Patent Application
20150268248
The instant application contains a Sequence Listing; the file, in ASCII format, is designated 3314011AWO_SequenceListing_ST25.txt and is 5.92 kilobytes in size. The file is hereby incorporated by reference in its entirety into the instant application.
The present invention relates generally to cell signaling, proteome labeling, and the differential labeling of cellular proteins in mixed cell populations.
Cell-to-cell communication, whether mediated by direct contact or through secreted factors, is fundamental to a range of biological phenomena including tissue development, homeostasis, and pathogenesis. Recent technological advances in mass spectrometry (MS) allow for the unbiased identification of hundreds to thousands of proteins in a single sample. Labeling techniques, such as SILAC and iTRAQ, further provide relative quantitation between samples. Several limitations of current methods pose significant challenges for detecting and quantifying the molecules, such as proteins, involved in cellular communication. For example, unfortunately, such quantitative methods require samples be grown and labeled in isolation, making studies of cell-to-cell cross communication difficult. Moreover, antibody-based methods for staining proteins of interest in individual cells are low-throughput and depend on prior knowledge. On the other hand, unbiased and high-throughput approaches, such as mass spectrometry (MS), cannot distinguish proteins derived from different cell-types and therefore require samples to be grown and labeled in isolation.
Protein signal transduction induced by cell-cell interactions is difficult to investigate with current research methods. Antibodies are widely used for identification and differentiation of proteins specific to different cell types in tissue or co-culture (e.g., immunostaining or FACS sorting), however antibody-based methodologies are relatively low throughput, vary in specificity, and are biased by preselection of protein readout and availability of reagents. High-throughput and unbiased methodologies, such as MS-based proteomics, may overcome these limitations. Unfortunately, as it is unable to differentiate proteins derived from different cell types in complex cell mixtures, MS is not well suited for cell-cell communication studies. A notable example of these limitations is the inability of any method to identify the cell-of-origin of growth factors, cytokines, and other secreted proteins.
In a different approach, each distinct cell type is labeled in isolation (e.g., SILAC using L-Lysine or L-Arginine isotopes), and the fully labeled cells are subsequently mixed. Peptides identified in MS/MS can then be assigned a source cell-type from the isotopic label status. Two recent reports demonstrate the feasibility of such an approach for identifying early ephrin signaling responses and determining proteins transferred between cell types. Unfortunately, these labels become diluted as cells grow in co-culture (approximately 50% with every cell division), making this experimental setup primarily useful for investigating early proteomic events. Given the caveats of each of these methodologies, in the field of cell signaling, there is a great need in the art for novel methods that overcome the limitations with current antibody and MS-based proteomics.
What is needed is a methodology that holds the potential to address a variety of questions not easily answered with current methods, including distinguishing the cell-of-origin for secreted factors in co-culture, identifying signaling pathway alterations in multicellular environments and identifying biomarkers in vivo that are relevant to disease by linking them to the cells from which they originate.
The present method, designated CTAP for Cell type specific labeling with Amino Acid Precursors, provides for the replacement of one or more essential amino acids required for cell growth, normally supplemented in the growth media, with stable isotopically labeled essential amino acid(s) that can be generated by the cell from stable isotopically labeled precursors of the essential amino acid. Transgenic expression by cells of interest of enzymes that catalyze substrate/precursor-to-amino acid reactions enables the selective and continuous labeling of those cells in culture or in vivo, for example, in a transgenic animal.
In one aspect, therefore, the invention relates to a method for labeling proteins in a vertebrate cell, the method comprising, exposing, under growth conditions, a transgenic vertebrate cell, i.e., one that has been engineered to express an exogenous enzyme that enables the cell to generate an essential amino acid from an amino acid precursor/substrate, to a composition comprising said amino acid precursor/substrate for a period of time sufficient for protein synthesis to occur. The substrate/precursor contains a stable isotope label, which is present in the resulting amino acid produced by the cell and ultimately, present in the proteome of the cell. Once labeled, recovery of labeled proteins from the cell(s), and evaluation of the proteins that contain the labeled amino acid facilitate investigations of the proteome of that cell and others in its environment.
In another aspect, the invention relates to a method for monitoring protein synthesis in a vertebrate cell, the method comprising a) exposing a transgenic vertebrate cell that expresses an exogenous enzyme, which enables the cell to generate an essential amino acid from the essential amino acid substrate/precursor to a composition comprising said amino acid substrate/precursor for a period of time sufficient for protein synthesis to occur, wherein said substrate/precursor for said essential amino acid comprises a stable-isotope label; b) isolating proteins from the cell; wherein proteins synthesized by said cell comprise the stable isotope-labeled essential amino acid.
In yet another aspect, the invention relates to a method for the differential labeling of cellular proteins in multiple cell types/populations, the method comprising, co-culturing a first transgenic vertebrate cell that expresses a first exogenous enzyme that can generate a first essential amino acid from a first amino acid precursor, and a second transgenic vertebrate cell that expresses a second exogenous enzyme that can generate a second essential amino acid from a second amino acid precursor, in a medium comprising a first essential amino acid precursor and a second essential amino acid, wherein said first and second essential amino acids differ only in mass, isolating proteins from said first and second vertebrate cells, and evaluating the proteins, wherein the protein can be attributed to the cell in which it was synthesized, based on its mass.
In a related aspect, the invention relates to method for differentiating proteins from a mixed population of vertebrate cells, the method comprising: (a) exposing (i) a first transgenic vertebrate cell that expresses an exogenous enzyme capable of converting a precursor/substrate for an essential amino acid to the essential amino acid; and (ii) a second vertebrate cell to a composition comprising said precursor/substrate for said essential amino acid, said precursor comprising a first stable isotope, and an essential amino acid comprising a second stable isotope for a period of time sufficient for protein synthesis to occur; (b) recovering proteins from said first and second vertebrate cells; (c) determining the amount of said first and second stable isotopes in said proteins to determine cell of origin, wherein a protein containing said first stable isotope was synthesized by said first transgenic vertebrate cell and a protein comprising said second stable isotope was synthesized by said second vertebrate cell. In some embodiments, the second vertebrate cell is also a transgenic cell that expresses an enzyme different from the enzyme expressed by the first vertebrate cell.
