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Proceedings of the National Academy of Sciences of the United States of America logoLink to Proceedings of the National Academy of Sciences of the United States of America
. 2002 Apr 2;99(8):5018–5023. doi: 10.1073/pnas.082644099

Inhibition of chymotrypsin through surface binding using nanoparticle-based receptors

Nicholas O Fischer †,‡, Catherine M McIntosh ‡, Joseph M Simard ‡, Vincent M Rotello †,‡,§
PMCID: PMC122714  PMID: 11929986

Abstract

Efficient binding of biomacromolecular surfaces by synthetic systems requires the effective presentation of complementary elements over large surface areas. We demonstrate here the use of mixed monolayer protected gold clusters (MMPCs) as scaffolds for the binding and inhibition of chymotrypsin. In these studies anionically functionalized amphiphilic MMPCs were shown to inhibit chymotrypsin through a two-stage mechanism featuring fast reversible inhibition followed by a slower irreversible process. This interaction is very efficient, with a KInline graphic = 10.4 ± 1.3 nM. The MMPC–protein complex was characterized by CD, demonstrating an almost complete denaturation of the enzyme over time. Dynamic light scattering studies confirm that inhibition proceeds without substantial MMPC aggregation. The electrostatic nature of the engineered interactions provides a level of selectivity: little or no inhibition of function was observed with elastase, β-galactosidase, or cellular retinoic acid binding protein.

Synthetic receptors targeted at protein surfaces provide an alternative paradigm to active site inhibition for regulating enzyme activity. Additionally, protein surface recognition provides a means for modulating the numerous protein–protein (1, 2) and protein–nucleic acid (3) interactions that are central to cellular processes. Optimal binding of the convex and solvent-exposed surface of proteins, however, requires large preorganized surfaces for the presentation of complementary recognition elements. These two factors make the design and synthesis of surface receptors a challenging goal, which has been addressed to date by using peptide and polymer systems (1, 2, 4–6), and large monomeric receptors (refs. 4 and 7 and refs. therein).

Mixed monolayer protected gold clusters (MMPCs) provide an effective scaffold for biomolecular binding. These nanoparticles are readily formed through reduction of tetrachloroauric acid in the presence of thiol capping agents (8) (Fig. 1). Through control of aurate-thiol stoichiometry, MMPCs featuring core diameters from 2 to 10 nm can be readily fabricated (9, 10), providing suitably scaled systems for biomacromolecular interactions. Nanoparticles featuring a wide variety of surface functionality can be created through use of functionalized thiols as capping agents (11). The diversity of surface functionality can be further enhanced through the introduction of additional substituents in subsequent place-displacement reactions, allowing divergent and potentially combinatorial synthesis of nanoparticle systems (12).

Figure 1.

Figure 1

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Fabrication of MMPCs by using the Brust reduction and the Murray place-displacement reaction to introduce additional functionalized thiols.

In addition to the size and diversity of surface functionality possible with nanoparticles, the surface properties of MMPCs provide unique opportunities for the control of surface interactions. First, the thiols are mobile on the surface of the MMPCs, a feature that has been used to create self-optimizing multivalent receptors (13). Additionally, the faceted surface of these nanoparticles results in a radial dependence of monolayer packing (9), allowing the further fine-tuning of monolayer structure through thiol chain length (14). These facets can also be approximated as two-dimensional self-assembled monolayers (9), facilitating the solution study of many biological systems and processes that are difficult to study with surface techniques.

In recent studies we have demonstrated the effective binding of amphiphilic cationic MMPCs to DNA and established the viability of these systems for gene delivery (15) and the inhibition of transcription (16). To establish the utility of nanoparticles for protein surface binding, we explored the interactions of the gold particles with α-chymotrypsin (ChT) (for a previous example of ChT inhibition using surface recognition, see ref. 17). The active site of ChT is immediately surrounded by hydrophobic residues and further by a ring of cationic residues (Fig. 2) (18), providing a target for the creation of complementary amphiphilic binding surfaces. Additionally, the enzymatic activity of ChT is well characterized (19, 20), making this protein a particularly attractive target for in vitro analysis.

Figure 2.

