Quantum dot-PNA conjugates for target-catalyzed RNA detection

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Quantum dot-PNA conjugates for target-catalyzed RNA detection Oleksandr Zavoiura, Ute Resch-Genger, and Oliver Seitz Bioconjugate Chem., Just Accepted Manuscript • DOI: 10.1021/acs.bioconjchem.8b00157 • Publication Date (Web): 25 Apr 2018 Downloaded from http://pubs.acs.org on April 26, 2018

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Bioconjugate Chemistry

Quantum dot-PNA conjugates for target-catalyzed RNA detection. Oleksandr Zavoiura,1,2,3 Ute Resch-Genger,1 and Oliver Seitz2 1

Division Biophotonics, Federal Institute for Materials Research and Testing (BAM), Richard-

Willstaetter Str. 11, 12489, Berlin, Germany, 2 Department of Chemistry, Humboldt University of Berlin, Brook-Taylor-Str. 2, 12489 Berlin, Germany,

3

School of Analytical Sciences

Adlershof, Humboldt University of Berlin, Unter den Linden 6, 10099, Berlin, Germany. KEYWORDS: QD, peptide nucleic acid, RNA, templated reactions, RNA detection, nanoparticle click chemistry. ABSTRACT: Detection of pathogenic nucleic acids remains one of the most reliable approaches for the diagnosis of a broad range of diseases. Current PCR-based methods require experienced personnel and cannot be easily used for point-of-care diagnostics, making alternative strategies for the sensitive, reliable, and cost-efficient detection of pathogenic nucleic acids highly desirable. Here, we report an enzyme-free method for the fluorometric detection of RNA that relies on a target-induced fluorophore transfer onto a semiconductor quantum dot (QD), uses PNA probes as selective recognition elements and can be readout with simple and inexpensive equipment. For QD-PNA conjugates with optimized PNA content limits of detection of dengue RNA in the range of 10 pM to 100 nM can be realized within 5 h in the presence of a high excess of non-complementary RNA.

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INTRODUCTION The detection of nucleic acids is essential to corroborate the presence of viral or bacterial species in a variety of biological samples. Today, polymerase chain reaction (PCR) – based techniques are considered a “gold standard” in the field of DNA and RNA diagnostics.1, 2 PCR relies on enzyme-induced replication of the DNA of interest prior to its detection (e.g., by means of fluorescence), thereby enabling target identification principally at the single molecule level. RNA diagnostics is accomplished via reverse transcription-PCR (RT-PCR).3 In this case, RNA needs to be reversely transcribed into its complementary DNA (cDNA) prior to the replication step. An important area of PCR application is the identification and detection of viruses in biological samples. RT-PCR has been successfully applied e.g. to the sensitive detection of RNA viruses such as the dengue virus (DENV).4, 5 The exceptional sensitivity achieved by enzymatic amplification makes PCR, however, susceptible also to cross-contamination by nucleic acids.6, 7 Furthermore, PCR methods are difficult to implement in point-of-care testing (POCT) schemes and are affected by chemicals such as solvents or detergents and therefore require highly trained personnel. Bypassing enzymatic pre-amplification could principally simplify the detection of viral RNA as required for POCT provided that alternative routes are found for the necessary high selectivity and sensitivity of detection. Such routes must be rapid, reliable, and enable simple, preferably optical read-out. One possible way to tackle these challenges is the use of oligonucleotide-templated reactions (OTR) that occur between two reactive hybridization probes and are catalyzed by a nucleic acid target,8,

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typically yielding a fluorescent product.10-21 A number of chemistries have been

successfully utilized to produce a fluorogenic product, such as the Staudinger ligation,14 Rucatalyzed

photoreduction

reactions,16-18,

native

chemical

ligation-based

reactions,22


2

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selenocystine-selenoester ligation20, tetrazine-mediated reactions,13,

21

and others. These

approaches and related modifications have been successfully used to detect nucleic acids in live cells,14,

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cell lysates,12,

21

live vertebrate23 and constitute a promising approach for a wide

spectrum of biological applications. OTR do not need enzymes and can be performed isothermally, hence requiring simpler equipment than PCR. The course of OTR can be conveniently monitored via Förster resonance energy transfer (FRET) using two labels with overlapping absorption and emission bands.19 This can enable ratiometric readout of the measured fluorescence. The vast majority of FRET detection schemes is confined to organic dyes as fluorescent reporters, which renders a relatively high probe concentration mandatory for signal generation due to the moderate brightness (brightness: product of fluorescence quantum yield (QY) and molar extinction coefficient) of these molecular emitters compared to e.g. brighter nanoparticles like semiconductor nanocrystals (quantum dots, QDs).24 Moreover, for organic dyes with their comparatively broad absorption and emission bands, a common drawback of this strategy can be spectral crosstalk, typically direct excitation also of the FRET acceptor. We envisioned that the use of a QD with its very large molar extinction coefficient (ε = 106 - 107 M-1cm-1), high QY, and relatively narrow emission band as FRET donor in conjunction with a near-infrared (NIR) organic dye as FRET acceptor should enhance the signal-to-background ratio and improve the detection sensitivity. In addition, due to their larger size, QDs can act as carrier for several recognition moieties24,

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making them ideal FRET donors for template-catalyzed detection systems. A recent proof-of-concept experiment has demonstrated that DNA-catalyzed dye transfer can proceed on a QD.26 However, the assay suffered from slow reaction kinetics and hence, long reaction times, and was restricted to the detection of the DNA target only in the nM


