Title of Invention: High-Throughput Cell-Based Screening Methodology For Evaluating Carbohydrate-Active Enzymes Abstract: The present disclosure relates, in one aspect, to the discovery of a high throughput screening (HTS) method to rapidly screen for GH/GS variants that are generated using directed evolution techniques and that can significantly enhance glycosynthase catalytic activity or product specificity. Detailed Description: CROSS REFERENCE TO RELATED APPLICATIONS This application claims priority under 35 U.S.C. § 119(e) to U.S. Provisional Patent Application No. 62/877,021, filed Jul. 22, 2019, which application is incorporated herein by reference in its entirety. STATEMENT REGARDING FEDERALLY SPONSORED RESEARCH OR DEVELOPMENT This invention was made with government support under grant numbers 1904890 and 1704679 awarded by the National Science Foundation. The government has certain rights in the invention. SEQUENCE LISTING The ASCII text file named “370602-7015US1(00056) Sequence,” created on Jul. 22, 2020, comprising 33.4 Kbytes, is hereby incorporated by reference in its entirety. BACKGROUND OF THE DISCLOSURE Synthesis of glycan-based polymers (oligosaccharides and polysaccharides) using engineered carbohydrate-active enzymes (CAZymes) offers exquisite regioselective and stereoselective control over traditional synthetic chemistry approaches, which are atom inefficient and involve multi-step transformations. Glycosyltransferases (GTs) are naturally occurring CAZymes that synthesize glycans but give poor heterologous expression yields, have narrow substrate specificity, and use expensive nucleotide sugars, limiting the scale-up of in vitro glycans synthesis. Chemoenzymatic synthesis using glycosyl hydrolases (GH) could permit production of complex glycans at high yields. GH are nature's antipodes of GT by hydrolyzing glycosidic linkages, but can also produce glycans via transglycosylation if the nucleophilic water is replaced by a sugar molecule as an acceptor. Unfortunately, transglycosylation suffers from low yields since the product is also a substrate for GH-mediated hydrolysis. However, most GH have plasticity in their structure, which allows for improving synthase activity. Interestingly, glycosynthases (GSs) offer an alternative biosynthetic approach to producing glycans in a facile manner. The GSs are mutants of readily available microbial glycosyl hydrolases (GHs), which are incapable of hydrolyzing glycosidic bonds, and can be engineered to specifically synthesize complex glycans. However, to date, only a limited number of GSs have been created from wild-type GHs using an inefficient empirical strategy that have limited biosynthetic activity. Unlike GTs, there is a much larger selection of GHs available that can be expressed readily in E. coli. Further, the active site GH nucleophile residue can be mutated to prevent product hydrolysis and improve product yields. However, the role of various accessory domains on the transglycosylation activity of mutant GH/GS is mostly unknown. Thus, there is a need in the art for a method of identifying mutant GH/GS enzymes that allow for glycan production. The present disclosure fulfills this need. BRIEF SUMMARY OF THE DISCLOSURE Disclosed herein is a method of determining if a protein has transglycosylase activity. In certain embodiments, the method comprises contacting the protein with an azido glycosyl donor and a glycosyl acceptor to form a system, and measuring any change in the azide concentration in the system. In certain embodiments, the azido glycosyl donor is substituted with an azido group at an anomeric carbon. In other embodiments, the azido glycosyl donor is substituted with an azido group at a non-anomeric carbon. In certain embodiments, the measurement of azide concentration comprises measurement of the concentration of an inorganic azide. In other embodiments, the measurement of azide concentration comprises the measurement of the concentration of an organic substituted azide, including azido glycosyl species. In certain embodiments, the measuring step comprises contacting the system with a reagent comprising a strained alkyne coupling to a dye, under conditions that allow for reaction of the strained alkyne with any azide or azido compound present in the system. In certain embodiments, the reagent comprises bicycle[6.1.o]nonyne (BCN), dibenzocyclooctyne (DBCO), or any other strained alkyne. In certain embodiments, the reagent comprises 5-carboxytetramethylrhodamine (5-TAMRA), 6-carboxytetramethylrhodamine (6-TAMRA), or any combinations thereof. In certain embodiments, the strained alkyne and the dye are covalently linked by a linker in the reagent. In certain embodiments, the linker comprises a polyethylene glycol linker. In certain embodiments, the measuring step uses a control protein that has no measurable transglycosylase activity or has a known transglycosylase activity. In other embodiments, the protein is a mutated glycosyl hydrolase (GH). In certain embodiments, the protein is expressed in a cell. In other embodiments, the cell comprises E. coli or P. pastoris. In certain embodiments, the system is within a cell (intracellular). In certain embodiments, the measuring step comprises monitoring the fluorescence of the system. In other embodiments, fluorescence activated cell sorting (FACS) is used to separate individual cells by measured fluorescence. In other embodiments, the FACS is configured for high-throughput screening. Disclosed herein are mutant polypeptide amino acid sequences of WT TmAfc-0306_(SEQ ID NO:1) comprising the mutation D224G (SEQ ID NO:2) and further comprising at least one additional mutation. In certain embodiments, the at least one additional mutation of the mutated construct (SEQ ID NO:2) is selected from the group consisting of L15K, N70D, A366V, T392S, K395N, D400A, T413P, I428T, and T429P. In other embodiments, the at least one additional mutation is selected from the group consisting of L15K-N7OD (SEQ ID NO:3); N70D-T392S (SEQ ID NO:4); N70D-T392S-A366V-K395N (SEQ ID NO:5); N70D-T392S-D400A (SEQ ID NO:6); N70D-T392-I428T (SEQ ID NO:7); and N70D-D400A-T413P-T429P (SEQ ID NO:8). BRIEF DESCRIPTION OF THE DRAWINGS The following detailed description of illustrative embodiments of the disclosure will be better understood when read in conjunction with the appended drawings. For the purpose of illustrating the disclosure, exemplary embodiments are shown in the drawings. It should be understood, however, that the disclosure is not limited to the precise arrangements and instrumentalities of the embodiments shown in the drawings. FIG. 1A: Two-step hydrolysis mechanism of WT Agrobacterium sp. β-glucosidase. First, the anomeric configuration of the sugar glycosidic bond (β linkage) is inverted to form the α-glycosyl-enzyme intermediate (GEI). The second step recovers the original anomeric configuration in the product (β-D-glucose). FIG. 1B: The E358A mutant glycosidase (or glycosynthase) cannot form a productive GEI. The enlarged binding site accommodates the α-glucosyl fluoride activated sugar donor. FIG. 2A: One-step hydrolysis mechanism of an inverting glycoside hydrolase. FIG. 2B: One-step synthesis mechanism of an inverting GT employing a divalent cation. FIG. 2C: Two-step synthesis mechanism of a retaining GT, showing the GEI. FIGS. 3A-3B: Two common human milk oligosaccharides (HMOs) used to test GH29 activity are (FIG. 3A) 2′-fucosyllactose (2′FL) and (FIG. 3B) 3′-fucosyllactose (3′FL). FIG. 4A: The structure of the TmAfcA (PDB ID: 10DU), showing a bound fucose product, acid/base residue G266 (upper rendered side chain), and nucleophilic residue D224 (lower rendered side chain). FIG. 4B: Structure of glycosynthase TmAfcA D224G co-crystalized with fucosyl-azide, revealing that the overall protein structure is maintained in the mutant. FIGS. 5A-5E: Computational methodology for uncovering CAZyme reaction mechanisms. FIG. 5A: If a solved crystal structure is unavailable, utilize homology modeling to predict CAZyme structure. FIG. 5B: Perform MD simulations to determine low energy conformations of reactant and product states. FIG. 5C: Informed by previously discovered transition state (TS) conformations, build multiple hypothesized TS structures. FIG. 5D: Using unbiased simulations, determine whether postulated TSs indeed lead to both reactant and product low-energy states. These are used to generate an ensemble of validated TSs to analyze for properties that correlate with reactivity, thus revealing the reaction coordinate. FIG. 5E: The resulting reaction coordinate allows determination of the energy required to overcome reaction barriers, which is converted to reaction rate coefficients that directly relate to experimentally observable activities. FIG. 6A: Donor and acceptor sugars docked within CelE active site. FIG. 6B: Agar-CMC plate based GH-GS zone-clearing method. FIG. 6C: Some CelE mutants can efficiently synthesize oligosaccharides. FIG. 6D: SAXS suggests dynamic interaction of CBM with CelE catalytic domain drives TG activity. FIG. 7: Phylogenetic tree of GH29 enzymes from published amino-acid sequences. The green arrows highlight the 3 enzymes to be studied to build the initial computational, predictive models. The enzymes highlighted in red are mutated, based on model predictions, to create new GSs. These genes were chosen to sample a diverse range of phylogenetically related sequences. FIG. 8: Snapshots of the elementary synthesis step by TmAfcA D224G, joining 1-Azido-β-L-Fucose (1AF) to 4-Nitrophenyl β-D-xylopyranoside (4NX). FIG. 9: Chemical rescue of TmAfcA mutants' hydrolytic activity on pNP-fucose using an external nucleophile confirms that the D224G mutant is a true GS. FIG. 10A: HT-cloning and CAZymes synthesis using wheat germ cell free expression system. CAZymes directly assayed for (FIG. 10B) binding or (FIG. 10C) catalytic activity on glycans in a combinatorial manner. FIG. 11A: Glycosynthase products docked in the active site of TmAfc-D224G based on quantum mechanical/molecular mechanics. FIG. 11B: Thin-layer chromatography (TLC) analysis of glycosynthase reaction of D224G with β-L-fucopyranosyl azide and pNP-β-D-xylopyranoside visualized with UV light. FIG. 11C: Reaction scheme of glycosynthase reaction of Tm0306-D224G with β-L-fucopyranosyl azide and pNP-β-D-xylopyranoside. FIG. 12: SDS-PAGE protein gel for wild-type TmAFc0306_(WT), mutant D224A TmAfc0306_(A), mutant D224S TmAfc0306_(S), mutant D224G TmAfc0306_(G), and several epPCR derived purified mutant proteins identified by screening. FIG. 13: TLC of glycosynthase reaction mixtures resulting from either wild-type (WT) or mutant D224G (Gly) with β-L-fucopyranosyl azide and pNP-β-D-xylopyranoside substrates. FIG. 14: FACS based directed evolution approaches utilizing azide detection for screening glycan synthesizing CAZymes. FIG. 15 illustrates the structure of fluorescent alkyne-DBCO-PEG4-FLUOR 545, as well as its reaction with sodium azide. FIG. 16 illustrates absorbance analysis for the reaction of alkyne-DBCO-PEG4-TAMRA (or alkyne-DBCO-PEG4-Fluor 545) with sodium azide and glucosyl azide. UV spectra shows the loss of absorbance at 309 nm over time for both reactions. FIG. 17 illustrates fluorescence analysis of click reaction of DBCO-PEG4-Fluor 545 and sodium azide and glucosyl azide in vitro. The alkyne to azide ratio is 1:2 in 1× PBS pH 7.4 buffer (37° C., 400 rpm, 5 hours). FIGS. 18A-18B illustrates the fluorescence analysis of the SPAAC reaction between DBCO-PEG4-FLUOR 545 (200 μM) with sodium azide and β-D-glucopyranosyl azide in vitro at a temperature with a temperature of (FIG. 18A) 25° C. and (FIG. 18B) 10° C. FIG. 19 provides the emission fluorescence as a function of excitation wavelength for SPAAC reaction products. FIGS. 20A-20D illustrate the impact of free azide and triazole moieties on Rhodamine B fluorescence. FIG. 20A provides the structures of Rhodamine B and Fluor 545. FIG. 20B provides the relative fluorescence traces for mixtures of Rhodamine B with either organic or inorganic azide. FIG. 20C provides UV spectra for SPAAC reactions confirming the formation of triazole products from organic and inorganic azides with DBCO-NHS. FIG. 20D provides the fluorescence of a mixture of Rhodamine B with different triazoles independently as a function of time. FIG. 21 provides a comparison of the fluorescence of the SPAAC reaction of glucosyl and fucosyl azide with DBCO-PEG4-Fluor 545 as a function of time. FIG. 22 provides the fluorescence of SPAAC reactions of organic, inorganic, and an equimolar mixture of inorganic and organic azides with DBCO-PEG4-FLUOR 545. FIG. 23A provides the protocol for sample preparation for confocal microscopy. FIG. 23B provides confocal microscopy images confirming permeation of DBCO-PEG4-FLUOR 545 dye into E. coli cells via Hoechst staining (0.1 μg/ml), bright field microscopy, and fluorescence with DBCO (0.1 mM). FIG. 24 provides a proof-of-concept of the use of fluorescence assisted cell sorting (FACS) to identify and separate cells on the basis of the difference in fluorescence observed from triazole products formed in a SPAAC reaction in vivo. FIGS. 25A-25B illustrates the in vivo click reaction of DBCO-PEG4-FLUOR 545 with sodium azide and β-L-fucopyranosyl azide, as monitored by FACS. FIG. 25A provides a scatter plot of the in vivo click reaction. FIG. 25B provides a fluorescence decay plot of the in vivo click reaction. FIG. 26 provides flow cytometry characterization of fucosynthase (D224G) and wild type (WT) TmAfc enzyme expressing E. coli populations. FIG. 27 provides FACS characterization of fucosynthase (D224G) and wild type (WT) TmAfc