Figure 2

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(a) Space-filling model of ChT. Surface binding of the protein by anionic MMPCs focuses on the ring of cationic residues situated around the active site. Functionally significant residues are noted. (b) Relative sizes of ChT and MMPC 1.

To explore the binding and concomitant inhibition of ChT by using nanoparticle-based receptors, we fabricated MMPCs 1-3 (Fig. 3) through place exchange of the corresponding thiols into octanethiol-capped nanoparticles (11). MMPCs 1-3 feature a 2-nm gold core, with an overall diameter of 6 nm; all three nanoparticles are amphiphilic in nature, with the monolayer composed of both charged and hydrophobic thiols. MMPCs 1 and 2 are functionalized with carboxylate groups designed to target the cationic surface residues of ChT. In this article, we report the highly efficient binding and inhibition of ChT by these anionic nanoparticles.

Figure 3.

Figure 3

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Anionic MMPCs 1 and 2 and cationic control 3.

Materials and Methods

General.

α-ChT (type II from bovine pancreas), elastase (type I from porcine pancreas), β-galactosidase (grade VI from Escherichia coli), N-succinyl-Ala-Ala-Ala-p-nitroanilide, o-nitrophenyl β-d-galactopyranoside, benzoyl tyrosine p-nitroanilide, and N-succinyl-Ala-Ala-Pro-Phe-p-nitroanilide (Suc-AAPF-pNA) were purchased from Sigma. Hydrogen tetrachloroauric acid was obtained from Alfa Aesar, Ward Hill, MA. All other chemicals were purchased from Aldrich and used as received.

Nanoparticle Fabrication.

The acid functionalized nanoparticles (MMPCs 1 and 2) were synthesized by using our previously published procedure (21): 11-mercaptoundecanoic acid was added in different amounts (500 and 250 mg for 1 and 2, respectively) to octanethiol-functionalized nanoparticles (50 mg) in 6 ml of tetrahydrofuran. Argon was bubbled through each solution for 10 min, and the reactions were stirred for 2 days under argon. The solutions were then reduced in vacuo and washed three times with 50 ml of dichloromethane, and the precipitate was collected by centrifugation. MMPC 3 was prepared as described (16). The functional group loading of each of the particles was determined by NMR endgroup analysis.

Activity Assays.

ChT was preincubated with varying concentrations of nanoparticles (50 nM–800 nM) in 225 μl of 5 mM sodium phosphate (pH 7.4). Activity assay was modified from a reported procedure (17). Enzyme concentration was kept constant at 800 nM. At established time points, 25 μl of benzoyl tyrosine p-nitroanilide stock solution was added, resulting in a final reaction concentration of 100 μM. Activity was followed by monitoring product formation every 15 s for 5 min at 405 nm with a MRX Revelation plate reader (Dynex Technologies, Chantilly, VA). Assays for cellular retinoic acid binding protein (22), elastase (23), and β-galactosidase (24) were modified from previously described procedures. Temperature was maintained at 25°C during assays and preincubation periods.

CD.

Samples were prepared in deionized water (pH 7.4, adjusted with NaOH). MMPC 1 was diluted from a stock solution of 43 μM to final concentrations between 84 nM and 2.1 μM. ChT was prepared from a dry powder to 3.2 μM final concentration. SDS was prepared from a dry powder to 10 mM final concentration. CD was performed on a Jasco 700 spectrophotometer, using a quartz cuvette with a 1-mm path length. Scans were taken from 190 to 250 nm at a rate of 5 nm/min, with a 0.1-nm step resolution and a 4-s response. Three scans were averaged at a constant temperature of 20°C, with a 5-min equilibration before the scans. Samples sat at room temperature overnight before reaching the 24-h time point. SDS samples were mixed approximately 30 min before scanning to allow equilibration. In a few cases, nanoparticle blanks that directly overlapped with another spectrum were measured at a rate of 10 nm/min for two scans (all other parameters were unchanged). Data obtained with high tension voltage (HT) over 600 V were not included in the analysis.

Dynamic Light Scattering.

