Patent Application
20250066746
The present disclosure relates to compositions and methods for making and using water-soluble glycosyltransferase fusion proteins.
Glycosylation—the process by which carbohydrate-based compounds known as glycans are covalently attached to acceptor molecules, typically proteins and lipids —is fundamental to all life (Varki, A., “Biological Roles of Glycans,” Glycobiology 27:3-49 (2017) and Varki, A. et al., “Essentials of Glycobiology,” (Cold Spring Harbor (NY) (2015)). Following conjugation to biomolecules, glycans add an additional layer of information and play important roles in numerous biological processes (Moremen et al., “Vertebrate Protein Glycosylation: Diversity, Synthesis and Function,” Nat. Rev. Mol. Cell Biol. 13:448-62 (2012)) including cell adhesion and signaling (Crocker, P. R., “Siglecs: Sialic-acid-binding Immunoglobulin-like Lectins in Cell-cell Interactions and Signaling,” Curr. Opin. Struct. Biol. 12:609-15 (2002) and Cummings, R. D., “Stuck on Sugars—How Carbohydrates Regulate Cell Adhesion, Recognition, and Signaling,” Glycoconj. J. 36:241-257 (2019)), cell growth and development (Haltiwanger and Lowe, “Role of Glycosylation in Development,” Annu. Rev. Biochem. 73:491-537 (2004)), and immune recognition/response (Zhou and Cobb, “Glycans in Immunologic Health and Disease,” Annu. Rev. Immunol. 39:511-536 (2021) and Rudd et al., “Glycosylation and the Immune System,” Science 291:2370-6 (2001)), among others. Moreover, structural remodeling of protein-linked glycans can improve therapeutic properties in a number of ways such as extending activity and stability both in vitro and in vivo (Sola and Griebenow, “Effects of Glycosylation on the Stability of Protein Pharmaceuticals,” J. Pharm. Sci. 98:1223-45 (2009) and Sinclair and Elliott, “Glycoengineering: The Effect of Glycosylation on the Properties of Therapeutic Proteins,” J. Pharm. Sci. 94:1626-35 (2005), modulating interactions with specific immune receptors (Rothman et al., “Antibody-dependent Cytotoxicity Mediated by Natural Killer Cells is Enhanced by Castanospermine-induced Alterations of IgG Glycosylation,” Molecular Immunology 26:1113-23 (1989)), and targeting to specific cells or tissues (Friedman et al., “A Comparison of the Pharmacological Properties of Carbohydrate Remodeled Recombinant and Placental-derived Beta-glucocerebrosidase: Implications for Clinical Efficacy in Treatment of Gaucher Disease,” Blood 93:2807-16 (1999)).
As appreciation for the biological roles and therapeutic potential of glycans continues to grow, so too does the need for reliable, user-friendly technologies that enable their synthesis and remodeling. However, quantitative preparation of structurally-defined glycans and glycoconjugates remains technically challenging and represents a critical technology gap that limits widespread access to this important biomolecule class (Transforming Glycoscience: A Roadmap for the Future (The National Academies Press) (2012)). A major reason for this difficulty is the lack of template encoding in glycan biosynthesis, which distances carbohydrate structure and function from gene sequence. Hence, unlike nucleic acids and proteins, glycans cannot be directly produced from recombinant DNA technology. Instead, glycan biosynthesis is controlled by the availability, abundance, and activities of glycoenzymes, in particular, glycosyltransferases (GTs) that catalyze formation of specific glycosidic linkages by transferring sugar molecules from donor substrates (e.g., nucleotide sugar or lipid-linked sugar) to hydroxyl groups of acceptor molecules (Lairson et al., “Glycosyltransferases: Structures, Functions, and Mechanisms,” Annu. Rev. Biochem. 77:521-55 (2008) and Taniguchi et al., “Handbook of Glycosyltransferases and Related Genes,” (Springer, Tokyo, Japan (2014)) and glycosyl hydrolases (GHs) that cleave glycan structures during oligosaccharide maturation (Davies and Henrissat, “Structures and Mechanisms of Glycosyl hydrolases,” Structure 3:853-9 (1995)).
GTs exhibit unique catalytic specificities for a wide range of sugar donors and acceptor substrates and generate products with distinct anomeric configurations, which helps to explain the vast structural diversity of “glycospace”. In mammals alone, it is estimated that there are approximately 7,000 oligosaccharide structures (Cummings, R. D., “The Repertoire of Glycan Determinants in the Human Glycome,” Mol. Biosyst. 5:1087-104 (2009)) whose generation involves more than 200 GTs (Moremen Et Al., “Expression System For Structural And Functional studies of Human Glycosylation Enzymes,” Nat. Chem. Biol. 14:156-162 (2018) from 45 different protein families that have been annotated in the carbohydrate-active enzymes (CAZy) database (Lombard et al., “The Carbohydrate-active Enzymes Database (CAZy) in 2013,” Nucleic Acids Res 42: D490-5 (2014)). Moreover, GTs are proficient at replicating the diversity of naturally occurring glycans and glycoconjugates in unnatural contexts, leading to their emergence as powerful synthetic tools for building complex glycomolecules in the laboratory. Much of the progress in this regard exploits sugar nucleotide-dependent GTs of mammalian and bacterial origin for synthesis of complex carbohydrates, glycoconjugates, and glycosylated natural products, which are generated by functionally reconstituting artificial networks of these glycoenzymes within model cellular systems (Clausen et al., “Glycosylation Engineering in Essentials of Glycobiology (eds. rd et al.) 713-728 (Cold Spring Harbor (NY) (2015); Natarajan et al., “Metabolic Engineering of Glycoprotein Biosynthesis in Bacteria,” Emerg. Top Life Sci. 2:419-432 (2018); Williams et al., “Metabolic Engineering of Capsular Polysaccharides,” Emerg. Top Life Sci. 2:337-348 (2018); and Pandey et al., “Metabolic Engineering of Glycosylated Polyketide Biosynthesis,” Emerg. Top Life Sci. 2:389-403 (2018)) or in cell-free, one-pot reaction systems (Na et al., “Recent Progress in Synthesis of Carbohydrates With Sugar Nucleotide-dependent Glycosyltransferases,” Curr. Opin. Chem. Biol. 61:81-95 (2021); Jaroentomeechai et al., “Cell-free Synthetic Glycobiology: Designing and Engineering Glycomolecules Outside of Living Cells,” Front. Chem. 8:645 (2020); and Kightlinger et al., “Synthetic Glycobiology: Parts, Systems, and Applications,” ACS Synth. Biol. 9:1534-1562 (2020)).
These developments notwithstanding, broad access to GTs for fundamental and applied research is bottlenecked by difficulties associated with their recombinant expression. A major reason for this difficulty is that many GTs catalyze reactions at membrane interfaces (e.g., between the cytoplasm and periplasm in Gram-negative bacteria or between the cytosol and endoplasmic reticulum (ER)/Golgi organelles within eukaryotes). As such, these enzymes are typically either secretory proteins or integral membrane proteins (IMPs) that need post-translational modifications (PTMs) (e.g., disulfide bonds, N-linked glycosylation) and/or specialized chaperones to achieve proper folding, membrane translocation/insertion, and function. Efforts to express GTs in the absence of required PTMs or chaperones, or in the presence of single- or multi-pass transmembrane domains (TMDs) or terminal signal peptides (e.g., N-terminal export signals, C-terminal retention signals), are often met with non-functional protein aggregates. This is particularly pronounced for expression of mammalian GTs in bacterial hosts, with successful reports often involving time- and labor-intensive searches for solubility-enhancing fusion partners and molecular chaperones, optimal host strains and culture conditions, and compatible detergents and denaturants for IMP solubilization and in vitro refolding from inclusion bodies, respectively (Skretas et al., “Expression of Active Human Sialyltransferase ST6GalNAcI in Escherichia coli,” Microb. Cell Fact 8:50 (2009); Rao et al., “Structural Insight Into mammalian Sialyltransferases,” Nat. Struct. Mol. Biol. 16:1186-8 (2009); and Ramakrishnan and Qasba, “Crystal Structure of Lactose Synthase Reveals a Large Conformational Change in its Catalytic Component, the Beta1,4-galactosyltransferase-I,” J. Mol. Biol. 310:205-18 (2001)).
For these reasons, functional expression of mammalian GTs in bacteria remains rare. Instead, eukaryotic cells remain the preferred host for producing recombinant glycoenzymes albeit with most studies involving small-scale expression of just one or a few GTs (Taniguchi et al., “Handbook of Glycosyltransferases and Related Genes,” (Springer, Tokyo, Japan (2014)). To date, there are only a few reports of larger-scale expression campaigns involving significant numbers of GTs: one such study describes expression of 51 human GTs as fusions to the yeast cell wall Pir proteins to enable immobilization on the surface of Saccharomyces cerevisiae (Shimma et al., “Construction of a Library of Human Glycosyltransferases Immobilized in the Cell Wall of Saccharomyces cerevisiae,” Appl. Environ. Microbiol. 72:7003-12 (2006)) while a second study describes expression of 339 human glycoenzymes as fusions to a solubility-enhancing GFP domain in either mammalian cells (HEK293) or baculovirus-infected insect cells (Moremen Et Al., “Expression System For Structural And Functional studies of Human Glycosylation Enzymes,” Nat. Chem. Biol. 14:156-162 (2018)). Interestingly, the authors of this latter study explore the potential of Escherichia coli for human glycoenzyme expression but report that all GTs expressed in this host accumulate as insoluble aggregates (Moremen Et Al., “Expression System For Structural And Functional studies of Human Glycosylation Enzymes,” Nat. Chem. Biol. 14:156-162 (2018)). Thus, the biosynthetic capacity and versatility of simple E. coli bacteria, one of the most important model organisms in biology and biotechnology (Blount, Z. D., “The Unexhausted Potential of E. coli,” Elife 4(2015)), is yet to be unlocked for functional expression of GTs on a large scale.
The present disclosure is directed to overcoming these and other deficiencies in the art.
A first aspect of the present disclosure relates to a nucleic acid construct. The nucleic acid construct includes a chimeric nucleic acid molecule encoding a tripartite glycosyltransferase fusion protein. The chimeric nucleic acid molecule includes a first nucleic acid moiety encoding an amphipathic shield domain protein; a second nucleic acid moiety encoding a glycosyltransferase; and a third nucleic acid moiety encoding a water soluble expression decoy protein. The first nucleic acid moiety is coupled to the second nucleic acid moiety's 3′ end and the third nucleic acid moiety is coupled to the second nucleic acid moiety's 5′ end. The coupling may be direct or indirect.
Another aspect of the present disclosure relates to an expression vector including the nucleic acid construct according to the present disclosure.
Another aspect of the present disclosure relates to a host cell comprising the nucleic acid construct of the present disclosure.
Another aspect of the present disclosure relates to a tripartite glycosyltransferase fusion protein produced by a host cell according to the present disclosure.
Another aspect of the present disclosure relates to a cell-free protein expression system. The cell-free protein expression system comprises a cell lysate or extract and a nucleic acid construct according to the present disclosure.
Another aspect of the present disclosure relates to a tripartite glycosyltransferase fusion protein produced by the cell-free expression system according to the present disclosure.
Another aspect of the present disclosure relates to a method of recombinantly producing a tripartite glycosyltransferase fusion protein in water soluble form. This method involves providing a host cell according to the present disclosure or a cell-free expression system according to the present disclosure. The method further involves culturing the host cell or using the cell-free expression system under conditions effective to express the tripartite glycosyltransferase fusion protein in a water soluble form within the host cell cytoplasm or the cell-free expression system.
Another aspect of the present disclosure relates to a tripartite glycosyltransferase fusion protein produced by the methods of recombinantly producing a tripartite glycosyltransferase fusion protein according to the present disclosure.
Another aspect of the present disclosure relates to a tripartite glycosyltransferase fusion protein comprising: an amino terminal water soluble expression decoy protein; a glycosyltransferase; and a carboxyl terminal amphipathic shield domain protein.
Another aspect of the present disclosure relates to a method of cell-free glycan remodeling. This method involves providing a glycan primer; providing one or more tripartite glycosyltransferase fusion protein(s) according to the present disclosure; and incubating the glycan primer with the one or more tripartite glycosyltransferase fusion protein(s) under conditions effective to transfer a glycosyl group to the glycan primer to produce a modified glycan structure.
The Examples of the present disclosure describe a generalizable workflow for efficient production of structurally diverse GTs using standard E. coli expression strains. At the heart of this workflow is a protein engineering method called SIMPLEx (solubilization of integral membrane proteins with high levels of expression) (Mizrachi et al., “Making Water-soluble Integral Membrane Proteins In Vivo Using an Amphipathic Protein Fusion Strategy,” Nat Commun 6:6826 (2015), which is hereby incorporated by reference in its entirety) that enables topological conversion of secretory and membrane-bound proteins into water-soluble variants. In the Examples of the present disclosure, this conversion is achieved for GTs by modifying their N-termini with a decoy protein that prevents membrane insertion and their C-termini with an amphipathic protein that effectively shields hydrophobic surfaces from the aqueous environment (
A first aspect of the present disclosure relates to a nucleic acid construct. The nucleic acid construct includes a chimeric nucleic acid molecule encoding a tripartite glycosyltransferase fusion protein. The chimeric nucleic acid molecule includes a first nucleic acid moiety encoding an amphipathic shield domain protein; a second nucleic acid moiety encoding a glycosyltransferase; and a third nucleic acid moiety encoding a water soluble expression decoy protein. The first nucleic acid moiety is coupled to the second nucleic acid moiety's 3′ end and the third nucleic acid moiety is coupled to the second nucleic acid moiety's 5′ end. The coupling may be direct or indirect.
Another aspect of the present disclosure relates to a tripartite glycosyltransferase fusion protein produced by the methods of recombinantly producing a tripartite glycosyltransferase fusion protein according to the present disclosure.
The nucleic acid molecules encoding the various polypeptide components of a tripartite glycosyltransferase fusion protein can be ligated together along with appropriate regulatory elements that provide for expression of the tripartite glycosyltransferase fusion protein. Typically, the nucleic acid construct encoding the chimeric protein can be inserted into any of the many available expression vectors and cell systems using reagents that are well known in the art and further described infra.
As used herein, “nucleic acid”, refers to a polymeric form of nucleotides of any length, either ribonucleotides or deoxynucleotides. Thus, this term includes, but is not limited to, single-, double-, or multi-stranded DNA or RNA, genomic DNA, cDNA, DNA-RNA hybrids, or a polymer comprising purine and pyrimidine bases or other natural, chemically or biochemically modified, non-natural, or derivatized nucleotide bases. The nucleic acid construct may be a synthetic nucleic acid construct. As used herein “synthetic” nucleic acid construct refers to a nucleic acid construct that is artificially produced and/or that does not exist in nature. As described in more detail herein, the nucleic acid constructs of the present disclosure are utilized to make water-soluble glycosyltransferases using an amphipathic protein fusion strategy. In particular, the nucleic acid constructs are part of a new strategy for the solubilization of glycosyltransferases based on the affinity for hydrophobic surfaces displayed by amphipathic proteins.
As used herein, the term “glycosyltransferase” (GT) includes an enzyme or fragment thereof which catalyzes the transfer of a donor glycosyl moiety from a glycosyl donor to an acceptor. Suitable glycosyl donors include, without limitation, CMP-sialic acid, GDP-fucose, GDP-mannose, UDP-glucose, UDP-galactose, UDP-xylose, UDP-N-acetylglucosamine, UDP-N-acetylgalactosamine, UDP-glucuronic acid, Dolichol-P-glucose, Dolichol-P-mannose, Dolichol-P-P-(glucose3-mannose9-GlcNAc2), and undecaprenyl-P—P—N-acetylmuramic acid-pentapeptide-GlcNAc). Suitable acceptor moieties include, without limitation, oligosaccharides, monosaccharides, polypeptides, proteins, lipids such as ceramides, small organic molecules, and nucleic acid molecules such as DNA.
