Detection of Ligand-Induced Conformational Changes in

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Detection of ligand-induced conformational changes in oligonucleotides by second-harmonic generation at a supported lipid bilayer interface Margaret T. Butko, Ben Moree, Richard B Mortensen, and Joshua Salafsky Anal. Chem., Just Accepted Manuscript • DOI: 10.1021/acs.analchem.6b02498 • Publication Date (Web): 04 Oct 2016 Downloaded from http://pubs.acs.org on October 6, 2016

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Analytical Chemistry

Detection of ligand-induced conformational changes in oligonucleotides by second-harmonic generation at a supported lipid bilayer interface Margaret T. Butko, Ben Moree, Richard B. Mortensen, Joshua Salafsky*

* [email protected], Biodesy, Inc. 384 Oyster Point Blvd Suite #8, South San Francisco, CA 95080

ACS Paragon Plus Environment


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Abstract There is a high demand for characterizing oligonucleotide structural changes associated with binding interactions, as well as identifying novel binders that modulate their structure and function. In this study, second-harmonic generation (SHG) was used to study RNA and DNA oligonucleotide conformational changes associated with ligand binding. For this purpose we developed an avidin-based biotin capture surface based on a supported lipid bilayer membrane. The technique was applied to two well-characterized aptamers, both of which undergo conformational changes upon binding either a protein or a small molecule ligand. In both cases, SHG was able to resolve conformational changes in these oligonucleotides sensitively and specifically, in solution and in real time, using nanogram amounts of material. In addition, we developed a competition assay for the oligonucleotides between the specific ligands and known, nonspecific binders, and we demonstrated that intercalators and minor groove binders affect the conformation of the DNA and RNA oligonucleotides in different ways upon binding, and subsequently block specific ligand binding in all cases. Our work demonstrates the broad potential of SHG for studying oligonucleotides and their conformational changes upon interaction with ligands. As SHG offers a powerful, high-throughput screening approach, our results here also open an important new avenue for identifying novel chemical probes or sequence-targeted drugs that disrupt or modulate DNA or RNA structure and function.

Introduction The range of central roles played by oligonucleotides in biology, particularly RNA, is widening as our understanding of how their structures and conformational changes modulate biological function grows. RNA adopts different secondary and tertiary structural motifs crucial for various functions in transcription, translation, and gene regulation, among others.1-7 Conversely, dysregulation or structural alteration of RNA and DNA can lead to cellular dysfunction and disease.8-14 Structural techniques such as NMR and X-ray crystallography have allowed a glimpse into the wide range of structures RNA can adopt.15-18 However, because


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RNA is relatively small, structurally dynamic, and chemically unstable, RNA targets can be difficult to study using conventional structural techniques.15, 19-22 In addition, these structural techniques can be laborious and are typically not well suited for screening ligands or drug candidates that can modulate RNA conformation. On the other hand, affinity assays such as surface plasmon resonance or isothermal titration calorimetry have provided insight into mechanisms governing ligand binding and are amenable to screening ligands,23-26 but typically cannot distinguish different conformational states of the oligonucleotide, for example when in the ligand-bound or ligand-free state, information crucial to understanding the mechanism of action of a ligand or drug.18, 27 Moreover, oligonucleotide binding assays are often confounded by promiscuous binders that bind nucleotide-based structures nonspecifically.28-30 A method to measure conformational changes in oligonucleotides sensitively in solution and in real time would be a useful complement to these methods for studying structure-activity relationships and for developing drugs that target oligonucleotides, including pathological RNA. In this work, we used second-harmonic generation (SHG), an intrinsically surface-selective technique, to study oligonucleotides at an interface. SHG is a nonlinear optical technique in which two photons of equal energy are combined by a nonlinear material or molecule to generate one photon with twice the energy.31 In isotropic materials and media, SHG occurs only at interfaces that break centrosymmetry, a prerequisite for the effect.32-38 Although biological molecules are not usually SH active, they can be rendered so through the incorporation of a SHactive dye molecule.39-40 Moreover, because the intensity of the SHG signal is highly sensitive to the angular orientation of the dye, the technique has been shown to be a powerful method for studying structure and conformational changes.41-45 Once tethered to a surface, a labeled SH-active biomolecule, irradiated by a fundamental light beam, produces an SHG signal whose intensity depends sensitively on the tilt angle of the dye with respect to the surface. When the biomolecule undergoes a conformational change upon ligand binding, this causes a change in the orientational distribution of the SH-active moiety, whether temporal, spatial, or both, leading to a change in the intensity of light generated and providing an