enzyme expressing E. coli populations. FIG. 28 provides FACS based sorting of D224G mutant epPCR library with variation of the excitation wavelength used. FIG. 29 illustrates the impact of free inorganic azide on initial E. coli growth rate. FIG. 30 provides a conceptual overview of click-chemistry based ultrahigh-throughput screening (uHTS) for in vivo detection and sorting of improved glycosynthases. FIG. 31A provides FACS scatter plots of the control and mutated libraries observed emplying an uHTS method. FIG. 31B provides a bar graph representing the chemical rescue activity for D224G cells and sorted mutants with improved GS activity. FIG. 31C provides the specific activity of the isolated M5 mutant (SEQ ID NO:6). FIG. 32 provides a bar graph representing the chemical rescue activity for D224G cells and sorted mutants with improved GS activity. FIG. 33 provides histograms showing the sampling along the reaction coordinate from each window in the umbrella sampling procedure described herein. FIG. 34A provides the root mean square fluctuation of the M5 mutant fucosynthase structural model. FIG. 34B provides QM/MM simulations predicted free energy profile of the M5 mutant fucosynthase reaction. FIG. 35A: Fucosylated glycans currently produced. FIG. 35B: Suite of complex fucosylated oligosaccharides. FIGS. 36A-36C provides FACS scatter plots of an epPCR mutant library sorted to screen for mutants showing activity with β-L-fucopyranosyl azide as a donor sugar and common acceptor sugars associated with fucose: (FIG. 36A) galactose, (FIG. 36B) lactose, and (FIG. 36C) N-acetyl galactosamine. DETAILED DESCRIPTION OF THE DISCLOSURE The present disclosure relates, in one aspect, to the discovery of a high throughput screening (HTS) method to rapidly screen for GH/GS variants that are generated using directed evolution techniques and that can significantly enhance glycosynthase catalytic activity or product specificity. A model fucosynthase from Thermotoga maritima has been developed for validation of this HTS method. Copper-free click chemistry reaction conditions were optimized for rapid quantification of azide-based products formed by active glycosynthase mutants using fluorescence. The difference in fluorescence profiles of wild type enzyme and mutants were analyzed using a flow cytometer. This click chemistry based screening technique was applied to the mutant library generated by random mutagenesis. In certain embodiments, this technique is a universal approach to screen for glycosynthases that have activated azide group on the donor substrate. Glycoside Hydrolases, Glycosyltransferases, and Glycosynthases GHs and GTs are ubiquitous enzymes found in all kingdoms due to the central role of carbohydrates in life processes, as various oligosaccharide structures are used in diverse biological functions including signaling, energy storage, and structural components. GH enzymes cleave glycosidic bonds that join monomeric sugars to create oligosaccharides, and are grouped into families by amino-acid sequence similarity, now numbering 156 families as curated on the Carbohydrate-Active enZyme (CAZyme) database. Enzymes of each family are classified as either retaining or inverting, depending on whether the stereochemistry at the anomeric carbon is preserved between the reactant and product. Retaining enzymes (FIG. 1A) require a two-step hydrolysis mechanism, with a glycosyl-enzyme intermediate (GEI) characterized by a covalent bond between the cleaved substrate and the protein in the alternate orientation (in FIG. 1A, the GEI has an α-bond, as opposed to the β-orientation of the reactant and product). In inverting enzymes (FIG. 2A), a water molecule attacks the substrate while an acidic protein residue donates a proton to the glycosidic bond and a basic protein residue accepts a proton from the attacking water molecule. GH enzymes are often secreted extracellularly by organisms, and industrial processes routinely use them to break down carbohydrates, such as in converting plant biomass into simple sugars. GTs fill an opposite function in nature, creating oligosaccharides by joining a sugar acceptor with an activated monomer, most commonly nucleotide diphosphate sugars such as UDF-glucose, UDP fructose, or GDP-mannose. in contrast to GHs, GTs operate mostly within cells (typically as membrane-associated proteins), and are less soluble and stable compared to GHs, and thus are less suited to industrial use. Further hampering their exploitation for production of oligosaccharides for research and industrial use is the high cost of generating sugar nucleotide substrates. In vivo synthesis can address some of these challenges by transferring or modifying the biosynthetic glycosylation pathways from desired eukaryotic or prokaryotic systems (e.g., Campylobacter jejuni) into genetically tractable and industrially relevant expression systems like E. coli or Pichia pastoris. However, glycosylation is an innately stochastic process leading to a complex milieu of glycoforms, making it challenging to produce a. defined library of glycans using such approaches alone. Due to the difficulties of using GTs to create oligosaccharides, Gas have been explored for their potential to build glycosidic bonds, exploiting the innate ability of some GB enzymes to act as transglycosylases (TGs). The general mechanism is similar to the retaining mechanism shown in FIG. 1A, except that rather than nucleophilic attack by water, the attack is instigated by a hydroxyl oxygen on a sugar. Some enzymes have a higher natural propensity to act as TGs, and this activity can he enhanced by high sugar concentrations. However, the products are still potential reactants that can either be hydrolyzed or undergo further transgiycosylation, leading to low yields and a mixture of products, limiting their utility for bespoke oligosaccharides synthesis. The insight behind GSs is to enhance the ability of GHs to create glycosidic bonds and remove their capacity for glycosidic bond cleavage. Thus, GSs were created to perform the function of GTs without the need for expensive substrates. Derived from GHs, they retain the favorable GH characteristics of ease of expression and higher stability. Most GSs have been created from retaining GHs, although some have been created from inverting GHs. In those created from retaining GHs, the general approach has followed from the first engineered GS, illustrated in FIG. 1B. Specifically, the nucleophi lie residue is mutated to one that can no longer accept a proton, such as mutation from aspartate or glutamate to alanine, glycine, serine, and cysteine. In certain embodiments, the size of the side chains used in place of the WI nucleophile is an important factor in proper substrate positioning. In addition, the role of other residues within the active site