GTs may be classified as (i) single-pass transmembrane proteins with C-termini in the cytoplasm (type I transmembrane protein); (ii) single-pass transmembrane proteins with N-termini in the cytoplasm (type II transmembrane protein); (iii) multi-pass transmembrane proteins; (iv) secretory proteins with N-terminal signal peptides and C-terminal ER retention domains; and (v) cytosolic proteins. In some embodiments, the glycosyltransferase is selected from the group consisting of (i) a single-pass transmembrane protein with C-terminus in cytoplasm (type I transmembrane protein); (ii) a single-pass transmembrane protein with N-terminus in cytoplasm (type II transmembrane protein); (iii) a multi-pass transmembrane protein; and (iv) a secretory protein with N-terminal signal peptide and C-terminal ER retention domain.
In some embodiments, the glycosyltransferase is a full-length glycosyltransferase. Accordingly the second nucleic acid moiety encodes a full-length glycosyltransferase. In accordance with such embodiments, the second nucleic acid moiety comprises a full-length GT gene.
For example, the full-length GT may contain an internal single-pass or multi-pass TMD (e.g., human Dol-P-Man:Man(5)GlcNAc(2)-PP-Dol alpha-1,3-mannosyltransferase (HsAlg3), human Dol-P-Man:Man(7)GlcNAc(2)-PP-Dol alpha-1,6-mannosyltransferase (HsAlg12), human GPI mannosyltransferase 1 (HsPIGM), human GPI mannosyltransferase 3 (HsPIGB), human GPI mannosyltransferase 4 (HsPIGZ), human dolichyl pyrophosphate Man9GlcNAc2 alpha-1,3-glucosyltransferase (HsAlg6), human probable dolichyl pyrophosphate Glc1Man9GlcNAc2 alpha-1,3-glucosyltransferase (HsAlg8), human Dol-P-Glc:Glc(2)Man(9)GlcNAc(2)-PP-Dol alpha-1,2-glucosyltransferase (HsAlg10), human ceramide glucosyltransferase (HsUGCG), E. coli undecaprenyl-phosphate alpha-N-acetylglucosaminyl 1-phosphate transferase (EcWecA), yeast beta-1,4-mannosyltransferase OS=Saccharomyces cerevisiae (ScAlg1), and yeast GDP-Man:Man(3)GlcNAc(2)-PP-Dol alpha-1,2-mannosyltransferase (ScAlg11) (
In other embodiments, the full-length GT may be a predicted cytosolic GT (e.g., human isoform 2 of putative UDP-N-acetylglucosamine transferase (HsAlg13), human dolichol-phosphate mannosyltransferase subunit 1 (HsDPM1), human glycogenin-1 (HsGLYG), Campylobacter jejuni CsTII (CjCsTII), Neisseria meningitidis polysialic acid O-acetyltransferase (NmPolysiaT), Campylobacter jejuni beta-1,3-galactosyltransferase (CjCgtB), Helicobacter pylori (strain 51) beta-4-galactosyltransferase (HpLgtB), Neisseria meningitidis serogroup B (strain MC58) lacto-N-neotetraose biosynthesis glycosyltransferase LgtB (NmLgtB), Neisseria gonorrhoeae lacto-N-neotetraose biosynthesis glycosyltransferase (NgLgtB), E. coli galactoside 2-alpha-L-fucosyltransferase WbgL (EcFUT), Legionella pneumophila subsp. Pneumophila subversion of eukaryotic traffic protein A (LpSetA), and Neisseria meningitidis Alpha-2,9-polysialyltransferase (NmSynE) (
As described infra N-/C-terminal transmembrane domains (TMDs) as well as C-terminal ER retention domains in mammalian GTs are used as membrane anchors and are dispensable for catalytic activity (Harduin-Lepers et al., “The Human Sialyltransferase Family,” Biochimie 83727-83737 (2001), which is hereby incorporated by reference in its entirety). Thus, in some embodiments, the glycosyltransferase is a truncated glycosyltransferase. The truncated glycosyltransferase may exclude a GT C-terminal ER retention domain, a terminal TMD anchor, or both a C-terminal ER retention domain and a terminal TMD anchor. In some embodiments, the truncated glycosyltransferase excludes an N-terminal signal peptides. Various exemplary truncated GTs are provided in
Glycosyltransferases play vital roles in glycosylation and glycan remodeling. The tripartite glycosyltransferase fusion proteins according to the present disclosure are water soluble following extraction from their native environment (e.g., a cellular membrane) without the use of detergents and/or detergent-like amphiphiles, overproduction using recombinant systems, protein engineering, and/or mutations to the GT itself, thereby allowing for improved functional and structural studies of GTs as well as in vitro reconstitution of enzymatic activity or in vitro reconstitution of a biological pathway involving water soluble GT enzymes and engineering of biological/metabolic pathways involving the water soluble GTs.
The GTs according to the present disclosure may be prokaryotic glycosyltransferases or eukaryotic glycosyltransferase (e.g., human glycosyltransferases, rodent glycosyltransferases, yeast glycosyltransferases). Suitable exemplary prokaryotic and eukaryotic glycosyltransferases are identified in
The glycosyltransferase may be selected from the group consisting of fucosyltransferases (FucTs), galactosyltransferases (Gals), glucosyltransferases (GlcTs), mannosyltransferases (ManTs), N-acetylgalactosyltransferases (GalNAcTs), N-acetylglucosaminyltransferases (GlcNAcTs), and sialyltransferases (SiaTs).
Fucosyltransferases (FucTs) catalyze the transfer a fucose sugar from a donor substrate to an acceptor substrate. Suitable FucTs include, without limitation, human galactoside 2-alpha-L-fucosyltransferase 1 (HsFUT1), human galactoside 2-alpha-L-fucosyltransferase 2 (HsFUT2), HUMAN Galactoside 3(4)-L-fucosyltransferase (HsFUT3), human alpha-(1,3)-fucosyltransferase 4 (HsFUT4), human alpha-(1,3)-fucosyltransferase 5 (HsFUT5), human alpha-(1,3)-fucosyltransferase 6 (HsFUT6), human alpha-(1,3)-fucosyltransferase 7 (HsFUT7), human alpha-(1,6)-fucosyltransferase (HsFUT8), human alpha-(1,3)-fucosyltransferase 9 (HsFUT9), human alpha-(1,3)-fucosyltransferase 10 (HsFUT10), human alpha-(1,3)-fucosyltransferase 11 (HsFUT11), and human GDP-fucose protein O-fucosyltransferase 1 (HsPOFUT1) (see, e.g.,
Galactosyltransferases (Gals) catalyze the transfer of a galactose sugar from a donor substrate to an acceptor substrate. Suitable Gals include, without limitation, human beta-1,3-galactosyltransferase 1 (HsB3GalT1), human beta-1,3-galactosyltransferase 2 (HsB3GalT2), human beta-1,4-galactosyltransferase 1 (HsB4GalT1), human beta-1,4-galactosyltransferase 2 (HsB4GalT2), human beta-1,4-galactosyltransferase 3 (HsB4GalT3), human beta-1,4-galactosyltransferase 4 (HsB4GalT4), human beta-1,4-galactosyltransferase 5 (HsB4GalT5), and human beta-1,4-galactosyltransferase 6 (HsB4GalT6) (see, e.g.,
Glucosyltransferases (GlcTs) catalyze the transfer of a glucose sugar from a donor substrate to an acceptor substrate. Suitable GlcTs include, without limitation, human dolichyl-phosphate beta-glucosyltransferase (HsAlg5), human dolichyl pyrophosphate man9GlcNAc2 alpha-1,3-glucosyltransferase (HsAlg6), human probable dolichyl pyrophosphate Glc1Man9GlcNAc2 alpha-1,3-glucosyltransferase (HsAlg8), human Dol-P-Glc:Glc2Man9GlcNAc2—PP-Dol alpha-1,2-glucosyltransferase (HsAlg10), human ceramide glucosyltransferase (HsUGCG), human beta-1,3-glucosyltransferase (HsB3GLCT), and human protein O-glucosyltransferase 1 (HsPOGLUT1) (see, e.g.,
Mannosyltransferases (ManTs) catalyze the transfer of a mannose sugar from a donor substrate to an acceptor substrate. Suitable ManTs include, without limitation, human chitobiosyldiphosphodolichol beta-mannosyltransferase (HsAlg1), human alpha-1,3/1,6-mannosyltransferase (HsAlg2), human Dol-P-Man:Man(5)GlcNAc(2)-PP-Dol alpha-1,3-mannosyltransferase (HsAlg3), human GDP-man:man(3)GlcNAc(2)-PP-Dol alpha-1,2-mannosyltransferase (HsAlg11), human dol-p-man:man(7)GlcNAc(2)-PP-Dol alpha-1,6-mannosyltransferase (HsAlg12), human dolichol-phosphate mannosyltransferase subunit 1 (HsDPM1), human GPI mannosyltransferase 1 (HsPIGM), human GPI mannosyltransferase 3 (HsPIGB), human GPI mannosyltransferase 4 (HsPIGZ), yeast beta-1,4-mannosyltransferase OS=Saccharomyces cerevisiae (ScAlg1), and yeast GDP-Man:Man(3)GlcNAc(2)-PP-Dol alpha-1,2-mannosyltransferase (ScAlg11) (see, e.g.,
N-acetylgalactosyltransferases (GalNAcTs) catalyze the transfer of an N-acetylgalactosamine to an acceptor substrate. Suitable GalNAcTs include, without limitation, human alpha-N-acetylgalactosaminide alpha-2,6-sialyltransferase 1 (HsST6GalNAc1), human alpha-N-acetylgalactosaminide alpha-2,6-sialyltransferase 2 (HsST6GalNAc2), human alpha-N-acetyl-neuraminyl-2,3-beta-galactosyl-1,3-N-acetyl-galactosaminide alpha-2,6-sialyltransferase (HsST6GalNAc4), human polypeptide N-acetylgalactosaminyltransferase 1 (HsppGalNAcT1), human polypeptide N-acetylgalactosaminyltransferase 2 (HsppGalNAcT2), human polypeptide N-acetylgalactosaminyltransferase 3 (HsppGalNAcT3), human polypeptide N-acetylgalactosaminyltransferase 4 (HsppGalNAcT4), human polypeptide N-acetylgalactosaminyltransferase 5 (HsppGalNAcT5), human polypeptide N-acetylgalactosaminyltransferase 6 (HsppGalNAcT6), human N-acetylgalactosaminyltransferase 7 (HsppGalNAcT7), human probable polypeptide N-acetylgalactosaminyltransferase 8 (HsppGalNAcT8), human polypeptide N-acetylgalactosaminyltransferase 9 (HsppGalNAcT9), human polypeptide N-acetylgalactosaminyltransferase 10 (HsppGalNAcT10), and human UDP-GalNAc:beta-1,3-N-acetylgalactosaminyltransferase 1 (HsB3GALNT1) (see, e.g.,
N-acetylglucosaminyltransferases (GlcNAcTs) catalyze the transfer of an N-acetylglucosamine to an acceptor substrate. Suitable GlcNAcTs include, without limitation, human alpha-1,3-mannosyl-glycoprotein 2-beta-N-acetylglucosaminyltransferase (HsGnTI/MGAT1), human alpha-1,6-mannosyl-glycoprotein 2-beta-N-acetylglucosaminyltransferase (HsGnTII/MGAT2), human beta-1,4-mannosyl-glycoprotein 4-beta-N-acetylglucosaminyltransferase (HsGnTIII/MGAT3), human alpha-1,3-mannosyl-glycoprotein 4-beta-N-acetylglucosaminyltransferase a (HsGnTIV/MGAT4), human beta-1,3-galactosyl-O-glycosyl-glycoprotein beta-1,6-N-acetylglucosaminyltransferase (HsGCNT1), human N-acetyllactosaminide beta-1,6-N-acetylglucosaminyltransferase (HsGCNT2), human N-acetyllactosaminide beta-1,3-N-acetylglucosaminyltransferase 2 (HsB3GNT2), human acetylgalactosaminyl-O-glycosyl-glycoprotein beta-1,3-N-acetylglucosaminyltransferase (HsB3GNT6), human phosphatidylinositol N-acetylglucosaminyltransferase subunit A (HsPIGA), Nicotiana tabacum alpha-1,3-mannosyl-glycoprotein 2-beta-N-acetylglucosaminyltransferase (NtGnTI), and Nicotiana tabacum alpha-1,6-mannosyl-glycoprotein 2-beta-N-acetylglucosaminyltransferase-like (NtGnTII) (see, e.g.,
Sialyltransferases (SiaTs) catalyze the transfer of sialic acid to an acceptor substrate. Suitable SiaTs include, without limitation, human CMP-N-acetylneuraminate-beta-galactosamide-alpha-2,3-sialyltransferase 1 (HsST3Gal1), human CMP-N-acetylneuraminate-beta-1,4-galactoside alpha-2,3-sialyltransferase (HsST3Gal3), human CMP-N-acetylneuraminate-beta-galactosamide-alpha-2,3-sialyltransferase 4 (HsST3Gal4), human type 2 lactosamine alpha-2,3-sialyltransferase (HsST3Gal6), human beta-galactoside alpha-2,6-sialyltransferase 1 (HsST6Gal1), human alpha-N-acetylgalactosaminide alpha-2,6-sialyltransferase 1 (HsST6GalNAc1), human alpha-N-acetylgalactosaminide alpha-2,6-sialyltransferase 2 (HsST6GalNAc2), human alpha-N-acetyl-neuraminyl-2,3-beta-galactosyl-1,3-N-acetyl-galactosaminide alpha-2,6-sialyltransferase (HsST6GalNAc4), human alpha-N-acetylneuraminide alpha-2,8-sialyltransferase (HsST8Sia1), human alpha-2,8-sialyltransferase 8b (HsST8Sia2), human sia-alpha-2,3-gal-beta-1,4-GlcNAc-R:alpha 2,8-sialyltransferase (HsST8Sia3), human CMP-N-acetylneuraminate-poly-alpha-2,8-sialyltransferase (HsST8Sia4), and Neisseria meningitidis alpha-2,9-polysialyltransferase (NmSynE) (see, e.g.,