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instantaneous, real-time read out. In addition, conformational changes can be resolved and classified by their response in magnitude and direction of the SHG signal change relative to baseline, thereby allowing for discrimination of the ligandinduced conformational states. Because of the structural plasticity of RNA and DNA, it was unclear whether an SHG-based approach would be useful for detecting ligand-induced changes in these molecules. The present work was motivated in part by an interest in testing the sensitivity of SHG to study this biologically important class of molecules. In our previous work with proteins we employed glass-supported lipid bilayer surfaces, which comprised lipids with Ni-nitrilotriacetic acid (NTA) head groups for capturing His-tagged proteins to the surface.41-42 However, it was more convenient for us to develop a biotin-based attachment strategy for oligonucleotides in which a lipid bilayer membrane doped with biotin-bearing lipids was used to couple neutravidin, which in turns allows for capture of biotinylated oligonucleotides. Biotin-based capture of molecules has been used previously by others for SHG measurements.34 Building on our prior work with proteins, our main aim here was to study oligonucleotide conformational changes in response to binding both specific and nonspecific ligands. For this purpose we chose two oligonucleotide aptamers as model systems, both well characterized by other techniques. First, we studied a 33base theophylline-binding RNA sequence that adopts a short hairpin and undergoes conformational change upon binding specific ligands.46 The structure of this RNA aptamer in the presence and absence of theophylline has been determined by NMR measurements.47 The NMR structure reveals that the top GAAA loop and the stems form without the ligand, while the inner bulges are not well structured. Addition of theophylline stabilizes the internal bulges and induces a conformational change. The second aptamer was a 31-base thrombin-binding DNA sequence that undergoes conformational change to form a G-quartet structure upon ligand binding,48-49 a common structural motif found in both DNA and RNA that determines physiological function.50-51 A biotin moiety was conjugated to both oligonucleotides along with an SH-active dye used previously for studies with proteins.41-42 Both the specific ligands, theophylline and thrombin, and two nonspecific binders, ethidium bromide


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(EtBr) and netropsin, were tested for their effect on oligonucleotide conformation. Both nonspecific molecules are known to bind to DNA or RNA independent of sequence and cause a stiffening of the oligonucleotide structure.52-58

Experimental Section Materials Oligonucleotides were synthesized, conjugated, and purified by TriLink Biosciences using the following sequences: 5’GGCGAUACCAGCCGAAAGGCCCUUGGCAGCGUC-3’ (theophylline aptamer) and 5’TAAGTTCATCTCCCCGGTTGGTGTGGTTGGT-3’ (thrombin aptamer). Biotin was conjugated to the 5’ end (Biotin BB) and the SH-active dye (Biodesy-1) was conjugated to the 3’ end of each oligonucleotide. Theophylline (Sigma T1633), caffeine (Sigma W222402), EtBr (Sigma E8751), and netropsin (Sigma N9653) were solubilized in endotoxin-free water or MilliQ-purified water. These compounds, as well as human α-thrombin (Haemtech HCT-0020), and dihydrofolate reductase (DHFR) (Biodesy 930-4) were diluted to the desired concentration in assay buffer before injection into the sample well on the Biodesy Delta system.

Surface preparation Biodesy’s Avidin membranes, used to immobilize biotinylated biomolecules, are comprised of biotinylated lipids doped into the lipid bilayer mixture. Upon bilayer formation, neutravidin can be captured by the surface, as demonstrated by the large increase in SHG signal upon injection of SHG dye-conjugated neutravidin onto the biotinylated bilayer surface (Figure 1). This increase in signal approaches a plateau indicating saturation at ~60 minutes.


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Average SHG intensity (cps)


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Figure 1. Neutravidin labeled with an SH-active dye produces a large increase in SHG signal on a biotin-doped supported lipid bilayer, indicating that neutravidin binds to and is ordered on the surface. SHG signal is photons or counts per second (cps). Error bars indicate standard deviation.

We used Biodesy’s Avidin lipid bilayer membranes for all experiments. In addition to biotinylated lipid, the membranes also contain a lipid coupled to rhodamine, which was used to monitor by fluorescence microscopy the uniformity and fluidity of the bilayer surfaces throughout the experiments. To prepare the supported lipid bilayer surfaces, Biodesy’s Avidin membranes were diluted 5-fold in Tris-buffered saline (TBS), added to wells on a Biodesy Read Plate (384-well format), incubated at room temperature for 30 minutes to allow bilayer formation, and washed with TBS to remove excess small unilamellar vesicles (SUVs). Notably, the washout procedure ensures that the well volume never falls below 10 μL to avoid disrupting the lipid bilayer membrane. After SUV washout, neutravidin was diluted in TBS, added to the wells, incubated at room temperature for one hour to allow specific attachment to the bilayer surface, and then washed with assay buffers previously verified to be compatible with these aptamers (RNA: 100 mM HEPES pH 7.3, 50 mM NaCl, 5 mM MgCl2; DNA: 100 mM Tris pH 7.4, 140 mM NaCl, 20 mM MgCl2, 20 mM KCl)46, 59 to remove unbound neutravidin while keeping the layer under aqueous buffer at all times. After neutravidin washout, the conjugated oligonucleotides were diluted to 2 μM in assay buffer, added to the corresponding


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wells (10 μL into 10 μL for a final well concentration of 1 μM), incubated at room temperature for one hour to allow capture to the tethered neutravidin, and washed with assay buffer to remove unbound oligonucleotide. A schematic of the surface architecture and ligand-induced conformational change is shown in Figure 2. The final volume in the plate before SHG measurement was 20 μL. The plate was then placed on Biodesy’s Delta system for SHG measurements.