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concentration range. This lack in sensitivity was primarily attributed to the relatively large distance between the QD and the dye due to the use of streptavidin-biotin conjugation chemistry, hampering FRET efficiency. Other drawbacks included the considerable hydrophobicity of the organic dye, favoring non-specific interactions, and crowding of the acceptor probes (termed in the following label acceptor probe) on the nanoparticle, imposing steric and kinetic constraints. Here, we describe the design of a FRET system for the rapid detection of RNA utilizing modified peptide nucleic acid (PNA) probes and a FRET-pair for signal generation. This FRET pair consists of a red emissive QD as donor and a NIR dye as fluorescent acceptor, optimized regarding FRET efficiency, hydrophilicity, and target selectivity. We chose PNA instead of conventional DNA or RNA-based probes because of the exceptional chemical and enzymatic stability of PNA as well as its high affinity and specificity for RNA.27-31 This allows the use of very specific short probes with high affinity that are not easily degraded by the components of biological systems like certain enzymes. PNA immobilization on the QD was achieved with a two-step strain-promoted azide-alkyne cycloaddition (SPAAC) procedure that yielded bioconjugates of relatively small sizes and allowed to control probe labeling density. For optimum assay sensitivity, subsequently QD-PNA conjugates with different labeling densities in the range of 4 to 59 PNA per QD were prepared and assessed regarding their performance in target-catalyzed RNA detection. RESULTS AND DISCUSSION Detection principle. Most QD Sandwich-type assays for nucleic acid detection rely on proximity-induced FRET induced by adjacent annealing of capture and reporter probes.32-39 In contrast, our detection strategy (Scheme 1) involves two reactive PNA probes and utilizes the ability of one RNA molecule to trigger multiple dye transfer processes, thereby providing a


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higher sensitivity. This is realized by immobilizing a controlled number of label acceptor probes bearing cysteine at the N-terminus on a QD that acts as a probe carrier and FRET donor. The label donor probes are functionalized via a thioester linkage with a Cy5 dye, the absorption band of which overlaps with the QD emission band as a prerequisite for efficient FRET (Supporting information (SI), Figure S8). Annealing of the probes with the target triggers the transfer of Cy5 onto the QD in a native chemical ligation manner. As a target sequence, we chose a highly conserved fragment within the 3’UTR of DENV genomic RNA40 in order to compensate for variations in the DENV genome. This RNA was used as a model nucleic acid target for the proof-of-concept experiments. The main challenges faced during the development of this assay, which are subsequently addressed, included i) the identification of a conjugation chemistry including suitable building blocks that provided control of the labeling density of the QD-PNA conjugates and enabled a high FRET efficiency as a prerequisite for the desired detection limits in the low pM, ii) the design of small PNA probes with high target affinity and solubility in common buffers, and iii) the prevention of non-specific binding between label acceptor and label donor PNA probes. As the biotin/streptavidin conjugation approach previously exploited by us to immobilize label acceptor probes on a QD26 produced nanobioconjugates with a relatively large size, that resulted in relatively low FRET efficiencies, we focused on QD-PNA conjugates with a small distance between FRET donor and acceptor. This was expected to drastically improve the sensitivity of the envisioned assay.


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Cy5

Cy5

Cy5

QD

QD

Cy5

QD

QD

Cy5

Cy5

Cy5

QD

Cy5

Scheme 1. RNA detection utilizing target-templated dye transfer.

Table 1. RNA and PNA sequences used in the study. RNA or PNA sequence

Designation

Sequence (RNA: 5→3 terminus, PNA: N→C terminus)

RNA target

RNAc

GGAAAGACCAGAGAUCCUGCUGUCUCCU

RNA non-complement

RNAnc

GUGCAAAUGGGACUUAGCCGCACUCCUA

RNA-Cy5 complement

RNA-Cy5c

GGAAAGACCAGAGAUC – Cyanine 5

RNA-Cy5 non-complement

RNA-Cy5nc

GACACAAUAGGCGAAG – Cyanine 5

Label acceptor PNA probe

QD-PNA(a-d) (immobilized on a QD)

H-Cys(StBu)-tctctggtctt-Glu-Glu-Orn(N3)-NH2

Label donor PNA probe

PNA2-Cy5

Ac-(Glu)3-Gly-gagacagca-Cys(Cy5-Gly)-NH2

Small gold-PNA nanobioconjugates have been previously prepared via direct modification of the nanoparticles with thiol-containing PNA oligomers.41,

42

This approach exploits the strong

gold-thiol bond. Thiol ligands also show a relatively high affinity towards Cd(II) and Zn(II), which present commonly encountered surface atoms for II/VI core-only and core-shell QDs, as reflected by the use of small thiol ligands such as mercaptopropionic acid (MPA), thioglycolic


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acid (TGA) or various zwitterionic ligands43-47 and DNA bearing thiol groups33,

48

for the

preparation of nanoparticles in water or the phase transfer of hydrophobic QDs to aqueous solution. This strategy, however, does not allow to attach PNA with a cysteine moiety required for acyl transfer to the QD surface, as it competes for the binding with the anchoring groups. Furthermore, preparation of such conjugates via commonly used ligand exchange procedures can lead to a decrease in QD QY.49 Also the colloidal stability of such systems, stabilized with coordinatively bound thiol surface ligands is expected to depend strongly on the dissociation rate of the respective ligand from the nanocrystal surface25 as well as their photoluminescence (PL) QY, which can be critical at low QD concentration. Instead, we used polymer-encapsulated QDs that are supposed to maintain their original brightness independent of QD concentration and microenvironment.25 We anticipated that despite the relatively large size imposed by the polymer shell, such QDs should nevertheless allow for a high FRET efficiency when combined with Cy5 due to the large Förster radius (R0) of this donor-acceptor pair (75 Å; see details on the calculations of R0 in the Experimental procedures section (see equations (eqs.) 1 and 2) and excellent spectral overlap between the emission of the chosen QD and the absorption spectrum of Cy5 given in the SI (Figure S8). Moreover, the presence of multiple acceptor dyes was expected to further boost the fluorescence signal. We decided to use an azide-alkyne cycloaddition as biorthogonal chemistry for PNA conjugation as the azido group is stable during Fmoc-based solid phase peptide synthesis (SPPS) and can be therefore directly incorporated into PNA. While conjugation of DNA probes to iron oxide50 and gold51 nanoparticles has been accomplished via Cu(I)-catalyzed azide-alkyne cycloaddition (CuAAC) in the past, a copper catalyst has been shown to induce irreversible quenching of QD luminescence,52 making this chemistry less desirable for the envisaged


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application. Instead, we developed a two-step SPAAC immobilization strategy that does not require a copper catalyst and proceeds smoothly in a buffer such as MOPS at neutral pH and room temperature (Scheme 2). In a first step, a bifunctional linker BCN-NH2 was used to immobilize the bicyclononyne (BCN) moiety onto the polymer-encapsulated QD via EDCpromoted formation of amide bonds. By varying the molar excess of BCN-NH2, several QDBCN conjugates were prepared and in a next step conjugated to the azido-functionalized label acceptor PNA probes. We intentionally left the N-terminal cysteine of the PNA probes protected in the form of a disulfide to prevent a possible thiol-yne addition.53 Copper-free click chemistry has been reported for attaching cycloalkyne-functionalized DNA to azide-modified nanoparticles,54 yet post-modification was used for the introduction of the cycloalkyne functionality into the DNA strand. In contrast, our approach allows to incorporate the azide group directly during SPPS, thereby eliminating the need for post-modification. Furthermore, our two-step procedure, which seems to be the first example of click chemistry-mediated functionalization of a nanoparticle with PNA probes, allows to control and fine-tune the number of PNA at the nanoparticle. This allowed us to prepare nanobioconjugates with different PNA labeling densities and compare their performance in RNA detection with respect to optimum target selectivity and sensitivity.


