Exploring the Motional Dynamics of End-Grafted DNA

As calculated from the HYTHER calculator (HYTHER version 1.0, Nicolas ..... the development of a diffusional model for the motion of the DNA borne Fc ...
0 downloads 0 Views 249KB Size
Download PDF
J. Phys. Chem. B 2007, 111, 6051-6058

6051

Exploring the Motional Dynamics of End-Grafted DNA Oligonucleotides by in Situ Electrochemical Atomic Force Microscopy Kang Wang, Ce´ dric Goyer, Agne` s Anne,* and Christophe Demaille* Laboratoire d’Electrochimie Mole´ culaire, Unite´ Mixte de Recherche UniVersite´ sCNRS No. 7591, UniVersite´ de Paris 7 - Denis Diderot, 2 place Jussieu, 75251 Paris Cedex 05, France ReceiVed: January 18, 2007; In Final Form: March 16, 2007

We introduce herein the use of atomic-force electrochemical microscopy (AFM-SECM) to simultaneously probe locally the conformation and motional dynamics of nanometer-sized single-stranded (ss) and doublestranded (ds) DNA oligonucleotides end-tethered to electrode surfaces. The ss-DNA system studied here consists of a low-density monolayer of (dT)20 oligonucleotides, 5′-thiol end-tethered onto a flat gold surface via a C6 alkyl linker and bearing at their free 3′-end a redox ferrocene label. It is shown that, as a result of the flexibility of the relatively long C6 linker, hinge motion, rather than elastic deformation of the DNA chain, is the major component of the dynamics of both the (dT)20 strand and its post-hybridized (dT-dA)20 duplex. DNA chain elasticity is nevertheless sufficiently contributing to the overall dynamics to result in ∼4 times slower dynamics for (dT-dA)20 than for (dT)20. Taking advantage of this dissimilar dynamical behavior of ss- and ds-DNA, it is demonstrated that hybridization can be easily locally detected at the scale of ∼200 molecules by AFM-SECM.

Introduction Understanding the behavior of DNA oligonucleotides endtethered to surfaces is of considerable importance owing to the fact that DNA-related biotechnologies, such as biosensing, rely entirely on the ability of end-attached DNA oligonucleotides (probes) to hybridize with target DNA strands in solution.1 Still, end-attachment of the DNA chain can significantly interfere with its molecular recognition capability.1a,2 In particular, interactions with the anchoring surface, for example, by adsorption of the DNA bases, can modify the conformation of end-attached DNA probes thus altering the biosensor performances.3 Therefore, being able to reliably assess the conformation adopted by endattached DNA chains is highly desirable. To this aim, a large amount of work has been devoted to the structural characterization of end-grafted DNA layers, using numerous techniques among which are optical methods,4 neutron reflectivity,5 atomic force6 or tunneling7 microscopy (AFM or STM), surface plasmon resonance (SPR),3c,8 surface force apparatus measurements (SFA),9 and X-ray photoelectron spectroscopy (XPS).10 Typically theses techniques provide a measurement of the overall thickness of the DNA layer. Since this thickness is usually found to be less than the full length of the DNA molecule, it is deduced that the end-grafted DNA strand is either coiled (for reputedly flexible single-stranded ssDNA chains) or tilted toward the surface (for more rigid double-stranded dsDNA chains). In this later case it is typically concluded that endgrafted DNA duplexes adopt a ∼45° orientation with respect to the grafting surface.5,6 Though this thickness measurementbased approach of DNA conformation yields useful results, it implicitly promotes the idea that end-grafted DNA strands adopt static conformations. However, DNA oligonucleotides are nanometer-sized objects and as such are necessarily animated by Brownian thermal motion. It should therefore be recalled that such measurements only yield time-averaged, and not actual, * Corresponding authors. E-mails: [email protected]; demaille@ paris7.jussieu.fr.

conformations of end-grafted DNA strands. This time averaging effect is explicitly taken into account in recent works using fluorescence based techniques, such as FRET,11 fluorescence self-interference12 or surface-quenching,13 which specifically address the position of the fluorophore-labeled free-end of grafted DNA oligonucleotides. These later works have the great merit of underlining the interplay between chain conformation and chain dynamics. Such fluorescence-based experiments were successfully used to characterize the average orientation of surface-bound oligonucleotides,13a and to monitor the kinetics of electric-field driven reorientation of grafted-DNA.13b,c However, to the best of our knowledge, neither these later techniques, nor any other, have allowed the underlying Brownian dynamics of short end-grafted DNA to be locally assessed. Yet, being able to quantify the dynamics of end-grafted DNA, in a local probe configuration, can be of particular importance to the active field of molecular electronics since STM investigations have revealed that conformational fluctuations of the DNA chain modulates the single-molecule conductance of surface-grafted oligonucleotides.14 In this context, we recently demonstrated15 that motional dynamics of nanometer-sized linear polymeric chains, anchored onto electrode surfaces, could be quantitatively probed by atomic-force electrochemical microscopy (AFM-SECM),16 a technique combining the capabilities of atomic force (AFM)17 and electrochemical microscopy (SECM),18 provided that the free end of the polymer chain is labeled to bear a redox moiety. The aim of the present work is thus to demonstrate the feasibility of using the unique capabilities of AFM-SECM to locally probe the conformation and motional dynamics of end-grafted, singlestranded DNA and its post-formed duplex, starting with a welldefined homopolymeric (dT)20 system. Experimental Methods Materials. Synthetic DNA oligonucleotides were purchased from Eurogentec (France/Belgium) as their sodium salts at a

10.1021/jp070432x CCC: $37.00 © 2007 American Chemical Society Published on Web 05/08/2007

