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Langmuir 2008, 24, 10786-10790
Nanoneedle Method for High-Sensitivity Low-Background Monitoring of Protein Activity Maryam Jouzi,† Matthew B. Kerby,‡,§ Anubhav Tripathi,*,‡,§ and Jimmy Xu†,‡ Department of Physics, DiVision of Engineering, and DiVision of Medical Science, Brown UniVersity, ProVidence, Rhode Island 02912 ReceiVed NoVember 20, 2007. ReVised Manuscript ReceiVed May 3, 2008 We describe a new method for measuring the activity of protein in miniscule quantities using a carbon nanotube nanoneedle. The unique features of this new method are (a) the immobilization of a few molecules; (b) subsequent translocation and isolation of them near the tip of a position-actuated nanoneedle; and (c) a fixed, defined, and unhindered molecular position to allow rapid real-time sensing and monitoring. The kinetic bioactivity of immobilized alkaline phosphatase (AP) molecules was measured as test model. Results show no decrease in enzymatic activity compared to that of the solution-phase enzyme reaction, suggesting that the immobilization provided unhindered access for ligand binding and minimal conformational modulation caused by undesired surface interactions.
1. Introduction Protein immobilization, isolation, and real-time activity monitoring are essential to many protein studies, including protein and peptide mapping,1-3 analysis of nucleic acids,4,5 determination of post-translational modifications such as phosphorylation, glycosylation, and lipidylation,6 combinatorial synthesis,7 and characterization of molecules or single cells. The immobilization of enzymes, proteins, or substrates allows for the sparing or repeated use of reagents. Proteins, RNA, and DNA from investigative samples are commonly produced or available only in small quantities. Because the purification of large quantities of these substances from cells can be prohibitively expensive or labor-intensive, a tool for rapid protein immobilization, translocation, and isolation, which enables biomolecular detection, identification, and characterization of function from limited samples, is desired. Immobilization, a powerful technique for the study of biochemical systems, allows for the continuous observation of dynamic behavior of a chosen target. However, methodical challenges remain in achieving suitable immobilization and isolation of biological molecules without compromising their structure, function, activity, or signal. The conventional immobilization of enzymes on a planar surface often results in reduced enzymatic activity as a result of altered protein conformation and/or steric hindrance of the catalytic site.8 In addition, a diffusion layer around the immobilized support creates resistance to mass transfer, which can limit the reaction rate.9 A new method that can provide immobilization for a few
Figure 1. (a) SEM image of a carbon nanotube nanoneedle made out of single-walled carbon nanotubes. (b) Schematic illustration of carbon nanoneedle manufacturing on top of microprobes using a solution of free carbon nanotubes and an electrical field.
* Corresponding author. Phone: 401-863-3063. E-mail: anubhav_tripathi@ brown.edu. † Department of Physics. ‡ Division of Engineering. § Division of Medical Science. (1) Palm, A. K.; Novotny, M. V. Rapid Commun. Mass Spectrom. 2004, 18, 1374–82. (2) Wu, H.; Zhai, J.; Tian, Y.; Lu, H.; Wang, X.; Jia, W.; Liu, B.; Yang, P.; Xu, Y.; Wang, H. Lab Chip 2004, 4, 588–97. (3) Gevaert, K.; Vandekerckhove, J. Electrophoresis 2000, 21, 1145–54. (4) Hashimoto, M.; Chen, P. C.; Mitchell, M. W.; Nikitopoulos, D. E.; Soper, S. A.; Murphy, M. C. Lab Chip 2004, 4, 638–45. (5) Lagally, E. T.; Scherer, J. R.; Blazej, R. G.; Toriello, N. M.; Diep, B. A.; Ramchandani, M.; Sensabaugh, G. F.; Riley, L. W.; Mathies, R. A. Anal. Chem. 2004, 76, 3162–70. (6) Nalivaeva, N. N.; Turner, A. J. Proteomics 2001, 1, 735–47. (7) Watts, P.; Haswell, S. J. Comb Chem High Throughput Screen 2004, 7, 397–405. (8) DeLouise, L. A.; Miller, B. L. Anal. Chem. 2005, 77, 1950–6.