In some embodiments, the glycosyltransferase is selected from the group consisting of human galactoside 2-alpha-L-fucosyltransferase 1 (HsFUT1), human galactoside 2-alpha-L-fucosyltransferase 2 (HsFUT2), HUMAN Galactoside 3(4)-L-fucosyltransferase (HsFUT3), human alpha-(1,3)-fucosyltransferase 4 (HsFUT4), human alpha-(1,3)-fucosyltransferase 5 (HsFUT5), human alpha-(1,3)-fucosyltransferase 6 (HsFUT6), human alpha-(1,3)-fucosyltransferase 7 (HsFUT7), human alpha-(1,6)-fucosyltransferase (HsFUT8), human alpha-(1,3)-fucosyltransferase 9 (HsFUT9), human alpha-(1,3)-fucosyltransferase 10 (HsFUT10), human alpha-(1,3)-fucosyltransferase 11 (HsFUT11), human GDP-fucose protein O-fucosyltransferase 1 (HsPOFUT1), human CMP-N-acetylneuraminate-beta-galactosamide-alpha-2,3-sialyltransferase 1 (HsST3Gal1), human CMP-N-acetylneuraminate-beta-1,4-galactoside alpha-2,3-sialyltransferase (HsST3Gal3), human CMP-N-acetylneuraminate-beta-galactosamide-alpha-2,3-sialyltransferase 4 (HsST3Gal4), human type 2 lactosamine alpha-2,3-sialyltransferase (HsST3Gal6), human beta-galactoside alpha-2,6-sialyltransferase 1 (HsST6Gal1), human alpha-N-acetylgalactosaminide alpha-2,6-sialyltransferase 1 (HsST6GalNAc1), human alpha-N-acetylgalactosaminide alpha-2,6-sialyltransferase 2 (HsST6GalNAc2), human alpha-N-acetyl-neuraminyl-2,3-beta-galactosyl-1,3-N-acetyl-galactosaminide alpha-2,6-sialyltransferase (HsST6GalNAc4), human alpha-N-acetylneuraminide alpha-2,8-sialyltransferase (HsST8Sia1), human alpha-2,8-sialyltransferase 8b (HsST8Sia2), human sia-alpha-2,3-gal-beta-1,4-GlcNAc-R:alpha 2,8-sialyltransferase (HsST8Sia3), human CMP-N-acetylneuraminate-poly-alpha-2,8-sialyltransferase (HsST8Sia4), human polypeptide N-acetylgalactosaminyltransferase 1 (HsppGalNAcT1), human polypeptide N-acetylgalactosaminyltransferase 2 (HsppGalNAcT2), human polypeptide N-acetylgalactosaminyltransferase 3 (HsppGalNAcT3), human polypeptide N-acetylgalactosaminyltransferase 4 (HsppGalNAcT4), human polypeptide N-acetylgalactosaminyltransferase 5 (HsppGalNAcT5), human polypeptide N-acetylgalactosaminyltransferase 6 (HsppGalNAcT6), human N-acetylgalactosaminyltransferase 7 (HsppGalNAcT7), human probable polypeptide N-acetylgalactosaminyltransferase 8 (HsppGalNAcT8), human polypeptide N-acetylgalactosaminyltransferase 9 (HsppGalNAcT9), human polypeptide N-acetylgalactosaminyltransferase 10 (HsppGalNAcT10), human UDP-GalNAc:beta-1,3-N-acetylgalactosaminyltransferase 1 (HsB3GALNT1), human beta-1,4 N-acetylgalactosaminyltransferase 1 (HsB4GALNT1), human histo-blood group ABO system transferase (Hs-A-group), human lactosylceramide 4-alpha-galactosyltransferase (HsA4GalT), human beta-1,3-galactosyltransferase 1 (HsB3GalT1), human beta-1,3-galactosyltransferase 2 (HsB3GalT2), human beta-1,4-galactosyltransferase 1 (HsB4GalT1), human beta-1,4-galactosyltransferase 2 (HsB4GalT2), human beta-1,4-galactosyltransferase 3 (HsB4GalT3), human beta-1,4-galactosyltransferase 4 (HsB4GalT4), human beta-1,4-galactosyltransferase 5 (HsB4GalT5), human beta-1,4-galactosyltransferase 6 (HsB4GalT6), human histo-blood group ABO system transferase (Hs-B-group), human 2-hydroxyacylsphingosine 1-beta-galactosyltransferase (HsUGT8), human glycoprotein-N-acetylgalactosamine 3-beta-galactosyltransferase 1 (HsC1GLT), human C1GALT1-specific chaperone 1 (HsCOSMC), human chitobiosyldiphosphodolichol beta-mannosyltransferase (HsAlg1), human alpha-1,3/1,6-mannosyltransferase (HsAlg2), human Dol-P-Man:Man(5)GlcNAc(2)-PP-Dol alpha-1,3-mannosyltransferase (HsAlg3), human GDP-man:man(3)GlcNAc(2)-PP-Dol alpha-1,2-mannosyltransferase (HsAlg11), human dol-p-man:man(7)GlcNAc(2)-PP-Dol alpha-1,6-mannosyltransferase (HsAlg12), human isoform 2 of putative UDP-N-acetylglucosamine transferase (HsAlg13), human UDP-N-acetylglucosamine transferase subunit alg14 homolog (HsAlg14), human dolichol-phosphate mannosyltransferase subunit 1 (HsDPM1), human GPI mannosyltransferase 1 (HsPIGM), human GPI mannosyltransferase 3 (HsPIGB), human GPI mannosyltransferase 4 (HsPIGZ), human dolichyl-phosphate beta-glucosyltransferase (HsAlg5), human dolichyl pyrophosphate man9GlcNAc2 alpha-1,3-glucosyltransferase (HsAlg6), human probable dolichyl pyrophosphate Glc1Man9GlcNAc2 alpha-1,3-glucosyltransferase (HsAlg8), human Dol-P-Glc:Glc2Man9GlcNAc2-PP-Dol alpha-1,2-glucosyltransferase (HsAlg10), human ceramide glucosyltransferase (HsUGCG), human beta-1,3-glucosyltransferase (HsB3GLCT), human glycogenin-1 (HsGLYG), human protein O-glucosyltransferase 1 (HsPOGLUT1), human alpha-1,3-mannosyl-glycoprotein 2-beta-N-acetylglucosaminyltransferase (HsGnTI/MGAT1), human alpha-1,6-mannosyl-glycoprotein 2-beta-N-acetylglucosaminyltransferase (HsGnTII/MGAT2), human beta-1,4-mannosyl-glycoprotein 4-beta-N-acetylglucosaminyltransferase (HsGnTIII/MGAT3), human alpha-1,3-mannosyl-glycoprotein 4-beta-N-acetylglucosaminyltransferase a (HsGnTIV/MGAT4), human beta-1,3-galactosyl-O-glycosyl-glycoprotein beta-1,6-N-acetylglucosaminyltransferase (HsGCNT1), human N-acetyllactosaminide beta-1,6-N-acetylglucosaminyl-transferase (HsGCNT2), human N-acetyllactosaminide beta-1,3-N-acetylglucosaminyltransferase 2 (HsB3GNT2), human acetylgalactosaminyl-O-glycosyl-glycoprotein beta-1,3-N-acetylglucosaminyltransferase (HsB3GNT6), human phosphatidylinositol N-acetylglucosaminyltransferase subunit A (HsPIGA), human xyloside xylosyltransferase 1 (HsXXLT1), human UDP-glucuronosyltransferase 1-1 (HsUGT1A1), human beta-1,4-glucuronyltransferase 1 (HsB4GAT1), human UDP-glucuronosyltransferase 1-3 (HsUGT1A3), Campylobacter jejuni CsTII (CjCstII), Neisseria meningitidis polysialic acid O-acetyltransferase (NmPst), Campylobacter jejuni beta-1,3-galactosyltransferase (CjCgtB), Helicobacter pylori (strain 51) beta-4-galactosyltransferase (HpLgtB), Neisseria meningitidis serogroup B (strain MC58) lacto-N-neotetraose biosynthesis glycosyltransferase LgtB (NmLgtB), Neisseria gonorrhoeae lacto-N-neotetraose biosynthesis glycosyltransferase (NgLgtB), E. coli galactoside 2-alpha-L-fucosyltransferase WbgL (EcWbgL), E. coli undecaprenyl-phosphate alpha-N-acetylglucosaminyl 1-phosphate transferase (EcWecA), Legionella pneumophila subsp. Pneumophila Subversion of eukaryotic traffic protein A (LpSetA), Neisseria meningitidis alpha-2,9-polysialyltransferase (NmSynE), yeast beta-1,4-mannosyltransferase OS=Saccharomyces cerevisiae (ScAlg1), yeast GDP-Man:Man(3)GlcNAc(2)-PP-Dol alpha-1,2-mannosyltransferase (ScAlg11), Nicotiana tabacum alpha-1,3-mannosyl-glycoprotein 2-beta-N-acetylglucosaminyltransferase (NtGnTI), Nicotiana tabacum alpha-1,6-mannosyl-glycoprotein 2-beta-N-acetylglucosaminyltransferase-like (NtGnTII), Bos taurus n-acetyllactosaminide alpha-1,3-galactosyltransferase (BtGGTA1), mouse n-acetyllactosaminide alpha-1,3-galactosyltransferase (MmGGTA1), rat n-acetyllactosaminide alpha-1,3-galactosyltransferase (RnGGTA1), and Bos taurus beta-1,4-galactosyltransferase 1 (BtB4GalT1) (see, e.g.,
In some embodiments, the nucleic acid molecule encodes a second nucleic acid moiety encoding a glycosyltransferase having the amino acid sequence of any one of SEQ ID NOs: 1-174 (see
The Examples of the present disclosure demonstrate the use of tripartite glycosyltransferase fusion proteins (e.g., Sx-CjCstII, Sx-Δ36HsFucT7, Sx-Δ34HsST3Gal1, Sx-Δ29HsGnTI, Sx-Δ29HsGnTII, Sx-Δ44Hsβ4GalT1, Sx-Δ26HsST6Gal1, Sx-Δ44HsFucT8) to catalyze the formation of a spectrum of homogenous N-glycan structures on intact glycoproteins. Thus, in some embodiments, the glycosyltransferase is selected from the group consisting of Campylobacter jejuni CsTII (CjCstII), human alpha-(1,3)-fucosyltransferase 7 (HsFUT7), human CMP-N-acetylneuraminate-beta-galactosamide-alpha-2,3-sialyltransferase 1 (HsST3Gal1), human alpha-1,3-mannosyl-glycoprotein 2-beta-N-acetylglucosaminyltransferase (HsGnTI/MGAT1), human alpha-1,6-mannosyl-glycoprotein 2-beta-N-acetylglucosaminyltransferase (HsGnTII/MGAT2), human beta-1,4-galactosyltransferase 1 (Hsβ4GalT1), human β-galactoside-α2,6-sialyltransferase 1 (HsST6Gal1), and human alpha-(1,6)-fucosyltransferase (HsFUT8).
As used herein, the term “amphipathic shield domain protein” includes any protein that displays both hydrophilic and hydrophobic surfaces and is often associated with lipids as membrane anchors or involved in their transport as soluble particles. The amphipathic shield domain protein, in one embodiment, serves as a molecular shield to sequester large lipophilic surfaces of the glycosyltransferase from water.
In various other embodiments, the amphipathic shield domain protein is selected from the group consisting of apolipoprotein A (ApoA), apolipoprotein B (ApoB), apolipoprotein C (ApoC), apolipoprotein D (ApoD), apolipoprotein E (ApoE), apolipoprotein H (ApoH), truncated human apolipoprotein A1 lacking its 43-residue globular N-terminal domain (ApoAI*), and a peptide self-assembly mimic (PSAM). In particular, the amphipathic shield domain protein may be apolipoprotein A1 (ApoAI). As used herein, ApoAI avidly binds phospholipid molecules and organizes them into soluble bilayer structures or discs that readily accept cholesterol. ApoAI contains a globular amino-terminal (N-terminal) domain (residues 1-43) and a lipid-binding carboxyl-terminal (C-terminal) domain (residues 44-243). In some embodiments, the amphipathic shield domain protein is human apolipoprotein A1. The apolipoprotein A1 may be a truncated human apolipoprotein A1. Truncated variants of ApoA1 include, but are not limited to, human ApoAI lacking its 43-residue globular N-terminal domain (ApoA1*).
As used herein, ApoA1 exhibits remarkable structural flexibility, and may adopt a molten globular-like state for lipid-free ApoAI under conditions that may allow it to adapt to the significant geometry changes of the lipids with which it interacts. The present disclosure provides tripartite fusion proteins in which, for example, ApoAI* may be genetically fused to the carboxyl terminus of a glycosyltransferase (or truncated glycosyltransferase). As described herein, expression of such tripartite glycosyltransferase fusion proteins may yield appreciable amounts of globular, water-soluble tripartite glycosyltransferase fusion proteins that are stabilized in a hydrophobic environment and retain structurally relevant conformations. The approach provides, inter alia, a facile method for efficiently solubilizing structurally diverse glycosyltransferases, for example in both prokaryotic and eukaryotic cells, without the need for detergents or lipid reconstitutions.
As used herein, the term “water soluble expression decoy protein” includes any protein which serves to direct an glycosyltransferase into cellular cytoplasm. The water soluble expression decoy protein may assist in “tricking” a hydrophobic glycosyltransferase into thinking that it is not hydrophobic. The water soluble expression decoy protein may be selected from the group consisting of outer surface protein (OspA) lacking its native export signal peptide, DnaB lacking its native export signal peptide, and maltose-binding protein (MBP) lacking its N-terminal signal peptide. In some embodiments, the water soluble expression decoy protein is maltose-binding protein (MBP) lacking its N-terminal signal peptide.
In some embodiments of the nucleic acid constructs and the tripartite glycosyltransferase fusion proteins according to the present disclosure, the amphipathic shield domain protein is truncated human apolipoprotein A1 lacking its 43-residue globular N-terminal domain (ApoAI*) and the water soluble expression decoy protein is maltose-binding protein (MBP) lacking its N-terminal signal peptide. For example, the nucleic acid construct may comprise a chimeric nucleic acid molecule comprising a first nucleic acid moiety encoding truncated human apolipoprotein A1 lacking its 43-residue globular N-terminal domain (ApoAI*), a second nucleic acid moiety encoding human β-galactoside-α2,6-sialyltransferase 1 (HsST6Gal1) or a truncated HsST6Gal1 variant in which 26 amino acids from the N-terminus of HsST6Gal1 comprising its CT and TMD were deleted (Δ26HsST6Gal1), and a third nucleic acid moiety encoding maltose-binding protein (MBP) lacking its N-terminal signal peptide (ΔspMBP). Such embodiments are described in Example 1, where ΔspMBP-HsST6Gal1-ApoAI* (abbreviated as Sx-HsST6Gal1) and ΔspMBP-Δ26HsST6Gal1-ApoAI* (abbreviated as Sx-Δ26HsST6Gal1) are shown to accumulate almost exclusively in the soluble cytoplasmic fraction of E. coli cells. The importance of the amphipathic shield domain and water soluble expression decoy proteins is evidenced by expression of unfused HsST6Gal1 and Δ26HsST6Gal1, which were not observed to accumulate in the soluble fraction and were only observed in minimal amounts in the insoluble and detergent-solubilized fractions.
In some embodiments, the construct further includes a promoter and a termination sequence, where the promoter and the termination sequence are operatively coupled to the chimeric nucleic acid molecule.
The chimeric nucleic acid molecules of the present disclosure include DNA molecules (e.g., linear, circular, cDNA, chromosomal, genomic, or synthetic, double stranded, single stranded, triple-stranded, quadruplexed, partially double-stranded, branched, hair-pinned, circular, or in a padlocked conformation) and RNA molecules (e.g., tRNA, rRNA, mRNA, genomic, or synthetic) and analogs of the DNA or RNA molecules of the described as well as analogs of DNA or RNA containing non-natural nucleotide analogs, non-native inter-nucleoside bonds, or both.
In some embodiments, the first nucleic acid moiety, the second nucleic acid moiety, and/or the third nucleic acid moiety may be free of naturally flanking sequences (i.e., sequences located at the 5′ and 3′ ends of the first nucleic acid moiety, the second nucleic acid moiety, and/or the third nucleic acid moiety) in the chromosomal DNA of the organism from which the first nucleic acid moiety, the second nucleic acid moiety, and/or the third nucleic acid moiety was derived, respectively.
In various embodiments, the first nucleic acid moiety, the second nucleic acid moiety, and/or the third nucleic acid moiety may contain less than about 10 kb, 5 kb, 4 kb, 3 kb, 2 kb, 1 kb, 0.5 kb, 0.1 kb, 50 bp, 25 bp or 10 bp of naturally flanking nucleotide chromosomal DNA sequences of the microorganism from which the first nucleic acid moiety, the second nucleic acid moiety, and/or the third nucleic acid moiety was derived, respectively.
