Molecular Interaction between Organophosphorus ... - ACS Publications

Department of Chemistry, University of Miami, Coral Gables, Florida 33124-0431, United States Army. Edgewood Chemical & Biological Center, Aberdeen ...
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Biomacromolecules 2005, 6, 1555-1560

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Molecular Interaction between Organophosphorus Acid Anhydrolase and Diisopropylfluorophosphate Jiayin Zheng,† Celeste A. Constantine,† Liang Zhao,† Vipin K. Rastogi,‡ Tu-Chen Cheng,§ Joseph J. DeFrank,‡ and Roger M. Leblanc*,† Department of Chemistry, University of Miami, Coral Gables, Florida 33124-0431, United States Army Edgewood Chemical & Biological Center, Aberdeen Proving Ground, Maryland 21010-5423, and GEO-Centers, Incorporated, Gunpowder Club, Aberdeen Proving Ground, Maryland 21010 Received December 17, 2004; Revised Manuscript Received January 26, 2005

Organophosphorus acid anhydrolases (OPAA; E.C.3.1.8.2) are a class of enzymes that hydrolyze a variety of toxic acetylcholinesterase-inhibiting organophosphorus (OP) compounds, including pesticides and fluorinecontaining chemical nerve agents. In this paper, subphase conditions have been optimized to obtain stable OPAA Langmuir films, and the diisopropylfluorophosphate (DFP) hydrolysis reaction catalyzed by OPAA in aqueous solution and at the air-water interface was studied. OPAA-DFP interactions were investigated utilizing different spectroscopic techniques, that is, circular dichroism and fluorescence in aqueous solution and infrared reflection absorption spectroscopies at the air-water interface. The characterization of OPAA and its secondary structure in aqueous solution and as a monolayer at the air-water interface in the absence and in the presence of DFP dissolved in aqueous solution or in the aqueous subphase demonstrated significantly distinctive features. The research described herein demonstrated that OPAA can be used in an enzyme-based biosensor for DFP detection. 1. Introduction Highly toxic organophosphorus (OP) compounds are probably the best known and most widely used agents in recent years as pesticides, insecticides, and chemical warfare (CW) nerve agents.1-3 They are very toxic to mammals, particularly when absorbed through the skin.2,4 Laboratorybased methods which are commonly used for detection and measurement of OP compounds include gas chromatography,5,6 high-performance liquid chromatography,7,8 and capillary electrophoresis.5,9 Although these methods can measure a wide variety of OP insecticides, they are typically expensive and time-consuming and are not well-suited to field applications or process control monitoring.5 To overcome these disadvantages, enzymatic biosensors have been designed for enhanced speed of detection, high efficiency, sensitivity, and cost effectiveness.10,11 In the past decade, two bacterial enzyme systems have been identified and characterized for use in the detoxification of a variety of neurotoxins, including pesticides and G-type and V-type chemical nerve agents.12,13 Organophosphorus hydrolase (OPH; E.C.3.1.8.1), originally isolated from Pseudomonas dininuta, is able to cleave P-O, P-F, P-S, and P-CN bonds via an SN2 mechanism, resulting in hydrolysis products which change the solution pH.14 Detailed studies on surface chemistry and spectroscopic properties of OPH in solution and after immobilization, as well as * To whom correspondence should be addressed. Tel.: (305) 284-2194. Fax: (305) 284-6367. E-mail: [email protected]. † University of Miami. ‡ United States Army Edgewood Chemical & Biological Center. § GEO-Centers, Incorporated.

