Rapid Profiling of Peptide Stability in Proteolytic Environments

Jan 21, 2009 - Hans H. Gorris, Steffen Bade, Niels Röckendorf, Eike Albers, M. Alexander Schmidt, Milan Fránek and Andreas Frey*. Division of Mucosa...
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Anal. Chem. 2009, 81, 1580–1586

Rapid Profiling of Peptide Stability in Proteolytic Environments Hans H. Gorris,†,‡,| Steffen Bade,† Niels Ro¨ckendorf,† Eike Albers,‡,⊥ M. Alexander Schmidt,‡ Milan Fra´nek,§ and Andreas Frey*,†

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Division of Mucosal Immunology, Research Center Borstel, Parkallee 22, 23845 Borstel, Germany, Institute of Infectiology, University of Mu¨nster, von-Esmarch-Strasse 56, 48149 Mu¨nster, Germany, and Department of Analytical Biotechnology, Veterinary Research Institute, Hudcova 70, 621 32 Brno, Czech Republic The notorious degradation susceptibility of peptides is a major obstacle to their use as medicinal drugs. Assays with which the stability of peptides in complex proteolytic environments can be determined are thus indispensable for peptide drug development. Herein, we describe a new peptide proteolysis assay that meets that demand. It unites the high-throughput capacity of heterogeneous with the well-defined kinetic characteristics of homogeneous assay formats and operates on the cleavage-caused loss of a detection handle. We have confirmed the assay’s accuracy with well-defined model interactions and proved its high versatility and robustness with a representative application where we determined the half-lives of 375 different peptides in a crude intestinal protease preparation. With this reliable, reproducible, and efficient assay the enzyme kinetics of any peptide-protease interaction is accessible even for complex protease solutions. Throughout the body, peptides act as key players in manifold regulatory and metabolic processes. Peptides owe this crucial role to a distinctive blend of properties. They combine small size and high diversity with high specificity and affinity. It is therefore not surprising that peptides draw increasing attention from drug manufacturers. Envisioned fields of application for peptide drugs include antibiotics, anticoagulants, hormone therapies, and cancer treatment. Peptides of at least 10 amino acids may be used as vaccines since they fulfill the minimal requirement for MHC-II binding,1 while peptides specifically binding to relevant diagnostic targets may be engineered into potent contrast agents for use in molecular imaging.2 Yet, although the therapeutic peptide market gains momentum, there are still hurdles to be overcome. One drawback is the limited stability of peptides which entails a quick * To whom correspondence should be addressed. Phone: + 49-4537-188 562. Fax: + 49-4537-188 693. E-mail: [email protected]. † Research Center Borstel. ‡ University of Mu ¨ nster. § Veterinary Research Institute. | Present address: Department of Chemistry, Tufts University, 62 Talbot Avenue, Medford, MA 02155. ⊥ Present address: Department of Clinical Chemistry and Pharmacology, University Hospital Bonn, Sigmund-Freud-Strasse 25, 53127 Bonn, Germany. (1) Siklodi, B.; Vogt, A. B.; Kropshofer, H.; Falcioni, F.; Molina, M.; Bolin, D. R.; Campbell, R.; Hammerling, G. J.; Nagy, Z. A. Hum. Immunol. 1998, 59, 463–471. (2) Zhou, M.; Ghosh, I. Biopolymers 2007, 88, 325–339.

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clearance from the body in general and, even more frustrating, a rapid digestion after oral uptake. This is not altogether surprising, since there are more than 550 putative proteases encoded by the human genome.3 Indeed, proteases are just as fundamentally important in every living organism as are peptides. One main characteristic of a protease is its ability to discriminate between many potential substrates, and the catalytic efficiency kcat/KM most accurately reflects the proteolytic power of a protease toward a given substrate.4 The action of proteases must therefore be taken into account when evaluating the quality of a peptidic drug or in vivo diagnostic. Hence, in order to render peptide drugs fit for the broad range of applications they may be suited for, screening techniques which allow the estimation of bioavailability and clearance behavior of peptide drugs and which can be used for identifying orally available digestion-resistant peptides are urgently required. In general, protease activities can be analyzed with homogeneous or heterogeneous assays. Homogeneous assays measure enzyme activities in solution and allow exact insights into enzyme kinetics. Fluorescence resonance energy transfer (FRET) is most widely employed for homogeneous assays. Yet, FRET requires spatial distances between donor and acceptor dyes of typically 20-60 Å,5 thus limiting the maximal peptide length that can be analyzed with this type of assay. Longer peptides can be analyzed by physical separation and detection of proteolysis fragments, e.g., by high-performance liquid chromatography (HPLC) or mass spectrometry, but these approaches are not suitable for multiplexed analyses. High-throughput analyses usually are hampered by peptide purification and detection, such that heterogeneous assays, which measure protease activities on surface-bound substrates, provide a better approach for high-throughput screening. By phage display, the prototype technique for peptide lead identification, very high numbers (107) of potential protease substrates can be tested concurrently.6 For a detailed characterization of selected substrates, however, other means are employed. Synthetic peptides can either be directly subjected to proteolysis on their synthesis support, e.g., on beads7 or on (3) Rawlings, N. D.; Morton, F. R.; Barrett, A. J. Nucleic Acids Res. 2006, 34, D270-D272. (4) Brot, F. E.; Bender, M. L. J. Am. Chem. Soc. 1969, 91, 7187–7191. (5) Selvin, P. R. Nat. Struct. Biol. 2000, 7, 730–734. (6) Matthews, D. J.; Wells, J. A. Science 1993, 260, 1113–1117. (7) Meldal, M.; Svendsen, I.; Breddam, K.; Auzanneau, F. I. Proc. Natl. Acad. Sci. U.S.A. 1994, 91, 3314–3318. 10.1021/ac802324f CCC: $40.75  2009 American Chemical Society Published on Web 01/21/2009