The chimeric nucleic acid molecules may further include one or more linker nucleic acid moieties coupling the first, second, and/or third nucleic acid moieties together.
The tripartite glycosyltransferase fusion proteins according to the present disclosure include a continuous polymer of amino acids which comprise the full or partial sequence of three or more distinct proteins. The construction of fusion proteins is well-known in the art. Two or more amino acids sequences may be joined chemically, for instance, through the intermediacy of a crosslinking agent. For example, a fusion protein may be generated by expression of a nucleic acid construct comprising a chimeric nucleic acid molecule according to the present disclosure in a host cell. Such nucleic acid constructs may generally also contain replication origins active in host cells and one or more selectable markers encoding, for example, drug or antibiotic resistance.
The tripartite glycosyltransferase fusion proteins of the present disclosure can be generated as described herein or using any other standard technique known in the art. For example, the tripartite glycosyltransferase fusion proteins can be prepared by translation of a chimeric nucleic acid molecule encoding a tripartite glycosyltransferase fusion protein according to the present disclosure. The chimeric nucleic acid molecule encoding a tripartite glycosyltransferase fusion protein is inserted into an expression vector which is used to transform or transfect a host cell.
Different chimeric nucleic acid molecules encoding unique tripartite glycosyltransferase fusion proteins may be present on separate nucleic acid constructs or on the same nucleic acid construct. Inclusion of different chimeric nucleic acid molecules encoding unique tripartite glycosyltransferase fusion proteins on the same nucleic acid molecule is advantageous, in that uptake of only a single species of nucleic acid by a host cell is sufficient to introduce sequences encoding the tripartite glycosystransferase(s) into the host cell. By contrast, when different chimeric nucleic acid molecules encoding unique tripartite glycosyltransferase fusion proteins are present on different nucleic acid constructs, both nucleic acid molecules are taken up by a particular host cell for the assay to be functional.
A nucleic acid construct comprising a chimeric nucleic acid molecule encoding a tripartite glycosyltransferase fusion proteins may be inserted into an expression system to which the nucleic acid construct is heterologous. The heterologous nucleic acid construct may be inserted into the expression system or vector in proper sense (5′-3′) orientation relative to the promoter and any other 5′ regulatory molecules, and correct reading frame. The preparation of the nucleic acid constructs can be carried out using standard cloning methods well known in the art as described by Sambrook et al., Molecular Cloning: A Laboratory Manual, Cold Springs Laboratory Press, Cold Springs Harbor, New York (1989), which is hereby incorporated by reference in its entirety. U.S. Pat. No. 4,237,224 to Cohen and Boyer, which is hereby incorporated by reference in its entirety, also describes the production of expression systems in the form of recombinant plasmids using restriction enzyme cleavage and ligation with DNA ligase.
Another aspect of the present disclosure is directed to tripartite glycosyltransferase fusion proteins produced by the host cells described herein.
As described herein, a variety of prokaryotic expression systems can be used to express the tripartite glycosyltransferase fusion proteins of the present disclosure. Expression vectors can be constructed which contain a promoter to direct transcription, a ribosome binding site, and a transcriptional terminator. Examples of regulatory regions suitable for this purpose in E. coli are the promoter and operator region of the E. coli tryptophan biosynthetic pathway (Yanofsky et al., “Repression is Relieved Before Attenuation in the trp Operon of Escherichia coli as Tryptophan Starvation Becomes Increasingly Severe,” J. Bacteria. 158:1018-1024 (1984), which is hereby incorporated by reference in its entirety) and the leftward promoter of phage lambda (N) (Herskowitz et al., “The Lysis-lysogeny Decision of Phage Lambda: Explicit Programming and Responsiveness,” Ann. Rev. Genet., 14:399-445 (1980), which is incorporated by reference in its entirety). Vectors used for expressing foreign genes in bacterial hosts generally will contain a sequence for a promoter which functions in the host cell. Plasmids useful for transforming bacteria include pBR322 (Bolivar et al., “Construction and Characterization of New Cloning Vehicles II. A Multipurpose Cloning System,” Gene 2:95-113 (1977), which is hereby incorporated by reference in its entirety), the pUC plasmids (Messing, “New M13 Vectors for Cloning,” Meth. Enzymol. 101:20-77 (1983), Vieira et al., “New pUC-derived Cloning Vectors with Different Selectable Markers and DNA Replication Origins,” Gene 19:259-268 (1982) which are hereby incorporated by reference in their entirety), and derivatives thereof. Plasmids may contain both viral and bacterial elements. Methods for the recovery of the proteins in biologically active form are discussed in U.S. Pat. No. 4,966,963 to Patroni and 4,999,422 to Galliher, which are incorporated herein by reference in their entirety. Suitable expression vectors include those which contain replicon and control sequences that are derived from species compatible with the host cell. For example, if E. coli is used as a host cell, plasmids such as pUC19, pUC18 or pBR322 may be used. Alternatively, plasmids such as pET28a and pMALc2× may be used. Other suitable expression vectors are described in Molecular Cloning: a Laboratory Manual: 3rd edition, Sambrook and Russell, 2001, Cold Spring Harbor Laboratory Press, which is hereby incorporated by reference in its entirety. Many known techniques and protocols for manipulation of nucleic acids, for example in preparation of nucleic acid constructs, mutagenesis, sequencing, introduction of DNA into cells and gene expression, and analysis of proteins, are described in detail in Current Protocols in Molecular Biology, Ausubel et al. eds., (1992), which is hereby incorporated by reference in its entirety.
Different genetic signals and processing events control many levels of gene expression (e.g., DNA transcription and messenger RNA (“mRNA”) translation) and subsequently the amount of fusion protein that is displayed on the ribosome surface. Transcription of DNA is dependent upon the presence of a promoter, which is a DNA sequence that directs the binding of RNA polymerase, and thereby promotes mRNA synthesis. Promoters vary in their “strength” (i.e., their ability to promote transcription). For the purposes of expressing a cloned gene, it is desirable to use strong promoters to obtain a high level of transcription and, hence, expression and surface display. Therefore, depending upon the host system utilized, any one of a number of suitable promoters may also be incorporated into the expression vector carrying the deoxyribonucleic acid molecule encoding the protein of interest coupled to a stall sequence. For instance, when using E. coli, its bacteriophages, or plasmids, promoters such as the T7 phage promoter, lac promoter, trp promoter, recA promoter, ribosomal RNA promoter, the PR and PL promoters of coliphage lambda and others, including but not limited, to lacUV5, ompF, bla, lpp, and the like, may be used to direct high levels of transcription of adjacent DNA segments. Additionally, a hybrid trp-lacUV5 (tac) promoter or other E. coli promoters produced by recombinant DNA or other synthetic DNA techniques may be used to provide for transcription of the inserted gene.
Translation of mRNA in prokaryotes depends upon the presence of the proper prokaryotic signals, which differ from those of eukaryotes. Efficient translation of mRNA in prokaryotes requires a ribosome binding site called the Shine-Dalgarno (“SD”) sequence on the mRNA. This sequence is a short nucleotide sequence of mRNA that is located before the start codon, usually AUG, which encodes the amino-terminal methionine of the protein. The SD sequences are complementary to the 3′-end of the 16S rRNA (ribosomal RNA) and probably promote binding of mRNA to ribosomes by duplexing with the rRNA to allow correct positioning of the ribosome. For a review on maximizing gene expression, see Roberts and Lauer, “Maximizing Gene Expression on a Plasmid Using Recombination In Vitro,” Methods in Enzymology 68:473-82 (1979), which is hereby incorporated by reference in its entirety.
In accordance with this and other aspects of the present disclosure, the amphipathic shield domain protein, glycosyltransferase, and/or water soluble expression decoy proteins are linked either directly or via a linker located adjacent to each other within the construct, coupled to each other in tandem or separated by at least one linker. In one embodiment, the chimeric nucleic acid molecule includes a linker coupling the nucleic acid moieties together. Likewise, the tripartite glycosyltransferase fusion proteins may include a linker coupling the amphipathic shield domain protein, the glycosyltransferase (or truncated glycosyltransferase), and the water soluble expression decoy protein together. The amphipathic shield domain protein, the glycosyltransferase (or truncated glycosyltransferase), and the water soluble expression decoy protein may be linked by a covalent linkage or may be linked by methods known in the art for linking peptides.
Linkers may include synthetic sequences of amino acids that are commonly used to physically connect polypeptide domains to each other or to biologically relevant moieties. Most linker peptides are composed of repetitive modules of one or more of the amino acids glycine and serine. Peptide linkers have been well-characterized and shown to adopt unstructured, flexible conformations. For example, linkers comprised of Gly and Ser amino acids have been found to not interfere with assembly and binding activity of the domains it connects. Freund et al., “Characterization of the Linker Peptide of the Single-chain Fv Fragment of an Antibody by NMR Spectroscopy,” FEBS 320:97 (1993), which is hereby incorporated by reference in its entirety.
The nucleic acid constructs and tripartite glycosyltransferase fusion proteins of the present disclosure may include a flexible polypeptide linker separating the amphipathic shield domain protein, glycosyltransferase (or truncated glycosyltransferase), and/or water soluble expression decoy proteins and allowing for their independent folding. The linker is optimally 15 amino acids or 60 Å in length (˜4 Å per residue) but may be as long as 30 amino acids but preferably not more than 20 amino acids in length. It may be as short as 3 amino acids in length, but more preferably is at least 6 amino acids in length. To ensure flexibility and to avoid introducing steric hindrance that may interfere with the independent folding of the fragment domain of reporter protein and the members of the putative binding pair, the linker should be comprised of small, preferably neutral residues such as Gly, Ala, and Val, but also may include polar residues that have heteroatoms such as Ser and Met, and may also contain charged residues. The first, second, and third proteins may be linked via a short polypeptide linker sequence. Suitable linkers include peptides of between about 2 and about 40 amino acids in length and may include, for example, glycine residues Gly185 and Gly186. Preferred linker sequences include glycine-rich (e.g. G30.5), serine-rich (e.g., GSG, GSGS, GSGSG, GSNG), or alanine rich (e.g., TSAAA) linker sequences. Other exemplary linker sequences have a combination of glycine, alanine, proline and methionine residues such as AAAGGM; AAAGGMPPAAAGGM (SEQ ID NO: 175); AAAGGM; and PPAAAGGMM. Linkers may have virtually any sequence that results in a generally flexible chimeric protein.
Another aspect of the present disclosure relates to an expression vector including the nucleic acid construct of the present disclosure. Suitable nucleic acid vectors include, without limitation, plasmids, baculovirus vectors, bacteriophage vectors, phagemids, cosmids, fosmids, bacterial artificial chromosomes, viral vectors (for example, viral vectors based on vaccinia virus, poliovirus, adenovirus, adeno-associated virus, SV40, herpes simplex virus, and the like), P1-based artificial chromosomes, yeast plasmids, yeast artificial chromosomes, and other vectors. In some embodiments of the present disclosure, vectors suitable for use in prokaryotic host cells. Accordingly, exemplary vectors for use in prokaryotes such as Escherichia coli include, but are not limited to, pACYC184, pBeloBac11, pBR332, pBAD33, pBBR1MCS and its derivatives, pSC101, SuperCos (cosmid), pWE15 (cosmid), pTrc99A, pBAD24, vectors containing a ColE1 origin of replication and its derivatives, pUC, pBluescript, pGEM, and pTZ vectors.
Another aspect of the present disclosure relates to a host cell comprising the nucleic acid construct of the present disclosure. In accordance with this and other aspects of the present disclosure, suitable host cells include both eukaryotic and prokaryotic cells.
In some embodiments, the host cell is eukaryotic. Eukaryotic host cells, include without limitation, animal cells, fungal cells, insect cells, plant cells, and algal cells. In some embodiments, the eukaryotic host cells are selected from the group consisting of human cells, yeast, cells, and cell lines. Suitable eukaryotic host cells include, but are not limited to, Pichia pastoris, Pichia finlandica, Pichia trehalophila, Pichia koclamae, Pichia membranaefaciens, Pichia opuntiae, Pichia thennotolerans, Pichia salictaria, Pichia guercuum, Pichia pijperi, Pichia stiptis, Pichia methanolica, Pichia sp., Saccharomyces cerevisiae, Saccharomyces sp., Hansenula polymorpha, Kluyveromyces sp., Kluyveromyces lactis, Candida albicans, Aspergillus nidulans, Aspergillus niger, Aspergillus oryzae, Trichoderma reesei, Chrysosporium lucknowense, Fusarium sp., Fusarium gramineum, Fusarium venenatum, Neurospora crassa, Chlamydomonas reinhardtii, and the like. In some embodiments, the eukaryotic host cell is a yeast cell and the yeast cell strain is SBY49. In some embodiments, the eukaryotic host cell is a human cell. Exemplary human cells lines include, without limitation, HEK293T (ATCC), FreeStyle™ 293-F (Thermo Fisher), and Expi293F™ GnTI- (Thermo Fisher).
In accordance with the present disclosure, the host cell may be prokaryotic, such as a bacterial cell. Such cells serve as a host for expression of recombinant proteins for production of recombinant therapeutic proteins of interest. Suitable microorganisms include Pseudomonas sp. such as Pseudomonas aeruginosa, Escherichia sp., Escherichia coli and other Enterobacteriaceae, Salmonella sp. such as Salmonella gastroenteritis (typhimirium), S. typhi, S. enteriditis, Shigella sp. such as Shigella flexneri, S. sonnie, S dysenteriae, Neisseria sp. such as Neisseria gonorrhoeae, N. meningitides, Haemophilus sp. including Haemophilus influenzae H. pleuropneumoniae, Pasteurella sp. including Pasteurella haemolytica, P. multilocida, Legionella sp. such as Legionella pneumophila, Treponema pallidum, T. denticola, T. orales, Borrelia burgdorferi, Borrelia spp. Leptospira interrogans, Klebsiella sp. such as Klebsiella pneumoniae, Proteus vulgaris, P. morganii, P. mirabilis, Rickettsia prowazeki, R. typhi, R. richettsii, Porphyromonas (Bacteroides) gingivalis, Chlamydia psittaci, C. pneumoniae, C. trachomatis, Campylobacter sp. such as Campylobacter jejuni, C. intermedis, C. fetus, Helicobacter sp. such as Helicobacter pylori, Francisella sp. such as Francisella tularenisis, Vibrio cholerae, Vibrio parahaemolyticus, Bordetella sp. including Bordetella pertussis, Burkholderia sp. such as Burkholderie pseudomallei, Brucella sp. including Brucella abortus, B. susi, B. melitens is, B. canis, Spirillum minus, Pseudomonas mallei, Aeromonas sp. such as Aeromonas hydrophila, A salmonicida, and Yersinia sp. such as Yersinia pestis. Additional microorganisms include Wolinella sp., Desulfovibrio sp. Vibrio sp., Bacillus sp., Listeria sp., Staphylococcus sp., Streptococcus sp., Peptostreptococcus sp., Megasphaera sp., Pectinatus sp., Selenomonas sp., Zymophilus sp., Actinomyces sp., Arthrobacter sp., Frankia sp., Micromonospora sp., Nocardia sp., Propionibacterium sp., Streptomyces sp., Lactobacillus sp., Lactococcus sp., Leuconostoc sp., Pediococcus sp., Acetobacterium sp., Eubacterium sp., Heliobacterium sp., Heliospirillum sp., Sporomusa sp., Spiroplasma sp., Ureaplasma sp., Erysipelothrix, sp., Corynebacterium sp. Enterococcus sp., Clostridium sp., Mycoplasma sp., Mycobacterium sp., Actinobacteria sp., Moraxella sp., Stenotrophomonas sp., Micrococcus sp., Bdellovibrio sp., Hemophilus sp., Proteus mirabilis, Enterobacter cloacae, Serratia sp., Citrobacter sp., Proteus sp., Acinetobacter sp., Actinobacillus sp., Capnocytophaga sp., Cardiobacterium sp., Eikenella sp., Kingella sp., Flavobacterium sp. Xanthomonas sp., Plesiomonas sp., and alpha-proteobacteria such as Wolbachia sp., cyanobacteria, spirochaetes, green sulfur and green non-sulfur bacteria, Gram-negative cocci, Gram negative bacilli which are fastidious, Enterobacteriaceae-glucose-fermenting gram-negative bacilli, Gram negative bacilli—non-glucose fermenters, Gram negative bacilli—glucose fermenting, oxidase positive. In some embodiments, the prokaryotic host cells is an E. coli cells such as DH5a, BL21 (DE3), SHuffle® T7 Express lysY, and Origami2(DE3) gmd::kan ΔwaaL.
