Methods for solid phase nanoextraction and desorption

Abstract

Multiplexed assays may be performed using extraction probe comprised of solid supports and extraction phases.

Description

FIELD OF THE INVENTION

The present invention relates generally to the separation and analysis of complex materials, specifically biological materials. More particularly, the present invention relates to methods for the multiplexed separation and/or characterization of components of complex biological mixtures utilizing solid phase extraction techniques, preferably on a micro- or nanoscale. In some preferred embodiments, the present invention employs combinatorially derived extraction phases on nanoparticles to extract analytes from a sample.

BACKGROUND OF THE INVENTION

A variety of methods have been developed for the separation of mixtures for analysis (e.g., filtration, chromatography, extraction, electrophoresis, etc.). However, these methods have not proven sufficient for the separation of biological samples (e.g., blood, plasma, serum, synovial fluid, cerebrospinal fluid, saliva, tears, bronchial lavages, urine, stool, excised organ tissue, bone marrow, etc.). Such samples are comprised of a complex and heterogeneous mixture of molecular and cellular material in which certain components may be quite abundant, while others are present in only trace amounts. The separation and analysis of these types of samples have presented challenges to scientists using conventional techniques.

For example, the currently preferred method of performing proteome analysis (“proteomics”) uses two-dimensional (2-D) gel electrophoresis to separate complex protein mixtures. After electrophoresis and staining, the revealed spots of the gel are excised. The protein is then separated from the gel and subjected to enzymatic digestion. The resulting peptide fragments are then typically characterized by mass spectrometry, such as Matrix-Assisted Laser Desorption Ionization Time-of-Flight (MALDI-TOF MS) or electrospray ionization (ESI-MS). The original protein structure is then reconstructed by matching the peptide masses against theoretical peptide masses for known proteins that can be found in protein sequence databases, such as SWISS-PROT. Shortcomings of this technology include the lack of reproducibility of the 2-D gel process, difficulties in protein quantitation, and sample loss when recovering the protein from the gel. 2-D gels also suffer from a separation bias against proteins (and other molecules) of very low and very high molecular weight, and against proteins with the same pI. Accordingly, 2-D gels cannot be used for profiling small organic molecules, chemokines, metabolites, and so on. Many molecules known to be important in various disease states (e.g., cholesterol, thyroid hormone, etc.) are, therefore, not detectable by this method.

Specific affinity binding is an additional technique used to capture specific target ligands from complex mixtures such as biological fluids. For example, monoclonal or polyclonal antibodies, may be immobilized on a surface. When the surface is contacted with the sample, the antibodies bind to components of the mixture. Analysis can be carried out via competitive binding, or in sandwich mode using a secondary antibody. In both modes, there is typically a tag (enzyme, radiolabel, fluorophore, etc.) that is used for detection and/or amplification. Specific affinity binding techniques have been applied to proteomics in order to characterize gene products. Although it is highly specific, such immunoseparation has many of the same drawbacks as other assays that take place in two dimensions. Moreover, immunoseparation fails when there is no high-affinity antibody available to components in the sample, which is often the case. In particular, immunoseparation fails for (i) unknown molecules, (ii) known protein molecules that are post-translationally modified at or near the high affinity epitope; and (iii) molecules too small to elicit a strong immune response.

One specific affinity binding approach to proteomics where the analysis is limited to known proteins (i.e., proteins for which antibodies are commercially available), is the state-of-the-art FlowMetrix system developed and commercialized by Luminex Corp. (Austin, Tex.). The FlowMetrix system uses microspheres as the solid support for performing multiplexed immunoassays. Currently Luminex offers 64 different bead sets. Each bead set can, in principle, support a separate immunoassay and the beads are read using an instrument similar to a conventional flow cytometer. A major limitation of the Luminex approach is that the frequency space of molecular fluorescence used both for microsphere tagging and detection is not wide enough to accommodate nearly as many different assays as would be desirable to fully realize the advantages of multiplexing.

Solid phase microextraction (SPME) is a solvent-free separation technique that combines sampling and analyte concentration. The basic process of solid phase extraction involves adsorption of one or more target analytes from a sample matrix into a solid “extraction” phase. During the extraction, exposure of the extraction phase to the sample leads to the partitioning of analyte between the sample and extraction phases. The amount that any particular analyte is extracted from the sample depends on a number of factors, including the partition coefficient.

A device for performing SPME was the subject of U.S. Pat. No. 5,691,206, entitled, “Method and Device for Solid Phase Microextraction and Desorption.” As described therein, a thin coat of polymer or other extraction phase is coated on a fused silica fiber. The coated fiber, or probe, is contained within a hollow needle extending from the barrel of a syringe-like apparatus and can be extended or retracted using a plunger. To extract analytes from a sample, the needle is inserted into the sample and the coated fiber is extended into the sample. The sample matrix containing the analytes can be a gaseous sample, a liquid sample, or even the headspace above a liquid sample. After the microextraction has been allowed to take place, the fiber is retracted and the needle is removed from the sample.

The extracted analytes can then be delivered to a suitable instrument for analysis. SPME has been successfully coupled to high pressure liquid chromatography (HPLC) and gas chromatography (GC). For analysis by mass spectrometry (MS), analytes adsorbed into the extraction phase may be thermally desorbed and studied by MALDI-MS or Surface Assisted Laser Desorption Ionization mass spectrometry (SALDI-MS), or the analytes may be ionized by electrospray techniques.

SPME has been used for numerous applications in pharmaceutical science, environmental science, biological science, and chemical science. In short, SPME can be used for any application in which chromatographic separation is desired. In many contexts, SPME is simpler, faster and produces extracts of greater purity than traditional solvent-solvent extractions. SPME has been successfully used, for example, to extract pyrazines from peanut butter, fatty acids from milk, and amphetamines from biological fluids.

As it is currently practiced, SPME has several important limitations. First, performing SPME using a single fiber does not allow for multiplexing. The single needle method described in the literature would be of limited value for larger scale efforts that require many experiments to be run simultaneously in the same sample. It would be impracticable, for example, for a full-scale proteomics effort to rely on existing SPME.

Second, the limited number of solid extraction phases currently available necessarily limits SPME's selectivity as a separation technique. In the original SPME literature, the extraction phase associated with the fiber probe was polydimethylsiloxane (PDMS) or polyacrylamide (PA). These materials possess the fundamental properties necessary to effect SPME—they are chemically stable; are able to be cast as a thin film; have a semi-porous or porous geometry, and have a reasonably high affinity for one or more classes of molecules. In particular, PDMS has a high affinity for non-polar organics and PA has a high affinity for polar organics. However, neither material exhibits particularly high affinity for water-soluble species. Efforts to address the limited selectivity of SPME extraction phases have met with only limited success—there are now roughly ten different commercially available extraction phases for use in SPME. However, considering the diversity of structure present in the proteome, as well as in the roughly 10,000 different low molecular weight species known to be present in blood, it is clear that SPME in its current method of practice—using single needle extractions and a small number of different extraction phases—is of limited utility for comprehensive profiling of biological samples.

In some cases, researchers have resorted to using two or more different separation methods in order to profile complex mixtures. However, such “hyphenated separation techniques” generally require increased sample volume and have been hampered by incompatibilities with respect to different separation techniques and the methods eventually used to analyze the separated analytes.

Superimposed on the challenges presented using conventional techniques to analyze biological samples, is the pressure to do so faster and with smaller sample sizes. Indeed, advances in medicine and biology have resulted in a paradigm change in what is traditionally defined as bioanalytical chemistry. A major focus of new technologies is to generate what could be called “increased per volume information content.” This term encompasses several approaches, from reduction in the volume of sample required to carry out an assay, to highly parallel measurements (“multiplexing”), such as those involving immobilized molecular arrays, to incorporation of second (or third) information channels, such as in 2-D gel electrophoresis or CE-electrospray MS/MS. It also encompasses efforts to achieve miniaturization of the machinery of analysis—as in Bio-Microelectromechanical systems (Bio-MEMS), microfabricated devices using silicon, glass and polymer substrates that have been utilized in electrophoresis, electrochemistry and chromatography to reduce sample volume and increase speed and throughput. (See, e.g., Manz, A., Becker, H. Eds., “Microsystem Technology in Chemistry and Life Science,” Springer-Verlag: Berlin (1998)).

Unfortunately, many of these seemingly revolutionary technologies are limited by a reliance on relatively pedestrian materials, methods, and analyses. For example, the development of DNA microarrays (“gene chips”) for analysis of gene expression and genotyping by Affymetrix, Inc., Incyte Genomics and others provides the wherewithal to immobilize up to 20,000 different fragments or full-length pieces of DNA in a spatially-defined 1 cm 2 array. At the same time, however, the use of these chips in all cases requires hybridization of DNA in solution to DNA immobilized on a planar surface, which is marked both by a low efficiency of hybridization (especially for cDNA) and a high degree of non-specific binding. It is unclear whether these problems can be completely overcome. Moreover, there is a general sense of disillusionment both about the cost of acquiring external technology and the lead-time required to develop DNA arraying internally.

A second example of how groundbreaking techniques can be slowed by inferior tools, is in pharmaceutical discovery by combinatorial chemistry. Solution phase, 5-10 μm diameter latex beads are used as sites for molecular immobilization in some protocols. Exploiting the widely adopted “split and pool” strategy, libraries of upwards of 100,000 compounds can be simply and rapidly generated. As a result, the bottleneck in drug discovery has shifted from the synthesis of candidates to screening, and equally importantly, to compound identification, (i.e., knowing which compound is on which bead). Current approaches to the latter problem include “bead encoding”, whereby each synthetic step applied to a bead is recorded by the parallel addition of an organic “code” molecule. Reading the code allows the identity of the drug lead on the bead to be identified. Unfortunately, the “code reading” protocols are far from optimal. In such strategies, the code molecule must be cleaved from the bead and separately analyzed by HPLC, mass spectrometry or other methods. In other words, there is at present no way to identify potentially interesting drug candidates by direct, rapid interrogation of the beads on which they reside, even though there are numerous screening protocols in which such a capability would be desirable.

Two alternative technologies with potential relevance both to combinatorial chemistry and genetic analysis involve “self-encoded beads”, in which a spectrally identifiable bead substitutes for a spatially defined position on a solid supporting chip. In the approach pioneered by Walt and co-workers, beads are chemically modified with a ratio of fluorescent dyes intended to uniquely identify the beads, which are then further modified with a unique chemistry (e.g., a different antibody or enzyme). The beads are then randomly dispersed on an etched fiber array so that one bead associates with each fiber. The identity of the bead is ascertained by its fluorescence readout, and analytes are detected by fluorescence readout at the same fiber in a different spectral region. The seminal reference (Michael et al., Anal. Chem., 22, 1242-1248 (1998)) describing this technology suggests that with 6 different dyes (15 combinations of pairs) and with 10 different ratios of dyes, 150 “unique optical signatures” could be generated, each representing a different bead “flavor”. A very similar strategy is used by at Luminex, that combines flavored beads ready for chemical modification (100 commercially available) with a flow cytometry-like analysis. (See, e.g., McDade et al., Med. Rev. Diag. Indust., 19, 75-82 (1997)). Once again, the particle flavor is determined by fluorescence, and once the biochemistry is put onto the bead, any spectrally distinct fluorescence generated due to the presence of analyte can be detected. Note that as currently configured, it is necessary to use one color of laser to interrogate the particle flavor, and another, separate laser to excite the bioassay fluorophores.

A significant limitation of self-encoded latex beads is that imposed by the wide bandwidth associated with molecular fluorescence. If the frequency space of molecular fluorescence is used both for encoding and for bioassay analysis, it is hard to imagine how, for example, up to 20,000 different flavors could be generated. This problem may be alleviated somewhat by the use of combinations of glass-coated quantum dots, which exhibit narrower fluorescence bandwidths. (See, e.g., Bruchez et al., Science, 281, 2013-2016 (1998)). If, however, it were possible to generate very large numbers of intrinsically-differentiable particles by some means, then particle-based bioanalysis would become exceptionally attractive, insofar as a single technology platform could then be considered for the multiple high-information content research areas; including combinatorial chemistry, genomics, and proteomics (via multiplexed immunoassays).