a cellulose membrane,8 or they may first be captured on a microtiter plate and then subjected to proteolysis.9 The extent of cleavage can be determined by assessing either the amount of uncleaved immobilized peptide or of immobilized product. Yet, the heterogeneous assay format cannot provide information about enzyme kinetics since diffusion processes and steric constraints impede the reaction of the enzyme with the surfacebound substrate. Here, we report a highly efficient new method that unites the advantages of both homogeneous and heterogeneous assay formats. It is suitable for automated high-throughput screening as combinatorial peptide libraries can be prepared by standard solid-phase peptide synthesis and a highly sensitive and robust ELISA-type procedure is used for the detection of proteolytic cleavage. Simultaneously, precise insights into enzyme kinetics are possible as the crucial proteolytic step takes place in solution. Thus, the method described in this paper enables the large-scale determination of enzyme kinetics of any peptide-protease interaction even in crude protease solutions. EXPERIMENTAL SECTION Buffers and Reagents. The following were used: L(ite)-PBS, 10 mM sodium phosphate, pH 7.0, 10 mM NaCl; D(ulbecco’s)PBS, 2.7 mM KCl, 1.5 mM KH2PO4, 136 mM NaCl, 8.1 mM Na2HPO4, pH 7.3; simulated intestinal fluid (SIF),10 8 mM phosphate, pH 7.2, 4.6 mM K+, 111.3 mM Na+, 101.5 mM Cl-; protease inhibitor cocktail, 308 nM aprotinin, 20 µM leupeptin, 400 µM 4-(2-aminoethyl)benzenesulfonyl fluoride, 2 mM EDTA in L-PBS. 11-(2-(2,4-Dichlorophenoxy)acetylamino)undecanoic acid was synthesized in our laboratory (unpublished results). The monoclonal anti-2,4-D antibody was generated against the 2,4-D hapten coupled to thyroglobulin according to standard procedures.11 Clone E2/G2 was used throughout all experiments. Intestinal fluids were harvested from excised murine intestines (unpublished results). For detection of horseradishperoxidase-labeled streptavidin (SA-HRP, Vector, Burlingame, CA) we employed a 3,3′,5,5′-tetramethylbenzidine (TMB) substrate system.12 Peptide Synthesis. Peptide libraries were SPOT-synthesized on a cellulose membrane support using standard fluorenylmethoxycarbonyl (Fmoc) amino acid protection chemistry as described by Frank.13 Briefly, peptide SPOTs were defined by automated application of a Boc-Lys (Fmoc) solution on a prolinederivatized cellulose membrane, establishing a cleavable ε-lysineproline anchor.14 Next, the first label, a biotin derivative with a poly(ethylene glycol) (PEG) spacer, was applied. The peptide was then assembled in a semiautomated cycle using an ASP 222 peptide synthesizer (Intavis AG, Cologne, Germany): (i) acetyl(8) Dekker, N.; Cox, R. C.; Kramer, R. A.; Egmond, M. R. Biochemistry 2001, 40, 1694–1701. (9) Gutierrez, O. A.; Salas, E.; Hernandez, Y.; Lissi, E. A.; Castrillo, G.; Reyes, O.; Garay, H.; Aguilar, A.; Garcia, B.; Otero, A.; Chavez, M. A.; Duarte, C. A. Anal. Biochem. 2002, 307, 18–24. (10) Lockwood, J. S.; Randall, H. T. Bull. N. Y. Acad. Med. 1949, 25, 228–243. (11) Franek, M.; Kolar, V.; Granatova, M.; Nevorankova, Z. J. Agric. Food Chem. 1994, 42, 1369–1374. (12) Frey, A.; Meckelein, B.; Externest, D.; Schmidt, M. A. J. Immunol. Methods 2000, 233, 47–56. (13) Frank, R. Tetrahedron 1992, 48, 9217–9232. (14) Bray, M. B.; Maeji, N. J.; Geysen, H. M. Tetrahedron Lett. 1990, 31, 5811– 5814.

