Patent Grant
7544518
Genomic technology has advanced to a point at which, in principle, it has become possible to determine complete genomic sequences and to quantitatively measure the mRNA levels for each gene expressed in a cell. For some species the complete genomic sequence has now been determined, and for one strain of the yeast Saccharomyces cervisiae, the mRNA levels for each expressed gene have been precisely quantified under different growth conditions (Velculescu et al., 1997). Comparative cDNA array analysis and related technologies have been used to determine induced changes in gene expression at the mRNA level by concurrently monitoring the expression level of a large number of genes (in some cases all the genes) expressed by the investigated cell or tissue (Shalon et al., 1996). Furthermore, biological and computational techniques have been used to correlate specific function with gene sequences. The interpretation of the data obtained by these techniques in the context of the structure, control and mechanism of biological systems has been recognized as a considerable challenge. In particular, it has been extremely difficult to explain the mechanism of biological processes by genomic analysis alone.
Proteins are essential for the control and execution of virtually every biological process. The rate of synthesis and the half-life of proteins and thus their expression level are also controlled post-transcriptionally. Furthermore, the activity of proteins is frequently modulated by post-translational modifications, in particular protein phosphorylation, and dependent on the association of the protein with other molecules including DNA and proteins. Neither the level of expression nor the state of activity of proteins is therefore directly apparent from the gene sequence or even the expression level of the corresponding mRNA transcript. It is therefore essential that a complete description of a biological system include measurements that indicate the identity, quantity, and the state of activity of the proteins which constitute the system. The large-scale (ultimately global) analysis of proteins expressed in a cell or tissue has been termed proteome analysis (Pennington et al., 1997).
At present no protein analytical technology approaches the throughput and level of automation of genomic technology. The most common implementation of proteome analysis is based on the separation of complex protein samples most commonly by two-dimensional gel electrophoresis (2DE) and the subsequent sequential identification of the separated protein species (Ducret et al., 1998; Garrels et al., 1997; Link et al., 1997; Shevchenko et al., 1996; Gygi et al. 1999; Boucherie et al., 1996). This approach has been revolutionized by the development of powerful mass spectrometric techniques and the development of computer algorithms which correlate protein and peptide mass spectral data with sequence databases and thus rapidly and conclusively identify proteins (Eng et al., 1994; Mann and Wilm, 1994; Yates et al., 1995). This technology has reached a level of sensitivity which now permits the identification of essentially any protein which is detectable by conventional protein staining methods including silver staining (Figeys and Aebersold, 1998; Figeys et al., 1996; Figeys et al., 1997; Shevchenko et al., 1996). However, the sequential manner in which samples are processed limits the sample throughput, the most sensitive methods have been difficult to automate and low abundance proteins, such as regulatory proteins, escape detection without prior enrichment, thus effectively limiting the dynamic range of the technique. In the 2DE/(MS)n method, proteins are quantified by densitometry of stained spots in the 2DE gels.
The development of methods and instrumentation for automated, data-dependent electrospray ionization (ESI) tandem mass spectrometry (MSn) in conjunction with microcapillary liquid chromatography (μLC) and database searching has significantly increased the sensitivity and speed of the identification of gel-separated proteins. As an alternative to the 2DE/MSn approach to proteome analysis, the direct analysis by tandem mass spectrometry of peptide mixtures generated by the digestion of complex protein mixtures has been proposed (Dongr'e et al., 1997). μLC-MS/MS has also been used successfully for the large-scale identification of individual proteins directly from mixtures without gel electrophoretic separation (Link et al., 1999; Opitek et al., 1997). While these approaches dramatically accelerate protein identification, the quantities of the analyzed proteins cannot be easily determined, and these methods have not been shown to substantially alleviate the dynamic range problem also encountered by the 2DE/MS/MS approach. Therefore, low abundance proteins in complex samples are also difficult to analyze by the μLC/MS/MS method without their prior enrichment.
It is therefore apparent that current technologies, while suitable to identify the components of protein mixtures, are neither capable of measuring the quantity nor the state of activity of the protein in a mixture. Even evolutionary improvements of the current approaches are unlikely to advance their performance sufficiently to make routine quantitative and functional proteome analysis a reality.
This invention provides methods and reagents that can be employed in proteome analysis which overcome the limitations inherent in traditional techniques. The basic approach described can be employed for the quantitative analysis of protein expression in complex samples (such as cells, tissues, and fractions thereof, the detection and quantitation of specific proteins in complex samples, and the quantitative measurement of specific enzymatic activities in complex samples.
In this regard, a multitude of analytical techniques are presently available for clinical and diagnostic assays which detect the presence, absence, deficiency or excess of a protein or protein function associable with a normal or disease state. While these techniques are quite sensitive, they do not necessarily provide chemical speciation of products and may, as a result, be difficult to use for assaying several proteins or enzymes simultaneously in a single sample. Current methods may not distinguish among aberrant expression of different enzymes or their malfunctions which lead to a common set of clinical symptoms. The methods and reagents herein can be employed in clinical and diagnostic assays for simultaneous (multiplex) monitoring of multiple proteins and protein reactions.
This invention provides analytical reagents and mass spectrometry-based methods using these reagents for the rapid, and quantitative analysis of proteins or protein function in mixtures of proteins. The analytical method can be used for qualitative and particularly for quantitative analysis of global protein expression profiles in cells and tissues, i.e. the quantitative analysis of proteomes. The method can also be employed to screen for and identify proteins whose expression level in cells, tissue or biological fluids is affected by a stimulus (e.g., administration of a drug or contact with a potentially toxic material), by a change in environment (e.g., nutrient level, temperature, passage of time) or by a change in condition or cell state (e.g., disease state, malignancy, site-directed mutation, gene knockouts) of the cell, tissue or organism from which the sample originated. The proteins identified in such a screen can function as markers for the changed state. For example, comparisons of protein expression profiles of normal and malignant cells can result in the identification of proteins whose presence or absence is characteristic and diagnostic of the malignancy.
In an exemplary embodiment, the methods herein can be employed to screen for changes in the expression or state of enzymatic activity of specific proteins. These changes may be induced by a variety of chemicals, including pharmaceutical agonists or antagonists, or potentially harmful or toxic materials. The knowledge of such changes may be useful for diagnosing enzyme-based diseases and for investigating complex regulatory networks in cells.
The methods herein can also be used to implement a variety of clinical and diagnostic analyses to detect the presence, absence, deficiency or excess of a given protein or protein function in a biological fluid (e.g., blood), or in cells or tissue. The method is particularly useful in the analysis of complex mixtures of proteins, i.e., those containing 5 or more distinct proteins or protein functions.
The inventive method employs affinity-labeled protein reactive reagents that allow for the selective isolation of peptide fragments or the products of reaction with a given protein (e.g., products of enzymatic reaction) from complex mixtures. The isolated peptide fragments or reaction products are characteristic of the presence of a protein or the presence of a protein function, e.g., an enzymatic activity, respectively, in those mixtures. Isolated peptides or reaction products are characterized by mass spectrometric (MS) techniques. In particular, the sequence of isolated peptides can be determined using tandem MS (MSn) techniques, and by application of sequence database searching techniques, the protein from which the sequenced peptide originated can be identified. The reagents also provide for differential isotopic labeling of the isolated peptides or reaction products which facilitates quantitative determination by mass spectrometry of the relative amounts of proteins in different samples. Also, the use of differentially isotopically-labeled reagents as internal standards facilitates quantitative determination of the absolute amounts of one or more proteins or reaction products present in the sample.
In general, the affinity labeled protein reactive reagents of this invention have three portions: an affinity label (A) covalently linked to a protein reactive group (PRG) through a linker group (L):
A-L-PRG
The linker may be differentially isotopically labeled, e.g., by substitution of one or more atoms in the linker with a stable isotope thereof. For example, hydrogens can be substituted with deuteriums or 12C with 13C.
The affinity label A functions as a molecular handle that selectively binds covalently or non-covalently, to a capture reagent (CR). Binding to CR facilitates isolation of peptides, substrates or reaction products tagged or labeled with A. In specific embodiments, A is a strepavidin or avidin. After affinity isolation of affinity tagged materials, some of which may be isotopically labeled, the interaction between A and the capture reagent is disrupted or broken to allow MS analysis of the isolated materials. The affinity label may be displaced from the capture reagent by addition of displacing ligand, which may be free A or a derivative of A, or by changing solvent (e.g., solvent type or pH) or temperature conditions or the linker may be cleaved chemically, enzymatically, thermally or photochemically to release the isolated materials for MS analysis.
Two types of PRG groups are specifically provided herein: (a) those groups that selectively react with a protein functional group to form a covalent or non-covalent bond tagging the protein at specific sites, and (b) those that are transformed by action of the protein, e.g., that are substrates for an enzyme. In specific embodiments, PRG is a group having specific reactivity for certain protein groups, such as specificity for sulfhydryl groups, and is useful in general for selectively tagging proteins in complex mixtures. A sulfhydryl specific reagent tags proteins containing cysteine. In other specific embodiments, PRG is an enzyme substrate that is selectively cleaved (leaving A-L) or modified (giving A-L-PRG′) by the action of an enzyme of interest.
Exemplary reagents have the general formula:
A-B1—X1—(CH2)n—[X2—(CH2)m]x—X3—(CH2)p—X4—B2-PRG
where:
One or more of the CH2 groups of the linker can be optionally substituted with small (C1-C6) alkyl, alkenyl, or alkoxy groups, an aryl group or can be substituted with functional groups that promote ionization, such as acidic or basic groups or groups carrying permanent positive or negative charge. One or more single bonds connecting CH2 groups in the linker can be replaced with a double or a triple bond. Preferred R and R′ alkyl, alkenyl, alkynyl or alkoxy groups are small having 1 to about 6 carbon atoms.
One or more of the atoms in the linker can be substituted with a stable isotope to generate one or more substantially chemically identical, but isotopically distinguishable reagents. For example, one or more hydrogens in the linker can be substituted with deuterium to generate isotopically heavy reagents.
In an exemplary embodiment the linker contains groups that can be cleaved to remove the affinity tag. If a cleavable linker group is employed, it is typically cleaved after affinity tagged peptides, substrates or reaction products have been isolated using the affinity label together with the CR. In this case, any isotopic labeling in the linker preferably remains bound to the protein, peptide, substrate, or reaction product.
Linker groups include among others: ethers, polyethers, ether diamines, polyether diamines, diamines, amides, polyamides, polythioethers, disulfides, silyl ethers, alkyl or alkenyl chains (straight chain or branched and portions of which may be cyclic), aryl, diaryl or alkyl-aryl groups. Aryl groups in linkers can contain one or more heteroatoms (e.g., N, O, or S atoms).