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QD

RAM

QD

QD

QD

Scheme 2. Immobilization of the label acceptor PNA probes on the QD via a two-step strain-promoted azide-alkyne cycloaddition (SPAAC)-based strategy.

Design of PNA probes. PNA is an inherently hydrophobic molecule and its hydrophilicity is primarily controlled by its length and purine content, with long and purine-rich probes known to be poorly water soluble. Preliminary experiments revealed that even short (7-mer) non-modified PNA probes are prone to non-specific interactions, limiting the achievable signal-noise-ratio in


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our assay. This was evident by a moderate FRET in the absence of RNA target that we attributed to non-specific binding of label donor PNA probes to the surface of QD, which contained hydrophobic domains created by label acceptor PNA molecules (see SI, Figure S7). We envisioned that the incorporation of negatively charged glutamic acid residues into PNA should enhance the water solubility of these PNA probes. More importantly, electrostatic repulsion was expected to reduce non-specific interactions between label acceptor and label donor PNAs that may lead to false-positive results in the assay. Thus, the label acceptor PNA probe was equipped with two glutamic acids adjacent to the C-terminus in order to decrease the hydrophobicity of the nanoconjugate. Because of its high purine content, the label donor PNA probe was modified with three glutamic acid units next to the N-terminus in addition to its functionalization with the negatively charged sulfo-Cy5 fluorophore. These chemical modifications considerably improved the water solubility of our probes and alleviated non-specific binding between them, resulting in reduced FRET in the absence of the RNA target and, consequently, a lower background. With this functionalization strategy at hand, we optimized the length of the PNA label acceptor and label donor probes with respect to the detection sensitivity for RNA and hybridization rate/assay duration. Experiments with short (7-mer label acceptor and 6-mer label donor) PNA probes revealed that such systems provide only limits of detection of RNA in the nM range even at long incubation times (data not shown). This indicated that the affinity of short PNA to the target was not sufficient for sensitive RNA detection, particularly at sub-nM concentrations of probes. Thus, we set out to enhance the affinity of PNA by increasing the length of probes and tested an 11-mer label acceptor probe and a 9-mer label donor probe (Table 1). This significantly accelerated the hybridization rate for RNA detection, making it possible to minimize the reaction time to 5 h and to decrease the LOD to the low pM range.


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Labeling density of the QD-BCN conjugates. The reactivity of the QD-immobilized BCN groups was corroborated by conjugating the nanoconjugates with azide-labeled Cy5 (Cy5-N3) (Figure 1, panel a) and subsequent absorption and fluorescence measurements. This revealed that we could incorporate different numbers of BCN groups in the range of 7 to 90 Cy5 per particle in a controlled manner which allowed to immobilize a defined number of PNA molecules onto the QD in the next step. Labeling density-dependent changes in the spectral shape of the absorption spectrum of Cy5 (Figure 1, panel b), i.e., the appearance of a new blue-shifted absorption band at the position of the vibronic shoulder of Cy5,55 indicated that surface-bound Cy5 dyes form Htype aggregates on the nanoparticle for systems containing more than 15 Cy5 molecules per particle. This results in a decrease in FRET-mediated Cy5 emission (Figure 1, panel c) as H-type aggregates are commonly barely or non-emissive and can act as energy sinks.56 In order to avoid an underestimation of the number of Cy5 molecules due to the formation of dye aggregates, we used integral absorption coefficients for the quantification of Cy5 per QD ratios except for QDBCN1-Cy5 (Table 2) based on a protocol previously derived by us for dye-protein bioconjugates.57 The procedure and formulas used for the determination of the number of dye molecules per QD are detailed in the Experimental procedures section. The aggregation-induced changes in the spectral overlap integral (J(λ)) in the commonly used formula for calculating FRET efficiencies (eq. 1 and eq. 2) were also subsequently considered (Table 2) utilizing this procedure).


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Figure 1. a) Quantification of QD-immobilized BCN groups with Cy5-azide. b) Normalized absorption spectra of exemplarily chosen QD-BCN-Cy5 conjugates, which underline the formation of non- or barely emissive H-type dimers with increasing number of dye molecules per QD, favoring dye-dye interactions; c) Emission spectra of the QD-BCN-Cy5 conjugates. The formation of H-type dimers with increasing dye labeling density accounts for the apparent diminution in Cy5 emission despite enhanced FRET.

Table 2. Photometric quantification of Cy5 per QD ratios, FRET efficiencies and Förster radii. Conjugate

Cy5/QD (UV-Vis)

J(λ), M-1cm-1nm4

R 0, Å

E(FRET)

rQD, Å*

QD-BCN1-Cy5

7

1.16·1016

74.7

0.89

73

QD-BCN2-Cy5

15

1.25·1016

75.7

0.91

81

QD-BCN3-Cy5

37

1.40·1016

77.2

0.97

79

QD-BCN4-Cy5

90

1.49·1016

77.9

0.99

77


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* rQD were calculated according to eq.5; more details on the calculations are provided in the Experimental procedures section.