MMPC 1 was diluted from a stock solution of 43 μM to a final concentration of 200 or 43 nM. ChT was prepared from a dry powder to a final concentration of 800 nM or 10 μM. Samples were diluted by using ddH2O filtered through Acrodisc 0.2-μm filters (Pall Gelman Laboratory, Ann Arbor, MI). MMPC 1, ChT, and MMPC 1–ChT solutions were examined with an argon laser tuned to 514 nm with ALV-5000 software. Data were collected for 30 s, and scans were repeated until scattering caused by transient dust was eliminated. Data were transformed by applying a regularized fit with the maximum data points acceptable for each scan. At least three scans were averaged for each sample.

Kinetic Assays.

Stock concentrations of ChT and Suc-AAPF-pNA were determined in H2O by using extinction coefficients (ChT, ɛ280 = 51,000; N-Suc, ɛ315 = 14,000) (25, 26). Progress curve reactions were buffered in 5 mM sodium phosphate (pH 7.4). A 15 mM stock of Suc-AAPF-pNA was prepared in DMSO. Final DMSO concentration did not exceed 1%. To initiate reactions, 100 μl of ChT was added to 900 μl of substrate-nanoparticle solution, resulting in final reaction concentrations of 10 nM ChT and 150 μM Suc-AAPF-pNA, respectively. Readings were taken in duplicate every 20 s for 6 min, conditions under which the control plot was linear. Temperature was held constant at 25°C.

Progress Curve Analysis.

Progress curves were used to analyze the kinetics of MMPC 1 inhibition (27). The apparent Ki was calculated from initial velocity studies at MMPC 1 concentrations between 1 and 10 nM. Progress curves were fit to the following equation for slow binding inhibitors that bind either very tightly or irreversibly:

graphic file with name M2.gif 1

where vs is the steady-state velocity, vi is the initial velocity, t is time, and kobs is the pseudo-first-order rate constant. Depletion of inhibitor and enzyme are accounted for by γ, given as:

graphic file with name M3.gif 2

where [Et] and [It] are total concentrations, and v0 is uninhibited velocity. Slow, tight binding inhibitors are described by the following scheme:

graphic file with name M4.gif 3

where E*I is the complex of isomerized enzyme and inhibitor. However, if k6 is very small compared with k5, or is zero, the reaction can be regarded as irreversible. To determine the apparent Ki, the following equation was used (28):

graphic file with name M5.gif 4

Results

Activity Assays.

The inhibitory effects of MMPCs 1-3 on ChT activity were determined after various preincubation periods with the MMPCs (Fig. 4a). The anionic nanoparticles 1 (92% carboxylate coverage) and 2 (68% coverage) strongly inhibited ChT activity, with MMPC 1 displaying more efficient inhibition. Cysteine-nanoparticle interactions will not interfere with the binding process because all cysteine residues in ChT are involved in disulfide bonds, which have been shown not to displace the thiol monolayer (12). The decreased binding of MMPC 2 is expected based on the lesser number of anions available to complement the ChT surface; although reorganization of the monolayer to optimize the binding may occur over the time scale of the experiment, the resultant nanoparticle would be expected to support a lower stoichiometry of binding, thus displaying less inhibition than MMPC 1 at the same concentration. Cationic MMPC 3, as expected, had no detectable inhibitory effects.

Figure 4.

Figure 4

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Time course of ChT activity after preincubation with MMPCs ([ChT] = 800 nM). (a) Anionic nanoparticles 1 and 2 and cationic control MMPC 3. (b) Concentration dependence of MMPC 1 inhibition. Lines are provided to lead the eye.

A relatively high degree of selectivity was obtained through electrostatic complementarity. For example, virtually no inhibition of elastase was observed by using MMPC 1 (less than 10% inhibition as compared with more than 80% with ChT at the same molar ratio) (17). Additionally, no initial inhibition occurred with β-galactosidase, with very slow loss of activity observed over extended periods, potentially suggesting a reorganization of the nanoparticle monolayer over the 24-h time course of the study. As a final direct probe of structural changes upon addition of MMPC 1, we examined the fluorescence of cellular retinoic acid binding protein (15 kDa). The maximum wavelength and intensity of tryptophan fluorescence and the binding of retinoic acid both provide sensitive probes to examine the function and conformational state of the protein (22). In our studies, addition of MMPC 1 caused only a slight shift in λmax at higher colloid/protein ratios (4- to 8-nm shift, as opposed to 20-nm shifts that occur upon denaturation). Moreover, binding of the retinoic acid ligand is unperturbed even after a 24-h incubation, indicating that the protein structure and function remain intact.