interaction between OPH and paraoxon, have been reported previously.11,13,15-18 Another well-characterized hydrolytic enzyme, organophosphorus acid anhydrolase (OPAA; E.C.3.1.8.2), has been demonstrated as unique for decontamination and detection of G-type nerve agents. OPAA was originally isolated from a halophilic bacterial isolate designated Alteromonas spJD6.5 and demonstrated to have high levels of hydrolytic activity against the P-F bond (such as diisopropylfluorophosphate, DFP) while catalytic hydrolysis of the P-O or P-S bond was minimal.19 Apparent kinetic parameters for soluble OPAA with DFP as a substrate, obtained by a pH electrode, were Vmax ) 2.70 ( 0.28 mV/s and Km ) 0.54 ( 0.08 mM.14 This unique substrate preference makes OPAA very attractive for discriminative detection of fluorine-containing OP compounds including CW G-type nerve agents. DFP is a phosphonofluoridate neurotoxin similar to the G-type nerve agents sarin and soman with significantly reduced toxicity. The structural similarity with the CW agents makes DFP an attractive analogue for the development of detection and decontamination technologies for CW agents.20 Despite the studies on preparation and characterization of Langmuir and Langmuir-Blodgett (LB) films of OPAA,13 the molecular interaction between OPAA and DFP still remain unclear. In this paper, spectroscopic methods such as circular dichroism (CD) and fluorescence and infrared reflection absorption spectroscopies (IRRAS) were applied to study the hydrolysis reaction of DFP catalyzed by OPAA in aqueous solution and as a Langmuir film for its potential application in developing an enzyme-based G-type agent detection sensor. The optimum conditions to get a stable

10.1021/bm049199o CCC: $30.25 © 2005 American Chemical Society Published on Web 03/23/2005

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OPAA monolayer at the air-water interface and the stability of the monolayer were also investigated. 2. Experimental Section 2.1. Materials. Purified OPAA (E.C.3.1.8.2) was obtained from the U.S. Army Laboratory (Edgewood Chemical and Biological Center, APG, MD) with a purity of 85-90%. OPAA aqueous stock solution (3.2 mg/mL) was kept in the refrigerator at 4 °C. The OPAA concentration used for Langmuir film preparation and IRRAS measurements was 0.64 mg/mL, whereas for CD and fluorescence measurements the concentration was 0.08 mg/mL. The water used as a subphase was purified by a Modulab 2020 water purification system (Continental Water Systems Corp., San Antonio, TX). The pure water has a specific resistance of 18 MΩ‚cm and a surface tension of 72.6 mN m-1 at 20 ( 1 °C. The spreading solvent for the enzyme was the same buffer used as the subphase in Langmuir film preparation. Buffers with different compositions and pHs were prepared. Phosphate buffer (pH ) 7.2) was prepared with 0.1 M KH2PO4 and 0.1 M NaOH; carbonate buffer (pH ) 8.3) was prepared with 0.5% ammonium carbonate; BTP buffer (pH ) 6.8) was prepared with 10 mM bis-tris-propane, 0.05 mM MnCl2, and 0.1 mM dithiothreitol. KCl was added into the buffer solution as an electrolyte, and the concentration was 0.5 M. All the chemicals at the highest purity including DFP and NaF were purchased from Sigma Chemical Co. (St. Louis, MO). They were used as received. 2.2. Methods. The secondary structure of OPAA was determined using Jasco J-810 spectropolarimeter fitted with a 150-W xenon lamp. Quartz cells of 1-mm path length were used for all CD measurements of OPAA solution in the absence and presence of DFP. Several sets of samples were prepared by diluting 500 µL of OPAA (0.16 mg/mL) with DFP [1.1 × 10-2 M, in CH3OH-H2O (1:1, v/v)] and water to a total volume of 1 mL. The reference samples were prepared by diluting 500 µL of OPAA solution to a total volume of 1 mL with water. The CD spectra were recorded in the far-UV region (185-260 nm) with a response time of 8 s and scan speed of 50 nm/min. Three scans were accumulated and averaged for each spectrum after the background of the buffer solution was subtracted. The CDPro software package was used to estimate the secondary structure fractions of OPAA from the CD spectrum. More information about CDPro is available at the website http:// lamar.colostate.edu/∼sreeram/CDPro. The assignment from CDPro gave six secondary structural classes: regular R-helix, RR; distorted R-helix, RD; regular β-strand, βR; distorted β-strand, βD; turns, T; and unordered, U. The samples for the fluorescence measurements were prepared in the same way as for CD measurements and then placed in a 10-mm optical path length quartz fluorescence cuvette. The emission spectrum of each sample was measured using a Spex Fluorolog 1680 spectrofluorometer (SPEX Industries, Edison, NJ). All samples were excited at 285 nm. Experiments were designed to determine the optimum conditions for developing a stable monolayer of OPAA at the air-water interface and the stability of this monolayer