blocking, (ii) Fmoc-deprotection with 20% piperidine, (iii) bromophenol blue staining for synthesis control, (iv) drying, (v) automatic application (3×) of 0.2 µL of diisopropylcarbodiimideactivated amino acid solutions (0.2 M Fmoc-protected amino acid and 0.35 M 1-hydroxybenzotriazole in methyl-2-pyrrolidone). As the last synthesis step, the second label 2,4-dichlorophenoxyacetic acid was attached. For side-chain deprotection and diketopiperazine formation of the ε-lysine-proline anchor the membrane was incubated in 50% trifluoroacetic acid, washed, and dried. The SPOTs were punched out and transferred individually to polypropylene tubes. Peptides were cleaved from the membrane in 0.1 M triethylammonium acetate, 20% ethanol at 30 °C. Lyophilized peptides were dissolved in 1.5 mL of L-PBS × 0.005% (w/v) Tween 20, snap-frozen in liquid N2, and stored at -80 °C. For a detailed description of the SPOT synthesis see the Supporting Methods in the Supporting Information. Capture ELISA. High-bind standard 96-well microtiter plates (Corning, Wiesbaden, Germany) coated overnight at 4 °C with 75 µL/well of 50 ng/mL anti-2,4-D antibody E2/G2 in L-PBS were washed three times with 300 µL/well of D-PBS × 0.05% (w/v) Tween 20 (D-PBST), blocked with 250 µL/well of 1% (w/v) casein/ D-PBS for 3-4 h at room temperature (RT), and again washed four times with D-PBST. Peptides were serially diluted in 75 µL/ well of SIF × 0.005% (w/v) Tween 20 (SIFT). Alternatively, to determine the peptide concentration or the influence of free label compounds, 2,4-D-aminoundecanoic acid, 2,4-D, or biotin was serially diluted in 75 µL/well of 67 pM peptide in SIFT on a 96well polypropylene microtiter plate and transferred to the antibodycoated microtiter plate. After 2 h and 30 min plates were washed four times with D-PBST and incubated for 60 min at RT with 75 µL/well of 1 µg/mL SA-HRP in 1% (w/v) casein in D-PBS. After six washes with D-PBST, plates were developed using the TMB substrate system. Proteolysis Assay. Proteolysis reactions were carried out in a 96-well polypropylene microtiter plate. A defined enzyme solution or a crude enzyme preparation was serially 1:3 diluted in 50 µL/ well of 67 pM peptide in SIFT × 1 mM CaCl2. For inhibition studies, aprotinin was added to the peptide solution. The sealed microtiter plate was incubated for 90 min at 37 °C before the enzyme reaction was stopped by addition of 50 µL/well of protease inhibitor cocktail. The microtiter plate was resealed, stored for 10 min on ice, heated for 10 min to 90 °C, and cooled on ice. From each cavity, 75 µL of the peptide solution was transferred into a corresponding cavity of a high-bind 96-well microtiter plate which beforehand had been coated with 75 µL/ well of 30 ng/mL anti-2,4-D antibody E2/G2, washed and blocked as described above. After 2 h and 30 min of incubation at RT the plate was washed four times with D-PBST and incubated for 60 min at RT with 75 µL/well of 1 µg/mL SAHRP in 1% (w/v) casein in D-PBS. After six washes with D-PBST, plates were developed with the TMB substrate system. RESULTS AND DISCUSSION Experimental Design. Peptides were synthesized in parallel on a cellulose membrane by standard Fmoc-protection chemistry.13 This so-called SPOT synthesis provides a combinatorial peptide library of up to 1700 different peptides for high-throughput screening in a nanomole scale. A drawback of the SPOT synthesis Analytical Chemistry, Vol. 81, No. 4, February 15, 2009

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Figure 1. Schematic representation of the proteolysis assay. (A) Peptide synthesis on a cellulose membrane support (1): cleavable anchor (2), biotin label (3), substrate sequence motif consisting of n amino acids (4), hapten label (5). (B) Release of peptides from the membrane and incubation with a protease (6) in solution. (C) Peptide detection in a microtiter plate (7): An antibody (8) captures the hapten label of the peptide irrespective of peptide cleavage. Only uncleaved peptides which are stable to proteolysis carry the second label that is detected by an appropriate receptor (9). The receptor is linked to a reporter enzyme (10) which catalyzes color development (11). The ratio of cleaved to uncleaved peptide defines the degree of proteolytic degradation.