6052 J. Phys. Chem. B, Vol. 111, No. 21, 2007 0.2-1 µmol scale and were Oligold purified grade products. The precursor sequence, HS-C6-5′-(dT)20-3′-NH2, used for chemical synthesis of 5′-C6-thiolated-3′-ferrocenylated oligonucleotide 3 (see below: Scheme 1) carries a (C7) aminolinker ) -(CH2)CHOHCH2O(CH2)3-NH2 at the 3′-end and a free thiol SH at the 5′-end. Note that protected forms of the free C6-thiol (such as C6-disulfide) were not available to us. All chemicals and solvents were analytical grade and used without further purification. Unless otherwise specified, all reactions were carried out in polypropylene tubes protected from light. All aqueous solutions were made with Milli-Q purified water (Millipore). Chromatography. Reversed-phase high-performance liquid chromatography (HPLC) was performed with a Gilson 305/306 HPLC pumps system equipped with a Gilson 119 UV-vis detector operating at 270 nm, using a Macherey-Nagel Nucleosil 120-5 C18 (4.6 mm ID × 15 cm) analytical column. Pump control and data processing used a Gilson Unipoint LC software (for Windows). The eluent conditions were as follows: solvent A, 10% acetonitrile in aqueous triethylammonium acetate, Et3NH+, AcO- (TEAA, 0.1 M) pH adjusted to 7.0 with acetic acid; solvent B, acetonitrile. Elutions were done in a three-step linear gradient: 5-19% B in 14 min, 19-50% B in 5 min, then isocratic. The flow rate was 0.8 mL/min. Spectrometries. Matrix-assisted laser desorption/ionization time-of-flight (MALDI-TOF) mass spectra were obtained from the Universite´ Pierre et Marie Curie, Laboratoire de Chimie Structurale Organique et Biologique (Paris, France) on a PerSeptive Biosystems Inc. (Framingham, MA) Voyager Elite instrument operating in the positive mode using 2,4,6-trihydroxyacetophenone in diammonium citrate as the matrix and 274 nm pulsed light. Internal calibration was carried out using the nucleotide sequence d(A11T7G5C5) (M 8604.6) in the above matrix. Syntheses. The two-step synthetic route we introduce herein for the preparation of the 3′-ferrocene-labeled oligonucleotide bearing a reactive 5′-SH thiol, 3, is shown in Scheme 1. This general procedure provides, in a first step, the simultaneous redox labeling of the 3′-end of the starting oligonucleotide and the protection of its 5′-free thiol under a disulfide form. More explicitly, the aerated aqueous mild basic conditions (pH 8) chosen for the ferrocene bioconjugation reaction were found to allow easy oxidation of the 5′-terminal thiol function. The thus obtained air-stable, bridged disulfide 3′-ferrocene-oligonucleotide dimer, 2, could be obtained as a pure compound after reverse-phase HPLC, and was stored before use for the TCEP cleavage step of the disulfide bond to 3. The ferrocene labeling reagent 1 (sulfo-N-hydroxysuccinimide of 3-ferrocenylpropanoic acid) was prepared as previously described.22b Bis(3′-Ferrocenylated-(dT)20-oligonucleotide-5′)-Bridged C6-Disulfide 2. A 20 mM solution of ferrocene sulfo NHS ester 1 in aqueous phosphate buffer (pH 8) containing 50% DMSO was prepared and filtered from residual urea through a Millipore 0.45 µm HV membrane before immediate use. In a typical experiment, approximately 30 OD260 units (∼0.18 µmol) of the sodium salt of 3′-amino-modified oligonucleotide, HS-C6-5′-(dT)20-3′-NH2 were dissolved in 380 µL of aerated phosphate buffer (pH 8.0). Then, 120 µL of the ferrocene sulfo NHS ester 1 solution (2.4 µmol) were added. The reaction tube was capped and the resulting mixture was allowed to react under stirring for 1.5 h. At this point, analytical HPLC on Nucleosil C18 showed the quantitative conversion of the starting oligonucleotide (tr ) 9.2 min) into the desired ferrocene-linked oligonucleotide dimer product 2 (tr ) 17.7 min), and a slower

Wang et al. SCHEME 1: Synthesis of Bis(3'-ferrocenylated-(dT)20 Oligonucleotide-5')-Bridged C6-Disulfide 2 and Its Subsequent Reductive Cleavage to 5'-C6-thiol 3'-ferrocenylated-(dT)20 Oligonucleotide 3a

a (a) Fc-NHS, oxidation conditions of the free thiol: aerated aq. phosphate, pH 8, r.t. 3 h, 30% isolated (HPLC purification). (b) TCEP: [tris(2-carboxyethyl)phosphine], Tris buffer, pH 7.0, r.t., 16 h, >90% (HPLC analysis).

CHART 1: Ferrocene Product 4 Formed during Preparation of 2, Resulting from the Coupling of Active Ester Coupling Agent 1 to Both 3′-NH2 and 5′-SH Groups of the Starting Oligonucleotide

eluting unwanted ferrocene product 4 (tr ) 20.5 min) (Figure 1S). As judged by comparison of HPLC peak areas (using the known molar extinction coefficient value 270 at 270 nm for (dT)20 (270 97800 M-1 cm-1) and for the ferrocene unit (270 of 6640 M-1 cm-1)), products 2 and 4 were formed in a ratio of approximately 1:1. The resulting mixture was directly purified by reversed phase HPLC. Appropriate fractions were dried by lyophilization and excess salt was removed by repeated lyophilization (×3) with deionized water, yielding the respective triethylammonium salt of product 2 and product 4. The identity and purity of products 2 and 4 were confirmed by mass spectrometry analyses. - Disulfide bis ferrocene oligonucleotide product 2: Average isolated yield 30%. MS (MALDI-TOF) (+) data: m/z (M+H)+, 13333.2, C452H602N82O292P42S2Fe2 requires 13332.8.