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was examined by the compression/decompression cycle. The surface chemistry experiments were done in a clean room class 1000, with a constant temperature of 20.0 ( 0.5 °C and a relative humidity of 50 ( 1%. A µ-S trough (Kibron, Inc., Fin00171, Helsinki, Finland) with an area of 115 cm2 (5.9 cm × 19.5 cm) was utilized for surface chemistry and IRRAS measurements. All the isotherms and spectra shown in the Results and Discussion section represent an average of at least three measurements. Molecular interaction of the OPAA with DFP was studied at the air-water interface by dissolving the DFP in the subphase at a concentration of 1.1 × 10-5 to 1.1 × 10-3 M and by spreading the enzyme as a monolayer. IRRAS measurements at the air-water interface with and without DFP were performed on the EQUINOX 55 spectrometer (Bruker, Boston, MA) equipped with a mercury cadmium telluride detector cooled by liquid nitrogen. The IR beam was conducted out of the spectrometer and focused onto the water surface of the Langmuir trough. The trough is enclosed in a poly(methyl methacrylate) cover with a nitrogen flux channel. The spectra were acquired with a resolution of 4 cm-1 by coaddition of 1200 (p polarization) scans after 20 min of nitrogen flux. A total of 40 µL of OPAA aqueous solution (0.64 mg/mL) was spread dropwise on the surface resulting in an initial surface pressure of 0 mN/m. After spreading the enzyme, 15 min were allowed for film equilibration. 3. Results and Discussion 3.1. OPAA Interaction with DFP in Aqueous Solution. 3.1.1. CD Spectroscopy. To get the information on the behavior of the OPAA interaction with DFP in aqueous solution, CD, a well-known spectroscoptic technique, was used to predict the secondary structure of the enzyme. CD is particularly good for studying the conformational stability of an enzyme under different environmental conditions. The secondary structure of enzymes can be determined by CD spectroscopy in the far-UV spectral region (185-260 nm)21-23 CD spectra of the OPAA aqueous solution (0.08 mg/mL), and the OPAA aqueous solutions in the presence of DFP at different concentrations are presented in Figure 1. The CD spectrum of teh OPAA aqueous solution showed two negative peaks near 210 and 221 nm, which were assigned to R-helix structure. The crossover point at 202 nm and the positive peak at 193 nm were also assigned to the R-helix structure. The presence of DFP led to a change in the intensity of the CD spectrum, reducing the contribution of the two negative R-helix peaks and the positive peak at 193 nm. The change in the shape of the peaks was not apparent when the DFP concentration was below 1.1 × 10-3 M while the secondary structure was almost totally lost when the concentration increased to 1.1 × 10-3 M. Table 1 shows the average fraction of the secondary structure of OPAA estimated from the CD data using the CDPro software package. The regular R-helix content of the OPAA aqueous solution was 22.8%, and the distorted R-helix content was 16.1%. In presence of DFP at a concentration of 1.1 × 10-3 M, these two helix structures were reduced to 6.0 and 8.4%,

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Molecular Interaction between OPAA and DFP

Figure 1. CD spectra of the OPAA aqueous solution in the absence and presence of DFP. The OPAA concentration is 0.08 mg/mL in all cases. (1) OPAA aqueous solution; (2-5) OPAA aqueous solution with 1.1 × 10-6, 1.1 × 10-5, 1.1 × 10-4, and 1.1 × 10-3 M DFP. The inset presents the CD spectra of the OPAA aqueous solution in the absence and presence of NaF. The concentration of NaF added is the same as that of DFP. Table 1. Average Estimation of Secondary Structure Fractions of the OPAA Aqueous Solution (0.08 mg/mL) in the Presence and Absence of DFP Using the CDPro Software Packagea fraction of secondary structure enzyme OPAA solution (0.08 mg/mL) OPAA with 10-6 M DFP OPAA with 10-5 M DFP OPAA with 10-4M DFP OPAA with 10-3 M DFP

RR % RD % βR % βD % T % U % 22.8 20.6 19.8 18.3 6.0

16.1 8.0 6.9 16.0 8.8 7.4 15.7 9.0 7.6 14.7 9.7 7.8 8.4 22.6 13.2

18.4 19.6 20.1 21.2 22.7

27.7 27.5 27.8 28.3 25.7

a R , regular R-helix; R , distorted R-helix; β , regular β-strand; β , R D R D distorted β-strand; T, turns; and U, unordered.