Methods for transforming/transfecting host cells with expression vectors are well-known in the art and depend on the host system selected as described in Sambrook et al., Molecular Cloning: A Laboratory Manual, Cold Springs Laboratory Press, Cold Springs Harbor, New York (1989), which is hereby incorporated by reference in its entirety.
The present disclosure is also directed to tripartite glycosyltransferase fusion proteins produced by the host cells of the present disclosure.
When the nucleic acid construct is assembled in a host cell, the host cell may be cultured in a suitable culture medium optionally supplemented with one or more additional agents, such as an inducer (e.g., where a nucleotide sequence encoding a chimeric protein is under the control of an inducible promoter). The inducer may be, for example, isopropyl-β-
In some embodiments of the present disclosure, the tripartite glycosyltransferase fusion protein is separated from other products, macromolecules, etc., which may be present in the cell culture medium, the cell lysate, or the organic layer. Separation of the tripartite glycosyltransferase fusion protein from other products that may be present in the cell culture medium, cell lysate, or organic layer is readily achieved using standard methods known in the art, e.g., standard chromatographic techniques. Several methods are readily known in the art, including ion exchange chromatography, high performance liquid chromatography, hydrophobic interaction chromatography, affinity chromatography (e.g., Ni2+ affinity chromatography), size exclusion chromatography, gel filtration, and reverse phase chromatography. The tripartite glycosyltransferase fusion protein is preferably produced in purified form (at least about 40% pure, at least about 50% pure, at least about 60% pure, at least about 70% pure, at least about 80% pure, at least about 90% pure, at least about 95% pure, at least about 98%, or more than 98% pure) by conventional techniques. Depending on whether the host cell is made to secrete the protein into growth medium (see U.S. Pat. No. 6,596,509 to Bauer et al., which is hereby incorporated by reference in its entirety), the protein can be isolated and purified by centrifugation (to separate cellular components from supernatant containing the secreted protein) followed by sequential ammonium sulfate precipitation of the supernatant. The fraction containing the protein can be subjected to gel filtration in an appropriately sized dextran or polyacrylamide column to separate the protein from other cellular components and proteins. If necessary, the protein fraction may be further purified by HPLC. Accordingly, the tripartite glycosyltransferase fusion protein produced by the present disclosure can be used to isolate and solubilize a glycosyltransferase in a purified form, e.g., “pure” in the context of a tripartite glycosyltransferase fusion protein that is free from other intermediate or precursor products, macromolecules, contaminants, etc.
Expression of soluble tripartite glycosyltransferase fusion proteins may be increased by at least about 10%, at least about 20%, at least about 25%, at least about 30%, at least about 40%, at least about 50%, at least about 60%, at least about 70%, at least about 80%, at least about 90%, at least about 100% (or two-fold), as compared to when the corresponding glycosyltransferase is expressed in the absence of the amphipathic shield domain protein and/or water soluble expression decoy protein. In other embodiments of the present disclosure, the expression of tripartite glycosyltransferase fusion proteins from the nucleic acid constructs disclosed herein is at least about 2.5-fold, at least about 3-fold, at least about 5-fold, at least about 7-fold, at least about 10-fold, at least about 15-fold, at least about 20-fold, at least about 50-fold, at least about 100-fold, or more, higher compared to the expression level of the glycosyltransferase from nucleic acid constructs lacking the first nucleic acid moiety encoding an amphipathic shield domain protein and/or the third nucleic acid moiety encoding a water soluble expression decoy protein. Likewise, the expression of tripartite glycosyltransferase fusion proteins from the nucleic acid constructs disclosed herein may be at least about 2.5-fold, at least about 3-fold, at least about 5-fold, at least about 7-fold, at least about 10-fold, at least about 15-fold, at least about 20-fold, at least about 50-fold, at least about 100-fold, or more, higher compared to the expression of a corresponding wild type glycosyltransferase protein, which is not fused to a heterologous amphipathic shield domain protein and/or a water soluble expression decoy protein.
Methods for transforming/transfecting host cells with expression vectors are well-known in the art and depend on the host system selected, as described in Sambrook et al., Molecular Cloning: A Laboratory Manual, Cold Springs Laboratory Press, Cold Springs Harbor, New York (1989), which is hereby incorporated by reference in its entirety. For eukaryotic cells, suitable techniques may include calcium phosphate transfection, DEAE-Dextran, electroporation, liposome-mediated transfection and transduction using retrovirus or other virus, e.g. vaccinia or, for insect cells, baculovirus. For bacterial cells, suitable techniques may include calcium chloride transformation, electroporation, and transfection using bacteriophage.
The simplest single-celled organisms are composed of central regions filled with an aqueous material and a variety of soluble small molecules and macromolecules. Enclosing this central region is a membrane which is composed of phospholipids arranged in a bilayer structure. In more complex living cells, there are internal compartments and structures that are also enclosed by membranes. There are many protein molecules embedded or associated within these membrane structures, and these membrane proteins are often the most important to determining cell functions including communication and processing of information and energy. The largest problem in studying membrane proteins is that the inside of the phospholipid bilayer is hydrophobic and the embedded or anchored part of the membrane protein is itself also hydrophobic. In isolating these membrane proteins from their native membrane environments, the present disclosure overcomes the difficult task of preventing recombinant glycosyltransferases from forming inactive aggregates while remaining in a native configuration. In one embodiment of the present disclosure, a tripartite glycosyltransferase fusion protein is encoded by the nucleic acid construct of the present disclosure and, preferably, the tripartite glycosyltransferase fusion protein is in water soluble form. The term “solubilizing” according to the present disclosure includes dissolving a molecule in a solution. This aspect of the disclosure is carried out in substantially the same way as described above.
The present disclosure is also directed to tripartite glycosyltransferase fusion proteins produced by the host cells of the present disclosure.
In addition to cell-based expression hosts/systems of the present disclosure, the tripartite glycosyltransferase fusion proteins may also be expressed using cell-free expression platforms. Thus, another aspect of the present disclosure relates to a cell-free protein expression system. The cell-free protein expression system comprises a cell lysate or extract and a nucleic acid construct according to the present disclosure. The cell lysate or extract may include a heterologous and/or recombinant RNA polymerase. In some embodiments, the cell lysate or extract is capable of (i) transcribing the nucleic acid construct or the vector to form a translation template and (ii) translating the translation template. In some embodiments, the cell lysate or extract is an E. coli lysate or extract. Examples of cell-free expression platforms include, but are not limited to, the PURExpress kit from NEB and S30 lysate high-expression kit from Promega, among others.
The present disclosure is also directed to tripartite glycosyltransferase fusion proteins produced by the cell-free protein expression systems of the present disclosure.
Another aspect of the present disclosure relates to a method of recombinantly producing a tripartite glycosyltransferase fusion protein in water soluble form. This method involves providing a host cell according to the present disclosure or a cell-free expression system according to the present disclosure. The method further involves culturing the host cell or using the cell-free expression system under conditions effective to express the tripartite glycosyltransferase fusion protein in a water soluble form within the host cell cytoplasm or the cell-free expression system.
In some embodiments, the method further includes recovering the tripartite glycosyltransferase fusion protein from the host cell or the cell-free expression system following the culturing or the using, respectively. The tripartite glycosyltransferase fusion protein may be recovered from the cell's cytoplasm. The recovery of the tripartite glycosyltransferase fusion protein from the host cell is consistent with the recovery of proteins discussed supra.
In some embodiments where the host cell is provided, the recovering involves lysing the cell to form a cell lysate comprising a water soluble fraction and subjecting the water soluble fraction of the cell lysate to chromatography to isolate the tripartite glycosyltransferase fusion protein.
In some embodiments where the cell-free expression system is provided, the recovering involves subjecting the water soluble fraction of the cell lysate to chromatography to isolate the tripartite glycosyltransferase fusion protein.
In one embodiment of this aspect of the present disclosure, the tripartite glycosyltransferase fusion proteins are provided in a purified isolated form.
The tripartite glycosyltransferase fusion protein can be synthesized using standard methods of protein/peptide synthesis known in the art, including solid phase synthesis or solution phase synthesis. Alternatively, the tripartite glycosyltransferase fusion proteins can be generated using recombinant expression systems and purified using any method readily known in the art, including ion exchange chromatography, hydrophobic interaction chromatography, affinity chromatography, gel filtration, and reverse phase chromatography.
Nucleotide sequences encoding the tripartite glycosyltransferase fusion proteins may be modified such that the nucleotide sequence reflects the codon preference for a particular host cell. For example, when yeast host cells are utilized, the nucleotide sequences encoding the chimeric proteins can be modified for yeast codon preference (see, e.g., Bennetzen and Hall, “Codon Selection in Yeast,” J. Biol. Chem. 257(6):3026-3031 (1982), which is hereby incorporated by reference in its entirety). Likewise, when bacterial host cells are utilized, e.g., E. coli cells, the nucleotide sequences encoding the chimeric biological pathway proteins can be modified for E. coli codon preference (see e.g., Gouy and Gautier, “Codon Usage in Bacteria: Correlation With Gene Expressivity,” Nucleic Acids Res. 10(22):7055-7074 (1982); Eyre-Walker et al., “Synonymous Codon Bias is Related to Gene Length in Escherichia coli: Selection for Translational Accuracy?,” Mol. Biol. Evol. 13(6):864-872 (1996) and Nakamura et al., “Codon Usage Tabulated From International DNA Sequence Databases: Status for the year 2000,” Nucleic Acids Res. 28(1):292 (2000), which are hereby incorporated by reference in their entirety).
A variety of genetic signals and processing events that control many levels of gene expression (e.g., DNA transcription and messenger RNA (“mRNA”) translation) can be incorporated into the nucleic acid construct encoding the chimeric proteins to maximize protein production. For the purpose of expressing a cloned nucleic acid sequence encoding the desired tripartite glycosyltransferase fusion protein, it is advantageous to use strong promoters to obtain a high level of transcription. Depending upon the host system utilized, any one of a number of suitable promoters may be used. For instance, when cloning in E. coli, its bacteriophages, or plasmids, promoters such as the T7 phage promoter, lac promoter, trp promoter, recA promoter, ribosomal RNA promoter, the PR and PL promoters of coliphage lambda and others, including but not limited, to lacUV5, ompF, bla, lpp, and the like, may be used to direct high levels of transcription of adjacent DNA segments. Additionally, a hybrid trp-lacUV5 (tac) promoter or other E. coli promoters produced by recombinant DNA or other synthetic DNA techniques may be used to provide for transcription of the inserted chimeric genetic construct. Common promoters suitable for directing expression in mammalian cells include, without limitation, SV40, MMTV, metallothionein-1, adenovirus Ela, CMV, immediate early, immunoglobulin heavy chain promoter and enhancer, and RSV-LTR. Common promoters suitable for directing expression in a yeast cell include constitutive promoters such as an ADH1 promoter, a PGK1 promoter, an ENO promoter, a PYK1 promoter and the like; or a regulatable promoter such as a GAL1 promoter, a GAL10 promoter, an ADH2 promoter, a PHO5 promoter, a CUP1 promoter, a GAL7 promoter, a MET25 promoter, a MET3 promoter, a CYC1 promoter, a HIS3 promoter, a PGK promoter, a GAPDH promoter, an ADC1 promoter, a TRP1 promoter, a URA3 promoter, a LEU2 promoter, an ENO promoter, a TP1 promoter, and a AOX1 promoter.
There are other specific initiation signals required for efficient gene transcription and translation in eukaryotic and prokaryotic cells that can be included in the nucleic acid construct to maximize chimeric protein production. Depending on the vector system and host utilized, any number of suitable transcription and/or translation elements, including constitutive, inducible, and repressible promoters, as well as minimal 5′ promoter elements, enhancers, or leader sequences may be used. For a review on maximizing gene expression see Roberts and Lauer, “Maximizing Gene Expression On a Plasmid Using Recombination In Vitro,” Methods in Enzymology 68:473-82 (1979), which is hereby incorporated by reference in its entirety.
A nucleic acid molecule encoding a tripartite glycosyltransferase fusion protein of the present disclosure, a promoter molecule of choice, including, without limitation, enhancers, and leader sequences; a suitable 3′ regulatory region to allow transcription in the host, and any additional desired components, such as reporter or marker genes, are cloned into a vector of choice using standard cloning procedures in the art, such as described in Sambrook et al., M
In some embodiments, the recovered tripartite glycosyltransferase fusion protein is conformationally correct.
Another aspect of the present disclosure relates to a tripartite glycosyltransferase fusion protein produced by the methods of recombinantly producing a tripartite glycosyltransferase fusion protein according to the present disclosure.
As will be apparent to one of skill in the art, the present disclosure allows for a broad range of in vivo or in vitro glycan remodeling. The constructs of the present disclosure allow for solubilized tripartite glycosyltransferase fusion proteins for use in methods of in vivo or in vitro glycan remodeling. Accordingly, another aspect of the present disclosure relates to a method of cell-free glycan remodeling. This method involves providing a glycan primer; providing one or more tripartite glycosyltransferase fusion protein(s) according to the present disclosure; and incubating the glycan primer with the one or more tripartite glycosyltransferase fusion protein(s) under conditions effective to transfer a glycosyl group to the glycan primer to produce a modified glycan structure.
The glycan primer may be a monosaccharide or an oligosaccharide. For example, the glycan primer may comprise Man3GlcNAc2 or Man5GlcNAc2.
In some embodiments, the glycan primer is attached to an amino acid residue such as an asparagine residue. In some embodiments, the glycan primer is attached to a protein. Accordingly, the glycan primer may be attached to a glycoprotein. The glycoprotein may comprise an N-glycosidic linkage. For example, the glycoprotein may comprises an N-acetylglucosamine (GlcNAc) linkage to asparagine.
The glycoprotein may be selected from the group consisting of an antibody or a hormone.
In some embodiments, the glycoprotein comprises an O-glycosidic linkage.
Suitably tripartite glycosyltransferase fusion proteins are described in detail supra. In some embodiments, the glycosyltransferase fusion protein is selected from the group consisting of Sx-Δ29HsGnTI, Sx-Δ29HsGnTII, Sx-Δ30HsFucT8, Sx-Δ44Hsβ4GalT1, Sx-Δ26HsST6Gal1, and combinations thereof.
In some embodiments, when the incubating step is carried out with a plurality of different tripartite glycosyltransferase fusion proteins, at least some of the different tripartite glycosyltransferase proteins being used sequentially during said incubating. In accordance with such embodiments, the incubating step produces a modified glycan primer. In some embodiments, the method may further involve incubating a modified glycan primer with one or more glycosyl hydrolases. In accordance with such embodiments, the one or more hydrolases may be used sequentially during said further incubating.
In some embodiments, when the incubating step is carried out with a plurality of different tripartite glycosyltransferase fusion proteins, at least some of the different tripartite glycosyltransferase proteins being used simultaneously during said incubating.