End-Grafted DNA Oligonucleotides - Unwanted ferrocene thiolester product 4 (Chart 1): MS (MALDI-TOF) (+) data: m/z (M+H)+, 6909.1, C239H314N41O147P21SFe2 requires 6907.5. Preparation of the Template-Stripped Gold Substrate Surface. The smooth template-stripped gold on mica substrates were prepared as described in the literature.19 Tapping-mode AFM characterization of the freshly stripped surface was then carried out using conventional AFM probes and only the surfaces having a peak-to-valley roughness of less than 1 nm were considered for assembly of the Fc-DNA layer. When required, evaluation of the effective area of the gold substrates was carried out as previously reported, by integration of the reduction peak of the gold oxide monolayer formed by cycling the substrate in a 0.5 M H2SO4 solution.15 Self-Assembly the Fc-(dT)20 Layer. In a typical procedure, reduction of the disulfide linkage of the 5-cystaminyl 20-mer oligonucleotide 2 (∼0.6 OD270 units, 3.8 nmol) was effected by treatment with tris(2-carboxyethyl)phosphine (TCEP, HCl) (30 nmol) in 8µL of 0.1M Tris buffer for 16h under argon. The thiol deprotected form 3 was purified from excess reagent by RP-HPLC, and eluted in ∼0.8 mL of a solution composed of ∼35% acetonitrile in 70 mM aqueous triethylammonium acetate, pH 7.0. The oligonucleotide solution was then concentrated by evaporation at room temperature under a flux of argon to a volume of ∼200 µL. The final, purely aqueous, assembly solution thus contained ∼ 30 µM 5′-thiol-terminated Fc-(dT)20 oligonucleotide in triethylammonium acetate, Et3NH+, AcO(TEAA, ∼0.35 M), pH 7.0. Self-assembly of the Fc-(dT)20 layer was carried out as described in the Results and Discussion section. Hybridization of the Fc-(dT)20 Surface. See Results and Discussion section. Dehybridization of the Fc-(dT-dA)20 Bearing Surface. Dehybridization of the end-grafted Fc-(dT-dA)20 strand was carried out by soaking the gold substrate in a 10 mM NaClO4 solution at room temperature for ∼12 h. This gentle dehybridization technique takes advantage of the lowering of the melting temperature of the DNA duplex at low Na+ concentration. As calculated from the HYTHER calculator (HYTHER version 1.0, Nicolas Peyret and John SantaLucia, Jr., Wayne State University),20 lowering the Na+ concentration from 1 M down to 10 mM is theoretically expected to lower the Tm of (dT-dA)20 in solution from ∼ 58 to ∼21 °C (for 0.25 µM (dT-dA)20) and thus to yield slow but complete dehybridization of the endgrafted Fc-dsDNA at room temperature. Comparison of the low scan rate cyclic voltammograms recorded before and after dehybridization revealed that no chain loss occurred during dehybridization. The intensity of the current approach curve recorded before hybridization of the Fc-(dT)20 surface was fully recovered upon dehybridization. Fabrication of the Combined AFM-SECM Probes (Tips). The combined AFM-SECM probes were fabricated and characterized as previously reported.21 Briefly a 60 µm gold microwire was bent at right angle ∼1 mm away from one of its ends while its other end was flattened between stainlesssteel plates to form a flexible cantilever arm. The other extremity of the wire was then etched electrochemically and its tip-end was locally melted, using a spark technique, to form a spherical electrode-tip ∼1 µm in diameter. The probe was then entirely insulated by deposition of electrophoretic paint, glued onto a standard AFM silicon chip and its spherical tip-end was selectively exposed by applying a high-voltage pulse. Calculation of the Spring Constant of the Combined Probes. The spring constant of the combined probe, kprobe, was

J. Phys. Chem. B, Vol. 111, No. 21, 2007 6053 estimated from the dimensions of the cantilever arm of the probe as described previously. 15a AFM-SECM Experiments. The AFM/SECM experiments were performed with a Molecular Imaging PICOSPM AFM microscope (Scientec, France) which was modified and operated as previously reported.15a The tip deflection was monitored via the reflection of the AFM laser onto the cantilever arm of the probe and the detection of the reflected beam on a position sensitive detector (PSD). The PSD was calibrated in situ using its linear response vs the piezo elongation. Eight different combined probes having similar tip sizes and spring constants were used in the experiments. For each experiment the force and current approach curves displayed the features discussed in the text, typical of either bare gold, Fc-ssDNA or Fc-dsDNA layers. Dozens of AFM-SECM approach curves were acquired at different locations on the substrate from which the experimental uncertainties on the values of the tip-substrate distance d, the tip deflection Ztip, and the tip current i were estimated to be respectively of (1 nm, (15%, (20%. Results and Discussion The DNA system studied here consists of a low-density monolayer of single-stranded oligonucleotides (dT)20, 3′-endlabeled to bear a low potential redox alkyl-ferrocene moiety (Fc) and end-grafted by their 5′-thiol end onto a flat templatestripped gold surface. We previously showed that this “model” redox-DNA system is extremely stable and suitable for the electrochemical characterization of the dynamics of end-grafted DNA.22 For the present work, the thiol functionality is linked to DNA by the commonly used, and rather long, C6 alkyl linker (See Figure 1a).

Figure 1. (a) Chemical structure of the Fc-(dT)20 oligonucleotide 5′C6-thiol end-grafted onto a gold substrate. (b) Solid lines: cyclic voltammogram at the gold substrate electrode bearing the Fc-(dT)20 chains, before hybridization (blue trace) and after hybridization by complementary unlabeled dA20 strands (red trace). Dotted line: background signal recorded after the Fc-ssDNA layer had been electrochemically stripped. The green and black traces are respectively, the raw and smoothed background-corrected signals recorded before hybridization. Fc-DNA surface concentration Γ ) 2.5 × 10-12 mol/ cm2. Scan rate V ) 2 V/s. Supporting electrolyte: 1 M NaClO4 + 25 mM sodium phosphate buffer pH 7.

Assembly and Characterization of the Fc-ssDNA Layer. Assembly of the Fc-(dT)20 layer on the flat template-stripped gold surface was carried out by reacting the freshly HPLCpurified 5′-thiol-terminated Fc-(dT)20 oligonucleotide with the