Determination of PNA/QD labeling ratio. The accessibility of QD-bound PNA for RNA hybridization is crucial for the assay performance of the nanobioconjugates. In order to verify the presence of PNA on the QD, the QD-PNA conjugates were incubated with the complementary Cy5-labeled RNA (RNA-Cy5c) (Figure 2, panel b). Formation of PNA/RNA duplexes on the QD surface was expected to lead to an increase in FRET. This assumption was also confirmed by positive control experiments with non-complementary Cy5-labeled RNA (RNA-Cy5nc), thereby excluding non-specific adsorption. The emission spectra of four QD-PNA conjugates in the presence of RNA-Cy5c, RNA-Cy5nc, and in the absence of RNA are illustrated in Figure 2, panel a. The number of PNA per particle was calculated from the measured PL intensities of the QD (see eqs. 3 and 6, Scheme S3). These measurements yielded PNA/QD ratios in the range of 4 to 59 for our systems. Amongst the systems studied, non-specific interactions between the QDPNA and RNA-Cy5nc were the weakest for QD-PNAa (Figure 2, panel a and d). Apparently, non-specific RNA binding correlates with the total concentration of PNA on the particle surface. This result agrees well with the study of non-Watson-Crick PNA/DNA interactions in solution reported by Tackett et al.58 These effects, however, do not hamper the sensitive detection of target RNA in the presence of non-complementary RNA sequences in our Sandwich-type assay as will be shown in a following section.


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Figure 2. a) Emission spectra of QD-PNA conjugates in the absence of RNA (red), in the presence of complementary RNA-Cy5 (RNA-Cy5c) (green), and in the presence of non-complementary RNA-Cy5 (RNA-Cy5nc) (blue). b) Principle of quantifying PNA labeling density via FRET. c) Schematic presentation of QD-PNA systems with 4 different labeling densities. d) FRET efficiencies of QD-PNA conjugates in the presence of RNA-Cy5c and RNA-Cy5nc. In order to confirm that FRET between the QD and Cy5 occurs, time-resolved luminescence measurements were performed representatively for QD-PNAd in the absence and in the presence of RNA-Cy5c and RNA-Cy5nc (Figure S9). The PNA and RNA sequences are summarized in Table 1.


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Impact of the temperature on assay performance. The temperature can affect not only the absorption and particularly the PL properties of the QD and dye in an emitter-specific manner, and hence signal intensity in an assay, but also the diffusion kinetics, the equilibrium of PNA/RNA binding, and the stability of the intramolecular RNA structures, respectively. The QY of both QD and Cy5 drop with increasing temperatures (Figure S10), as observed for most fluorophores, making measurements at lower temperatures favorable from the spectroscopic point of view. The temperature affects also the hybridization efficiency of PNA/RNA. At low temperatures, most RNA molecules are expected to hybridize with the PNA probes, with the adjacent annealing of label acceptor PNA (LAPNA) and label donor PNA (LDPNA) bringing the reactive groups of PNA probes in close proximity, resulting in dye transfer and in FRET. However, in order for RNA to facilitate multiple dye transfers it needs to dissociate from the OTR products and hybridize with a fresh set of probes, thereby enabling RNA turnover and triggering multiple chemical reactions between the reactive probes. Increased temperatures favor the dissociation of the RNA template from its ternary complex with the probes thereby establishing a dynamic equilibrium between the bound and unbound RNA molecules. Such a balance is crucial to facilitate OTR. Another important feature of RNA, that needs to be considered in the context of its detection, is its tendency to form secondary structures through intramolecular base-pairing interactions. This makes the target sequence less available for the hybridization with recognition elements. Higher temperatures tend to disrupt the hydrogen bonds responsible for RNA folding and improve thus the accessibility of the RNA target. These factors call for a careful optimization of the temperature regime chosen for efficient OTR. We investigated the impact of temperature on OTR using 0.1 nM and 10 nM of target RNA (RNAc) to catalyze the dye transfer. The best QD-PNA conjugate was expected to provide


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substantial signal enhancements regardless of the RNA concentration applied. QD-PNA conjugates with four labeling densities were used at different QD concentrations. As a measure for the efficiency of dye transfer, the ratio between the intensity of the background-corrected FRET-sensitized emission of Cy5 in the presence and in the absence of RNAc was plotted as a function of temperature (Figure 3). This reveals a notable increase in the efficiency of the OTR for each QD-PNA conjugate with increasing temperature, with the maximum signal-tobackground ratio occurring at temperatures in the range of 44 - 48 °C and 40 - 44 °C for 10 nM RNA and 0.1 nM RNA, respectively. A similar behaviour has been reported previously.22 The higher efficiency of the templated transfer reaction with increasing temperature originates from the reduced affinity of the PNA probes to RNA and the concomitant acceleration of strand exchange reactions. Additionally, the RNA secondary structure becomes more unstable at higher temperatures, which improves its availability for PNA binding. However, temperatures ≥ 52 °C significantly decrease the affinity of the PNA probes and accelerate the hydrolysis of the thioester, thereby offering lower signal-to-background ratios. Interestingly, we found that the QD surface load has an effect on the temperature dependence of the dye transfer efficiency. With QD-PNAa and QD-PNAb the temperature dependence of dye transfer efficiency was more pronounced than when QD-PNAc and QD-PNAd were used. At present, we have no explanation for this behavior. Regardless of the mechanistic origin, our temperature screening studies revealed that for 10 nM RNA, the best signal-to-background ratio was obtained with the system QD-PNAb, while in the case of 0.1 nM RNA, the most sensitive response was achieved with QD-PNAd. Therefore, we subsequently focused on these two systems for assessing the assay sensitivity.


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Figure 3. RNA-catalyzed dye transfer in the presence of 10 nM and 0.1 nM of RNAc at different temperatures for 4 QD-PNA conjugates; the measurements were performed 2.5 h after PNA2-Cy5 addition. The blank is defined as buffer that only contains QD-PNA and PNA2-Cy5. Conditions: Citrate buffer: 40 mM citrate, 2 mM TCEP, 50 mM NaCl, 0.05% (w/v) TWEEN20, pH 7.0. a) C(QD-PNAa ) = 350 pM, C(PNA2-Cy5) = 3.5 nM, b) C(QD-PNAb) = 250 pM, C(PNA2-Cy5) = 3.5 nM, c) C(QD-PNAc) = 50 pM, C(PNA2-Cy5) = 1 nM, d) C(QD-PNAd) = 25 pM, C(PNA2-Cy5) = 1 nM. All experiments were performed in duplicate; the error bars represent standard deviations. These experiments were performed for screening purposes to identify the temperature range providing optimum assay results.