or its vicinity on glycosynthase activity is even more poorly understood, GSs are characterized by the ability for catalytic rescue of hydrolytic activity in the presence of high concentrations (>1M) of an external nucleophile (e.g., azide or formate). To perform synthesis, GSs are supplied with sugars whose structures mimic the GEI. Like the native intermediates, these sugars will have the opposite stereochemistry at the anomeric center from the native reactant, and the anomeric carbon will be bonded to a leaving group such as fluoride or azide. These “donor” sugars are less expensive to produce than the activated, diphosphate sugars required by GTs. These donor sugars are paired with “acceptor” sugars that have high affinity for the binding site not occupied by the GEI mimic (termed the acceptor site in GSs), such as the phenyl-P-β-glucopyranoside shown in FIG. 1B. Since the native ability to hydrolyze glycosidic bonds is minimized for GS, there is no loss of product by subsequent cleavage. The requirement for the donor sugar to have an electrophilic leaving group, which is removed in the course of oligosaccharide synthesis, limits which species can serve as reactants, thus providing more control of the resulting product slate as compared to transglycosylation product distributions. To date, only a limited number of GSs have been created, with between one and six members of any one family having been converted, and only from 17 GH families. The general empirical strategy has been to a) determine the nucleophilic catalytic residue, b) mutate that residue to alanine, glycine, serine, and/or cysteine, c) test for hydrolytic chemical rescue using external nucleophiles, and d) perform activity tests. This empirical approach is cumbersome and lacks the ability to screen growing genomic databases of CAZymes to identify the best targets using a theoretically based first-principles methodology. GH29 Enzymes Of the 156 currently designated GH families, only two families contain α-fucosidases: family 29 (retaining hydrolases) and family 95 (inverting). As retaining enzymes have been more amenable to conversion into GSs, GH29 enzymes provide a promising route for creating enzymes to produce specific fucosylated oligosaccharides. The CAZyme database currently lists over 3,000 protein sequences classified as GH29 enzymes, with additional sequences continually deposited. The enzyme sources span archaea., bacteria, and eukaryota (from fungi to human). Of these, 33 have been characterized and show only α-fucosidase activity, breaking α-1,2-fucoside linkages (as in 2′-fucosyllacose, FIG. 3A) and/or α-1,3-fucoside linkages (as in 3′-fucosyllacose, FIG. 3B). The first structure to be solved was for TmAfcA in 2004, an α-1,2-fucoside (FIG. 4A). Since that time, structures have been solved for GH29 enzymes from seven additional organisms, and a structure of the GS TmAfcA D224G co-crystalized with α-L-Fuc-(1-2)-β-L-Fuc-N3 in the donor site (FIG. 4B; Cobucci-Ponzano, et al, 2009, Chem. Biol. 16:1097-1108). The GSs TmAfcA D224G and SsFucAl D242S displayed varying activity with a β-L-Fuc-N3 donor and a variety of acceptors, demonstrating that one GS can be used to create multiple oligosaccharides. Depending on the acceptor, between 1 and 5 different products were observed for a particular combination of donor-acceptor. In contrast, BfoAfcB D703 S produced only one reaction product for the five donor-acceptor pairs (same acceptors, but a β-L-Fuc-F donor) for which a synthesized compound was identified. Biological Role of Fucosylated Glycans and Current Synthesis Demand The determination of finictional roles of glycans has been enabled by their commercial availability, but only a few such glycans are available, resulting in a limited understanding of glycans in living systems. Even so, it has become clear that fucosylated glycans play many key roles in biology, including mammalian use in ABO blood group antigens, host cells-gut microbe interactions, and selectin-dependent leukocyte adhesion. Also, non-digestible dietary glycans, together with mammalian gut host cells-produced glycans, represent critical energy sources that modulate the survival and proliferation of many microbial components of the gut microbiota. Creating specific glycans by standard chemical synthesis is painstaking and expensive. Thus, biological routes are being pursued. As noted earlier, GSs have advantages for in vitro use, including lower cost compared to GTs and higher yield compared to GHs. Transglycosylation reactions using fucosidases has yielded inefficient routes (<5% yield) to synthesize fucosylated glycans. Much higher product yields of model fucosylated di-, tri-, and tetrasaccharides (30-50%) have been recently shown to be formed using β-fucosyl fluoride sugar donors and GS derived from both GH families 29 and 95. Due to the poor stability of β-glycosyl fluoride (vs. its α-anomer), there has been recent interest in exploring novel activated glycosyl donor sugars like β-fucosyl azides to produce glycans instead. Catalytic efficiency of GS employing non-native activated glycosyl donor substrates requires engineering of the active site residues, that cannot yet be predicted a priori using rational engineering approaches. GHs are being rapidly discovered through cheaper sequencing of isolated microbial, microbiome, and metagenomic sources. These GHs offer a large library of enzymes that have not yet been exploited for engineering more effective and highly selective GSs. Directed evolution of GSs can be used to increase reaction rate and introduce novel substrate specificity. Additionally, isolated novel extremophilic GHs offer an opportunity to develop novel GSs with higher specific activity in non-aqueous solvents that would favor glycan (or glycoconjugate) synthesis and improve reactant and/or substrate solubility. However, one of the major challenges identified has been the lack of suitable high-throughput screening (HTS) methods for screening large GS libraries (>106 mutants/day). A two-plasmid HTS method has been disclosed wherein one plasmid contains the GS gene while the other contains a screening enzyme that only releases a fluorophore from the product of the GS reaction but not the reactants (Bode, et al., 2016, Nutr. Rev. 74:635-644). Similarly, chemical complementation using a yeast three-hybrid system was used to link GS activity to the transcription of a reporter gene, making cell growth dependent on product formation (Lin, et al., 2004, J. Am. Chem. Soc. 126(46):15051-15059). Both of these approaches are highly specific to individual GS family and have narrow applicability to screen for novel substrate specificity. The first universal method to screen GS libraries (˜104/day) using glycosyl fluoride as the sugar donor was a pH based assay (Ben-David, et al., 2008, Chem. Biol. 