6054 J. Phys. Chem. B, Vol. 111, No. 21, 2007 gold substrate for about 16 h at ambient temperature in a deaerated solution protected from light (see Experimental methods). The Fc-(dT)20-modified-gold surface was then carefully washed with 1 M aqueous NaClO4. The ferrocene heads borne by the chains were electrochemically detected by cyclic voltammetry of the functionalized substrate. As seen in Figure 1b, at scan rate V < 10 V/s, a reversible faradaic signal is obtained whose features are as expected for an ideal Nernstian surface bound Fc-ssDNA:22 the peak heights are proportional to the scan rate and the peak-topeak separation is less than 10 mV. Such a voltammetric behavior insures that all the ferrocene heads are given ample time to reach the substrate and to reversibly exchange an electron with it.23 A value of E° ) 130 ( 10 mV/SCE was determined for the surface standard potential of the DNA-borne Fc/Fc+ redox couple from the average value of the anodic and cathodic peak potentials, this later value being in agreement with earlier reported values.22 Integration of the surface signal gave access to Γ, the surface coverage of grafted Fc heads, i.e., of DNA chains, each DNA chain bearing only one Fc head. From the typical value of Γ ≈ 2 × 10-12 mol/cm2 we obtained, the average distance between the anchoring points of the DNA strands can be calculated by ∼1/xNΓss ≈ 8 nm (N is the Avogadro number), which is close to the contour length of (dT)20 (Lss ≈ 10 nm).24 Consequently, it can be safely concluded on the basis of the above discussion that there is little lateral interaction between neighboring grafted strands for the present system.13b Moreover it should also be emphasized that, considering the high ionic strength of the 1M NaClO4 aqueous electrolyte solution used in the present study, reorientation of the end-anchored DNA, as a function of the electrode potential, should be negligible here.13b,d The homemade probe (tip) used for the AFM-SECM characterization of the DNA layer consists in a bent gold microwire, flattened to act as a flexible cantilever, and bearing a very smooth spherical tip-end of micrometer dimension, acting as a current-sensing microelectrode (see Experimental Methods). When, in a pH 7 buffered solution containing solely the 1M NaClO4 supporting electrolyte, such a combined probe, biased at a potential positive enough with respect to the standard potential of the ferrocene head is approached from a surface bearing Fc-DNA chains, biased negatively with respect to Fc head standard potential, the simultaneously acquired deflection and current approach curves shown in Figure 2 are obtained. The deflection and current approach curves can be delineated into 3 zones, as follows: in zone I the tip is far away from the surface, and no deflection or current is detected. In zone II the tip starts to interact with the DNA layer and the cantilever arm supporting the tip bends upward. Within this zone, the deflection curve recorded on the DNA surface differs significantly from the one recorded over a bare gold substrate (see lower inset in Figure 2a) and unambiguously reflects the compression of the DNA layer. Concomitantly a current begins to flow through the tip and increases while the tip is further pressed against the DNA layer, until a broad current peak is reached. Further compression of the Fc-DNA layer results in a sudden jump of current as zone III was reached, which is attributed to tunneling. At the same time the tip versus piezo movement plot becomes linear indicating that the tip has made hard contact with the substrate. When the tip is pulled away from the surface (blue curves), the then recorded retraction curves are similar to the approach curves, the observed offset being due to the piezo hysteresis. The first indication that the current approach curve recorded here is due to the tip electrochemically addressing the

Wang et al.

Figure 2. AFM-SECM approach curves. (a) Approach (red) and retraction (blue) deflection curves recorded with an AFM-SECM tip over a gold surface bearing end-grafted Fc-(dT)20 chains. The tip deflection Ztip, tracked by reflecting the alignment laser of the AFMsetup on the tip-arm, is plotted as a function of Zpiezo, the elongation of the piezo-tube supporting the tip. (b) Simultaneously recorded current approach and retraction curves. The green dotted line is the current approach curve recorded over a gold surface bearing unlabeled (dT)20 chains. In zone I the tip is too far away from the surface to interact with the Fc-DNA layer. In zone II the tip is compressing the layer. In zone III the tip has made hard contact with the underlying substrate surface. The tip and surface are respectively biased at potentials of +0.25 and -0.1 V vs SCE. Supporting electrolyte: 1 M NaClO4 + 25 mM sodium phosphate buffer pH 7. The upper inset in panel a depicts schematically the compression of the Fc-DNA layer by the combined tip, and shows graphically the definition of Ztip, Zpiezo, and the tip-tosubstrate distance d. The lower inset in panel a shows the typical deflection approach curve recorded upon approaching of a combined tip toward a bare gold substrate.

DNA-borne Fc heads is brought by the fact that, whenever unlabeled (dT)20 chains are grafted instead of Fc-DNA chains, although a deflection curve similar to the one presented in Figure 2a is recorded (data not shown), the current approach curve shows no current in zone II. Only the abrupt tunneling current rise of zone III, also recorded over a bare gold surface, is then recorded (see green dotted line in Figure 2b). The deflection approach curves were converted into force (F) curves using the value of the spring constant kprobe of the AFMSECM combined probe, determined as previously described (see Experimental Methods),15a since: F ) kprobe Ztip. Moreover, knowing the piezo elongation Zpiezo and the tip deflection Ztip, yields access to d, the tip-substrate distance by : d ) Ztip+Zpiezo. The approach curves can then be re-plotted as a function of d, as shown in Figure 3 (continuous blue curves). Compared to bare gold, a notable increase in both the force and current curves is observed below a tip-substrate separation of d ≈ 12 nm. The origin of the recorded tip-current can be fully characterized by using the positioning capabilities of the AFM apparatus

End-Grafted DNA Oligonucleotides

J. Phys. Chem. B, Vol. 111, No. 21, 2007 6055

Figure 3. AFM-SECM approach curves. The simultaneously recorded tip-deflection (a) and tip-current (b) approach curves are plotted vs the tip-to-substrate distance d for: a bare gold substrate (green line), a gold substrate bearing a 5′-end-grafted layer of Fc-(dT)20, before hybridization (blue trace) and after hybridization (red dashed line) by complementary unlabeled (dA)20 strands. Insets: (a) force curves for the Fc-DNA layer rescaled by dividing the actual d scale by the overall length of the ss or dsDNA strand (respectively Lss) 10 nm, Lds) 7 nm, 1 nm for the linker). (b) Log plot of the tip current vs d for the Fc-(dT)20 layer (upper blue curve) and the Fc-(dT-dA)20 layer (lower red curve). Fc-DNA surface coverage: ∼2.5 × 10-12 mol/cm2. Supporting electrolyte: 1M NaClO4 + 25 mM sodium phosphate buffer pH 7. Substrate potential, -0.1 V/SCE; tip bias, + 0.25 V/SCE. Approach rate: 10 nm/s. Estimated accuracy on the tip-substrate distance ≈ (1 nm. Averaging multiple curves ) 10.