In one aspect, the invention provides a mass spectrometric method for identification and quantitation of one or more proteins in a complex mixture which employs affinity labeled reagents in which the PRG is a group that selectively reacts with certain groups that are typically found in peptides (e.g., sulfhydryl, amino, carboxy, homoserine lactone groups). One or more affinity labeled reagents with different PRG groups are introduced into a mixture containing proteins and the reagents react with certain proteins to tag them with the affinity label. It may be necessary to pretreat the protein mixture to reduce disulfide bonds or otherwise facilitate affinity labeling. After reaction with the affinity labeled reagents, proteins in the complex mixture are cleaved, e.g., enzymatically, into a number of peptides. This digestion step may not be necessary, if the proteins are relatively small. Peptides that remain tagged with the affinity label are isolated by an affinity isolation method, e.g., affinity chromatography, via their selective binding to the CR. Isolated peptides are released from the CR by displacement of A or cleavage of the linker, and released materials are analyzed by liquid chromatography/mass spectrometry (LC/MS). The sequence of one or more tagged peptides is then determined by MSn techniques. At least one peptide sequence derived from a protein will be characteristic of that protein and be indicative of its presence in the mixture. Thus, the sequences of the peptides typically provide sufficient information to identify one or more proteins present in a mixture.
Quantitative relative amounts of proteins in one or more different samples containing protein mixtures (e.g., biological fluids, cell or tissue lysates, etc.) can be determined using chemically identical, affinity tagged and differentially isotopically labeled reagents to affinity tag and differentially isotopically label proteins in the different samples. In this method, each sample to be compared is treated with a different isotopically labeled reagent to tag certain proteins therein with the affinity label. The treated samples are then combined, preferably in equal amounts, and the proteins in the combined sample are enzymatically digested, if necessary, to generate peptides. Some of the peptides are affinity tagged and in addition tagged peptides originating from different samples are differentially isotopically labeled. As described above, affinity labeled peptides are isolated, released from the capture reagent and analyzed by (LC/MS). Peptides characteristic of their protein origin are sequenced using MSn techniques allowing identification of proteins in the samples. The relative amounts of a given protein in each sample is determined by comparing relative abundance of the ions generated from any differentially labeled peptides originating from that protein. The method can be used to assess relative amounts of known proteins in different samples. Further, since the method does not require any prior knowledge of the type of proteins that may be present in the samples, it can be used to identify proteins which are present at different levels in the samples examined. More specifically, the method can be applied to screen for and identify proteins which exhibit differential expression in cells, tissue or biological fluids. It is also possible to determine the absolute amounts of specific proteins in a complex mixture. In this case, a known amount of internal standard, one for each specific protein in the mixture to be quantified, is added to the sample to be analyzed. The internal standard is an affinity tagged peptide that is identical in chemical structure to the affinity tagged peptide to be quantified except that the internal standard is differentially isotopically labeled, either in the peptide or in the affinity tag portion, to distinguish it from the affinity tagged peptide to be quantified. The internal standard can be provided in the sample to be analyzed in other ways. For example, a specific protein or set of proteins can be chemically tagged with an isotopically labeled affinity tagging reagent. A known amount of this material can be added to the sample to be analyzed. Alternatively, a specific protein or set of proteins may be labeled with heavy atom isotopes and then derivatized with an affinity tagging reagent.
Also, it is possible to quantify the levels of specific proteins in multiple samples in a single analysis (multiplexing). In this case, affinity tagging reagents used to derivatize proteins present in different affinity tagged peptides from different samples can be selectively quantified by mass spectrometry.
In this aspect of the invention, the method provides for quantitative measurement of specific proteins in biological fluids, cells or tissues and can be applied to determine global protein expression profiles in different cells and tissues. The same general strategy can be broadened to achieve the proteome-wide, qualitative and quantitative analysis of the state of modification of proteins, by employing affinity reagents with differing specificity for reaction with proteins. The method and reagents of this invention can be used to identify low abundance proteins in complex mixtures and can be used to selectively analyze specific groups or classes of proteins such as membrane or cell surface proteins, or proteins contained within organelles, sub-cellular fractions, or biochemical fractions such as immunoprecipitates. Further, these methods can be applied to analyze differences in expressed proteins in different cell states. For example, the methods and reagents herein can be employed in diagnostic assays for the detection of the presence or the absence of one or more proteins indicative of a disease state, such as cancer.
In a second aspect, the invention provides an MS method for detection of the presence or absence of a protein function, e.g., an enzyme activity, in a sample. The method can also be employed to detect a deficiency or excess (over normal levels) of protein function in a sample. Samples that can be analyzed include various biological fluids and materials, including tissue and cells. In this case, the PRG of the affinity labeled reagent is a substrate for the enzyme of interest. Affinity labeled substrates are provided for each enzyme of interest and are introduced into a sample where they react to generate affinity labeled products, if the enzyme of interest is present in the sample. Products or unreacted substrate that are tagged with the affinity label are isolated by an affinity isolation method, e.g., affinity chromatography, via their selective binding to the CR. The isolated tagged substrates and products are analyzed by mass spectrometry. Affinity labeled products include those in which the substrate is entirely cleaved from the linker or in which the substrate is modified by reaction with a protein of interest. Detection of the affinity-labeled product indicates the protein function is present in the sample. Detection of little or no affinity labeled product indicates deficiency or absence, respectively, of the protein function in the sample.
The amount of selected protein, e.g., measured in terms of enzyme activity, present in a sample can be measured by introducing a known amount of an internal standard which is an isotopically labeled analog of the expected product of the enzymatic reaction of the reagent substrate. The internal standard is substantially chemically identical to the expected enzymatic reaction product, but is isotopically distinguishable therefrom. The level of protein function (e.g., enzymatic activity) in a given sample can be compared with activity levels in other samples or controls (either negative or positive controls). The procedure therefore can detect the presence, absence, deficiency, or excess of a protein function in a sample. The method is capable of quantifying the velocity of an enzymatic reaction since it enables the amount of product formed over a known time period to be measured. This method can be multiplexed, by simultaneous use of a plurality of affinity labeled substrates selective for different protein functions and if quantitation is desired by inclusion of the corresponding internal standards for expected products, to analyze for a plurality of protein functions in a single sample.
The methods of this invention employ affinity tagged protein reactive reagents in which the affinity tag is covalently attached to a protein reactive group by a linker. The linker can be isotopically labeled to generate pairs or sets of reagents that are substantially chemically identical, but which are distinguishable by mass. For example a pair of reagents, one of which is isotopically heavy and the other of which is isotopically light can be employed for the comparison of two samples one of which may be a reference sample containing one or more known proteins in known amounts. For example, any one or more of the hydrogen, nitrogen, oxygen or sulfur atoms in the linker may be replaced with their isotopically stable isotopes: 2H, 13C, 15N, 17O, 18O or 34S.
Suitable affinity tags bind selectively either covalently or non-covalently and with high affinity to a capture reagent (CR). The CR-A interaction or bond should remain intact after extensive and multiple washings with a variety of solutions to remove non-specifically bound components. The affinity tag binds minimally or preferably not at all to components in the assay system, except CR, and does not significantly bind to surfaces of reaction vessels. Any non-specific interaction of the affinity tag with other components or surfaces should be disrupted by multiple washes that leave CR-A intact. Further, it must be possible to disrupt the interaction of A and CR to release peptides, substrates or reaction products, for example, by addition of a displacing ligand or by changing the temperature or solvent conditions. Preferably, neither CR nor A react chemically with other components in the assay system and both groups should be chemically stable over the time period of an assay or experiment. The affinity tag preferably does not undergo peptide-like fragmentation during (MS)n analysis. The affinity label is preferably soluble in the sample liquid to be analyzed and the CR should remain soluble in the sample liquid even though attached to an insoluble resin such as Agarose. In the case of CR term soluble means that CR is sufficiently hydrated or otherwise solvated such that it functions properly for binding to A. CR or CR-containing conjugates should not be present in the sample to be analyzed, except when added to capture A.
Examples of A and CR pairs include:
d-biotin or structurally modified biotin-based reagents, including d-iminobiotin, which bind to proteins of the avidin/streptavidin, which may, for example, be used in the forms of strepavidin-Agarose, oligomeric-avidin-Agarose, or monomeric-avidin-Agarose;
any 1,2-diol, such as 1,2-dihydroxyethane (HO—CH2—CH2—OH), and other 1,2-dihydroxyalkanes including those of cyclic alkanes, e.g., 1,2-dihydroxycyclohexane which bind to an alkyl or aryl boronic acid or boronic acid esters, such as phenyl-B (OH)2 or hexyl-B(OEthyl)2 which may be attached via the alkyl or aryl group to a solid support material, such as Agarose;
maltose which binds to maltose binding protein (as well as any other sugar/sugar binding protein pair or more generally to any ligand/ligand binding protein pairs that has properties discussed above);
a hapten, such as dinitrophenyl group, for any antibody where the hapten binds to an anti-hapten antibody that recognizes the hapten, for example the dinitrophenyl group will bind to an anti-dinitrophenyl-lgG;
a ligand which binds to a transition metal, for example, an oligomeric histidine will bind to Ni(II), the transition metal CR may be used in the form of a resin bound chelated transition metal, such as nitrilotriacetic acid-chelated Ni(II) or iminodiacetic acid-chelated Ni(II);
glutathione which binds to glutathione-S-transferase.
In general, any A-CR pair commonly used for affinity enrichment which meets the suitability criteria discussed above. Biotin and biotin-based affinity tags are preferred. Of particular interest are structurally modified biotins, such as d-iminobiotin, which will elute from avidin or strepavidin columns under solvent conditions compatible with ESI-MS analysis, such as dilute acids containing 10-20% organic solvent. It is expected that d-iminobiotin tagged compounds will elute in solvents below pH 4. d-Iminobiotin tagged protein reactive reagents can be synthesized by methods described herein for the corresponding biotin tagged reagents.
A displacement ligand, DL, is optionally used to displace A from CR. Suitable DLs are not typically present in samples unless added. DL should be chemically and enzymatically stable in the sample to be analyzed and should not react with or bind to components (other than CR) in samples or bind non-specifically to reaction vessel walls. DL preferably does not undergo peptide-like fragmentation during MS analysis, and its presence in sample should not significantly suppress the ionization of tagged peptide, substrate, or reaction product conjugates.
DL itself preferably is minimally ionized during mass spectrometric analysis and the formation of ions composed of DL clusters is preferably minimal. The selection of DL, depends upon the A and CR groups that are employed. In general, DL is selected to displace A from CR in a reasonable time scale, at most within a week of its addition, but more preferably within a few minutes or up to an hour. The affinity of DL for CR should be comparable or stronger than the affinity of the tagged compounds containing A for CR. Furthermore, DL should be soluble in the solvent used during the elution of tagged compounds containing A from CR. DL preferably is free A or a derivative or structural modification of A. Examples of DL include, d-biotin or d-biotin derivatives, particularly those containing groups that suppress cluster formation or suppress ionization in MS.