15(6):546-551). Here, hydrofluoric acid, a by-product of the GS reaction, was detected by a pH sensitive color indicator. A chemical probe that reacts specifically to the fluoride anion to generate a fluorophore has been used recently to screen small GS libraries (˜102/day) (Andres, et al., 2014, Biochem. J. 458(2):355-363). However, to increase the probability of finding rarer GS mutants, screening techniques capable of handling much larger mutant libraries (106-103 mutants) are necessary. In one aspect, FACS based HTS methods alleviate the need to lyse cells, isolate plasmids, and retransform cells for iterative screening of much larger libraries. Directed evolution experiments for GSs are necessary to identify mutations both within and outside the active site region that can increase catalytic efficiency by >102-103 fold. The challenge is to use substrates without directly incorporating fluorophore tags to monitor GS reactions that typically bias substrate specificity. Computational Design of Engineered Enzymes While computation is not required for design of novel enzyme, a semi-rational approach, combining computational insight into reaction mechanisms with experimental methods (such as directed evolution and mutational screening) can be used (FIG. 5A-5E). This combined approach can increase the success rate of experimental efforts. Computational approaches can harness the variety of experimental data available, from genomic sequences to three-dimensional structural characterization. The genomic data available has led to a variety of enzyme classification tools based on sequence similarity, which are harnessed in databases such as the CAZyme database to group enzymes into families that are likely to have similar activities, and building phylogenetic trees that can be used to postulate enzyme evolution and their implications for enzyme design. However, sequence information in isolation is generally insufficient to predict enzyme function. Structural information, usually from X-ray crystallography of the enzyme to be studied or built from a solved structure of a homologous enzyme, greatly aids in proper enzyme characterization, improving sequence alignment and providing clues as to which residues perform different functions. However, crystallization requires modifications of the environment and native substrates cannot be trapped in active conformations with WT enzymes. Instead, substrate mimics and/or inactive mutants can be co-crystalized to aid in active-site identification. Computational atomistic models can be generated using this experimental data, and the substrates and proteins modified to represent native molecules. These models of the native system can be used in molecular dynamics (MD) simulations to elucidate mechanisms and advance our understanding of structure-function relationships, using experimental activity studies to validate the models. Such simulations offer advantages over static models (such as Rosetta-based or docking models), as dynamic models can account for how the interactions between proteins and substrates lead enzymes to adapt to new, low energy conformations over the course of reaction, and thus provide more accurate predictions to direct targeted experiments for engineering more efficient enzymes. Additionally, the validated models can be used as a basis to test dynamic behavior of new mutants and/or non-native substrates can then be performed with the validated models. In certain embodiments, a traditional Congo red dye assay for carboxymethyl cellulose (CMC) added to agar plates can be used for HTS of E. coil colonies that express active vs. inactive GHs. This allows one to identify protein mutants that have significant transglycosylation vs. hydrolytic activity on CMC based on ‘zone clearing’ about the colonies. However, this method cannot be used for screening GS capable of using activated sugar donors like β-glycosyl azide to synthesize non-glucosyl glycans. Aspects of the present disclosure are described elsewhere herein. Definitions As used herein, each of the following terms has the meaning associated with it in this section. Unless defined otherwise, all technical and scientific terms used herein generally have the same meaning as commonly understood by one of ordinary skill in the art to which this disclosure belongs. Generally, the nomenclature used herein and the laboratory procedures in animal pharmacology, pharmaceutical science, and molecular biology are those well-known and commonly employed in the art. It should be understood that the order of steps or order for performing certain actions is immaterial, so long as the present teachings remain operable. Any use of section headings is intended to aid reading of the document and is not to be interpreted as limiting; information that is relevant to a section heading may occur within or outside of that particular section. All publications, patents, and patent documents referred to in this document are incorporated by reference herein in their entirety, as though individually incorporated by reference. In the application, where an element or component is said to be included in and/or selected from a list of recited elements or components, it should be understood that the element or component can be any one of the recited elements or components and can be selected from a group consisting of two or more of the recited elements or components. In the methods described herein, the acts can be carried out in any order, except when a temporal or operational sequence is explicitly recited. Furthermore, specified acts can be carried out concurrently unless explicit claim language recites that they be carried out separately. For example, a claimed act of doing X and a claimed act of doing Y can be conducted simultaneously within a single operation, and the resulting process will fall within the literal scope of the claimed process. In this document, the terms “a,” “an,” or “the” are used to include one or more than one unless the context clearly dictates otherwise. The term “or” is used to refer to a nonexclusive “or” unless otherwise indicated. The statement “at least one of A and B” or “at least one of A or B” has the same meaning as “A, B, or A and B.” As used herein, the term “about” will be understood by persons of ordinary skill in the art and will vary to some extent on the context in which it is used. As used herein, “about” when referring to a measurable value such as an amount, a temporal duration, and the like, is meant to encompass variations of ±20%, ±10%, ±5%, ±1%, or ±0.1% from the specified value, as such variations are appropriate to perform the disclosed methods. The following notation conventions are applied to the present disclosure for the sake of clarity. In any case, any teaching herein that does not follow this convention