Assay sensitivity. We assessed the systems QD-PNAb and QD-PNAd on the basis of their assay kinetics and target sensitivity/LOD. We deliberately performed these experiments in a buffer containing 100 nM of non-complementary RNA (RNAnc) in order to estimate the impact of foreign RNA on the detection efficiency. As illustrated in Figure 4 (panels a and b), RNAc in the concentration range of 0.1 nM to 100 nM can be detected with QD-PNAb and QD-PNAd within


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80 and 30 min despite the presence of high concentration of RNAnc, respectively. These findings confirm the formation of specific PNA/RNA duplexes on the nanoparticle surface and minimal interferences from RNAnc. As shown in Figure 4, for all concentrations used, the S/N ratios reach a plateau after 150 min, but even for a relatively high concentration of RNA (1 nM) the reaction is not quantitative (no completion reached). This results partly from the hydrolysis of the thioester (see SI, Figure S11), a competitive reaction that renders the LDPNA probe unreactive by cleaving off the fluorophore. Furthermore, the hybridization on the nanoparticle is typically slower compared to that in the solution, thereby requiring more time for the formation of a ternary complex between the PNA probes and RNA. While the S/N starts to decrease (QDPNAb, Figure 4a) or remains constant (QD-PNAd, Figure 4b) after 150 min, for both systems, the intensity of the emission of Cy5 continuously rises after longer reaction times (see SI, Figure S13). The diminished signal enhancement at longer incubation times originates from the increased background signal, induced by non-specific interactions between the label acceptor and the label donor PNA, that does not reach a plateau, but rather increases linearly. This has a more pronounced effect on the S/N ratio for the detection of high RNA concentrations (10 and 100 nM) compared to lower ones (1 and 0.1 nM), respectively. We therefore decided to subsequently perform all assay experiments for a period of 5 h to achieve the broadest possible detection range, since after this time all RNA concentrations assessed induced a signal enhancement exceeding that of the matrix/background and were clearly distinguishable from the latter. A noteworthy observation is that 100 nM RNA produced less signal than 10 nM RNA. At 100 nM the concentration of RNA is 100-fold higher than the concentration of PNA probes (Figure 4b). As a result, the probability that both QD-LAPNA and LDPNA hybridize with the same RNA strand is low. Without an OTR, the FRET efficiency would be extremely low and it would


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be impossible to detect RNA at such high concentrations. However, under conditions of dynamic strand exchange – that is when probes are in an equilibrium between bound and unbound states – the OTR leads to product formation. Next, we performed experiments with a broader range of RNA concentrations. By using conjugate QD-PNAb, 40 pM RNA can be clearly distinguished from the matrix, yet only concentrations exceeding 0.1 nM were above the LOD. The latter was defined here as the sum of the signal induced by 100 nM of RNAnc and three times its standard deviation (Figure 4, panel c). In contrast, the system QD-PNAd offered a wider detection range of 10 pM to 100 nM (Figure 4, panel d), which, combined with the rapid kinetics, makes this system the best candidate for RNA detection. Notably, not only sub-stochiometric amounts of the RNAc, but also a 100-fold excess of this target relative to the label donor probe could be detected with QDPNAd. To corroborate that the detection of RNA was achieved via OTR and not by adjacent annealing of PNA probes, control experiments were performed (see SI, Scheme S4, Figures S12 and S14). Briefly, the on-QD transfer reaction was quenched after 110 min by adding base. Under these conditions, the LDPNA probes are subject to hydrolysis of the thioester, thereby cleaving off the fluorophore (Figure S12). Without the occurrence of a transfer, this treatment would lead to a decrease of the intensity of the QD-induced Cy5 emission. This was not the case (Figure S14) and we concluded that the Cy5 dye was linked to the nanoparticle surface via a stable amide bond.


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Figure 4. a,b) Kinetics of RNA detection for different concentrations of RNAc obtained for QD-PNAb and QD-PNAd in the presence of 100 nM RNAnc. Conditions: a) C(QD-PNAb) = 250 pM, C(PNA2-Cy5) = 3.5 nM. b) C(QD-PNAd) = 25 pM, C(PNA2-Cy5) = 1 nM. Cy5 emission was corrected for the emission of QD only and divided by the signal induced by RNAnc. c,d) Detection of RNAc in the range of concentrations in the presence of 100 nM RNAnc. Cy5 emission was measured after 5 h and plotted without correction. Conditions: c) C(QD-PNAb) = 250 pM, C(PNA2-Cy5) = 3.5 nM. Experiments were performed in duplicate; the error bars represent the standard deviations. d) C(QD-PNAd) = 25 pM, C(PNA2-Cy5) = 1 nM. Experiments were performed at least three times; the error bars represent the standard deviations. LOD was calculated as: matrix signal + 3*standard deviation. Matrix: 100 nM RNAnc in citrate buffer (40 mM citrate, 2 mM TCEP, 50 mM NaCl, 0.05% (w/v) TWEEN20, pH 7.0) and T = 44°C were used for all experiments.

The advancement regarding enzyme-free RNA detection provided by the QD-based OTR assay follows immediately from a comparison with other OTR assays. Although organic dyebased systems developed in the Seitz group gave similar LOD, this required much longer


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reaction times of one day.10 A recently reported fluorophore-based detection strategy that relies on two templated reactions12 yields a similar sensitivity, yet our approach needs only two probes, is more simple, and offers the common benefits of PNA over DNA probes. Rapid templated reactions reported by Shibata et al.11 and Sadhu et al.16 provided lower LODs of 0.5 pM and 5 pM respectively, yet required also more time than our detection scheme, that can be accomplished within a (3 times and 5 times, respectively) shorter time period. Moreover, we defined the LOD based on the signal induced by a high concentration of non-complementary RNA, while all the aforementioned systems used the signal generated by the probes in buffer only without any nucleic acids to define the background. Compared to the majority of previously reported QD-based systems, that do not use templated chemistry as signal enhancement strategy,32, 34, 36, 38, 39 our assay offers a better sensitivity and a wider detection range. A LOD in the fM range for nucleic acids has been only achieved with FRET to a non-emissive quencher.33 A loss in signal as read out for a fluorophore-quencher pair is, however, more prone to unspecific quenching effects that are not related to the target analyte, as the dual fluorophore signaling strategy used by us, as well as to instrumental effects like fluctuations in the excitation light intensity. This can considerably affect assay precision and reliability. Such limitations are elegantly circumvented by our approach using an emissive acceptor. The FRET assay reported by Zhang et al.,35 which utilizes fluorescence measurements of single QDs in a continuous flow, allowed for ultra-sensitive DNA detection in the low fM range. Assay performance in this case required, however, a more complex instrumentation than the assay developed by us, namely a custom-made setup for confocal fluorescence measurements. In contrast, our assay can be performed with standard and easily minituarizable equipment available in every laboratory performing fluorescence assays.