Scheme 1. Hydrolysis Reaction of DFP Catalyzed by OPAA

respectively. For this solution, the regular and disordered β-strands of OPAA increased from 8.0 and 6.9% to 22.6 and 13.2%, respectively. The percentage of β-turn and unordered structure did not change much with the concentration of DFP. The CD data indicate that there was a change in the secondary structure during the hydrolysis reaction represented in Scheme 1, and the β-strand structure may be more stable than R-helix as a result of the hydrolysis of DFP. The DFP-induced changes in the CD spectra may be affected by the presence of fluoride ion released after the hydrolysis reaction. This was tested using NaF instead of DFP. NaF was added into OPAA aqueous solution (0.08 mg/

mL) at the same concentration as DFP. The inlet of Figure 1 showed some minor decrease in the intensities of the two R-helix peaks upon addition of NaF, while the intensity of the strong positive peak around 193 nm was almost the same. Thus, we can conclude that the fluoride ion did not have a large influence on the secondary structure of OPAA. 3.1.2. Fluorescence Spectroscopy. The interaction between OPAA and DFP was also tested in aqueous solution by measuring the emission spectrum. Among the amino acids found in enzymes and proteins, phenylalanine, tyrosine, and tryptophan possess measurable fluorescence and can be used as intrinsic probes. Tryptophan possesses the most intense fluorescence in the native state.24 OPAA contains amino tryptophan groups which have been used to study the nature of the OPAA-DFP interaction by fluorescence spectroscopy. The tryptophan moiety has an emission at 350 nm when excited at 285 nm. As we can see in Figure 2, upon addition of DFP (1.1 × 10-6 M) to the OPAA aqueous solution, the emission intensity is quenched. Increasing the concentration of the DFP causes no further decrease of the intensity. The fluorescence measurements support the above CD results, indicating that there is a change in the secondary structure of the enzyme to favor the catalytic hydrolysis reaction of DFP. This change may cause quenching of the tryptophan emission intensity. The inlet of Figure 2 showed the emission spectra in the absence and presence of NaF. The fluorescence intensity did not change when the NaF concentration was 1.1 × 10-6 M. But when NaF concentration increased to 1.1 × 10-5 M, some fluorescence quenching was noticed. Increasing the concentration of the DFP causes no further decrease of the intensity either. The explanation of the

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Figure 2. Emission spectra of the OPAA aqueous solution (0.08 mg/mL) in phosphate buffer (1) and in the presence of DFP: (2) 1.1 × 10-6; (3) 1.1 × 10-5; and (4) 1.1 × 10-4 M. The inset presents the emission spectra of the OPAA aqueous solution in the absence and presence of NaF. The concentration of NaF added is the same as that of DFP.

Figure 3. Surface pressure-area isotherms of OPAA Langmuir films on (1) ammonium buffer, pH 8.3; (2) BTP buffer, pH 6.8; and (3) phosphate buffer, pH 7.2. All buffers contained 0.5 M KCl.