A method for determining the proteome of origin for proteins from a mixed cell culture, the method comprising: (a) exposing (i) a first transgenic vertebrate cell that expresses an exogenous enzyme capable of converting an essential amino acid substrate/precursor to an essential amino acid; and (ii) a second vertebrate cell; to a composition comprising said essential amino acid and said amino acid substrate/precursor for a period of time sufficient for protein synthesis to occur; wherein each of said essential amino acid and said essential amino acid substrate/precursor is labeled a different stable isotope; (b) recovering proteins from the co-cultured cells; c) determining the amount of each of said stable isotopes in the proteins; wherein proteins from said first transgenic vertebrate/mammalian cell exhibits a different mass than the proteins from said second vertebrate cells; and d) evaluating the proteins that comprise the labeled amino acid.
In yet another aspect, the invention relates to a method for the differential labeling of proteins in more than one cell population, the method comprising: (a) providing a first vertebrate cell population capable of synthesizing a first essential amino acid from a first amino acid precursor and a second vertebrate cell population capable of synthesizing a second essential amino acid from a second amino acid precursor; (b)
co-culturing said first and second vertebrate cell populations in a medium comprising said first and second essential amino acid precursors for a time sufficient for protein synthesis to occur; (c) recovering proteins from said cells; and (d) determining the amount of protein comprising said first essential amino acid in and the amount of protein comprising said second essential amino acid, wherein protein comprising the first essential amino acid was synthesized by said first cell population and protein comprising the second essential amino acid was synthesized by said second cell population. First and second precursors have different masses, for example, heavy and light lysine precursors so as to be distinguishable once incorporated into protein.
In yet another aspect, the invention relates to vertebrate cells that have been transiently or stably transfected to express an enzyme capable of producing a labeled essential amino acid from its labeled substrate/precursor, as well as novel cells and vectors containing nucleic acids encoding an exogenous enzyme for transfecting the cells.
In another aspect, the invention relates to vectors useful for the production of transgenic cells that express an exogenous enzyme that generates an essential amino acid from an essential amino acid substrate/precursor, and stable isotopically-labeled essential amino acid substrate/precursors.
In a related aspect, the invention relates to kits for labeling proteins and monitoring protein synthesis, the kit comprising vectors for the transfection of vertebrate cells so that the cells express an exogenous enzyme that generates an essential amino acid from an essential amino acid substrate/precursor and/or stable isotopically-labeled essential amino acid substrate/precursors.
In related aspects, the invention relates to methods for labeling proteins, monitoring protein synthesis and differentiating proteins in different cell types in mammalian cells. Mammalian cells are typically transiently or stably transfected to express an exogenous enzyme that produces an essential amino acid from a substrate/precursor molecule. By supplying a substrate/precursor for the essential amino acid that is labeled, not only can the transfected cell generate its own source of essential amino acid, proteins produced by these cells are labeled during synthesis.
All publications, patents and other references cited herein are incorporated by reference in their entirety into the present disclosure.
In practicing the present invention, many conventional techniques in molecular biology are used. Such techniques are well known and are explained in, for example, Sambrook et al., 2001, Molecular Cloning: A Laboratory Manual, Third Edition, Cold Spring Harbor Laboratory Press, Cold Spring Harbor, N.Y.; DNA Cloning: A Practical Approach, Volumes I and II, 1985 (D. N. Glover ed.); Oligonucleotide Synthesis, 1984 (M. L. Gait ed.); Nucleic Acid Hybridization, 1985, (Hames and Higgins, eds.); Transcription and Translation, 1984 (Hames and Higgins, eds.); Animal Cell Culture, 1986 (R. I. Freshney ed.); Immobilized Cells and Enzymes, 1986, (IRL Press); Perbas, 1984, A Practical Guide to Molecular Cloning; the series, Methods in Enzymology (Academic Press, Inc.); Gene Transfer Vectors for Mammalian Cells, 1987 (J. H. Miller and M. P. Calos eds., Cold Spring Harbor Laboratory); and Methods in Enzymology Vol. 154 and Vol. 155 (Wu and Grossman, and Wu, eds., respectively); Current Protocols in Molecular Biology, John Wiley & Sons, Inc. (1994), and all more current editions of these publications. The contents of these references and other references containing standard protocols, widely known to and relied upon by those of skill in the art, including manufacturers' instructions are hereby incorporated by reference as part of the present disclosure.
In the description that follows, certain conventions will be followed as regards the usage of terminology.
The term “expression” refers to the transcription and translation of a structural gene (coding sequence) so that a protein (i.e. expression product) having biological activity is synthesized. It is understood that post-translational modifications may remove portions of the polypeptide that are not essential and that glycosylation and other post-translational modifications may also occur.
The term “transfection,” as used herein, refers to the uptake, integration and expression of exogenous DNA by a host cell, and includes, without limitation, transfection with plasmids, episomes, other circular DNA forms and other vectors and transfectable forms of DNA known to those of skill in the art. The expression vector may be introduced into host cells via any one of a number of techniques known in the art including but not limited to viral infection, transformation, transfection, lipofection or other cationic lipid based transfection, calcium phosphate co-precipitation, gene gun transfection, and electroporation. These techniques are well known to persons of skill in the art.
Atoms with the same atomic number (proton number) but different mass numbers (the sum of the proton and neutron numbers) are called isotopes. Isotopes result from the presence in an atom of additional neutrons and include radioactive and stable isotopes. A “stable isotope,” therefore, refers to an isotope of an element that is not radioactive. Examples of stable isotopes include, for example, stable isotopes of carbon (e.g. 13C), hydrogen (e.g. 1H and 2H, deuterium), oxygen (e.g., 17O and 18O), nitrogen (15N) and sulfur (33S, 34S, 38S.) Stable isotopes are used in the method of the invention to impart a detectable difference in mass to the protein/proteome in which the isotope becomes incorporated.
A “stable isotope-” or “stable isotopically-labeled” amino acid or amino acid precursor, therefore, is an analog of the amino acid/precursor which incorporates a stable isotope. Examples of labeled substrate/precursors include without limitation, light (unlabeled) meso-2,6-diaminopimelate (DAP0, Sigma), light (unlabeled) D-Lysine (Sigma), medium [2H4]D-Lysine (DLYS4, C/D/N Isotopes), heavy [2H8]D-Lysine (DLYS8, C/D/N Isotopes), heavy labeled [13C6, 15N2] Z-Lysine etc.
“Relative abundance” as that term is known in mass spectrometry is a method of reporting the amount of each Mass to Charge measurement (m/z) after assigning the most abundant ion 100%. All of the other peaks are reported as a relative intensity to the largest peak.
The present method represents a technological advance in that it allows researchers to distinguish cell-types (and their proteomes) in a mixture of cells by engineering certain of the cells for continuous and specific metabolic labeling by introducing a nucleic acid encoding an amino acid-producing enzyme, thereby allowing the cell to overcome its normal auxotrophic state. Stable isotope labeling by amino acid precursors in vivo or in cell culture is a simple and straightforward approach for incorporation of a label into proteins of the transgenic cells for mass spectrometry (MS)-based quantitative proteomics.
