Multiple Fishhooks - American Chemical Society

Jun 12, 2012 - interpretation have often been a great hurdle for its widespread application. Here, we present a much simplified experimental scheme by...
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A Versatile “Multiple Fishhooks” Approach for the Study of Ligand− Receptor Interactions Using Single-Molecule Atomic Force Microscopy Xiaotian Han, Meng Qin, Hai Pan, Yi Cao,* and Wei Wang* National Laboratory of Solid State Microstructure, Department of Physics, Nanjing University, 22 Hankou Road, Nanjing, Jiangsu 210093, People’s Republic of China S Supporting Information *

ABSTRACT: Despite the powerfulness of atomic force microscopy (AFM)-based single-molecule force spectroscopy in the study of ligand− receptor interactions, complicated cantilever functionalization and data interpretation have often been a great hurdle for its widespread application. Here, we present a much simplified experimental scheme by using a “multiple fishhooks” approach. In this strategy, multiple ligands are labeled on a single polymer chain, which forms complexes with receptors anchored on the substrate surface. Therefore, multiple singlebond rupture events can be captured in the same force−extension curves, similar to the widely used polyprotein approach. This method also allows nonsingle-molecule events and nonspecific interactions between cantilever and surface to be readily excluded from real data pool and greatly increases the quality and quantity of single-molecule data. The biggest advantage of our approach over the previously reported one is the choice of a naturally occurring polysaccharide, hyaluronan, the conformation of which in solution can be fine-tuned by pH, as the polymer backbone of the “multiple fishhooks” handle. Furthermore, our approach greatly simplifies the chemical synthesis of the polymer handle, allowing bioactive molecules to be easily one-step labeled on the handles in aqueous solution. We validate this strategy using the widely studied streptavidin−biotin system, and our single-molecule AFM results are in good agreement with previously reported ones. We anticipate that this novel strategy can be used as a versatile tool to study other complex and challenging ligand−receptor interactions.

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INTRODUCTION Atomic force microscopy (AFM)-based single-molecule force spectroscopy (SMFS) has enjoyed great success in quantifying mechanical strength of ligand−receptor bonds, elucidating the binding mechanism, and exploring the free energy landscape underlying the binding/unbinding events.1−13 A great challenge in SMFS of ligand−receptor interaction is to identify real single-molecule ligand−receptor dissociation events against multiple unbinding events and false-positive results from nonspecific interactions between the cantilever tip and the surface. A tentative solution to this challenge is to reduce the density of ligands and receptors on the cantilever tip and the substrate and to coat them with inert biomacromolecules.14−16 To further avoid nonspecific cantilever−surface interaction, a polymer handle approach is typically used, in which one part of the ligand−receptor pair is hung on the cantilever tip through a long polymer linker, such as polyethylene glycol (PEG).17−21 Although these methods greatly improved the quality of SMFS data, it remains a big challenge to fully discard false-positive results. Some research toward further improving singlemolecule ligand−receptor interactions currently is underway.22−26 © 2012 American Chemical Society

Recently, Samori and co-workers have pioneered a multivalent handle approach,23 which allows sequentially breaking ligand−receptor pairs in the same force−extension trajectory, mimicking the multiple-domain-protein or polyprotein approach typically used in protein folding/unfolding studies.8,27−35 Because all rupture events originate from the same polymer, they should carry the same mechanical features (e.g., persistence length, chair-boat transition pattern, or hydrophobic collapse);36−47 the nonspecific interactions that occur at short separation and the multiple unbinding events that show shorter persistence length can be easily rejected in the data analysis. Therefore, this method can potentially simplify the interpretation of SMFS data using the fingerprint from multiple rupture events. Moreover, this method can greatly increase the successful rate of picking single molecules, enhancing both the quantity and the quality of single-molecule data. However, there are still some fundamental difficulties averting practical application of this technique. First, the chemical synthesis of multivalent handle requires multiple steps and involves organic Received: May 9, 2012 Revised: June 11, 2012 Published: June 12, 2012 10020

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Figure 1. The synthetic scheme of biotin-functionalized HA.

