Open Access

Nanomechanical characterization of chemical interaction between gold nanoparticles and chemical functional groups

  • Gyudo Lee1, 2,
  • Hyungbeen Lee2,
  • Kihwan Nam1, 2,
  • Jae-Hee Han3,
  • Jaemoon Yang4,
  • Sang Woo Lee2,
  • Dae Sung Yoon2,
  • Kilho Eom1, 2 and
  • Taeyun Kwon1, 2Email author
Nanoscale Research Letters20127:608

DOI: 10.1186/1556-276X-7-608

Received: 15 September 2012

Accepted: 19 October 2012

Published: 31 October 2012


We report on how to quantify the binding affinity between a nanoparticle and chemical functional group using various experimental methods such as cantilever assay, PeakForce quantitative nanomechanical property mapping, and lateral force microscopy. For the immobilization of Au nanoparticles (AuNPs) onto a microscale silicon substrate, we have considered two different chemical functional molecules of amine and catecholamine (here, dopamine was used). It is found that catecholamine-modified surface is more effective for the functionalization of AuNPs onto the surface than the amine-modified surface, which has been shown from our various experiments. The dimensionless parameter (i.e., ratio of binding affinity) introduced in this work from such experiments is useful in quantitatively depicting such binding affinity, indicating that the binding affinity and stability between AuNPs and catecholamine is approximately 1.5 times stronger than that between amine and AuNPs. Our study sheds light on the experiment-based quantitative characterization of the binding affinity between nanomaterial and chemical groups, which will eventually provide an insight into how to effectively design the functional material using chemical groups.


Au nanoparticle Dopamine Surface chemistry Atomic force microscopy Lateral force microscopy


Surface chemistry has played a critical role in designing functional nanomaterials for their biological or medical applications such as drug delivery, molecular therapeutics, and diagnostics [1, 2]. In particular, the surface modification of a nanoparticle is of great importance to enhancing functionality in terms of target affinity [35], imaging contrast [3, 4, 6, 7], and curative power [8]. For instance, magnetic nanoparticles chemically modified with chemical functional groups or moieties (e.g., ligand and receptor) have been utilized for high-resolution MRI, which is useful in cancer diagnostics since the chemical modification using chemical functional groups or moieties leads to improved targetability and imaging contrasts [3, 6, 7]. Moreover, gold nanoparticles (AuNPs) functionalized with chemical functional groups or moieties have been recently used to enhance photocatalytic performance [9], to form 3D networks of functionalized AuNPs [10], and to sensitively detect specific biological molecules (e.g., DNA) [1113] and cancerous single cells [14, 15].

Dopamine hydrochloride (DOPA) has recently been considered as a chemical linker that allows for efficient surface chemistry useful in not only inorganic materials (e.g., nanoparticles) but also biological materials (e.g., tissue) due to its excellent adhesive property and biocompatibility [16, 17]. In particular, DOPA has been reported as a chemical linker that is useful not only in the chemical modification of the surfaces of nanomaterials such as nanoparticles [18, 19], graphene oxide sheet [20], and carbon nanotubes [21], but also in improving binding affinities such as protein-peptide cross-linking [22], cellular adhesion to substrate [23], osteoconduction [24], and hemostatic adhesive in segmentectomy [25]. Despite the broad application of DOPA to surface chemistry using mutual interaction between DOPA and nanomaterials (e.g., nanoparticle), such an interaction has been poorly understood and not yet studied thoroughly. Since the surface modification of nanomaterials using DOPA typically employs a noncovalent conjugation (e.g., coordinate bonding, hydrophobic and electrostatic interactions, etc.) [6, 26], it is essential to establish an experimental framework that allows for measuring a weak binding affinity corresponding to such a noncovalent conjugation, which is useful in the development of drug carrier due to the fact that noncovalent conjugation enables the excretion of waster matter from the human body after the drug carrier completes the function of drug delivery or bioimaging [6, 7, 27, 28].

In this work, we have quantitatively studied a chemical interaction between nanoparticles and chemical functional groups (e.g., DOPA and amine functional group) using experimental toolkits such as cantilever bioassay [2932], PeakForce Quantitative Nanomechanical Property Mapping (PeakForce QNM) [33, 34], as well as lateral force microscopy (LFM) [3538]. In a recent decade, cantilever bioassays have been widely utilized for quantitative understanding of molecular interactions on the surface by measuring the bending deflection change [39, 40] and/or shifts in resonance [29, 41]. Moreover, a cantilever has been also employed to measure physical quantities such as temperature [42], quantum state [43], and surface stress [29, 44]. We have shown that a cantilever whose surface is functionalized with specific chemical functional groups (DOPA or amine functional group) allows us to quantitatively characterize the binding affinity between nanoparticles and such chemical functional groups. Furthermore, LFM has recently been taken into account for deciphering the molecular interactions by estimating a frictional force that occurs due to breakage of such molecular interactions [35, 38]. In our study, we have employed LFM enabling the movement of a nanoparticle, which is chemically interacting with chemical functional groups on the surface, in order to quantitatively understand the binding affinity between nanoparticle and chemical functional groups by measuring the frictional forces required to break the binding between the nanoparticle and chemical functional groups. In addition, we have also measured the adhesion force between nanoparticles and chemical functional groups using atomic force microscopy (AFM), particularly the PeakForce QNM module. We have shown that the noncovalent interaction between nanoparticles and specific chemical functional groups can be quantitatively studied using the aforementioned experimental techniques (i.e., cantilever assay, PeakForce QNM, and LFM) and that catecholamine (i.e., DOPA) is a chemical functional group useful in the surface modification of nanomaterials (e.g., nanoparticle) due to its excellent binding affinity.