Basically, the method relies on metabolic incorporation of a given ‘light’ or ‘heavy’ form of an amino acid into the proteins. The method relies on the incorporation into the cell's proteins of amino acids with substituted stable isotopic nuclei (e.g. deuterium, 13C, 15N etc.) that are produced by the cell from a stable isotopically-labeled amino acid precursor.
One or more cell populations that exist in the same environmental niche or which are co-cultured in vitro are exposed to different amino acid precursors that contain a different makeup of stable isotopes (e.g., light vs. heavy precursors, the end-product of which becomes 12C and 13C labeled L-lysine) so that the amino acids generated from them are distinguishable by mass spectrometry because they have different masses. When the labeled analog of an amino acid precursor is supplied to cells that have the ability to produce the amino acid from a stable isotopically-labeled precursor, the corresponding stable isotopically-labeled amino acid is incorporated into all newly synthesized proteins. After a number of cell divisions, each instance of this particular amino acid will be replaced by its isotope labeled analog.
In the presence of stable isotopically-labeled substrate/precursor for the essential amino acid, selective labeling of these cells in conjunction with modern tandem (LC-MS/MS) facilitates the differentiation, identification, and quantification of proteins derived from each cell type in a mixed population of cells.
In one aspect, the invention relates to a method for labeling proteins in a vertebrate cell, the method comprising, exposing, under conditions permitting growth/protein synthesis, a vertebrate cell that has been engineered to be able to generate an essential amino acid from its amino acid precursor/substrate, to a composition comprising said amino acid precursor/substrate for a period of time sufficient for protein synthesis to occur. The substrate/precursor contains a stable isotope label, which is present in the resulting amino acid and ultimately in proteins synthesized in the presence of the labeled amino acid. Once labeled, recovery of the proteins from the cell, and evaluation of the proteins that comprise the labeled amino acid are possible. In one embodiment, the essential amino acid is lysine and substrate/precursors therefore include diaminopimelate (DAP), D-lysine and Z-lysine. Lysine substrate/presursors contain at least one stable isotope of carbon, hydrogen, oxygen, and/or nitrogen.
In one aspect, therefore, the invention relates to a method for labeling proteins in a vertebrate cell, the method comprising, exposing, under growth conditions, a vertebrate cell that has been engineered to be able to generate an essential amino acid from its amino acid precursor/substrate, to a composition comprising said amino acid precursor/substrate for a period of time sufficient for protein synthesis to occur. The substrate/precursor contains a stable isotope label, which is present in the resulting amino acid produced by the cell and ultimately the proteome of the cell. Once labeled, recovery of labeled proteins from the cell(s), and evaluation of the proteins that contain the labeled amino acid facilitate investigations of the proteome of that cell and others in its environment.
In one embodiment, the essential amino acid is lysine and substrate/precursors therefore include without limitation diaminopimelate (DAP), D-lysine and Z-lysine. Lysine substrate/precursors contain at least one stable isotope (or no stable isotopes in the case of light label) of carbon, hydrogen, oxygen, and/or nitrogen. Any combination of stable isotopes may be present in a particular form of the essential amino acid, as long as each amino acid has a different mass and is therefore distinguishable, for example, by mass spectrometry analysis, from other forms of the same essential amino acid. Examples of labeled substrate/precursors include without limitation, light meso-2,6-diaminopimelate (DAP0, Sigma), heavy [2H8]D-Lysine (DLYS8, C/D/N Isotopes), heavy labeled [13C6, 15N2] Z-Lysine etc.
In certain embodiments, a vertebrate cell is transiently or stably transfected to express one or more enzyme components of the synthetic pathway for the essential amino acid. Enzymes may be encoded by nucleic acids from an exogenous source including bacteria, fungi, plants etc. Exemplary enzymes include without limitation, diaminopimelate carboxylase (DDC) from, for example, Arabidopsis thaliana or Escherichia coli, lysine racemase (lyr) from, for example, Proteus mirabilis and CBZcleaver, for example, from Sphingomonas paucimobilis.
The following represent examples of applications of the disclosed technology. Numerous other applications of the disclosed method are feasible.
In this work, the validity and feasibility of the CTAP method for cell-selective proteome labeling in multicellular systems is demonstrated. Using precursors of the essential amino acid L-Lysine and enzymes that catalyze its synthesis, this disclosure shows that canonical amino acids can be isotopically labeled in specific cell types in co-culture. Cell types of both mouse and human origin successfully overcome L-Lysine auxotrophy in the presence of specific enzyme-precursor pairs involved in the production of L-Lysine. The work demonstrates that there are limited molecular and phenotypic consequences of culturing DDC or lyr-expressing cells in L-Lysine free conditions on DAP or D-Lysine, respectively. Mass spectrometry analysis of enzyme-expressing cells in monoculture shows complete molecular labeling by L-Lysine derived from precursor. Differential-labeling of individual cell types in co-culture can be achieved using a dual-enzyme-precursor pair setup in the absence of L-Lysine, allowing all identified proteins to be assigned relative-quantitated values in each cell type. Supporting these data, it was also found that CTAP is applicable for labeling a specific cell-type of interest in a mixed cell culture system using only one enzyme-precursor pair, although titrating down the amount on L-Lysine in the media is necessary (for example). Finally, analyzing the supernatant of cells in co-culture, cell-of-origin of secreted proteins can be readily established.
There are several features of the CTAP system that collectively distinguish it from other cell-selective protein labeling approaches. First, the products of enzymatic catalysis are canonical amino acids allowing mature proteins to maintain their normal structure and avoiding possible functional alterations when using amino acid analogs. Second, CTAP allows individual cell populations to be continuously labeled as they are grown and passaged over extended periods of time. Third, the genetic requirement of enzyme activity to overcome essential amino acid auxotrophy makes labeling controllable by limiting transgenic expression. Fourth, utilizing multiple enzyme-precursor pairs permits differential labeling of multiple distinct cell types simultaneously. Fifth, CTAP can distinguish proteins from different cell types of the same organism rather than relying on artificial inter-species experimental setups. Finally, CTAP makes use of the same previously developed data-analysis workflows as the widely used SILAC method. To the best of our knowledge, CTAP is the only method in which the proteome of specific cell populations can be labeled continuously and differentially by canonical amino acids in a complex mixture of cells.