Surface derivatized probes consisting of self-assembled monolayers (SAMs) terminated with ionic functional groups also have been used for extracting peptides/proteins. (Warren et al., Anal. Chem., 70, 3757-3761 (1998)).

SPME followed by CE as the second dimension has been used to analyze a mixture of peptides from a proteolytic digest. (Tong et al., Anal. Chem., 71, 2270-2278 (1999)). Although the SPME-CE/MS improved the concentration detection limit by more than two orders of magnitude when compared to CE-MS alone, the large electro-osmotic force of the aminopropylsilane (APS) coated capillary tended to elute all the peptides in a relatively short period of time. This presents the possibility of confounding results owing to the co-elution of compounds.

A strategy has been used for the separation of MHC class I peptides, several thousand peptides at sub-femtomolar concentrations. The literature reports immuno-affinity concentration followed by reverse phase, and subsequently concentrated on specially designed membranes capillaries. (Tomlinson et al., J. Caromator. A, 744, 237-78 (1996)). In addition, a comprehensive two-dimensional separation technique has been described for profiling proteins. (Opiteck et al., Anal. Chem., 62, 1518-1524 (1997)).

There is a need for analytical methods of high sensitivity and selectivity that have the power to resolve and profile different components of a complex mixture, such as a biological fluid. At the same time, there is a need for such methods to be able to identify and preferably quantitate minute quantities of biomolecules in small sample sizes, potentially even in single cells.

There is also a need for streamlined and automated methods for analyte capture that are compatible with sophisticated separation and detection technologies, such as HPLC, GC, CE, and MS.

There is also a need for methods of rapidly interrogating a biological sample that can be multiplexed. In particular, there is a need to have methods for separation and analysis of low-molecular weight organic molecules, peptides, and larger proteins simultaneously in a microvolume samples.

There is a need for combinatorially-derived extraction phases to extract analytes from a sample. In particular, there is a need for such surfaces that can be used in multiplexed analyses.

SUMMARY OF THE INVENTION

The present invention relates generally to methods for multiplexed separation and analysis of biological materials. More particularly, the present invention relates to methods for multiplexed characterization of components of biological materials utilizing solid phase extraction techniques, preferably on a micro- or nanoscale. The solid phase extraction methods of the present invention are accomplished using solid supports that have been coated or are otherwise associated with an extraction phase. In some preferred embodiments, the present invention employs combinatorially derived extraction phases on nanoparticle supports. In other preferred embodiments, the solid supports for the extraction phases are arrays of fibers. In some preferred embodiments, the present invention relates to methods and materials for performing solid phase extraction and analysis using nanoparticles coated with an extraction phase to extract analytes from a sample.

The resent invention includes a method for performing solid phase extraction using particles as the stationary support or probe. In preferred embodiments, such particles are differentiable from each other. In some preferred embodiments, the particles are nanobarcodes which allow extremely high-level assay multiplexing in solution, essentially combining the advantages of arrays (e.g., gene and/or protein chips) with the advantages of solution-based assays. Nanobarcodes may be used according to the present invention to simultaneously perform, for example, thousands of chemically and biochemically selective nanoscale extractions on samples, and then analyzing the extracted molecules, including by using mass spectrometry and/or fluorescence. Although not necessary to achieve the benefits of the present invention, the use of nanobarcode technology increases the power of the analytical separation methods described herein.

In some preferred embodiments, the present invention includes methods of carrying out solid-phase nanoextractions (SPNE). Such methods preferably employ particles that are distinguishable from one another. Other solid supports included within the scope of the invention are beads. Also included within the scope of the invention are methods for the simultaneous use of a plurality of differentiable solid supports, each associated with a different extraction phase for solid phase nanoextraction.

BRIEF DESCRIPTION OF THE FIGURES

FIG. 1 illustrates examples of protein-protein assays on nanobarcodes.

FIG. 2 illustrates the preparation of nanobarcodes possessing self-assembled monolayers terminating in carboxyl functionalities.

FIG. 3 illustrates the preparation of a library of gamma-hydroxy amides of dextran coated nanobarcodes.

FIG. 4 illustrates the preparation of streptavidin-coated nanobarcodes.

FIG. 5 illustrates multiplexed molecular recognition with microparticles.

DETAILED WRITTEN DESCRIPTION OF THE INVENTION

The present invention is directed to novel separation and analytical methodologies. In particular, the present invention is directed to methods for separation and analysis that can be multiplexed. Most generally, multiplexing refers to multiple measurements on the same sample. These measurements can be carried out simultaneously or in rapid succession; preferably, they are carried out simultaneously. These measurements may be carried out in a single sample aliquot, or in a divided sample; preferably, the measurements are carried out in the same sample volume. Thus, multiplexing covers the range from multiple measurements taken simultaneously in a single sample, to multiple measurements carried out in different locations on a single sample (e.g., a gene chip) to multiple measurements carried out in succession on different sample aliquots. The common thread of multiplexing is multiple measurements on a sample.

Measurement, as used herein, may refer to any information about a sample, any operation performed on said sample mixture, e.g., the separation of components, the differential or non-differential concentration of analytes, or determining the existence and/or quantity of any analyte or class of analytes within such sample mixture. The methods of the present invention utilize extraction probes comprised of solid supports that are partially or totally coated, physically or chemically attached to, or in some way associated with an extraction phase.

In certain preferred embodiments the extraction probes of the present invention are uniquely derivatized nanoparticles. Such nanoparticle extraction probes may be introduced into the sample where they can independently assort in three-dimensions, allowing the extraction phase associated with each nanoparticle to contact and interact with analytes present in the sample. Once retrieved from the sample the nanoparticles can be directly interrogated via analytic methods. The retrieval may be by any means, including filtration, centrifugation, or magnetic means. As described more fully below, the method of the present invention is rapid and can be automated and multiplexed; in addition, the incorporation of the extraction phase into the nanoparticles also may be accomplished on an automated basis, using combinatorial methods to synthesize the extraction phase.

In other preferred embodiments, the extraction probes are arrays of derivatized fibers. The fibers can be sized in the range of 100 μm diameter or, preferably, they can be of less than 1 micron diameter. Configured as arrays, the fibers (or “needles”) can be coated with extraction phase and exposed to the sample in a multiplexed fashion that lends itself to automation. Furthermore, presentation to a suitable analytic instrumentation is also facilitated, when the extraction probe needles are configured in arrays.

In some preferred embodiments, the extraction phase is comprised of combinatorially-derived materials (e.g., those derived from a split/pool synthesis). (See Schultz et al., U.S. Pat. No. 5,985,356, entitled, “Combinator Synthesis of Novel Materials” and Wu et al., U.S. Pat. No. 6,045,671, entitled, “Systems and methods for the combinatorial synthesis of novel materials,” both incorporated herein in their entirety by reference.) The combinatorially-derived extraction phase is preferably a polymer. However, there are a number of other materials (e.g., inorganic materials, metal alloys, oxides, glasses, ceramics, zeolites, polyelectrolyte multilayers) or combinations thereof, that may be useful as combinatorially-derived extraction phases. Furthermore, the combinatorially-derived extraction phase may be generated randomly, synthesized in a more controlled manner, or chosen from available materials. For example, a series of extraction phases chosen from the available chromatographic literature is contemplated as being “combinatorially” derived. Likewise, a single extraction phase, used in different ways (i.e., at different densities, porosities, ph values, etc.) is contemplated as being “combinatorially” derived. In this way, a library of combinatorially-derived extraction phases is generated, selected, purchased, prepared or obtained by one means or another, such that each member will differ (to a greater or lesser extent) from the others, e.g., by their physical, chemical or functional properties. Accordingly, the various combinatorially-derived surfaces can be used to increase the selectivity of separation and analytical methodologies.

According to the present invention, the extraction probes are comprised of solid supports coated or otherwise associated with an extraction phase. In those embodiments of the invention comprising an array of extraction probes, each solid support may be associated with an extraction phase that may interact with the sample and the analytes in the sample differently. For example, assume a sample contained only three analytes, A, B and C. Extraction probe I is comprised of an extraction phase that will extract a quantity of analyte A from the sample, but will not extract any of analytes B or C. Extraction probe H comprises an extraction phase that extracts a quantity of analytes A and B from the sample, but not analyte C. Extraction probe III, however, only extracts analyte C. By the use of multiplexing, i.e., simultaneously contacting the sample with extraction probes I, II and III, one can detect for the presence of analytes A, B and C. By expanding this example to, for example, a system whereby 10,000 different extraction probes are utilized, it can be shown how a complex sample, such as a biological material, can be profiled. Some analytes within a sample will be extracted into a large number of the extraction probes, while other analytes will only be extracted by a few extraction probes. In addition, it will not just be a question of whether or not the analyte has been extracted—there will be degrees of extraction. By detecting the presence and quantity of any analytes extracted from each of the extraction probes, it is possible to profile virtually all of the analytes within a highly complex sample. By performing the extraction simultaneously, it is possible to generate a huge amount of information about a sample in any extremely short period of time.

The concentration of analytes via extraction into the extraction phase serves as a discrete separation process. The power of the invention arises from multiplexing—many of thousands of these discrete separations can be accomplished simultaneously.

Freestanding nanoparticles are the preferred solid supports to achieve the multiplexed separation and analysis of the present invention. In general, such a multiplexed separation and analysis would proceed as follows: The nanoparticles individually can be coated or otherwise associated with an extraction phase and then placed in contact with the sample. Because of the small size of the nanoparticles, only a small amount of sample is required; in some instances merely a few drops of whole blood. Then, after sufficient time has been allowed for the extraction to take place, the nanoparticles are recovered from the sample. Recovery may be accomplished by a number of known means (e.g., centrifugation, filtration, etc.). In some embodiments, the nanoparticle extraction probes can be made magnetic to facilitate recovery. Once recovered, the analytes extracted from the sample can be detected by a suitable analytic device; for example, by a mass spectrometer. In certain embodiments, the nanoparticle may be delivered to the analytic device by microfluidic means. Coated with combinatorially-derived extraction phases (as described more fully below), the nanoparticles allow a combinatorial separation (“combi-sep”) to proceed on a nanoscale.

In this embodiment, nanoparticle-based assays utilize the combination of solid phase extraction techniques, microfluidics and mass spectrometric techniques to accomplish differential proteomic and small organic molecule (“orgeomic”) profiling as well as detection of other parameters (e.g., presence of a pathogen) in a biological sample. In a series of preferred embodiments, this may include (i) placing a combinatorial self-assembled monolayer extraction phase on the nanoparticle for the purpose of extraction of any type of molecule or other analyte from a complex biological fluid; and (ii) using a nanoparticle/mass spectrometry interface to analyze the surface adsorbed molecules with nanoparticle specific ESI or matrix-free ionization-time of flight (SALDI-TOF) techniques.

Mass spectrometry is preferentially selected as a detection element for broad-based differential, comprehensive molecular analysis (differential molecular phenotyping) because of its applicability for both high and low molecular weight species. Indeed, it is presently the only technique capable of both furnishing molecular identification of peptide fragments associated with large proteins and molecular weight/identification of molecules in the 100-500 amu range. Moreover, insofar as several masses can be identified simultaneously, mass spectrometry is an inherently multiplexed detection technique.

Mass spectrometry is rapidly becoming the tool of choice for detailed identification and analysis of polypeptides and proteins. There are two widely-used methods for biomolecular sample introduction in mass spectrometry: Electrospray and matrix-assisted laser desorption/ionization (MALDI). In MALDI, the analyte of interest is embedded into a solid ultraviolet-absorbing organic matrix that vaporizes upon pulsed-laser irradiation, carrying with it the analyte. (See, e.g., Karas et al., Anal. Chem. 60, 2299-2301 (1988)). During this process the energy absorbed by the matrix is transferred to the analyte that is ionized. The gas phase analyte ion is then sent to the Time-Of-Flight (TOF) mass analyzer. MALDI-TOF is currently successfully utilized for the analysis of proteins, polypeptides and other macromolecules. Even though the introduction of an organic matrix to transfer energy to the analyte has advanced tremendously the field of desorption mass spectrometry, MALDI-TOF still has some limits. For instance, the detection of small molecules is not practical because of the presence of background ions from the matrix. Also, MALDI experiments are inherently sensitive to matrix choice—matrix type as well as matrix amounts must often be tailored to the nature of the analyte (a severe limitation to the analysis of complex mixtures).