Materials and sample preparation

All materials including gold nanoparticle (G1652, approximately 20 nm in size) and dopamine hydrochloride ((HO)2C6H3CH2NH2·HCl) were purchased from Sigma-Aldrich (St. Louis, MO, USA). A silicon (Si) microcantilever (TESP, Bruker, Madison, WI, USA) was first rinsed by piranha solution (50% of sulfuric acid and 50% of hydrogen peroxide). The cantilever was immersed for 25 min into a 3-aminopropyltrimethoxysilane (APTMS) solution (200 μl/ethanol of 5 ml) for amine functionalization and then carefully washed by ethanol and pure water. The aminated surface of the cantilever (SA) was immersed into the AuNP suspension (approximately 0.01% as HAuCl4) for 30 min for the preparation of AuNP-SA (i.e., AuNP attached to amine-modified surface). In the case of DOPA-functionalized surface (SD), the aminated microcantilever was treated with glutaraldehyde (GA, 10% in phosphate-buffered saline (PBS)) for 30 min for surface activation and then immersed into the DOPA solution (65 mM in PBS at pH 7.4) for 10 h [16]. Consequently, the DOPA-functionalized cantilever was immersed into the AuNP-dissolved solution for the preparation of AuNP-SD (i.e., AuNP bound to DOPA-functionalized surface). All experiment was conducted at room temperature.

Analysis of surface chemistry

Scanning electron microscopy (SEM) imaging was obtained using JSM-6500 F (JEOL, Tokyo, Japan). The number of AuNPs in the SEM images was accurately counted by ImageJ software (NIH, Bethesda, MD, USA). X-ray photoelectron spectroscopy (XPS) analysis was implemented with Escalab 220i-XL (Thermo VG, Hastings, UK). The sampling area was 5 mm × 5 mm in a vacuum of 1.0 × 10−9 mbar with calibration of C 1 s (285 eV). To measure the resonant frequency shift of the cantilever due to AuNP binding onto the cantilever surface, the samples were dried overnight in each fabrication process. The resonant frequency of the cantilever is measured using the Nanoscope V controller (Veeco, Santa Barbara, CA, USA).

Measurement of adhesion/friction forces

PeakForce QNM was used to measure the adhesion between AuNPs and chemically functionalized surface using the BioScope Catalyst (Veeco). For PeakForce QNM imaging, we have used a cantilever, particularly ScanAsyst Air probes (kN = 0.58 N/m; Bruker) in 22.2°C and 38% humidity. For LFM imaging, we have employed various AFM cantilever tips (i.e., SNL-10, ScanAsyst Air, ScanAsyst Fluid, Bruker) with their stiffness in the range of 0.1 to 1 N/m. LFM images were obtained by scanning the sample in contact mode with a scan size of 2 × 2 μm2, scan rate of 0.5 Hz, and a set point of 1 V. The detached AuNPs from the surface was confirmed by using PeakForce QNM imaging. All AFM, LFM, and PeakForce QNM images were analyzed with NanoScope Analysis software (Bruker).

We have prepared the silicon surface onto which the AuNPs were attached using chemical functional group (i.e., amine or DOPA), as shown in Figure 1, in order to study the chemical interaction between the nanoparticle and chemical functional group. In particular, we have studied such chemical interaction using various experimental tools such as cantilever assay, LFM, and PeakForce QNM as described above.
Figure 1

AuNPs attached onto silicon substrate using chemical functional group. Schematic illustration of the AuNP-coating procedure on the Si substrate and characterization regimes for the chemical interaction between AuNPs and chemical functional groups. (I) AuNPs immobilized on aminated surface (AuNP-SA); (II) AuNPs immobilized on catecholic surface (AuNP-SD) functionalized with dopamine (DOPA). Each sample is quantitatively characterized by experimental toolkits such as cantilever bioassay, PeakForce QNM, as well as lateral force microscopy (LFM).

Results and discussion

Characterization of the AuNPs and surface chemistry of SAand SD

The size distribution of AuNPs that were used in our experiment was obtained based on transmission electron microscopy (TEM) images as shown in Figure 2a. In particular, the mean diameter of the AuNPs is given as 19.4 nm, and the standard deviation is estimated as 2.2 nm. Moreover, we have confirmed the surface chemistry of the substrate, i.e., chemical functional groups formed on a silicon surface, by using XPS analysis. Figure 2b,c shows the XPS survey of the formation of -NH2-Si (panel I in Figure 1) and DOPA-Si (panel II in Figure 1). The N 1 s signal appeared in the survey spectrum as the -NH2 monolayer was being fabricated on the Si substrate. In the following step, the DOPA was chemically adsorbed onto the amine-functionalized surface and polymerized to form the dopamine layer [45]. The XPS survey shows N 1 s and C 1 s peaks (Figure 2c) that are higher than those of the -NH2-Si sample (Figure 2b). Also, Si 2 s and Si 2p peaks disappeared in the DOPA-Si sample because of the thickness of the DOPA film, implying the well-formed DOPA layer with self-assembled monolayer [4547].
Figure 2

Characteristics of the AuNPs and surface chemistry of S A and S D . Histogram of AuNP diameters (a) and XPS survey spectra of (b) the aminated Si substrate (SA) and (c) DOPA-functionalized Si substrate (SD). Inset in (a): TEM images of the AuNPs.