CTAP can be quickly adaptable across many cell types without phenotypic or molecular disturbance. Cell lines that are suitable for use in practicing the method of the invention include, but are not limited to mouse fibroblast 3T3 cells, mouse melanoma, B16 cells, human embryonic kidney (HEK) cells, human mammary adenocarcinoma cells, MDA-MB-321, etc.
Focusing on the DDC/DAP and lyr/D-Lysine enzyme-precursor pairs, the results indicate that cells behave similarly when cultured on their specific precursor relative to L-Lysine, however, these similarities were measured after a period of growth that varied in length depending on the cell type tested.
The principle of proteome labeling by amino acids produced from stable isotope-labeled precursors was demonstrated in mono-culture. Although this labeling was complete for both precursor-enzyme pairs (approximately 95%,
It is anticipated that CTAP will be an important tool for gaining insight into intercellular signaling in fundamental processes of but not limited to organogenesis, maintenance, and disease development. For example, in various cancers the interaction between malignant cells and the surrounding stromal tissue has been shown to be important for disease progression, maintenance, and altered drug efficacy (19-21). How stromal cells affect these processes is unclear, partly due to inadequate techniques for assaying their roles. The use of CTAP may address these limitations and offer an opportunity to understand the molecular mechanisms by which surrounding stroma alter tumor growth and response to treatment. Once precursor delivery, tolerance, and enzyme expression are optimized, another possible application of CTAP will be identification of disease biomarkers in vivo. Current approaches for biomarker identification are limited by their inability to classify whether a potential marker originates from the diseased tissue itself or from normal tissue. Using the described technique we can circumvent these limitations, as proteins from specific cell types of interest can in principle be labeled continuously in vivo. Any labeled protein identified in the serum or proximal fluids will originate from the cell type of interest.
Utilization of exogenous amino acid biosynthesis components allows for continuous cell-selective metabolic labeling of proteins. Furthermore, the principle behind CTAP can be applied to essential amino acids other than L-Lysine. CTAP therefore, represents a significant step forward in the field of proteomics, allowing unbiased and high-throughput MS/MS to differentiate peptides derived from distinct cells in complex cellular mixtures. The method is a powerful tool which will allow researchers to probe a variety of questions regarding cell-cell communication and cell-specific origin of biomarkers not easily accessible with other methodologies.
Investigating protein signal transduction induced by secreted factors and cell-cell interactions is limited by current research methods. A notable example of these limitations is the inability of any current method to identify the cell-of-origin of growth factors, cytokines, and other secreted proteins. Antibodies are widely used for identification and differentiation of proteins specific to different cell types in tissue or co-culture (e.g., immunostaining or fluorescence-activated cell sorting, FACS), however antibody-based methods are relatively low throughput, vary in specificity, and are biased by preselection of protein readout and availability of reagents. High-throughput and unbiased methods, such as quantitative mass spectrometry (MS) based proteomics (1-3), might overcome some of these limitations. However, as MS is unable to differentiate from which cell-type proteins originate in complex cell mixtures, it is not well suited for cell-cell communication studies. Research in cell-cell communication would greatly benefit from methods that overcome the complimentary limitations with current antibody and MS-based proteomics.
Several recent efforts have been made to differentiate the proteome of distinct cell types in co-culture. In one such approach each distinct cell type is labeled in isolation (e.g., using heavy stable isotope-labeled L-Lysine or L-Arginine), and the fully labeled cells are subsequently mixed. Peptides identified in liquid chromatography tandem mass spectrometry (LC-MS/MS) can then be assigned a source cell-type from the isotopic label status. Two recent reports demonstrate the feasibility of such an approach for identifying early ephrin signaling responses (4) and determining proteins transferred between cell types (5). Unfortunately, these labels become rapidly diluted as cells grow and divide in co-culture, making this experimental setup primarily useful for investigating early proteomic events. In a different approach, protein sequence differences between species are used to determine cell-of-origin in cross-species co-cultures and xenografts (6; 7). Although this approach has the ability to distinguish proteins between cell types, the major drawbacks are that only a subset of peptides can be differentiated, established same-species co-culture models cannot be used, and the findings from mixed-species models may not be physiologically relevant. Yet another technique utilizes tRNA-synthetases that specifically recognize and incorporate amino acid analogs into proteins (8; 9). Using certain tRNA-synthetase/amino-acid-analog pairs, this method provides for both proteomic incorporation that is specific to transgenic cells as well as the ability to perform affinity enrichment on chemical moieties (e.g., azides). However, structural differences between the analogs and canonical amino acids might cause unpredictable functional alterations in mature proteins (10). Given the caveats of each of these methods, a novel method for continuous cell-specific labeling with canonical amino acids would be valuable.
The present invention provides a method for cell-selective proteomic labeling that overcomes the problems of throughput and specificity of antibody-based cell staining, possible functional perturbations induced by amino acid analogs, physiological relevance of cross-species models, and the requirement of short co-culture time frames for cells labeled in isolation. This technique allows the proteome of distinct cell-types growing together either in vivo or in co-culture to be differentially labeled by canonical amino acids, which leads to naturally folded proteins and avoids the use of amino acid analogs. Our method utilizes the inability of vertebrate cells to synthesize certain amino acids required for growth and homeostasis. These “essential” amino acids are produced in some plants, bacteria, and lower eukaryotes, and must be supplemented to the media of vertebrate cultured cells or obtained in the diet of animals (11). Using transgenic expression of enzymes that synthesize essential amino acids, vertebrate cells are able to overcome auxotrophy by producing their own amino acids from supplemented precursors. These precursors can be isotopically-labeled, allowing cell-of-origin of proteins to be determined by label status identified by LC-MS/MS. For these studies we focus on L-Lysine, as the biosynthesis of this essential amino acid is well studied and it is commonly used in quantitative proteomic methods such as stable isotope labeling by amino acids in cell culture (SILAC) (2). In this work, we test the validity and feasibility of the CTAP method and demonstrate its viability for continuous and differential metabolic labeling of cells in co-culture. Using this novel method, we are able to determine relative protein expression between two cell types in co-culture and identify cell-of-origin of secreted proteins.
The invention, having been generally described, may be more readily understood by reference to the following examples, which are included merely for purposes of illustration of certain aspects and embodiments of the present invention, and are not intended to limit the invention in any way.
By engineering vertebrate cells to produce their own supply of L-Lysine from labeled precursors, it is possible to achieve differential proteomic tagging of specific cell types in co-culture (
To investigate the candidate precursors and eliminate those that autonomously rescue L-Lysine auxotrophy, growth rates in SILAC media supplemented with L-Lysine, various precursors, or in L-Lysine-free conditions were examined. With the exception of N2-acetyl-L-Lysine, the tested precursors alone had little or no effect on growth in wild-type cells (FIG. S2a-e).