fluorescence quenching in the presence of NaF could be interpreted as a change of the polarity of the environment of the tryptophan moiety, causing a diminution of the intensity of the fluorescence. This experiment showed that the fluoride ion released after the DFP hydrolysis reaction could be another explanation of the fluorescence quenching. 3.2. OPAA as Langmuir Films Interacting with DFP in the Subphase. 3.2.1. Surface Pressure-Area Isotherm. Experiments were carried out to determine the optimal conditions of OPAA Langmuir film formation. Because of their solubility in water, enzymes have been spread on an aqueous subphase containing salt that helps to stabilize the

monolayers at the air-water interface by precluding dissolution.15,25,26 The solubility of the enzyme is also affected by the pH of the subphase, and this can be shown in the surface pressure-area isotherm. Figure 3 shows the surface pressure-area isotherms of the OPAA monolayer on different buffer subphases. When phosphate buffer was used, the OPAA monolayer was compressed at nil surface pressure to a larger surface area per molecule (3500 Å2 molecule-1) and a higher surface pressure (33 mN m-1) than using ammonium or BTP buffer. The results indicated that the most insoluble OPAA monolayer can be obtained on phosphate buffer at pH 7.2. This

Molecular Interaction between OPAA and DFP

Figure 4. Compression/decompression cycle of the OPAA monolayer at the pH ) 7.2 phosphate buffer subphase in the presence of 0.5 M KCl from nil to 5, 10, 15, and 20 mN/m surface pressure. The compression cycle starts from zero surface pressure, and the decompression cycle ends at zero surface pressure.

result is different compared to the one published recently.13 Because OPAA displays catalytic activity in a wide range of pHs (6.0-9.5) and temperatures (10-65 °C),27 we attribute this difference to the level of purity of the enzyme from one preparation to the other. For the purified sample that we used in this paper, the phosphate buffer at pH 7.2 is the most suitable one for the formation of a stable OPAA monolayer. The stability of the OPAA monolayer at the air-water interface was examined by the compression/decompression cycle under the optimal conditions of the subphase; the result is shown in Figure 4. No significant decrease in the apparent limiting molecular area was observed during the compression and decompression cycles at surface pressures of 5, 10, 15, and 20 mN m-1. This suggests the formation of a stable OPAA monolayer under those surface pressures at the airwater interface. When the surface pressure was set up at 20 mN m-1, it showed some hysteresis in the compression/ decompression cycle. This result is explained by the rearrangement of the OPAA molecules at the interface when the monolayer was decompressed. The surface pressure-area isotherm of OPAA monolayer in the presence of DFP (1.1 × 10-5 M) dissolved in the subphase has been investigated, and the results were compared to the isotherm of OPAA in absence of DFP. As indicated in Figure 5, the apparent limiting molecular area of the OPAA in the presence of DFP was 2800 compared to 3500 Å2/molecule in the absence of DFP. The collapsed surface pressure of the OPAA monolayer in absence and presence of DFP was 33 mN m-1 and 26 mN m-1, respectively. Again, there is some disparity from what was reported previously,13 where the mean molecular area increases significantly when DFP (10-4 M) is present (in Figure 6 of ref 13). Though this phenomenon was explained by the hydrolysis of DFP releasing fluoride ions, comparison of the two figures in ref 13 will lead to a conclusion that Figure 6 was mislabeled. Figure 2 in ref 13 showed the surface pressure-area isotherms of OPAA on a BTP buffer with a limiting molecular area of 3500 Å2/molecule, whereas

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Figure 5. Surface pressure-area isotherms of OPAA on the phosphate buffer subphase in the absence (1) and in the presence of DFP (1.1 × 10-5 M; 2).

Figure 6. IRRAS spectra of the OPAA monolayer collected at 15 mN m-1. (1) OPAA monolayer (subphase: phosphate buffer, pH ) 7.2, 0.5 M KCl); (2) OPAA monolayer on the phosphate buffer subphase in the presence of DFP at 1.1 × 10-5; (3) 1.1 × 10-4; and (4) 1.1 × 10-3 M.