Indirect measurement of the binding affinity between AuNPs and chemical functional groups

We have investigated the binding affinity between AuNPs and the silicon surface chemically modified with a functional group (i.e., amine or DOPA) by measuring the number of AuNPs attached on the chemically modified silicon surface. Here, the DOPA-modified surface is denoted as SD, whereas we denote the amine-functionalized surface as SA. It should be also noted that the AuNPs attached to SD exhibit uniform shape and size, as shown in Figure 3a,b,c. It is interestingly shown that the AuNPs attached to SA were locally aggregated, while the AuNPs immobilized on SD were distributed with relatively high uniformity (Figure 3d,e), which suggests that the AuNPs were functionalized as a uniform monolayer onto SD. It is attributed to the fact that the local aggregation of AuNPs onto SA is highly related to the molecular structure of APTMS, which leads to the formation of amine functional group as a disordered layer on the surface and, consequently, the decrease in the uniformity of surface functionalization and large variation of surface density of AuNPs [48]. On the other hand, a uniform attachment of AuNPs onto SD is attributed to GA acting as a linker molecule between the surface and DOPA such that the linker molecule allows for an ordered formation of DOPA on a silicon surface. The uniform distribution of the attached AuNPs on the surface is due to the electrostatic repulsion between nanoparticles. Meanwhile, the uniformity of the functionalized molecules is a critical factor in determining the binding affinity of AuNPs [48] because the uniform distribution of functional molecules is a priori requisite to optimize the binding affinity between the surface and AuNPs. Although there are small aggregates of AuNPs locally even in the AuNP-SD samples, the binding affinity between the surface and AuNPs is clearly shown in the electron microscope imaging assay. The number of AuNPs attached onto either SA or SD (denoted as NA or ND, respectively) can be used as a quantity that represents the binding affinity. Based on the SEM images of AuNPs attached to either SA or SD, it is found that NA = 503 ± 54 (mean ± standard deviation) per unit area of 1 μm2, whereas ND = 798 ± 75 per unit area of 1 μm2 (Figure 3f). This clearly elucidates that SD exhibits higher binding affinity to AuNPs than SA. For quantitative comparison, we have introduced a dimensionless measure defined as RN= ND/NA. This RN ratio can be used as a dimensionless quantity useful in representing the binding affinity (for more detail, see below).
Figure 3

Characterization of the surface chemistry of the samples such as AuNP- S A and AuNP- S D . (a, b) SEM images of the microcantilever functionalized with DOPA molecules used in the cantilever bioassay. The dimensions of the microcantilever such as stylus height, cantilever length, and width are shown. (c) The magnified image of the stylus vertex shows uniformly coated AuNPs on the entire microcantilever. (d, e) SEM images of the AuNPs immobilized on the Si substrate functionalized with amino groups and DOPA molecules, respectively. (f) Plot of the average number of AuNPs from the SEM images (n = 5). (g, h) Resonant frequency shift of the microcantilever in every step of the fabrication procedure is measured in air (see Table 1). (i) Plot of the total mass of AuNPs bound to the chemically modified surface measured from the frequency shift of a cantilever due to AuNP binding (n = 3).

Now, we have studied binding affinity using cantilever assay that allows for measuring the total mass of AuNPs attached to the surface of the microcantilever. For such a study, we have prepared cantilevers whose surfaces are chemically modified by an amine group or DOPA, respectively. It is shown in Figure 3g,h that the surface modification of cantilevers using amine group or DOPA reduces their resonant frequencies, which is attributed to the weight of the functionalized chemical groups (i.e., amine or DOPA). We have observed that the binding of AuNPs onto the amine- or DOPA-immobilized surface of the cantilever significantly decreases the resonant frequency of such a cantilever (Table 1). The total weight of AuNPs chemically bound to the cantilever can be estimated from the measured frequency shift due to AuNP binding to the cantilever. In particular, the relationship between the total mass of AuNPs bound to the cantilever and the frequency shift is represented in the form Δω/ω0 = (1/2)(ΔM/Mc), where Δω is the resonant frequency shift due to AuNP binding onto the cantilever, ΔM is the total mass of AuNPs chemically attached to the cantilever, and ω0 and Mc represent the resonant frequency and mass, respectively, of the microcantilever whose surface is chemically modified. It is found that the total mass of AuNPs attached to the amine-modified cantilever surface is estimated as ΔMA = 488 ± 10 pg, while the total mass of AuNPs bound to the DOPA-functionalized cantilever surface is measured as ΔMD = 630 ± 27 pg (Figure 3i). This clearly demonstrates that the DOPA-modified surface exhibits stronger binding affinity to AuNPs than the amine-modified surface. As in the previous paragraph, we have introduced the dimensionless quantity RM defined as RM = ΔMDMA, which allows for quantitative comparison. It is interestingly shown that RM is very close to RN, as anticipated (i.e., RM = approximately 1.3 and RN = approximately 1.6). This confirms that the dimensionless quantities RM and RN are useful parameters that allow for quantifying the binding affinity between the chemically modified surface and AuNPs.
Table 1

The resonant frequency shift of AFM cantilevers in cantilever assay





AuNP coating

AuNP-SA (kHz)

300.2 ± 32.3

299.8 ± 31.7


298.4 ± 32.6

AuNP-SD (kHz)

279.8 ± 3.5

279.5 ± 3.8

276.9 ± 4.0

274.2 ± 4.0

Standard deviations are derived from three independent measurements.