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Figure 2. Schematic of a biotinylated and labeled oligonucleotide tethered to a supported lipid bilayer membrane. The biotinylated lipid-containing bilayer (pink) captures neutravidin (blue-grey), which is used to capture biotinylated (dark grey), SHG dye-labeled (blue) oligonucleotides. The intensity of the SHG light signal is highly dependent on the time and space-averaged tilt angle (orientational distribution) of the SH-active dye (blue) conjugated to the oligonucleotide. Upon ligand (green) binding, changes in the mean tilt angle or the orientational width of the dye angular distribution produces a change in SHG intensity, which is measured instantaneously.

Instrumentation The Biodesy Delta system is a 384-well microplate-based instrument, combining optical detection of the SHG signal with automated liquid handling to enable detection of conformational change in real-time, synchronized with ligand injection. The optical subsystem comprises a femtosecond Ti:S laser as the source of the fundamental beam at 800 nm. The fundamental is directed to an arrangement of



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prisms integrated with the custom, glass-bottomed Biodesy Delta Read Plates, where it undergoes total internal reflection to generate the evanescent field at the glass-lipid bilayer interface. This field interacts with the SH-active probe attached to the bilayer-tethered protein to generate the SH light at 400 nm, which emerges from the interface in a collimated beam nearly collinear with the fundamental beam. The fundamental light is filtered out leaving only the SH light, which is detected by a photomultiplier tube and processed with custom electronics. The SHG signal is the intensity of the SH light in photons or counts per second (cps).

SHG Measurements Compound or control buffer injections and SHG detection were carried out on the Biodesy Delta system with the following injection and mix protocol: after baseline signal is read, 20 μL of the compound at 2x the desired final concentration is injected into 20 μL of solution volume in the well. After one mixing cycle, 20 μL is then removed from the well. SHG signal was read at multiple time points after injection at user-defined intervals to ensure the plateau of the response was reached. For competition measurements, the first compound was injected as described above. After the response from the first compound plateaued, a mixture of the second compound at 2x well concentration and the first compound at 1x well concentration were injected to keep the concentration of the first compound constant during the second injection. For washout measurements, after the compound response plateaued, 50 μL of assay buffer was added and mixed, and 50 μL was then removed from the well. This procedure was repeated four more times for a total dilution of 17.5-fold before the SHG signal was measured again after washout.

Data Analysis Percent change in SHG intensity (%∆) at a given time point t was calculated as: %∆ = ((It-It0)/It0) x 100


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where It is the SHG intensity at time t and It0 is the SHG intensity at time 0 before injection. Dose-response curves were analyzed using Prism software (GraphPad Software, Inc., La Jolla, CA) and were fitted with a y = A1 + (A2-A1)/(1 + 10(Log(x0-x)*p) equation, where Log(x0) is the EC50 value for the given inhibitor, p is a Hill slope, and A1 and A2 reflect the minimum and maximum of the observed signal.

Results and Discussion To explore SHG’s sensitivity to conformational changes in a DNA oligonucleotide, we tethered the SHG dye-labeled thrombin aptamer to the avidin surface via its 5’ biotin linkage and a baseline signal was established. Thrombin (1 μM), a specific ligand for the oligonucleotide, was injected into the well resulting in an increase in the SHG intensity (Figure 3a), indicating that a conformational change in the DNA oligonucleotide occurs upon binding, as expected. This conformational change was specific to the DNA oligonucleotide, as no response was detected on surfaces prepared identically but pre-incubated with excess, blocking biotin in an additional step prior to capturing the oligonucleotide. As a control, dihydrofolate reductase (DHFR), a protein not expected to bind, did not induce conformational change in the oligonucleotide upon injection. The thrombin-induced conformational change was dose-dependent (Figure 3b), and the measured EC50 of the interaction was measured to be 185 nM. Previous affinity measurements have indicated a binding dissociation constant (Kd) of 103 nM for this interaction.59-60 The SHG signal monitors the observed conformational change associated with ligand binding in real time, and at a saturating thrombin concentration (1 μM) this response occurs over a timescale of minutes, plateauing in ~10 – 15 minutes (Figure 3c). Previous measurements of this conformational change to a G-quartet structure have also indicated that this change occurs on a timescale of minutes.59, 61


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Figure 3. Oligonucleotide response to target molecules. (a) Thrombin but not a control protein, DHFR, induces an increase in SHG intensity for the DNA oligonucleotide, but not on surfaces blocked by pre-incubation with excess free biotin. (b) The thrombin response is dose-dependent with an EC50 of 185 nM for the interaction. (c) At a saturating dose (1 μM) the thrombin response takes ~10 – 15 minutes to plateau. (d) Theophylline but not caffeine induces an increase in SHG intensity for the RNA oligonucleotide but not on surfaces blocked by pre-incubation with excess free biotin. (e) The theophylline response is dose-dependent with an EC50 of 2.5 μM for this interaction. (f) At a saturating dose (30 μM) the theophylline


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response takes less than a minute to reach a plateau. Error bars indicate standard deviation.