Figure 2. (a) Schematic illustration of the immobilization and isolation protocol using a carbon nanotube nanoneedle. (b) Illustrations of protein (not to scale) immobilization on a nanoneedle tip (upper) versus a planar surface (lower).
molecules supplying uncompromised biomolecular functionality would be highly desirable. In this work, we introduce such a method based on the assembly of carbon nanotubes into a nanoneedle device, which, through a model case study, shows
10.1021/la703630a CCC: $40.75 2008 American Chemical Society Published on Web 08/28/2008
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Figure 3. (a) Bright-field and fluorescent microscope images (∼1700× magnification) of fluorescein-conjugated AP bound to the tip of a nanoneedle made of multiwalled carbon nanotubes. Dashed lines are to guide the eye alongside the surface-protected, nonfluorescent region of the nanoneedle. (b) Schematic diagram of immobilized AP enzymes converting DiMFUp molecules (red spheres) to fluorescent DiMFU molecules (green spheres).
but carries a significant time cost due to all of the complicated target tracking and programming procedures needed to evaluate the data. In addition to optical limitations, the Brownian motion of molecules complicates the tracking of the target molecules, which move in and out of the field of view.17 Alternative monitoring approaches that isolate a single molecule for efficient and unambiguous probing using an immobilization platform11,14,16,18 often carry the significant penalty of denaturation, which results from interaction with the surface, or the chemical modifications used to immobilize them on the surface.11 The nanoneedle method introduced in this work offers essentially unhindered access to the proteins, elimination of background signal from the otherwise coexisting molecules, and a fixed, defined molecular location to allow rapid, real-time detection for reaction kinetics or activity studies.
2. Experimental Methods and Details
Figure 4. Schematic illustration of proteins (not to scale) bound to the nanoneedle tip. It is important to note that the 15 nm tip was likely to be composed of tips of a few single-walled carbon nanotubes of diameter close to 1 nm. Hence, such tips were sharp enough to provide singlepoint contacts for AP molecules (∼5 nm).
Materials. The alkaline phosphatase enzyme was chosen as a representative reactive protein model to assess the potential of the nanoneedle for biomolecule immobilization and kinetics studies. Specifically, the kinetic bioactivity of immobilized biotin-conjugated alkaline phosphatase (biotin-AP) was measured by the dephosphorylation of 6,8-difluoro-4-methylumbelliferyl/phosphate (DiFMUp) to fluorescent 6,8-difluoro-4-methylumbelliferone (DiFMU). Calf intestinal alkaline phosphatase biotin (AP-biotin) was obtained from Rockland Immunochemical (Gilbertsville, PA). DiFMUp (14 000 cm-1 M-1 at 320 nm) and fluorescent DiFMU (18 000 cm-1 M-1 at 358 nm)19 were purchased from Invitrogen (Carlsbad, CA). Both were freshly reconstituted, calibrated by UV absorbance (ND-1000,
the promise of high-sensitivity monitoring of enzymatic activity and the preservation of protein viability. Although not yet achieved in this work, the results presented here represent a significant advance toward reaching the ultimate potential offered by this methodsperforming single-enzyme investigations. Nearly all single-molecule detection techniques are based on optical methods such as fluorescent resonance energy transfer (FRET) using confocal or wide-field microscopy,10-16 but in all of these methods, the area illuminated by excitation and