Direct measurement of the binding affinity between AuNPs and chemical functional groups

While measurement of the number of attached AuNPs on the surface or mass of the AuNPs bound to the surface is an indirect method to quantify the binding affinity between AuNPs and chemically functionalized surface, we have taken into account the direct method for quantitative characterization of such binding affinity. Here, we have employed a novel scanning technique, namely PeakForce QNM [49], that is useful in measuring the adhesion force between AuNPs and chemically modified surface. Figure 4a,b shows the AFM topography images of AuNPs attached to either SA or SD. It is shown that the AFM height for the AuNPs attached to SD is measured as approximately 14.2 nm, whereas the AFM height for the AuNPs bound to SA is measured as approximately 15.5 nm. This is attributed to the size of the functionalized molecules such that the chain length of DOPA is much larger than that of the amine group [48]. As shown in Figure 4g,h, the AuNP is more likely to be embedded in DOPA, that is, more number of DOPA molecules (than that of amine molecules) is likely to be involved in AuNP binding. This suggests that DOPA molecules may allow for establishing the stable, reliable adhesion of nanoparticles. Notably, we found that the width of the AuNPs bound to SD (99.1 ± 12.3) in both topology and adhesion map is larger by the amount of approximately 7 nm than that bound to SA (92.2 ± 19.7), as shown in Figure 4a,b. This result seems to contradict the fact that the AuNPs in SD are more deeply embedded than those in SA. It may be attributed to the fact that AFM indentation may induce the significant motion of AuNPs, which may distort the size of the AuNPs. In particular, a previous study [50] reports that there is greater energy dissipation at the edge of a nanoparticle than at its center, implying that the nanoparticle would be wobbled during AFM indentation, whereas the nanoparticle would not be moved during tapping mode AFM imaging. As shown in the AFM deformation images (Figure 4e,f), there is a larger deformation change of AuNPs in AuNP-SA than in AuNP-SD during PeakForce QNM. This indicates a stronger binding of AuNPs onto SD rather than on SA, which is attributed to the narrow structural dimension of AuNPs immobilized on SA in comparison with those on SD. Moreover, we have also considered the adhesion map for AuNPs attached to SA or SD. It is found that the adhesion force difference between silicon nitride (Si3N4) AFM tip and the surface (i.e., SA or SD) is <5 nN and that the adhesion force is not significantly dependent on the type of surface chemistry (i.e., whether the surface is functionalized with amine group or DOPA). This indicates that the interaction between the Si3N4 AFM tip and the surface is not critical when we measure the adhesion force between AuNPs and surface. It is shown that the adhesion force between the Si3N4 AFM tip and chemically modified surface to which the AuNPs do not adhere is measured as approximately 10 nN. Nevertheless, the adhesion force map obtained from PeakForce QNM is insufficient to distinguish the binding affinity between AuNPs and SD from that between AuNPs and SA, while the AFM height in topology and the width in the adhesion map for AuNPs bound to SA or SD allow for the distinction between such binding affinities as described earlier.
Figure 4

PeakForce QNM analysis of AuNP-immobilized surfaces. (a, b) Topographic AFM images of 20 nm of AuNPs immobilized on aminated surface and DOPA-functionalized surface, respectively. (c, d) Adhesion images of the samples show relative adhesion interaction between the bare Si AFM tip and AuNPs or other regions outside the AuNPs. All scale bars are 200 nm. The dashed line is a depiction of AFM stylus trajectory. (e, f) AFM deformation images show a larger deformation change near the edge than at the center of the nanoparticles. (g, h) Schematic diagram of the geometric design depicted AuNPs immobilized on the substrates functionalized with amino group and DOPA molecules, respectively. The terms wtopology and htopology indicate the measured width and height, respectively, of the AuNPs in the aspect of topology (a, b).

Another way to directly measure the binding affinity between AuNPs and chemically functionalized surface is to utilize LFM that enables the measurement of friction force between the AFM tip and the sample surface (Figure 5). For LFM imaging, we have utilized a triangular-shaped microcantilever whose normal spring constant knorm is in the range of 0.16 to 1 N/m, suitable for contact mode AFM imaging. In general, the normal spring constant of a cantilever depends on its shape and material [51]. In our study, we have used Si3N4 cantilevers so that the normal spring constant of a microcantilever is determined from its shape. The lateral spring constant klat is related to the normal spring constant given by the following equation [35]:
k lat = 2 6 cos 2 θ + 3 1 + v sin 2 θ L H 2 k norm
where θ is the angle between the base arms of the triangular cantilever, v is the Poisson ratio for silicon nitride, L is the length of the cantilever beam, and H is the tip vertical height (see Table 2). With the estimation of klat from Equation 1, the lateral force (Flat) can be calculated from the measurement of the lateral force signal in LFM analysis such as [36]
F lat = k lat × S lat × Δ V
Figure 5

Lateral force microscopy analysis of binding affinity between AuNPs and chemically functionalized surfaces. (a, b) AFM topographic images and lateral force images of AuNPs immobilized on Si substrates, respectively, functionalized with (a) amino groups (AuNP-SA) and (b) catecholamine molecules (AuNP-SD) by different normal spring constants (kN) of microcantilevers. The inset shows left embankment (indicated by a red arrowhead) due to swept AuNPs formed by scanning the surface in AFM contact mode with kN = 0.7 N/m of microcantilever. All scale bars including that of the inset are 250 nm. (c) Line profiles corresponding to the white arrow in each image of (a) and (b) show the curves of scanner retracting distance versus the AuNP lateral displacement and the lateral force versus the lateral displacement, respectively. (d) Graph of the average lateral force of >100 AuNPs in (a) and (b) (the asterisk indicates p < 0.001) was extracted and calculated from the line profiles of lateral forces (c). (e) The model illustrates the physical interaction between the AFM tip and AuNPs attached on the chemically functionalized substrate (i.e., AuNP-SA and AuNP-SD).

Table 2

Summary of triangular microcantilever parameters (SNL and ScanAsyst Fluid) used in LFM study


ScanAsyst Fluid


L (μm)



w (μm)



t (μm)



H (μm)



E (GPa)









knorm (N/m)



Snorm (nm/V)



klat (N/m)



Slat (nm/V)



L, cantilever length; w, cantilever width; t, cantilever thickness; H, stylus height; E, Young's modulus of Si; v, Poisson's ratio of Si; θ, angle between the base arms of the triangular cantilever; knorm, measured cantilever's normal spring constant; Snorm, measured cantilever's normal sensitivity; klat, lateral spring constant calculated from the knorm; Slat lateral normal sensitivity calculated from Snorm[35, 36].