Because of the substantial degree of ChT inhibition, MMPC 1 was chosen for further characterization. Varying concentrations of this nanoparticle were preincubated with ChT to determine the effects of concentration and preincubation time on enzyme activity (Fig. 4b). The addition of equimolar concentrations of MMPC 1 resulted in near complete inhibition in less than 600 min. Complete inhibition was also observed at 48 h at a 1:5 MMPC 1/ChT ratio, indicating that MMPC 1 was capable of concurrently binding five molecules of ChT.

Structural and Conformational Analysis of MMPC 1–ChT Assemblies.

CD was performed to determine the effects of surface binding on ChT conformation (Fig. 5). The spectrum observed for wild-type ChT was the same as reported (29). The conformation of 3.2 μM ChT was measured after initial mixing with MMPC 1 and a 24-h incubation at room temperature. Addition of either 0.8 or 2.1 μM MMPC 1 effected a substantial increase in the intensity of the minimum at 202 nm. Upon further incubation, this minimum was shifted to lower wavelengths, indicating a conformational shift toward random coil and a loss of native secondary structure. In addition, the characteristic minimum at 232 nm was lost upon mixing with the higher concentration of nanoparticle, an effect also observed upon thermal denaturation of the protein. The loss of this feature was observed to a lesser extent in the 0.8 μM MMPC 1 sample. The complete loss of structure observed for ChT upon incubation may be facilitated by the fact that 70% of surface-exposed ChT residues are nonpolar, as compared with ≈55% in most other proteins (30). The exposure of these additional hydrophobic groups to the buried alkane chains in the monolayer may increase the driving force for denaturation.

Figure 5.

Figure 5

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CD of ChT ([ChT] = 3.2 μM). (a) After initial mixing. (b) After 24-h incubation and thermal denaturation (green trace).

Dynamic light scattering was performed to determine the extent of aggregation of the ChT–MMPC 1 assemblies. The hydrodynamic radius of the nanoparticles at a 1:4 MMPC 1/ChT ratio (200 nM:800 nM) was the same within experimental error in the absence or presence of ChT. The nanoparticle is at the limit of size scale for the dynamic light scattering, so quantification of size changes upon protein binding was inconclusive. After an extended incubation of 330 min, the radius was unchanged, indicating that size of the ChT–MMPC 1 complex is not time dependent. The absence of large assemblies in these studies suggest that aggregation is not a factor in the inhibition observed.

Kinetic Assays.

With evidence pointing to irreversible or multistep time-dependent inhibition of ChT by carboxylate functionalized nanoparticles, the inhibition kinetics of the nanoparticle were determined. The activity assays, normalized to the control, indicate that there is an initial inhibition followed by a second phase of time-dependent inhibition. To investigate this further, progress curves were studied at different concentrations of MMPC 1 (Fig. 6a). Suc-AAPF-pNA was chosen as a more sensitive substrate to study initial product formation. Using this substrate to determine inhibition in the first 6 min of interaction, the pseudo first-order rate constant (kobs) was found for each concentration of nanoparticle while keeping ChT and substrate concentrations fixed at 10 nM and 150 μM, respectively. For purposes of calculation, each nanoparticle was treated as five independent inhibitors, as there was no evidence for positive or negative cooperativity in the kinetic data. Fitting the data to Eq. 4, the apparent Ki of the nanoparticle was 10.4 ± 1.3 nM with a rate constant (k5) of 0.0743 s−1 (Fig. 6b). The zero intercept of this fitted curve indicates, as expected, that the inhibition is either irreversible or that the reversible step (k6) is infinitely small compared with the forward reaction. A double reciprocal plot of these data results in a nonzero intercept (Fig. 7). This finding is indicative of a two-step mechanism of inactivation; the initial binding step is followed by a slower inactivation event (28).

Figure 6.