is still part of the present disclosure, and can be fully understood in view of the context in which the teaching is disclosed. Protein symbols are disclosed in non-italicized capital letters. As non-limiting example, “CelE” refer to the protein. Notations about mutations are shown as uppercase text. As non-limiting example, “E316G” refer to mutated site 316, where in a glutamic acid residue is replaced with a glycine residue. As used herein the terms “alteration,” “defect,” “variation,” or “mutation” refer to a mutation in a gene in a cell that affects the function, activity, expression (transcription or translation) or conformation of the polypeptide it encodes, including missense and nonsense mutations, insertions, deletions, frameshifts and premature terminations. As used herein, the terms “conservative variation” or “conservative substitution” as used herein refers to the replacement of an amino acid residue by another biologically similar residue. Conservative variations or substitutions are not likely to change the shape of the peptide chain. Examples of conservative variations, or substitutions, include the replacement of one hydrophobic residue such as isoleucine, valine, leucine or methionine for another, or the substitution of one polar residue for another, such as the substitution of arginine for lysine, glutamic for aspartic acid, or glutamine for asparagine. As used herein, the terms “effective amount,” refer to a nontoxic but sufficient amount of an agent to provide the desired results. That result may be enhancing the rate of reaction, increasing purity of the product, increasing the yield of the product As used herein, the term “fragment,” as applied to a nucleic acid, refers to a subsequence of a larger nucleic acid. A “fragment” of a nucleic acid can be at least about 15, 50-100, 100-500, 500-1000, 1000-1500 nucleotides, 1500-2500, or 2500 nucleotides (and any integer value in between). As used herein, the term “fragment,” as applied to a protein or peptide, refers to a subsequence of a larger protein or peptide, and can be at least about 20, 50, 100, 200, 300 or 400 amino acids in length (and any integer value in between). “Instructional material,” as that term is used herein, includes a publication, a recording, a diagram, or any other medium of expression that can be used to communicate the usefulness of the nucleic acid, peptide, and/or compound of the disclosure in the kit for identifying or alleviating or treating the various diseases or disorders recited herein. “Isolated” means altered or removed from the natural state. For example, a nucleic acid or a polypeptide naturally present in a living animal is not “isolated,” but the same nucleic acid or polypeptide partially or completely separated from the coexisting materials of its natural state is “isolated.” An isolated nucleic acid or protein can exist in substantially purified form, or can exist in a non-native environment such as, for example, a host cell. An “oligonucleotide” or “polynucleotide” is a nucleic acid ranging from at least 2, in certain embodiments at least 8, 15 or 25 nucleotides in length, but may be up to 50, 100, 1000, or 5000 nucleotides long or a compound that specifically hybridizes to a polynucleotide. As used herein, the term “polypeptide” refers to a polymer composed of amino acid residues, related naturally occurring structural variants, and synthetic non-naturally occurring analogs thereof linked via peptide bonds. As used herein, “substantially purified” refers to being essentially free of other components. For example, a substantially purified polypeptide is a polypeptide that has been separated from other components with which it is normally associated in its naturally occurring state. Non-limiting embodiments include 95% purity, 99% purity, 99.5% purity, 99.9% purity and 100% purity. As used herein, the term “wild-type” refers to a gene or gene product isolated from a naturally occurring source. A wild-type gene is most frequently observed in a population and is thus arbitrarily designed the “normal” or “wild-type” form of the gene. In contrast, the term “modified” or “mutant” refers to a gene or gene product that displays modifications in sequence and/or functional properties (i.e., altered characteristics) when compared to the wild-type gene or gene product. Naturally occurring mutants can be isolated; these are identified by the fact that they have altered characteristics (including altered nucleic acid sequences) when compared to the wild-type gene or gene product. Ranges: throughout this disclosure, various aspects of the present disclosure can be presented in a range format. It should be understood that the description in range format is merely for convenience and brevity and should not be construed as an inflexible limitation on the scope of the present disclosure. Accordingly, the description of a range should be considered to have specifically disclosed all the possible subranges as well as individual numerical values within that range. For example, description of a range such as from 1 to 6 should be considered to have specifically disclosed subranges such as from 1 to 3, from 1 to 4, from 1 to 5, from 2 to 4, from 2 to 6, from 3 to 6 etc., as well as individual numbers within that range, for example, 1, 2, 2.7, 3, 4, 5, 5.3, and 6. For example, a range of “about 0.1% to about 5%” or “about 0.1% to 5%” should be interpreted to include not just about 0.1% to about 5%, but also the individual values (e.g., 1%, 2%, 3%, and 4%) and the sub-ranges (e.g., 0.1% to 0.5%, 1.1% to 2.2%, 3.3% to 4.4%) within the indicated range. The statement “about X to Y” has the same meaning as “about X to about Y,” unless indicated otherwise. Likewise, the statement “about X, Y, or about Z” has the same meaning as “about X, about Y, or about Z,” unless indicated otherwise. This applies regardless of the breadth of the range. The disclosure provides a method of determining if a protein has transglycosylase activity. In certain embodiments, the method comprises contacting the protein with an azido glycosyl donor and a glycosyl acceptor to form a system, and measuring any change in azide concentration in the system. In certain embodiments, the azido glycosyl donor is substituted with an azido group at an anomeric carbon. In other embodiments, the azido glycosyl donor is substituted with an azido group at a non-anomeric carbon. In certain embodiments, the measurement of azide concentration comprises measurement of the concentration of an inorganic azide. In other embodiments, the measurement of azide concentration comprises the measurement of the concentration of an organic substituted azide, including azido glycosyl species. In certain embodiments, the measuring step comprises contacting the system with a reagent comprising a strained alkyne