Recently, Sunner et al. have introduced the term SALDI for Surface-Assisted Laser Desorption/Ionization (Sunner et al., Anal. Chem. 67, 4335 (1995)). This technique is matrix-free, allows analysis of small organic molecules and yields performances similar to MALDI. Noble metal nanoparticles are a vastly superior choice for laser-based ionization, for two reasons. First, colloidal noble metal nanoparticles exhibit very large extinction coefficients in the visible and near IR. This contrasts with organic matrices. Second, irradiation of Au nanoparticles is known to lead to dramatic enhancements in electric field strength at the particle surface. This leads to increased ionization efficiencies and is the basis of surface-enhanced Raman scattering. Moreover, combined with nanobarcode technology, SALDI-MS becomes a powerful molecular fingerprinting tool.

The technical hurdle in the art associated with mass spectrometry-based approaches to differential molecular analysis concerns sample separation—that is, how to convert exceedingly complex samples containing thousands of species into simpler mixtures containing a dozen or fewer analytes—the problem to which the present invention is directed.

As more fully described below, nanoparticle extraction probes offer an important means to perform solid phase extraction on the nanometer scale, and to perform combinatorial extractions, whereby thousands of chemically distinct extractions are performed simultaneously.

Clearly, the ability to use distinctly coded particles significantly increases the power of the present invention. Perhaps most importantly, the use of such nanoparticle extraction probes allows the separation and analytical processes to be multiplexed. Each type (“flavor”) of nanoparticle can be provided with a unique extraction phase. Given the diversity of structure present in the proteome, as well as in the orgeome (low molecular weight species), the greater the number of differentiable particles possible the more utility they would have for comprehensive profiling of biological samples. Combining a large number of differentiable nanoparticles as solid supports complements the ability of combinatorial chemistry to create an equally large number of materials for inclusion in extraction phases. Such combinatorially-derived extraction phases could capture several analytes simultaneously on distinctly coded particles from biological samples.

Such combinatorially-derived extraction phases can be made of any material. In certain preferred embodiments they are polymers. However, they can be composed of any material, e.g., oxides, glasses, ceramics, clays, zeolites, dendimers, oligomers, macromolecular complexes, supramolecular assemblies. To illustrate: A library of synthetic organic polymers could be obtained using combinatorial chemistry techniques, each member having its own particular properties. Then, the encoded nanoparticle solid supports could be uniquely coated with one member of the combinatorially-derived polymer library. An assembly or array of nanoparticles could be prepared, each nanoparticle with its own extraction phase coating and associated properties. This results in a library of unique nanoparticle extraction probes. Because multiple nanoparticle extraction probes (with different extraction phases) can then interact with the sample at the same time, and because it is known (or can be determined) what extraction phase was attached to a given nanoparticle solid support, differential analysis is possible.

The analysis of complex mixtures, such as biological fluids usually involves the removal and concentration of the analytes of interest. Solid supports, such as nanobarcodes can be coupled to a large number of methods for analyte capture. Most require derivatization of the solid support with an extraction phase which has an affinity for a component or a set of components which may be present in the sample. Thus, for example, a solid support could be derivatized with a thin coating of polydimethylsiloxane (PDMA), an extraction phase that is used to coat the fiber in some conventional SPME applications. The PDMA-coated solid support when put in contact with the sample will selectively adsorb a set of non-polar organic molecules. Similarly, in another experiment, a nanobarcode solid support could be coated with polyacrylamide (PA). Indeed, nanobarcodes coated with SPME polymers would allow a number of extraction phases for concentration of the analytes on the particle and delivery of the analytes to be resolved after desorption. Because nanobarcodes are differentiable, experiments could be done simultaneously in the same sample. In other words, nanobarcode extraction probes allow the SPME process to be multiplexed.

The wider the range of extraction phases used, the more powerful the separation. By appropriately selecting and designing distinct extraction phases, there is an opportunity to create a large library of different extraction probes. The extraction phases can be prepared by a combinatorial process. Such combinatorially-derivatized nanoparticles will provide surfaces with varying characteristic extraction phases that will allow extraction of a wide variety of molecules present in the biological sample. Being able to sample a much larger fraction of the number of molecules in a sample would significantly increase the utility of SPME as an analytic technique. In this way, multiple SPME extractions would occur, either in parallel or in series, each sampling some region of “molecular structure space.” For example, a set of SPME experiments could be performed in which each nanoparticle would be comprised of an extraction phase with one of a spectrum of different organic phase polymers.

This concept can be further illustrated by the following examples. At one extreme, the extraction phases could be highly specific for a certain analyte. Thus, monoclonal antibodies could be used as part of the extraction phase, either bound directly to the solid support or affixed to a polymer on the solid support. In theory, the extraction phase in this instance would exhibit high affinity for one and only one molecule. In real biological systems, however, antibodies to particular proteins will capture the protein of interest as well as any post-translationally modified species for which the modification does not significantly abrogate the antibody-antigen interaction. To capture a large fraction of molecules using this approach would require a large number of parallel or serial experiments, each employing a different antibody. A fundamental limitation of this approach is the finite number of monoclonal antibodies available and—equally significantly—the inability to capture completely new or novel proteins (i.e., those for which no antibodies are available).

At the other extreme, the extraction phase could have a low affinity for a large number of molecules, e.g., a C18 reversed phase typically used for HPLC. Here, few species will bind with high affinity, but it will be possible in principle to extract new substances, that are present at a sufficiently high concentration, and have a partition coefficient appreciable enough to lead to non-negligible amounts being concentrated in the stationary phase.

It should be noted that from a fundamental perspective, there is no difference between a “partition coefficient,” as the term is used in standard chromatographic and separation science texts, and an “affinity constant,” as that terms is used in bioanalytical work—the only difference is one of degree. In essence, both the partition coefficient and the affinity constant, in conjunction with other useful (and likewise analogous) parameters such as loading or capture reagent concentration/surface coverage, allow for the accurate prediction of what will happen to a particular molecule in the presence of an extraction phase. Immobilized high-affinity receptors carry out the exact same chemistry as traditional SPME extraction phases, although with higher affinity for a particular species and with greater selectivity. In certain applications, this increased affinity and selectivity can be advantageous; in others, a low affinity, low selectivity extraction phase is preferred. In still others, it will be a combination of high affinity and low affinity, high selectivity and low selectivity extraction phases that can be used to maximum impact.

Nanoparticle solid supports may be associated with different extraction phases for the capture of analytes (e.g., proteins, organic molecules, or other components of the biologic matrix). The range of extraction phases is broad and can include, without limitation, hydrophobic materials, hydrophilic materials, acids, bases, polyclonal or monoclonal antibodies, aptamers, small molecule receptors, and combinatorial chemistry libraries. As described above, the nanoparticle extraction probes are mixed with the sample matrix under conditions that allow for analyte capture. The mechanism of capture depends on the nature of the extraction phase. For example, if the extraction phase is an SPME polymer, the analyte is captured by means of microextraction into the SPME polymer, if the extraction phase comprises an antibody, the analyte is captured by means of specific binding to the antibody.

The use of extraction probes that are differentiable from one another allows this process to be multiplexed. Rod-shaped nanoparticles have been prepared whose composition varies along the length of the rod. These freestanding particles are referred to as nanoparticles or “nanobarcodes,” though in reality some or all dimensions may be in the micron size range. These nanobarcodes comprise a plurality of segments or stripes which may be comprised of different materials and may be functionalized on selected or all segments. The types of particles are differentiable based on the length, width, shape and/or composition of the particles. This allows a plurality of assays or measurements of analyte concentrations or activities to a plurality of analytes comprising contacting a solution that may contain said analytes with a plurality of nanoparticle extraction probes, wherein each nanoparticle extraction probe comprises an extraction (e.g., molecule, species or other material) that interacts with one of said analytes bound to the nanoparticle.

Thus, besides panning the biological sample for small organic molecules, peptides, and nucleotides, it is possible to take advantage of the nanobarcode technology to multiplex assays (e.g., immunoassays). The combination of nanobarcode technology, SALDI and fluorescence based immunoassays into one platform, as described below, for example, enables the generation of highly sensitive, quantitative, multiplexed immunoassays for known proteins. The ability to merge selectivity, sensitivity, multiplexing, quantitation and mass analysis in the same measurement offers, among other benefits, a minimum of 100-fold increase in sensitivity.

The protocol by which this advance is achieved is outlined below. First, a specific immunoassay is associated to each “flavor” of nanobarcode solid support by attaching a specific capture antibody as the extraction phase. The analyte is bound to the antibody-coated nanobarcode and is detected with a second antibody tagged with a fluorescent dye, which recognizes a different epitope on the analyte. Similarly, an analyte bound to its receptor could be detected with an appropriate second antibody tagged with a fluorescent dye which recognizes an epitope on the receptor.

This process can be done in the same sample for as many proteins as there are both capture and detection antibodies. Several hundred pairs of antibodies are currently available. Thus, this process makes it possible to simultaneously interrogate a biological sample for the presence of all known proteins for which matched antibody pairs are available. Moreover, only one fluorophore needs to be used for the entire panel of assays run in the sample.

Since the potential number of flavors of nanorods far exceeds the number of available reagents, the same solid support platform may also be able to detect post-translationally modified proteins, which are good candidates for new disease markers. Any protein can be subjected to co- and post-translational modifications. These modifications have an influence on the charge, hydrophobicity, and conformation with respect to the “parent protein”, and can occur at different levels. Modifications such as acetylation, phosphorylation, methylation, hydroxylation, N- and O-glycosylation, can occur at the cellular level as well as in extracellular fluids.

To detect post-translational modification, polyclonal antibodies raised against a protein are conjugated to a selected barcode solid support. The polyclonal antibody will capture not only the protein against which it has been raised, but also protein isoforms (i.e., proteins that share similar epitopes but are modified at different sites). If the isoform is recognized by the detection antibody, it will be quantitated along with the “parent protein” (i.e. by the fluorescence immunoassay). If post-translational modification has affected the epitope that is recognized by the detection antibody, the isoform will not be quantitated by fluorescence. If both the capture antibody and the detection antibody are polyclonal antibodies, at least a great number of the modified proteins will be quantified. After the fluorescence measurement, the nanorod extraction probes are subjected to mass spectrometry analysis.

Characterization of the proteins by SALDI-MS also identifies the different post-translation modifications. The SALDI-MS laser energy ruptures all non-covalent bonds allowing for detection of any molecule complexed on the extraction probes, including even the protein sandwiched by two antibodies. This highlights the importance of the ability to tag an assay with a specific nanobarcode extraction probe. The code on the extraction probe will be associated with a specific protein having a specific molecular weight. When this nanorod is analyzed, instead of a full scan analysis, the mass spectrometer may be tuned to concentrate on a particular mass by using a technique called Single Ion Monitoring (SIM). SIM mode will allow faster acquisition of data and will increase the analytical sensitivity (up to 1000-fold enhancement in detecting an ion in SIM mode versus detecting this same ion in full scan mode). With the knowledge of the expected mass (and the sequence) of the analyte, the mass analyzer may be focused on a mass range allowing the detection of all the possible isoforms related to the parent protein. The monitoring range may be set to the molecular weight of the parent +/−500 Da, for example. Determination of the molecular weight of the isoform reveals immediately the modifications that the parent protein has suffered. Thus, the combination of nanorod solid support and polyclonal antibodies has the advantage of localizing the parent proteins as well as the corresponding isoforms on one flavor of nanorod extraction probe. Thus, the nanorods allow for a connection between the “parent protein” and the corresponding isoforms. This is not the case for 2-D gel electrophoresis where the post-translationally modified protein can “appear” in a different place of the gel if the charge has changed (following phosphorylation, for example). In short, 2-D gel requires a large amount of additional effort (such as sequence determination) for the identification of the modified protein because the connection between proteins of the same family is missing.

