation (NSF) Divisionof Molecular and Cellular BiosciencesAwards. ‘‘National Science Foundation:

Where Discoveries Begin’’. 2012. http://w w w . n s f . g o v / a w a r d s e a r c h /showAward?AWD_ID=1052733 [accessedNov 12 2012].

289. National Science Foundation (NSF)-FundedCollaborative Research Project QSTORM.‘‘Switchable Quantum Dots and AdaptiveOptics for Super-Resolution Imaging.’’ 2012.www.qstorm.org [accessed Nov 12 2012].

290. Y. Chen, L. Shao, Z. Ali, J. Cai, Z.W. Chen.‘‘NSOM/QD-Based Nanoscale Immunofluo-rescence Imaging of Antigen-Specific T-CellReceptor Responses During an in VivoClonal Vc2Vd2 T-Cell Expansion’’. Blood.2008. 111(8): 4220-4232.

291. J. Chen, Y. Wu, C. Wang, J. Cai. ‘‘NanoscaleOrganization of CD4 Molecules of Human THelper Cell Mapped by NSOM and QuantumDots’’. Scanning. 2008. 30(6): 448-451.

292. J. Chen, Y. Pei, Z. Chen, J. Cai. ‘‘QuantumDot Labeling Based on Near-Field OpticalImaging of CD44 Molecules’’. Micron. 2010.41(3): 198-202.

293. L. Zhong, G. Zeng, X. Lu, R.C. Wang, G.Gong, L. Yan, D. Huang, Z.W. Chen.‘‘NSOM/QD-Based Direct Visualization ofCD3-Induced and CD28-Enhanced Nano-spatial Coclustering of TCR and Coreceptorin Nanodomains in T Cell Activation’’. PLoSOne. 2009. 4(6): e5945.

294. G.W. Walker, V.C. Sundar, C.M. Rudzinski,A.W. Wun, M.G. Bawendi, D.G. Nocera.‘‘Quantum-Dot Optical TemperatureProbes’’. Appl. Phys. Lett. 2003. 83(17):3555-3557.

295. L.M. Maestro, C. Jacinto, U.R. Silva, F.Vetrone, J.A. Capobianco, D. Jaque, J.G.Sole. ‘‘CdTe Quantum Dots as Nanothermom-eters: Towards Highly Sensitive ThermalImaging’’. Small. 2011. 7(13): 1774-1778.

296. P. Haro-Gonzalez, L. Martınez-Maestro, I.R.Martın, J. Garcıa-Sole, D. Jaque. ‘‘High-Sensitivity Fluorescence Lifetime ThermalSensing Based on CdTe Quantum Dots’’.Small. 2012. 8(17): 2652-2658.

297. R.K. Pai, M. Cotlet. ‘‘Highly Stable, Water-Soluble, Intrinsic Fluorescent Hybrid Scaf-folds for Imaging and Biosensing’’. J. Phys.Chem. C. 2011. 115(5): 1674-1681.

298. M.J. Ruedas-Rama, A. Orte, E.A.H. Hall,J.M. Alvarez-Pez, E.M. Talavera. ‘‘A Chlo-ride Ion Nanosensor for Time-ResolvedFluorimetry and Fluorescence Lifetime Im-aging’’. Analyst. 2012. 137(6): 1500-1508.

299. P.P. Provenzano, K.W. Eliceiri, P.J. Keely.‘‘Multiphoton Microscopy and FluorescenceLifetime Imaging Microscopy (FLIM) toMonitor Metastasis and the Tumor Microen-vironment’’. Clin. Exp. Metastas. 2009.26(4): 357-370.

300. Y. Sun, J. Phipps, D.S. Elson, H. Stoy, S.Tinling, J. Meier, B. Poirier, F.S. Chuang,D.G. Farwell, L. Marcu. ‘‘FluorescenceLifetime Imaging Microscopy: In VivoApplication to Diagnosis of Oral Carcino-ma’’. Opt. Lett. 2009. 34(13): 2081-2083.

301. D. Shcherbo, E.A. Souslova, J. Goedhart,T.V. Chepurnykh, A. Gaintzeva, T.W. She-miakina, II, J. Gadella, S. Lukyanov, D.M.Chudakov. ‘‘Practical and Reliable FRET/FLIM Pair of Fluorescent Proteins’’. BMCBiotechnol. 2009. 9: 24.


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