Figure 6 in ref 13 showed 1300 Å2/molecule on BTP buffer. Replacing 1300 Å2 in Figure 6 with 3500 Å2 in Figure 2 will lead to a decrease in mean molecular area with the presence of DFP, which confirms our result here. We believe that the observations of the change on molecular area and the collapsed surface pressure of the enzyme in the presence of DFP are due to a change of conformation of the enzyme at the air-water interface, leading to the solubility of the enzyme into the subphase. The data of CD and fluorescence in solution show a secondary structure change after the hydrolysis reaction. Since we cannot do the same measurements at the air-water interface, what we observe by surface pressure-area isotherm confirms a structural change of the enzyme monolayer. 3.2.2. IRRAS. IRRAS spectra of the OPAA monolayer at the air-water interface in absence and in the presence of DFP are shown in Figure 6. On the basis of the surface chemistry results shown above, all the spectra were collected at 15 mN m-1 and the polarization angle was 41°. We chose 15 mN m-1 because the OPAA monolayer is stable during the whole compression procedure. Careful examination of

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the amide I (1700-1600 cm-1) and amide II (1600-1500 cm-1) regions in Figure 6 (1) revealed several overlapping resonances at 1542, 1560, 1652, 1673, and 1683 cm -1. On the basis of experimental obervations28,29 and theoretical calculations,30,31 the 1542, 1560, and 1652 cm-1 frequencies were assigned to the R-helix conformation, whereas the 1673 and 1683 cm-1 were assigned to β-sheet structure. There is no obvious change in the band positions and shape, although the intensities decreased when the DFP concentration was set at 1.1 × 10-5 and 1.1 × 10-4 M, indicating a change in the conformation of the OPAA. However, at a concentration of DFP of 1.1 × 10-3 M, the fingerprint of the spectrum was completely destroyed in the regions of amide I and amide II. The OPAA secondary structure was totally lost at high concentrations of DFP, especially in the amide I region. The 1652 cm-1 band which is associated with R-helix conformation disappeared, and the 1673 and 1683 cm-1 bands assigned to β-sheet structure became two very small shoulders. From Figure 5, it was noted that, at a concentration of 1.1 × 10-5 M, there was a change in the surface pressure-area isotherm, but the change in secondary structure by IRRAS was only noted at a higher concentration of DFP (1.1 × 10-3 M). Thus, the enzyme-substrate reaction resulted in a completely unfolded enzyme when the substrate concentration is high by causing disruption of the hydrogen bonding of the enzyme, and the reduction of the OPAA interfacial concentration might result from the change of the enzyme conformation. 4. Conclusion We have reported the molecular interaction between OPAA and DFP in aqueous solution and at the air-water interface as well as the optimum conditions for stable Langmuir film formation of OPAA. The characterization of OPAA aqueous solution and OPAA monolayer during DFP hydrolysis by spectroscopic methods revealed a change in OPAA secondary structure. The CD and IRRAS spectra indicate that a definite loss of secondary structure was observed at 1.1 × 10-3 M of DFP. The quenching of fluorescence intensity can be used as an effective biosensor with a much lower detection limit of 10-6 M of DFP because fluorescence spectroscopy is a more sensitive system of detection relative to CD and IRRAS spectroscopies. Acknowledgment. This work was supported by a grant from the U.S. Army Research Office (DAAD 19-03-1-0131). References and Notes (1) Thust, M.; Mulchandani, A.; Wang, J.; Arzdorf, M.; Mulchandani, P.; Chen. W.; Schoning, M. J. Tech. Mess. 2003, 12, 561-564. (2) Karalliedde, L.; Wheeler, H.; Maclehose, R.; Murray, V. Public Health 2000, 114, 238-248.