The linker group (L) should be soluble in the sample liquid to be analyzed and it should be stable with respect to chemical reaction, e.g., substantially chemically inert, with components of the sample as well as A and CR groups. The linker when bound to A should not interfere with the specific interaction of A with CR or interfere with the displacement of A from CR by a displacing ligand or by a change in temperature or solvent. The linker should bind minimally or preferably not at all to other components in the system, to reaction vessel surfaces or CR. Any non-specific interactions of the linker should be broken after multiple washes which leave the A-CR complex intact. Linkers preferably do not undergo peptide-like fragmentation during (MS)n analysis. At least some of the atoms in the linker groups should be readily replaceable with stable heavy-atom isotopes. The linker preferably contains groups or moieties that facilitate ionization of the affinity tagged reagents, peptides, substrates or reaction products.
To promote ionization, the linker may contain acidic or basic groups, e.g., COOH, SO3H, primary, secondary or tertiary amino groups, nitrogen-heterocycles, ethers, or combinations of these groups. The linker may also contain groups having a permanent charge, e.g., phosphonium groups, quaternary ammonium groups, sulfonium groups, chelated metal ions, tetralky or tetraryl borate or stable carbanions.
The covalent bond of the linker to A or PRG should typically not be unintentionally cleaved by chemical or enzymatic reactions during the assay. In some cases it may be desirable to cleave the linker from the affinity tag A or from the PRG, for example to facilitate release from an affinity column. Thus, the linker can be cleavable, for example, by chemical, thermal or photochemical reaction. Photocleavable groups in the linker may include the 1-(2-nitrophenyl)-ethyl group. Thermally labile linkers may, for example, be a double-stranded duplex formed from two complementary strands of nucleic acid, a strand of a nucleic acid with a complementary strand of a peptide nucleic acid, or two complementary peptide nucleic acid strands which will dissociate upon heating. Cleavable linkers also include those having disulfide bonds, acid or base labile groups, including among others, diarylmethyl or trimethylarylmethyl groups, silyl ethers, carbamates, oxyesters, thiesters, thionoesters, and -fluorinated amides and esters. Enzymatically cleavable linkers can contain, for example, protease-sensitive amides or esters, -lactamase-sensitive-lactam analogs and linkers that are nuclease-cleavable, or glycosidase-cleavable.
The protein reactive group (PRG) can be a group that selectively reacts with certain protein functional groups or is a substrate of an enzyme of interest. Any selectively reactive protein reactive group should react with a functional group of interest that is present in at least a portion of the proteins in a sample. Reaction of PRG with functional groups on the protein should occur under conditions that do not lead to substantial degradation of the compounds in the sample to be analyzed. Examples of selectively reactive PRGs suitable for use in the affinity tagged reagents of this invention, include those which react with sulfhydryl groups to tag proteins containing cysteine, those that react with amino groups, carboxylate groups, ester groups, phosphate reactive groups, and aldehyde and/or ketone reactive groups or, after fragmentation with CNBr, with homoserine lactone.
Thiol reactive groups include epoxides, -haloacyl group, nitriles, sulfonated alkyl or aryl thiols and maleimides. Amino reactive groups tag amino groups in proteins and include sulfonyl halides, isocyanates, isothiocyanantes, active esters, including tetrafluorophenyl esters, and N-hydroxysuccinimidyl esters, acid halides, and acid anyhydrides. In addition, amino reactive groups include aldehydes or ketones in the presence or absence of NaBH4 or NaCNBH3.
Carboxylic acid reactive groups include amines or alcohols in the presence of a coupling agent such as dicyclohexylcarbodiimide, or 2,3,5,6-tetrafluorophenyl trifluoroacetate and in the presence or absence of a coupling catalyst such as 4-dimethylaminopyridine; and transition metal-diamine complexes including Cu(II)phenanthroline
Ester reactive groups include amines which, for example, react with homoserine lactone.
Phosphate reactive groups include chelated metal where the metal is, for example Fe(III) or Ga(III), chelated to, for example, nitrilotriacetiac acid or iminodiacetic acid.
Aldehyde or ketone reactive groups include amine plus NaBH4 or NaCNBH3, or these reagents after first treating a carbohydrate with periodate to generate an aldehyde or ketone.
PRG groups can also be substrates for a selected enzyme of interest. The enzyme of interest may, for example, be one that is associated with a disease state or birth defect or one that is routinely assayed for medical purposes. Enzyme substrates of interest for use with the methods of this invention include, acid phosphatase, alkaline phosphatase, alanine aminotransferase, amylase, angiotensin converting enzyme, aspartate aminotransferase, creatine kinase, gamma-glutamyltransferase, lipase, lactate dehydrogenase, and glucose-6-phosphate dehydrogenase which are currently routinely assayed by other methods.
The requirements discussed above for A, L, PRG, extend to the corresponding segments of A-L-PRG and the reaction products generated with this reagent.
Internal standards, which are appropriately isotopically labeled, may be employed in the methods of this invention to measure absolute quantitative amounts of proteins in samples. Internal standards are of particular use in assays intended to quantitate affinity tagged products of enzymatic reactions. In this application, the internal standard is chemically identical to the tagged enzymatic product generated by the action of the enzyme on the affinity tagged enzyme substrate, but carries isotope labels which may include 2H, 13C, 15N, 17O 18O, or 34S, that allow it to be independently detected by MS techniques. Internal standards for use in the method herein to quantitate one or several proteins in a sample are prepared by reaction of affinity labeled protein reactive reagents with a known protein to generate the affinity tagged peptides generated from digestion of the tagged protein. Affinity tagged peptides internal standards are substantially chemically identical to the corresponding affinity tagged peptides generated from digestion of affinity tagged protein, except that they are differentially isotopically labeled to allow their independent detection by MS techniques.
The method of this invention can also be applied to determine the relative quantities of one or more proteins in two or more protein samples, the proteins in each sample are reacted with affinity tagging reagents which are substantially chemically identical but differentially isotopically labeled. The samples are combined and processed as one. The relative quantity of each tagged peptide which reflects the relative quantity of the protein from which the peptide originates is determined by the measurement of the respective isotope peaks by mass spectrometry.
The methods of this invention can be applied to the analysis or comparison of multiple different samples. Samples that can be analyzed by methods of this invention include cell homogenates; cell fractions; biological fluids including urine, blood, and cerebrospinal fluid; tissue homogenates; tears; feces; saliva; lavage fluids such as lung or peritoneal ravages; mixtures of biological molecules including proteins, lipids, carbohydrates and nucleic acids generated by partial or complete fractionation of cell or tissue homogenates.
The methods of this invention employ MS and (MS)n methods. While a variety of MS and (MS)n are available and may be used in these methods, Matrix Assisted Laser Desorption Ionization MS (MALDI/MS) and Electrospray Ionization MS (ESI/MS) methods are preferred.
Quantitative Proteome Analysis
This method is schematically illustrated in Scheme 1 using a biotin labeled sulfhydryl-reactive reagent for quantitative protein profile measurements in a sample protein mixture and a reference protein mixture. The method comprises the following steps:
Reduction. Disulfide bonds of proteins in the sample and reference mixtures are reduced to free SH groups. The preferred reducing agent is tri-n-butylphosphine which is used under standard conditions. Alternative reducing agents include mercaptoethylamine and dithiothreitol. If required, this reaction can be performed in the presence of solubilizing agents including high concentrations of urea and detergents to maintain protein solubility. The reference and sample protein mixtures to be compared are processed separately, applying identical reaction conditions;
Derivatization of SH groups with an affinity tag. Free SH groups are derivatized with the biotinylating reagent biotinyl-iodoacetylamidyl-4,7,10 trioxatridecanediamine the synthesis of which is described below. The reagent is prepared in different isotopically labeled forms by substitution of linker atoms with stable isotopes and each sample is derivatized with a different isotopically labeled form of the reagent. Derivatization of SH groups is preferably performed under slightly basic conditions (pH 8.5) for 90 min at RT. For the quantitative, comparative analysis of two samples, one sample each (termed reference sample and sample) are derivatized as illustrated in Scheme 1 with the isotopically light and the isotopically heavy form of the reagent, respectively. For the comparative analysis of several samples one sample is designated a reference to which the other samples are related to. Typically, the reference sample is labeled with the isotopically heavy reagent and the experimental samples are labeled with the isotopically light form of the reagent, although this choice of reagents is arbitrary. These reactions are also compatible with the presence of high concentrations of solubilizing agents;
Combination of labeled samples. After completion of the affinity tagging reaction, defined aliquots of the samples labeled with the isotopically different reagents (e.g., heavy and light reagents) are combined and all the subsequent steps are performed on the pooled samples. Combination of the differentially labeled samples at this early stage of the procedure eliminates variability due to subsequent reactions and manipulations. Preferably equal amounts of each sample are combined;
Removal of excess affinity tagged reagent. Excess reagent is adsorbed; for example, by adding an excess of SH-containing beads to the reaction mixture after protein SH groups are completely derivatized. Beads are added to the solution to achieve about a 5-fold molar excess of SH groups over the reagent added and incubated for 30 min at RT. After the reaction the beads are be removed by centrifugation;
Protein digestion. The proteins in the sample mixture are digested, typically with trypsin. Alternative proteases are also compatible with the procedure as in fact are chemical fragmentation procedures. In cases in which the preceding steps were performed in the presence of high concentrations of denaturing solubilizing agents the sample mixture is diluted until the denaturant concentration is compatible with the activity of the proteases used. This step may be omitting in the analysis of small proteins;
Affinity isolation of the affinity tagged peptides by interaction with a capture reagent. The biotinylated peptides are isolated on avidin-agarose. After digestion, the pH of the peptide samples is lowered to 6.5 and the biotinylated peptides are immobilized on beads coated with monomeric avidin (Pierce). The beads are extensively washed. The last washing solvent includes 10% methanol to remove residual SDS. Biotinylated peptides are eluted from avidin-agarose, for example, with 0.3% formic acid at pH 2;
Analysis of the isolated, derivatized peptides by μLC-MSn or CE-MSn with data dependent fragmentation. Methods and instrument control protocols well-known in the art and described, for example, in Ducret et al., 1998; Figeys and Aebersold, 1998; Figeys et al., 1996; or Haynes et al., 1998 are used.
In this last step, both the quantity and sequence identity of the proteins from which the tagged peptides originated can be determined by automated multistage MS. This is achieved by the operation of the mass spectrometer in a dual mode in which it alternates in successive scans between measuring the relative quantities of peptides eluting from the capillary column and recording the sequence information of selected peptides. Peptides are quantified by measuring in the MS mode the relative signal intensities for pairs of peptide ions of identical sequence that are tagged with the isotopically light or heavy forms of the reagent, respectively, and which therefore differ in mass by the mass differential encoded within the affinity tagged reagent. Peptide sequence information is automatically generated by selecting peptide ions of a particular mass-to-charge (m/z) ratio for collision-induced dissociation (CID) in the mass spectrometer operating in the MSn mode. (Link, A. J. et al., 1997; Gygi, S. P., et al. 1999; and Gygi, S. P. et al., 1999). The resulting CID spectra are then automatically correlated with sequence databases to identify the protein from which the sequenced peptide originated. Combination of the results generated by MS and MSn analyses of affinity tagged and differentially labeled peptide samples therefore determines the relative quantities as well as the sequence identities of the components of protein mixtures in a single, automated operation.