Next, whether transgenic expression of L-Lysine biosynthesis enzymes would allow cells to acquire the ability to grow on precursors was investigated. Genes encoding the enzymes diaminopimelate decarboxylase (DDC) from Arabidopsis thaliana and Lysine racemase (lyr) from Proteus mirabilis were stably expressed in several cell lines (Table 1).
(P. mirabilis)
(A. thaliana)
(A. thaliana)
Abbreviations used in Table 1 are as follows: lyr=Lysine Racemase, DDC=diaminopimelate decarboxylase, DAP=meso-2,6-diaminopimelate. Da=Dalton. * indicates the heavy form of the precursor (deuterated, 3,3,4,4,5,5,6,6-d8).
Additionally, 3T3 and HEK293 cells were produced that express CBZcleaver and truncated lyr, respectively. Other transgenic cells generated that successfully overcome lysine auxotrophy include B16 expressing either DDC or truncated lyr, and MDA-MB-231 cells that express DDC or truncated lyr.
The gene for diaminopimelate decarboxylase (DDC) from Arabidopsis thaliana was cloned directly from A. thaliana cDNA using the primers in Table 2. The oligonucleotide sequence for DDC is given in SEQ ID NO: 10. For more information about DDC in A. thaliana, see AT3G14390 at the Arabidopsis Information Resource (TAIR).
Three PCR reactions were performed to generate pLM-GFP-P2A-DDC for insert into pLM using the Agel and Sall restriction enzymes. In the first reaction, a GFP-P2A oligonucleotide fusion that began with an Agel site was created. The second reaction generated a PCR fragment of P2A-DDC flanked by Sall. Finally, an overlapping PCR reaction created Agel-GFP-P2A-DDC-Sall. This sequence was then ligated into the Agel-Sall digested pLM vector.
GCC
ctcgagATGGCGGCAGCTACTCAAT
CGCgaattcGTTCATAGACCTTCAAAGAAACGC
ATCgaattcATGGCGGCAGCTACTCAAT
Results from initial attempts to produce a cell expressing lysine racemase (lyr) from P. mirabilis suggested that the enzyme was being secreted by the transfected cell. In view of a determination by SignalP (data not shown) that lyr contained a signal peptide, constructs for truncated forms of the enzyme, including T18 (N-terminal 18 amino acids removed) and T12 (N-terminal 12 amino acids removed) were designed.
The oligonucleotide sequence for truncated lyr from Proteus mirabilis, as synthesized for use in some embodiments of the present method, is given in SEQ ID NO: 11. The amino acid sequence of T18 with a His-tag is given in SEQ ID NO: 14.
Three PCR reactions were performed to generate pLM-GFP-P2A-lyr for insert into pLM using the Agel and Sall restriction enzymes. Primers used are shown in Table 3. In the first reaction, a mCherry-P2A oligonucleotide fusion that began with an Agel site was created. The second reaction generated a PCR fragment of P2A-lyr flanked by Sall. Finally, an overlapping PCR reaction created Agel-mCherry-P2A-lyr-Sall. This sequence was then ligated into the Agel-Sall digested pLM vector.
DDC-expressing mouse 3T3 and HEK293T cells, along with lyr-expressing human MDA-MB-231 cells, exhibited growth rates in media supplemented with the precursors meso-2,6-diaminopimelate (DAP) and D-Lysine, respectively, comparable to those in media containing L-Lysine (
Cell-Selective Incorporation of L-Lysine Produced from Precursors
Although the phenotypic data served as a proxy for L-Lysine availability, they did not directly show molecular precursor-based incorporation. To investigate whether L-Lysine is directly produced by enzymatic-turnover of the supplemented precursors, we applied the SILAC principle of exchanging the isotopic label status of amino acids from one form to another (e.g., light L-Lysine to heavy L-Lysine) (2). At the beginning of the experiments, DDC-expressing 3T3 cells were labeled with heavy [13C6, 15N2]L-Lysine (H) and lyr-expressing MDA-MB-231 cells were labeled with light L-Lysine (L). These cells were then grown in monoculture for 13 days (3 passages) in L-Lysine-free that contained unlabeled DAP (L), heavy-labeled [2H8]D-Lysine (H), or both precursors. Protein from cell lysate was trypsin-digested, submitted to high resolution LC-MS/MS, and H/L ratio for each peptide was determined by MaxQuant (14).
In the presence of light-labeled DAP alone, peptides identified in DDC-expressing 3T3 cells switched from being predominantly labeled heavy (95%, median peptide) to light (97%) (
Next, whether cells behave similarly when grown on precursors compared to L-Lysine was investigated. Cells were cultured for 3 days in media containing L-Lysine, precursor, or neither (starved, positive control for perturbed state) and mRNA expression levels were profiled using microarrays (
After demonstrating the principle of precursor-based L-Lysine production and incorporation in mono-culture, the next step was to test whether the same cells could be differentially labeled in co-culture with each population utilizing a distinct enzyme-precursor pair. To assess the specificity of labeling, we took advantage of species-specific sequence differences to compare label status between the enzyme-expressing mouse 3T3 and human MDA-MB-231 cell lines. Labeling each cell type in isolation, the 3T3 cells were initially cultured in heavy L-Lysine (H) and the MDA-MB-231 cells in light L-Lysine (L). A sample was harvested and combined 1:1 to verify the ability to differentiate label status based on species-specific peptide classification. As expected, labels of mouse-specific and human-specific peptides at the start of the experiment were confirmed to be primarily heavy and light, respectively (
With the expectation that each cell type would exchange label status, the pre-labeled cells were then combined in co-culture into media containing both DAP (L) and D-Lysine (H). After three passages, with near equal growth rates of each cell population (Figure S9), the two cell types switched labels (
Having validated continuous and differential labeling of human-mouse cells in co-culture, the next step was to determine whether the CTAP method could differentiate the proteome of a same-species co-culture system. DDC-expressing GFP+ HEK293T cells were plated together with lyr-expressing mCherry+ MDA-MB-231 cells. After five days of growth in DAP (L) and D-Lysine (H), a co-culture sample was sorted for mCherry and GFP+ cells by FACS (Supplementary FIG. S11) and each of the sorted populations was separately subjected to LC-MS/MS. Analysis of protein from the GFP+ and mCherry+ cells showed similar labeling efficiency to that seen in the human-mouse co-culture, with each cell population exhibiting distinct H/L ratios (
To test the unique potential of the CTAP method to discriminate the cell-of-origin of secreted factors, supernatant was collected from the same human-mouse co-culture setup as the previous section. Prior to harvesting, the cells were grown for 24 hours in serum-free media to avoid overloading the sample with serum proteins. Secreted proteins were concentrated by ultracentrifugation, precipitated by methanol-chloroform, and subjected to LC-MS/MS. Focusing on proteins identified only by species-specific peptides, nearly all species-specific proteins could be completely distinguished by label alone (
Applying a similar approach for analyzing secreted factors in a same-species co-culture, supernatant was collected and subjected to LC-MS/MS from the same co-cultured DDC-expressing HEK293T and lyr-expressing MDA-MB-231 cells as previously used. Quantitative analysis of the H/L ratios of 245 identified proteins spanned a similar range as those detected intracellularly with the tails of the distribution representing proteins primarily expressed in one cell type (
The following are representative applications of the CTAP methodology disclosed herein.