We next tested our method with a well-characterized RNA aptamer that binds theophylline. The SHG dye-conjugated RNA oligonucleotide was tethered to the avidin surface via its 5’ biotin linkage, and changes in SHG intensity were measured upon addition of theophylline (30 μM). When the oligonucleotide was tethered to the surface, an increase in SHG was measured upon theophylline injection (30 μM) (Figure 3d). This conformational response did not occur on surfaces prepared by pre-incubating the tethered neutravidin with excess free biotin. The response was also specific for theophylline because caffeine, a structurally similar molecule that differs from theophylline by a methyl group and is known not to bind to the oligonucleotide, did not produce a change in the SHG signal. The theophylline response was dose-dependent (Figure 3e), and the measured EC50 for the interaction was 2.5 μM. Previous affinity measurements have indicated a binding dissociation constant of 320 nM for this interaction.46 In comparison to the thrombin response on the DNA oligonucleotide, the theophylline response on the RNA oligonucleotide was rapid at a saturating theophylline concentration (30 μM)(Figure 3f), reaching a plateau by the first time point (Figure 3f), similar to the response of protein conformational changes measured previously by SHG.41-42 It is well established that nonspecific intercalators and minor groove binders can rigidify oligonucleotides upon binding.52-58 Minor groove binders offer the highest sequence specificity for disrupting gene expression, and these molecules are being investigated as anti-cancer agents.62-68 We studied the effects of nonspecific oligonucleotide binders on the conformation of these oligonucleotides. EtBr, an intercalator, and netropsin, a minor groove binder, were injected at 30-fold molar excess in separate experiments on surfaces containing the DNA thrombin-binding oligonucleotide, the RNA theophylline-binding oligonucleotide, or blank bilayer containing no bound oligonucleotides. EtBr (30 μM) did not induce a detectable change in SHG for the DNA thrombin-binding oligonucleotide, but it did produce a



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small signal change in SHG for the RNA theophylline-binding oligonucleotide (Figure 4a), and this concentration was saturating for the latter response (Figure 4d). Whereas theophylline injection induced an increase in SHG intensity, EtBr injection resulted in a decrease in SHG intensity, indicating a different movement of the dye with respect to the surface, providing evidence that EtBr binds to a different structural state than theophylline. Netropsin (30 μM) produced no detectable change in SHG signal for the theophylline-binding RNA oligonucleotide as expected because it is known not to bind to double-stranded RNA.69 However, netropsin (30 μM) did cause a large increase in SHG intensity upon binding the thrombin-specific DNA oligonucleotide (Figure 4a), and this concentration was saturating for the response (Figure 4c). The netropsin response on the DNA oligonucleotide was different in magnitude from the thrombin response, providing evidence for a different conformation from the one stabilized by binding thrombin. Neither EtBr nor netropsin induced a change in SHG on the blank bilayer surface, indicating that the responses observed were specific for the tethered oligonucleotides.


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Figure 4. Effect of nonspecific DNA binders on oligonucleotide conformation. (a) EtBr induces a small decrease in SHG intensity for the RNA oligonucleotide, distinct from the theophylline response, but causes no response for the DNA oligonucleotide or bilayer alone. In contrast, netropsin induces a large increase in SHG intensity for the DNA oligonucleotide, which is distinct from the thrombin response, but no response for the RNA oligonucleotide or bilayer alone. In the presence of high doses of EtBr (b) or netropsin (c), the thrombin response for the DNA oligonucleotide is blocked. Similarly, in the presence of high doses of EtBr (d) or netropsin (e), the theophylline response for the RNA oligonucleotide is blocked. Dose-response curves



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for these interactions are included in Figure S1. Error bars indicate standard deviation.

Next we explored whether the presence of nonspecific binders would prevent subsequent specific ligand binding. Therefore, we established a competition assay between EtBr and netropsin and either thrombin or theophylline for the DNA and RNA assays, respectively. To test whether EtBr competes with thrombin binding, a saturating dose of thrombin (1 μM) was injected onto the surfaces containing the thrombin-binding DNA oligonucleotide pre-incubated with EtBr at a range of doses. At high EtBr concentrations (> 3 μM), the thrombin response was completely blocked, and this inhibition was dose-dependent, with almost no inhibition at 0.1 μM (Figure 4b, Figure S1a). The results indicate that although there was no detectable conformational change upon injection of EtBr (Figure 4a), the DNA oligonucleotide bound EtBr and this interaction subsequently blocked its interaction with thrombin, presumably by stabilizing the EtBr-bound conformation and preventing conformational change to the thrombin-bound form. Netropsin, a classical minor groove binder used for anti-cancer studies,58, 62, 66 was tested in a similar manner, and it also blocked the thrombin response in a dose-dependent manner (Figure 4c, Figure S1b). However, netropsin required much higher concentrations for inhibition than EtBr, suggesting that netropsin is less potent at blocking this interaction. EtBr and netropsin were also tested for an ability to compete with theophylline in the RNA oligonucleotide. Similar to the DNA oligonucleotide, theophylline binding was inhibited by both EtBr (Figure 4d, Figure S1c) and netropsin (Figure 4e, Figure S1d) in a dose-dependent manner. EtBr appears to be more potent than netropsin at inhibiting theophylline ligand binding as well. Furthermore, these data indicate that although there was no detectable conformational change upon injection of netropsin on the RNA oligonucleotide (Figure 4a), netropsin bound to this molecule because the theophylline response was partially blocked in the presence of high doses of netropsin. These data demonstrate that nonspecific binders can cause structural changes on their own and once bound, can prevent the ligand-induced conformational changes that