The above disclosure generally describes the present disclosure. A more specific description is provided below in the following examples. The examples are described solely for the purpose of illustration and are not intended to limit the scope of the present disclosure. Changes in form and substitution of equivalents are contemplated as circumstances suggest or render expedient. Although specific terms have been employed herein, such terms are intended in a descriptive sense and not for purposes of limitation.
The bacterial, yeast, and mammalian cells used in Examples 1-9 are listed in
To facilitate high-throughput cell growth measurements, three individual colonies corresponding to each construct were seeded into 96-deep well plates (Eppendorf) where each well contained 100 μL LB media. Culture plates were then sealed using plate sealer and placed in an incubator shaker at 37° C. for 16 hours. Then, 5 μL of the overnight culture was subcultured into fresh 100 μL LB media and incubated for 8 hours, after which IPTG was supplemented to a final concentration of 0.1 mM. Protein expression proceeded at 16° C. for 18 hours. To measure OD600, 10 μL of each sample was mixed with 90 μL DI water in a Costar 96-well assay plate (Corning) and OD600 of all samples was measured in an Infinite M1000Pro spectrophotometer (Tecan).
All plasmids used in this study are listed in
Plasmids encoding Sx-GT and unfused GT constructs were used to transform either E. coli strain BL21(DE3) for GTs containing no disulfide bonds or SHuffle T7 Express lysY for GTs contain predicted or confirmed to contain disulfide bonds. Small 5-mL LB cultures of E. coli harboring either a Sx-GT or GT plasmid were grown to an optical density at 600 nm (OD600) of approximately 0.6-0.8 and then induced with IPTG to a final concentration of 0.1 mM. Protein expression proceeded for 18 hours at 16° C., after which culture volumes equivalent to OD600 of 2.0 were harvested. Media was removed by centrifugation and the resulting cell pellet was resuspended in 1 mL phosphate buffer saline (PBS). Cells were lysed using a Q125 Sonicator (Qsonica) with a 3.175-mm diameter probe at a frequency of 20 kHz and 40% amplitude. Lysate was first centrifuged at 15,000×g for 30 minutes at 4° C. Supernatant was collected and centrifuged at 100,000×g for 1 hour at 4° C. The supernatant from this ultracentrifugation step was collected as the soluble fraction. Pellet was then resuspended in 1 mL PBS containing 1% (v/v) Triton X-100. The suspension was incubated for 1 hour at 4° C. to allow partitioning of membrane proteins into Triton X-containing buffer. Following ultracentrifugation at 100,000×g for 1 hour at 4° C., supernatant was collected as the detergent-solubilized fraction, while the pellet was taken as the insoluble fraction.
A single colony of E. coli harboring plasmid DNA encoding a specific glycoenyzme was selected from a transformation plate and grown overnight in LB media at 37° C. The next day, cells were subcultured 5% into 1 L of fresh LB media. Cells were grown at 37° C. until OD600 reached approximately 0.6-0.8, after which IPTG was supplemented into culture at 0.1 mM final concentration. Protein expression proceeded at 16° C. for 18 hours. Unless otherwise noted, all purification procedures were performed at 4° C. Cells were harvested, resuspended in PBS supplemented with 10% (v/v) glycerol, and lysed by passing the cell suspension through an Emulsiflex C5 homogenizer (Avestin) twice at 15,000 psi maximum pressure. Supernatant was collected following centrifugation at 15,000×g for 30 minutes and then incubated with 300 μL pre-washed HisPur™ Ni-NTA resin (Thermo Fisher Scientific) at 4° C. for 1 hour. The suspension was loaded onto an Econo-Pac© gravity flow chromatography column (Bio-Rad) and resin was washed with 6 column volumes HisPur wash buffer (50 mM NaH2PO4, 300 mM NaCl, 10 mM imidazole, pH 8.0). The target protein was eluted with HisPur elusion buffer (50 mM NaH2PO4, 300 mM NaCl, 300 mM imidazole, pH 8.0). Sample was then buffer exchanged into PBS using Zeba spin desalting columns, 7K MWCO (Thermo Fisher Scientific). Protein concentration was determined using Bradford assay (Bio-Rad). Purified protein fractions were subjected to standard Coomassie-blue staining of SDS-PAGE gels and purity of each was determined by densitometry analysis using BioRad Image Lab software (version 6.1.0 build 7), whereby the intensity of the band corresponding to the full-length Sx-GT construct was normalized to the intensity of all bands that appeared in the same lane of the gel. In general, purity of isolated Sx-GTs was approximately 50-80% following just a single-step Ni-NTA purification. Final yield values were tabulated based on both total protein concentration and purity, and were representative of three biological replicates starting from freshly transformed cells.
All other purification was performed as described above but with amylose resin (NEB) instead of Ni-NTA resin. Clarified lysate was incubated with 300 μL pre-washed amylose resin with rotation for 2 hours at 4° C. The suspension was loaded onto an Econo-Pac© gravity column (Bio-Rad) and resin was washed with 6 column volumes of amylose column buffer (20 mM Tris-HCl, 200 mM NaCl, 1 mM EDTA, pH 7.4). The target protein was eluted with amylose elusion buffer (10 mM maltose in column buffer). Protein purity and concentration were determined by Coomassie staining and Bradford assay (both from Bio-Rad), respectively. Proteins were kept at 4° C. for 2 weeks. For longer term storage at −80° C., protein solution was supplemented with 10% (v/v) glycerol and 0.02% (w/v) sodium azide as a cryogenic agent and bacteriostat, respectively.
For human MAN2A1 expression and purification, an expression construct encoding the truncated catalytic domain of human MAN2A1 (UniProt Q16706, residues 27-1144) was used (Moremen Et Al., “Expression System For Structural And Functional studies of Human Glycosylation Enzymes,” Nat. Chem. Biol. 14:156-162 (2018), which is hereby incorporated by reference in its entirety). This recombinant human MAN2A1 construct was expressed by transient transfection of suspension culture HEK293F cells, with soluble recombinant human MAN2A1 expressed as a soluble secreted product that was purified as described (Kadirvelraj et al., “Human N-acetylglucosaminyltransferase II Substrate Recognition Uses a Modular Architecture That Includes a Convergent Exosite,” Proc. Natl. Acad. Sci. USA 115:4637-4642 (2018), which is hereby incorporated by reference in its entirety). Briefly, the conditioned culture medium was loaded on a Ni2+-NTA Superflow column (Qiagen) equilibrated with 20 mM HEPES, 300 mM NaCl, 20 mM imidazole, pH 7.4, washed with column buffer, and eluted successively with column buffers containing stepwise increasing imidazole concentrations (40-300 mM). The eluted fusion protein was pooled, concentrated, and concurrently mixed with recombinant TEV protease and EndoF1 at ratios of 1:10 relative to the GFP-MGAT2 for each enzyme, respectively, and incubated at 4° C. for 36 hours to cleave the tag and glycans. Dilution to lower the imidazole concentration was followed by passing the sample through a Ni2+-NTA column to remove the fusion tag and His-tagged TEV protease and EndoF1. The protein was further purified on a Superdex 75 gel filtration column (GE Healthcare) and peak fractions of MGAT2 were collected. The protein buffer was exchanged by ultrafiltration and adjusted to 1 mg/mL with buffer containing 20 mM HEPES, 100 mM NaCl, pH 7.0, 0.05% sodium azide, and 10% glycerol and stored at −80° C. until use.
For antibody expression and purification, glycoengineered HEK293F GnTI− cells were used as follows. After at least three passages, cells were washed and resuspended at 3 million cells per mL concentration. Plasmid pVITRO1-Trastuzumab-IgG1/κ (Addgene #61883) was prepared from E. coli culture and the purified plasmid was flowed through an endotoxin removal column to remove contaminating endotoxin. Plasmid DNA-cationic lipid complex was then generated using Lipofectamine™ Transfection Reagent (Thermo Fisher Scientific) and was slowly added into the culture media with gentle mixing. The amount of DNA, cationic-lipid reagents, and cells were scaled linearly according to the manufacturer's protocol. Cells were maintained in a 37° C. incubator shaker for 24 hours prior to being supplemented with Expression Enhancer Reagents (Thermo Fisher Scientific). Cell cultures were maintained at the same condition for another 5 days to allow antibody accumulation in the culture supernatant. Cells were then removed by centrifugation at 1,000×g for 5 minutes and supernatant was filtered through a 0.2-micron bottle-top filter. Supernatant was then mixed with 1×PBS at a 1:1 (v/v) ratio. This solution was flowed through MabSelect SuRe resin (Sigma-Aldrich) twice to allow antibody capture on protein A/G beads. Following extensive washing with 1×PBS, captured antibodies were eluded using glycine solution (pH 2.0) directly into neutralizing buffer (Tris-HCl pH 8.5). The antibody product was then buffer exchanged into 1×PBS supplemented with 0.01% sodium azide. Antibody was stored at 4° C. and was stable at the described conditions for at least a month.
Prior to electrophoretic separation, samples were combined with NuPAGE™ 4× LDS Sample Buffer (Invitrogen) supplemented with 2.5% β-mercaptoethanol and then boiling at 100° C. for 10 minutes. Samples equivalent to OD600 of 0.375 for small-scale expression or 15 μL of CFPS reaction were loaded into each well of Bolt™ 8% Bis-Tris Plus Gels (Thermo Fisher Scientific). Following electrophoretic separation and transfer to Immobilon-P polyvinylidene difluoride (PVDF) membranes (0.45 m), blots were washed with TBS buffer (80 g/L NaCl, 20 g/L KCl, and 30 g/L Tris-base) followed by a 1-hour incubation in blocking solution (50 g/L non-fat milk in TBS supplemented with 0.05% (v/v %) Tween-20; TBST). Blots were then washed 4 times with TBST in 10-minute intervals and probed with primary antibodies including rabbit polyclonal antibody to 6×His epitope tag (Thermo Fisher Scientific; Cat #PA1-983B; 1:5,000 dilution), mouse monoclonal anti-GAPDH clone 6C5 (Calbiochem; Cat #CB1001; 1:10,000 dilution), rabbit polyclonal anti-GroEL (Sigma-Aldrich; Cat #G6532; 1:20,000 dilution), and rabbit anti-alpha tubulin clone EPR13799 (Abcam; Cat #ab184970; 1:10,000 dilution). Secondary antibodies were used as needed and these include goat anti-rabbit IgG H&L (HRP) (Abcam; Cat #ab6721; 1:5,000 dilution), rabbit anti-mouse IgG H&L (HRP) (Abcam; Cat #ab6728; 1:5,000 dilution), and ExtrAvidin®-Peroxidase (Sigma-Aldrich; Cat #E2886; 1:4,000 dilution). Blots were then washed as above. Imaging of blots was performed using a ChemiDoc™ XRS+ System following a brief incubation with Western ECL substrate (Bio-Rad).
Kinetic analysis of sialytransferases was performed using a commercial sialytransferase activity kit (R&D Systems, Cat #EA002) according to manufacturer's protocols. Briefly, assays used 2 g/mL of purified Sx-Δ26HsST6Gal1 or commercial human ST6Gal1 (amino acids 44-406) (R&D Systems; Cat #7620-GT-010), 1.0 mg/mL of asialofetuin (Sigma-Aldrich; Cat #A4781-50MG) as acceptor substrate, and 0.02-0.8 mM of CMP-Neu5Ac as donor substrate. All reactions were incubated for 15 minutes at 37° C. Values for Vmax and Km were determined using Prism 9 for MacOS version 9.2.0. A conversion factor used for calculating the amount of enzymatically released inorganic phosphate from CMP-Neu5Ac was determined to be 3,833.5 pmol/OD620 using the phosphate standards included in the kit and was used for all data analysis. Specific activity was calculated using 0.1 mM of CMP-Neu5Ac, 1.0 mg/mL of asialofetuin, and 0.04-0.23 g of Sx-Δ26HsST6Gal1. A linear plot of absorbance (OD620) versus amount of Sx-Δ26HsST6Gal1 was generated (
Strain-promoted alkyne-azide cycloaddition was used to assess the ability of Sx-GTs to chemoenzymatically remodel glycoprotein substrates. In a typical reaction, a 1.5-mL microcentrifuge tube was charged with 20 μL of reaction mixture consisting of 1 μg purified Sx-GT or 50 μg cell lysate, 3 μg purified acceptor glycoprotein substrate, and 10 mM nucleotide-activated monosaccharide donor modified with an azide functional group. Depending on the GT reactions, the nucleotide-activated monosaccharide donors included UDP-GlcNAz, UDP-GalNAz, GDP-AzFuc, and CMP-AzNeu5Ac (all from R&D Systems). Following an incubation in a 37° C. water bath for 1 hour, reaction mixtures were supplemented with 2-iodoacetamide (Sigma-Aldrich) at 100 mM final concentration and incubated in the dark at room temperature for 1 hour. Then, 100 mM final concentration of carboxyrhodamine 110 or biotin(PEG)4 conjugated dibenzocyclooctyne-amines (Click Chemistry Tools) in N,N-dimethylformamide (DMF) was supplemented into the reaction mixture. Strain-promoted click reactions were carried out at 37° C. for 2 hours. Samples were then combined with 4×LDS Sample Buffer (Invitrogen) supplemented with 2.5% β-mercaptoethanol and heated at 65° C. for 5 minutes. Following SDS-PAGE analysis, in-gel fluorescence from carboxyrhodamine110-linked glycans on glycoproteins was measured using a ChemiDoc™ MP Imaging System (Bio-Rad) with 501/523 nm λex/λem. Biotin-linked glycans on glycoproteins were analyzed following immunoblot analysis using horseradish peroxidase conjugated streptavidin (Sigma-Aldrich) in a similar manner as described above for immunoblot analysis.
E. coli lysate was prepared according to an established protocol (Kwon and Jewett, “High-throughput Preparation Methods of Crude Extract for Robust Cell-free Protein Synthesis,” Scientific Reports 5:8663 (2015), which is hereby incorporated by reference in its entirety). Briefly, E. coli strain BL21(DE3) was cultured in 2×YTPG media (16 g/L tryptone, 10 g/L yeast extract, 5 g/L NaCl, 7 g/L potassium phosphate monobasic, 3 g/L potassium phosphate dibasic and 18 g/L glucose) at 37° C. with 0.5 mM IPTG until OD600 reached approximately 1.0. Cells were then harvested and washed twice with cold S30 buffer (10 mM tris-acetate pH 8.2, 14 mM magnesium acetate and 60 mM potassium acetate). The resulting pellet was stored at −80° C. until used. To prepare crude extract, pellets were thawed on ice and resuspended with S30 buffer (1 mL per gram cell pellet). Cells were lysed using a Q125 Sonicator with a 3.175-mm diameter probe at a frequency of 20 kHz and 40% amplitude until the total energy input reached 1500 J. Lysate was then centrifuged twice at 30,000×g at 4° C. for 30 minutes. Supernatant was then collected, aliquoted, and stored at −80° C. until used. Cell-free synthesis of Sx-GT and unfused GT constructs was performed using the modified PANOx-SP system (Jewett and Swartz, “Mimicking the Escherichia coli Cytoplasmic Environment Activates Long-lived and Efficient Cell-free Protein Synthesis,” Biotechnology and Bioengineering 86:19-26 (2004), which is hereby incorporated by reference in its entirety). Specifically, S30 lysate was pre-conditioned with 750 M iodoacetamide in the dark at room temperature for 30 minutes and then lysate was supplemented with 200 mM glutathione at a 3:1 ratio between oxidized and reduced forms. Then, 200 ng plasmid DNA was introduced into cell-free protein synthesis reaction containing 30% (v/v) S30 lysate and the following: 12 mM magnesium glutamate, 10 mM ammonium glutamate, 130 mM potassium glutamate, 1.2 mM adenosine triphosphate (ATP), 0.85 mM guanosine triphosphate (GTP), 0.85 mM uridine triphosphate (UTP), 0.85 mM cytidine triphosphate (CTP), 0.034 mg/mL folinic acid, 0.171 mg/mL E. coli tRNA (Roche), 2 mM each of 20 amino acids, 30 mM phosphoenolpyruvate (PEP, Roche), 0.33 mM nicotinamide adenine dinucleotide (NAD), 0.27 mM coenzyme-A (CoA), 4 mM oxalic acid, 1 mM putrescine, 1.5 mM spermidine, and 57 mM HEPES. The synthesis reaction was carried out at 30° C. for 6 hours, after which the sample was centrifuged at 15,000×g for 30 minutes at 4° C. Supernatant was collected and stored at −20° C. until further analysis.