Microenvironment-mediated drug resistance is understudied and likely plays an important role in the failure of many therapies. For example, studies have implicated bone marrow cells as playing an important role in multiple myeloma resistance to the glucocorticoid, dexamethasone. Response to other drugs, such as DNA intercalating agent doxorubicin, have been less clear, showing enhanced effects in certain tumor-stromal contexts and are attenuated effects in others.
In vitro co-culture models of stroma-tumor interactions have been developed for cancer drug screening, however, these models are largely limited to phenotypic end-points such as cell growth or death. The molecular mechanisms of the cell-cell interactions are under-appreciated, partly due to the fact that current methods are unable to discriminate proteins originating from the different cell types. The CTAP method can facilitate the development of targeted therapies directed at malignant tumor-stroma interactions as well as help understand the mechanisms leading to stromal mediated drug resistance or sensitivity.
The CTAP methodology is also applicable in vivo as the enzyme can be expressed in a tissue or cell-specific manner in genetically modified animals. A particular cell type of interest is engineered to express the enzyme using cell-specific promoters, and a labeled precursor is administered to the animal, leading to selective labeling of the transgenic cells. Labeled proteins secreted from these cells can be detected in proximal fluids or in the serum and thus serve as unambiguous cell-specific biomarkers. By focusing on proteins that come from diseased tissue, it is more likely that markers that are indicative of disease development, maintenance, or outcome can be found. Current biomarker discovery techniques, which rely solely on statistical methods to prioritize proteins important for diagnosis or prognosis do not have this advantage as they are unable to determine from what cell-type the biomarker originates.
The following methodology is used in practicing the disclosed invention.
The L-Lysine producing enzymes used in this study were DDC, lyr, and CBZcleaver. DDC was directly amplified by PCR from Arabidopsis Thaliana cDNA (TAIR id=AT3G14390, primer sequences shown in Table S1). The lyr and CBZcleaver constructs were synthesized by GeneArt with the amino acid sequence specified by Kuan et al. [22] and Naduri et al. [23] respectively, with nucleotide sequences optimized for expression in mouse. Sequences were verified for all plasmids by the Sanger method of sequencing.
Two MSCV based retroviral vector backbones, one expressing GFP (pMIG) and the other mCherry (pMIC), were used to infect mouse cells. For insert into pMIG, the PCR product of DDC was cloned into the EcoRI site of the vector. CBZcleaver was directly subcloned from the GeneArt supplied vector pMA-RQ into pMIC using EcoRI and Xhol restriction sites. Viral supernatants for pMIG and pMIC were produced by transfecting Phoenix cells with each plasmid and the supernatant was used to infect 3T3 cells 48 hours later as previously described [24; 25].
The lentiviral backbone pLM was used to infect human cells. Overlapping PCR was performed to generate eGFP-DDC and mCherry-lyr constructs that were linked by a P2A peptide preceded by a Gly-Ser-Gly linker [26]. The pLM-P2A-enzyme virus was packaged by calcium phosphate transfection of the HEK293T packaging cell line using 10 μg of transfer vector, 6.5 μg of CMV6R8.74, and 3.5 μg of the VSV.G plasmid. MDA-MB-231 and HEK293T cells were then infected with lentiviral supernatant produced from the pLM construct 48 hours post-transfection of the packaging line.
Cell lines were grown in Dulbecco's modified Eagle's medium (DMEM) without L-Lysine and L-Arginine (SILAC-DMEM, Thermo Fisher Scientific) supplemented with 10% dialyzed FBS, antibiotics, and L-glutamine. For mono-culture growth assays, 1 mM L-Arginine was added to the media and cells were seeded in 200 μL in 96-well plates with 4000 or 5000 cells per well in different concentrations of L-Lysine, meso-2,6-diaminopimelate (DAP, Sigma, 33240), D-Lysine HCL (Sigma, L5876), N-α-Cbz-L-Lysine (Z-Lysine, BaChem, C-2200), or N2-acetyl-L-Lysine (N2A, Sigma, A2010). Cell viability was measured using either the metabolic-activity based Resazurin (Sigma) reagent or the impedance-based xCELLigence system (Roche). For Resazurin experiments, 25 μL of the Resazurin reagent was added to each well and cellular growth was estimated after two to three hours of incubation at 37° C. as described by the manufacturer. For xCELLigence experiments, cells were plated in either 16 or 96-well E-plates, allowed to settle for 30 minutes at room temperature, and then placed in the RTCA DP or RTCA MP analyzer where impedance was measured every 15 minutes for 96-120 hours. At least three replicates were performed for each condition.
Measuring the percentage of mCherry+ and GFP+ cells in co-culture was performed by either flow cytometry (BD LSR II) or Tali image-based cytometry (Invitrogen). For flow cytometric assays, 25,000 cells from each cell line were seeded together in 6-well plates in 3-4 mL media supplemented with different concentrations of L-Lysine and/or L-Lysine precursors. After 72 hours, cells were trypsinized, washed, and resuspended in 200 μL PBS containing 2% dialysed FBS and 0.1% NaN3. 20 μL was used for estimating total cell numbers using the ViaCount assay (Guava Technologies/Millipore) as described by the manufacturer. The remaining 180 μL was mixed with an equal volume of 2% paraformaldehyde. The percentage of GFP+ and mCherry+ cells in each sample was analyzed by flow cytometry. At least two replicates were performed for each condition. For Tali assays, cells were trypsinized, resuspended in media, 25 μL of co-culture cell suspension was used to determine the percentage of GFP+ and RFP+ cells in biological triplicate.
mRNA Microarray Expression Profiling
Cells were seeded at equal densities into SILAC media containing 798 μM K0, 798 μM K4, 4 mM D-Lysine HCl, or 10 mM DAP. After 72 hours, cells were washed, trypsinized, pelleted, and frozen at −80° C. RNA was extracted using the RNeasy mini kit (Qiagen), labeled, and hybridized to Illumina mouseref-8 or Human HT-12 microarrays. After median centering the probe intensities for each array, moderated t-statistics and false discovery rate calculations for multiple hypothesis correction were performed using the eBayes method provided in LIMMA (27; 28).