The nanobarcode extraction probe platform will also enable the investigation of protein-protein interactions by incorporating specific proteins in the extraction phase on nanorods for screening the biological sample for potential entities capable of molecular recognition. Nanorod extraction probe technology combined to fluorescence-based quantitation and mass spectrometer-based identification also allows investigation of specific protein-protein interactions. The scheme in FIG. 1 depicts different assay formats to interrogate a biological sample for the presence of free analyte A ( FIG. 1A ) or for the presence of the free receptor R for analyte A (FIG. 1 C). These two formats have been described above. Another set of nanorod extraction probes labeled with antibody directed against A can be used to quantify analyte A bound to its receptor R by using a detection antibody directed against the receptor(FIG. 1 B). Similar assays can be set-up in which free auto-antibodies ( FIG. 1E ) and auto-antibodies binding analyte A ( FIG. 1D ) can be quantified using a fluorescent anti-Fc antibody, for example.

As described above, the nanorod extraction probes may be analyzed by SALDI-TOF MS to allow identification of the different isoforms present. However, even for cases where no detection antibodies are available, SALDI-TOF is still able to identify and characterize the different components. Quantitation by fluorescence will be missing, but identification by mass spectrometry of the captured analyte will still be possible. Alternatively a nanorod with an extraction phase comprising any protein can be used to pan the biological sample for the presence of a protein or any other entity having any affinity for said conjugated protein. The presence of the bound protein may be detected by mass spectrometric analysis.

All conceivable ligands that have been used in affinity chromatography can be used in extraction phases, and all the extraction probes may be combined in one single tube containing a minimum volume of biological sample. Accordingly, sample size will be greatly reduced compared to currently developed protein arrays.

The present invention also allows integration of various separation techniques with detection systems. As discussed above, in order to maximize the utility of available separation techniques, researchers have resorted to using two or more different separation methods to obtain separation. However, difficulties are often encountered in integrating the various separation techniques with each other and with the detection system. For example, it is often difficult to maintain resolution upon transfer to the second dimension. Another major difficulty is that there is often a lack of compatibility between the mobile phases and the detection system. For example, salts and detergents in the eluant are incompatible with electrospray mass spectrometry. As another example, additional procedures must be taken to “clean-up” a sample before MALDI-MS analysis is often necessary because the use of certain preservatives (e.g., chaotropes and solubilizing agents) suppress ionization.

Additionally, biological sources often contain a complex mixture of inorganic salts, buffers, chaotropes, preservatives, and other additives—often at higher concentrations to the molecules of interest—some of which are detrimental to MALDI MS. The approach used by the present invention is not only capable of generating as many extraction phases as the number of molecules present in a biological sample, it can also bind the molecules of interest to be analyzed by mass spectrometry. Accordingly, it provides an excellent mode of sample preparation prior to a analysis.

Nanoparticles as Solid Supports

Nanoparticles of the type which can be used as solid supports in the present invention are described in detail in U.S. application Ser. No. 09/598,395, filed Jun. 20, 2000, and its continuation in part, U.S. application Ser. No. 09/677,198, filed Oct. 2, 2000, “Colloidal Rod Particles as Nanobar Codes ” both incorporated herein in their entirety by reference. Also incorporated herein in their entirety by reference, are two U.S. Utility applications entitled “Methods of Manufacture of Colloidal Rod Particles as Nanobar Codes” and U.S. Application Ser. No. 09/676,890 “Methods of Imaging Colloidal Rod Particles as Nanobar Codes” each filed Oct. 2, 2000.

Because bar coding is so widely-used in the macroscopic world, the concept has been translated to the molecular world in a variety of figurative manifestations. Thus, there are “bar codes” based on analysis of open reading frames, bar codes based on isotopic mass variations, bar codes based on strings of chemical or physical reporter beads, bar codes based on electrophoretic patterns of restriction-enzyme cleaved mRNA, bar-coded surfaces for repeatable imaging of biological molecules using scanning probe microscopies, and chromosomal bar codes (a.k.a. chromosome painting) produced by multi-chromophore fluorescence in situ hybridization. All these methods comprise ways to code biological information, but none offer the range of advantages of the bona fide bar codes transformed to the nanometer scale.

The particles to be used as solid supports according to this embodiment of the invention are alternately referred to as nanoparticles, nanobarcodes, nanobar codes, nanorods rods and rod shaped particles. To the extent that any of these descriptions may be considered as limiting the scope of the invention, the label applied should be ignored. For example, although in certain embodiments of the invention, the particle's composition contains informational content, this is not true for all embodiments of the invention. Likewise, although nanometer-sized particles fall within the scope of the invention, not all of the particles of the invention fall within such size range.

In preferred embodiments of the present invention, the nanobarcode solid supports are made by electrochemical deposition in an alumina or polycarbonate template, followed by template dissolution, and typically, they are prepared by alternating electrochemical reduction of metal ions, though they may easily be prepared by other means, both with or without a template material. Typically, the nanobarcodes have widths between 30 nm and 300 nanometers, though they can have widths of several microns. Likewise, while the lengths (i.e. the long dimension) of the materials are typically on the order of 1 to 15 microns, they can easily be prepared in lengths as long as 50 microns, and in lengths as short as 10 nanometers. In some embodiments, the nanobarcodes comprise two or more different materials alternated along the length, although in principle as many as dozens of different materials could be used. Likewise, the segments could consist of non-metallic material, including but not limited to polymers, oxides, sulfides, semiconductors, insulators, plastics, and even thin (i.e., monolayer) films of organic or inorganic species.

When the nanoparticle solid supports of the present invention are made by electrochemical deposition the length of the segments can be adjusted by controlling the amount of current passed in each electroplating step; as a result, the rod resembles a “bar code” on the nanometer scale, with each segment length (and identity) programmable in advance. The same result could be achieved using another method of manufacture in which the length or other attribute of the segments can be controlled. While the diameter of the rods and the segment lengths are typically of nanometer dimensions, the overall length is such that in preferred embodiments it can be visualized directly in an optical microscope, exploiting the differential reflectivity of the metal components.

The synthesis and characterization of multiple segmented particles is described in Martin et al., Adv. Materials, 11, 1021-25 (1999). The article is incorporated herein by reference in its entirety. Also incorporated herein by reference in their entirety are U.S. Provisional Application Serial. No. 60/157,326, filed Oct. 1, 1999, entitled “Self Bar-coded Colloidal Metal Nanoparticles”; U.S. Provisional Application Ser. No. 60/189,151, filed Mar. 14, 2000, entitled “Nanoscale Barcodes”; U.S. Provisional Application Ser. No. 60/190,247, filed Mar. 17, 2000, entitled “Colloidal Rod Particles as Barcodes”; and U.S. Provisional Application Ser. No. 60/194,616, filed Apr. 5, 2000, entitled “Nanobarcodes: Technology Platform for Phenotyping.”

In certain preferred embodiments, the nanoparticle solid supports of the present invention are defined in part by their size and by the existence of at least 2 segments. The length of the particles can be from 10 nm up to 50 μm. In preferred embodiments the particle is 500 nm-30 μm. In the most preferred embodiments, the length of the particles of this invention is 1-15 μm. The width, or diameter, of the particles of the invention is within the range of 5 nm-50 μm. In preferred embodiments the width is 10 nm-1 μm, and in the most preferred embodiments the width or cross-sectional dimension is 30 nm-500 nm.

As discussed above, in certain preferred embodiments, the nanoparticle solid supports of the present invention are characterized by the presence of at least two segments. A segment represents a region of the particle that is distinguishable, by any means, from adjacent regions of the particle. Segments of the particle bisect the length of the particle to form regions that have the same cross-section (generally) and width as the whole particle, while representing a portion of the length of the whole particle. In preferred embodiments of the invention, a segment is composed of different materials from its adjacent segments. However, not every segment needs to be distinguishable from all other segments of the particle. For example, a particle could be composed of 2 types of segments, e.g., gold and platinum, while having 10 or even 20 different segments, simply by alternating segments of gold and platinum. A particle of the present invention contains at least two segments, and as many as 50. The particles of the invention preferably have from 2-30 segments and most preferably from 3-20 segments. The particles may have from 2-10 different types of segments, preferably 2 to 5 different types of segments.

A segment of the particle is defined by its being distinguishable from adjacent segments of the particle. The ability to distinguish between segments includes distinguishing by any physical or chemical means of interrogation, including but not limited to electromagnetic, magnetic, optical, spectrometric, spectroscopic and mechanical. In certain preferred embodiments of the invention, the method of interrogating between segments is optical (reflectivity).

Adjacent segments may even be of the same material, as long as they are distinguishable by some means. For example, different phases of the same elemental material, or enantiomers of organic polymer materials can make up adjacent segments. In addition, a rod comprised of a single material could be considered to fall within the scope of the invention if segments could be distinguished from others, for example, by functionalization on the surface, or having varying diameters. Also particles comprising organic polymer materials could have segments defined by the inclusion of dyes that would change the relative optical properties of the segments.

The composition of the nanoparticle solid supports of the present invention is best defined by describing the compositions of the segments that make up the particles. A particle may contain segments with extremely different compositions. For example, a single particle could be comprised of one segment that is a metal, and a segment that is an organic polymer material.

The segments may be comprised of any material. In preferred embodiments of the present invention, the segments comprise a metal (e.g., silver, gold, copper, nickel, palladium, platinum, cobalt, rhodium, iridium); any metal chalcognide; a metal oxide (e.g., cupric oxide, titanium dioxide); a metal sulfide; a metal selenide; a metal telluride; a metal alloy; a metal nitride; a metal phosphide; a metal antimonide; a semiconductor; a semi-metal. A segment may also be comprised of an organic mono- or bilayer such as a molecular film. For example, monolayers of organic molecules or self assembled, controlled layers of molecules can be associated with a variety of metal surfaces.

A segment may be comprised of any organic compound or material, or inorganic compound or material or organic polymeric materials, including the large body of mono and copolymers known to those skilled in the art. Biological polymers, such as peptides, oligonucleotides and carbohydrides may also be the major components of a segment. Segments may be comprised of particulate materials, e.g., metals, metal oxide or organic particulate materials; or composite materials, e.g., metal in polyacrylamide, dye in polymeric material, porous metals. The segments of the particles of the present invention may be comprised of polymeric materials, crystalline or non-crystalline materials, amorphous materials or glasses.

Segments may be defined by notches on the surface of the particle, or by the presence of holes or perforations into the particle. In embodiments of the invention where the particle is coated, for example with a polymer or glass, the segment may consist of a void between other materials.

The length of each segment may be from 10 nm to 50 μm. In preferred embodiments the length of each segment is 50 nm to 20 μm. The interface between segments, in certain embodiments, need not be perpendicular to the length of the particle or a smooth line of transition. In addition, in certain embodiments the composition of one segment may be blended into the composition of the adjacent segment. For example, between segments of gold and platinum, there maybe a 5 to 50 nm region that is comprised of both gold and platinum. This type of transition is acceptable so long as the segments are distinguishable. For any given particle the segments may be of any length relative to the length of the segments of the rest of the particle.

As described above, the nanoparticle solid supports can have any cross-sectional shape. In preferred embodiments, the particles are generally straight along the lengthwise axis. However, in certain embodiments the particles may be curved or helical. The ends of the particles of the present invention may be flat, convex or concave. In addition, the ends may be spiked or pencil tipped. Sharp-tipped embodiments of the invention may be preferred when the particles are used in Raman spectroscopy applications or others in which energy field effects are important. The ends of any given particle may be the same or different.

In preferred embodiments of the invention, an array or assembly of solid supports is prepared. In certain embodiments, the members of the array are identical, while in other embodiments, the array is comprised of a plurality of different types of particles. In embodiments of the invention comprising assemblies of identical particles, the length of particles for particles in the 1 μm-15 μm range may vary up to 50%. Segments of 10 nm in length will vary ±5 nm while segments in 1 μm range may vary up to 50%. The width of such particles may vary between 10 and 100% preferably less than 50% and most preferably less than 10%.