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solvent, preventing the attachment of protein or other bioactive ligands. Second, the ideal molecular weight for polymers suitable for SMFS study is more than 20 kDa, which is challenging for routine polymer synthesis protocols. Third, and more importantly, this approach requires the formation of multiple ligand−receptor pairs between the polymer and the surface. However, the polymer handle chosen in their approach (PEG) is of short persistence length and generally adopts a coiled structure in solution.48 This makes many ligands on the polymer buried inside and inaccessible to the receptors on the surface. Therefore, the number of ligand−receptor pairs formed between the multivalent handle and the surface is very limited (no more than four unbinding events in each trace).23 To circumvent these hurdles and make this approach more practical and efficient, here we introduce a novel strategy, which we name as “multiple fishhooks” approach, for the detection of ligand−receptor interaction. This strategy greatly simplifies the chemistry of the synthesis and improves the data quality. The biggest improvement of our approach relies on our careful choice of the backbone polymer, which significantly facilitate the formation of ligand−receptor pairs between the handle and the surface. The conformation of HA can be tuned by the pH of the solution. It adopts more extended structures at a basic pH of 9, facilitating the formation of ligand−receptor pairs between the handle and the surface (Figure S1). However, at neutral pH, HA is less charged and behaves like a random coil (Figure S1). Therefore, the ligand−receptor interactions in the system are not affected by the charges on the polymer backbone. Using the widely studied biotin−streptavidin system as a mode system,14,49−54 we demonstrate two ways to perform single-molecule AFM experiments using this approach. Our experimental data are in good agreement with the published results in literature, indicating the reliability of this novel approach. In addition, we show that this method can greatly avoid the contamination of experimental data from nonspecific tip−surface interaction and from simultaneous multiple unbound events.

EXPERIMENTAL SECTION

Synthesis of Multivalent Polymer Handle. HA−biotin handle was prepared as follows. First, 60 mg of HA (MW: 150 kDa, Freda Biopharm, Shangdong, China) was first dissolved in 1 mL of Milli-Q water. Second, 46.5 mg (0.243 mmol) of EDC (Sigma-Aldrich) was added to HA solution and stirred for 0.5 h to convert the carboxyl groups of HA to amine-reactive acylisourea intermediates. Next, 45.7 mg (0.177 mmol) of biotin hydrazide (Sigma-Aldrich), predissolved in 20 μL of DMSO (Sigma-Aldrich), was added to the same solution to conjugate biotin to HA. The reaction was preceded overnight at room temperature under magnetic stirring. The unreacted biotin hydrazide and other byproducts were removed by dialyzing against excess MilliQ water using dialysis tubing with the molecular weight cutoff of 50 kDa. The final product was lyophilized and stored at −20 °C for subsequent single-molecule AFM experiments. The conjugation rate was estimated to be ∼40% on the basis of the NMR spectrum of HA− biotin (Figure S2). Surface Modification of the Substrate. First, the glass coverslips (Fisher Scientific) were place in a warm chromium acid solution for 3 h to remove residual organic matter. Next, the coverslips were rinsed with Milli-Q water and dried under a stream of nitrogen. Subsequently, each dried coverslip was silanized with 20 μL of pure 3aminopropyl dimethyl ethoxysilane for 30 min. Next, the coverslips were rinsed with Milli-Q water, dried under a stream of nitrogen, and further incubated in a drying oven at 80 °C for 1 h. These aminated coverslips were stored at room temperature under nitrogen protection for further use. 100 μL of a mixture of biotin-PEG-SVA (MW 5000 Da, Laysan Bio Inc.) and mPEG-SVA (MW 5000 Da, Laysan Bio Inc.), at a total PEG concentration of 50 mM in the same sodium bicarbonate buffer, was added on each coverslip and allowed to incubate at room temperature for 1 h. The biotin-PEG-SVA/mPEGSVA ratio varied between 1:4 and 1:8 to control the density of biotin on the surface. The coverslips were then rinsed with Milli-Q water and used in single-molecule AFM experiments. Single-Molecule Force Spectroscopy. Single-molecule AFM experiments were carried out on a commercial AFM (JPK NanoWizard II). In the force spectroscopy experiments on the biotinfunctionalized glass, 50 μL of streptavidin solution (0.5 mg mL−1 streptavidin in a buffer containing 25 mM Tris and 150 mM NaCl, pH 7.4) was first placed on the functionalized glass and allowed to adsorb to surface for 15 min. The floating streptavidin was then removed by extensively rinsing the surface using the rinsing buffer (25 mM Tris and 150 mM NaCl, pH 9.0). Next, HA−biotin polymer was deposited on the surface and allowed adsorbing for 15−30 min. HA is more 10021