coupled to a dye, under conditions that allow for reaction of the strained alkyne with any azide or azido compound present in the system. In certain embodiments, the reagent comprises bicyclo[6.1.0]nonyne (BCN), dibenzocyclooctyne (DBCO), or any other strained alkyne. In certain embodiments, the reagent comprises 5-carboxytetramethylrhodamine (5-TAMRA), 6-carboxytetramethylrhodamine (6-TAMRA), or any combinations thereof. In certain embodiments, the strained alkyne and the dye are covalently linked by a linker in the reagent. In certain embodiments, the linker comprises a polyethylene glycol linker. In certain embodiments, the measuring step uses as a control a protein that has no measurable transglycosylase activity or has a known transglycosylase activity. In certain embodiments, the protein is a mutated glycosyl hydrolase (GH). In certain embodiments, the protein is expressed in a cell. In certain embodiments, the cell comprises E. coli or Pichia pastoris. In certain embodiments, the system is within the cell (intracellular). In certain embodiments, the measuring step comprise monitoring fluorescence of the system. In certain embodiments, fluorescence activated cell sorting (FACS) is used to separate individual cells by measured fluorescence. In certain embodiments, the method is configured for high-throughput screening. In certain embodiments, disclosed herein, are mutant polypeptide amino acid sequences of WT TmAfc-0306_(SEQ ID NO:1) comprising the mutation D224G (SEQ ID NO:2) and further comprising at least one additional mutation. In certain embodiments, the at least one additional mutation of the mutated construct (SEQ ID NO:2) is selected from the group consisting of L15K, N70D, A366V, T392S, K395N, D400A, T413P, I428T, and T429P. In other embodiments, the at least one additional mutation of D224G (SEQ ID NO: 2) is selected from the group consisting of L15K-N7OD (SEQ ID NO:3); N70D-T392S (SEQ ID NO:4); N70D-T392S-A366V-K395N (SEQ ID NO:5); N70D-T392S-D400A (SEQ ID NO:6); N70D-T3925-1428T (SEQ ID NO:7); and N70D-D400A-T413P-T429P (SEQ ID NO:8). Those skilled in the art will recognize, or be able to ascertain using no more than routine experimentation, numerous equivalents to the specific procedures, embodiments, claims, and examples described herein. Such equivalents were considered to be within the scope of this disclosure and covered by the claims appended hereto. For example, it should be understood, that modifications in reaction and assaying conditions with art-recognized alternatives and using no more than routine experimentation, are within the scope of the present application. It is to be understood that wherever values and ranges are provided herein, all values and ranges encompassed by these values and ranges, are meant to be encompassed within the scope of the present disclosure. Moreover, all values that fall within these ranges, as well as the upper or lower limits of a range of values, are also contemplated by the present application. The following examples further illustrate aspects of the present disclosure. However, they are in no way a limitation of the teachings or disclosure of the present disclosure as set forth herein. EXPERIMENTAL EXAMPLES The disclosure is further described in detail by reference to the following experimental examples. These examples are provided for purposes of illustration only, and are not intended to be limiting unless otherwise specified. Thus, the disclosure should in no way be construed as being limited to the following examples, but rather, should be construed to encompass any and all variations which become evident as a result of the teaching provided herein. Without further description, it is believed that one of ordinary skill in the art can, using the preceding description and the following illustrative examples, practice the claimed methods of the present disclosure. The following working examples therefore, specifically point out the preferred embodiments of the present disclosure, and are not to be construed as limiting in any way the remainder of the disclosure. Methods Gene Synthesis and Cloning A model GH family 29 fucosidase enzyme, referred to as Tm-alpha-fucosidase (TmAfc) from a hyperthermophile Thermotoga maritima was selected for modification. The native (or wild type) gene Tm0306_that encodes TmAfc was optimized for E. coli expression and custom synthesized with AsiSI and BamH1 restriction sites specific flanking residues in pUC57 by Genscript Biotech Corporation (Piscataway, N.J.). The Tm0306_gene was sub-cloned from Genscript's pUC57 vector into our customized pEC vector (with T5 promotor & Kanamycin selection marker) using standard restriction cloning. The catalytic nucleophile of Tm0306_ (D224) was independently mutated into alanine (D224A), serine (D224S), and glycine (D224G) using standard site-directed mutagenesis protocols. For mutagenesis, 0.5 μM of forward and reverse primers for mutagenesis were mixed with 20 ng of plasmid DNA in a 10 μl reaction volume. The reaction was carried out using 1× Master Mix (Phusion DNA polymerase, 200 μM dNTPs, 1× Phusion HF buffer, 1.5 mM MgCl2) with 5% DMSO and the reaction volume was made up to 10 μl by adding nuclease free PCR water. Amplification was confirmed by gel electrophoresis before the PCR amplified reaction mixtures were digested with 10 U of Dpn1 enzyme (New England Biolabs) at 37° C. for 1 hour. The Dpn1 digested mixture was transformed into E. Cloni 10 g competent cells (Lucigen, Wis.) using the Zymo transformation kit and plated onto LB agar plates with appropriate selection marker (Kanamycin). Several random colonies were selected, plasmid DNA was extracted, and verified by DNA sequencing (Genscript, Piscataway, N.J.). Tm Primer name Primer sequence (° C.) Tm0306_D224A_Forward (SEQ ID NO: 9) 69 GATGTTCTGTGGAACGCC ATGGGTTGGCCGGAG Tm0306_D224A_Reverse (SEQ ID NO: 10) 69 CTCCGGCCAACCCATGGC GTTCCACAGAACATC Tm0306_D224S_Forward (SEQ ID NO: 11) 67.3 GATGTTCTGTGGAACTCC ATGGGTTGGCCGGAG Tm0306_D224S_Reverse (SEQ ID NO: 12) 67.3 CTCCGGCCAACCCATGGA GTTCCACAGAACATC Tm0306_D224G_Forward (SEQ ID NO: 13) 69 GATGTTCTGTGGAACGGC ATGGGTTGGCCGGAG Tm0306_D224G_Reverse (SEQ ID NO: 14) 69 CTCCGGCCAACCCATGCC GTTCCACAGAACATC Protein Expression and Purification Sequence verified wild type (Tm0306_WT) and corresponding nucleophile mutant (Tm0306_D224A/S/G) DNA plasmids were transformed into E. coli BL21 (DE3) competent cells and plated onto LB agar plates with 50 μg/ml kanamycin. Individual colonies were picked to inoculate a 50 ml starter culture of LB media supplemented with kanamycin antibiotic (50 μg/ml) and incubated at 37° C. for 12-16 hours. Overnight grown cultures were transferred into 1000 ml LB media containing 50 μg/ml kanamycin and grown at 37° C. until the culture density reached an OD600 of 0.4-0.8. The protein expression