is a substantial amount of synthesis impurities, mainly truncated peptides resulting from unwanted new synthesis starts during peptide assembly cycles or premature termination of the peptide chain. This drawback, however, is circumvented by a double purification strategy outlined in Figure 1: Only peptides which feature the C-terminal anchor are released into solution by specific anchor cleavage. After proteolysis, only peptides and cleavage products carrying the N-terminal label are immobilized on a microtiter plate for detection with an ELISA-type assay, whereas all peptides synthesized incompletely are not immobilized. This headfirst peptide recapture renders additional purification steps dispensable and is especially useful if crude protease preparations are examined as all components that may interfere with peptide detectionsa notorious issue especially when dealing with fluorogenic substratessare removed by washing. Only the proteolytic reaction itself takes place in solution, which is necessary for the determination of enzyme kinetics. Assay Sensitivity and Robustness. Peptide recapturing required N- and C-terminal peptide labels with high affinity and specificity for their respective receptors to minimize interferences with other components. Furthermore, the labels had to be Fmoc chemistry compatible and ought to be small to minimize steric hindrance. As one label, we chose biotin to bind a streptavidincoupled detection system as this pair has the highest known binding constant15 of biologicals and is already well established in ELISA-type assays. The peptide’s water solubility and accessibility was improved by a PEG spacer between biotin and the sequence motif. An appropriate binding pair for the second label, however, was not easily at hand. We therefore devised a novel label system which is based on the hapten 2,4-dichlorophenoxyacetic acid (2,4-D) and a respective anti-2,4-D antibody.11 Here we demonstrate that this new label system constitutes a sensitive binding pair which is excellently suited for peptide capturing. To optimize peptide detection we made use of the so-called bridge (15) Green, N. M. Methods Enzymol. 1990, 184, 51–67.

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Figure 2. (CH2)n-spacers linked to 2,4-D enhance peptide detection. Pseudopeptides consisting of alanine and a short poly(ethylene glycol) stretch were N-terminally labeled with 2,4-D linked to various aliphatic aminocarboxylic acids and C-terminally with biotin. The peptides were serially diluted in a microtiter plate coated with 50 ng/mL anti-2,4-Dantibody and detected via their biotin label. Error bars indicate the SE of triplicate measurements performed with peptides from three different syntheses.

binding effect. Earlier observations with 2,4-D16 and other haptens17 had shown that linkage of a (CH2)n-spacer to the hapten results in an increased antibody affinity. Since this effect is deleterious for the hitherto predominant usage of anti-2,4-D antibodiessthe detection of the herbicide 2,4-D in the environment with competitive ELISAssthe bridge binding effect has never been investigated in detail before. In our optimization procedure we now tested a variety of spacers and found that aminoundecanoic acid resulted in the highest increase in affinity (Figure 2). The enormous improvement in sensitivity in comparison with the unmodified peptide is reflected in a 13-fold increase of ODmaxsthe maximum optical density obtained with antibody-saturating peptide amountssand a 34-fold increase of the test midpointsthe peptide concentration where half of the binding sites of the capture antibody are occupied. In this way, the detection limit for peptides was pushed beyond 1 fmol. The average SPOT synthesis yield of peptides with random sequence motifs carrying biotin as well as 2,4-D-aminoundecanoic acid label was determined with a competitive ELISA. From the test midpoint, which represents equal amounts of competitor and peptide, the total peptide yield could be determined to be about 1 nmol/peptide spot (for details, see Supporting Information Table S1). To minimize the influence of variations in peptide synthesis yields and to enhance the sensitivity of our assay we chose to employ 3/1000 of a peptide spotsabout 3 pmol of peptidesfor each proteolysis experiment which resulted in an antibody saturation level of more than 99.5%, thus approximating ODmax. Free 2,4-D or biotin in crude protease preparations might compete with the detection of their respective peptide labels and thus impair peptide detection. However, we found that concentrations of at least 1 µM 2,4-D or 50 µM biotin are required for assay interference. These amounts by far exceed the concentrations to be expected in biological samplessthe maximum admissible concentration of 2,4-D in tap water being 0.45 nM in the European Union, the highest concentration of biotin in biological samples amounting to around 5 µM (liver). Thus, the assay is highly robust toward biotin and 2,4-D contaminations. (16) Hatzidakis, G. I.; Tsatsakis, A. M.; Krambovitis, E. K.; Spyros, A.; Eremin, S. A. Anal. Chem. 2002, 74, 2513–2521. (17) Franek, M. J. Steroid Biochem. 1987, 28, 95–108.