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In our study, we chose DENV RNA as an exemplarily target. The concentration of DENV RNA in the blood of an infected individual is significantly influenced by the day of illness and severity of the disease.59, 60 It has been reported that the concentration of DENV RNA in plasma may be as low as 103.4 RNA copies/ml (corresponding to 4·10-18 M)61 or as high as 109.3 RNA copies/ml (corresponding to 3·10-12 M)59, respectively. Our method may prove suitable for the detection of DENV RNA in samples with high viral titer, provided the target is pre-concentrated prior to its detection. Though further optimization is required, we notice that RNA extracts obtained from 12 ml of human plasma with 107 RNA copies/ml and dissolved in 10 µl a buffer that contains PNA probes provide already a target concentration of 2·10-11 M which exceeds our current limit of detection. CONCLUSIONS AND OUTLOOK. We developed a rapid and sensitive nanoparticle-based system for enzyme-free RNA detection utilizing small QD-PNA nanoconjugates, prepared via copper-free click chemistry. The broadest detection range was achieved with the QD-PNA nanoconjugate that had the highest PNA labeling density, allowing for the detection of RNA in the range of 10 pM to 100 nM within 5 h in the presence of 100 nM of non-complementary RNA sequence. We expect that the limit of detection can be further improved by including denaturating agents which disrupt the secondary structure of RNA and hence allow for the detection at room temperature that would also enhance the QY of the dye and hence boost the luminescence signal. In principle, the assay can be readout with very simple instrumentation (e.g., a filter-based fluorescence sensor) and has the potential to be implemented in POCT detection schemes. Moreover, the assay strategy should be applicable to any kind of polymer-coated nanoparticle, including Cd-free QDs that could ultimately lead to environmentally friendly POCT kits with advanced detection capabilities.


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Furthermore, the QD lifetimes in the range of a few ten nanoseconds can be utilized for timegated emission, which is ideal for the suppression of scattered excitation light and short lived autofluorescence from organic compounds present in complex matrices. The unprecedented brightness of QDs interfaced with amplification by OTR allows to generate optical signals at low probe concentrations. This makes this assay strategy a cost-efficient alternative to traditional enzyme-free nucleic acid detection schemes as the amount of hybridization probes can be reduced. Though the sensitivity of our assay would be sufficient for the detection of high titers of the dengue virus in blood of infected persons, improvements would be required for the diagnosis at frequently occurring low−medium virus titers. We are currently working on approaches to enhance the affinity of the probes and consider the use of faster OTR chemistries to make our approach a powerful stand-alone method for the detection of nucleic acids.

EXPERIMENTAL PROCEDURES Materials. Quantum dots Qdot® 605 VIVID carboxyl were purchased from Thermo Fischer Scientific. RNA oligonucleotides were obtained from Biomers.net, resins for SPPS from Rapp polymer and Novabiochem, and protected amino acids from Novabiochem, Iris Biotech, and Bachem, respectively. PNA monomers were purchased from Link technologies. Cy5-carboxylic acid (product number S 2405) was purchased from FEW chemicals and Cy5-N3 (sulfo-Cy5azide) from Jena Bioscience. Illustra MicroSpin G-50, PD MiniTrap G-10 and G-25 columns were obtained from GE Healthcare, Amicon Ultra-0.5 centrifugal filter devices with NMWL 10 kDa from Merck Millipore, and ultrapure DNase/RNase free water, used for the preparation of buffer solutions, from Thermo Fischer Scientific, respectively. Other chemicals were purchased from Roth, Sigma-Aldrich, Carbolution Chemicals and TCI.


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Modification of carboxyl QDs with BCN groups. Attachment of BCN groups on the surface of QDs was performed according to the following procedure. Carboxyl QDs were mixed with 30 (QD-BCN1), 70 (QD-BCN2), 110 (QD-BCN3) or 200 (QD-BCN4) equivalents of BCN-amine linker (BCN-NH2), and 2,000 equivalents of EDC·HCl in borate buffer (20 mM, pH 7.4). Then, the mixture was thoroughly vortexed and left shaking for 40 min at 25 ˚C (300 rpm). Concentration of QDs during the reaction was kept at 1.2 μM. Next, a 3.5-fold (v/v) excess of borate buffer (50 mM, pH 8.5) was added and solution was left in the dark for additional 40 min in order to hydrolyze the residual active esters. The conjugate was then purified by SEC using Sephadex G25 as an immobile phase and borate buffer (50 mM, pH 8.5) as an eluent. Finally, the buffer was exchanged into MOPS (50 mM, pH 7.2) via ultrafiltration (AmiconUltra, 10 kDa). Three buffer exchange cycles were performed. The retained fraction was then down-spun at 1,000 G for 2 min (20 ˚C) and stored at +4 ˚C. Immobilization of label donor PNA probes. In a typical procedure, QD-BCN conjugate was mixed with a 100-fold (QD-PNAa) or 150-fold (QD-PNAb - QD-PNAd) molar excess of label acceptor PNA probe, vortexed for 2 min and left to react for 3 days at 25°C (300 rpm). Concentration of QD was kept at 1 μM for all conjugates except QD-PNAb (0.6 μM). Afterward, a 3,500-fold excess of 2-azidoacetic acid was added to cap any residual BCN groups (1 day). QD-PNA conjugates were purified using SEC (G50). The recovery of conjugate was performed at 500 G for 2 min (20°C). The conjugates were stored at +4 ˚C with the concentration of stock solutions in the range 0.14 - 0.44 μM. The conjugates can be either reduced prior to the templated transfer reaction for 5 min by the addition of 100-fold (v/v) excess of 15 mM TCEP in MOPS buffer (50 mM, pH 7.2) or reduced in a separate step, purified via SEC G50 and stored in a deprotected form in Ar-saturated buffer.