emission light tends to be large, as dictated by the law of diffraction, and thus the background tends to overwhelm the signal from single target molecules. Whereas the use of pinholes in confocal microscopes or evanescent waves in total internal reflection (TIR) microscopy was devised to improve the signalto-noise ratio, the minimal spot size in such far-field imaging settings is inherently many times larger than the target molecule and therefore vulnerable to unwanted signals from adjacent sources. Dilution of the target solution can be used effectively
(9) Fogler, H. S., Elements of Chemical Reaction Engineering. 3rd ed.; PrenticeHall PTR: Upper Saddle RiVer, NJ, 1999. (10) Ha, T. J.; Ting, A. Y.; Liang, J.; Caldwell, W. B.; Deniz, A. A.; Chemla, D. S.; Schultz, P. G.; Weiss, S. Proceedings of the National Academy of Sciences of the United States of America 1999, 96, 893–898. (11) Lesoine, J.; Pal, P.; Lieb, A.; Holmberg, B.; Novotny, L.; Knauf, P. Biophys. J. 2005, 88, 665A–665A. (12) Lu, H. P.; Xun, L. Y.; Xie, X. S. Science 1998, 282, 1877–1882. (13) Michalet, X.; Weiss, S.; Jager, M. Chemical ReViews 2006, 106, 1785– 1813. (14) Rhoades, E.; Gussakovsky, E.; Haran, G. Proceedings of the National Academy of Sciences of the United States of America 2003, 100, 7418–7418. (15) Talaga, D. S.; Lau, W. L.; Jia, Y. W.; DeGrado, W. F.; Hochstrasser, R. M. Biophys. J. 2000, 78, 401A–401A. (16) Zhuang, X. W.; Bartley, L. E.; Babcock, H. P.; Russell, R.; Ha, T. J.; Herschlag, D.; Chu, S., A single-molecule study of RNA catalysis and folding. Science 2000, 288, 2048. (17) Sabanayagam, C. R.; Eid, J. S.; Meller, A. Appl. Phys. Lett. 2004, 84, 1216–1218. (18) Boukobza, E.; Sonnenfeld, A.; Haran, G. Journal of physical chemistry B 2001, 105. (19) Sun, W. C.; Gee, K. R.; Haugland, R. P. Bioorg. Med. Chem. Lett. 1998, 8, 3107–10.
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Figure 5. (a) Fluorescent intensity measurements as a function of time for increasing DiMFUp concentrations. (b) Lineweaver-Burke analysis calculating the free biotin-AP enzymatic kinetic parameters (R2 ) 0.9985).
Figure 6. (a) Fluorescence intensity measurements in consecutive DiMFUp concentrations. (b) Lineweaver-Burke analysis measuring nanoneedlebound AP enzymatic kinetic parameters (R2 ) 0.9984).
NanoDrop Wilmington, DE), and frozen as aliquots at -20 °C until needed. A priming buffer of 50 mM Tris at pH 8, 2% DMSO, 0.01% Tween 20, and 0.1% BSA was used to prevent the nonspecific adhesion of AP-biotin to the channel walls. These reagents were obtained from Sigma-Aldrich (St. Louis, MO). All kinetic measurements were conducted in 50 mM Tris at pH 8. Fabrication of Nanoneedles. The fundamentals of forming nanoneedle structure came from the observation of the strong dielectrophoretic response of carbon nanotubes in aqueous environments. When a voltage is applied across a solution of carbon nanotubes, they align themselves along the direction of the electric field. Principal to the joint problems of isolation, immobilization, and subsequent controlled stimulation and real-time observation of single proteins is the utilization of minimally perturbing nanoscale probes, or nanoneedles, whose reliable fabrication and usefulness have recently been developed.20 These needles are composed of carbon nanotubes (CNTs),21 and the manufacturing protocol of these nanoneedles was similar to that reported in earlier work.20 Multiwalled carbon nanotubes were fabricated through the chemical vapor (20) Kouklin, N. A.; Kim, W. E.; Lazareck, A. D.; Xu, J. M. Appl. Phys. Lett. 2005, 87, 173901–173903. (21) Iijima, S. Nature 1991, 354, 56–58.