In addition to adverse effects caused by contaminating tags the peptide sequence motif itself may influence peptide detection. To improve solubility we provided each peptide with an additional PEG moiety and added the detergents Tween 20, Triton X-100, or cholic acid in low concentrations to the peptide solutions. Neither detergents nor various blocking reagents, however, could prevent that some peptides bound in an antibody-independent way to the microtiter plate (Figure 3B). This nonspecific binding, which seemingly is sequence-dependent, must be prevented in order to distinguish between cleaved and uncleaved peptides after proteolysis. We investigated systematically the nonspecific binding properties of all 732 overlapping 10-mer, 13-mer, and 16-mer sequence motifs (frame shift of two amino acids) from the model protein ovalbumin intended for proteolysis experiments. Correlation analyses of nonspecific binding versus amino acid classes in the sequence motif surprisingly revealed that nonspecific binding was not mediated by hydrophobicity (Figure 3C) as might be expected since microtiter plates consist of polystyrene. Although polar amino acids reduced nonspecific binding (Figure 3D), a significant positive correlation was observed between nonspecific binding and increasing net positive charge of the sequence motif (Figure 3E). Attachment of peptides onto polystyrene surfaces via positively charged amino acids has been described previously.18 This binding can be attributed to a noncovalent and mostly electrostatic interaction between a cation and a π-system.19 We could confirm the electrostatic character of this cation-π interaction on polystyrene surfaces as binding was abolished if the peptides contained an excess of negatively charged amino acids. On the basis of this finding, we deduced a strategy to prevent nonspecific binding. To each peptide we annexed flanking regions consisting of negatively charged amino acidssglutamate and the double negatively charged carboxyglutamatesand a short hydrophilic PEG. Using these constructs, we could show that the nonspecific binding is for the most part charge-dependent (Figure 3F). It was completely or almost completely prevented with two carboxyglutamate or glutamate moieties, on each side of the sequence motif, whereas PEG alone was hardly capable of reducing nonspecific binding. Thus, we had identified a means for reducing nonspecific binding of the peptides to microtiter plates, but it remained to be tested whether substrate degradation is influenced by negatively charged flanks. Assay Validation. Once we had established a sensitive and robust peptide detection system it was readily applicable for proteolysis experiments. For proteolysis, conditions for a pseudofirst-order reaction were chosen. In a first-order reaction the halflife is not dependent on the amount of reactants; thus, it is not necessary to determine the exact amounts of substrate peptide obtained in different syntheses. Proteolysis was performed on a nonbinding 96-well microtiter plate, such that after termination of the proteolysis reaction the samples were directly transferable to the antibody-coated microtiter plate. Although cleaved and uncleaved peptides compete with their 2,4-D label for the binding sites of the coating antibody, which captures a representative fraction of cleaved and uncleaved peptide, only uncleaved peptides generate a signal via their biotin label. By means of the coating (18) Loomans, E. E.; Petersen-van Ettekoven, A.; Bloemers, H. P.; Schielen, W. J. Anal. Biochem. 1997, 248, 117–129. (19) Ma, J. C.; Dougherty, D. A. Chem. Rev. 1997, 97, 1303–1324.

Figure 3. Nonspecific binding of peptides to polystyrene microtiter plates. The 16-mer peptides were C-terminally labeled with biotin and N-terminally with 2,4-D (b) or phenoxyacetic acid (O). The latter 2,4-D surrogate is not captured by anti-2,4-D antibodies but exhibits a similar chemical structure. Peptides were serially diluted in an anti-2,4-D antibody-coated microtiter plate and detected via their biotin label. (A-B) Binding behavior of two different peptides. While one sequence motif (A) did not bind nonspecifically to the microtiter plate, nonspecific binding was observed with the second sequence motif (B). (C-E) Test of 732 peptides for their propensity to bind nonspecifically to a microtiter plate. The polystyrene plate was coated with a polyclonal IgG preparation not directed against 2,4-D, and 2 pmol of peptide was employed. No significant correlation (F-test, p > 0.05) was observed between nonspecific binding and the presence of hydrophobic amino acids (C) in the sequence motif. Hydrophilic/uncharged amino acids (D) reduced the nonspecific binding significantly (negative correlation: F-test, p < 0.0001). Nonspecific binding increased with net positive charge (E) (significant positive correlation: F-test, p < 0.0001). (F) Reduction of nonspecific binding by various flanking motifs. A peptide sequence motif exhibiting high nonspecific binding (xxx) was provided with different combinations of a hydrophilic residue (poly(ethylene glycol) diglycolic acid (n ) 2), P), a single negative charge (D-glutamate, -) and a double negative charge (carboxyglutamate, )), and nonspecific binding to microtiter plates was analyzed as above. The lowest nonspecific binding was achieved with negatively charged building blocks on both sides of the sequence motif. Error bars represent the geometric mean ( SE of triplicate measurements.

concentration an optimal signal span can be adjusted. The span boundaries are defined by the signal of an entirely uncleaved peptide yielding ODmax and the background ODbg obtained after complete proteolysis. For data analysis we fitted the raw data of the pseudo-first-order proteolysis reaction to an exponential function 1: Analytical Chemistry, Vol. 81, No. 4, February 15, 2009