Here, Slat is the lateral sensitivity of the cantilever defined as Slat = PHSnorm/aR*L[36], where P is a proportionality factor (≈2.5 for the triangular cantilever), Snorm is the vertical deflection sensitivity of the cantilever, a is the amplification factor of the lateral signal measured, and R* is the ratio of the beam height to the beam width (R* = 0.5) [36]. ΔV is the measured value in LFM analysis, which is extracted from the LFM images. In general, the longer and larger the cantilever, the lower is its normal spring constant (i.e., more flexible in normal deflection), but the larger is its lateral spring constant (klat). We can control the exerted applied force using different spring constants of cantilevers (knorm) under an identical deflection set point (1 V) rather than a set point control with an identical AFM tip in order to avoid damage to the samples and a subsidiary frictional noise.

In LFM imaging for the measurement of friction force between the two surfaces (i.e., surface of the AuNPs and the substrate functionalized with chemical groups), one has to be cautious in selecting a cantilever; in particular, a cantilever with a stiffness of <0.16 N/m is too flexible to scan our sample, whereas a cantilever with a stiffness of ≥1 N/m is too stiff to measure the friction force in our sample. As shown in Figure 6, the AFM imaging of our sample using a cantilever with a stiffness of 1 N/m leads to the detachment of AuNPs from the surface during the imaging, which implies the difficulty in accurately measuring the friction force between AuNPs and surface (i.e., SA or SD). Figure 5a,b shows the AFM/LFM images of AuNPs attached to SD or SA. It is shown in Figure 5a that during AFM imaging using a cantilever with a stiffness of knorm = 0.16 N/m, AuNPs are detached from SA (i.e., AFM image shows a scratched pattern corresponding to the imaged AuNPs), while the detachment of AuNPs from SD does not occur. Moreover, it is found that AuNPs are still bound to SD even when AFM and LFM imaging were implemented using a cantilever with a stiffness of knorm = 0.7 N/m (Figure 5b). Figure 5c shows the section profile extracted from the AFM/LFM images (as indicated by a white arrow). It is shown that in the AFM height profile, as anticipated, the AFM height of the AuNPs bound to SA is close to that of the AuNPs attached to SD. On the other hand, in the lateral force profile extracted from the LFM image, we can find the significant differences between the durability of two samples, i.e., AuNPs bound to SA and SD, respectively. This is attributed to the fact that during imaging of AuNPs bound to SA, the twist of the cantilever tip is not significant, which leads to low signals in the LFM image, while the binding between AuNPs and SD (stronger than that between AuNPs and S A ) leads to more twist of the cantilever tip and consequently produce a large signal in LFM imaging [37] (Figure 5e). Based on the LFM images with Equations 1 and 2, we have measured the lateral force between AuNPs and chemically modified surface. It is found that the mean lateral force between AuNPs and SA is measured as 660 nN, while the mean lateral force between AuNPs and SD is estimated to be 1.2 μN (Figure 5d). For quantitative comparison, we have introduced a dimensionless parameter defined as RF = FD/FA, where FA indicates the lateral force between AuNPs and SA, and FD represents the lateral force between AuNPs and SD. It is interestingly found that the dimensionless parameter RF (=1.7) is very close to the aforementioned dimensionless parameters RN and RM (Figure 7). This suggests that the binding affinity between AuNPs and chemically functionalized surface can be quantitatively understood by using either of the indirect experimental methods such as cantilever assay or direct force measurement such as LFM imaging. It should be noted that the binding force between AuNPs and chemically functionalized surface could be measured using AFM pulling experiments [35, 52], which enables the measurement of the normal force required to break a chemical bond. In general, the normal adhesion force driven by the mechanical detachment of AuNPs from the surface might be much lower than the shear adhesion force between AuNPs and the surface. It is attributed to the fact that a shear force required to break chemical bonds is much larger than a normal force that leads to breakage of chemical bonds [53, 54]. This indicates that LFM imaging-based measurement allows for estimating the maximum strength of chemical bonds between nanostructure and chemical functional group. Moreover, AFM pulling experiment-based measurement of normal force required to break chemical bonds requires statistical analysis (based on repetitive experiments due to the effect of thermal fluctuation on force-driven bond rupture [5557]), while LFM imaging-based measurement of shear force for breaking bonds does not require repetitive experiments because LFM imaging enables the parallel measurement of shear forces required to break chemical bonds in the scanned area of a sample. In other words, LFM imaging enables the simultaneous measurement of shear forces (with more than 100 times) required to break chemical bonds, which results in an effective statistical analysis based on only a single LFM image.
Figure 6

The PeakForce QNM analysis of the AuNP- S D sample. AFM topology, peak force error, and adhesion images (3 × 3 μm2) of the embankments composed of AuNPs swept by the 2 × 2 μm2 scanning of a microcantilever with kN = 1 N/m in LFM.
Figure 7

Plot of binding affinity ratio between AuNPs and chemically functionalized surfaces (AuNP- S A and AuNP- S D ). The binding affinity ratios are obtained from experiments such as SEM image, cantilever assay, and LFM. This analysis indicates that DOPA molecules are approximately 1.5 times stronger than amino groups in their adhesion property for the immobilization of AuNPs.


In conclusion, we have demonstrated a quantitative characterization of the binding affinity between AuNPs and chemically modified surface using various experimental techniques such as SEM image analysis, cantilever assay, PeakForce QNM, and LFM image analysis. It is shown that the DOPA-modified surface is an effective conjugation method for functionalization of nanoparticles onto the surface when compared with amine-modified surface, as anticipated, from our various experiments. More remarkably, we have shown that dimensionless parameters (i.e., RN, RM, and RF) introduced in this work are useful in quantifying the binding affinity between nanoparticle and chemical functional groups, and that these dimensionless parameters are consistent regardless of experiments, i.e., RN, RM, and RF are almost identical to each other, implying that the binding affinity between nanostructure and chemical group can be quantitatively studied using either indirect method (i.e., SEM image analysis and cantilever assay) or direct method (i.e., lateral force measurement). Our study sheds light on how to quantitatively study the binding affinity between nanostructure and chemical functional group, which can provide the design principles for nanoparticle-based systems such as nanomedicine and nanobiosensor.