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Quantification of BCN/QD ratio. In order to estimate the number of BCN groups per nanoparticle, each QD-BCN conjugate was reacted with a 100-500-fold molar excess of Cy5-N3 (depending on the conjugate) at a C(QD) in the range of 0.5-1.0 μM in MOPS buffer (50 mM, pH 7.2) for 7 days at 25 °C. The completion of reaction was monitored via FRET. The prepared QD-BCN-Cy5 conjugates were then purified via SEC (G50). The removal of excess Cy5-N3 was performed at 735 G for 2 min (20°C). In order to confirm that Cy5 is completely removed during the purification process, QD-carboxyl was mixed with the Cy5-N3 and purified in the same fashion as QD-BCN-Cy5 conjugates. Absorption spectra were recorded on a NanoDrop ND1000 spectrometer. Luminescence spectra were recorded on a NanoDrop ND-3300 fluorometer. λex. = 435 nm. For luminescence measurements, the concentration of each conjugate was adjusted to 10 nM with 50 mM borate buffer, pH 8.5. PNA/RNA hybridization on the surface. In order to confirm that QD-immobilized PNA are available for hybridization, QD-PNA conjugates were incubated with 150-fold excess of complementary RNA-Cy5c. RNA-Cy5nc was used as a positive control. Completion of hybridization was monitored via FRET and final photoluminescence spectra were recorded after the emission of FRET donor ceased to decrease (24-28 hs). QD-PNA concentration was kept at 20 pM for all conjugates. All experiments were carried out in borate buffer (50 mM borate, 5 mM TCEP, 50 mM NaCl, pH 8.8) at 25°C in the dark. Emission spectra were recorded on a Fluoromax 4 (Horiba scientific) fluorescence spectrometer. RNA-templated dye transfer. Temperature optimization. Templated dye transfer reactions were performed in 1.5 ml low-binding plastic tubes on a thermo-shaker. First, an aliquote of QDPNA was mixed with a specific amount of RNA in citric acid buffer (40 mM citrate buffer, 2 mM TCEP, 50 mM NaCl, 0.05% (w/v) TWEEN20, pH 7.0) and heated to an indicated


25


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temperature for 25 minutes. Next, PNA label donor probe was added and the reaction was allowed to proceed for 150 min. Afterward the solution was transferred into the microtiter plate and emission spectra were measured on an Infinite® M200pro (Tecan) fluorescence reader. λex = 435 nm. T(meas.) = 31 ± 1 °C. The emission intensity in the Cy5 channel was then integrated in the range of (655-710) nm for QD-PNA(a-c) and (655-680) nm for QD-PNAd) and corrected for the background-generated signal (QD only). Concentration series and kinetics experiments. Fluorescence measurements with the concentration series and kinetics measurements were performed similar to experiments described in the section above, but the luminescence spectra were recorded on Infinite® F200pro (Tecan) fluorescence reader. Band-pass filters λex = 465 (35) nm and λem = 670 (25) nm were used. The temperature during the measurement was kept at 32 ± 1 °C for all samples. QY measurements. The QY of the QD was measured absolutely with a Quantaurus Spectrometer C11347 (Hamamatsu). QY of Cy5 was calculated relative to an Oxazine1 dye (Lambda Physics) (QY = 0.15 in ethanol)62 using a calibrated FLS920 (Edinburgh Instruments) fluorescence spectrometer. Absorption measurements were performed on a Specord 210 Plus (Analytik Jena) spectrometer. The luminescence of the QD and Cy5 were recorded at different temperatures on a Fluoromax 4 (Horiba scientific) fluorescence spectrometer.

Calculations of Förster radii, separation distances and number of acceptors. Förster radii (R0) expressed in Å were calculated according to the formula63 given in equation 1 (eq. 1): = 0.211( (/

(eq. 1)


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Here, κ2 is an orientation factor for randomly oriented transition dipoles (2/3), n is the refractive index of water (1.33), QD the quantum yield of FRET donor (0.80), and J(λ) the overlap integral between the donor´s emission and the acceptor´s absorption band, expressed in M-1cm-1nm4, respectively. The overlap integral was calculated according to equation 2 (eq. 2).

( = ( ( λ

(eq. 2)

Here, FD(λ) is the total emission intensity of the FRET donor normalized to unity, εA(λ) the wavelength-dependent molar extinction coefficient of the FRET acceptor expressed in M-1cm-1, and λ is the wavelength in nm, respectively. The FRET efficiency (E(FRET)) was calculated according to equation 3 (eq. 3). ( ! = 1 −

#$% #$

(eq. 3)

Here, FD and FDA are the emission intensities of the QD in the absence and presence of the acceptor dye, respectively. Based upon the formula for the FRET efficiency in the presence of multiple acceptors (eq. 4), ( ! =

&'()

&'() *+ )

(eq. 4)

equation 5 (eq. 5) for the calculation of the (average) distance separating donor and acceptor was derived. )

, = -

&( . .

(eq. 5)

Here, n is the number of FRET acceptors and r the separation distance between the center of the QD and the FRET acceptor. The number of acceptors was estimated from absorption


27


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measurements and the FRET efficiencies were calculated from the fluorescence spectra of the conjugates (Figure 1 and Table 2) according to eq. 3. On the basis of these parameters, the separation distance between the center of the QD and surface-bound Cy5 was calculated for the QD-BCN-Cy5 conjugates and averaged to estimate the radius of the QD (QD-BCN1-Cy5 was excluded from the averaging procedure for higher precision). The obtained value rav. = 78.8 Å was used for the subsequent calculations. The number of FRET acceptors was calculated from luminescence measurements according to equation 6 (eq. 6). =

.+ )

(eq. 6)

'() ( .