Figure 4. Cyclic voltammetry within the Fc-DNA layer. The AFMSECM tip is positioned at a tip-to-substrate distance d from the FcssDNA bearing substrate and the substrate potential is scanned at 20 mV/s. The tip potential is held at -0.1 V/SCE. Supporting electrolyte: 1M NaClO4 + 25 mM sodium phosphate buffer pH 7.

to insert the tip within the nanometer thick Fc-DNA layer at a predefined distance d from the substrate surface, and cycling the tip potential while recording the tip current. However scanning the tip potential has the disadvantage of generating a nonspecific capacitive tip current. We therefore preferred an alternative albeit equivalent configuration which consists in scanning the substrate potential while maintaining the tip at a constant potential of -0.1 V/SCE. One can see from the resulting voltammogram presented in Figure 4a that subpicoamp current signals can then be detected. It is also seen that when the tip is positioned ∼20 nm away from the surface a flat and featureless background signal is recorded. But when the tip is positioned closer form the surface (e.g., at d ≈ 5 nm) the voltammogram develops a typical sigmoid shape. The S-shaped voltammogram is centered at a potential close to the ferrocene head standard potential thus demonstrating the specific electrochemical detection of the Fc head by the combined probe.15a We therefore assign the current approach curves recorded here to the occurrence of the back and forth movement of the Fc heads between the tip and substrate as schematized in Figure 5. The ferrocene heads are alternatively oxidized at the tip and reduced at the substrate thus giving rise to the tip current in a SECM positive feedback process.18 Provided that electron-

Figure 5. Depiction of the tip-to-substrate back and forth motion, and associated electron transfers, of the DNA-borne ferrocene Fc heads giving rise to the SECM positive feedback current. For clarity the tip is not on scale.

transfer steps are fast enough, which can be experimentally ascertained by biasing the tip and substrate at respectively positive and negative enough potentials, the intensity of the feedback current solely depends on the rate of the ferrocene head cycling motion, i.e., on the motional dynamics of the DNA chain. The first insight into the behavior of end-anchored DNA chains brought by the AFM-SECM technique comes from the analysis of the recorded force curves, which provide specific information regarding the conformation of the DNA strand. Like for any other end-grafted linear polymer chain, the conformation of (dT)20 is predicted to be governed by the ratio of its full contour length L over its persistence length, lp, which characterizes the intrinsic flexibility of the chain (the shorter lp the more flexible the chain).25 For L . lp the chain coils and adopts a mushroom conformation and the height of the DNA layer is then much less than L. At the opposite for L , lp the chain rather behaves as a rigid rod chain, and the height of the DNA layer tends toward L. Examination of the force approach curves reported in Figure 2a reveals that the tip starts to compress the oligonucleotide layer for a tip-substrate distance of ∼ 12 nm, which is consistent with the overall length of the 20-bases C6linked Fc-ssDNA (∼11 nm, Lss ≈ 10 nm for (dT)20,24 + 1 nm for the C6-linker). From this result, it may be inferred that the 5′-anchored (dT)20 strands are not lying flat on the surface, as often suspected for single-stranded DNA on bare gold,26 but

6056 J. Phys. Chem. B, Vol. 111, No. 21, 2007

Figure 6. Schematic depiction of the types of motion of the end anchored DNA rod allowing the tip-to-substrate travel of the Fc head. (a) Bending motion of the rod. (b) Rotational motion of the rod around its flexible anchoring point (C6 linker).

also that (dT)20 adopts a rod rather than a mushroom conformation. This indicates that the persistence length of (dT)20 is sufficiently close to its contour length for the end-attached chain to behave like a rod, a result consistent with the relatively high value of ∼ 3 nm estimated for the persistence length of dT homopolymers.24 However, the (dT)20 rod is not static but undergoes fast thermal motion and this motion, associated with the redox cycling of the Fc head, is generating the tip current recorded in the AFM-SECM experiment. Inspection of the tip current approach curve reported in Figure 3b (blue continuous line) reveals that the tip starts to electrochemically address the DNAborne Fc heads for a tip-to-substrate distance of d ≈ 10 nm. As d is made smaller, the recorded current increases, since the time required for the tip-to-substrate travel of the Fc head becomes shorter, until a current-peak is reached in the d ≈ 1-2 nm region. The occurrence of such a peak in the current approach curve, at very close tip-substrate separation, is indicative of lateral surface displacement and/or lateral elongation of the overcompressed Fc-(dT)20 chains.15b Two main components of the motional dynamics of (dT)20, allowing the tip-to-substrate motion of the Fc head, have to be considered in order to account for the recorded tip current. On the one hand we expect elastic bending fluctuations of the ssDNA rod (Figure 6a) but also, on the other hand, we expect fast rotational dynamics of the chain around its anchoring point, since it consists in a rather long and flexible C6 alkyl linker (Figure 6b). Taking advantage of the large difference in flexibility between ss and ds DNA (lp for dsDNA ≈ 50 nm),25b hybridization of the surface grafted ssDNA was used as a mean to decipher between these two types of chain motion. More precisely, by stiffening the Fc-DNA strand, hybridization is expected to decrease the contribution of bending elasticity to the overall DNA chain dynamics, thus allowing its assessment. Characterization of the Post-Hybridized Fc-DNA Layer. We previously showed that, within such a low-density grafted DNA system, the molecular recognition capability of the DNA chains was kept intact,22 so that the Fc-(dT)20 strands could be quantitatively hybridized by exposing the ssDNA-bearing surface to a ∼10 µM solution of the full complementary strand (dA)20 in 1 M NaClO4, 50 mM sodium phosphate buffer pH 7.0, for 1.5 h at 18 °C. The surface was then thoroughly washed with a 1M NaClO4 solution. The cyclic voltammogram of the hybridized surface, recorded at low scan rate, is very similar to the one recorded before hybridization (see Figure 1b), showing that no chain loss occurred during hybridization. The hybridization efficiency for our DNA system was estimated by high scan rate cyclic voltammetry at gold microelectrodes bearing a FcDNA layer. As described previously, at high scan rates (V > 1000 V/s) the contribution of ss and ds Fc-DNA chains to the voltammetric response can be differentiated and their relative