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accompany binding of specific ligands (theophylline or thrombin). These data also reveal that the technique only detects binding that leads to conformational change, which can be limiting in studies focused on binding per se but advantageous for those focused on ligand-induced conformational changes that typically accompany functional changes. The reversibility of the interactions was also studied by performing buffer washout experiments following compound injection. To test the reversibility of the DNA-thrombin interaction, thrombin was first injected at a saturating concentration (1 μM), the response was allowed to plateau after about 12 minutes, and then wells were washed with fresh assay buffer, diluting thrombin to a final concentration of approximately 50 nM, well below the EC50 (Figure 5a). Washout partially reversed the thrombin response, indicating reversibility of this interaction. Furthermore, reinjection of thrombin after washout restored the thrombin response. The reversibility of the RNA-theophylline interaction was measured in a similar experiment (Figure 5b), and the response was partially reversed by washout and restored upon re-injection of theophylline.

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Figure 5. Washout of ligands reverses the conformational response to ligand binding. (a) Thrombin induces an increase in SHG signal for the DNA oligonucleotide, and this response was partially reversed upon compound washout and restored upon re-injection of thrombin. (b) Theophylline induces an increase in SHG signal for the RNA oligonucleotide, and this response was partially reversed upon compound washout and restored upon re-injection of theophylline. Error bars



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indicate standard deviation. The arrows and dashed lines denote ligand addition or the beginning of a washout sequence.

Summary Our results demonstrate SHG detection of RNA and DNA oligonucleotide conformational changes sensitively and in real time upon binding either protein or small molecule ligands. SHG provides a rapid means to sensitively probe ligandinduced conformational changes as a function of oligonucleotide sequence to dissect a ligand’s mechanism of action. The data presented in this paper demonstrate that ligand-induced conformational changes upon binding to biotinylated DNA or RNA oligonucleotides can be detected and resolved by SHG using avidin capture to a biotinylated supported lipid bilayer. The measurements can be used to characterize ligand interactions by dose-dependence, kinetics, and reversibility via the detected oligonucleotide conformational changes. While aptamers generally undergo dramatic conformational changes upon ligand binding, we expect SHG’s sensitivity will allow for detection of much more subtle conformational changes as well.42 The nonspecific ligands EtBr and netropsin were shown to bind both the DNA and RNA oligonucleotides, producing significant or relatively small conformational responses depending on the system, and they subsequently blocked the conformational changes associated with binding specific protein or small molecule ligands. Importantly, netropsin and EtBr induced detectable conformational responses upon binding to DNA and RNA, respectively, but these responses are distinct from those produced by the specific ligands for these oligonucleotides (thrombin or theophylline), providing evidence that the specific ligands bind to different conformational states than the nonspecific ones. Conformational selection or an induced fit mechanism must determine nonspecific ligand binding to the oligonucleotides. SHG is a straightforward method for characterizing ligand-induced structural changes of DNA and RNA oligonucleotides, which requires comparatively little material, assay development effort, and measurement time. By implementing our measurements in a 384-well plate, only 20 picomoles of oligonucleotide were used


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per well. Furthermore, different characteristics of an interaction can readily be resolved, including distinct conformational changes upon ligand binding, competition studies between ligands, and ligand binding reversibility. Our work here provides a foundation for studying any oligonucleotide-ligand interaction, as well as a method for identifying novel conformational modulators of oligonucleotides at high throughput for chemical biology studies and for sequencespecific drugs for personalized medicine.

Author Contributions M.B., B.M., R.M., and J.S. designed the experiments. M.B. performed the experiments and analyzed the data. M.B. and J.S. wrote the manuscript. J.S. supervised the work.

Conflicts of Interest M.B., B.M., R.M., and J.S. are employees of Biodesy, Inc. J.S. is the founder and chief scientific officer of Biodesy, Inc.

Acknowledgements The authors gratefully acknowledge Razvan Nutiu, Kirk Wright, Jaclyn Greimann, and Sam Kintz for helpful suggestions and discussions.

Supporting Information IC50’s of nonspecific binders measured by %inhibition of the thrombin or theophylline SHG response are provided in the supporting information.

References 1. Cruz, J. A.; Westhof, E., The dynamic landscapes of RNA architecture. Cell 2009, 136 (4), 604-9. 2. Dethoff, E. A.; Chugh, J.; Mustoe, A. M.; Al-Hashimi, H. M., Functional complexity and regulation through RNA dynamics. Nature 2012, 482 (7385), 32230. 3. Kozak, M., Regulation of translation via mRNA structure in prokaryotes and eukaryotes. Gene 2005, 361, 13-37.