The present invention includes imaging and distinguishing between members of arrays or assemblies of nanobarcodes made up of a plurality of nanoparticle solid supports that are differentiable from each other. The arrays of the present invention can include from 2 to 10 12 different and identifiable nanoparticles. Preferred assemblies include more than 10, more than 100, more than 1,000 and, in some cases, more than 10,000 different flavors of nanoparticles. The particles that make up the assemblies or collections of the present invention are segmented in most embodiments. However, in certain embodiments of the invention the particles of an assembly of particles do not necessarily contain a plurality of segments.

By “freestanding” it is meant that nanoparticle solid supports that are produced by some form of deposition or growth within a template have been released from the template. Such particles are typically freely dispensable in a liquid and not permanently associated with a stationary phase. Nanoparticles that are not produced by some form of deposition or growth within a template (e.g., self-assembled nanobarcodes) may be considered freestanding even though they have not been released from a template. The term “free standing” does not imply that such nanoparticles must be in solution (although they may be) or that the particles can not be bound to, incorporated in, or a part of a macro structure.

In the embodiments of the present invention where the nanobarcodes contain some informational content, or where an array of nanobarcodes contain a plurality of types of particles, the types of particles are differentiable apart from the nature of the extraction phase associated with each particle type. In this invention, the ability to differentiate particle types or to interpret the information coded within a particle is referred to as “interrogating” or “reading” or “differentiating” or “identifying” the nanoparticle. Such differentiation of particles may be read by any means, including optical means, electronic means, physical means, chemical means and magnetic means. The particle may even contain different sections that will be interrogated or read by different means. For example, one half of a particle may be comprised of segments whose pattern and shapes can be read by optical means, and the other half may be comprised of a segment whose pattern and shapes may be read by magnetic means. In another example, two different forms of interrogation may be applied to an entire particle, e.g., the shape or length of the particle may be read by optical means and the segment patterns by magnetic means.

A key property of certain embodiments of the nanoparticle solid supports of the present invention is that when the particles are segmented, differences in the reflectivities of the component metals can be visualized by optical microscopy. Thus, in a segmented Au/Pt/Au rod of 200 nm in diameter and 4-5 microns in overall length, the segments are easily visualized in a conventional optical microscope, with the Au segments having a gold lustre, and the Pt segments having a more whitish, bright lustre. Another key property of the materials is that the length of the segments, when they are prepared by alternating electrochemical reduction of two or more metal ions may be controlled (and defined) by (a) the composition of the solution and (b) the number of Coulombs of charge that are passed in each step of an electrochemical reduction. Thus, the widths and the number of the segments can be varied at will.

The ability to identify a nanobarcode via its reflectivity and the ability to modify their surfaces with biomolecules allows the nanobarcodes to be used as solid supports and optical tags simultaneously.

What distinguishes nanobarcodes from other types of optical tags, or indeed from any type of tag ever applied to a molecular system (including isotopic tags, radioactive tags, molecular tags for combinatorial beads, fluorescence-based tags, Raman-based tags, electrochemical tags, and other tags known to those of skill in the art,) is the essentially unlimited variability. With the ability to use 7 or more different metals, 20 or more different segments, and 4 or more different segment lengths, and with 3 or more different rod widths, there are essentially an infinite number of different nanobarcodes that can be prepared. Even with just two types of metals and just 10 segments, with just one segment length, and with just one rod width, over a thousand different types (“flavors”) of nanobarcodes can be prepared.

The nanoparticle solid supports of the present invention can be read using existing. instrumentation, e.g., chemical force microscopy, optical readers, etc. However, instrumentation and software specifically designed to identify nanobarcodes are also within the scope of this invention. Specifically included within the scope of the invention are modified Micro Volume Laser Scanning Cytometry (MLSC) apparatus and modified flow cytometer apparatus that can be used to image or read nanobarcodes.

It should be noted, however, that a variety of detection mechanisms for reading the nanoparticle solid support can be used, including but not limited to optical detection mechanisms (absorbance, fluorescence, Raman, hyperRaman, Rayleigh scattering, hyperRayleigh scattering, CARS, sum frequency generation, degenerate four wave mixing, forward light scattering, back scattering, or angular light scattering), scanning probe techniques (near field scanning optical microscopy, AFM, STM, chemical force or lateral force microscopy, and other variations), electron beam techniques (TEM, SEM, FE-SEM), electrical, mechanical, and magnetic detection mechanisms (including SQUID).

The sensitivity of hyperRaleight scattering may make it a particularly useful interrogation technique for reading the nanoparticle solid supports of the present invention. For example, see, Johnson et al., The Spectrum, 13, 1-8 (2000), incorporated herein in its entirety by reference.

In one embodiment of the invention the nanoparticle solid supports are not comprised of segments, but are differentiable based on their size, shape or composition. In this embodiment, each particle in an array or assembly of particles has at least one dimension that is less than 10 μm. In preferred embodiments, the particles have one dimension less than 500 nm, and more preferably less than 200 nm.

Such an array of particles, which can be made up of any material, is comprised of at least 2, preferably at least 3, and most preferably at least 5 types of particles, wherein each type of particle is differentiable from each other type of particle. In the preferred embodiment, since the types of particles may be comprised of a single material and since different types of particles may be comprised of the same material as other types of particles in the assembly, differentiation between the types is based on the size or shape of the particle types. For example, an assembly of particles of the present invention may be comprised of 5 different types of gold rod-shaped nanoparticles. Although, each type of rod-shaped particle has a width or diameter of less than 10 μm, the different types of particles are differentiable based on their length. In another example, 7 types of spherical silver particles make up an assembly. The different types of particles are differentiable based on their relative size. In yet another example, 8 types of rod-shaped particles, all composed of the same polymeric material, make up an assembly, although each type of rod-shaped particles have the same length, but they are differentiable based on their diameter and/or cross-sectional shape.

A further example of an array of nanoparticle solid supports that fall within the scope of this embodiment of the invention is an assembly of particles, each type of which may have the same size and shape (with at least one dimension less than 10 μm) where the particle types are differentiable based on their composition. For example, an array of particles of the present invention may be comprised of 5 different rod-shaped nanoparticles of the same size and shape. In this example, the different types of particles are differentiable based on the material from which they were made. Thus, one type of nanorod is made from gold, another from platinum, another from nickel, another from silver, and the remaining type from copper. Alternatively, each particle type may contain a different amount of a dye material, or a different percentage of magnetizable metal. In each case, a given particle type would be differentiable from the other particle types in the assembly or collection.

Of course, this embodiment of the invention includes arrays in which combinations of size, shape and composition are varied. The critical aspect of the array of particles of this embodiment is the fact that all particle types have at least one dimension less than 10 μm and that the particle types are differentiable, by any means, from the other particle types in the assembly. In this embodiment, the different types of particles may be associated with an extraction phase and the differentiable characteristics of the type of particles encode the nature of the extraction phase. By encoding the nature of the extraction phase, it is meant that the specific identifiable features of the nanoparticle can be attached selectively to a specific extraction phase, so that a key or log can be maintained wherein once the specific particle type has been identified, the nature of the associated extraction phase is known (or can be determined).

Needles as Solid Supports

Another means of achieving multiplexing of solid phase extraction on the micro- or nanoscale, is to employ arrays of extraction probes comprised of fibers (“needles”) as the solid support. Indeed, configured as arrays, the fibers can be coated with extraction phase and exposed to the sample in a multiplexed fashion that lends itself to automation. Furthermore, presentation to a suitable analytic instrumentation is also facilitated. When the needle solid supports are configured in arrays, solid phase microextraction may occur simultaneously. Needle arrays have never been used for SPME.

For example, the present invention contemplates a set of extraction probe needles being simultaneously inserted into an apparatus that provides a variable delay into a single analytic instrument for the analysis of analytes. This variable delay allows for sequential analysis; in other words, it is not necessary to have “N” analytical instruments for “N” needles; N extraction probe needles can be analyzed with a single instrument, if the analyses can be time-staggered. This approach is utilized for LC interfaces to mass spectrometry. The same concept would also work for introduction into LC, CE or GC analytic instrumentation.

The present invention includes an array of solid needles or rods that are coated with an extraction phase to provide for separation of analytes by differential affinity with respect to the extraction phase. The array may be comprised of solid supports that are needles, pinheads, rods or any other suitable solid objects. Extraction phases include, but are not limited to, C18, C8, hydroxyapatite, anion or cation exchange resins or material used for affinity chromatography, inorganic materials, metal alloys, oxides, glasses, ceramics, zeolites, polyelectrolyte multilayers or combinations thereof, etc.

An example protocol of the method of the present invention is as follows: The array of extraction probes is washed with a suitable reagent/solvent to disengage all bound molecules. The array is equilibrated in an appropriate solution to assure correct conditions for binding in the next step. The array is soaked with mild stirring in a mixture of analytes containing one or more components that may bind to the resin in stationary phase. The array is lifted from the mixture and introduced into a wash solution such that unbound analytes may be washed away. This step may be repeated several times. For extraction of bound analytes, the array is introduced into a solvent/solution that allows disengagement of the bound analytes into the extraction solvent. The extraction solvent then contains the analytes preferentially bound to the arrayed separation phase.

Experiments using the arrays can be designed in a number of ways. The experiments can use the same extraction phase in the array with different mixture samples. Alternatively, the extraction phase can be used to isolate different analytes from the same sample by employing different extraction phases on each rod/needle.

For example, DNA may be separated from salts using C18 resin as the extraction phase as follows. First, a 96-well array of C-18 coated needles is washed three times with Acetonitrile and then equilibrated in 50 mM triethylammonium acetate buffer, pH 6.5. Next, the array is soaked in a mixture of DNA and salts, allowing the DNA to bind to the C18 resin. Then the array is removed from the mixture and washed three times in the equilibration buffer (TEAA) to remove the unbound salts. Finally, the bound DNA is extracted into 50% Acetonitrile.

To further achieve the advantages of combinatorial SPME (“combiSPME”) of the present invention, it is important to have as large a number of extraction phases available for extraction as possible. This can be accomplished by increasing the number of arrayed extraction probes that are contacted with the sample. Because up to thousands of parallel experiments are contemplated for the preferred embodiments of the present invention, it would be impractical to use fibers of the size conventionally used in solid phase extraction techniques. Thus the arrayed extraction probes of the present invention can be reduced in size so that they are on the micron or submicron (i.e., nanometer) scale.

As conventionally practiced, the needle-shaped fibers used in SPME are injected into HPLC, CE, or GC instrumentation using traditional injection ports. However, the use of microfluidic devices enables the use of miniaturized collection means as an alternate detection/analysis embodiment. Current microfluidic devices typically may use anywhere from nanoliters to femtoliters of solvent. Such devices may be coupled to nanofiber extraction probes for analyte desorption and delivery. For such embodiments, conventionally sized needle-shaped fibers, on the other hand, would deliver far too much material to be practical.

Microfluidic devices may be replicated precisely and in large numbers at low cost. Because microfluidic-based separations occur in devices that have small physical dimensions, they may be stacked together, or otherwise concatenated. In fact, 1000 or more microfluidic devices may be clustered, each capable of extracting the contents of an individual nanofiber extraction probe. In contrast, it would be impractical to carry out 1000 sequential separations using one CE instrument, or even 100 with 10 CE instruments.