Wang et al. surface coverage estimated.22 We found than more than 99% (i.e., virtually all) of the DNA chains on the surface were hybridized. When the same AFM-SECM tip, used to characterize the Fc(dT)20 surface, is approached from the hybridized surface the then recorded force curve is qualitatively similar to the one obtained for ssDNA, as seen in Figure 3a (red dashed curve). The only noticeable difference is that the onset of the force approach curve is located ∼2 nm closer from the surface for dsDNA than for ssDNA. This reduction in range is compatible with the known reduction in DNA chain contour length upon hybridization (for a 20mer dT strand: Lss ) 10 nm, Lds ) 7 nm) (see above). Furthermore, when d is normalized by the estimated overall lengths of the anchored strands (Lss or ds + 1 nm for the C6 linker), the force approach curves recorded before and after hybridization fully coincide (see inset in Figure 3a). These results indicate that the nature of the force required to compress the ssDNA and dsDNA layers is not only the same but is also independent of the intrinsic elasticity of the strand. Considering the high ionic strength of the surrounding solution (1M), electric (double-layer) interactions between the tip and the DNA strands should only extent over a distance of ∼0.3 nm,27 and can consequently be excluded as the origin of the recorded force. The measured force is therefore attributed to the theoretically predicted,28 but rarely experimentally measured,9 steric (entropic) repulsive interaction between the endgrafted DNA strands and the incoming tip. This force opposes the confinement of the DNA chain within the tip-substrate gap which, by going against thermal motion of the strand, tends to decrease the entropy of the system. Compression by a spherical tip of a simple end-tethered rod, undergoing free hinge-motion around its anchoring point, is predicted to generate a steric force in the order of F ≈ 2πRtip ΓRT ≈ 0.2 nN (taking Rtip ≈ 500 nm and Γ ) 2.5 10-12 mol/cm2),28 which is comparable to the magnitude of the forces measured here. The above force curve analysis thus provides evidence that both ss and dsDNA are similarly animated by free rotational (hinge) motion around their C6 anchoring point rather than by elastic bending fluctuations, but yields no information regarding the dynamics of this motion. Motional dynamics of end-grafted chains can however be addressed by analyzing the experimental AFM-SECM current approach curve as detailed below. At the opposite of what was observed for the force curves, hybridization of the DNA layer resulted in a drastic change in the current approach curves (see Figure 2b): the intensity of the current approach curve for the Fc-DNA surface decreased spectacularly upon hybridization.29 However, as evidenced by the log plot presented in the inset of Figure 3b, the functional shape of the current approach curve remained the same. Considering the inherent sensitivity of SECM positive feedback approach curves to the geometry of the “diffusing” field of the current-generating redox species, this later result indicates that the motional path of the Fc head is similar whether it is borne by a ss- or a dsDNA chain.30 The measured decrease in intensity of the current approach curve upon hybridization therefore simply reflects the fact that the tip-to-substrate motional dynamics of the Fc head is faster for Fc-(dT)20 than for the Fc-(dT-dA)20 duplex. More quantitatively, we reproducibly observed that, upon hybridization, the intensity of the current approach curve for Fc-(dT)20 was divided by a typical extinction factor of ∼4 ( 1 (i.e., idsDNA(d) ) issDNA(d)/4, ∀ d). This value seems quite modest as compared to the very large difference in flexibility between ss and ds DNA, making unlikely that the tip-substrate motion of the Fc head is controlled by the elasticity

End-Grafted DNA Oligonucleotides of the DNA chain. Indeed if the dynamics of the DNA rod is modeled considering only the elastic bending of the chain, an extinction factor larger than 100 is theoretically predicted.31 At the other extreme, if only free rotation of the DNA chain around the C6 anchor is considered, the extinction factor should simply correspond to the ratio of the rotational diffusion coefficents of dT20 and -(dT-dA)20, which is not expected to differ much from unity.32 These results quantitatively support the notion that rotational (hinge) motion of both ss and ds DNA is the major component of the dynamics of the Fc head cycling motion, i.e., of the thermal motion of end-anchored (dT)20 and (dT-dA)20. However, the observed slower dynamics for (dT-dA)20 indicates that the contribution of DNA chain elasticity to the Fc-head motional dynamics is small but nevertheless sufficient to allow the dynamical behavior of ss and dsDNA to be discriminated by AFM-SECM current feedback measurements. A more detailed quantitative analysis of the current approach curves would require the development of a diffusional model for the motion of the DNA borne Fc head, which is beyond the scope of the present paper, but is expected to shine light on the Brownian dynamics of end-attached DNA strands under nanometric confinement.31 Another interesting aspect of the interaction between the (ss and ds) DNA layers and the tip is revealed by comparing the respective ranges of the simultaneously recorded force and current approach curves reported in Figure 3. It can be seen that the tip-to-substrate distance at which the incoming tip starts to sense a repulsive force is consistently 1-2 nm greater than both the contour length of the DNA strand and the distance corresponding to the onset of tip current (d ≈ 10 nm). The absence of tip current for d > 10 nm indicates that, beyond this distance, the DNA rod is not able to physically contact the tip and therefore that the then detected force acts through the solvent. A likely candidate for this force is the so-called hydration force which is a solvation shell-mediated repulsive interaction occurring between charged surfaces.27b Such hydration forces were previously observed between DNA strands,33a proteins33b or charged membrane surfaces33c separated by less than ∼2 nm. For d e 10 nm, the simple fact that an electrochemical current is recorded shows that the Fc head is then contacting the tip and therefore that the DNA rod is sterically interacting with the tip. It is noteworthy that being able to distinguish between “contact” and through-solvent forces in such a way is a specific benefit of the combined AFM-SECM configuration. The maximum number of DNA chains addressed by the AFM-SECM tip, nc, is worth estimating. It can be evaluated using simple geometric considerations: taking for the tip radius Rtip ) 500 nm and for the overall length of the Fc-DNA strand a value of L ≈ 10 nm, yields nc ≈ πΓN Rtip L ≈ 200 DNA chains. This number could be easily reduced to a few tens of chains by decreasing the tip size and/or using lower Fc-DNA surface concentration, at the expense of the intensity of the recorded approach curves. The above geometric reasoning also yields the conclusion that the tip actually probes a disk-shaped area on the substrate which is, at most ∼xRtipL ) 70 nm in radius. The fact that the force and current approach curves recorded at different locations on the substrate were found to be reproducible in intensity within ( 20% tend to demonstrate that the DNA chains were homogeneously distributed on the surface, at least at the above lateral scale.