Results of applying this method using the biotinylated sulfhydryl reagent and to the quantitative analysis of synthetic peptide samples, to the relative quantitation of the peptides in a protein digest and the tandem mass spectral analysis of a derivatized peptide are shown in
This method can also be practiced using other affinity tags and other protein reactive groups, including amino reactive groups, carboxyl reactive groups, or groups that react with homoserine lactones.
The approach employed herein for quantitative proteome analysis is based on two principles. First, a short sequence of contiguous amino acids from a protein (5-25 residues) contains sufficient information to uniquely identify that protein. Protein identification by MSn is accomplished by correlating the sequence information contained in the CID mass spectrum with sequence databases, using sophisticated computer searching algorithms (Eng, J. et al., 1994; Mann, M. et al., 1994; Qin, J. et al., 1997; Clauser, K. R. et al., 1995). Second, pairs of identical peptides tagged with the light and heavy affinity tagged reagents, respectively, (or in analysis of more than two samples, sets of identical tagged peptides in which each set member is differentially isotopically labeled) are chemically identical and therefore serve as mutual internal standards for accurate quantitation. The MS measurement readily differentiates between peptides originating from different samples, representing for example different cell states, because of the difference between isotopically distinct reagents attached to the peptides. The ratios between the intensities of the differing weight components of these pairs or sets of peaks provide an accurate measure of the relative abundance of the peptides (and hence the proteins) in the original cell pools because the MS intensity response to a given peptide is independent of the isotopic composition of the reagents (De Leenheer, A. P. et al (1992). The use of isotopically labeled internal standards is standard practice in quantitative mass spectrometry and has been exploited to great advantage in, for example, the precise quantitation of drugs and metabolites in bodily fluids (De Leenheer, A. P. et al., 1992).
In another illustration of the method, two mixtures consisting of the same six proteins at known, but different, concentrations were prepared and analyzed. The protein mixtures were labeled, combined, and treated as schematically illustrated in Scheme 1. The isolated, tagged peptides were quantified and sequenced in a single combined μLC-MS and μLC-MSn experiment on an ESI ion trap mass spectrometer. All six proteins were unambiguously identified and accurately quantified (Table 2). Multiple tagged peptides were encountered for each protein. The differences between the observed and expected quantities for the six proteins ranged between 2 and 12%.
The process is further illustrated for a single peptide pair in
Several beneficial features of the this method are apparent. First, at least two peptides were detected from each protein in the mixture. Therefore, both quantitation and protein identification can be redundant. Second, the identified peptides all contained at least one tagged cysteinyl residue. The presence of the relatively rare cysteinyl residue in a peptide adds an additional powerful constraint for database searching (Sechi, S. et al., 1998). Third, tagging and selective enrichment of cysteine-containing peptides significantly reduced the complexity of the peptide mixture generated by the concurrent digestion of six proteins. For this protein mixture, the complexity was reduced from 293 potential tryptic peptides to 44 tryptic peptides containing at least one cysteinyl residue. Fourth, the peptide samples eluted from the avidin affinity column are directly compatible with analysis by μLC-MSn.
Quantitative Analysis of Protein Expression in Different Cell States
The protein reactive affinity reagent strategy was applied to study differences in steady-state protein expression in the yeast, S. cerevisiae, in two non-glucose repressed states (Table 3). Cells were harvested from yeast growing in log-phase utilizing either 2% galactose or 2% ethanol as the carbon source. One hundred μg of soluble yeast protein from each cell state were labeled independently with the isotopically different affinity tagged reagents. The labeled samples were combined and subjected to the strategy described in Scheme 1. One fiftieth (the equivalent of approximately 2 μg of protein from each cell state) of the sample was analyzed.
Glucose repression causes large numbers of proteins with metabolic functions significant to growth on other carbon sources to be minimally expressed (Ronne, H., 1995; Hodges, P. E. et al., 1999). Growth on galactose or ethanol with no glucose present results in the expression of glucose repressed genes. Table 3 presents a selection of 34 yeast genes encountered in the analysis, but it contains every known glucose-repressed genes that was identified (Mann, M. et al., 1994). Each of these genes would have been minimally expressed in yeast grown on glucose. Genes specific to both growth on galactose (GAL1, GAL10) as well as growth on ethanol (ADH2, ACH1) were detected and quantitated.
The quantitative nature of the method is apparent in the ability to accurately measure small changes in relative protein levels. Evidence of the accuracy of the measurements can be seen by the excellent agreement found by examining ratios for proteins for which multiple peptides were quantified. For example, the five peptides found from PCK1 had a mean ratio ±95% confidence intervals of 1.57±0.15, and the percent error was <10%. In addition, the observed changes fit the expected changes from the literature (Ronne, H., 1995; Hodges, P. E. et al., 1999). Finally, the observed changes are in agreement with the changes in-staining intensity for these same proteins examined after two-dimensional gel electrophoresis (data not shown).
The alcohol dehydrogenase family of isozymes in yeast facilitates growth on either hexose sugars (ADH1) and ethanol (ADH2). The gene ADH2 encodes an enzyme that is both glucose- and galactose-repressed and permits a yeast cell to grow entirely on ethanol by converting it into acetaldehyde which enters the TCA cycle (
The results described above illustrate that the method of this invention provides quantitative analysis of protein mixtures and the identification of the protein components therein in a single, automated operation.
The method as applied using a sulfhydryl reactive reagent significantly reduces the complexity of the peptide mixtures because affinity tagged cysteine-containing peptides are selectively isolated. For example, a theoretical tryptic digest of the entire yeast proteome (6113 proteins) produces 344,855 peptides, but only 30,619 of these peptides contain a cysteinyl residue. Thus, the complexity of the mixture is reduced, while protein quantitation and identification are still achieved. The chemical reaction of the sulfhydryl reagent with protein can be performed in the presence of urea, sodium dodecyl sulfate (SDS), salts, and other chemicals that do not contain a reactive thiol group. Therefore, proteins can be kept in solution with powerful stabilizing agents until they are enzymatically digested. The sensitivity of the μLC-MSn system is dependent of the sample quality. In particular, commonly used protein solubilizing agents are poorly compatible or incompatible with MS. Affinity purification of the tagged peptides completely eliminates contaminants incompatible with MS. The quantitation and identification of low abundance proteins by conventional methods requires large amounts (milligrams) of starting protein lysate and involves some type of enrichment for these low abundance proteins. Assays described above, start with about 100 μg of protein and used no fractionation techniques. Of this, approximately 1/50 of the protein was analyzed in a single μLC-MSn experiment. This system has a limit of detection of 10-20 fmol per peptide (Gygi, S. P. et al. (1999)). For this reason, in the assays described which employ μLC-MSn only abundant proteins are detected. However, the methods of this invention are compatible with any biochemical, immunological or cell biological fractionation methods that reduce the mixture complexity and enrich for proteins of low abundance while quantitation is maintained. This method can be redundant in both quantitation and identification if multiple cysteines are detected. There is a dynamic range associated with the ability of the method to quantitate differences in expression levels of affinity tagged peptides which is dependent on both the intensity of the peaks corresponding the peptide pair (or set) and the overall mixture complexity. In addition, this dynamic range will be different for each type of mass spectrometer used. The ion trap was employed in assays described herein because of its ability to collect impressive amounts of sequencing information (thousands of proteins can potentially be identified) in a data-dependent fashion even though it offers a more limited dynamic quantitation range. The dynamic range of the ion trap (based on signal-to-noise ratios) varied depending on the signal intensity of the peptide pair and complexity of the mixture, but differences of up to 100-fold were generally detectable and even larger differences could be determined for more abundant peptides. In addition, protein expression level changes of more than 100-200-fold still identify those proteins as major potential contributors of the phenotypic differences between the two original cell states. The method can be extended to include reactivity toward other functional groups. A small percentage of proteins (8% for S. cerevisiae) contain no cysteinyl residues and are therefore missed by analysis using reagents with sulfhydryl group specificity (i.e., thiol group specificity). Affinity tagged reagents with specificities toward functional groups other than sulfhydryl groups will also make cysteine-free proteins susceptible to analysis.
The methods of this invention can be applied to analysis of low abundance proteins and classes of proteins with particular physico-chemical properties including poor solubility, large or small size and extreme p/values.
The prototypical application of the chemistry and method is the establishment of quantitative profiles of complex protein samples and ultimately total lysates of cells and tissues following the preferred method described above. In addition, the reagents and methods of this invention have applications which go beyond the determination of protein expression profiles. Such applications include the following:
Application of amino-reactive or sulfhydryl-reactive, differentially isotopically labeled affinity tagged reagents for the quantitative analysis of proteins in immuno precipitated complexes. In the preferred version of this technique protein complexes from cells representing different states (e.g., different states of activation, different disease states, different states of differentiation) are precipitated with a specific reagent, preferably an antibody. The proteins in the precipitated complex are then derivatized and analyzed as above.
Application of amino-reactive, differentially isotopically labeled affinity tagged reagents to determine the sites of induced protein phosphorylation. In a preferred version of this method, purified proteins (e.g., immunoprecipitated from cells under different stimulatory conditions) are fragmented and derivatized as described above. Phosphopeptides are identified in the resulting peptide mixture by fragmentation in the ion source of the ESI-MS instrument and their relative abundances are determined by comparing the ion signal intensities of the experimental sample with the intensity of an included, isotopically labeled standard.
Amino-reactive, differentially isotopically labeled affinity tagged reagents are used to identify the N-terminal ion series in MSn spectra. In a preferred version of this application, the peptides to be analyzed are derivatized with a 50:50 mixture of an isotopically light and heavy reagent which is specific for amino groups. Fragmentation of the peptides by CID therefore produces two N-terminal ion series which differ in mass precisely by the mass differential of the reagent species used. This application dramatically reduces the difficulty in determining the amino acid sequence of the derivatized peptide.
Quantitative Analysis of Surface Proteins in Cells and Tissue
The cell exterior membrane and its associated proteins (cell surface proteins) participate in sensing external signals and responding to environmental cues. Changes in the abundance of cell surface proteins can reflect a specific cellular state or the ability of a cell to respond to its changing environment. Thus, the comprehensive, quantitative characterization of the protein components of the cell surface can identify marker proteins or constellations of marker proteins characteristic for a particular cellular state, or explain the molecular basis for cellular responses to external stimuli. Indeed, changes in expression of a number of cell surface receptors such as Her2/neu, erbB, IGFI receptor, and EGF receptor have been implicated in carcinogenesis and a current immunological therapeutic approach for breast cancer is based on the infusion of an antibody (Herceptin, Genentech, Palo Alto, Calif.) that specifically recognizes Her2/neu receptor.