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ODt ) (ODmax - ODbg)e-tk + ODbg

(1)

where t is the reaction time and k is the rate constant. As substrate catalysis obeys Michaelis-Menten kinetics k can be substituted by [E]tkcat/KM, and if the total enzyme concentration [E]t is taken as constant, the catalytic efficiency (kcat/KM) of substrate proteolysis can be determined as a curve-fit parameter. However, if the proteolysis assay is applied for the high-throughput determination of peptide stability, one must presume a wide range of kcat/KM. Therefore, it is unfavorable to determine substrate decay as a function of incubation time. In order to circumvent this problem, we exchanged the independent variable t with [E]t to obtain eq 2: OD[E] ) (ODmax - ODbg)e-[E]t t(kcat/KM) + ODbg

(2)

By keeping the incubation time constant and varying the enzyme concentration, a wide range of kcat/KM values can be covered easily. Parts A and B of Figure 4 show that varying time or enzyme concentration yields the same kcat/KM. In case the stability measurements should be expanded into inhibition studies, only a second enzyme dilution with a defined concentration of competitive inhibitor is required. Under pseudofirst-order conditions the inhibition constant Ki can be directly determined according to eq 3: v0 [I]t )1+ vi Ki

(3)

where v0 and vi denote the steady-state rates, kcat/KM, in the absence and presence of the inhibitor and [I]t is the total inhibitor concentration. In order to validate this approach experimentally, we determined the inhibition constant Ki of the competitive inhibitor aprotinin for the degradation of GPARLA by trypsin to be (1.5 ± 0.8) × 10-9 M and for the degradation of GVPFGP by chymotrypsin to be (4.6 ± 0.7) × 10-7 M (geometric mean ± SE of triplicate measurements). For high-throughput analyses we prevented nonspecific peptide binding to the microtiter plate by flanking the sequence motif with negatively charged amino acids. It turned out, however, that kcat/ KM is strongly influenced by different flanking moieties (Figure 4, parts C and D). Negatively charged flanks rendered the substrate up to 3 orders of magnitude more stable to proteolysis, regardless whether trypsin or chymotrypsin was employed. The effect that negatively charged amino acids near the cleavage site reduce kcat/KM has been described before and may be explained by the influence of negatively charged amino acids on the catalytic machinery of serine proteases.20 We were able to avoid this drawback by inserting a short PEG moiety as an “insulator” between the sequence motif and the negatively charged amino acids. As an outcome of our stringent optimization process we finally chose the construct - -PxxxP- - depicted in Figure 4E as the optimal design for high-throughput screening. To juxtapose our new proteolysis assay to different established methods we synthesized various peptide substrates with and without the optimal flanking motif and subjected them to our (20) Coombs, G. S.; Dang, A. T.; Madison, E. L.; Corey, D. R. J. Biol. Chem. 1996, 271, 4461–4467.

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Figure 4. Determination of kcat/KM. Either 67 pM substrate (GPARLA) was incubated with 2 nM enzyme (trypsin) for 10, 30, 90, or 270 min at 37 °C (A) or the enzyme was serially diluted in 67 pM substrate and incubated for 90 min at 37 °C (B). After stopping the enzyme reaction, the peptide solutions were transferred to an antibody-coated microtiter plate, where uncleaved peptides were detected via their biotin label by enzyme-coupled signal amplification. Plotting substrate degradation against incubation time (A) or enzyme concentration (B) yielded both pseudo-first-order reaction kinetics, and eq 2 was employed for nonlinear curve fitting. Error bars indicate the SE of fivefold measurements. kcat/KM was 1.9 × 106 ( 0.2 × 106 M-1 s-1 (A) or 2.0 × 106 ( 0.4 × 106 M-1 s-1 (B) (geometric mean ( SE). Both values did not differ significantly from each other (p > 0.05, Student’s two-tailed t test). Comparable results were obtained with the enzyme chymotrypsin (see Supporting Information Figure S2). (C and D) Influence of flanking motifs on kcat/KM. The 16-mer substrate sequence motifs for trypsin (AEAGVDAASVSEEFRA) (C) and chymotrypsin (MLVLLPDEVSGLEQLE) (D) were combined with the best flanking motifs for preventing nonspecific binding (see Figure 3F), and kcat/KM was determined for all combinations. Error bars indicate the SE of triplicate measurements. Negatively charged building blocks rendered the peptides more stable to proteolysis than peptides without such flanking regions, but this effect was prevented by inserting PEG between sequence motif and negatively charged building blocks (for more examples see Supporting Information Figure S3). (E) Final peptide construct, - -PxxxP- -, optimized for minimal nonspecific binding and no interference of flanking motifs with the proteolytic reaction: biotin (1), PEG (2), 2× D-glutamate (3), sequence motif (4), aminoundecanoic acid (5), and 2,4-D (6).