J. Phys. Chem. B, Vol. 111, No. 21, 2007 6057 Conclusion In conclusion, we have introduced the use of atomic force electrochemical microscopy (AFM-SECM) to simultaneously locally probe the conformation and motional dynamics of short DNA strands end-anchored onto surfaces. The presented results lead to the conclusion that thermally activated hinge motion is the prevailing component of the dynamics of nanometer-sized DNA chains grafted onto surfaces via long C6 spacers. This result could have been expected for the relatively rigid doublestranded (dT-dA)20 chain but is shown to also apply to (dT)20, even though single-stranded DNA is usually thought-of as a relatively flexible chain. As a result elastic bending dynamics of the DNA chains cannot be accessed since it is masked by the flexibility of the long C6 anchor. This situation contrasts to what was previously observed by high scan rate cyclic voltammetry for redox-dsDNA chains end-anchored to electrodes via a short C2 linker, since in the later case chain bending was shown to control the dynamics of the end-grafted strands.22 This difference most probably arises from the fact that, because of a stronger steric hindrance between the lowest part of the DNA rod and the surface, hinge motion is largely hindered for DNA chains anchored via short-linkers. As a result DNA chain deformability plays a much larger role in the overall dynamics of shortly linked surface DNA systems. Such a result emphasizes the decisive role of the flexibility (and length) of the anchoring linker in the dynamic behavior of end-grafted DNA oligonucleotides. This observation may be relevant to the design of electrochemical DNA sensors which exploit the dissimilar physical behavior of ss and ds DNA oligonucleotide monolayers end-tethered to electrode surfaces to detect hybridization.1 The here-reported results also point to the fact that global motion of anchored DNA chains is a very efficient electron transport mechanism in redox DNA-films and should definitely be taken into account no matter the details of the DNA-based system. Finally, it was also demonstrated here that hybridization of nanometer sized end-grafted DNA strands can be easily detected at the scale of ∼200 molecules by AFM-SECM, as a benefit of the high sensitivity of the tip current to the hybridization event. Interestingly, force measurement is, at the opposite, almost insensitive to hybridization, a feature due to the steric nature of the tip-DNA interactions involved here. This result could be applied to the design of an electrochemical DNA hybridization sensor based on the same working principle but implemented in a more robust nanogap configuration. Acknowledgment. Kang Wang was supported by a Research Grant from the City of Paris (France) for hosting foreign researchers. References and Notes (1) (a) Tarlov, M. J.; Steel, A. B. In Biomolecular Films; Rustling, J. F., Ed.; Marcel Dekker Inc.: New-York, 2003; Vol. 111. (b) Heller, M. J. Annu. ReV. Biomed. Eng. 2002, 4, 129-153. (c) Thorp, H. H. Trends Biotechnol. 2003, 21, 522-524. (d) Drummond, T. G.; Hill, M. G.; Barton, J. K. Nat. Biotechnol. 2003, 21, 1192-1199. (e) Gooding, J. J.; King, G. C. J. Mater. Chem. 2005, 15, 4876-4880. (2) Peterson, A. W.; Wolf, L. K.; Georgiadis, R. M. J. Am. Chem. Soc. 2002, 142, 14601-14607. (3) (a) Storhoff, J. J.; Elghanian, R.; Mirkin, C. A.; Letsinger, R. L. Langmuir 2002, 18, 6666-6670. (b) Kimura-Suda, H.; Petrovykh, D. Y.; Tarlov, M. J.; Whitman, L. J. J. Am. Chem. Soc. 2003, 125, 9014-9015. (c) Wolf, L. K.; Gao, Y.; Georgiadis, R. M. Langmuir 2004, 20, 33573360. (4) Gray, D. E.; Case-Green, S. C.; Fell, T. S.; Dobson, P. J.; Southern, E. M. Langmuir 1997, 13, 2833-2842.

6058 J. Phys. Chem. B, Vol. 111, No. 21, 2007 (5) Levicky, R.; Herne, T. M.; Tarlov, M. J.; Satija, S. K. J. Am. Chem. Soc. 1998, 120, 9787-9792. (6) (a) Kelley, S. O.; Barton, J. K.; Jackson, N. M.; McPherson, L. D.; Potter, A. B.; Spain, E. M.; Allen, M. J.; Hill, M. G. Langmuir 1998, 24, 6781-6784. (b) Dong, L. Q.; Zhou, J. Z.; Wu, L. L.; Dong, P.; Lin, Z. H. Chem. Phys. Lett. 2002, 354, 458-465. (c) Zhang, Z-L.; Pang, D.-W.; Zhang, R.-Y.; Yan, J.-W.; Mao, B.-W.; Qi, Y.-P. Bioconjugate Chem. 2002, 13, 104-109. (7) (a) Zhou, D.; Sinniah, K.; Abell, C.; Rayment, T. Langmuir 2002, 18, 8278-8281. (b) Patole, S. N.; Pike, A. R.; Connolly, B. A.; Horrocks, B. R.; Houlton, A. Langmuir 2003, 19, 5457-5463. (c) Wackerbarth, H.; Grubb, M.; Zhang, J.; Hansen, A. G.; Ulstrup, J. Langmuir 2004, 20, 16471655. (8) Peterlinz, K. A.; Georgiadis R. M.; Herne, T. M.; Tarlov, M. J. J. Am. Chem. Soc. 1997, 3401-3402. (9) Cho, Y.-K.; Kim, S.; Lim, G.; Granick, S. Langmuir 2001, 17, 7732-7734. (10) (a) Herne, T. M.; Tarlov, M. J. J. Am. Chem. Soc. 1997, 119, 89168920. (b) Petrovykh, D. Y.; Kimura-Suda, H.; Tarlov, M. J.; Whitman, L. J. Langmuir 2004, 20, 429-440. (c) Petrovykh, D. Y.; Pe´rez-Dieste, V.; Opdahl, A.; Kimura-Suda, Sullivan, J. M. H.; Tarlov, M. J.; Himpsel, F. J.; Whitman, L. J. J. Am. Chem. Soc. 2006, 128, 2-3. (11) (a) Charreyre, M.-T.; Tcherkasskaya, O.; Winnik, M. A.; Hiver, A.; Delair, T.; Cros, P.; Pichot, C.; Mandrand, B. Langmuir 1997, 13, 31033110. (b) Murphy, M. C.; Rasnkic, I.; Cheng, W.; Lohman, T. M.; Ha, T. Biophys. J. 2004, 86, 2530-2537. (12) Moiseev, L.; U ¨ nlun, M. S.; Swan, A. K.; Goldberg, B. B.; Cantor, C. R. Proc. Natl. Acad. Sci. U.S.A. 2006, 103, 2623-2628. (13) (a) Rant, U.; Arinaga, K.; Fujita, S.; Yokoyama, N.; Abstreiter, G.; Tornow, M. Langmuir 2004, 20, 10086-10092. (b) Rant, U.; Arinaga, K.; Fujita, S.; Yokoyama, N.; Abstreiter, G.; Tornow, M. Nano Lett. 2004, 4, 2441-2445. (c) Rant, U.; Arinaga, K.; Tornow, M.; Kim, Y. W.; Netz, R. R.; Fujita, S.; Yokoyama, N.; Abstreiter, G. Biophys. J. 2006, 90, 36663671. (d) Rant, U.; Arinaga, K.; Fujita, S.; Yokoyama, N.; Abstreiter, G.; Tornow, M. Org. Biomol. Chem. 2006, 4, 3448-3455. (e) Sendner, C.; Kim, Y. W.; Rant, U.; Arinaga, K.; Tornow, M.; Netz, R. R. phsy. Stat. Sol. a 2006, 14, 3476-3491. (14) van Zalinge, H.; Schiffrin, D. J.; Bates, A. D.; Haiss, W.; Ulstrup, J.; Nichols, R. J. ChemPhysChem 2006, 7, 94 -98. (15) (a) Abbou, A.; Anne, A.; Demaille, C. J. Am. Chem. Soc. 2004, 126, 10095-10108. (b) Abbou, J.; Anne, A.; Demaille, C. J. Phys. Chem. B 2006, 110, 22664-22675. (16) (a) Macpherson, J. V.; Unwin, P. R.; Hillier, A. C.; Bard, A. J. J. Am. Chem. Soc 1996, 118, 6445-6452. (b) Macpherson, J. V.; Unwin, P. R. Anal. Chem. 2000, 72, 276-285. (c) Macpherson, J. V.; Webb, M. A.; Unwin, P. R. Anal. Chem. 2001, 73, 550-557. (d) Kranz, C.; Friedbacher, G.; Mizaikoff, B.; Lugstein, A.; Smoliner, J.; Bertagnolli, E. Anal. Chem. 2001, 73, 2491-2500. (e) Kranz, C.; Kueng, A.; Mizaikoff, B.; Lugstein, A.; Bertagnolli, E. Angew. Chem., Int. Ed. 2003, 42, 3238-3240. (17) Binnig, G.; Quate, C. F.; Gerber, C. Phys. ReV. Lett. 1986, 56, 930-933.