As technology advances, microfluidic devices become smaller, and it is useful for the extraction probe needles to become smaller. Thus, another aspect of the present invention is the use of needle extraction probe arrays that cannot be prepared by conventional means owing to their small size. For example, needles with a diameter of less than 1 micron (μm) may be prepared photolithographically. Using deep-ultraviolet or projection lithography, features as small as 100 nm are easily attainable. Through the use of this or equivalent technology, needle extraction probe arrays may be synthesized that are of nanometer dimensions. Likewise, a plate of wells—in which the well-well spacing is identical to spacing between needles—can also be fabricated. As described earlier for larger scale extraction probes, this would allow each needle to be coated with a different extraction phase, by filling each well with a different chemistry. This may be accomplished, for example, by putting into each well (of a multi-well plate) a single bead from a split pool synthesis. The compound on the bead may be known or unknown. The compound is then released from the bead by one of a number of means known in the art, and then reacted with a functional group on the surface of the solid support array of needles. In this way, each needle would harbor a different ligand. This needle extraction probe array could then be introduced into wells of another multi-well plate containing a sample (dividing the sample into aliquots in each well). The receptors in the sample with affinity (medium or high) for a particular needle-associated ligand will be extracted by the corresponding extraction phase. Alternatively, the needles from the array could be reacted with the same functional group so that each needle would harbor the same ligand. This needle extraction probe array could then be introduced into wells of another multi-well plate in which each well contains a different sample.

Another example of the application of ultra-miniature extraction probe needle arrays involves the use of lower affinity materials. As described below, libraries of 10,000 polymers may be generated using the technique of combinatorial materials synthesis wherein each of the polymers has a different structure and a different affinity for molecules or classes of molecules. Indeed, the synthesis could be designed to yield highly diverse properties for these materials. Alternatively, the combinatorially generated polymers could be selected (perhaps based on empirical tests) for their diverse properties. Subsequent immobilization of these combinatorially obtained polymers onto ultraminiature solid support needle arrays would enable a set of parallel SPNE experiments that broadly sampled the molecular structure space contained in a complex sample mixture. These experiments will involve the use of “nano-needles,” because the analyte volume will be split up into many thousand identical aliquots. For example, with 10,000 needles, and 10 microliters of total sample, each well would contain one nanoliter of solution. This translates to a cube of 0.01 cm on each side. In other words, the extraction probe needle would have to be no more than 10 microns long, and about 1 micron wide.

Needle extraction probe arrays do not require the use of combinatorially-derived libraries to furnish useful extraction phase chemistries. To the contrary, such needle arrays can exploit any collection or combination of extraction phase, from commercially available chromatographic resins to monoclonal or polyclonal antibodies to oxide materials.

The present invention allows several methods for solid phase micro- or nanoextraction of analytes in parallel. For example, chromatographic media could be placed as a micro column within a pipette tip called a Zip-Tip (Millipore) or, alternatively, coated on the inner surface of a pipette tip such as Supro Tip and Pro Tip (Amika Corporation/Harvard Apparatus). These are hollow objects with chromatographic media coated or as a plug in the hollow structure. A multichannel pipettor would allow parallel processing using these tips.

In addition to coated solid objects, the arrayed objects may be porous and comprised of packed chromatographic media in the pores. Indeed, in certain of such embodiments, each needle may also be attachable to a charge source that allows a range of voltages to be applied to any given needle in the array. In this way, the needle can differentially extract components of the sample based on charge. These arrayed, immobilized stationary chromatographic media can be used for isolation from a mixture of analytes that may preferentially bind to the selected media

Bead-based Solid Supports

Another aspect of this invention includes using a collection of beads as the solid support for producing a collection of bead-based extraction probes for performing SPME. The bead extraction probes are contacted with the sample to be analyzed and subsequently collected and separated (e.g., by centrifugation). The bound analyte can then be desorbed and analyzed.

In a simple embodiment of this approach, a collection of beads with the same extraction phase on each bead is used. This solution-based approach has several advantages over simple one-channel SPME as it is currently practiced. One important advantage is that equilibration time is shortened using a collection of beads. Because both the extraction phase and the analyte are mobile, encounters between them occur more frequently and the capture of analyte molecules is more rapid. This is of particular importance for applications in which the sample volume is large (e.g., in environmental samples), or where analyte is present in sufficiently low concentration to warrant concentration.

Another important advantage to using bead-based extraction probes is the increase in surface area available with small, three-dimensional particles compared to that of a fiber or needle. To illustrate: The surface area of a 5 mm cylinder of 100 micron diameter is approximately 1.57 mm 2 . When the same volume is made up from a collection of 6,000 beads of 5 micron diameter, the total surface area is 47.1 mm 2 , a factor of 30 greater. The increased surface area in the bead-based approach provides another factor that will lead to a more rapid equilibration. Equally important, an increase in the surface area means that the capacity of the extraction phase will increase, because a given volume is more accessible in the form of spherical particles, where 3-D diffusion is allowed, than in a monolithic solid.

Another advantage to using a bead-based extraction probe is the ability to access samples that would otherwise be difficult to obtain. For example, if one wanted to carry out solid phase micro- or nanoextraction on whole blood in circulation, one could use beads smaller than the diameters of capillaries. Alternatively, larger solid support beads could also be used if the animal were to be sacrificed at the end of the experiment, or if the bead extraction probes could be localized within certain body compartments (either natural or artificially created). To collect these beads, one could make them magnetic, thus allowing them to be readily removed from an organism after a certain time. Beads can be made magnetic by incorporation of magnetic material on the interior, exterior, or both. Magnetic retrieval allows the bead extraction probes to be isolated with minimal sample perturbation, for example, in applications where centrifugation is disfavored (i.e., in whole blood). Of course, numerous alternative methods of bead retrieval are available, including without limitation, centrifugation, gravity-based particle settling (in a non-gradient containing or gradient containing column, or even in solution), optical methods (e.g., optical trapping), and bead based flow/sorting methods (e.g., using cytometry and fluorescence-activated cell sorting (FACS)).

In addition, bead-based extraction probes may be collected using microfluidic devices using one of a number of different methods, including isoelectric focusing, dielectrophoresis, acoustic focusing, among a number of others. In addition to being collected, particles can also be sorted by dielectrophoretic trapping (see, e.g., Proc. SPIE, 4177, 164-173 (2000)) or by a number of analogous methods.

Bead-based SPME separations may be conducted in a number of ways. In one embodiment, numerous beads are used, each with the same extraction phase. More powerful is the embodiment where subsets of beads have different extraction phases, such as those described above with respect to nanobarcodes and needle arrays. For example, one could use eight different kinds of bead extraction probes, with high affinities for (i) carbohydrates, (ii) acids, (iii) bases, (iv) hydrophobic compounds, (v) hydrophilic compounds, (vi) aromatic compounds, (vii) metal cations, and (viii) inorganic anions. This list could be expanded, of course, to hundreds or thousands of different extraction phases as needed, encompassing specific capture agents such as oligonucleotides and/or antibodies as well as ligands for particular receptors, cofactors for proteins, and so forth. It will be appreciated by those skilled in the art that pairs of interacting molecules can be exploited in two ways: (1) with an extraction phase to capture a “ligand”, and (2) with an extraction phase to capture a counterligand “receptor.” The table below lists examples of ligands for inclusion in extraction phases to capture/extract specific molecules (counterligands) from biological samples.

LIGAND               COUNTERLIGAND
Cofactors            Enzymes
Lectins              Polysaccharides, glycoproteins
Nucleic acid         Nucleic acid binding protein
                     (enzyme or histone)
Biomimetic dyes      Kinases, phosphatases,
                     Dehydrogenases etc . . .
Protein A, Protein G Immunoglobulins
Metals ions          Most proteins can form complexes
                     with metal ions
Enzymes              Substrate, substrate analogues,
                     inhibitor, cofactors
Phage displays       Proteins, peptides, any type of protein
DNA libraries        Complementary DNA
Aptamers             Proteins, peptides, any type of protein
Antibody libraries   Any type of protein
Carbohydrates        Lectins
ATP                  Kinases
NAD                  Dehydrogenases
Benzamide            Serine Protease
Phenylboronic acid   Glycoproteins
Heparin              Coagulation proteins and other
                     plasma proteins
Receptor             Ligand
Antibody             Virus

It should be understood that countless other examples of specific interactions are known and may be exploited. Moreover, subsets of bead extraction probes may bind with high affinity to unknown partners as well (e.g., using a library of small molecule ligands in the extraction phase to target a receptor).

The different extraction phases that can be used on subsets of beads can be “low affinity” as well, including the standard extraction phases commonly used in SPME, and also including designed low affinity stationary phases. Furthermore, combinations of high affinity and low affinity bead extraction probes could be instrumental in sampling a large fraction of a complex sample.

While this section has focused on bead-based extraction probes, which are typically understood to be spherical, the present invention is not so limited. Beads, particles, or objects of any shape or size, can be used and are contemplated by the present invention as solid supports, so long as they can be made to be coated with, attached to or associated with an extraction phase. Thus, collections of spherical nanoparticles of any dimension could be used, as could cylindrically-shaped particles of any size. Indeed, a collection of particles of different sizes, shapes and compositions could be used as well. For example, one could use 10 nm diameter metal nanoparticles with a first stationary phase, 50 nm diameter latex particles with second extraction phase, 3 micron cylindrical oxide particles with a third extraction phase, and so on. In short, any single size, shape or composition of particles, or any combination of sizes, shapes, and compositions can be used. Note that when nanoparticles of any size or shape are used, the method may be referred to as solid-phase “nanoextraction” (SPNE). By analogy, the same is true for needle extraction probe arrays on the submicron scale.

It is not necessary that the beads or particles used for bead-based SPME or SPNE be encoded, that is, contain information that can be used to differentiate or identify the beads or particles. However, when such encoding is used, significant benefits are achieved. In particular, it allows identification of the extraction phase used to extract a compound or compounds of interest. While bead- or particle-based SPME or SPNE can be carried out with identical, non-distinguishable beads or particles, it complicates analysis in two significant respects. First, it impairs the ability to match the proper analysis to the proper analyte. To illustrate: The non-distinguishable particles are each coated with one of a number of extraction phases, each extraction phase having a different affinity and/or partition coefficient, and possibly with a different selectivity as well. When these bead extraction probes are contacted with a complex molecular mixture, each bead “samples” a different fraction. In such a circumstance, it is expected that analysis of the material bound to each particle (or some fraction of the particles) subsequently will be performed. Because the particles, each with a different extraction phase, capture different classes of analytes, the preferred analysis conditions could be completely different, and in unencoded extraction probes, this information is not available. For example, one would use very different ionization conditions (voltage, flow rate, etc.) for electrospray ionization (ESI) mass spectrometric analysis of positively and negatively charged species. However, without the information provided by the encoded extraction probe, such decisions cannot be made. Thus, even in this very simple case, where positively charged species are captured on one extraction probe, and negatively charged species on another, the use of unencoded extraction probe is a significant disadvantage. Mass spectrometry literature contains dozens of different analyte-specific ionization conditions, further increasing the value of particle encoding.

A second benefit of extraction probe encoding involves the subsequent use of the information gained following de-extraction. For example, if analysis of one particular extraction probe leads to the identification of a material of interest, it may be desirable to design an extraction phase that could bind more (or less) of the analyte. In such cases, knowledge of the identity of the extraction phase itself, as could be obtained using encoded extraction probes, is essential. In this example, exact identification of the analyte has occurred by mass spectrometry (or some other method) that furnishes chemical structure information (e.g., NMR, Raman, IR). The use of an encoded extraction probe is even more critical if detailed analysis of extracted compounds is not carried out. For example, if all that is known about a particular analyte is that is was partitioned onto an extraction probe, and upon extraction and chromatographic or electrophoretic separation by GC, LC, or CE exhibited a retention time of, say, 10 minutes (as detected by visible ultraviolet or fluorescence or any other method), the information has limited utility—it cannot be repeated, since it was not known from which bead the compound was eluted. More importantly, the experiment cannot be correlated to another experiment, because the specific identity of the extraction probe is unknown.

In contrast, if the bead extraction probe identity is known (say Bead A), the intensity of the peak at issue from Bead A in Sample 1 can be directly compared to the corresponding intensity from bead A in Sample 2. The ability to compare beads from different samples is essential to comprehensive phenotypic analysis via combinatorial separation, where it will likely be the case that the precise identification of every individual compound in a complex mixture, like blood, is less important (and less feasible) that a comparative sample to sample analysis, where the goal is to detect changes in individual species, or more likely, patterns of changes in species. In this case, which is envisioned as one of the primary ways in which combinatorial separations may be practiced, the use of multiple, distinguishable stationary phases is of limited benefit without a corresponding mechanism for extraction phase identification, e.g., via bead encoding.