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charged in the rinsing buffer of pH 9.0; therefore, the HA−biotin chain is more extended and allowed multiple biotin−streptavidin interactions to be formed between the polymer and the surface. Next, the fluid chamber was filled in with ∼1 mL of buffer containing 25 mM Tris, 150 mM NaCl at pH 7.4. The AFM experiments were carried out after allowing the mixture to equilibrate for 30 min. For the experiments using unfunctionalized surface, 50 μL of premixed streptavidin and HA−biotin diluted solution (50 μg mL−1 streptavidin with 5.4 μg mL−1 biotin) was placed on a freshly cleaned glass coverslip and allowed to adsorb for 15 min before each experiment. The rest of the conditions were the same as for the first method. Cantilevers (TR400 from Olympus or MLCT from Bruker) of typical spring constant of 20−100 pN nm−1 were used for all experiments and calibrated using the equipartition theorem before each experiment. The force−extension traces were recorded using commercial software from JPK and analyzed using home written protocol in igor 6.0 (Wavemetrics, Inc.).

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RESULTS AND DISCUSSION We choose a natural polysaccharide, hyaluronan (HA), as the backbone of the “multiple fishhooks” handle (Figure 1). Mechanical properties of individual HA have been studied in detail using SMFS.55 HA is more expanded in solution thanks to the formation of an intramolecular hydrogen-bond network.55−57 Moreover, the carboxyl groups allow HA to be easily functionalized with receptors of interest. We graft biotin hydrazide to HA according to well-established procedures (Experimental Section). Varying the ratio of HA and biotin leads to different grafting densities, which can be verified using NMR spectra (Figure S2). We choose the HA−biotin sample of 40% grafting density throughout this Article. The HA−biotin sample was extensively dialyzed against deionized water and then lyophilized before use. We first establish this method using a strategy similar to that first presented by Samori and co-workers (Figure 2a).23 We immobilized streptavidin on a biotin/PEG-functionalized surface. The floating streptavidin was removed by extensively rinsing the surface using Tris buffer at pH 9. Next, HA−biotin polymer was deposited on the surface and allowed to adsorb for 15−30 min. The conformation of HA is more extended at this pH due to intramolecular hydrogen bonds in HA and repulsive forces between carboxyl groups. Therefore, the biotins on the HA backbone are more exposed and can easily interact with streptavidin on the substrate. Afterward, we switch buffer to pH 7.4 for force-spectroscopy measurements. In a typical experiment, the cantilever was brought to the surface with a constant speed of 1000 nm s−1 and held on the surface at constant forces of 1.5 nN for 1 s. The HA−biotin molecule was picked up from the surface due to physical adsorption of the molecule to the cantilever. We kept the pickup rate as low as around 2% to minimize tip−surface adhesion and to avoid picking up multiple molecules simultaneously. Next, the cantilever was retracted at the same speed to achieve multiple unbinding events in a single force−extension trace. Two representative traces are shown in Figure 2b. We observed multiple sawtooth like peaks at ∼85 pN and of varying spacing. We assigned these peaks as the breaking of biotin−streptavidin interactions. Because biotin was grafted to HA at random positions, such a variable spacing is expected and can be controlled by the grafting density of biotin to HA and the amount of streptavidin on the surface. It is worth noting that the last peaks in these traces are not at elevated forces, distinct from typical force− extension traces obtained in polyprotein experiments. This is because breaking the physical adsorption of HA−biotin from