assay. Table 1 shows the catalytic efficiency obtained with the new assay in comparison to literature data. Good agreement is observed with the kinetic data from a FRET assay, which is most widely employed and commonly seen as the most accurate method for kinetic studies. Analysis of Peptide Half-Lives in a Crude Intestinal Protease Preparation. We used crude murine small intestinal lavage, which represents the entire complex digestive enzyme mixture present in the gut, to determine the stability of peptides in the small intestine. Since this preparation may contain, e.g., biotinidases capable to hydrolyze the peptide label, we first tested the peptide’s framework integrity under assay conditions by applying the enzyme preparation to “peptides” consisting of a single D-alanine sequence motif that does not constitute a substrate for intestinal enzymes. The peptide framework was completely stable to hydrolysis and is therefore amenable for proteolysis experiments.

Table 1. Comparison of the Proteolysis Assay with Literature Dataa kcat/KM [M-1 s-1] protease trypsin trypsin chymotrypsin

substrate

no flank

GGSGPFGRSALVPEE GPARLAIG PAPFAAA

(6.6 ± 1.5) × 10 (1.3 ± 1.2) × 106 (1.4 ± 0.6) × 105 6

- -PxxxP- -

literature data

(2.2 ± 1.7) × 10 (6.7 ± 3.5) × 105 (1.4 ± 0.6) × 105

HPLC (ref 20): 2.0 × 106 FRET (ref 29): 1.2 × 106 chromatography (ref 22): 4.5 × 104

6

a Proteases were serially diluted in 67 pM peptide (no flank and - -PxxxP- -) and incubated for 90 min at 37 °C. After stopping the enzyme reaction, the peptide solutions were transferred to an antibody-coated microtiter plate, where uncleaved peptides were detected via their biotin label. The data are the geometric mean ± SE of triplicate measurements.

the half-lives of peptides in the intestinal protease preparation by transforming eq 2 to

( ) + OD -

ODx ) (ODmax - ODbg) 0.5

Figure 5. Profiles of peptide stability in the small intestine: halflives of overlapping 10-mer (left side) and 16-mer (right side) ovalbumin-derived peptides in an undiluted small intestinal protease preparation (for a complete list of half-lives see Supporting information Table S2). The sequence motif of every fourth peptide is shown. Peptide stabilities were measured using the optimized peptide construct described in Figure 4E and murine intestinal lavage of known dilution. Calculated half-lives were extrapolated to undiluted enzyme solutions. Data represent the geometric mean ( SE of triplicate measurements.

Peptide sequences may contain several cleavage sites, and the enzyme activity in the intestinal preparation is composed of various interacting enzymes. It is therefore not feasible to determine kcat/ KM for single cleavage-site-enzyme pairs. Thus, we calculated

tx t1⁄2

bg

(4)

where x is the dilution factor of the enzyme solution and t1/2 is the half-life. Although the sampling of the intestinal lavage required an enzyme dilution, our protocol allows for the determination of the dilution factor (unpublished results). The lavage contained about 1 µM each of trypsin, chymotrypsin, and elastase and, for poor substrates, had to be applied in high concentrations. With the substrate concentration in our proteolysis assay being 67 nM peptide, the steady-state assumption in Michaelis-Menten kinetics that requires an excess of substrate [S] over enzyme [E] seems to be violated. This condition, however, is unnecessarily stringent.21 The steadystate assumption also holds if [E] , [S] + KM. As KM values for good trypsin and chymotrypsin substrates are about 50 µM,20,22 and correspondingly higher for poor substrates, the steady-state assumption is assured for the whole range of peptide half-lives. To confirm that the enzyme activity of the intestinal protease preparation remained stable during the entire 90 min proteolysis time, we first diluted the protease preparation in the microtiter plate, preincubated it for 0, 30, 60, 90, or 120 min at 37 °C, and added peptide substrates with cleavage motives for trypsin or chymotrypsin after the preincubation time (see Supporting Information Figure S1). No significant differences in half-lives due to incubation times of up to 120 min were detectable (triplicate measurements, one-way ANOVA, Fisher’s PLSD test, p > 0.05). The stability of peptides in the small intestine is a fundamental requirement if these peptides are to be used as oral drug or vaccine candidates. Oral vaccines are much easier and safer to administer than parenteral vaccines and, in addition to systemic immunity, provide protection against mucosal pathogens,23 which induce the vast majority of infectious diseases.24 In oral vaccinations antigens are most efficient if transported across the epithelium by M cells of Peyer’s patches, hence via the same route as authentic pathogens,25 before a classical immune response is launched with antigen presentation of MHC-II-bound peptides. (21) Segel, L. A. Bull. Math. Biol. 1988, 50, 579–593. (22) Bauer, C. A. Biochim. Biophys. Acta 1976, 438, 495–502. (23) Lue, C.; van den Wall Bake, A. W.; Prince, S. J.; Julian, B. A.; Tseng, M. L.; Radl, J.; Elson, C. O.; Mestecky, J. Clin. Exp. Immunol. 1994, 96, 356– 363. (24) Sirard, J. C.; Niedergang, F.; Kraehenbuhl, J. P. Immunol. Rev. 1999, 171, 5–26. (25) Frey, A.; Giannasca, K. T.; Weltzin, R.; Giannasca, P. J.; Reggio, H.; Lencer, W. I.; Neutra, M. R. J. Exp. Med. 1996, 184, 1045–1059.