Cell surface proteins are also experimentally accessible. Diagnostic assays for cell classification and preparative isolation of specific cells by methods such as cell sorting or panning are based on cell surface proteins. Thus, differential analysis of cell surface proteins between normal and diseased (e.g., cancer) cells can identify important diagnostic or therapeutic targets. While the importance of cell surface proteins for diagnosis and therapy of cancer has been recognized, membrane proteins have been difficult to analyze. Due to their generally poor solubility they tend to be under-represented in standard 2D gel electrophoresis patterns and attempts to adapt 2D electrophoresis conditions to the separation of membrane proteins have met limited success. The method of this invention can overcome the limitations inherent in the traditional techniques.
The analysis of membrane proteins is challenging because they generally are difficult to maintain in solution under conditions that are compatible with high sensitivity analytical instruments such as mass spectrometers. The application of the methods of the present invention to the analysis of membrane proteins is exemplified using human T cell lymphoma cell line Jurkat for membrane protein labeling and extraction and the well characterized human prostate epithelial cell line P69SV40T and two P69SV40T sublines which differ in IGF-1 receptor expression by factor of 10 to exemplify quantitative, differential analysis of membrane proteins.
Jurkat cells are an appropriate model system because the cells are easy to grow in large numbers and because the modulation of cell surface proteins in response to different stimuli and experimental conditions has been well characterized in T lymphocytes. Commercially available biotinylating reagents or more general affinity tagging reagents are employed to derivatize lysine residues and the free N-termini. Water soluble biotinylating reagents such as Sulfo-NHS (N-hydroxy succinimide) biotin and analogs (Sulfosuccinimidyl-6-(biotinamido)-hexanoate, Pierce, Rockford, Ill.) which have been used extensively for labeling cell surface proteins can be employed. The reaction of NHS esters with primary amines is best at neutral pH values and above and is compatible with the presence of organic solvent such as DMSO or DMF. Biotinylation of cell surface proteins from the Jurkat cells is carried out in PBS buffer at pH 7.2. Cells (1×107) are washed with PBS buffer to remove contaminating serum and other proteins from the culture medium. The cells are resuspended at 25×106 cell/ml and reacted with 0.5 mg/ml of Sulfo-NHS-Biotin (Pierce, Rockford, Ill.) for 30 min at RT. The labeled cells are washed twice with cold PBS to remove unreacted biotinylating reagent. Biotinylated cells are solubilized at 5×107 cells/ml in lysis buffer containing 1% Triton X-114. Triton X-114 has the property of phase-partitioning into detergent phase and aqueous phase at 30□C. Following the phase partitioning, detergent phase is removed from the aqueous phase by centrifugation at 300×g. Phase partitioning has previously been successfully used to enrich cell membrane. Also, this technique was found to enrich membrane proteins from Jurkat cell lysates. Triton phase is diluted 1:5 (v/v) using 50 mM ammonium bicarbonate buffer, pH 8.5, and high-purity, modified porcine-trypsin is added to digest the proteins at a concentration of 12.5 ng/ml for overnight at 37° C. Trypsin is neutralized by the addition of a cocktail of serine protease inhibitors and tryptic peptides are isolated by the avidin affinity chromatography techniques. Eluted peptides are separated e.g., by μLC methods and identified by searching peptide sequence databases, using for example, the Sequest program.
The human prostate epithelial cell line P69SV40T which was immortalized with SV 40 T antigen has been well characterized. This cell line is immortal but not tumorigenic and expresses type 1 insulin like growth factor receptor (IGF-1R) at 2×104 receptors per cell. A subline, called M12, was derived from P69SV40T by sequential passage in male athymic nude mice. This cell line is highly tumorigenic and metastatic and expresses 1.1×103 IGF-1R per cell. The relative difference in the abundance of IGF-1R in the cell lines P69SV40T and M12 can be quantitatively determined using methods of this invention adapted for application to membrane proteins. Since the number of IGF-1R for these cell lines has already been determined, this well characterized system can provide a reference to validate the efficiency of the quantitative methods of this invention
P69SV40T cells (1×107) are biotinylated with an isotopically heavy biotin tagged amino reactive reagent and the M12 cells (1×107) are biotinylated with a corresponding isotopically light amine reactive biotin tagged amino reactive reagent. IGF-1R is then immunoprecipitated from the combined lysate of both cell lines using an antibody against human IGF-1 R and the total mass of immunoprecipitated proteins is digested with trypsin. Trypsin is then neutralized, e.g., by the addition of inhibitors and tagged peptides are purified by biotin-avidin affinity chromatography. The eluted peptides are analyzed by LC-MS and LC-MSn for peptide quantitation and identification, respectively, as has been described above. Quantitation in this experiment is facilitated by the option to use selective ion monitoring in the MS. In this mode only the masses of tagged peptide ions expected to derive from IGF-1R need be monitored.
The described technique can be applied to compare the differences in relative abundance of cell surface proteins between parental prostate cell line (P69SV40T) and M12 cells to detect and identify those cell surface proteins whose expression level is different in the two cell lines and which may be characteristic of the different cell states. Using the methods described herein, global, relative quantitation of the cell surface proteins in any two or more cell lines can be analyzed to detect and identify those cell surface proteins characteristic of the different cell states. Results can be independently confirmed using procedure such as 1D or 2D gels, if applicable, or quantitative western blotting to confirm quantitation results.
It is expected that the experimental variability of quantitation of cell surface proteins will be considerably better than the accuracy of quantitation achieved by currently available cDNA array technology. In addition to relative protein quantity and identity, the method can also be used to reveal the orientation of the protein in the membrane, based on the presumption that intact, alive cells will exclude the biotinylating reagent.
Alternative methods can be applied to enhance the selectivity for tagged peptides derived from cell surface proteins. For example, tagged cell surface proteins can be trypsinized directly on the intact cells to generate tagged peptides, purified and analyzed as discussed. In addition, traditional cell membrane preparations may be used as an initial step to enrich cell surface proteins. These methods can include gentle cell lysis with a dounce homogenizer and series of density gradient centrifugations to isolate membrane proteins prior to proteolysis. This method can provide highly enriched preparations of cell surface proteins. Affinity tagged proteins may also be isolated by affinity chromatography prior to proteolysis as well as after proteolysis. This chromatography can be performed in the presence of surfactants such as TX-100, NP-40 or Tween-20 to maintain protein solubility. The sequential application of affinity chromatography steps (one for the intact protein and one for the tagged peptide fragments) provides a high degree of selectivity. These alternative methods are easily scalable for the detection of low abundance membrane proteins and the relative quantity of tagged peptides tagged is maintained through the selective enrichment steps.
In the application of the methods of this invention to cell surface proteins, once the tagged proteins are fragmented, the tagged peptides behave no differently from the peptides generated from more soluble samples.
Synthesis of Affinity Tagged Protein Reactive Reagents that are Selective for Certain Protein Groups
Synthetic routes of exemplary affinity tagged reagents suitable for use in the methods of this invention are provided in Schemes 2-3 where well-known synthetic techniques are employed in synthesis of the non-deuterated and deuterated reagents.
Biotinyl-iodoacetylamidyl-4,7,10 trioxatridecanediamine 4 (Scheme 3) consists of a biotin group, a chemically inert spacer of capable of being isotopically labeled with stable isotopes and an iodoacetamidyl group, respectively. The biotin group is used for affinity enrichment of peptides derivatized with the reagent, the ethylene glycol linker is differentially isotopically labeled for mass spectral analysis and the iodoacetamidyl group provides specificity of the reagent for sulfhydryl-containing peptides. The reagent can be synthesized in an all hydrogen form (isotopically light form) with and with 1-20, and preferably 4-8 deuterium atoms in the linker (isotopically heavy forms).
Analysis of Velocities of Multiple Enzymes in Cell Lysates
Monitoring enzyme functions by biochemical assays is an essential diagnostic tool that employs a multitude of analytical techniques including spectrophotometric, fluorometric, and radiometric detection of products. However, current methods are difficult to use for assaying several enzymes simultaneously in a single sample. Mass spectrometry for quantification of a collection of metabolites in biological fluids has emerged as a powerful approach for the analysis of birth defects (Morris et al., 1994), but this analytical technique has not been developed for the direct analysis of rates of individual enzymatic steps. The analytical method described herein for monitoring and quantification of enzymatic activities in cell homogenates and other biological samples permits simultaneous (multiplex) monitoring of multiple reactions, and can be readily automated.
A feature of the method of this invention as applied to enzyme assays is the use of electrospray ionization mass spectrometry (ESI-MS) (Cole et al., 1997) for the simultaneous detection of enzymatic products and chemically identical internal standards, which are distinguished by stable isotope (deuterium) labeling. A second feature is the use of affinity tagged reagents containing an enzyme substrate which when combined with affinity purification provide for facile capture of enzymatic products from crude biological fluids. The affinity tagged reagents are designed to contain a target substrate for an enzyme of interest that is covalently attached to an affinity tag via a linker. Action of the enzyme of interest on the substrate conjugate causes cleavage or other modification that changes its molecular mass (Scheme 4). The change of mass is detected by ESI-MS. The linker and affinity tag used preferably facilitate ionization by ESI, block action of other enzymes in the biological fluid, and allow highly selective capture from the complex matrix for facile purification.
An example of this approach is the design and synthesis of affinity tagged enzyme substrate reagents 1 and 2 (Scheme 5) to simultaneously assay lysosomal β-galactosidase and N-acetyl-α-D-glucosaminidase, respectively. Deficiency of the former enzyme results in one of the lysosomal storage diseases, GM1-gangliosidosis, a condition that occurs in the population with a frequency of about 1 in 50,000 and leads to early death of affected children. Deficiency of N-acetyl-R-D-glucosaminidase results in the rare lysosomal storage disorder Sanfilippo syndrome type B. This example has been described in Gerber et al. (1999) J. Amer. Chem. Soc. 121: 1102-1103 which is incorporated by reference herein in its entirety.
Conjugates 1 and 2 consist of biotin as an affinity tag, which is coupled to sarcosine. Biotin allows highly specific capture of the substrate conjugate through non-covalent binding to streptavidin immobilized on agarose beads (Bayer et al., 1990). Sarcosine provides an N-methylated amide linkage to biotin to block the enzyme biotinidase, which is often present in the cellular fluids and could cause cleavage of the conjugate molecule during the assay (Wilbur et al., 1997). In addition, it was found that biotinyl-sarcosine conjugates could be displaced from streptavidin by addition of biotin. The N-biotinylsarcosine block is linked to a polyether diamine, the length of which can be varied to avoid mass/charge overlaps of products and internal standards. The linker also allows facile introduction of multiple deuterium atoms (i.e., 8 deuteriums in 5 and 4 in 6, Scheme 5) to permit the synthesis of internal standards. The d8-linker was made by reacting DOCH2CH2OCH2CH2OD with CD2=CDCN in benzene with catalytic NaOD (Ashikaga, K., et al., 1988) and the resulting dinitrile was reduced to the diamine with Ra—Ni. The d4-linker was made in the same way using ethylene glycol and CD2=dCDCN in CH3CN and catalytic NaOH.