Combinatorial SPME

One of the embodiments of the present invention is referred to herein as combinatorial SPME. That is, SPME in which a large number of empirically-chosen microextractions are carried out in parallel (or serially in rapid succession). This number can be as few as 3, and as great as 10,000,000. Useful embodiments include numbers between 4 and 100,000, and also between 10 and 1,000.

Combinatorial approaches have met with success in several fields. For example, combinatorial synthesis of possible drug candidates has found acceptance in medicinal chemistry. Likewise, combinatorial discovery of materials has become popular via approaches that lead to thousands to millions of unique compositions of polymers, oxides, ceramics, etc. These advantages can be brought to the generation of analytical surfaces, to be used as extraction phases.

Thus, a number of nanoparticles can be coated, each with different extraction phases, including, for example, an antibody or other solid phases resulting in a library of unique nanoparticle extraction probes. Multiple nanoparticle extraction probes can interact with the sample at the same time. Differential analysis is possible because it is known (or can be determined) what extracting phase was associated with a given nanoparticle.

The coating of the solid support may be accomplished using an automated approach. The components to be coated on the solid support can be obtained by combinatorial methods known in the art. Indeed, the synthesis of substituents comprising the extraction phase on the nanoparticle lends itself to combinatorial approaches where the automated, parallel synthesis of thousands, or even hundreds of thousands, of chemical variations is possible. The resulting diversity increases the resolution that can be obtained in the analysis of the biological sample. The greater number of extraction phases means that there is a greater chance that any given analyte will interact with an extraction probe. The large number of differentiable nanoparticles possible allows a diverse population of probes. Thus, the present invention allows for the creation of a vast library of nanoparticle extraction probes, with varying affinities for different molecules. When this diverse set of probes is added to a biological sample, incubated, washed, and analyzed by, for example, SALDI-MS, each nanoparticle probe will represent a particular extraction phase. As a whole, the ensemble will provide a fingerprint of the sample.

Such combinatorial SPME is distinct from multiplexed SPME. Multiplexed SPME refers to a number of SPME measurements that are carried out in parallel (or serially in rapid succession). Combinatorial SPME, in contrast, refers to an empirical approach to synthesis that emphasizes generation of a large number of random or semi-random structures in hopes of finding one or more with desired properties. Often, combinatorial syntheses are multiplexed in the sense that they are often carried out simultaneously in the same sample. For example, immunoassays in which multiple immunoassays are carried out simultaneously in the same sample volume are multiplexed, but are not combinatorial because they target a well-defined set of molecules. Similarly, when the species attached to the array surface in “gene chips” or oligonucleotide arrays are selected for the purpose of quantitation of complementary sequences, they may be multiplexed, but cannot be considered combinatorial.

Combinatorial SPME employs a large variety of stationary extraction phases with the goal of extracting as many species as possible from a complex mixture. The extraction phases contemplated by the present invention need not be limited to the type traditionally used for SPME needle experiments. Indeed, the extraction phases can run the gamut from monoclonal antibodies and oligonucleotides (that have high affinity for few species) to those used in chromatography or traditional SPME (i.e., with low affinity for a large number of species).

Examples of extraction phases that can be used in the combinatorial methods of the present invention include, but are not limited to, polymers, block copolymers, self-assembled monolayers and derivatives thereof, molecularly-imprinted polymers, hyperbranched polymers, dendrimers, polyelectrolytes, gels, glasses, oxides, ceramics, semiconductors, amorphous materials, nucleic acids, oligonucleotides, carbohydrates, polysaccharides, peptides, proteins, lipids, and other biological molecules. Additional examples include all known stationary phases that have been used in paper, thin-layer, liquid and gas chromatography. Additional examples also include individual members of combinatorial libraries or multiple members thereof. For example, if SPME could be carried out with an extraction phase comprising latex modified with particular organic compounds. That molecule in the extraction phase would exhibit a variable range of affinities for analytes in a sample. This could be replicated for numerous compounds from a library on numerous solid supports, each with a slightly different affinity.

Those skilled in the art will recognize that the extraction phase of a combinatorial SPME experiment can encompass virtually all known chemical structures, whether molecular or non-molecular (e.g., supramolecular, solid-state, etc.). Moreover, those skilled in the art will recognize that there are an arbitrarily large number of possible combinations of these SMPE extraction phases.

With the numbers of possible nanobarcodes being enormous, combinatorial chemistry allows creation of an equally large number of compounds for inclusion in the extraction phase. Thus, it becomes important to be able to attach these compounds to (or otherwise associate these compounds with) nanobarcodes in order to create combinatorially designed surfaces for analyte capture. Several methods are known in the art that could accomplish this attachment. They include: Self-assembled monolayers, monolayers that are not self-assembled, partial layers, deposited films and materials (including from gas phase and/or solution phase), multilayers, grown materials (i.e., deposited materials that are chemically transformed), etc. It should further be clear that the extraction phase could result from tranformation of the material intrinsic to the nanobarcode. In other words, the outermost portion an Al stripe can easily be converted to Al 2 O 3 , and this can be used as an extraction phase. Likewise, Si can converted to SiO 2 , W to WO 3 , and so forth. Indeed, an entire segment could be used as an extraction phase. For example, if one of the segments in a nanobarcode were porous glass, the entire segment itself could be used as an extraction phase. Moreover extraction phases could be prepared by addition of materials to the segments themselves. For example, a SiO 2 segment, upon treatment with a chiral reagent, could serve as an effective extraction phase for certain classes of chiral compounds. Finally, it should be clear to those skilled in the art that combinations of extraction phases on nanoparticles are both feasible, owing to the differentiable chemistry of non-identical segments, and desirable, insofar as a combination of extractions from various portions of an individual particle might comprise an improved separation relative to a single extraction from a single particle, or from a combination of single extractions from multiple particles. By the same token, it should be recognized that different amounts of the same extraction phase on different particles comprise distinct extraction phases, to the extent that they will bind different amounts of materials.

One means to generate these extraction phases is to use self-assembled monolayers (SAMs) terminated with reactive functional groups. These SAMs may be derivatized with libraries of reagents to give nanobarcode extraction probes with extraordinary variety in surface chemistry.

There are alternatives to using SAMs. For example, nanoparticles could be coated with polymers; the polymers could be synthetic organic polymers. The inventors have prepared nanoparticles that have at least one polymer attached to them. Each polymer coat serving as an extraction phase can have selected properties. Monomers may be attached to the nanoparticles and the polymerization reaction conducted directly on the surface of the nanoparticle. (E.g., Mirkin, “Preparation of Nanoparticles with Polymer Shells for Use in Assays.”)

Useful Polymers may be inorganic such as an amorphous silica (i.e., glass) coat that may be polymerized on top of a thioalkyl silane SAM. Resulting amorphous silica can possess chemically active functional groups. Polymers could be directly adsorbed to the nanoparticle surface. This polymer layer would be stabilized through multipoint attachment (non-covalent). Examples include polylysines, aminodextrans, or selected proteins.

Ideally, each extraction phase is uniquely designed to capture only one class of molecules. Such classes may include large molecules, such as proteins.

Self-assembled monolayers formed from co-carboxy substituted alkanethiols on the surface of gold have been used as model surfaces to study the interactions of proteins with surfaces. (Mrksich et al., JACS, 117, 12009 (1995)). Derivatization of such nanoparticles may be achieved by various chemical means. One way involves “capping” with water soluble mercapto derivatives, typically mercapto carboxylic acid or amines. The carboxyl or amines are subsequently used to covalently label proteins, peptides or nucleic acids to give biomolecular conjugates of these particles that can be used in biological assays. This is discussed in, for example, Spinke et al., Langmuir, 2, 1821 (1993); Willner et al., J. Am. Chem. Soc., 114, 10965 (1992); and Mrksich et al., J. Am. Chem. Soc., 117, 12009 (1995).

Mixed SAMs formed from hydrophobic (alkyl, phenyl) and hydrophilic (hydroxyl, oligo ethylene glycol), positively charged (quaternary ammonium) and negatively charged (carboxylate, phosphate, sulphonate) species can be used in extraction phases to recognize and bind various molecules. Mixed SAMs have been used to study the adsorption of fibrinogen, lysozyme, pyruvate kinase, RNAse and carbonic anhydrase. (See, Lahiri et al., Anal. Chem., 71, 777 (1999); Prime et al., Science, 252, 1164 (1991)).

According to the present invention, nanorod solid supports are prepared that possess SAMs that terminate with carboxyl functionality. This is achieved by reacting the nanoparticles with ω-carboxy alkanethiols. The carboxyl functionality is then activated to an anhydride for further reaction with a wide variety of amines with diverse functional groups. This is illustrated in FIG. 2 .

Another class of derivatives that would provide amine reactive functionality as well as prevent non-specific interactions with proteins is dextran lactones. These can be prepared from carboxymethyl dextran.

The initial derivatization of the nanoparticles can be accomplished with 3-mercapto propyl(trimethoxy)silane. Then, the silane alkoxy is exchanged with the free hydroxyls of a carboxymethyl dextran derived lactone. Subsequent cleavage of the lactone with amines carrying diverse functional groups will yield a library of gamma-hydroxy amides of dextran coated nanoparticles. These methods provide a common reactive intermediate that is easily prepared. This is illustrated in FIG. 3 .

The dextran-coated or hydrophilic SAMs simultaneously provide a surface that is resistant to non-specific interaction between the nanorod extraction probes and proteins having a wide range of molecular weights and isoelectric points. By appropriately choosing and designing structurally distinct amine reactants for derivatization, there is an opportunity to create a vast library of extraction phases. This may be prepared using a combinatorial process. These combinatorially-derivatized nanoparticles would present extraction phases with varying avidity for binding to the wide variety of molecules present in a biological sample. These will be expected to be provide much greater efficacy than has been described for Surface Enhanced Laser Desorption Ionization Mass Spectrometry (SELDI-MS). Furthermore, compared to protein “chip” technology, the use of nanobarcodes as extraction probes will provide access to a greater number of different extraction phases. In addition, nanobarcodes extraction probes will be able to achieve the interrogation using smaller sample volume, and will have a kinetic advantage (i.e., the small size of nanobarcodes make interaction with biological sample almost homogenous compared to planar surface). In the protein “chip” technology—with which SELDI-MS is used—the protein probes are immobilized on a planar surface. Compared to the three-dimensional assortment of free nanoparticles extraction probes, a two-dimensional approach is hampered by a decrease in the efficiency of interaction and a greater degree of non-specific binding. It is unclear whether these problems can be completely overcome.

In contrast to systems based on “chips,” affinity capture techniques using nanoparticles extraction probes will use off-line incubation steps for capturing the analytes (i.e., the nanobarcode will go into the sample while the sample will go onto the “chip”). Using nanoparticles as the extraction probes is inherently superior from a kinetic viewpoint because it results in more rapid capture of analytes. In addition, this approach is advantageous from mass action perspective to drive binding—the density of the binding determinant (i.e., ligand or capture agent) on the nanoparticle can be varied to accommodate the wide range of analyte concentrations that are encountered in a biological fluid.

Carbohydrate derivatized SAMs with varying densities have been used to address issues involving cell-surface carbohydrate-protein interactions. These surfaces can be tailored to recognize free saccharides and, at the same time, are designed to take advantage of multiple binding determinants for carbohydrates in glycoproteins, for example. This approach will provide extraction phases capable of binding a wide spectrum of molecules, from low molecular weight organic compounds to large proteins, which are addressable and amenable to analysis.

