Printing Small Molecules as Microarrays and Detecting Protein−Ligand Interactions en Masse

The ability to identify small molecule ligands for any protein of interest has far-reaching implications, both for the elucidation of protein function and for the development of novel pharmaceuticals. With the introduction of the split-pool strategy for synthesis1 and the development of appropriate tagging technologies,2 chemists are now able to prepare large collections of natural product-like compounds immobilized on polymeric synthesis beads.3 These libraries provide a rich source of molecules for the discovery of new protein ligands. With such libraries in hand, the availability of efficient methods for screening these compounds becomes imperative. One method that has been used is the on-bead binding assay.4 An appropriately tagged protein of interest is mixed with the library and beads displaying cognate ligands are subsequently identified by a chromagenic or fluorescence-linked assay. Despite the utility of this approach, it is limited by the small number of proteins that can be screened efficiently. In principle, the beads can be stripped of one protein and re-probed with another; however, this serial process is slow and limited to only a few iterations. To identify a specific small molecule ligand for every protein in a cell, tissue, or organism, high-throughput assays that enable each compound to be screened against many different proteins in a parallel fashion are required. To address this issue, we have developed a technique that we refer to as small molecule printing (SMP). First, synthesis beads are distributed into polypropylene microtiter plates at a density of one bead per well. The attached compounds are then released from their beads and dissolved in a small volume of a suitable solvent. Due to the minute quantities of compound present on each bead, extreme miniaturization of the subsequent assay is an absolute requirement. Taking our cue from cDNA microarray technology, we use a high-precision robot5 to pick up a small volume of dissolved compound from each well and repetitively deliver approximately 1 nL of solution to defined locations on a series of chemically derivatized glass microscope slides (∼150 slides per print run). This results in the formation of microscopic spots of compound on the slides (200-250 μm in diameter). Each compound contains a common functional group that mediates covalent attachment to the slide surface. In this way, compounds are arrayed and subsequently immobilized on glass slides at extremely high spatial densities (>1000 spots per cm2). Each slide can then be probed with a different tagged protein and binding events can be detected by a fluorescence-linked assay. While the presence of a linker connecting the small molecule to the slide reduces the number of binding modes available to each compound in the library, it also enables the selection of compounds with known sites of attachment. This information can facilitate the preparation of small-molecule dimerizers, the purification of target proteins by affinity chromatography, and the measurement of binding kinetics by surface plasmon resonance. Successful implementation of this technique requires robust attachment chemistry. We have explored several approaches and currently favor the use of a Michael addition. Plain glass slides (2.5 cm × 7.5 cm) are derivatized as shown in Figure 1 to give surfaces that are densely functionalized with maleimide groups.6 Thiol-containing compounds readily attach to the surface upon printing, presumably via the expected thioether linkage. A critical feature of the binding assay is the observed lack of nonspecific protein-binding to the slide surface in aqueous buffer, which we attribute to the hydrophilicity of the maleimide functionality. To test the idea of SMP, we chose three unrelated molecules for which specific protein receptors are available. Compound 1 (R ) OH) is the vitamin biotin, which is recognized by the bacterial protein streptavidin.7 Compound 2 (R ) OH) is a derivative of the steroid digoxigenin and is recognized by the mouse monoclonal antibody DI-22 (Sigma). Finally, compound 3 (R ) OH) is a synthetic pipecolyl R-ketoamide, which was designed to be recognized by the human immunophilin FKBP12.8 Each of these compounds was attached to 425 μm diameter polystyrene beads via a 6-aminocaproic acid linker and either 4-methoxytrityl-protected cysteine (X ) S(Mmt)) or alanine (X ) H; negative control). To create reference points on the slides, we also prepared beads with a thiol-labeled derivative of tetramethylrhodamine (4a). Individual beads were placed in 28 separate wells of a 96-well plate and the compounds were deprotected, cleaved, and subsequently dissolved in 5 μL of DMF. The released compounds were then arrayed robotically onto a series of maleimide-derivatized slides with a distance of 300 μm between the centers of adjacent spots. Each slide was printed according to the pattern illustrated in Figure 3D. After 12 h, the slides were blocked with 2-mercaptoethanol, washed extensively, and probed with different proteins. The slide in Figure 3A was probed with Cy5-conjugated streptavidin, washed, and scanned using an ArrayWoRx fluorescence slide scanner. The slide was scanned for tetramethylrhodamine fluorescence (false-colored green) and Cy5 fluorescence (false-colored red). As anticipated, only the spots containing 1a were visible when scanned for Cy5 fluorescence, indicating that localization of streptavidin on these spots was both specific for biotin and dependent on the thiol functionality. Using a twostep detection method, the slide in Figure 3B was probed first with DI-22 and then with a Cy5-conjugated goat-anti-mouse (1) (a) Furka, A.; Sebestyen, F.; Asgedom, M.; Dibo, G. Int. J. Pept. Protein Res. 1991, 37, 487-493. (b) Lam, K. S.; Salmon, S. E.; Hersh, E. M.; Hruby, V. J.; Kazmierski, W. M.; Knapp, R. J. Nature 1991, 354, 82-84. (2) Nestler, H. P.; Bartlett, P. A.; Still, W. C. J. Org. Chem. 1994, 59, 4723-4724. (3) Tan, D. S.; Foley, M. A.; Shair, M. D.; Schreiber, S. L. J. Am. Chem. Soc. 1998, 120, 8565-8566. (4) Lam, K. S.; Lebl, M.; Krchnak, V. Chem. ReV. 1997, 97, 411-448. (5) Schena, M.; Shalon, D.; Davis, R. W.; Brown, P. O. Science 1995, 270, 467-470. (6) Experimental details are available via the Web at http://wwwschreiber.chem.harvard.edu/home/protocols/SMP.html. (7) Chaiet, I.; Wolf, F. J. Arch. Biochem. Biophys. 1964, 106, 1-5. (8) Holt, D. A.; Luengo, J. I.; Yamashita, D. S.; Oh, H.-J.; Konialian, A. L.; Yen, H.-K.; Rozamus, L. W.; Brandt, M.; Bossard, M. J.; Levy, M. A.; Eggleston, D. S.; Liang, J.; Schultz, L. W.; Stout, T. J.; Clardy, J. J. Am. Chem. Soc. 1993, 115, 9925-9938. Figure 1. Preparation of maleimide-derivatized glass slides. 7967 J. Am. Chem. Soc. 1999, 121, 7967-7968