Figure 6

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Kinetic analysis of MMPC 1 inhibition. (a) Progress curves of ChT activity illustrating the decrease in activity and product formation over time as MMPC 1 concentration is increased. Concentration of ChT was fixed at 10 nM. Chromogenic substrate N-Suc-AAPF-pNA concentration was kept at 150 μM. MMPC 1 concentrations were 1, 2, 5, and 10 nM. (b) Plot of kobs vs. MMPC 1 concentration, from which KInline graphic was calculated by nonlinear curve fitting. Calculations were completed by assuming a 1:5 nanoparticle/ChT binding ratio.

Figure 7.

Figure 7

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Reciprocal plot of 1/kobs vs. 1/[MMPC 1], showing nonzero intercept indicative of a two-step inactivation mechanism.

Discussion

Gold colloids have been conjugated to biomolecules and used for a variety of applications (31–33). There have been, however, few examples where the particle has been used as a scaffold for the multivalent presentation of binding elements (15, 16). The design of the anionic nanoparticle presented here applies simple electrostatic and hydrophobic interactions to explore fundamental aspects of binding, providing a starting point for the creation of more complex and selective receptors.

The size and shape of the colloid are similar to detergent micelles, with the polar head groups coupled with the nonpolar interior of the alkane chains providing structural similarities to our particles. However, many of the proposed modes of SDS–protein interaction (ref. 34 and references therein, and ref. 35) are not possible in MMPC systems because of the higher order of preorganization provided by the nanoparticle core. The insertion of single detergent molecules into the protein interior to yield SDS clusters, for example, requires an inherent flexibility that the MMPC system does not display. An alternative SDS model, termed the “protein-decorated micelle” model, indicates that an intact micelle is encased by the hydrophilic protein (35). Although this model may describe the gross features of MMPC–ChT binding, the nanoparticles display several features that still distinguish them from SDS denaturation. Each nanoparticle behaves as a single macromolecule independent of concentration or availability of a protein target, allowing control over the macroscopic features offered to a selected guest. SDS nondiscriminately denatures all proteins (36), whereas MMPC 1 shows selectivity in its interaction with several proteins (see above). In addition, the ability of the colloids to be multifunctionalized extends their potential applications beyond that of simple electrostatic pairing.

Structural changes occurring upon denaturation of ChT confirm that the mechanism of the nanoparticle–protein interaction is distinct from that of SDS denaturation. The CD of ChT shows a decrease in α-helical content and an increase in random coil either upon binding to MMPC 1 or thermal denaturation. In contrast, SDS-mediated denaturation dramatically increases α-helix content (37). The relative changes in helical and random coil content in the presence of MMPC 1 and SDS are mirrored in the spectra of ChT adsorbed onto hydrophilic and hydrophobic colloids (29), indicating that the conformational change must be driven by differing forces. Although the initial mechanism likely in each of the two systems would suggest that an alternate mode of denaturation is occurring, we cannot conclude based on the current data that the kinetics observed in the two systems are different (38).

Noncovalent colloidal hosts will not only allow the development of a new class of protein inhibitors both for traditional enzymes and protein–protein interactions, but will also provide a tool for studying protein adsorption at surfaces (39, 40). Because self-assembled monolayer (SAM) protected gold colloids approximate two-dimensional SAMs (9), they can be used to study the mechanisms of protein adsorption and the resultant conformational and functional changes by using solution techniques difficult or impossible to adapt to surface studies (e.g., NMR and CD spectroscopy). In addition, the MMPC platform offers advantages over other colloidal systems because of the ease of functionalization and the spectral handles available to observe the colloids specifically.

In summary, we have shown that anionic functionalized nanoparticles are highly effective surface-based inhibitors of ChT by a two-step inactivation mechanism. This inhibition occurs through electrostatic complementarity between the carboxylate endgroups and the halo of cationic residues located around the periphery of the active site; cationic MMPCs showed no activity with ChT. Complete inhibition of enzyme activity was observed at a 1:5 nanoparticle to ChT ratio, arising from the large area of the anionic nanoparticle surface (≈110 nm2).

Acknowledgments

We are grateful for assistance from U. Serdar Tülü for preliminary studies. Cellular retinoic acid binding protein and retinoic acid were the generous gift of Jennifer Habink, Univ. of Massachusetts. This research was supported by the National Science Foundation (Materials Research Science and Engineering Center instrumentation) and the National Institutes of Health (GM 62998). V.M.R. acknowledges support from the Alfred P. Sloan Foundation, Research Corporation, and the Camille and Henry Dreyfus Foundation. N.O.F. and C.M.M. acknowledge support from National Institutes of Health Chemistry-Biology Interface Training Grant GM 08515.