Yeast cells were transformed with plasmid pYS338 encoding Δ26HsST6Gal1 using the LiAc/single stranded carrier DNA/PEG method (Gietz and Schiestl, “High-efficiency Yeast Transformation Using the LiAc/SS Carrier DNA/PEG Method,” Nat. Protoc. 2:31-4 (2007), which is hereby incorporated by reference in its entirety). For yeast expression, SBY49 cells were grown in-URA media at 30° C. until OD600 reached approximately 0.6-0.8, after which protein expression was induced with galactose to a final concentration of 2% (w/v). Protein expression was performed for 22 hours at 30° C. Yeast cells were lysed by vortexing the cell suspension with glass beads in PBS containing zymolyase enzyme. For mammalian cell expression, 2.0 mL of HEK293T cells at approximately 80% confluency in a 6-well plate were transfected with 2 g plasmid DNA using jetPRIME® transfection reagent (Polyplus Transfection). After transfection, cells were maintained in an incubator at 37° C. with 5% CO2 and 90% relative humidity for 36 hours, after which they were harvested. HEK293T cells were lysed by tip sonication. Subcellular fractionation analysis for yeast and HEK293T cells was performed similarly as described above. All samples were stored at −20° C. until further analysis.
All glycans and nucleotide-activated sugar substrate solutions were prepared in sterile DI water and stored at −20° C. Glycan 1 was prepared as described (Hamilton et al., “A Library of Chemically Defined Human N-glycans Synthesized From Microbial Oligosaccharide Precursors,” Sci. Rep. 7:15907 (2017), which is hereby incorporated by reference in its entirety). Briefly, dried cell pellets from a 250-mL culture of E. coli Origami2(DE3) gmd::kan ΔwaaL cells carrying plasmid pConYCGmCB (Glasscock et al., “A Flow Cytometric Approach to Engineering Escherichia coli for Improved Eukaryotic Protein Glycosylation,” Metab. Eng. 47:488-495 (2018), which is hereby incorporated by reference in its entirety) were resuspended in 2:1 chloroform: methanol, sonicated, and the remaining solids collected by centrifugation. This pellet was sonicated in water and collected by centrifugation. The resulting pellet was sonicated in 10:10:3 chloroform: methanol:water to isolate the lipid-linked oligosaccharides (LLOs) from the inner membrane. The LLOs were purified using acetate-converted DEAE anion exchange chromatography as they bind to the anion exchange resin via the phosphates that link the lipid and glycan. The resulting compound was dried and treated by mild acid hydrolysis to release glycans from the lipids. The released glycans were then separated from the lipid by a 1:1 butanol:water extraction, wherein the water layer contains the glycans. The glycans were then further purified with a graphitized carbon column using a 0-50% water: acetonitrile gradient. Following this procedure, approximately 750 g of glycan 1 that was well resolved from contaminant peaks was reproducibly obtained (
Unless noted otherwise, all glycoprotein remodeling reactions were performed at 37° C. for 1 hour prior to bioorthogonal labeling reaction as described above. The sialytransferase activity of Sx-CjCstII was assessed using human A1AT as glycoprotein acceptor substrate. A total of 3 g of recombinant A1AT (R&D Systems) was treated with 20 U/L a2-3,6,8,9 neuraminidase A (NEB) in a 10-μL reaction at 37° C. for 2 hours to remove terminal sialic acid residues on A1AT glycans. Reaction mixtures were then heated at 85° C. for 15 minutes to inactivate neuraminidase A. Neuraminidase A-treated A1AT was then incubated with Sx-CjCstII and CMP-AzNec5Ac in SiaT buffer in a 37° C. water bath for 1 hour. Sialyltransferase activity of Sx-Δ34HsST3Gal1 was evaluated in a similar manner but neuraminidase-treated bovine submaxillary glands mucin (Sigma-Aldrich) was used as the glycoprotein substrate. N-acetylglucosaminyltransferase activity of Sx-Δ29HsGnTI was assessed using MBP-GCGDQNAT a fusion between E. coli MBP and human glucagon (residues 1-29) followed by a C-terminal DQNAT glycosylation tag (Glasscock et al., “A Flow Cytometric Approach to Engineering Escherichia coli for Improved Eukaryotic Protein Glycosylation,” Metab. Eng. 47:488-495 (2018), which is hereby incorporated by reference in its entirety). The MBP-GCGDQNAT construct was glycosylated with Man3GlcNAc2 using glycoengineered E. coli as described (Glasscock et al., “A Flow Cytometric Approach to Engineering Escherichia coli for Improved Eukaryotic Protein Glycosylation,” Metab. Eng. 47:488-495 (2018), which is hereby incorporated by reference in its entirety). Briefly, Origami2(DE3) gmd::kan ΔwaaL cells carrying plasmid pConYCGmCB along with plasmid pMAF10 (Feldman et al., “Engineering N-linked Protein Glycosylation With Diverse O Antigen Lipopolysaccharide Structures in Escherichia coli,” Proc Natl Acad Sci USA 102:3016-21 (2005), which is hereby incorporated by reference in its entirety) and pTrc-spDsbA-MBP-GCGDQNAT (Glasscock et al., “A Flow Cytometric Approach to Engineering Escherichia coli for Improved Eukaryotic Protein Glycosylation,” Metab. Eng. 47:488-495 (2018), which is hereby incorporated by reference in its entirety) were grown in 100 mL of LB at 37° C. until OD600 reached ≈1.5. Culture temperature was reduced to 30° C. and allowed to grow overnight at 30° C. The next day, cells were induced with 0.1 mM isopropyl β-D-1-thiogalactopyranoside (IPTG) to initiate synthesis of the MBP-GCGDQNAT acceptor protein. Protein expression proceeded for 8 hours at 30° C. Cells were then harvested and subjected to subcellular fractionation. This involved pelleting and washing 100 mL of IPTG-induced culture with subcellular fractionation buffer (0.2 M Tris-Ac (pH 8.2), 0.25 mM EDTA, and 0.25 M sucrose, and 160 g/mL lysozyme). Cells were resuspended in 1.5 mL subcellular fractionation buffer and then incubated for 5 minutes on ice and spun down. After addition of 60 μL of 1M MgSO4, cells were incubated for 10 minutes on ice. Cells were spun down, and the supernatant was taken as the periplasmic fraction. To isolate glycoproteins, periplasmic fractions were subjected to affinity chromatography using HisPur™ Ni-NTA resin (Thermo Fisher Scientific). Eluates were collected, solubilized in Laemmli sample buffer containing 5% β-mercaptoethanol, and resolved on SDS-polyacrylamide gels. Purified MBP-GCGDQNAT was incubated with Sx-Δ29HsGnTI and UDP-GlcNAz in GnT buffer in a 37° C. water bath for 1 hour. Fucosyltransferase activity was evaluated by incubating A1AT or neuraminidase A-treated A1AT with Sx-Δ36HsFucT7 and GDP-AzFuc in FucT buffer in a 37° C. water bath for 1 hour.
In a sterile Eppendorf microcentrifuge tube, 1 g of purified trastuzumab bearing Man5GlcNAc2 glycan was incubated with: (i) Streptococcus pyogenes Endo S2 (Genovis #AO-GL8-020) in Glycobuffer 1 (NEB #B1727SVIAL); (ii) Elizabethkingia meningosepticum Endo F1 (Sigma-Aldrich #324725) in GlycoBuffer 4 (NEB #B1703); (iii) Elizabethkingia miricola Endo F3 (NEB #P0771S) in GlycoBuffer 4; or (iv) PBS control. Reaction mixtures were incubated at 37° C. for 16 hours and the product was analyzed by LC-MS using intact protein MS mode.
Glycan remodeling on full-length mAb was performed in an on-column mode. 50 g purified trastuzumab bearing Man5GlcNAc2 glycan was first incubated with MabSelect SuRe resin (Sigma-Aldrich) for 10 minutes to allow antibody capture on protein A/G beads. This mixture was then transferred to a spin column, followed by washing twice with PBS. The bottom of the spin column was then capped with rubber cap. In a separate tube, 50 μL of a specific glycan remodeling reaction mixture was prepared. For preparing N-acetylglucosaminyltransferase, galactosyltransferase, fucosyltransferase, and sialyltransferase reaction mix. UDP-GlcNAz substrate was used at the same concentration as UDP-GlcNAc. Reaction using β-N-acetylglucosaminidase S (NEB #P0744S) was performed in Glycobuffer 1 (NEB) at 37° C. for 4 hours. Reactions using human Man2A1 mannosidase were performed in 50 mM sodium acetate buffer (pH 5.5) 1 mM ZnCl2 at 37° C. for 16 hours. Following each reaction step, the reaction mixture was removed by centrifugation at 300×g for 2 minutes. Resin was then washed twice with PBS using the same centrifugation setting. In general, approximately 80-90% recovery yield of IgG was observed following purification as determined by NanoDrop spectrophotometer. Subsequent reaction mixture was then added to the column and the clean-up process was repeated for each reaction step. Final IgG product was eluted using glycine solution (pH 2.0) and analyzed immediately by LC-MS.
Hydrophilic interaction liquid chromatography (HILIC) was carried out using an Exion HPLC system with built-in autosampler (SCIEX). The free glycan samples were reconstituted in buffer A (80%: 20% acetonitrile: water), filtered with 0.22 μm spin filter (Corning) and loaded onto a Kinetex HILIC column (2.6 μm, 2.6×150 mm; Phenomenex) with 80% ACN/20% water as buffer A and 50 mM NH4FA with pH 4.4 as buffer B. LC was performed using a 7-min gradient from 80 to 0% of buffer B at a flow rate of 400 μL/min.
All LC-MS/MS analysis was carried out using an X500B QTOF (SCIEX) mass spectrometer equipped with an electrospray ion source and coupled with an Exion HPLC system. Each reconstituted sample was injected onto a Kinetex HILIC column (2.6 μm, 2.6×150 mm; Phenomenex). The free glycans were eluted in a 9-min gradient of 80% to 0% (80% ACN/20% water) at 400 nL/min followed by a 3-minute hold at 80% (80% ACN/20% water) for re-equilibration. The instrument was operated in positive ion mode with ESI voltage set at 5.0 kV, ion source gas 1, gas 2=50 psi, curtain gas=35 and CAD gas=7, and source temperature of 350° C. Calibration was done using positive calibrant with CDS system. For free glycan analysis, the instrument was operated in MS full-scan mode from m/z range from 2,00-2,000 followed by multiple reaction monitoring high-resolution (MRM-HR) scan from 0-12 minutes at two different collision energies of 20 and 35 V with DP=20 V and accumulation time of 0.25 s. MS survey scans were performed for the mass range of m/z 2,00-2000 with DP=20 V, CE=7 V and accumulation time of 0.25 s and MS/MS MRM-HR scans were at the same DP voltage and CE=20 V and with Q1 unit resolution. All MS and MS/MS raw spectra from each sample obtained by MRM-HR scan were analyzed by SCIEX OS 1.4 data analysis system. XIC spectra were extracted from MS full-scan with each MRM transition. The glycan structure was annotated manually using GlycanMass-ExPAsy tool.
The name, amino acid sequence, structure availability (full-length or partial), and predicted post-translational modifications (i.e., disulfide bonds, glycosylation) for each GT enzyme were retrieved from the UniProt database (UniProt, C., “UniProt: A Worldwide Hub of Protein Knowledge,” Nucleic Acids Res 47:D506-D515 (2019), which is hereby incorporated by reference in its entirety). GT family members were annotated from the CAZy database (Lombard et al., “The Carbohydrate-active Enzymes Database (CAZy) in 2013,” Nucleic Acids Res 42:D490-5 (2014), which is hereby incorporated by reference in its entirety). Amino acid sequences of full length, truncated, and SIMPLEx-fused GTs were compiled in FASTA format. The Mw and pI were calculated using the ExPASy Bioinformatics resource portal in average resolution setting (Wilkins et al., “Protein Identification and Analysis Tools in the ExPASy Server,” Methods Mol. Biol. 112:531-52 (1999), which is hereby incorporated by reference in its entirety). Solubility prediction score was calculated using CamSol Intrinsic version 2.1 (Sormanni et al., “The CamSol Method of Rational Design of Protein Mutants With Enhanced Solubility,” J. Mol. Biol. 427:478-90 (2015), which is hereby incorporated by reference in its entirety). The expression scores for all constructs were annotated based on immunoblots in
To ensure robust reproducibility of all results, experiments were performed with at least three biological replicates and at least three technical measurements. Sample sizes were not predetermined based on statistical methods but were chosen according to the standards of the field (at least three independent biological replicates for each condition), which gave sufficient statistics for the effect sizes of interest. All data were reported as average values with error bars representing standard error of the mean (SEM). Statistical significance was determined by Welch's t-test and p-values of <0.05 were considered significant. All graphs were generated using Microsoft Excel, Prism 9 for MacOS version 9.2.0, or R software version 3.4.2. No data were excluded from the analyses. The experiments were not randomized. The Investigators were not blinded to allocation during experiments and outcome assessment.