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

4. Mauger, D. M.; Siegfried, N. A.; Weeks, K. M., The genetic code as expressed through relationships between mRNA structure and protein function. FEBS Lett 2013, 587 (8), 1180-8. 5. Mortimer, S. A.; Kidwell, M. A.; Doudna, J. A., Insights into RNA structure and function from genome-wide studies. Nat Rev Genet 2014, 15 (7), 469-79. 6. Sharp, P. A., The centrality of RNA. Cell 2009, 136 (4), 577-80. 7. Simon, A. E.; Gehrke, L., RNA conformational changes in the life cycles of RNA viruses, viroids, and virus-associated RNAs. Biochim Biophys Acta 2009, 1789 (910), 571-83. 8. Bernat, V.; Disney, M. D., RNA Structures as Mediators of Neurological Diseases and as Drug Targets. Neuron 2015, 87 (1), 28-46. 9. Cheng, J.; Guo, J. M.; Xiao, B. X.; Miao, Y.; Jiang, Z.; Zhou, H.; Li, Q. N., piRNA, the new non-coding RNA, is aberrantly expressed in human cancer cells. Clin Chim Acta 2011, 412 (17-18), 1621-5. 10. Di Leva, G.; Garofalo, M.; Croce, C. M., MicroRNAs in cancer. Annu Rev Pathol 2014, 9, 287-314. 11. Esteller, M., Non-coding RNAs in human disease. Nat Rev Genet 2011, 12 (12), 861-74. 12. Han Li, C.; Chen, Y., Small and Long Non-Coding RNAs: Novel Targets in Perspective Cancer Therapy. Curr Genomics 2015, 16 (5), 319-26. 13. Tsai, M.-C.; Spitale, R. C.; Chang, H. Y., Long intergenic non-coding RNAs – New links in cancer progression. Cancer research 2011, 71 (1), 3-7. 14. Wapinski, O.; Chang, H. Y., Long noncoding RNAs and human disease. Trends Cell Biol 2011, 21 (6), 354-61. 15. Bothe, J. R.; Nikolova, E. N.; Eichhorn, C. D.; Chugh, J.; Hansen, A. L.; AlHashimi, H. M., Characterizing RNA dynamics at atomic resolution using solutionstate NMR spectroscopy. Nat Methods 2011, 8 (11), 919-31. 16. Montange, R. K.; Batey, R. T., Structure of the S-adenosylmethionine riboswitch regulatory mRNA element. Nature 2006, 441 (7097), 1172-5. 17. Serganov, A.; Huang, L.; Patel, D. J., Structural insights into amino acid binding and gene control by a lysine riboswitch. Nature 2008, 455 (7217), 1263-7. 18. Serganov, A.; Huang, L.; Patel, D. J., Coenzyme recognition and gene regulation by a flavin mononucleotide riboswitch. Nature 2009, 458 (7235), 233-7. 19. Felden, B., RNA structure: experimental analysis. Curr Opin Microbiol 2007, 10 (3), 286-91. 20. McFadden, E. J.; Hargrove, A. E., Biochemical Methods To Investigate lncRNA and the Influence of lncRNA:Protein Complexes on Chromatin. Biochemistry 2016, 55 (11), 1615-30. 21. Reyes, F. E.; Garst, A. D.; Batey, R. T., Strategies in RNA crystallography. Methods Enzymol 2009, 469, 119-39. 22. Zhang, J.; Ferre-D'Amare, A. R., New molecular engineering approaches for crystallographic studies of large RNAs. Curr Opin Struct Biol 2014, 26, 9-15. 23. Buurma, N. J.; Haq, I., Advances in the analysis of isothermal titration calorimetry data for ligand-DNA interactions. Methods 2007, 42 (2), 162-72.