deposition method and single-walled nanotubes were obtained commercially from Nanolab (Newton, MA). The following describes the construction procedure for nanoneedles. A droplet of carbon nanotube solution (water) in 15% ethanol was placed upon a polydimethylsiloxane (PDMS) slide. Here, carbon nanotube solutions with concentrations of ∼1 nM were used for the efficient production of carbon nanotube nanoneedles. Two tungsten microelectrodes were inserted into the solution and positioned parallel to each other (tip to tip). Tungsten microprobes were obtained from MicroManipulator, (Carson City, NV). They were tapered and 1 µm in diameter at the tip. After voltage (an ac frequency of 3 MHz, 4 V peak-to-peak combined with a dc offset of 6 V, 3 V peak-to-peak) was applied, one of the electrodes was slowly pulled apart from the other one and out of the solution. The existing ethanol in the carbon nanotube solution reduced the aggregation of carbon nanotubes. This in turn facilitated the production of smooth nanoneedles with small diameters. It is important to note that the voltage applied across the microprobes through the whole manufacturing process was a combination of ac and dc voltages. Whereas the frequency and amplitude of the applied ac voltage affected the diameter of the nanoneedle produced, the amplitude of the dc voltage and the carbon nanotube solution concentration affected the overall length of the final structure. The
Nanoneedle Method for Protein ActiVity Monitoring quality of the nanoneedles was based on their straightness, sharpness, and lack of kinks as visually observed from many SEM images of manufactured nanoneedles under different conditions. Figure 1a shows an SEM image of a sharp nanoneedle device with an aspect ratio of about 1500 (l/d ) 20 µm/13 nm ≈ 1500) assembled from carbon nanotubes dispersed in solution and anchored atop a microprobe for ease of handling. Figure 1b shows that as the two microprobes were slowly pulled apart from each other the surface tension of the liquid and the hydrophobicity of the carbon nanotubes coerce the tubes into a needlelike structure. It was observed that by using a strong acid solution of concentrated nitric acid and sulfuric acid in a 1:3 ratio that the carbon nanotubes were made completely water-soluble. It was difficult to make nanoneedles out of such solutions. Because the carbon nanotube water solubility perhaps resulted from a loss of hydrophobicity, it disallowed an effective aggregation of nanotubes necessary for the formation of nanoneedles. Such aggregation may have been caused by strong hydrophobic interaction between carbon nanotubes in solution. Immobilization of AP on a Nanoneedle Tip. The carboxylic groups (COOH) created on the truncated tips of the carbon nanotubes comprise the nanoneedle link to and the means for the immobilization of the biomolecules by amine-carboxylic reaction chemistry. The carboxylic groups on the nanoneedle tip were acid activated (in a solution of 12 mg/mL N-hydroxy sulfosuccinimide sodium salt (NHS) and 8 mg/mL 1-ethyl-3-(3-dimethylaminopropyl)-carbodiimide hydrochloride (EDC) in a solution of 0.1 M 2-(Norpholino) ethanesulfonic acid (MES) and 0.5 M sodium chloride at pH 6 in 1:1:2 ratio) for later enzyme conjugation. This step was followed by adding streptavidin (2 mg/mL in 100 mM sodium phosphate buffer at pH 7.0, New England Biolabs, Beverly, MA) so that streptavidin was covalently bonded to the activated nanoneedle. Then the streptavidin-attached nanoneedle was dipped in a solution of biotin-AP, which rapidly forms a tight biotin-streptavidin linkage and functionalizes the nanoneedle with AP. The high conductivity of the nanotube nanoneedle permits the use of an electric field to attract enzymes to the tip where the electrical field is the strongest, as shown by the inverse relationship between the field and the square of the radius of curvature, and thereby further enhances tip-bound immobilization. This provides us an additional degree of freedom that one can exploit, as we did here. Specifically, a positive voltage of 100 mV was applied to the nanoneedle to attract negatively charged proteins in the solution in contact with the tip via an electrostatic field. Whereas the electrostatic field was initially used for the first few seconds to attract the AP molecules to the nanoneedle, the electrostatic field was removed for rest of the AP immobilization process. The nanoneedle with attached molecules was then washed away using many consecutive washes. Hence, it is likely that the only remaining immobilized AP molecules were covalently bound to the nanoneedle. As illustrated in Figure 2a by using a nanoneedle, protein immobilization and the subsequent isolation of a few molecules at the needle tip can be accomplished. Shown in the schematics of Figure 2b is the near “point contact” topology that helps keep the proteins in their natural conformation. Unlike immobilization schemes on a planar surface with only top-entry access of a limited “solid angle”,11 in this nanoneedle setting substrates can access the enzymes