Atomic force microscopy




Gold nanoparticle


Dopamine hydrochloride

PeakForce QNM: 

PeakForce quantitative nanomechanical property mapping


Lateral force microscopy


Amine-functionalized surface


DOPA-modified surface


Scanning electron microscope


X-ray photoelectron spectroscopy.



This work is supported by the National Research Foundation (NRF) of Korea (under grant nos. NRF-2010-0009428, 2010–0027238, 2011–0009885, and 2012R1A2A2A04047240).

Authors’ Affiliations

Institute for Molecular Sciences
Department of Biomedical Engineering, Yonsei University
Department of Energy IT, Gachon University
Department of Radiology, College of Medicine, Yonsei University


  1. Rosi NL, Mirkin CA: Nanostructures in biodiagnostics. Chem Rev 2005, 105: 1547. 10.1021/cr030067fView ArticleGoogle Scholar
  2. Schroeder A, Heller DA, Winslow MM, Dahlman JE, Pratt GW, Langer R, Jacks T, Anderson DG: Treating metastatic cancer with nanotechnology. Nat Rev Cancer 2012, 12: 39.View ArticleGoogle Scholar
  3. Lee J-H, Huh Y-M, Jun Y-w, Seo J-w, Jang J-t, Song H-T, Kim S, Cho E-J, Yoon H-G, Suh J-S, Cheon J: Artificially engineered magnetic nanoparticles for ultra-sensitive molecular imaging. Nat Med 2007, 13: 95. 10.1038/nm1467View ArticleGoogle Scholar
  4. Chen J, Saeki F, Wiley BJ, Cang H, Cobb MJ, Li Z-Y, Au L, Zhang H, Kimmey MB, Li X, Xia Y: Gold nanocages: bioconjugation and their potential use as optical imaging contrast agents. Nano Lett 2005, 5: 473. 10.1021/nl047950tView ArticleGoogle Scholar
  5. Yang P-H, Sun X, Chiu J-F, Sun H, He Q-Y: Transferrin-mediated gold nanoparticle cellular uptake. Bioconjug Chem 2005, 16: 494. 10.1021/bc049775dView ArticleGoogle Scholar
  6. Ling D, Park W, Park YI, Lee N, Li F, Song C, Yang S-G, Choi SH, Na K, Hyeon T: Multiple-interaction ligands inspired by mussel adhesive protein: synthesis of highly stable and biocompatible nanoparticles. Angew Chem Int Ed 2011, 50: 11360. 10.1002/anie.201101521View ArticleGoogle Scholar
  7. Lim E-K, Huh Y-M, Yang J, Lee K, Suh J-S, Haam S: pH-triggered drug-releasing magnetic nanoparticles for cancer therapy guided by molecular imaging by MRI. Adv Mater 2011, 23: 2436. 10.1002/adma.201100351View ArticleGoogle Scholar
  8. Lee J-H, Jang J-t, Choi J-s, Moon SH, Noh S-h, Kim J-w, Kim J-G, Kim I-S, Park KI, Cheon J: Exchange-coupled magnetic nanoparticles for efficient heat induction. Nat Nanotechnol 2011, 6: 418. 10.1038/nnano.2011.95View ArticleGoogle Scholar
  9. Mingce L, Jingjing J, Yan L, Ruqiong C, Liying Z, Weimin C: Effect of gold nanoparticles on the photocatalytic and photoelectrochemical performance of Au modified BiVO4. Nano-Micro Lett 2011, 3: 171.View ArticleGoogle Scholar
  10. Vitale F, Fratoddi I, Battocchio C, Piscopiello E, Tapfer L, Russo M, Polzonetti G, Giannini C: Mono- and bi-functional arenethiols as surfactants for gold nanoparticles: synthesis and characterization. Nanoscale Res Lett 2011, 6: 103. 10.1186/1556-276X-6-103View ArticleGoogle Scholar
  11. Jin R, Wu G, Li Z, Mirkin CA, Schatz GC: What controls the melting properties of DNA-linked gold nanoparticle assemblies? J Am Chem Soc 2003, 125: 1643. 10.1021/ja021096vView ArticleGoogle Scholar
  12. Storhoff JJ, Elghanian R, Mucic RC, Mirkin CA, Letsinger RL: One-pot colorimetric differentiation of polynucleotides with single base imperfections using gold nanoparticle probes. J Am Chem Soc 1959, 1998: 120.Google Scholar
  13. Zhang Y, Li B, Chen X: Simple and sensitive detection of dopamine in the presence of high concentration of ascorbic acid using gold nanoparticles as colorimetric probes. Microchimica Acta 2010, 168: 107. 10.1007/s00604-009-0269-5View ArticleGoogle Scholar
  14. Yang J, Eom K, Lim E-K, Park J, Kang Y, Yoon DS, Na S, Koh EK, Suh J-S, Huh Y-M, Kwon TY, Haam S: In situ detection of live cancer cells by using bioprobes based on au nanoparticles. Langmuir 2008, 24: 12112. 10.1021/la802184mView ArticleGoogle Scholar
  15. Li J-L, Wang L, Liu X-Y, Zhang Z-P, Guo H-C, Liu W-M, Tang S-H: In vitro cancer cell imaging and therapy using transferrin-conjugated gold nanoparticles. Cancer Lett 2009, 274: 319. 10.1016/j.canlet.2008.09.024View ArticleGoogle Scholar
  16. Lee H, Dellatore SM, Miller WM, Messersmith PB: Mussel-inspired surface chemistry for multifunctional coatings. Science 2007, 318: 426. 10.1126/science.1147241View ArticleGoogle Scholar