For exchange-of-label experiments (all monocultures, all human-mouse co-cultures, and Supplementary FIG. S4), cells were first metabolically labeled by growth for at least 10 cellular doublings in SILAC DMEM containing 798 μM light L-Lysine (K0), medium [2H4]L-Lysine (K4), or heavy [13C6, 15N2]L-Lysine (K8) (Cambridge Isotopes). Cells were then seeded in mono- or co-culture with 10 mM light meso-2,6-diaminopimelate (DAP0, Sigma), 2.5 mM or 4 mM heavy [2H8]D-Lysine (DLYS8, C/D/N Isotopes), 2.5 mM heavy labeled [13C6, 15N2]Z-Lysine (Z8, Figure S4), or both DAP0 and DLYS8. For experiments that maintained label (all human-human co-cultures), cells were initially grown for at least 10 cellular doublings in their respective precursors (DDC-expressing in DAP0, lyr-expressing in DLYS8). Populations were then combined in 10 mM DAP and 3 mM DLYS8 and grown together for 5 days in co-culture (approximately 4 cellular doublings). All cell lines were passaged 1:10-1:15 at 95% confluence.
For cultured media samples, cells were washed three times with PBS and supplied with serum-free SILAC DMEM 24 hours prior to supernatant sample collection. Media was collected, filtered with a 0.22 μm filter, and proteins were concentrated to around 1 mg/mL using a 3 KDa Amicon Ultra Centrifuge filter (Millipore) as described by the manufacturer. For harvesting of cell lysate, cells were trypsinized, resuspended in SILAC DMEM, washed three times in ice cold PBS, and cell pellets frozen at −80° C. For FACS samples, co-cultures of GFP+ and mCherry+ cells were trypsinized, washed, and resuspended in PBS with 20% media (2% FBS) to a concentration of approximately 2×107 cells/mL. Cells were then sorted into single GFP+ and mCherry+ populations on a MoFlo cell sorter (Dako), washed twice with ice cold PBS, and cell pellets were stored at −80° C. for further analysis.
Cell pellets were resuspended with Denaturation buffer (6 M Urea/2 M thio Urea in 10 mM Tris), 1 μL of benzonase was added, followed by incubation for 10 minutes at room temperature. Cellular debris was removed by centrifugation at 4000 g for 30 min. For the supernatant samples, the secreted proteins were precipitated by chloroform/methanol extraction. Protein concentration was assessed by the Bradford assay (Bio-Rad). Crude protein extracts were subjected to either GelC or in-solution digest. For the GeLC-MS analysis, protein extracts were cleaned on a 10 cm, 4-12% gradient SDS-PAGE gel (Novex). The resulting lane was cut from the gel and subjected to in-gel digestion with trypsin as described previously (29). Upon gel extraction, peptides were cleaned using Stage-tips and analyzed by nano-LC-MS. For in-solution digestion, proteins from the crude extract were reduced with 1 mM dithiothreitol (DTT), alkylated with 5 mM iodoacetamide, predigested with the endoproteinase Lys-C (Wako) for 3 h, and further digested with trypsin overnight (30). The resulting peptide mixture was cleaned using Stage-tips (31) and subjected to nano-LC-MS without prior peptide separation.
All samples were analyzed by online nanoflow liquid chromatography tandem mass spectrometry (LC-MS/MS) as previously described (32) with a few modifications. Briefly, nanoLC-MS/MSexperiments were performed on an EASY-nLC™ system (Proxeon Biosystems, Odense, Denmark) connected to an LTQ-Orbitrap XL or LTQ-Orbitrap Elite (Thermo Scientific, Bremen, Germany) through a nanoelectrospray ion source. Peptides were auto-sampled directly onto the 15 cm long 75 mm-inner diameter analytical column packed with reversed-phase C18 Reprosil AQUA-Pur 3 mm particles at a flow rate of 500 nl/min. The flow rate was reduced to 250 nl/min after loading, and the peptides were separated with a linear gradient of acetonitrile from 545% in 0.5% acetic acid for either 100, 150, or 240 minutes. Eluted peptides from the column were directly electrosprayed into the mass spectrometer. For the LTQ-Orbitrap XL analyses, the machine was operated in positive ion mode, with the following acquisition cycle: a full scan recorded in the orbitrap analyzer at resolution R 120,000 was followed by MS/MS (CID) of the top 10 most intense peptide ions in the LTQ analyzer. The total acquisition gradient was either 150 or 240 minutes. For LTQ-Orbitrap Elite data acquisition the machine was operated in the positive ion mode, with the following acquisition cycle: a full scan recorded in the orbitrap analyzer at reso-lution R 120,000 was followed by MS/MS (CID Rapid Scan Rate) of the 20 most intense peptide ions in the LTQ analyzer. The total acquisition gradient was either 100 or 240 minutes depending on the method of sample preparation. Mono-enzyme co-culture samples were measured on the LTQ-Orbitrap XL with slight modifications: a full scan recorded in the orbitrap analyzer at resolution R 120,000 was followed by MS/MS (CID) of the top 5 most intense peptide ions, with a total acquisition gradient of 95 minutes.
The MaxQuant software package (version 1.2.2.9) with the Andromeda search engine was used to identify and quantify proteins in cellular lysates and media (14; 33). Mouse and human IPI protein databases (both version 3.84, http://www.ebi.ac.uk/IPI/) plus common contaminants were used. With the exception of “second peptides”, which was deselected, default parameters were selected. For L-Lysine derived from precursors DAP, Z8, and DLYS8, variable labels were specified as K0, K8, and a custom modification (8 deuterium atoms for L-Lysine), respectively. Detection of non-precursor-based L-Lysine was specified as K0, K4, and K8.
Peptide and protein statistics (e.g., sequences, H/L ratios, intensities) were extracted from MaxQuant exported peptides.txt and proteingroups.txt, respectively. Peptides were determined to be species-specific if they only appeared in either one of the human or mouse IPI protein databases. Percent heavy label was calculated from the H/L ratio (HtoL) as =100*HtoL/(HtoL+1). In order to determine the overlap of H/L ratios between the human and mouse sequence-specific peptides, the median H/L ratio of each species was first determined. Next, the average of these two median values was used as a separator for each cell type and the miscategorizations were determined by the percentage of misclassified peptides on either side of this separator.