In addition, the linker is hydrophilic to ensure good water solubility of the substrate conjugate, and it has basic groups which are efficiently protonated by ESI and thus ensure sensitive detection by mass spectrometry. The target carbohydrate substrates are attached to the polyether linker by a β-alanine unit (Scheme 5). The enzymatic product conjugates 3 and 4 are also shown Scheme 5. Conjugates 1 and 2 were prepared as shown in Scheme 5. All reagents were purified to homogeneity by reverse-phase HPLC and characterized by high-field 1H-NMR and ESI-MS. The substrate was linked to the diamine spacer by Michael addition of the latter onto the p-acryloylamidophenyl glycoside, (Romanowska et al., 1994) and the intermediate was coupled with the tetrafluorophenyl ester of N-biotinylsarcosine (Wilbur et al., 1997).
The ESI-MS assay of -galactosidase and N-acetyl-R-D-glucosaminidase is based on enzymatic cleavage of the glycosidic bond to release monosaccharide and conjugates 3 and 4 (mass differences are 162 and 203 Da, respectively). In a typical procedure, 0.2 mM 1 and 0.3 mM 2 were incubated with sonicated cultured fibroblasts from individual patients with -galactosidase deficiency and with fibroblasts cultured from unaffected people. After incubation, labeled internal standards 5 and 6 were added, and the biotinylated components were captured on streptavidin-agarose beads. Quantitative strepavidin capture efficiency from a cell homogenate was observed with model reagents. After purification by multiple washings to remove nonspecifically bound components, the biotinylated products were released by free biotin, and the eluant was analyzed by ESI-MS. About 85% release of the biotinylated products was observed after incubation with excess biotin for 90 min. A blank was obtained by quenching the assay with all components present at time zero.
A typical procedure, cell protein (75 μg) in 15 μL of water was added to 15 μL buffer (0.1 M Na citrate, pH 4.25) containing 2 (0.3 mM) and 1 (0.2 mM, added 5 h after addition of cell protein). After incubation for 5.5 h at 37° C., the reaction was quenched by addition of 200 μL of 0.2 M glycine carbonate buffer, pH 10.3, and 5 and 6 (1 nmol each) were added. After centrifugation to remove cell debris, the supernatant was loaded onto a bed of streptavidin-agarose (7 nmol biotin binding capacity, Pierce) in a small filtration device (micro BioSpin, Bio-Rad). After 5 min, filtration was effected by centrifugation, and the gel bed was washed with 0.1% Triton X-100 (about 1 min incubation, then spin) and then six times with purified water (Milli-Q, Millipore). Elution was carried out in 25 μL of 50% methanol containing 56 nmol of free biotin (1-10 h incubation, then spin). Filtrate was diluted 4-fold with 50% methanol/water, and 1 μL was analyzed by ESI-MS.
The ESI-MS spectrum of the blank (
ESI-MS was carried out on a Finnigan LCQ ion trap instrument. Data were collected in full scan mode from m/z 625 to 875 by direct infusion at 1.5 μL/min. Specific activities were obtained from the ratio of product to internal standard ion peak areas (averaged over 30 scans).
The approach described for assaying enzymes using substrate reagents and ESI-MS can be broadly applied. The multiplex technique can be expanded to assay dozens or more enzymes simultaneously in a single reaction, obviating the need for multiple assays to assist in confirming diagnoses of rare disorders. The method can be used to measure several enzymes simultaneously when evaluating the rate of chemical flux through a specific biochemical pathway or for monitoring biochemical signaling pathways. The affinity tag-capture reagent method for isolation of affinity tagged reaction products and substrates from complex mixtures is technically simple and can be readily automated, particular when biotin-strepavidin is employed. Because of the high sensitivity of the ESI-MS detection employed, which requires only sub-microgram quantities of the substrate reagents per assay, the synthesis of several hundred substrate reagents on a low-gram scale becomes practical and economical. Since most enzyme active sites are exposed to solvent, it is possible to attach an affinity tagged linker to most enzyme substrates while preserving enzymatic activity. Scheme 6 provides the structures of several additional enzyme substrates, suitable for use in this method, indicating by arrows allowable positions for tag attachment sites. Allowable tag sites for additional enzyme substrates can be determined by review of X-ray crystal structures of enzyme-substrate or enzyme-substrate analog structures. Using a standard computer graphics program, available X-ray data and by attaching an extended chain butyl group (as a model for the affinity tagged linker) to potential tag attachment sites, suitable attachment sites that show there are no enzyme-atoms in van der Waals overlap with the model tag can be predicted.
Analogous methods to those described above can be applied to the analysis of enzymes associated with other Sanfillipo Syndromes (A, C, and D). SFA is associated with heparan sulfamidase, SFC is associated with acetyl-CoA-alpha-glucosaminide N-acetyltransferase, and SFD is associated with N-acetylglucosamine 6-sulfatase. Exemplary affinity tagged enzyme substrate reagents useful in the analysis of these enzymes and the diagnosis of these disorders is provided below. The methods can also be applied of the diagnosis of Niemann-Pick Type A and B disease by assaying for acid sphingomyelinase and to the diagnosis of Krabbe disease by assaying for galactocerebroside beta-galactosidase. These enzymes are currently assayed employing fluorphore-derivatived reagents as indicated in Scheme 7. Enzyme substrate reagents for assay of these enzymes in the methods herein can be readily prepared by replacement of the fluorophore with an A-L group herein. This approach to preparation of affinity tagged enzyme substrates is generally applicable to any known fluorophore-derivatized enzyme substrate or substrate analog.
Table 4 provides exemplary enzymes that are associated with certain birth defects or disease states. These enzymes can be assayed by the methods described herein.
Assaying Enzymatic Pathways for Carbohydrate-Deficient Glycoprotein Syndromes (CDGS)
The methods and reagents of this invention can be employed to quantify the velocities of multiple enzymes pertinent to diagnosis of CDGS diseases.
CDGS Type Ia and Ib are caused by the deficiency or absence of the enzymes phosphomannoisomerase (PMIb) (Type Ib) and phosphomannomutase (PMM2) (Type Ia) which are part of a multistep pathway (Scheme 8) for conversion of glucose to mannose-1-phosphate (Freeze, 1998). The monosaccharide substrates involved in the pathway are fructose-6-phosphate, mannose-6-phosphate, and mannose-1-phosphate. These monosaccharides can be somewhat difficult to convert to substrate conjugates because it is not a priori clear which atom on the sugar should be conjugated with the linker without impairing enzyme activity. PMIb and PMM2 can, however, be assayed indirectly. Mammalian cell microsomes contain dolichol-P-mannose synthase which catalyzes the reaction of dolichol-phosphate with GDP-mannose to form dolichol-P-mannose and GDP (Scheme 8, Chapman et al. 1980). This synthase can be assayed using the methods of this invention, specifically with a biotin-linker substrate. Microbial PMM and the enzyme which makes GDP-mannose from GTP and mannose-1-P, GDP-mannose pyrophosphorylase, are readily purified from bacteria and yeast (Glaser, 1966, Preiss, 1966), and these enzymes can be supplied exogenously to the enzyme assay. If PMIb activity is deficient, little or no mannose-6-P will be made when the reaction sequence is started by addition of fructose-6-P. Without mannose-6-P, mannose-1-P and GDP-mannose will not be made, and thus no conjugated-dolichol-P-mannose will be detected by ESI-MS. Exogenous GTP is supplied as a requirement for the GDP-mannose pyrophosphorylase step, and ATP is omitted so that mannose-6-P cannot be made from mannose. To assay PMM2, the reaction sequence is initiated with mannose-6-P, and PMM2 deficiency results in the failure to make conjugated-dolichol-P-mannose.
The carrier dolichol is a ˜60- to 105-carbon isoprenoid. Evidence is accumulating that many enzymes that operate on carbohydrates attached to dolichol chains are tolerant to significant shortening of the dolichol chain; even 10- and 15-carbon dolichols are tolerated (Rush and Wachter, 1995). It appears that such enzymes act on the water-soluble carbohydrate portion of the dolichol conjugate and thus have little or no requirement to bind the dolichol anchor. Based on this, an affinity labeled substrate for the direct assay of dolichol-P-mannose synthase and the indirect assay of PMIb and PMM2 is prepared by attaching an affinity labeled linker to the non-polar end of a short dolichol, such as the 10-carbon dolichol analog citronellol.
The synthesis of a biotinylated dolichol10-substrate conjugate containing a sarcosinyl linker (B-S-Dol10-P) is shown in Scheme 9. Protected citronellol (R=t-BuSiMe2) is regioselectively oxidized at the terminal alyllic methyl group (McMurry and Kocovsky, 1984), and the allylic alcohol is coupled with biotinylsarcosine active ester (R═CH3). The citronellol 1-hydroxy group is subsequently deprotected and phosphorylated with POCl3 (Rush and Wachter, 1995). In a parallel synthesis, d5-sarcosine, CD3NHCD2COOH, is used to prepare the isotopically labeled (heavy) reagent for use as an internal standard. d5-Sarcosine is readily prepared from commercially available materials (BrCD2COOD and CD3NH2) using standard synthetic techniques.
The deuterated internal standard, B-d5-S-Dol10-P-Mannose, is synthesized enzymatically by incubating hen oviduct microsomes with GDP-mannose and the synthetic B-d5-S-Dol10-P substrate conjugate (Rush and Waechler, 1995). An added advantage of the B—S-conjugate is that it allows for a facile affinity purification of the microsomal mannosylated product by specific capture on agarose-streptavidin beads followed by elution with free biotin.
This method employing affinity tagged short dolichol analogues is generally applicable for assaying other enzymes that operated on dolichol anchored carbohydrates. Such an approach is useful for the subsequent identification of enzyme deficiencies present in other types of CDGS that have not been yet identified.