EXAMPLES

Example 1

Preparation of Carboxy Terminated Dextran Coated Nanobarcodes for Protein Conjugation

A solution of aminodextran (Molecular Probes, Eugene Oreg.; 10 kD, 1 amine/10sugar residues) was made by dissolving 100 mg of the solid in 1 mL of phosphate buffer (pH. 8.0, 10 mM). To this solution was added 375 μl of a 32 mM freshly made solution of SPDP (Pierce, Rockford, Ill.) in DMSO. The solution was vortexed and allowed to shake in an end over end shaker for 12 h. The SPDP derivatized aminodextran was then reduced with triscarboxyethyl phosphine (TCEP, Molecular Probes, Eugene, Oreg.) by adding 400 μL of a 32 mM TCEP solution. The mercapto aminodextran (80 mg/mL, 100 μL) thus obtained was incubated with 2×10 8 nanorods (Ag—Au—Ag stripes) with end over end shaking for 15 h. The particles were made to 1.0 ml by diluting with water, centrifuged at 14 k for 2 min, and the supernatant discarded. The particles were resuspended in pH 8.0 phosphate buffer (1.0 mL) by sonication and then washed by centrifugation at 14 k rpm.

The supernatant was discarded and the particles were subjected to a final resuspension, centrifugation followed by a final suspension in 100 μL of phosphate buffer (pH 8.0. 100 mM). A solution of succinic anhydride in DMSO (100 mg/mL) was prepared and added dropwise to the suspension of the dextran coated nanobarcodes. The succinic anhydride solution was added in 10 μL aliquots followed by 10 μL aliquots of 1.0(M) NaOH between each addition. A total of 10 additions were made over a period of 30 minutes. The particles were washed by the usual centrifugation, removal of supernatant followed by resuspension for a total of 3×1 mL washes. The carboxy-terminated dextran coated nanobarcodes were then stored in 100 μL of water. The presence of dextran was qualitatively determined by the anthrone test (Anal. Biochem, 68, 332-335 (1975)).

Example 2

Coupling of Streptavidin to Carboxy-Terminated Dextran Coated Nanobarcodes

A suspension of the carboxy-terminated dextran coated nanobarcodes in 100 μL of MES buffer (pH 6.1, 10 mM) was treated with 10 μL of sulfo N-hydroxysuccinimide (10 mg/mL, Pierce) followed by 10 μL of ethyl dimethylaminopropyl carbodiimide (10 mg/mL in H 2 O, Pierce). The mixture was shaken in an end over end shaker for 30 minutes and then gradually added to 50 μl of a 5 mg/mL solution of streptavidin in phosphate buffer (pH 8.0, 100 mM). The solution pH was adjusted to 8.0 with a few drops of 1.0M NaOH. The reaction mixture was shaken in an end over end shaker for 14 h. The volume was made to 1.0 mL with water and the nanobarcodes washed by repeated centrifugation, removal of supernatant and resuspension in a fresh buffer for a total of three cycles. The streptavidin coated nanobarcodes were stored in 100 μL of water at a concentration of 2×10 9 particles per ml. The streptavidin number per particle was determined from the depletion of fluorescence when different concentrations of particles were incubated with a fixed concentration of the biotin-fluorescein conjugate.

Example 3

Preparation of Oligonucleotide Particles

The oligonucleotide particles were prepared by incubating biotinylated (dT)21 mer (0.3 nmoles, HPLC purified from IDT Inc) with streptavidinated particles (2×10 8 particles, 1×10 5 streptavidin per particle) in a TRIS/EDTA buffer (100 μL, pH 8.0). The particles and the oligonucleotides were shaken in an end over end shaker for 1 h, after which it was washed by centrifugation, removal of supernatant and resuspension in water for a total of three cycles.

Example 4

Coupling Aminophenylboronic Acid to NHS-Linked 8.6 μm Beads

NHS-Ester linked beads (10 mg) were taken up in pH 9, TAPS buffer in a 1.5 ml Eppendorf tube. To the bead solution, 20 μl of aminophenylboronic acid (10 mg/ml) was added. The solution was rotated overnight. The beads were then centrifuged and the supernatant removed. The beads were then resuspended in fresh TAPS buffer and washed two more times before they were suspended to a final concentration of 50 mg/ml in TAPS buffer.

Example 5

Preparation of Streptavidin Coated Particles

A suspension of carboxy modified latex particles (8.6 microns, Bangs lab) was constituted to 20 mg/ml in MES buffer (pH 6.0, 10 mM). To a 1.0 ml solution was added 10 μl of a freshly prepared solution of EDAC (10 mg/ml) in water. The reaction mixture was incubated in an end over end shaker for 15 minutes. The activated bead suspension was then added to a streptavidin solution (100 μl of 0.5 mg/ml) and the pH adjusted to 7.5 with phosphate buffer. The coupling reaction was allowed to proceed for 2 hours and then quenched with 1 mL of 1M glycine (pH 9.0). The particles were then washed by centrifugation at 5 K for 10 minutes, decanting the supernatant and resuspending in water (1 mL). The centrifugation, decantation and resuspension were repeated for a total of three cycles. This is illustrated in FIG. 4 .

Example 6

Alternative Preparation of Streptavidin Coated Nanorods

SAM-coating of Nanobarcodes. A Nanobarcode solution (10 μl,˜1×10 8 rods/ml) was added to 500 μl of SAM solution (@100 mM in ethanol, freshly made). The solution was incubated overnight using either a stir bar on a stirring plate or the rotator. Then the solution was rinsed with 200 μl of ethanol and H 2 O mixtures at ratios of 1:0, 3:1, 1:1, 1:3, 0:1, respectively.

Example 7

Combinatorial Separation with Derivatized Particles

The following particles were used: 10 mg (8.6 microns, 3×10 7 particles) of boronic acid derivatized magnetic particles in 100 μL of TAPS buffer, 100 μg of streptavidinated particles (3×10 5 particles, 10 5 streptavidin per particle) in 100 μL of water and 10 mg of carboxy modified latex particles in 100 μL of water. The particles were added to a solution 15 μL of 1 mg/mL glucose+25 μL of 250 nM biotin-fluorescein (Molecular Probes) conjugate and 100 μL of 1 mM dioxadodecanediamine (Aldrich). The mixture was vortexed and allowed to shake in an end over end shaker for 12 h. The suspension was centrifuged and the supernatant analyzed for depletion of the analytes.

Biotin-Flourescein Conjugate Depletion Was Determined By Fluorescence (490 Excitation, 520 Emission) 58597 Units For Control And 3369 In Supernatant. Glucose depletion was determined by the anthrone test, (Anal. Biochem, 68, 332-335 (1975)) absorbance at 626 nm was 0.121 for control and 0.06 in the supernatant Diamine depletion was determined by the TNBS assay, absorbance at 415 nm was 0.620 for control and 0.332 in the supernatant. The control numbers in the above experiments refer to the analyte concentration measure before extraction, 300 μL of water was used instead of the particles.

Example 8

Reacting Boronic Acid Linked Beads with Glucose Reacting Streptavidin Beads with Biotin-Fluorescein Conjugate

To a 1.5 ml Eppendorf tube, 20 μl of streptavidin linked bead solution (5 mg/ml, 100 ug) was added. To the same tube, 200 μl of boronic acid linked bead solution (50 mg/ml, 10 mg) in pH 9 TAPS buffer was also added. To the dual bead solution, 15 μL of glucose solution (1 mg/ml) was added, and 25 μl of biotin/fluorescein solution (250 nM) was added. The tube was rotated for approximately two hours. The beads were then centrifuged and the supernatant was removed from the precipitated beads.

The supernatant was then tested for fluorescein depletion and glucose depletion. For fluorescein, compared to a standard solution representing 0% depletion (58597 units), the experimental value for the supernatant indicated over 95% depletion of fluorescein (3369 unit). For glucose, compared to a standard solution representing 0% depletion (A=0.121), the experimental value for the supernatant indicated 30% depletion (A=0.110) (The standard and experimental glucose solutions were increased in concentration by a factor of 3, so a 10% decrease in absorbance is really representative of 30% depletion). This is illustrated in FIG. 5 .

The foregoing examples are presented as illustrations and should in no way be considered as limiting the scope of this disclosure.

Claims

1. A method for extracting a plurality of analytes from a sample, comprising the steps of:providing a position-addressable array of extraction probes, each probe comprising a solid support and an extraction phase; providing an array of capillaries addressable by the array of extraction probes, the capillaries containing aliquots of the sample; contacting the array of extraction probes with the array of capillaries such that the extraction probes are positioned within the capillaries; separating the array of extraction probes from the array of capillaries, such as that the extraction probes are separated from the sample.

2. The method of claim 1 wherein each capillary tube comprises a different sample.

3. An assembly of extraction probes, each extraction probe comprising a solid support and a combinatorially derived extraction phase, wherein said extraction probes comprise a plurality of different types of extraction phases, and wherein the nature of the extraction phases are encoded by the solid supports.

4. A method for extracting a plurality of analytes from a sample, comprising the steps of:providing a plurality of extraction probes capable of adsorbing analytes, wherein each extraction probe comprises a solid support and a combinatorially-derived extraction phase, wherein said extraction probes are differentiable, wherein said extraction probes are encoded, and wherein said extraction probes are distinguished in dependence on said encoding; distinguishing between at least two different separated extraction probes; contacting said extraction probes with a sample suspected of comprising at least one of the analytes; and separating said extraction probe from the sample.

5. The method of claim 4 wherein said separated extraction probes are distinguished by an optical method.

6. The method of claim 5 wherein said separated extraction probes are distinguished by a method selected from the group consisting of absorbance, fluorescence, Raman, hyperRaman, Rayleigh scattering, hyperRayleigh scattering, CARS, sum frequency generation, degenerate four wave mixing, forward light scattering, back scattering, and angular light scattering.

7. The method of claim 4 wherein said separated extraction probes are distinguished by a method selected from the group consisting of near field scanning optical microscopy, atomic force microscopy, scanning tunneling microscopy, chemical force microscopy, lateral force microscopy, transmission electron microscopy, scanning electron microscopy, field emission scanning electron microscopy, electrical methods, mechanical methods, magnetic detection methods, and SQUID.

8. A method for extracting a plurality of analytes from a sample, comprising the steps of:providing a plurality of extraction probes capable of adsorbing analytes, wherein each extraction probe comprises a solid support and a combinatorially-derived extraction phase, and wherein at least one of said extraction phases comprises a self-assembled monolayer; contacting said extraction probes with a sample suspected of comprising at least one of the analytes; and separating said extraction probe from the sample.

9. An assembly of extraction probes, each extraction probe comprising a solid support and a combinatorially derived extraction phase, wherein said extraction probes comprise a plurality of different types of extraction phases, and wherein at least one of said extraction phases is a self-assembled monolayer.

10. A method for extracting a plurality of analytes from a sample, comprising the steps of:providing a plurality of extraction probes capable of adsorbing analytes, wherein each extraction probe comprises a solid support and a combinatorially-derived extraction phase, and wherein said extraction phases are selected from a combinatorial library; contacting said extraction probes with a sample suspected of comprising at least one of the analytes; and separating said extraction probe from the sample.

11. A method for extracting a plurality of analytes from a sample, comprising:providing a position-addressable array of extraction probes, each comprising a fiber and an extraction phase, wherein each extraction probe is capable of adsorbing an analyte; contacting said array of extraction probes with sample aliquots suspected of comprising at least one of said analytes; and separating said array of extraction probes from said sample aliquots.

12. The method of claim 11 wherein each extraction probe comprises a different extraction phase.

13. The method of claim 11 wherein each sample aliquot is different.

14. The method of claim 11 wherein each fiber has a diameter of less than 100 microns.

15. The method of claim 14 wherein each fiber has a diameter of less than 1 micron.

16. A method for extracting a plurality of analytes from a sample, comprising the steps of:providing a plurality of extraction probes capable of adsorbing analytes, wherein each extraction probe comprises a freestanding particle having a composition that varies along its length and an extraction phase; contacting said extraction probes with a sample suspected of comprising at least one of the analytes, wherein the sample and the freestanding particle are in solution and the freestanding particle is not bound to, incorporated in, or part of a macrostructure; allowing the particles and the sample to exist in solution; and separating said extraction probes from the sample, wherein said extraction probes are differentiable, and wherein the method further comprising distinguishing between at least two different separated extraction probes, and wherein said extraction probes are encoded, and wherein said separated extraction probes are distinguished in dependence on said encoding.