The raw data associated with this study will be released upon manuscript acceptance.
Cells were seeded in 96-well plates (2000 cells/well) and grown to 40% confluence in SILAC media containing 798 μM Ko or 10 mM DAP DMEM with 10% dialyzed fetal bovine serum (FBS). Cells were then inhibited with eight different drug concentrations (2 fold dilution) in eight replicates. Drugs used were Stattic (STAT3 inhibitor), PI3K-IV (PI3K inhibitor), AKT-VIII (AKT inhibitor), and SL327 (MEK inhibitor). After 48 hours drug treatment cell viability was measured by Resazurin (Sigma) as described by manufacturer. Cell viability relative to untreated cells was calculated to obtain dose-response curves.
Frozen cell pellets were thawed and lysed for 20 minutes with NP40 lysis buffer, which contained 1% Nonidet P-40, 1 mM sodium orthovanadate, and Complete protease inhibitors (Roche Diagnostics) in PBS. Protein concentrations were determined by the Bradford assay (BioRad) and adjusted to 1-1.5 mg/mL. Protein was then denatured in 2% SDS for 5 minutes at 95° C. Approximately 20 μg of each sample was then separated by SDS-PAGE, transferred to PVDF membrane, and immunoblotted using primary and secondary antibodies. All antibodies were from Cell Signaling. Chemoluminescence visualization was performed on Kodak or HyBlotCL films and films were scanned by a microTEK scanner at 600 d.p.i. in gray scale. The membranes were stripped and reprobed with anti-GAPDH (Cell Signaling) to test for protein loading.
Synthesis of Z-Lysine [Nα-Cbz-lysine (K8)]
To a solution of saturated aqueous NaHCO3 (1.25 mL) and L-lysine.2HCl (250 mg, 1.11 mmol, 1.00 equiv) was added solid NaHCO3 (105 mg, 1.13 equiv, 1.25 mmol) followed by aqueous CuSO4 (1.5 mL, 0.50 M, 0.68 mmol 0.60 equiv), immediately forming a blue copper complex. After stirring for 10 min, di-tert-butyl dicarbonate (325 mg, 1.49 mmol, 1.35 equiv) was added in 1 mL acetone. After stirring for 16 h, additional di-tert-butyl dicarbonate solid (150 mg, 0.621 equiv, 0.690 mmol) was added. After 24 h, the reaction was quenched with methanol (1 mL) and stirred for an additional 16 h. Ethyl acetate (1 mL) and water (1 mL) were added and the heterogeneous suspension was filtered. The recovered blue solid was taken up in H2O (3 mL), sonicated for 30 s, and filtered. After air drying, the Nε-Boc-protected copper complex was collected as a fine periwinkle blue powder (235 mg, 0.423 mmol, 74.2% yield), which was used without further purification.
To a suspension of Nε-Boc-protected copper complex (235 mg, 0.417 mmol, 1.00 equiv) in acetone (1.5 mL) was added 8-hydroxyquinoline (130 mg, 0.900 mmol, 2.13 equiv) and 10% Na2CO3 (1.8 mL). After 1 h, N-(Benzyloxycarbonyloxy)succinimide (205 mg, 0.821 mmol, 1.97 equiv) in 1 mL acetone was added dropwise over 10 min and stirred for 1 h. The reaction mixture was filtered, and the residue washed with water (3×1 mL). The pale green filtrate was acidified carefully with 1 N HCl to a pH of 2, and extracted with ethyl acetate (2×5 mL). The combined organics were washed with brine, dried over sodium sulfate, filtered, and concentrated by rotary evaporation to afford crude Nε-Boc-Nα-Cbz-L-lysine(K8) (148 mg, 45.7% yield, 0.381 mmol).
To a solution of crude Nε-Boc-Nα-Cbz-L-lysine(K8) (148 mg, 0.381 mmol, 1.00 equiv) in acetone (1.7 mL) was added TSOH.H2O (145 mg, 0.762 mmol, 2.00 equiv). After 16 h, crystals were collected by vacuum filtration and washed sparingly with cold acetone, giving Nα-Cbz-lysine(K8).TSOH (124 mg, 71.0% yield, 0.270 mmol).
Crude Nα-Cbz-lysine(K8).TSOH was dissolved in 1.0 mL 5% acetonitrile (v/v in water), treated with triethylamine (37.5 μL, 0.269 μmol, 1.00 equiv), and purified on a 5.5 g C-18 ISCO RediSep Gold column (5→90% acetonitrile in H2O). Lyophilization furnished Nα-Cbz-lysine(K8) as a fluffy white amorphous solid (77 mg, 0.27 mmol, 99% yield).
1H NMR (D2O, 600 MHz) (δ 7.25-7.35 (m, 5H), 5.04 (d, J=12.5 Hz, 1H), 4.97 (d, J=12.5 Hz, 1H), 3.83 (dm, JCH=140.4 Hz, 1H), 2.84 (dm, JCH=142.8 Hz, 2H), 1.66 (dm, JCH=128.4 Hz, 1H), 1.53 (dm, JCH=131.4 Hz, 3H), 1.28 (dm, JCH=132.6 Hz, 2H); 13C-NMR (D2O, 151 MHz) δ 179.8 (d, J=8.4 Hz), 179.5 (d, J=8.4 Hz), 128.7 (s), 128.2 (s), 127.6 (s), 66.8 (s), 56.2 (ddd, J=138.0, 46.2, 14.4 Hz), 55.8 (ddd, J=139.2, 46.8, 15.0 Hz), 34.2 (dt, J=161.0, 18.6 Hz), 31.1 (td, J=
138.6, 18.0 Hz), 23.2 (td, J=138.6, 18.8 Hz), 22.0 (t, J=137.4 Hz); [α]19D: −12.50±0.04° (c=2.00, 0.2 N HCl); FTIR (solid, cm−1) 3306, 3031, 2931, 1717, 1654, 1497, 1402, 1369, 1344, 1232; ESI-HRMS (m/z): calcd for C8 13C6H21 15N2O4 (M+H)+289.1643. found 289.1650.
This application claims the priority of U.S. provisional application No. 61/697,584 filed Sep. 6, 2012, the contents of which are incorporated herein by reference in their entirety into the present disclosure.
| Filing Document | Filing Date | Country | Kind |
|---|---|---|---|
| PCT/US13/58212 | 9/5/2013 | WO | 00 |
| Number | Date | Country | |
|---|---|---|---|
| 61697584 | Sep 2012 | US |