Towards the goal of developing a versatile and universal approach for large-scale GT production, it was hypothesized that SIMPLEx could relieve bottlenecks that have hampered GT expression in E. coli. The rationale for this hypothesis was based on two observations. First, the SIMPLEx strategy has previously been shown as a promising technique for converting IMPs into water-soluble proteins with retention of biological function (Mizrachi et al., “Making Water-soluble Integral Membrane Proteins In Vivo Using an Amphipathic Protein Fusion Strategy,” Nat. Commun. 6:6826 (2015) and Mizrachi et al., “A Water-soluble DsbB Variant That Catalyzes Disulfide-bond Formation In Vivo,” Nat. Chem. Biol. 13:1022-1028 (2017), which are hereby incorporated by reference in their entirety). Second, SIMPLEx was able to rescue soluble expression of a diverse panel of globular proteins that were previously reported to be recalcitrant to soluble expression in E. coli (Dyson et al., “Production of Soluble Mammalian Proteins in Escherichia coli: Identification of Protein Features That Correlate With Successful Expression,” BMC Biotechnol. 4:32 (2004), which is hereby incorporated by reference in its entirety) (
To see if the benefits of SIMPLEx could be leveraged for GT expression, the human β-galactoside-α2,6-sialyltransferase 1 (HsST6Gal1), a sialytransferase belonging to the GT29 family, was chosen as a model GT for proof-of-concept experiments. HsST6Gal1 consists of a short N-terminal cytoplasmic tail (CT), a transmembrane domain (TMD), a stem region that serves as a linker, and a large C-terminal catalytic domain that adopts a variant GT-A fold containing a seven-stranded central 3-sheet flanked by α-helices (
To demonstrate the importance of the decoy and shield domains, chimeras lacking each of these elements were also expressed. When the decoy protein was omitted, Δ26HsST6Gal1-ApoAI* partitioned almost entirely in the insoluble fraction (
To determine whether soluble Sx-Δ26HsST6Gal1 was biologically active, the enzyme was purified (
Upon confirming that Sx-Δ26HsST6Gal1 was enzymatically active, it was next sought to demonstrate its practical utility for chemoenzymatic remodeling of N-linked glycans present on glycoprotein substrates. To this end, a bioorthogonal click chemistry-based assay for quantifying sialyltransferase-mediated chemoenzymatic modification was developed (
Using clarified lysate generated from E. coli cells expressing Sx-Δ26HsST6Gal1 as a catalyst source, a strong fluorescence from the treated A1AT was detected (
Encouraged by the ability of SIMPLEx to promote soluble expression of HsST6Gal1 in E. coli while preserving its biological activity, whether the strategy could be extended to a larger collection of structurally diverse GTs was next investigated. To this end, a library of 98 GT genes from diverse prokaryotic and eukaryotic organisms was compiled, with an emphasis placed on those of human origin (
Another advantage of expressing GTs in the SIMPLEx framework is the potential to relieve cellular stress that arises from high-level accumulation of severely misfolded proteins (e.g., inclusion bodies) or destabilization of the cytoplasmic membrane caused by high-level expression of membrane proteins, phenomena that are both well-known to negatively impact cell growth and productivity. Indeed, cultures expressing Sx-GTs were consistently observed to reach higher final cell densities than those expressing unfused GTs (
It was next sought to identify the protein features that correlated with soluble protein expression by comparing physicochemical properties of the proteins including molecular weight (Mw), isoelectric point (pI), and amino acid content. This involved assigning an expression score to each of the Sx-GT and GT constructs based on their soluble expression profiles (
This observation prompted further investigation of the relationship between soluble expression of the protein and its Mw. To this end, all GTs were categorized into one of three size groups: small (Mw<40 kDa), medium (Mw=40-60 kDa), and large (Mw>60 kDa). The average expression score (
To further expand the utility of the platform and demonstrate its universality, SIMPLEx fusions in other popular expression platforms including: (i) E. coli-based cell-free protein synthesis (CFPS); (ii) Saccharomyces cerevisiae strain SBY49; and (iii) human embryonic kidney (HEK) 293T cells were produced. Using appropriate expression vectors for each system, significant accumulation of the Sx-Δ26HsST6Gal1 construct in the soluble fractions derived from each of these three systems was observed (
To date, a growing number of cell-free bio/chemoenzymatic synthesis strategies have been reported that provide access to large repertoires of pure and chemically-defined glycans, especially complex structures that are otherwise difficult to obtain by conventional chemical synthesis (Hamilton et al., “A Library of Chemically Defined Human N-glycans Synthesized From Microbial Oligosaccharide Precursors,” Sci. Rep. 7:15907 (2017); Li and Wang, “Chemoenzymatic Methods for the Synthesis of Glycoproteins,” Chem. Rev. 118:8359-8413 (2018); and Li et al., “Strategies for Chemoenzymatic Synthesis of Carbohydrates,” Carbohydr. Res. 472:86-97 (2019), which are hereby incorporated by reference in their entirety). Because these approaches generally depend on the availability of glycoenzymes, many of which cannot be recombinantly expressed or purified at scale, it was sought to demonstrate the practical utility of Sx-GTs as biocatalysts for constructing customized glycan structures via a previously described bioenzymatic synthesis approach (Hamilton et al., “A Library of Chemically Defined Human N-glycans Synthesized From Microbial Oligosaccharide Precursors,” Sci. Rep. 7:15907 (2017), which is hereby incorporated by reference in its entirety). To this end, two multi-GT enzyme pathways for de novo biosynthesis of a library of human hybrid- and complex-type N-glycans starting from a mannose3-N-acetylglucosamine2 (Man3GlcNAc2) primer were devised (
Using 1 as a primer, glycan elaboration with GlcNAc was carried out by sequential treatment with purified Sx-Δ29HsGnTI and Sx-Δ29HsGnTII, yielding hybrid-type glycan 2 (also known as G0-GlcNAc) and complex-type glycan 3 (G0), respectively, as evidenced by MALDI-TOF MS analysis of each reaction (
Overall, enzymatic conversion in each of these reactions was at or near 100% except in the cases involving the Sx-Δ26HsST6Gal1-catalyzed sialyation reactions. However, because the unstable nature of sialic acid-containing glycans in MALDI-TOF MS may have confounded the sialylation analysis, nano-scale reverse phase chromatography and tandem MS (nano LC-MS/MS) analysis were performed to confirm the abundance and identity of the sialylated glycans 5, 6, 9, and 10. While both mono- and di-sialylated products were clearly detected, this analysis revealed an approximate 5:1 ratio between the G2S1 and G2S2 glycans as well as the G2S1F and G2S2F glycans (
Glycoform manipulation is an emerging strategy for improving pharmacokinetics and pharmacodynamics of therapeutic glycoproteins (Wang et al., “Glycoengineering of Antibodies for Modulating Functions,” Annu. Rev. Biochem. 88:433-459 (2019) and Wang and Lomino, “Emerging Technologies for Making Glycan-defined Glycoproteins,” ACS Chem. Biol. 7:110-22 (2012), which are hereby incorporated by reference in their entirety). The remodeling of protein-linked glycans can be readily achieved using one or more GTs; however, the limited availability of requisite enzymes for customizing glycan structures represents a barrier to widespread adoption. To address this technology gap, members from the disclosed library of SIMPLEx-reformatted GTs were employed to alter the glycan profiles on several biomedically-relevant glycoproteins. Remodeling reactions included: (i) Sx-CjCstII-mediated α2,3-sialylation of the N-glycoforms on α1-antitrypsin (A1AT), a serpin used in prophylactic treatment of the genetic disorder α1-antitrypsin deficiency; (ii) Sx-Δ36HsFucT7-mediated fucosylation of the N-glycoforms on A1AT; (iii) Sx-Δ34HsST3Gal1-mediated α2,3-sialylation of the O-glycoforms on bovine submaxillary mucin (BSM), a glycoprotein with potential uses as a biocompatible material and drug delivery vehicle; and (iv) Sx-Δ29HsGnTI-catalyzed GlcNAc transfer onto Man3GlcNAc2 glycans present on a neoglycoprotein variant of human glucagon (GCG). In all cases, Sx-GTs readily remodeled their glycoprotein substrates, installing respective monosaccharides in 1-hour reactions that were monitored using bioorthogonal click chemistry-based assays with either a fluorophore or biotin reporter for glycan labeling (
N-glycans present on the Fc domain of IgG antibodies play a critical role in the structure and function of these important proteins, but understanding of how discrete glycan structures affect IgG behavior remains limited due to naturally occurring microheterogeneity. Hence, strategies for generating structurally-defined N-glycans on IgG-Fc are expected to improve the understanding of the roles played by these structures in human immunity and to open the door to creating better medicines through glycoengineering. To this end, members from the disclosed library of Sx-GTs were leveraged to generate a homogenously glycosylated variant of trastuzumab (
In addition to producing authentic, homogeneous human N-glycans, whether Sx-GTs could generate IgG-Fc bearing unnatural glycan structures was also investigated. To this end, Sx-Δ29HsGnTI was used to elaborate trastuzumab N-glycans with N-azidoacetylglucosamine (GlcNAz), a synthetic monosaccharide containing an azide moiety (
Examples 1-9 describe the creation of a universal expression platform for producing nearly 100 different GTs, predominantly of human origin, at relatively high titers (approximately 5-10 mg/L) using standard bacterial culture. This platform leverages SIMPLEx to engineer GT chimeras that are rendered highly soluble in the cytoplasm of E. coli cells. Consistent with earlier works (Mizrachi et al., “Making Water-soluble Integral Membrane Proteins In Vivo Using an Amphipathic Protein Fusion Strategy,” Nat. Commun. 6:6826 (2015) and Mizrachi et al., “A Water-soluble DsbB Variant That Catalyzes Disulfide-bond Formation In Vivo,” Nat. Chem. Biol. 13:1022-1028 (2017), which are hereby incorporated by reference in their entirety), SIMPLEx-reformatted GTs retained biological activity as exemplified by the human ST6Gal1 chimera that exhibited activity that was similar to a commercially sourced enzyme. The ability to solubilize such a large set of GTs without compromising function made it possible to remodel the structures of different free and protein-linked glycans including those found on the monoclonal antibody trastuzumab. Overall, the platform described infra represents a versatile addition to the synthetic glycobiology toolkit, providing easy access to a vast collection of transformative reagents that are expected to find use in structure-function studies of GTs and to fuel myriad applications where complex glycomolecules are featured.
Previous studies revealed the capacity of SIMPLEx to broadly transform all major classes of IMPs into water-soluble molecules (Mizrachi et al., “Making Water-soluble Integral Membrane Proteins In Vivo Using an Amphipathic Protein Fusion Strategy,” Nat. Commun. 6:6826 (2015) and Mizrachi et al., “A Water-soluble DsbB Variant That Catalyzes Disulfide-bond Formation In Vivo,” Nat. Chem. Biol. 13:1022-1028 (2017), which are hereby incorporated by reference in their entirety). These IMPs included proteins having both bitopic and polytopic α-helical structures such as glutamate receptor (GluA2) and bacteriorhodopsin (bR) as well as polytopic β-barrel structures such as voltage-dependent anion channel 1 (VDAC1). Here, this solubilization capacity was broadened to include polytopic α-helical GTs with multiple TMDs such as found in human mannosyltransferases Alg2, Alg3, and Alg12 and human glucosyltransferases Alg6, Alg8, and Alg10 as well as monotopic α-helical GTs with single-pass internal TMDs that could not be easily removed such as Alg2 and PigA. For these complex integral membrane proteins, introduction of an N-terminal decoy protein, MBP, prevented co-translational insertion of the polypeptide into the inner membrane through the signal recognition particle (SRP) pathway (Luirink and Sinning, “SRP-mediated Protein Targeting: Structure and Function Revisited,” Biochim. Biophys. Acta. 1694:17-35 (2004), which is hereby incorporated by reference in its entirety) while the amphipathic ApoAI* domain effectively shielded the hydrophobic TMDs from the aqueous environment.
It is noteworthy that most of the GTs investigated (72 out of 98 total) were simpler type II transmembrane proteins. Type II GTs such as HsST6Gal1 possess just a single-pass TMD at their N- or C-termini (
Importantly, the SIMPLEx architecture enabled soluble expression for nearly 100 GTs (>95% “hit” rate) under standard, identically matched conditions without any optimization, thereby offering a universal solution to GT production in E. coli that has not been possible with stand-alone fusion tags such as MBP or other expression optimization techniques (Wagner et al., “Rationalizing Membrane Protein Overexpression,” Trends Biotechnol. 24:364-71 (2006), which is hereby incorporated by reference in its entirety). An additional layer of universality stems from the compatibility of SIMPLEx-mediated GT solubilization with other commonly used expression hosts such as yeast and HEK293 cells as well as with E. coli-based cell-free protein synthesis (CFPS). Such platform flexibility is significant for several reasons. For one, each of these platforms is amenable to high-throughput profiling of protein expression and production that can be scaled up to larger volumes (Subedi et al., “High Yield Expression of Recombinant Human Proteins with the Transient Transfection of HEK293 Cells in Suspension,” J. Vis. Exp. e53568 (2015) and Spirin, A. S., “High-throughput Cell-free Systems for Synthesis of Functionally Active Proteins,” Trends Biotechnol. 22:538-45 (2004), which are hereby incorporated by reference in their entirety). Moreover, in the case of yeast and HEK293, the compatibility of SIMPLEx-reformatted GTs in these well-established eukaryotic hosts may provide access to protein folding networks and post-translational modifications including N- and O-linked glycosylation that may be important for the biological function of a subset of GTs (Mikolajczyk et al., “How Glycosylation Affects Glycosylation: The Role of N-glycans in Glycosyltransferase Activity,” Glycobiology 30:941-969 (2020), which is hereby incorporated by reference in its entirety) but are natively lacking in standard E. coli strains. In the case of E. coli-based CFPS, the “open” nature and multiplexability of these systems, combined with their speed and simplicity, should provide opportunities for high-throughput screening of GT function (Kightlinger et al., “Design of Glycosylation Sites by Rapid Synthesis and Analysis of Glycosyltransferases,” Nat. Chem. Biol. 14(6):627-635 (2018), which is hereby incorporated by reference in its entirety) as well as rapid discovery, prototyping, and optimization of glycomolecule synthesis pathways (Karim and Jewett, “A Cell-free Framework for Rapid Biosynthetic Pathway Prototyping and Enzyme Discovery,” Metab. Eng. 36:116-126 (2016) and Kightlinger et al., “A Cell-free Biosynthesis Platform for Modular Construction of Protein Glycosylation Pathways,” Nat. Commun. 10:5404 (2019), which are hereby incorporated by reference in their entirety).
As proof of concept for the utility of the disclosed SIMPLEx pipeline, some of the solubilized products were used in coordinated cell-free reaction networks to catalyze the formation of chemically-defined N-glycans. In one instance, it was possible to transform quantitative amounts of a simple paucimannose precursor N-glycan, Man3GlcNAc2 derived from glycoengineered E. coli (Valderrama-Rincon et al., “An Engineered Eukaryotic Protein Glycosylation Pathway in Escherichia coli,” Nat. Chem. Biol. 8:434-6 (2012) and Glasscock et al., “A Flow Cytometric Approach to Engineering Escherichia coli for Improved Eukaryotic Protein Glycosylation,” Metab. Eng. 47:488-495 (2018), which are hereby incorporated by reference in their entirety), into complex biantennary N-glycans including those containing core-fucose and sialic acid caps using a set of SIMPLEx-reformatted GTs. This workflow to efficiently generate a library of complex N-glycans, starting from expression and purification and then finally utilization of SIMPLEx-reformatted GTs, could be completed in less than one week. Using an identical strategy, it was possible to generate a spectrum of homogenous N-glycan structures on intact glycoproteins including trastuzumab, a mAb therapy used to treat breast and stomach cancers. Akin to earlier engineering of an artificial cytoplasmic disulfide formation pathway involving a water-soluble SIMPLEx variant of DsbB (Mizrachi et al., “A Water-soluble DsbB Variant That Catalyzes Disulfide-bond Formation In Vivo,” Nat. Chem. Biol. 13:1022-1028 (2017), which is hereby incorporated by reference in its entirety), ensembles of SIMPLEx-reformatted GTs could similarly be assembled into designer pathways, either in vitro or in living cells, for the on-demand biosynthesis of important glycans and glycoconjugates. Looking forward, it is anticipated that the constructs, expression systems, and workflows for glycoenzyme production described herein will find widespread use by those seeking to push the boundaries of our knowledge of glycobiology and glycochemistry and its application in health, energy, and materials science.
Although preferred embodiments have been depicted and described in detail herein, it will be apparent to those skilled in the relevant art that various modifications, additions, substitutions, and the like can be made without departing from the spirit of the invention and these are therefore considered to be within the scope of the invention as defined in the claims which follow.
This application claims the benefit of U.S. Provisional Patent Application Ser. No. 63/297,419, filed Jan. 7, 2022, which is hereby incorporated by reference in its entirety.
This invention was made with government support under HDTRA1-14-10052 awarded by Defense Threat Reduction Agency; CBET-1159581, CBET-1264701, CBET-1936823, and MCB 1413563 awarded by National Science Foundation; and 1R01GM137314, and 1R01GM127578 awarded by National Institutes of Health. The government has certain rights in the invention.
| Filing Document | Filing Date | Country | Kind |
|---|---|---|---|
| PCT/US2023/010330 | 1/6/2023 | WO |
| Number | Date | Country | |
|---|---|---|---|
| 63297419 | Jan 2022 | US |