immobilized on the tip within a solid angle that is twice as large. Whereas planar surface immobilization reduces enzyme activity (as evidenced by elevated Michaelis constants when compared to that of a solution-phase enzyme reaction), we think that this is much less the case for proteins immobilized on the nanoneedle tip. We verify this hypothesis by measuring the kinetic properties of nanoneedleimmobilized enzymes and then comparing to a solution-phase enzyme reaction. Figure 3a shows the fluorescent microscope image of immobilized fluorescein-conjugated AP molecules on the tip of a nanoneedle made of multiwalled carbon nanotubes. To avoid a possible false impression, we note that the fluorescence image itself was taken prior to the subsequent step of multiple washes. It helped to give a clearer image of the whereabouts of the immobilization, but without clarification it could also lead to the false impression of a large
Langmuir, Vol. 24, No. 19, 2008 10789 amount of immobilized enzymes, most of which would be washed off after several washing cycles. Note that the number of attached fluorescent AP molecules left after washing was too small to be detected with fluorescence imaging, even using confocal imaging. Because the kinetic experiments showed the enzyme activity in many consecutive trials in and out of solution using the same tip, it was established that the AP molecules not only were on the nanoneedle but also were firmly immobilized and were successfully transferred to different solutions using the nanoneedle. It is important to note that the nanoneedle is an assembly of many carbon nanotubes and the needle apex consists of multiple opened ends of integrated carbon nanotubes. Hence, the activated carbon groups were likely to exist on the tips of the carbon nanotubes, and by going through multiple washing steps, most of the nonspecific absorbed proteins bound to the side walls were removed. Hence, the remaining proteins were most likely immobilized on and surrounding the tips of the component carbon nanotubes. Note that the attachment of the AP molecules on the defect sites still cannot be excluded, but the exposed carboxylic bonds on the tip at least suggest the preferential attachment of AP molecules on the tip. Given the size of the singlewalled carbon nanotube tip (∼1 nm) or wall thickness of multiwalled carbon nanotubes (1-5 nm) with a typical protein size (∼5 nm) (specifically, 6 nm for the AP proteins used in this study), the point access immobilization (Figure 4) is consistent with findings from prior and independent experiments.22 As a negative control, it was verified that when carboxylic groups were capped, nonspecifically adsorbed AP was not detected, as evidenced by the lack of a fluorescent output signal. Conversely, the nanoneedle with reactive carboxylic end groups produced a fluorescent signal for 20 consecutive kinetic trials, thus supporting the notion of immobilization on the terminal group of the individual nanotubes that comprise the nanoneedle. The nonspecific adsorption of proteins to the sidewall of the nanotube nanoneedle was reduced by coating the side walls with blocking agents (bovine serum albumin and arabic gum22). This further disallowed the attachment of AP molecules to the side wall of the nanoneedle. Figure 3b illustrates the method used in this work to measure the activity of immobilized biomolecules on the nanoneedle by dipping it into the solution of the target reactant molecules. Here, the enzymeconjugated needle was removed from biotin-AP solution bath, washed multiple times, and then transported and placed in a separate cuvette of DiFMUp solution.
3. Results and Discussion The Michaelis constants for both free and nanoneedle-bound biotin-AP were estimated using Lineweaver-Burke analyses. As a control for the kinetics assay, solution-phase kinetics was first assayed at a fixed enzyme concentration of 360 pM in seven serial dilutions of DiFMUp from 100 nM to 100 µM in NEB3 buffer (100 mM NaCl, 50 mM Tris-HCl, 10 mM MgCl2, 1 mM dithiothreitol, New England Biolabs, Beverly, MA) containing 1% BSA at pH 8. The upper and lower limits were chosen to capture zeroth- and first-order reaction data at low conversion for the Lineweaver-Burke analysis. Baseline fluorescence of the DiFMUp substrate was measured in a 50 µL quartz cuvette (16.50F-Q-10/Z8.5, Starna cells, Atascadero, CA) filled to 60 µL. Because we use biotin-streptavidin biochemistry to attach AP to the nanoneedle tip, we measured the activity of free biotinconjugated AP proteins as the control. The cuvette was maintained at 25 °C using a digital temperature controller. The temperature was kept constant at 25 C to minimize the evaporation effects. Note that the experiments can be performed at any other temperature. The fluorometer reading was paused, the DiMFUp substrate was removed and mixed with 5 µL of biotin-AP, and the entire well-mixed volume was returned to the cuvette for (22) Withey, G. D.; Lazareck, A. D.; Tzolov, M. B.; Yin, A.; Aich, P.; Yeh, J. I.; Xu, J. M. Biosensors & Bioelectronics 2006, 21, 1560–1565.