Figure 2. Single-molecule experiments using HA−biotin and the biotin-functionalized surface. (a) Experimental scheme. (b) Two representative sawtooth like force−extension traces at a pulling speed of 1000 nm s−1. Each peak corresponds to a rupture event between biotin and streptavidin. Red lines correspond to worm-like chain (WLC) fitting to the rupture events using the same persistence length of ∼0.4 nm. (c) The rupture force histogram obtained at a pulling speed of 1000 nm s−1. Red line corresponds to a Gaussian fit.

the cantilever typically requires much higher force (typically ∼200−1000 pN) than the biotin−streptavidin interaction. Therefore, the last peak can still be assigned as the rupture of the biotin−streptavidin interaction instead of the detachment of the nonspecific physical adsorption of HA−biotin from the cantilever. To make sure these events are indeed resulted from breaking single HA−biotin bonds, we used the worm-like chain model (WLC) of polymer elasticity to fit each individual peaks (Figure 2b). It is evidenced that we can use the same persistence length of ∼0.4 nm to fit all peaks in the same trace. The histogram of the persistence length for all traces is centered at ∼0.4 nm, as shown in Figure S3a. Because such a persistence length is characteristic for HA at neutral condition, we were certain that only a single HA−biotin molecule was picked up.23 To further confirm that the sawtooth peaks we observed truly correspond to the rupture of the biotin−streptavidin interaction, we performed two control experiments on the PEG−biotin 10022

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molecules that have been functionalized on the glass surface, and the HA−biotin molecules on the PEG−biotin functionalized surface. Two representative traces for each control experiment are shown in Figures S4 and S5, respectively. Clearly, no such sawtooth-like peaks were observed in both experiments. To evaluate the quality of the data obtained using this method, we measured the rupture force distribution of biotin− streptavidin at a pulling speed of 1000 nm s−1 (Figure 2c). The average unbinding force is 84 ± 23 pN, which is in good agreement with the number reported in literature.14,49−54 It is worth noting that the rupture force histogram obtained using this method is rather clean without any high force tails due to simultaneously rupturing two or more ligand−receptor bonds, which are frequently found in typical SMFS measurements on ligand−receptor interactions. We also measured the rupture forces at different loading rates (Figure 3). It is well-known that

Figure 3. Loading rate dependence of the rupture force of the biotin− streptavidin pairs studied using the “multiple fishhooks” approach. Clearly, there are two regions with distinct loading-rate dependence behaviors. Fitting the experimental data using Bell−Evans model yields the spontaneous dissociation rates at zero force of 0.002 ± 0.003 and 8.8 ± 2.9 s−1 for the first and second barriers, respectively, and the distances to the transition state of 0.62 ± 0.10 and 0.12 ± 0.01 nm for the first and second barriers, respectively.

Figure 4. Single-molecule experiments using preformed HA−biotin and streptavidin complex. (a) Experimental scheme. (b) Two representative sawtooth like force−extension traces at a pulling speed of 1000 nm s−1. Each peak, except the last one, corresponds to a rupture event between biotin and streptavidin. Red lines correspond to worm-like chain fitting to the rupture events using the same persistence length of ∼0.4 nm. (c) The rupture force histogram obtained at a pulling speed of 1000 nm s−1. Red line corresponds to a Gaussian fit.

the rupture force of biotin-streptavidin shows two distinct loading-rate dependences at different loading rate regions due to its unique two-barrier free energy landscape.49−54 At loading rates lower than 5 × 103 pN s−1, the rupture forces increase slowly with respect to the loading rate, while at loading rates higher than 5 × 103 pN s−1, the rupture forces rise much more steeply. By fitting these two regions using Bell−Evans model58,59 (gray lines in Figure 3), the spontaneous unbinding rate and the barrier width were estimated to be 0.002 ± 0.003 s−1 and 0.62 ± 0.10 nm for the first transition, and 8.8 ± 2.9 s−1 and 0.12 ± 0.01 nm for the second transition, respectively. Our data are consistent with the results reported in the literature using standard experimental schemes,54 providing additional evidence of the reliability of this new method. We envisioned that the biotin−streptavidin interaction could be even probed using a simplified scheme, as shown in Figure 4a. Because streptavidin forms a mechanically stable tetramer that can bind up to 4 biotin molecules, we were able to directly prepare HA−biotin in complex with streptavidin in dilute solution. Unfolding such complexes could result in a series of rupture events of individual biotin−streptavidin pairs, similar to the polyprotein approach typically used to study protein mechanics.28−32 This new method does not need cantilever or