Abbreviations

ChT

chymotrypsin

MMPC

mixed monolayer protected gold cluster

Suc-AAPF-pNA

N-succinyl-Ala-Ala-Pro-Phe-p-nitroanilide

Footnotes

This paper was submitted directly (Track II) to the PNAS office.

References

  • 1.Ghosh I, Chmielewski J. Chem Biol. 1998;5:439–445. doi: 10.1016/s1074-5521(98)90160-0. [DOI] [PubMed] [Google Scholar]
  • 2.Schramm H J, de Rosny E, Reboud-Ravaux M, Buttner J, Dick A, Schramm W. Biol Chem. 1999;380:593–596. doi: 10.1515/BC.1999.076. [DOI] [PubMed] [Google Scholar]
  • 3.Nadassy K, Wodak S J, Janin J. Biochemistry. 1999;38:1999–2017. doi: 10.1021/bi982362d. [DOI] [PubMed] [Google Scholar]
  • 4.Zutshi R, Brickner M, Chmielewski J. Curr Opin Chem Biol. 1998;2:62–66. doi: 10.1016/s1367-5931(98)80036-7. [DOI] [PubMed] [Google Scholar]
  • 5.Hayashi T, Hitomi Y, Ogoshi H. J Am Chem Soc. 1998;120:4910–4915. [Google Scholar]
  • 6.Strong L E, Kiessling L L. J Am Chem Soc. 1999;121:6193–6196. [Google Scholar]
  • 7.Peczuh M W, Hamilton A D. Chem Rev. 2000;100:2479–2494. doi: 10.1021/cr9900026. [DOI] [PubMed] [Google Scholar]
  • 8.Brust M, Walker M, Bethell D, Schiffrin D J, Whyman R J. Chem Commun. 1994;7:801–802. [Google Scholar]
  • 9.Hostetler M J, Wingate J E, Zhong C-J, Harris J E, Vachet R W, Clark M R, Londono J D, Green S J, Stokes J J, Wignall G D, et al. Langmuir. 1998;14:17–30. [Google Scholar]
  • 10.Stavens K B, Pusztay S V, Zou S H, Andres R P, Wei A. Langmuir. 1999;15:8337–8339. [Google Scholar]
  • 11.Hostetler M J, Green S J, Stokes J J, Murray R W. J Am Chem Soc. 1996;118:4212–4213. [Google Scholar]
  • 12.Hostetler M J, Templeton A C, Murray R W. Langmuir. 1999;15:3782–3789. [Google Scholar]
  • 13.Boal A K, Rotello V M. J Am Chem Soc. 2000;122:734–735. [Google Scholar]
  • 14.Boal A K, Rotello V M. Langmuir. 2000;16:9527–9532. [Google Scholar]
  • 15.Sandhu K K, McIntosh C M, Simard J M, Smith S A, Rotello V M. Bioconjug Chem. 2002;13:3–6. doi: 10.1021/bc015545c. [DOI] [PubMed] [Google Scholar]
  • 16.McIntosh C M, Esposito E A, Boal A K, Simard J M, Martin C T, Rotello V M. J Am Chem Soc. 2001;123:7626–7629. doi: 10.1021/ja015556g. [DOI] [PubMed] [Google Scholar]
  • 17.Park H S, Lin Q, Hamilton A D. J Am Chem Soc. 1999;121:8–13. [Google Scholar]
  • 18.Capasso C, Rizzi M, Menegatti E, Ascenzi P, Bolognesi M. J Mol Recognit. 1997;10:26–35. doi: 10.1002/(SICI)1099-1352(199701/02)10:1<26::AID-JMR351>3.0.CO;2-N. [DOI] [PubMed] [Google Scholar]
  • 19.Barrett A J. In: Proteinase Inhibitors. Barrett A J, Salvesen G, editors. Amsterdam: Elsevier; 1986. pp. 3–9. [Google Scholar]
  • 20.Powers J C, Harper J W. In: Proteinase Inhibitors. Barrett A J, Salvesen G, editors. Amsterdam: Elsevier; 1986. pp. 55–152. [Google Scholar]