was then induced using 0.5 mM Isopropyl β-D-1-thiogalactopyranoside (IPTG) and cultures were incubated at 25° C. for 20 hours. The cell pellets were recovered by centrifugation and stored in freezer until needed. The cell pellets were suspended in lysis buffer (20 mM sodium phosphate, 500 mM NaCl and 20% glycerol, pH: 7.4) in a 1:5 ratio of cells to buffer solution (total weight basis), along with protease inhibitor cocktail (1 μM E-64, 0.5 mM benzamidine and 1 mM EDTA) and lysozyme (10 μg/m1) and lysed by sonication on ice. The lysed pellets were then centrifuged and the cell lysate supernatant enriched in the desired soluble protein was recovered. The N-terminal his-tagged proteins of interest were separated from the other undesired E. coli proteins using an IMAC (Ni-immobilized metal affinity chromatography) column using the NGC-FPLC system (Bio Rad, Hercules, Calif.). Briefly, the Ni-IMAC column was equilibrated with the IMAC binding buffer (100 mM MOPS, 10 mM imidazole, 500 mM NaC1, pH: 7.4). Next, the cell lysate supernatant was loaded onto the column and the IMAC binding buffer was run through the column to remove any non-specifically bound proteins from the column. The protein of interest was next eluted with the IMAC elution buffer (100 mM MOPS, 500 mM imidazole, 500 mM NaC1, pH 7.4). The protein was buffer exchanged using desalting columns (GE Healthcare, Catalog number: 17-0851-01) into 10 mM of 2-morpholin-4-ylethanesulfonic acid or IVIES at pH 6. The purified protein concentration was estimated using the Spectradrop UV spectrophotometer (SpectraMax M5e) based on 280 nm absorbance. Purity of all enzymes was confirmed by SDS-PAGE based on gel densitometric analysis using pre-cast stain-free (Bio-Rad) protein electrophoresis gels. Fucosidase Activity and Chemical Rescue Assays The activity of the purified enzymes Tm0306_WT, Tm0306_D224A, Tm0306_D224S, and Tm0306_D224G was evaluated using pNP-F (4-nitrophenol α-fucopyranoside) as substrate procured from Carbosynth Limited. In each experiment, 1 μg of protein was added to 2 mM pNP-F added in a reaction buffer containing 50 mM MES pH 6 and incubated at 60° C. for 1.5 hours. Blank wells with pNP-F alone were taken as buffer/substrate but without the added proteins as controls. Three replicates were taken for each reaction mixture. After 1.5 hours of the reaction, 100 μL of the reaction mixture was transferred to a transparent 96-well microplate along with 100 μL of 1 M NaOH and the absorbance was measured at 410 nm using a UV/Vis spectrophotometer (SpectraMax M5e) to determine total released pNP absorbance upon substrate hydrolysis. A pNP calibration curve was built to find the relationship between the measured absorbance and estimated concentration. In order to recover or ‘rescue’ the hydrolytic activity of the hydrolytically inactive nucleophile mutants, high concentrations of external nucleophiles like sodium azide and sodium formate (2 M each) were additionally added to reaction mixtures and incubated at 60° C. for 2 hours. After the reaction was completed, 30 μL of the reaction mixture was transferred to a transparent 96-well microplate and mixed with 70 μL of DI water and 100 μL of 0.1 M NaOH. The absorbance was measured at 410 nm using a UV/Vis spectrophotometer (SpectraMax M5e). Glycosynthase In Vitro Activity Assays For evaluating the glycosynthase activity of Tm0306_WT and Tm0306_D224G, 40 μg of the protein was added to a mixture of 10 mM β-L-fucopyranosyl azide (Catalog number: 66347-26-0, Chemily Glycosciences) and 50 mM pNP-0-D-Xylose (Carbosynth Limited) and incubated at 60° C. for 24 hours in 50 mM IVIES buffer pH 6.0. Two replicates were taken for each reaction mixture. The reaction mixture was then analyzed using Thin Layer Chromatography (TLC) using Silica Gel 60 F254 TLC plates from Merck. The mobile phase used for TLC was ethyl acetate: methanol: water (at 70:20:10 v/v ratios). Standards were also run on the TLC plate to determine the unknown detected spots in reaction sample based on retention factor (Rf) value. The plate was epi-illuminated and directly imaged under UV light at wavelength λ=305 nm to visualize pNP and pNP-containing compounds. The plates were then sprayed with visualization solution containing 0.1% orcinol dye in 10% H2SO4, then dried and heated at 100° C. for 15 min to visualize reducing sugars and acid-labile sugars. In Vitro Strain Promoted Azide-Alkyne Cycloaddition (SPAAC) Reaction DBCO-PEG4-Fluor, a commercially available click-chemistry reagent comprising a red fluorophore dye and a dibenzyocyclooctyne moiety connected by a PEGylated linker, was reacted with either sodium azide (inorganic azide) or β-D-glucopyranosyl azide (organic azide) at 37° C. in 1× pH 7.4 PBS buffer for a total reaction time of 5 hours. The strain-promoted azide-alkyne cycloaddition (SPAAC) reaction kinetics were monitored continuously during the 5 hour period. Exogenous Free Azide Rhodamine-B Fluorophore Studies 200 μM Rhodamine-B was mixed with 400 μM of azide (independently sodium azide and β-D-glucopyransoyl azide) in 1× PBS buffer pH=7.4. Separately, 200 μM Rhodamine-B in 1× PBS buffer pH=7.4 without an azide was taken as the Rhodamine-B only control. An azide only control was prepared with 400 μM of an azide independently in 1× PBS buffer pH=7.4 without Rhodamine-B. Each reaction was incubated at 37° C. for 3 hours and the fluorescence spectra for the each respective solution was recorded every 30 minutes at 550 nm excitation, 570 nm auto cutoff and 590 nm emission in a UV spectrophotometer SpectraMax M5e. Respective azides were mixed with Rhodamine-B and the mixture fluorescence was recorded at various time points at 550 nm excitation, 570 nm auto cutoff and 590 nm emission using UV spectrophotometer Spectra Max M5e. Exogenous Free Triazole Rhodamine-B Fluorophore Studies 200 μM of DBCO-NHS was mixed with 400 μM azides (sodium azide and β-D-glucopyransoyl azide) in 1× PBS buffer pH=7.4 to allow the SPAAC reaction to take place. Here, 200 μM of DBCO-NHS with 1× PBS buffer pH=7.4 without azides was taken as the DBCO-NHS control. Azides were taken with 1× PBS buffer pH=7.4 without DBCO-NHS as azide controls. Only 1× PBS buffer pH=7.4 was taken as the blank for the reaction. The reaction was incubated at 37° C. for 200 min at 400 rpm. The SPAAC reaction was monitored by absorbance at 309 nm at varioius timepoints. After 200 mins total reaction time, 200 μM of Rhodamine-B dye was added to all wells, including controls, and incubated at 37° C. while constantly measuring fluorescence at 550 nm excitation, 570 nm auto cutoff and 590 nm emission using a spectrophotometer SpectraMax M5e for various incubation times ranging from 0-120 minutes from the point of addition of the dye. Glucosyl- and Fucosyl-Azide
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