Page 18 of 22

Page 19 of 22

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


24. Chang, A. L.; McKeague, M.; Liang, J. C.; Smolke, C. D., Kinetic and equilibrium binding characterization of aptamers to small molecules using a label-free, sensitive, and scalable platform. Anal Chem 2014, 86 (7), 3273-8. 25. Feig, A. L., Applications of isothermal titration calorimetry in RNA biochemistry and biophysics. Biopolymers 2007, 87 (5-6), 293-301. 26. Polonschii, C.; David, S.; Tombelli, S.; Mascini, M.; Gheorghiu, M., A novel lowcost and easy to develop functionalization platform. Case study: aptamer-based detection of thrombin by surface plasmon resonance. Talanta 2010, 80 (5), 215764. 27. Liberman, J. A.; Wedekind, J. E., Riboswitch structure in the ligand-free state. Wiley Interdiscip Rev RNA 2012, 3 (3), 369-84. 28. Maiti, M.; Nauwelaerts, K.; Herdewijn, P., Pre-microRNA binding aminoglycosides and antitumor drugs as inhibitors of Dicer catalyzed microRNA processing. Bioorg Med Chem Lett 2012, 22 (4), 1709-11. 29. Shortridge, M. D.; Varani, G., Structure based approaches for targeting noncoding RNAs with small molecules. Curr Opin Struct Biol 2015, 30, 79-88. 30. Tran, T.; Childs-Disney, J. L.; Liu, B.; Guan, L.; Rzuczek, S.; Disney, M. D., Targeting the r(CGG) repeats that cause FXTAS with modularly assembled small molecules and oligonucleotides. ACS Chem Biol 2014, 9 (4), 904-12. 31. Heinz, T. F., Second-Order Nonlinear Optical Effects at Surfaces and Interfaces. Elsevier: Amsterdam, 1991. 32. Campen, R. K.; Zheng, D.-s.; Wang, H.-f.; Borguet, E., Second Harmonic Generation as a Probe of Multisite Adsorption at Solid−Liquid Interfaces of Aqueous Colloid Suspensions. The Journal of Physical Chemistry C 2007, 111 (25), 8805-8813. 33. Corn, R. M.; Higgins, D. A., Optical second harmonic generation as a probe of surface chemistry. Chemical Reviews 1994, 94 (1), 107-125. 34. Nguyen, T. T.; Sly, K. L.; Conboy, J. C., Comparison of the energetics of avidin, streptavidin, neutrAvidin, and anti-biotin antibody binding to biotinylated lipid bilayer examined by second-harmonic generation. Anal Chem 2012, 84 (1), 201-8. 35. Reeve, J. E.; Corbett, A. D.; Boczarow, I.; Wilson, T.; Bayley, H.; Anderson, H. L., Probing the orientational distribution of dyes in membranes through multiphoton microscopy. Biophys J 2012, 103 (5), 907-17. 36. Salafsky, J. S.; Eisenthal, K. B., Second harmonic spectroscopy: detection and orientation of molecules at a biomembrane interface. Chemical Physics Letters 2000, 319 (5-6), 435-439. 37. Svechkarev, D.; Kolodezny, D.; Mosquera-Vazquez, S.; Vauthey, E., Complementary surface second harmonic generation and molecular dynamics investigation of the orientation of organic dyes at a liquid/liquid interface. Langmuir 2014, 30 (46), 13869-76. 38. Zhuang, X.; Miranda, P. B.; Kim, D.; Shen, Y. R., Mapping molecular orientation and conformation at interfaces by surface nonlinear optics. Physical Review B 1999, 59 (19), 12632-12640. 39. Licari, G.; Brevet, P. F.; Vauthey, E., Fluorescent DNA probes at liquid/liquid interfaces studied by surface second harmonic generation. Phys Chem Chem Phys 2016, 18 (4), 2981-92.



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

40. Salafsky, J. S., 'SHG-labels' for detection of molecules by second harmonic generation. Chemical Physics Letters 2001, 342 (5-6), 485-491. 41. Moree, B.; Connell, K.; Mortensen, R. B.; Liu, C. T.; Benkovic, S. J.; Salafsky, J., Protein Conformational Changes Are Detected and Resolved Site Specifically by Second-Harmonic Generation. Biophys J 2015, 109 (4), 806-15. 42. Moree, B.; Yin, G.; Lazaro, D. F.; Munari, F.; Strohaker, T.; Giller, K.; Becker, S.; Outeiro, T. F.; Zweckstetter, M.; Salafsky, J., Small Molecules Detected by SecondHarmonic Generation Modulate the Conformation of Monomeric alpha-Synuclein and Reduce Its Aggregation in Cells. J Biol Chem 2015, 290 (46), 27582-93. 43. Salafsky, J. S., Detection of protein conformational change by optical secondharmonic generation. J Chem Phys 2006, 125 (7), 074701. 44. Salafsky, J. S., Second-harmonic generation for studying structural motion of biological molecules in real time and space. Phys Chem Chem Phys 2007, 9 (42), 5704-11. 45. Salafsky, J. S.; Cohen, B., A second-harmonic-active unnatural amino acid as a structural probe of biomolecules on surfaces. J Phys Chem B 2008, 112 (47), 151037. 46. Zimmermann, G. R.; Wick, C. L.; Shields, T. P.; Jenison, R. D.; Pardi, A., Molecular interactions and metal binding in the theophylline-binding core of an RNA aptamer. RNA 2000, 6 (5), 659-67. 47. Zimmermann, G. R.; Jenison, R. D.; Wick, C. L.; Simorre, J. P.; Pardi, A., Interlocking structural motifs mediate molecular discrimination by a theophyllinebinding RNA. Nat Struct Biol 1997, 4 (8), 644-9. 48. Macaya, R. F.; Schultze, P.; Smith, F. W.; Roe, J. A.; Feigon, J., Thrombin-binding DNA aptamer forms a unimolecular quadruplex structure in solution. Proc Natl Acad Sci U S A 1993, 90 (8), 3745-9. 49. Schultze, P.; Macaya, R. F.; Feigon, J., Three-dimensional solution structure of the thrombin-binding DNA aptamer d(GGTTGGTGTGGTTGG). J Mol Biol 1994, 235 (5), 1532-47. 50. Bugaut, A.; Balasubramanian, S., 5'-UTR RNA G-quadruplexes: translation regulation and targeting. Nucleic Acids Res 2012, 40 (11), 4727-41. 51. Jodoin, R.; Bauer, L.; Garant, J. M.; Mahdi Laaref, A.; Phaneuf, F.; Perreault, J. P., The folding of 5'-UTR human G-quadruplexes possessing a long central loop. RNA 2014, 20 (7), 1129-41. 52. Benevides, J. M.; Thomas, G. J., Jr., Local conformational changes induced in BDNA by ethidium intercalation. Biochemistry 2005, 44 (8), 2993-9. 53. Goodwin, K. D.; Long, E. C.; Georgiadis, M. M., A host-guest approach for determining drug-DNA interactions: an example using netropsin. Nucleic Acids Res 2005, 33 (13), 4106-16. 54. Nafisi, S.; Saboury, A. A.; Keramat, N.; Neault, J.-F.; Tajmir-Riahi, H.-A., Stability and structural features of DNA intercalation with ethidium bromide, acridine orange and methylene blue. Journal of Molecular Structure 2007, 827 (1–3), 35-43. 55. Neugebauer, U.; Schmid, U.; Baumann, K.; Simon, H.; Schmitt, M.; Popp, J., DNA tertiary structure and changes in DNA supercoiling upon interaction with ethidium