  • 21.Simard J, Briggs C, Boal A K, Rotello V M. Chem Commun. 2000;19:1943–1944. [Google Scholar]
  • 22.Clark P L, Weston B F, Gierasch L M. Fold Design. 1998;3:401–412. doi: 10.1016/s1359-0278(98)00053-4. [DOI] [PubMed] [Google Scholar]
  • 23.Bieth J, Spiess B, Wermuth C G. Biochem Med. 1974;11:350–357. doi: 10.1016/0006-2944(74)90134-3. [DOI] [PubMed] [Google Scholar]
  • 24.Naider F, Bohak Z, Yariv J. Biochemistry. 1972;11:3202–3207. doi: 10.1021/bi00767a010. [DOI] [PubMed] [Google Scholar]
  • 25.Bru R, Walde P. Eur J Biochem. 1991;199:95–103. doi: 10.1111/j.1432-1033.1991.tb16096.x. [DOI] [PubMed] [Google Scholar]
  • 26.DelMar E G, Largman C, Brodrick J W, Geokas M C. Anal Biochem. 1979;99:316–320. doi: 10.1016/s0003-2697(79)80013-5. [DOI] [PubMed] [Google Scholar]
  • 27.Morrison J F, Walsh C T. In: Advances in Enzymology. Meister A, editor. Vol. 61. New York: Interscience; 1988. pp. 201–301. [DOI] [PubMed] [Google Scholar]
  • 28.Copeland R A. Enzymes: A Practical Introduction to Structure, Mechanism, and Data Analysis. New York: Wiley; 2000. pp. 318–349. [Google Scholar]
  • 29.Norde W, Zoungrana T. Biotechnol Appl Biochem. 1998;28:133–143. [PubMed] [Google Scholar]
  • 30.Janin J, Chothia C. J Biol Chem. 1990;265:16027–16030. [PubMed] [Google Scholar]
  • 31.Mirkin C A, Letsinger R L, Mucic R C, Storhoff J J. Nature (London) 1996;382:607–609. doi: 10.1038/382607a0. [DOI] [PubMed] [Google Scholar]
  • 32.Alivisatos A P, Johnsson K P, Peng X G, Wilson T E, Loweth C J, Bruchez M P, Schultz P G. Nature (London) 1996;382:609–611. doi: 10.1038/382609a0. [DOI] [PubMed] [Google Scholar]
  • 33.Niemeyer C M. Angew Chem Int Ed. 2001;40:4128–4158. doi: 10.1002/1521-3773(20011119)40:22<4128::AID-ANIE4128>3.0.CO;2-S. [DOI] [PubMed] [Google Scholar]
  • 34.Rao K S, Prakash V. J Biol Chem. 1993;268:14769–14775. [PubMed] [Google Scholar]
  • 35.Ibel K, May R P, Kirschner K, Szadkowski H, Mascher E, Lundahl P. Eur J Biochem. 1990;190:311–318. doi: 10.1111/j.1432-1033.1990.tb15578.x. [DOI] [PubMed] [Google Scholar]
  • 36.Jirgensons B. Biochim Biophys Acta. 1973;131:131–138. doi: 10.1016/0005-2795(73)90205-5. [DOI] [PubMed] [Google Scholar]
  • 37.Greenfield N, Fasman G D. Biochemistry. 1969;8:4108–4116. doi: 10.1021/bi00838a031. [DOI] [PubMed] [Google Scholar]
  • 38.Steinhardt J, Stocker N. Biochemistry. 1973;12:1789–1797. doi: 10.1021/bi00733a020. [DOI] [PubMed] [Google Scholar]
  • 39.Prime K L, Whitesides G M. Science. 1991;252:1164–1167. doi: 10.1126/science.252.5009.1164. [DOI] [PubMed] [Google Scholar]
  • 40.Mrksich M, Whitesides G M. Annu Rev Biophys Biomol Struct. 1996;25:55–78. doi: 10.1146/annurev.bb.25.060196.000415. [DOI] [PubMed] [Google Scholar]

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