CDGS Type II results from defective GlcNAc transferase II (GlcNAc-T II) which transfers GlcNAc from UDP-GlcNAc to the 2-position of a mannose residue in the intermediate branched oligosaccharide (the Core Region) in the process of building up the disialo-biantennary chain (Scheme 10) (Schachter, 1986, Brockhausen et al, 1989). GlcNAc transferase II is one of the six known enzymes that mediate highly regiospecific glycosylation of the mannose residues in the Core Region. The Core Region is anchored at the reducing end of chitobiosylasparagine, where the asparagine residue is part of the peptide chain of the glycosylated protein. The latter structure unit in the substrate can be replaced by a hydrophobic chain without loss of enzyme activity (Kaur et al, 1991). Thus, the substrate conjugate for CDGS Type II is assembled by linking an affinity-labeled linker group to the reducing end of chitobiosylasparagine. However, the latter structure unit in the substrate can be replaced by a hydrophobic chain without loss of enzyme activity (Kaur et al, 1991). For example, commercially available α-D-manno-pyranosylphenylisothiocyanatecan be coupled to a biotin-labeled linker and the 5,6-hydroxyls are selectively protected as illustrated in Scheme 11 (Paulsen and Meinjohanns, 1992). Coupling of the equatorial 3-OH with per-O-acetylmannosyl-1-trichloroacetamidate (Paulsen et al, 1993) will provide a disaccharide conjugate (Scheme 12 ). If a minor amount of coupling occurs at the axial 2-OH group, the products can be separated by HPLC. After deprotection, the primary 6-OH is coupled with a second equivalent of per-O-acetylmannosyl-1-trichloroacetamidate to yield the Core Region conjugate. Deprotection of the O-acetyl groups yields the substrate conjugate for GlcNAc transferase I which can be converted to the GlcNAc-T II substrate by enzymatic glycosyl transfer using a Triton X-100 rabbit liver extract, a reaction that has been carried out on a preparative scale (Kaur et al. 1981).
The synthesis of the deuterium labeled derivative needed for the internal standard is performed in parallel by using a labeled PEG-diamine building block (Gerber et al., 1999). The biotinylated trisaccharide is converted to the tetrasaccharide (product of GlcNAc-T II) by incubation with UDP-GlcNAc and transferase II (Kaur and Hindsgaul, 1991; Tan et al., 1996) and utilizing the B—S handle for affinity purification of the enzymatic products.
CDGS Type V
The lipid-linked oligosaccharide (LLO) that is transferred to the Asn residue of the glycosylated protein is composed of 2 GlcNAc, 9 mannoses, and 3 glucoses. It has recently been shown that microsomes from CDGS type V patients are greatly deficient in the enzyme that transfers one or more glucose residues during LLO biosynthesis (Korner et al, 1998). Since the transferase that attaches the carbohydrate unit of LLO to the Asn residue discriminates against the glucose-deficient LLO, CDGS Type V patients have fewer numbers of carbohydrate units attached to glycoproteins, such as transferrin (Korner et al., 1998). However, the few carbohydrate units that are present are full-length, demonstrating that residual glucosyl transfer occurs in type V CDGS patients (Korner et al., 1998). Thus, quantification of the rate of Asn glycosylation by ESI-MS would constitute a viable assay of CDGS Type V syndrome.
Synthetic peptides with 3-7 amino acid residues containing the Asn-Xaa-Ser/Thr sequence have been shown to be good substrates for glycosylation (Ronin et al., 1981). The strategy for the ESI-MS assay of the oligosaccharide transferase relies on a B-S conjugate of an appropriate peptide containing the Asn-Xaa-Ser/Thr sequence (Scheme 13). A heptapeptide, NH2-Tyr-Gln-Ser-Asn-Ser-Thr-Met-NH2 (SEQ ID NO. 1) has shown high activity in a previous study (Ronin et al, 1981). The peptide is readily available by standard peptide synthesis using an in-house automatic synthesizer. The heptapeptide and its glycoconjugates can be ionized by ESI to provide stable singly-charged ions. Coupling of BS-tetrafluorophenyl ester with NH2-Tyr-Gln-Ser-Asn-Ser-Thr-Met-NH2 (SEQ ID NO. 1) will directly yield the substrate for the transferase. Several products are expected from the enzymatic glycosylation and subsequent modifications of the oligosaccharide antenna. The products can be prepared enzymatically by incubating thyroid rough microsomes with BS-Tyr-Gln-Ser-Asn-Ser-Thr-Met-NH2 and Dol-P-Glu (Ronin et al, 1981a), followed by affinity purification of the biotinylated products. Product distribution due to different degrees of glycosylation can be monitored by ESI-MS, and the major components can be purified by HPLC. An analogous procedure using a B—N(CD3)CD2CO— conjugate is used to prepare deuterated internal standards.
The molecular masses of the ionized substrate conjugates for the set of enzymes assayed for CDGS Ia, Ib, II, and V syndromes, as well as products and internal standards are compiled in Table 5, which shows that no isobaric overlaps among the (M+H)+ species occur. The close spacing between the (M+Na)+ ion from the Type Ia,b product and the (M+H)+ ion of the demannosylated B—(N—C2D5)-2,2-D2-Gly-Dol10-P internal standard can be readily avoided by adjusting the ESI-MS conditions by addition of Na+ ions to generate the gas phase ions as Na-adducts.
All three of the targeted enzymes can be analyzed simultaneously in a single biological sample, such as a cell lysate. The PMM2 and PMIb cannot be assayed simultaneously because they require the addition of different exogenous substrates. Nevertheless, two assays using identical ESI-MS techniques can be used for diagnosing the various CDGS types instead of relying on a battery of different methods.
Clinical Enzymology Assays
A fibroblast cell pellet is thawed on ice. Sufficient 0.9% NaCl is added to give a final protein concentration in the lysate of ˜5mg/mL (typically 100 mcL), and the cell pellet is sonicated in ice water 5 times for 2 seconds each at moderate power. Total protein is determined spectrophotometrically using the BCA reagent (BCA Protein Assay kit, Pierce).
The total enzyme reaction volume is 20-30 mcL. The substrate stock solutions are maintained at concentrations of 3 mM (SFB) and 2 mM (GM1). These concentrations were measured by 1H-NMR signal integration versus an internal standard (formamide proton of DMF). Final concentration of substrates is 0.3 and 0.2 mM, respectively. A volume of reaction buffer (e.g. 200 mM sodium citrate, pH 4.5) equal to the difference of the substrate addition (2-3 mcL) plus sufficient cell sample volume to equal 50-75 mcg total protein from 20-30 mcL is added to a 0.5 mL Eppendorf tube, followed by substrate. The sample is cooled on ice, and patient cell sample is added. The reaction is initiated by incubation at 37 deg C.
For SFB: The reaction is allowed to proceed for 4.5-6 hours, after which GM1 substrate can be added or the reaction can be quenched with 100 mcL of 200 mM glycine-carbonate buffer, pH 10.5.
For GM1: The reaction is allowed to proceed for 0.5 hours. Quenching is as for SFB.
After quenching, the samples are placed on ice. Internal standards are added (1 nmol each, i.e. 50 mcL of a 0.02 mM solution). The samples are microfuged at ˜15,000 rpm for 2 min at room temperature to pellet cell debris. Streptavidin-Agarose beads (Immunopure immobilized streptavidin, Pierce) are placed in a micro bio-spin chromatography column (Bio-Rad). Sufficient beads are added to give a total biotin binding capacity of 5 nmol (typical binding capacity 100 pmol per mcL of beads as determined by Pierce). The sample supernatant is transferred to the bio-spin tube and allowed to bind for 10 minutes at room temperature. The sample is spun at ˜3,000 rpm to remove excess supernatant, then washed once with 0.01% Triton X-100 and at least five times with purified water, spinning the tube in-between to remove solution. For each wash, sufficient wash solution is added to fill the bio-spin tube.
The purified beads are then treated with 30 mcL purified water, followed by 10 mcL of a 4 mM biotin solution. The tubes are capped at the bottom to prevent leakage and allowed to incubate at 2-8□C for 2-12 hours. The samples are spun at ˜3,000 rpm to elute the sample into a clean Eppendorf tube.
The sample is then diluted with 60 mcL of 50% methanol/water and infused into the ion-trap mass spectrometer. The ESI-MS spectrum is tuned to reduce non-specific cleavage of the samples by first analyzing a blank sample (cell lysate added after reaction quench). The infused sample is analyzed by ion chromatogram integration of a 1 amu-wide window about the (M+H+)+ ions of product and internal standard.
Results are reported in nmol product formed/hour of incubation/milligrams total protein in reaction mixtures.
Clinical Analysis of Patient Samples for GM1 and SFB
Patient skin fibroblasts were obtained as frozen pellets, and stored at −20° C. until use. Two GM1 affected samples and six normal controls were analyzed.
50 mcL of 0.9% NaCl was added to each patient cell pellet. The pellets were lysed by sonication in ice water 5× for 2 seconds each at moderate sonication power, chilling the microtip in ice water in between sonications.
Samples were quantitated by BCA (Pierce) assay as follows:
Reagent A and B were mixed in 50:1 ratio as described. A protein standard curve was prepared using bovine serum albumin as a standard at concentrations of 2,1, 0.5, 0.2, and 0.05 mg/ mL. A portion of the patient sonicates were diluted 1:15 in water, and 5 mcL of each diluted patient sample and standard curve point was added to separate glass culture tubes containing 200 mcL water, in duplicate. The samples were then diluted with 1 mL of the mixed BCA reagent, vortexed to mix, and incubated at 37° C. for 60 minutes. The samples were allowed to cool to room temperature, and analyzed against a blank containing only 200 mcL water. The samples were analyzed by monitoring absorbance at 562 nm in polystyrene cuvettes. Average patient absorbance values were blank corrected and compared to standards via linear regression.
The patient protein concentrations were determined to be:
1.(Affected) 12.2 mg/mL, 2. (Normal) 10.8 mg/mL, 3. (Normal) 11.9 mg/ mL, 4. (Normal) 12.1 mg/ mL, 5. (Normal) 10.3 mg/mL, 6. (Normal) 7.79 mg/mL, 7. (Normal) 15.7 mg/mL, 8. (Affected) 11.4 mg/mL
Incubations were performed in a total of 30 mcL of total volume. The amount of reaction buffer (200 mM sodium citrate, pH 4.25) added to blank Eppendorf tubes was the difference of the substrate volume (3 mcL of each substrate stock solution, 2 mM for GM1 and 3 mM for SFB, for a total of 6 mcL) plus the volume of cell lysate required to equal 75 mcg total protein, from 30 mcL. For example, patient 1. incubation mixture initially contained 3 mcL of SFB substrate solution, 6.14 mcL patient cell lysate, and 17.86 mcL reaction buffer. The GM1 substrate was added later in the incubation (see below).
Each patient sample was analyzed in triplicate. The reaction mixtures were kept on ice during preparation, and the reaction was initiated by transfer to a 37° C. water bath. 5.5 hr into the incubation, 3 mcL GM1 substrate was added to each reaction, and after an additional 0.5 hours the reactions were placed on ice and quenched with 200 mcL of a 200 mM glycine-carbonate buffer, pH 10.25.
The purification and analysis procedures are as described in Clinical Enzymology Assay (Typical).
The resultant enzyme activities, as an average standard deviation nmol product hour incubation/mg total protein:
The following synthetic methods refer to Schemes 14-23.
Synthesis for GM1-gangliosidosis (beta-D-galactosidase deficiency)
Synthesis is based on chemistry previously described (Kataky, R. et. al., 1990), with minor modifications and an additional two steps. As an example, deviations from the established procedure as well as exact details for the additional steps are outlined below for the starting material diethylene glycol.