The distance between the center of the QD and Cy5 was calculated as a sum of QD radius (described above) and the contribution of PNA/RNA-Cy5c duplex to the overall size of the conjugate (Scheme S3). The latter was calculated to be 24 Å based on the increase per base pair distance for the PNA/RNA duplexes.64 Thus, the overall separation distance between the center of the QD and Cy5 was calculated to be 102.8 Å. This value was used to calculate the number of PNA per QD according to eq. 6. Quantification of Cy5 to QD ratios based on integral absorbance measurements. Equation 7 (eq. 7) was used to estimate the number of QD-bound Cy5 molecules. /05⁄2 = /456 ⁄/7

(eq. 7)

Here CCy5 and CQD are the concentrations of Cy5 and QD, respectively. The QD concentration was calculated according to equation 8 (eq. 8):

/7 = 89: ; ∙=

(eq. 8),

89:


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Here, A349 is the absorbance at 349 nm, ε349 the extinction coefficient at 349 nm (1.14·107 M1

cm-1), and l the optical pathway (given by the length of the cuvette), respectively. The integral

absorbance of Cy5 was calculated according to equation 9 (eq. 9): ?

456 = @@ >(

(eq. 9)

Here, A(λ) is the absorbance of Cy5, corrected for the absorbance of the QD. In order to correlate the absorbance of Cy5 at its absorption maximum with the integral absorbance the coefficient K was introduced (eq. 10). A = /456B&. ∙ 456 (BCD ∙ E⁄456B&.

(eq. 10)

Here, CCy5mon is the concentration of monomeric Cy5, εCy5(λmax) the molar extinction coefficient at the (longest wavelength) absorption maximum (2.51·105 M-1cm-1), and 456B&. the integral absorbance of monomeric Cy5 calculated according to eq. 9. Finally, the concentration of Cy5 was calculated using equation 11: /456 = A ∙ ;

#FGH

FGH (IJKL ∙=

(eq. 11)

Acknowledgment. The authors thank the School of Analytical Sciences Adlershof (SALSA), the Excellence Initiative of the German Research Foundation (DFG) (GSC 1013), and the DFG (grant RE1203/17-01) for the financial support of this work. Associated Content Available Supporting information Synthesis of the PNA probes and their analytical data, RNA detection with short PNA probes, spectral overlap between the QD and Cy5, time-resolved measurements, temperature-dependent spectroscopic features of the QD and Cy5, kinetics of the thioester hydrolysis, kinetics of RNA detection at different concentrations (raw data), confirmation of reaction mechanism are provided in the SI.


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AUTHOR INFORMATION Corresponding Authors Oliver Seitz: [email protected]; and Ute Resch-Genger: [email protected]. Author Contributions The manuscript was written through contributions of all authors. All authors have given approval to the final version of the manuscript. ABBREVIATIONS BCN, bicyclononyne; CuAAC, Cu(I)-catalyzed azide-alkyne cycloaddition; DENV, dengue virus; FRET, Förster resonance energy transfer; LAPNA, label acceptor PNA; LDPNA, label donor PNA; LOD, limit of detection; NIR, near-infrared; PCR, polymerase chain reaction; PL, photoluminescence; PNA, peptide nucleic acid; POCT, point-of-care testing; QD, quantum dot; QY, quantum yield; R0, Förster radius; RT-PCR, reverse transcription-polymerase chain reaction; SPAAC, strain-promoted azide-alkyne cycloaddition; SPPS, solid phase peptide synthesis. REFERENCES (1)

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39

Bioconjugate Chemistry 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41

Page 40 of 44

Cy5

Cy5

QD

Cy5

Cy5

QD


Page 41 of 44 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43


a) QD

Cy5 Cy5

QD

Cy5

c) QD only QD-BCN1-Cy5 QD-BCN2-Cy5 QD-BCN3-Cy5 QD-BCN4-Cy5

1.0 0.8 0.6 0.4 0.2 0.0 400

480

560  (nm)

QD only QD-BCN1-Cy5 QD-BCN2-Cy5 QD-BCN3-Cy5 QD-BCN4-Cy5

3x104

Emission intensity (a.u.)

Absorbance (normalized)

b)

640

Cy5/QD 4

2x10

1x104

0 ACS Paragon Plus Environment 720 560

600

640  (nm)

680

720



a

1.0x105

b

PNA/QD = 4 8.0x104

QD-PNA only QD-PNA + RNA-Cy5c QD-PNA + RNA-Cy5nc RNA-Cy5c only

6.0x104

Cy5

Cy5

QD

QD

4.0x104 2.0x104

0.0 560

600

640

680

720

(nm)

Cy5

5

1.0x10


8.0x104

Cy5

QD

QD

PNA/QD = 11 QD-PNA only QD-PNA + RNA-Cy5c QD-PNA + RNA-Cy5nc RNA-Cy5c only

6.0x104 4.0x104

c

2.0x104

0.0 560

600

640

680

QD-PNA conjugates with 4 labeling densities

720

QD

(nm)

QD

QD

QD

11

33

59

1.0x105


PNA/QD = 33 8.0x104

QD-PNA only QD-PNA + RNA-Cy5c QD-PNA + RNA-Cy5nc RNA-Cy5c only

6.0x104 4.0x104

PNA/QD

d

2.0x104

0.0 560

600

640

680

4

1.0

0.8

720

RNA-Cy5c RNA-Cy5nc

PNA/QD = 59 1.0x105 QD-PNA only QD-PNA + RNA-Cy5c QD-PNA + RNA-Cy5nc RNA-Cy5c only

8.0x104 6.0x104

E(FRET)

(nm) 1.2x105


1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 42 of 44

0.6

0.4

0.2

4.0x104

0.0 QD-PNAa

4

2.0x10

QD-PNAb

QD-PNAc

Conjugate 0.0 560

600

640 (nm)

680

720 ACS Paragon Plus Environment

QD-PNAd

Page 43 of 44

12

8 10 nM 4

b

QD

F(Cy5)RNA/F(Cy5)blank


16

a

QD

12 10 nM

8

4 0.1 nM

0.1 nM 0

0 36

40

44

48

52

56

36

40

T (°C)

44

48

52

56

T (°C)

c

QD

9


9


1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41


6

10 nM 3

d

QD

6

10 nM

3

0.1 nM

0.1 nM

0

0 36

40

44

48

T (°C)

52

56

36

40

44

48

T (°C)


52

56

Bioconjugate Chemistry 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47


Page 44 of 44

Quantum dot-PNA conjugates for target-catalyzed RNA detection

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