  17. Lee H, Lee BP, Messersmith PB: A reversible wet/dry adhesive inspired by mussels and geckos. Nature 2007, 448: 338. 10.1038/nature05968View ArticleGoogle Scholar
  18. Lu C-C, Zhang M, Li A-J, He X-W, Yin X-B: 3,4-Dihydroxy-l-phenylalanine for preparation of gold nanoparticles and as electron transfer promoter in H2O2 biosensor. Electroanalysis 2011, 23: 2421. 10.1002/elan.201100291View ArticleGoogle Scholar
  19. Black KCL, Liu Z, Messersmith PB: Catechol redox induced formation of metal core − polymer shell nanoparticles. Chem Mater 2011, 23: 1130. 10.1021/cm1024487View ArticleGoogle Scholar
  20. Kim S, Ku SH, Lim SY, Kim JH, Park CB: Graphene–biomineral hybrid materials. Adv Mater 2009, 2011: 23.Google Scholar
  21. Ryu S, Lee Y, Hwang J-W, Hong S, Kim C, Park TG, Lee H, Hong SH: High-strength carbon nanotube fibers fabricated by infiltration and curing of mussel-inspired catecholamine polymer. Adv Mater 1971, 2011: 23.Google Scholar
  22. Burdine L, Gillette TG, Lin H-J, Kodadek T: Periodate-triggered cross-linking of dopa-containing peptide − protein complexes. J Am Chem Soc 2004, 126: 11442. 10.1021/ja045982cView ArticleGoogle Scholar
  23. Ku SH, Ryu J, Hong SK, Lee H, Park CB: General functionalization route for cell adhesion on non-wetting surfaces. Biomaterials 2010, 31: 2535. 10.1016/j.biomaterials.2009.12.020View ArticleGoogle Scholar
  24. Yang HS, Park J, La WG, Jang H-k, Lee M, Kim B-S: 3,4-Dihydroxyphenylalanine-assisted hydroxyapatite nanoparticle coating on polymer scaffolds for efficient osteoconduction. Tissue Eng Part C Methods 2011, 18: 1.Google Scholar
  25. Baik SH, Kim JH, Cho HH, Park S-N, Kim YS, Suh H: Development and analysis of a collagen-based hemostatic adhesive. J Surg Res 2010, 164: e221. 10.1016/j.jss.2010.08.004View ArticleGoogle Scholar
  26. Bishop KJM, Wilmer CE, Soh S, Grzybowski BA: Nanoscale forces and their uses in self-assembly. Small 2009, 5: 1600. 10.1002/smll.200900358View ArticleGoogle Scholar
  27. Cheng YC, Samia A, Meyers JD, Panagopoulos I, Fei B, Burda C: Highly efficient drug delivery with gold nanoparticle vectors for in vivo photodynamic therapy of cancer. J Am Chem Soc 2008, 130: 10643. 10.1021/ja801631cView ArticleGoogle Scholar
  28. Kim C-k, Ghosh P, Rotello VM: Multimodal drug delivery using gold nanoparticles. Nanoscale 2009, 1: 61. 10.1039/b9nr00112cView ArticleGoogle Scholar
  29. Eom K, Park HS, Yoon DS, Kwon T: Nanomechanical resonators and their applications in biological/chemical detection: nanomechanics principles. Phys Rep 2011, 503: 115. 10.1016/j.physrep.2011.03.002View ArticleGoogle Scholar
  30. Buchapudi KR, Huang X, Yang X, Ji H-F, Thundat T: Microcantilever biosensors for chemicals and bioorganisms. Analyst 2011, 136: 1539–1556. 10.1039/c0an01007cView ArticleGoogle Scholar
  31. Anja B, Søren D, Stephan Sylvest K, Silvan S, Maria T: Cantilever-like micromechanical sensors. Rep Prog Phys 2011, 74: 036101. 10.1088/0034-4885/74/3/036101View ArticleGoogle Scholar
  32. Datar R, Kim S, Jeon S, Hesketh P, Manalis S, Boisen A, Thundat T: Cantilever sensors: nanomechanical tools for diagnostics. MRS Bull 2009, 34: 449. 10.1557/mrs2009.121View ArticleGoogle Scholar
  33. Sweers K, van der Werf K, Bennink M, Subramaniam V: Nanomechanical properties of alpha-synuclein amyloid fibrils: a comparative study by nanoindentation, harmonic force microscopy, and Peakforce QNM. Nanoscale Res Lett 2011, 6: 270. 10.1186/1556-276X-6-270View ArticleGoogle Scholar
  34. Young TJ, Monclus MA, Burnett TL, Broughton WR, Ogin SL, Smith PA: The use of the PeakForce TM quantitative nanomechanical mapping AFM-based method for high-resolution Young's modulus measurement of polymers. Meas Sci Technol 2011, 22: 125703. 10.1088/0957-0233/22/12/125703View ArticleGoogle Scholar
  35. Noy A, Frisbie CD, Rozsnyai LF, Wrighton MS, Lieber CM: Chemical force microscopy: exploiting chemically-modified tips to quantify adhesion, friction, and functional group distributions in molecular assemblies. J Am Chem Soc 1995, 117: 7943. 10.1021/ja00135a012View ArticleGoogle Scholar
  36. Tocha E, Schönherr H, Vancso GJ: Quantitative nanotribology by AFM: a novel universal calibration platform. Langmuir 2006, 22: 2340. 10.1021/la052969cView ArticleGoogle Scholar
  37. Song J, Wang X, Riedo E, Wang ZL: Elastic property of vertically aligned nanowires. Nano Lett 1954, 2005: 5.Google Scholar
  38. Landherr LJT, Cohen C, Agarwal P, Archer LA: Interfacial friction and adhesion of polymer brushes. Langmuir 2011, 27: 9387. 10.1021/la201396mView ArticleGoogle Scholar