Page 20 of 22

Page 21 of 22

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


bromide and gyrase monitored by UV resonance Raman spectroscopy. Journal of Raman Spectroscopy 2007, 38 (10), 1246-1258. 56. Reinert, K. E., Adenosine-thymidine cluster-specific elongation and stiffening of DNA induced by the oligopeptide antibiotic netropsin. J Mol Biol 1972, 72 (3), 593-607. 57. Reinert, K. E., DNA stiffening and elongation caused by the binding of ethidium bromide. Biochim Biophys Acta 1973, 319 (2), 135-9. 58. Tse, W. C.; Boger, D. L., Sequence-selective DNA recognition: natural products and nature's lessons. Chem Biol 2004, 11 (12), 1607-17. 59. Xiao, Y.; Lubin, A. A.; Heeger, A. J.; Plaxco, K. W., Label-free electronic detection of thrombin in blood serum by using an aptamer-based sensor. Angew Chem Int Ed Engl 2005, 44 (34), 5456-9. 60. Pasternak, A.; Hernandez, F. J.; Rasmussen, L. M.; Vester, B.; Wengel, J., Improved thrombin binding aptamer by incorporation of a single unlocked nucleic acid monomer. Nucleic Acids Res 2011, 39 (3), 1155-64. 61. Li, B.; Wei, H.; Dong, S., Sensitive detection of protein by an aptamer-based label-free fluorescing molecular switch. Chem Commun (Camb) 2007, (1), 73-5. 62. Baraldi, P. G.; Bovero, A.; Fruttarolo, F.; Preti, D.; Tabrizi, M. A.; Pavani, M. G.; Romagnoli, R., DNA minor groove binders as potential antitumor and antimicrobial agents. Med Res Rev 2004, 24 (4), 475-528. 63. Chenoweth, D. M.; Dervan, P. B., Allosteric modulation of DNA by small molecules. Proc Natl Acad Sci U S A 2009, 106 (32), 13175-9. 64. Doss, R. M.; Marques, M. A.; Foister, S.; Chenoweth, D. M.; Dervan, P. B., Programmable oligomers for minor groove DNA recognition. J Am Chem Soc 2006, 128 (28), 9074-9. 65. Gearhart, M. D.; Dickinson, L.; Ehley, J.; Melander, C.; Dervan, P. B.; Wright, P. E.; Gottesfeld, J. M., Inhibition of DNA binding by human estrogen-related receptor 2 and estrogen receptor alpha with minor groove binding polyamides. Biochemistry 2005, 44 (11), 4196-203. 66. Nanjunda, R.; Wilson, W. D., Binding to the DNA minor groove by heterocyclic dications: from AT-specific monomers to GC recognition with dimers. Curr Protoc Nucleic Acid Chem 2012, Chapter 8, Unit8 8. 67. Raskatov, J. A.; Meier, J. L.; Puckett, J. W.; Yang, F.; Ramakrishnan, P.; Dervan, P. B., Modulation of NF-kappaB-dependent gene transcription using programmable DNA minor groove binders. Proc Natl Acad Sci U S A 2012, 109 (4), 1023-8. 68. Van Vliet, L. D.; Ellis, T.; Foley, P. J.; Liu, L.; Pfeffer, F. M.; Russell, R. A.; Warrener, R. N.; Hollfelder, F.; Waring, M. J., Molecular recognition of DNA by rigid [N]-polynorbornane-derived bifunctional intercalators: synthesis and evaluation of their binding properties. J Med Chem 2007, 50 (10), 2326-40. 69. Zimmer, C.; Wahnert, U., Nonintercalating DNA-binding ligands: specificity of the interaction and their use as tools in biophysical, biochemical and biological investigations of the genetic material. Prog Biophys Mol Biol 1986, 47 (1), 31-112.



For TOC only:

Ligand Ligand

Buffer

SHG

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Bilayer Glass Time


Detection of Ligand-Induced Conformational Changes in

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