surface modification, greatly simplifying the SMFS procedures. As shown in Figure 4b, stretching HA−biotin in complex with streptavidin resulted in sawtooth-like patterns similar to that obtained using surface-immobilized streptavidin method. Each peak, except the last one, corresponds to a rupture event of biotin−streptavidin pair. The last peaks in the traces obtained using this method typically occur at more elevated forces, which correspond to the detachment of physically adsorbed HA− biotin molecules either from the cantilever tip or from the substrate. WLC fitting of each individual peak yielded persistence lengths of 0.4 nm, similar to that obtained using previous method, suggesting that single HA−biotin molecules were stretched (Figure S3b). The rupture force histogram at 1000 nm s−1 obtained in this method centers at 81 ± 22 pN (Figure 4c), in good agreement of that obtained using previous methods,49−54 indicating the validity of this new SFMS procedure. It is worth noting that, because of the high binding 10023

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(5) Jiang, Z.; Zhang, Y.; Yu, Y.; Wang, Z.; Zhang, X.; Duan, X.; Wang, S. Study on intercalations between double-stranded DNA and pyrene by single-molecule force spectroscopy: toward the detection of mismatch in DNA. Langmuir 2010, 26, 13773−7. (6) Zhang, Y.; Liu, C.; Shi, W.; Wang, Z.; Dai, L.; Zhang, X. Direct measurements of the interaction between pyrene and graphite in aqueous media by single molecule force spectroscopy: understanding the pi-pi interactions. Langmuir 2007, 23, 7911−5. (7) Muller, D. J.; Dufrene, Y. F. Atomic force microscopy: a nanoscopic window on the cell surface. Trends Cell Biol. 2011, 21, 461−9. (8) Kedrov, A.; Janovjak, H.; Sapra, K. T.; Muller, D. J. Deciphering molecular interactions of native membrane proteins by single-molecule force spectroscopy. Annu. Rev. Biophys. Biomol. Struct. 2007, 36, 233− 60. (9) Muller, D. J.; Krieg, M.; Alsteens, D.; Dufrene, Y. F. New frontiers in atomic force microscopy: analyzing interactions from single-molecules to cells. Curr. Opin. Biotechnol. 2009, 20, 4−13. (10) Sotres, J.; Lostao, A.; Wildling, L.; Ebner, A.; Gomez-Moreno, C.; Gruber, H. J.; Hinterdorfer, P.; Baro, A. M. Unbinding molecular recognition force maps of localized single receptor molecules by atomic force microscopy. ChemPhysChem 2008, 9, 590−9. (11) Zhang, W.; Cui, S.; Fu, Y.; Zhang, X. Desorption force of poly(4-vinylpyridine) layer assemblies from amino groups modified substrates. J. Phys. Chem. B 2002, 106, 12705−12708. (12) Liu, N.; Peng, B.; Lin, Y.; Su, Z.; Niu, Z.; Wang, Q.; Zhang, W.; Li, H.; Shen, J. Pulling genetic RNA out of tobacco mosaic virus using single-molecule force spectroscopy. J. Am. Chem. Soc. 2010, 132, 11036−8. (13) Cui, S.; Liu, C.; Zhang, W.; Zhang, X.; Wu, C. Desorption force per polystyrene segment in water. Macromolecules 2003, 36, 3779− 3782. (14) Moy, V. T.; Florin, E. L.; Gaub, H. E. Intermolecular forces and energies between ligands and receptors. Science 1994, 266, 257−9. (15) Florin, E. L.; Moy, V. T.; Gaub, H. E. Adhesion forces between individual ligand-receptor pairs. Science 1994, 264, 415−7. (16) Marshall, B. T.; Long, M.; Piper, J. W.; Yago, T.; McEver, R. P.; Zhu, C. Direct observation of catch bonds involving cell-adhesion molecules. Nature 2003, 423, 190−3. (17) Hinterdorfer, P.; Baumgartner, W.; Gruber, H. J.; Schilcher, K.; Schindler, H. Detection and localization of individual antibody-antigen recognition events by atomic force microscopy. Proc. Natl. Acad. Sci. U.S.A. 1996, 93, 3477−81. (18) Conti, M.; Falini, G.; Samori, B. How strong is the coordination bond between a histidine tag and Ni-nitrilotriacetate? An experiment of mechanochemistry on single molecules. Angew. Chem., Int. Ed. 2000, 39, 215−218. (19) Lee, H.; Scherer, N. F.; Messersmith, P. B. Single-molecule mechanics of mussel adhesion. Proc. Natl. Acad. Sci. U.S.A. 2006, 103, 12999−3003. (20) Sonnenberg, L.; Luo, Y.; Schlaad, H.; Seitz, M.; Colfen, H.; Gaub, H. E. Quantitative single molecule measurements on the interaction forces of poly(L-glutamic acid) with calcite crystals. J. Am. Chem. Soc. 2007, 129, 15364−71. (21) Barattin, R.; Voyer, N. Chemical modifications of AFM tips for the study of molecular recognition events. Chem. Commun. 2008, 1513−32. (22) Jung, Y. J.; Hong, B. J.; Zhang, W.; Tendler, S. J.; Williams, P. M.; Allen, S.; Park, J. W. Dendron arrays for the force-based detection of DNA hybridization events. J. Am. Chem. Soc. 2007, 129, 9349−55. (23) Valle, F.; Zuccheri, G.; Bergia, A.; Ayres, L.; Rowan, A. E.; Nolte, R. J.; Samori, B. A polymeric molecular “handle” for multiple AFMbased single-molecule force measurements. Angew. Chem., Int. Ed. 2008, 47, 2431−4. (24) Leitner, M.; Mitchell, N.; Kastner, M.; Schlapak, R.; Gruber, H. J.; Hinterdorfer, P.; Howorka, S.; Ebner, A. Single-molecule AFM characterization of individual chemically tagged DNA tetrahedra. ACS Nano 2011, 5, 7048−54.