Zheng et al. (3) Mulchandani, A.; Mulchandani, P.; Chen, W. Field Anal. Chem. Technol. 1998, 2, 363-369. (4) Marrs, T. C.; Maynard, T. J.; Sidell, F. R. Chemical warfare agents: Toxicology and Treatment; John Wiley and Sons: Chichester, 1996. (5) Rogers, K. R.; Wang, Y.; Mulchandani, A.; Mulchandani, P.; Chen, W. Biotechnol. Prog. 1999, 15, 517-521. (6) Mendoza, C. E. Thin layer chromatography. In Pesticide Analysis; Dumas, K. G., Ed.; Marcel Dekker: New York, 1981; pp 144. (7) Hanks, A. R.; Colvin, B. M. High-performance liquid chromatography. In Pesticide Analysis; Dumas, K. G., Ed.; Marcel Dekker: New York, 1981; pp 99-174. (8) Barcelo, D.; Lawrence, J. F. Residue analysis of organophosphorus pesticides. In Emerging Strategies for Pesticide Analysis; Charins, T., Sherma, J., Eds.; CRC Press: Boca Raton, FL, 1992; pp 127150. (9) Clement, R. E.; Eiceman, G. A.; Koester, C. J. Environmental Analysis. Anal. Chem. 1995, 67, 221R-255R. (10) Dumschat, C.; Muller, H.; Stein, K.; Schwede, G. Anal. Chim. Acta 1991, 252, 7-9. (11) Constantine, C. A.; Mello, S. V.; Dupont, A.; Cao, X. H.; Santos, D.; Oliveira, O. N.; Strixino, F. T.; Pereira, E. C.; Cheng, T. C.; DeFrank, J. J.; Leblanc, R. M. J. Am. Chem. Soc. 2003, 125, 18051809. (12) Cheng, T.-C.; DeFrank, J. J.; Rastogi, V. P. Chem. Biol. Int. 1999, 119, 455-462. (13) Mello, S. V.; Mabrouki, M.; Cao, X. H.; Leblanc, R. M.; Cheng, T.-C.; DeFrank, J. J. Biomacromolecules 2003, 4, 968-973. (14) Simonian, A. L.; Grimsley, J. K.; Flounders, A. W.; Schoeniger, J. S.; Cheng, T.-C.; DeFrank, J. J.; Wild, J. R. Anal. Chim. Acta 2001, 442, 15-23. (15) Mello, S. V.; Coutures, C.; Cheng, T. C.; Rastogi, V. P.; DeFrank, J. J.; Leblanc, R. M. Talanta 2001, 55, 881-887. (16) Constantine, C. A.; Gattas-Asfura, K. M.; Mellow, S. V.; Crespo, G.; Rastogi, V.; Cheng, T. C.; DeFrank, J. J.; Leblanc, R. M. Langmuir 2003, 19, 9863-9867. (17) Constantine, C. A.; Gattas-Asfura, K. M.; Mello, S. V.; Crespo, G.; Rastogi, V.; Cheng, T. C.; DeFrank, J. J.; Leblanc, R. M. J. Phys. Chem. B 2003, 107, 13762-13764. (18) Cao, X.; Mello, S. V.; Sui, G.; Mabrouki, M.; Rastogi, V. K.; Cheng, T.-C.; DeFrank, J. J.; Leblanc, R. M. Langmuir 2002, 18, 76167622. (19) DeFrank, J. J.; Cheng, T.-C. J. Bacteriol. 1991, 173, 19381943. (20) Simonian, A. L.; diSioudi, B. D.; Wild, J. R. Anal. Chim. Acta 1999, 388, 189-196. (21) Sreerama, N.; Venyaminov, S. Y.; Woody, R. W. Anal. Biochem. 2000, 287, 243-251. (22) Sreerama, N.; Woody, R. W. Anal. Biochem. 2000, 287, 252-260. (23) Hirst, J. D.; Colella, K.; Gilbert, A. T. B. J. Phys. Chem. B 2003, 107, 11813-11819. (24) Lakowicz, J. R. Principles of Fluorescence Spectroscopy, 2nd ed.; Plenum Publishers: New York, 1999. (25) Gaines, G. L., Jr. Insoluble monolayers at liquid-gas interfaces; Wiley: New York, 1996. (26) MacRitchie, F. Chemistry at interfaces; Academic Press: New York, 1990. (27) Cheng, T.-C.; Calomiris, J. J. Enzyme Microb. Technol. 1996, 18, 597-601. (28) Pelletier, I.; Laurin, I.; Buffeteau, T.; Desbat, B.; Pezolet, M. Langmuir 2003, 19, 1189-1195. (29) Axelsen, P. H.; Citra, M. J. Prog. Biophys. Mol. Biol. 1996, 66, 227253. (30) Pauling, L.; Corey, R. B. Proc. Natl. Acad. Sci. U.S.A. 1951, 37, 29-36. (31) Krimm, S. J. Mol. Biol. 1962, 4, 528-532.

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