1,11-Dicyano-3,6,9-trioxaundecane (13) To a stirred solution of 2% (w/v) sodium hydroxide (5 mL) and diethylene glycol (5.3 g, 50 mmol) was added acrylonitrile (7.95 g, 150 mmol). The mixture was stirred at room temperature overnight and additioned with 50 mL dichloromethane. The organic layer was washed 2× with brine and dried (MgSO4). The solvent was removed by rotary evaporation. The oily residue was treated with 200 proof ethanol, and the solvent was removed by rotary evaporation. This was repeated 2× to remove excess unreacted acrylonitrile. The product was used without further purification.
Diethyl 4,7,10-trioxatridecane-1,13-dioate (14) 2 g (9.4 mmol) 13, was dissolved in 5 mL ethanol. 1 g conc. sulfuric acid was added slowly, over 5 minutes. The reaction was heated to reflux overnight. The reaction was extracted with 40 mL methylene chloride, washed once with 10 mL water and 3× with 10 mL dilute brine solution. The organic layer was dried (MgSO4) and solvent was removed to yield an oil. The final product was purified by silica chromatography (methylene chloride/ethyl acetate).
1,13-dihydroxy-4,7,10-trioxatridecane (15) Prepared exactly as described, using tetrahydrofuran as solvent. (1.7 g, 5.5 mmol 14, 50 mL distilled [CaH2] THF, 0.66 g, 16.5 mmol lithium aluminum hydride). Once addition was complete, excess LAH was quenched with ethanol, and the salts precipitated by dropwise addition of saturated sodium sulfate solution until a white precipitate formed. The solvent was removed, the precipitate washed 6×30 mL with THF and the combined organic extracts were evaporated to yield an oil. Final product is purified by silica chromatography (first with methylene chloride then with ethyl acetate and finally with acetone).
1,13-dichloro4,7,10-trioxatridecane (analog using P2) (16) 1.1 g, (4.9 mmol) 15 was added to 1.15 g (14.6 mmol) distilled pyridine in 30 mL dry benzene with stirring, followed by 1.8 g (14.6 mmol) thionyl chloride. The mixture was heated to reflux for 6 hours. After cooling in an ice bath, 5 mL 3M HCl was added with vigorous stirring. The organic layer was separated, washed 3× with a dilute brine solution, and dried (NaSO4) to yield a yellowish oil. After washing and removal of solvent, the dichloride was used without further purification.
Additional Steps:
1,13-dicyano4,7,10-trioxatridecane) (17) To a stirred solution of 0.78 g (15.5 mmol) sodium cyanide in 4 mL dimethyl sulfoxide at 80□C was added 1 g (3.9 mmol) of 16. After 2 hours, the reaction was additioned with 10 mL of saturated sodium chloride solution, 5 mL of water, and 50 mL ethyl acetate. The organic layer was washed 3× with a brine solution as before, after which the organic layer was dried (Na2SO4) and the solvents removed. The final product was purified by silica chromatography (methlyene chloride/ethyl acetate). ESI-MS: predicted, 240.1; observed, 241.1 (M+H+)+.
1,1 5-diamino-5,8, 11-trioxapentadecane (18) A stirred solution of 50 mL dry THF containing 0.42 g (10.4 mmol) fresh LAH was heated to gentle reflux under argon for 15 minutes. 0.5 g (2 mmol) 17 in 15 mL dry THF was added dropwise over 20 minutes, maintaining a gentle reflux. The unreacted LAH was quenched with ethanol, and the mixture was treated with dropwise addition of saturated sodium sulfate under efficient stirring until a white precipitate formed. The mixture was filtered, and the precipitate was washed 6×30 mL with THF. The organic extracts were combined and the solvent was removed by rotary evaporation to yield an oil. ESI-MS: predicted, 248.1; observed, 249.1 (M+H+)+.
Deuteration
Deuterium has been incorporated into the diamine linker by reduction of 14 and 17 using lithium aluminum deuteride (98% D) to achieve a d-8 deuterated diamine. No other aspects of the synthesis were changed for this procedure. These diols are used in the construction of the SFD conjugates as described later.
Clinical Substrate Synthesis for Sanfilippo Syndrome, type B (N-D-glucosaminidase deficiency).
N-(d-Biotinyl-sarcosinyl)-12-aminododecanoic acid (36). Compound 4 (32.2 mg, 0.102 mmole) was dried overnight in vacuo (with P2O5). Dry DMF (2 mL) was added and the mixture was stirred with warming to affect dissolution under nitrogen. Triethylamine (34 mcL) was added followed by 1 (20.4 mcL, 0.115 mmole) added in two 10.2 mcL portions, 5 min apart. The mixture was stirred for 1 hr at room temperature under nitrogen. 12-Aminododecanic acid (24.1 mg, 0.112 mmole, Sigma) was added in one portion, and the mixture was stirred at room temperature for 2 hr under nitrogen. CHCl3 (80 mL) was added, and the organic solution was washed with two 10 mL portions of 1 M HCl. CHCl3 was removed by rotary evaporation, and residual DMF was removed by vacuum centrifugation. The compound was dissolved in methanol and purified by HPLC (Vydac 218TP prep column). Solvent program is: 0-10 min, water with 0.06% TFA; 10-55 min, 0-100% methanol with 0.06% TFA, flow rate is 6 mL/min. Yield 31.7 mg. 1H-NMR. ESI-MS, calculated 513.4, observed 513.4 (M+H)+
N-hydroxysuccinimidyl ester of 36 (37). Compound 36 (9.8 mg, 19 mcmole) is dissolved in 100 mcL of dry DMF under nitrogen. N-hydroxysuccinimide (2.2 mg, 19 mcmole) was added followed by dicyclohexylcarbodiimide (3.9 mg, 19 mcmole). The mixture was stirred at room temperature for 60 h in the dark. Solvent was removed by vacuum centrifugation, and the residue was submitted to flash chromatography on silica gel using a gradient of CHCl3/CH3OH (15/1) to CHCl3/CH3OH (12/1). Yield 9.8 mg. 1H-NMR. ESI-MS, calculated 610.8, observed 609.7 (M+H)+.
N-(N-(d-Biotinyl-sarcosinyl)-12-aminododecanoyl)-pyschosine (38). Compound 37 (6.2 mg, 10 mcmole) and pyschosine (4.7 mg, 10 mcmole, Sigma) were dissolved in 200 mcL of dry DMF under nitrogen. Diisoproylethylamine (5 mcL) was added, and the mixture was stirred under nitrogen for 2 days in the dark. The compound was injected directly onto the HPLC column (Vydac 218TP semi-prep), and the column was developed at 2 mL/min with 0-20 min, water with 0.06% TFA, then 20-80 min, 0-100% methanol with 0.06% TFA. Yield 3.8 mg. 1H-NMR. ESI-MS, calculated 957.3, observed 956.8 (M+H)+.
N—(N-(d-Biotinyl-sarcosinyl)-12-aminododecanoyl)-sphingosylphosphorylcholine (39). Sphingosylphosphorylcholine (4.0 mg, Sigma) was mixed with 1 mL dry DMF and solvent was removed by vacuum centrifugation. This was repeated two more times. The final dried residue weighed 2.5 mg (5.4 mcmole). To this residue was added 3.3 mg of 37 (5.4 mcmole), 150 mcL of dry DMF, and 2.5 mcL of diisoproylethylamine. The mixture was stirred under nitrogen in the dark for 3 days. The compound was injected directly onto the HPLC column (Vydac 218TP semi-prep), and the column was developed at 2 mL/min with 0-20 min, water with 0.06% TFA, then 20-80 min, 0-100% methanol with 0.06% TFA. Yield 3.8 mg. 1H-NMR. ESI-MS, calculated 960.3, observed 958.7 (M+H)+.
Conjugate of d-biotin with 1,13-diamino4,7,10-trioxatridecane (40). Compound 2 was reacted with 1,13-diamino4,7,10-trioxatridecane (Fluka) essentially as described for the synthesis of 3. The product was purified by HPLC (Vydac 218TP, semi-prep) using 0-100% methanol with 0.06% TFA over 30 min at 1.5 mL/min.
Iodoacetylated 40 (41). Compound 40 was treated with 5 equivalents of iodoacetic anhydride (Aldrich) in dry DMF with stirring under nitrogen for 4 h at room temperature. The product was purified on HPLC as for 40. The structure was confirmed by ESI-MS.
Octadeuterated 41 (42). The title compound was prepared as for the 40 using 9 instead of 1,13-diamino4,7,10-trioxatridecane.
Octadeuterated 42 (43). The title compound was prepared from 42 as for 41. The structure was confirmed by ESI-MS.
Exemplary MSN Techniques and Instrumentation
An automated LC-MS/MS system for the identification of proteins by their amino acid sequence has been developed. A schematic representation is shown in
Identification of Proteins at Sub-Femtomole Sensitivity by Solid-Phase Extraction Capillary Electrophoresis Tandem Mass Spectrometry (SPE-CE-MS/MS)
Protein identification by this method is based on the same principle as described above, except that peptide separation and ionization are performed at significantly higher sensitivity.
aIsotope ratio was not analyzed because on a 4− peptide the isotope patterns were highly overlapping due to differences of only 2 amu between heavy and light ions.
†Ratios were calculated for each peptide as shown in FIG.3.
=Expected ratios were calculated from the known amounts of proteins present in each mixture.
†Protein expression ratios were calculated as described in FIG. 3.
=Carbon source for yeast growth was 2% ethanol (Eth) or 2% galactose (GAL0.
§Gene is known to be galactose- or glucose-repressed (19).
¶Eight other ribosomal proteins were detected at similar gene expression levels.
aCalculated for the GlcNAc-T II product and internal standard contain in a GlcNAc-GlcNAc-Mannose-(Mannose-GlcNAc)2 residue.
All references cited herein are incorporated by reference in their entirety herein.
This application is a continuation of U.S. patent application Ser. No. 09/839,884, filed Apr. 20, 2001, now U.S. Pat. No. 6,852,544, which is a continuation of U.S. patent application Ser. No. 09/383,062, filed Aug. 25, 1999, now U.S. Pat. No. 6,670,194, which takes priority under 35 U.S.C. §119(e) from U.S. provisional application Ser. No. 60/097,788, filed Aug. 25, 1998, and Ser. No. 60/099,113, filed Sep. 3, 1998, all of which are incorporated in their entirety by reference herein.
This invention was made through funding from the National Science Foundation Science and Technology Center for Molecular Biotechnology (grant BIR9214821) and the National Institutes of Health (NIH grants RR11823, T32HG00035, HD-02274 and GM60184). The United States government has certain rights in this invention.
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| Number | Date | Country | |
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| 20050233399 A1 | Oct 2005 | US |
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| 60097788 | Aug 1998 | US |
| Number | Date | Country | |
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| Parent | 09839884 | Apr 2001 | US |
| Child | 10994815 | US | |
| Parent | 09383062 | Aug 1999 | US |
| Child | 09839884 | US |