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continued measurement. Following each measurement, the cuvette was thoroughly washed with DI water, rinsed with 95% ethanol, and dried under nitrogen such that the cuvette windows were free from streaks. Figure 5a shows fluorescence versus time plots for various runs. The fluorescence increase is proportional to the rate of production of the fluorescent DiFMU molecules. The data were collected over the first 6 min. The initial linear enzymatic reaction velocities (ν ) νmax[DiMFUp]/(Km + [DiMFUp])) were then calculated using the data from the first 100 s of each curve. Figure 5b shows the Lineweaver-Burke analysis23 where the inverse reaction velocities are plotted against the inverse substrate (DIFMUp) concentrations. Each experiment was carried out three times to obtain the respective error bars. The Michealis constant for solution-phase AP-biotin was evaluated to be Km ) 5.83 ( 0.23 µM. Finally, the immobilized phase kinetics was assayed using an enzyme-conjugated nanoneedle in five serial dilutions of DIFMUp. The “loaded” nanoneedle was moved into different concentrations of DiMFUp solution for consecutive enzyme reaction measurements. The translocation proceeded from low to high concentration (five consecutive concentrations from 0.1 to 100 µM) to avoid any carryover. The success of this procedure also validated that the proteins were indeed immobilized onto the tip by conjugation because loosely adsorbed proteins would likely be lost in the process. Figure 6a shows the fluorescence intensity versus time measurements of produced DiMFU after inserting the nanoneedle into different solutions of DiMFUp. The data were collected over the first 5 min. The initial linear enzymatic reaction velocities were then calculated using the data from the first 100 s of each curve. Similar to solution-phase (control) experiments, the Lineweaver-Burke analysis determined a Km of 5.06 ( 0.2 µM for the nanoneedle-bound proteins (Figure 6b). This value is remarkably close to that obtained for free biotin-AP in the solution phase. This indicates no apparent degradation in the activity of the protein immobilized either through steric hindrance of the catalytic site or deactivation at the tip of the nanoneedle. This suggests that the enzyme molecules on the nanoneedle surface were accessible to the reactant molecules. In contrast, immobilized AP proteins bound to the (23) Segel, I. H., Enzyme Kinetics: BehaVior and Analysis of Rapid Equilibrium and Steady State Enzyme Systems Wiley: New York, 1993.
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surface of microbeads had a Km value that was 6 times greater than that of free AP measured under the same conditions.24 The experiments reported here were conducted using an ensemble of proteins immobilized on the nanoneedle tip. Physically, the number of immobilized proteins in this nanoneedle platform can approach a few proteins with a shorter exposed length at the tip after the coating, or shorter loading time during the immobilization steps, or a smaller voltage applied to the tip, or a lower concentration of the target protein solutions. Although the number of immobilized biotin-AP molecules could be lowered all the way down to the single-molecule level, in this work the number of immobilized molecules was limited by the sensitivity of the fluorometer apparatus and by the finite viable lifetime or stability of the alkaline phosphatase enzyme. In this particular setting, a single enzyme would be unable to catalyze the production of a sufficient quantity of DiMFU for a detectable fluorescent signal in a 60 µL test volume during the protein viability period.
4. Conclusions To summarize, we describe a novel nanoneedle platform to provide near-ideal immobilization and isolation conditions that offer the potential for single protein investigations. By comparing the reaction kinetics of the test proteins (biotin-AP) in free unbounded and bounded conditions, we found that the immobilization and isolation of a few proteins near the nanoneedle tip are not only feasible but also free of planarsurface-induced denaturation. Hence, high-sensitivity, lowbackground, real-time enzymatic reactions can be performed by incorporating this platform into existing detection devices without inducing unwanted conformational changes and modifications to the proteins under study. Acknowledgment. We thank Jin Ho Kim for his extremely helpful suggestions and insights. The part of this work performed by M.J. and J.X. is made possible by support from the AFOSR MURI. A.T. acknowledges the financial support of the National Science Foundation (grant no. BES-0555874) for this research. LA703630A (24) Kerby, M. B.; L, R. S.; Tripathi, A. Anal. Chem. 2006, 78, 8273–8280.