affinity of biotin−streptavidin pair, the supramolecular complex formed by HA−biotin and streptavidin could be purified by gel filtration chromatography prior to the SMFS experiments, leading to further improved data quality. However, for monovalent ligand−receptor pairs, chemical cross-linking is required for the utilization of this method.

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CONCLUSION In this Article, we present a more practical and efficient “multiple fishhooks” strategy to study ligand−receptor interactions. We demonstrate that the choice of the polymer backbone is crucial for the realization of this strategy. By using a more rigid polysaccharide, HA, as the polymer backbone of the “multiple fishhooks” handle, the quality and quantity of the SMFS data obtained using this method are much improved. Moreover, this method also greatly simplified the synthesis protocols for the ligand-labeled polymer handles without the need of sophisticated polymer synthesis procedures. We hope this strategy can be widely used as a general method to study various ligand−receptor interactions. Moreover, we provide a new way to study ligand−receptor interactions by using preformed complexes of a ligand-labeled polymer and multiple receptors. This method is extremely suitable for studying ligand−receptor interactions that involve receptors that are not suitable for immobilization on surface, such as single cells or single nanoparticles. Currently, we are working on different polymer backbones, such as positive charged polymers and polymers that are suitable to be used in organic solvents, to further explore this new strategy.

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ASSOCIATED CONTENT

* Supporting Information S

Data analysis protocols and four supporting figures. This material is available free of charge via the Internet at http:// pubs.acs.org.

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AUTHOR INFORMATION

Corresponding Author

*E-mail: [email protected] (Y.C.); [email protected] (W.W.). Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS This work is supported by the National Natural Science Foundation of China under Grant nos. 11074115, 81121062, 10834002, 10904064, and 31170813, the Natural Science Foundation of Jiangsu Province under Grant No.BK2009008, the program for New Century Excellent Talents in University, and the Priority Academic Program Development of Jiangsu Higher Education Institutions.

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dx.doi.org/10.1021/la301903z | Langmuir 2012, 28, 10020−10025