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Since peptides with a length of 10 amino acids constitute the minimal requirement and those with 15-16 amino acids the maximum allowance for MHC-II binding,1 we chose to determine in vivo half-lives of model antigen peptides of 10 as well as 16 amino acids length. We synthesized the sequence of the model antigen ovalbumin26 in fragments of overlapping 10-mer and 16mer peptides with a frame shift of two amino acids and created a portrayal of the ex vivo stability of all of these potential antigenic determinants (Figure 5). The stabilities of neighboring peptides with a large overlap are similar, but in general, stabilities differed in a range of 5 orders of magnitude with a maximal half-life of 40 s. A similar digestion profile is obtained with 10-mer (Figure 5, left) and 16-mer peptides (Figure 5, right), albeit 16-mer peptides are in general degraded more rapidly (median, 15 ms) than 10mer peptides (median, 52 ms), probably due to the longer sequence motifs bearing more potential cleavage sites. With this study we could for the first time substantiate the generally low stability of short unfolded peptides in the small intestine. This finding stresses the importance to stabilize oral peptide drugs and vaccines composed of natural amino acids.27 In addition to accurately determining peptide half-lives encompassing a time range from milliseconds to minutes, we could also demonstrate that peptides spanning a sequence motif of 16 amino acids between the detection labels are valid substrates for the new proteolysis assay, whereas this length is not feasible in FRETbased tests. Analysis of the Z′-Factor. To compare the novel assay with current methods for high-throughput screening, we used the Z-factor statistical analysis method that is an indicator of the signal dynamic range of an assay system.28 The Z′-factor ranges from 1 (ideal assay) to 0, and an assay displaying a value of g0.5 is considered excellent. The Z′-factor of an assay is calculated according to eq 5 using the mean and standard deviation for signals from both the positive and negative control: (26) Nisbet, A. D.; Saundry, R. H.; Moir, A. J. G.; Fothergill, L. A.; Fothergill, J. E. Eur. J. Biochem. 1981, 115, 335–345. (27) Brayden, D. J.; O’Mahony, D. J. Pharm. Sci. Technol. Today 1998, 1, 291– 299. (28) Zhang, J. H.; Chung, T. D.; Oldenburg, K. R. J. Biomol. Screening 1999, 4, 67–73. (29) Grahn, S.; Ullmann, D.; Jakubke, H. Anal. Biochem. 1998, 265, 225–231.

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Z'-factor ) 1 -

3σc+ + 3σc|µc+ - µc-|

(5)

where σc+ and σc- denote the standard deviations and µc+ and µc- are the means of the positive or negative control, respectively. For our assay, we designated the signals of not degraded peptides (ODmax) as the negative and the background signals (ODbg), representing total proteolysis, as the positive controls. The signals were normalized to ODmax′ ) ODmax/(ODmax + ODbg) and ODbg′ ) ODbg/(ODmax + ODbg). Using eq 5 we determined a Z′-factor of 0.77 for our assay, which confirms that the novel assay is reproducible, robust, and reliable for highthroughput screening purposes. CONCLUSION We have developed a new analytical assay for evaluating the degradation resistance of peptide drug candidates. By measuring the stability of several hundred peptides in a crude intestinal fluid we have demonstrated that our new peptide proteolysis assay combines ease of handling with a high robustness and the exact determination of kinetic features. It therefore is a suitable tool for creating peptide proteolysis profiles which can aid in the design of digestion-resistant pharmaceuticals for oral administration. ACKNOWLEDGMENT This work was funded in part by Grants from the German Ministry for Education and Research (BMBF, Grants 01KO0113 and 13N8473) and from the German Research Council (DFG, Grant Fr 958/4-1). NOTE ADDED AFTER ASAP PUBLICATION This article was released ASAP on January 21, 2009 with minor errors in equation 4. The correct version was posted on January 27, 2009. SUPPORTING INFORMATION AVAILABLE Additional information as noted in text. This material is available free of charge via the Internet at http://pubs.acs.org. Received for review December 26, 2008. AC802324F

November

4,

2008.

Accepted