  39. Lang HP, Hegner M, Gerber C: Cantilever array sensors. Mater Today 2005, 8: 30.View ArticleGoogle Scholar
  40. Wu G, Ji H, Hansen K, Thundat T, Datar R, Cote R, Hagan MF, Chakraborty AK, Majumdar A: Origin of nanomechanical cantilever motion generated from biomolecular interactions. Proc Natl Acad Sci 2001, 98: 1560. 10.1073/pnas.98.4.1560View ArticleGoogle Scholar
  41. Hamdy Y, Khaled E: State-space approach to vibration of gold nano-beam induced by ramp type heating. Nano-Micro Lett 2010, 2: 139.View ArticleGoogle Scholar
  42. Lin YH, McConney ME, LeMieux MC, Peleshanko S, Jiang C, Singamaneni S, Tsukruk VV: Trilayered ceramic–metal–polymer microcantilevers with dramatically enhanced thermal sensitivity. Adv Mater 2006, 18: 1157. 10.1002/adma.200502232View ArticleGoogle Scholar
  43. Chan J, Alegre TPM, Safavi-Naeini AH, Hill JT, Krause A, Groblacher S, Aspelmeyer M, Painter O: Laser cooling of a nanomechanical oscillator into its quantum ground state. Nature 2011, 478: 89. 10.1038/nature10461View ArticleGoogle Scholar
  44. Haiss W: Surface stress of clean and adsorbate-covered solids. Rep Prog Phys 2001, 64: 591. 10.1088/0034-4885/64/5/201View ArticleGoogle Scholar
  45. Ou J, Wang J, Liu S, Zhou J, Ren S, Yang S: Microtribological and electrochemical corrosion behaviors of polydopamine coating on APTS-SAM modified Si substrate. Appl Surf Sci 2009, 256: 894. 10.1016/j.apsusc.2009.08.081View ArticleGoogle Scholar
  46. Graf N, Yeğen E, Lippitz A, Treu D, Wirth T, Unger WES: Optimization of cleaning and amino-silanization protocols for Si wafers to be used as platforms for biochip microarrays by surface analysis (XPS, ToF-SIMS and NEXAFS spectroscopy). Surf Interface Anal 2008, 40: 180. 10.1002/sia.2621View ArticleGoogle Scholar
  47. Liu Y, Yu B, Hao J, Zhou F: Amination of surfaces via self-assembly of dopamine. J Colloid Interface Sci 2011, 362: 127. 10.1016/j.jcis.2011.06.007View ArticleGoogle Scholar
  48. Aureau D, Varin Y, Roodenko K, Seitz O, Pluchery O, Chabal YJ: Controlled deposition of gold nanoparticles on well-defined organic monolayer grafted on silicon surfaces. J Phys Chem C 2010, 114: 14180. 10.1021/jp104183mView ArticleGoogle Scholar
  49. Adamcik J, Berquand A, Mezzenga R: Single-step direct measurement of amyloid fibrils stiffness by peak force quantitative nanomechanical atomic force microscopy. Appl Phys Lett 2011, 98: 193701. 10.1063/1.3589369View ArticleGoogle Scholar
  50. Duner G, Thormann E, Dedinaite A, Claesson PM, Matyjaszewski K, Tilton RD: Nanomechanical mapping of a high curvature polymer brush grafted from a rigid nanoparticle. Soft Matter 2012, 8: 8312. 10.1039/c2sm26086gView ArticleGoogle Scholar
  51. Palacio MLB, Bhushan B: Normal and lateral force calibration techniques for AFM cantilevers. Crit Rev Solid State Mater Sci 2010, 35: 73. 10.1080/10408430903546691View ArticleGoogle Scholar
  52. Stroh C, Wang H, Bash R, Ashcroft B, Nelson J, Gruber H, Lohr D, Lindsay SM, Hinterdorfer P: Single-molecule recognition imaging microscopy. Proc Natl Acad Sci USA 2004, 101: 12503. 10.1073/pnas.0403538101View ArticleGoogle Scholar
  53. Kufer SK, Puchner EM, Gumpp H, Liedl T, Gaub HE: Single-molecule cut-and-paste surface assembly. Science 2008, 319: 594. 10.1126/science.1151424View ArticleGoogle Scholar
  54. Qu L, Dai L, Stone M, Xia Z, Wang ZL: Carbon nanotube arrays with strong shear binding-on and easy normal lifting-off. Science 2008, 322: 238. 10.1126/science.1159503View ArticleGoogle Scholar
  55. Eom K, Makarov DE, Rodin GJ: Theoretical studies of the kinetics of mechanical unfolding of cross-linked polymer chains and their implications for single-molecule pulling experiments. Phys Rev E 2005, 71: 021904.View ArticleGoogle Scholar
  56. Evans E: Probing the relation between force—lifetime—and chemistry in single molecular bonds. Annu Rev Biophys Biomol Struct 2001, 30: 105. 10.1146/annurev.biophys.30.1.105View ArticleGoogle Scholar
  57. Evans E, Ritchie K: Dynamic strength of molecular adhesion bonds. Biophys J 1997, 72: 1541. 10.1016/S0006-3495(97)78802-7View ArticleGoogle Scholar


© Lee et al.; licensee Springer. 2012

This article is published under license to BioMed Central Ltd. This is an Open Access article distributed under the terms of the Creative Commons Attribution License (, which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.