Wang et al. (18) In Scanning Electrochemical Microscopy; Bard, A. J., Mirkin, M. V., Eds.; Marcel Dekker: New York, 2001. (19) (a) Hegner, M.; Wagner, P.; Semenza, G. Surf. Sci. 1993, 291, 3946. (b) Wagner, P.; Hegner, M.; Gu¨ntherodt, H.-J.; Semenza, G. Langmuir 1995, 11, 3867-3875. (20) (a) SantaLucia, J. Proc. Natl. Acad. Sci. U.S.A. 1998, 95, 14601465. (b) Peyret, N.; Seneviratne, P. A.; Allawi, H. T.; SantaLucia, J. Biochemistry 1999, 38, 3468-3477. (21) Abbou, J.; Demaille, C.; Druet, M.; Moiroux, J. Anal. Chem. 2002, 74, 6355-6363. (22) (a) Anne, A.; Bouchardon, A.; Moiroux, J. J. Am. Chem. Soc. 2003, 125, 1112-1113. (b) Anne, A.; Demaille, C. J. Am. Chem. Soc. 2006, 128, 542-557. (23) Laviron, E. In Electroanalytical Chemistry; Bard, A. J., Ed.; Marcel Dekker: New York, 1982; Vol. 12, pp 53-157. (24) Mills, J. B.; Vacano, E.; Hagerman, P. J. J. Mol. Biol. 1999, 285, 245-257. (25) (a) Grosberg, A. Y.; Khokhlov, A. R. Statistical Physics of Macromolecules; American Institute of Physics: New York, 1994. (b) Bloomfield, V. A.; Crothers, D. M.; Tinoco, I. Nucleic Acids: Structures, Properties and Functions; University Science Books: Sausalito, CA, 2000. (26) This results falls in line with the low affinity of (dT) for gold (ref 3). (27) (a) Israelachvili, J. Intermolecular and Surface Forces, 2nd ed.; Academic Press: San Diego, 1992; p 238. (b) Israelachvili, J. Intermolecular and Surface Forces, 2nd ed.; Academic Press: San Diego, 1992; p 275. (28) Miller, I. C. B.; Williams, D. R. M. Phys. ReV. E 2000, 61, R4706R4709. (29) The intensity of the current approach curve recorded before hybridization of the Fc-dT20 surface was fully recovered upon dehybridization of the end-grafted Fc-(dTdA)20 strand, carried out under mild conditions (see Experimental Methods). This result evidences that the intensity of the approach curve is specifically (and reversibly) modulated by the hybridization/dehybridation of the anchored Fc-DNA chain. (30) As an illustration, for a redox species freely diffusing in solution, the curvature of SECM positive feedback approach curves was shown to sharply depend on the tip geometry, i.e., on the geometry of the diffusion field (path) around the microelectrode. See: (a) Davis, J.; Fan, F-R. F.; Bard, A. J. J. Electroanal. Chem. 1987, 238, 9-31. (b) Mirkin, M. V.; Fan, F-R. F.; Bard, A. J. J. Electroanal. Chem. 1992, 328, 47-62. (31) Wang, K.; Anne, A.; Demaille C. in press. (32) Ortega, A.; Garcı´a de la Torre, J. J. Chem. Phys. 2003, 119, 99149919. (33) (a) Rau, D. C.; Lee, B.; Parsegian, V. A. Proc. Natl. Acad. Sci. U.S.A. 1984, 81, 2621-2625. (b) Valle-Delgado, J. J.; Molina-Bolı´var. J. A.; Galisteao-Gonza´lez, F.; Ga´lvez-Ruiz, M. J.; Feiler, A.; Rutland, M. W. J. Phys. Chem. B 2004, 108, 5365-5371. (c) LeNeveu, D. M.; Rand, R. P.; Pargesian, V. A. Nature (London) 1976, 259, 601-603.