This chapter surveys various approaches for determining the structures of complex glycans. Specific chemical, enzymatic, and other analytical strategies as well as mass spectrometric (MS) and nuclear magnetic resonance (NMR) spectroscopic methods that lead to complete glycan sequence determination are described. Finally, methods for solving the three-dimensional structures of glycans are considered.
The primary structure of a glycan is defined not only by the nature and order of constituent monosaccharides, but also by the configuration and position of glycosidic linkages and the nature and location of nonglycan substituents (see Chapter 2). For a typical mammalian glycoprotein, the aim is often to identify the correct structure from a range of known or predictable candidate structures. For glycans from bacteria or less well-characterized organisms, it is hard to make predictions, and structural determinations are performed without any assumptions. Choice of methodology is often dictated by the amount and purity of material available and its source (e.g., from tissue samples or cultured cell lines). If quantities are not limiting, the complete primary structure may be determined. In most situations where purity and/or amounts are not optimal, partial characterization will usually be possible. The sensitivity of methods for glycan structural analysis continues to increase with technological advances. Many of these techniques are referred to by acronyms (Table 47.1).
DETECTION OF GLYCANS
Methods for initial glycan detection in glycoconjugates include direct chemical reactions with constituent monosaccharides, metabolic labeling with either radioactive or chemically reactive monosaccharides, and detection with specific lectins or antibodies. A general method for detecting the presence of glycans on proteins involves periodate oxidation of vicinal hydroxyl groups followed by Schiff base formation with amine- or hydrazide-based probes (see Chapter 2 for an example of this reaction). This chemical modification procedure, also referred to as the periodic acid–Schiff (PAS) reaction, can be used to identify glycoproteins in gels. Commercially available kits allow detection of 5–10 ng of a glycoprotein using the periodate reaction with subsequent amplification by means of biotin hydrazide/streptavidin-alkaline phosphatase.
Lectin overlays of blots of SDS-PAGE gels can also be used to detect the presence of specific glycans with comparable sensitivity and greater specificity. For example, the agglutinins from Sambucus nigra (SNA) bind to glycans that terminate in α2-6-linked sialic acid. Lectins specific for terminal fucose, galactose, N-acetylgalactosamine, and N-acetylglucosamine are also commercially available (see Chapter 45).
Metabolic labeling of glycoconjugates with radioactive sugars is another powerful tool for detecting glycans and determining the composition of their attached glycans (Figure 47.1). Cells incubated in medium containing 3H- or 14C-labeled monosaccharides will incorporate the label into the glycan chains of glycoconjugates. The radiolabeled molecules can be detected following gel electrophoresis (SDS-PAGE) or thin-layer chromatography (TLC) by autoradiography or fluorography. Proteins with a glycosylphosphatidylinositol (GPI) anchor may also be specifically labeled with radioactive precursors such as myo-inositol or ethanolamine (see Chapter 11). Glycosaminoglycan (GAG) chains on proteoglycans (see Chapter 16) can be metabolically labeled with 35 SO4- or [3H]-glucosamine and separated from other glycoproteins by ion-exchange chromatography or cetylpyridinium chloride/ethanol precipitation.
Another approach is to transfer a radioactive label in vitro from a radioactive nucleotide sugar to glycans on glycoproteins or cells using a purified glycosyltransferase. For example, the O-GlcNAc modification (see Chapter 18) may be detected by the transfer of labeled galactose using β4-galactosyltransferase and UDP-[3H]-galactose. Purified glycosyltransferases may also be used to label cells by modifying terminal sugars exposed on the glycans of cell-surface glycoproteins.
Radiolabels can also be introduced by chemical reactions that are selective for the structural features of specific glycan types. For example, terminal sialic acids have a unique arrangement of hydroxyl groups on their glycerol side chain that distinguishes this monosaccharide from the others found in mammalian glycans. Mild periodate oxidation of such terminal sialic acid side chains creates aldehyde groups that can be reduced by subsequent treatment with sodium [3H]-borohydride, thereby labeling the sialic acids (Figure 47.1).
The use of radiolabeling strategies for structural analysis also has the advantage of being easy to perform and monitor. Labeled glycans can be isolated by the same methods used for unlabeled sugars with the advantage that radiometric purity is much easier to achieve, and sufficient incorporation is usually obtained for further analysis. However, the information obtained from such analyses is limited, and a complete structural identification generally requires the isolation of unlabeled material.
Metabolic labeling can also be performed with synthetic monosaccharides that are modified with chemically reactive groups. For example, the azido monosaccharide N-azidoacetylmannosamine is converted by cells to N-azidoacetyl sialic acid, which is incorporated into sialylated glycans in place of the natural sialic acid residue. The azide group can then be selectively reacted with phosphine or alkyne reagents (see Chapter 49) that introduce a fluorescent dye or an affinity probe such as biotin, thereby enabling detection of the sialic acid residue. Commercially available azido analogs of N-acetylgalactosamine and N-acetylglucosamine can be used to label O-GalNAc glycans (see Chapter 9) and O-GlcNAc-modified cytoplasmic and nuclear proteins (see Chapter 18), respectively.
Glycoproteins and Mucins
During gel electrophoresis, a glycosylated protein typically presents as one or more diffuse bands, which result from heterogeneity in the glycan. Visualized by protein-staining reagents, this phenomenon is often the first indication of the presence of glycans. Some mucins of very high molecular weight do not enter ordinary gels or, if they do, they migrate as heterogeneous smears. Agarose gels or combination polyacrylamideagarose gels may be useful in this situation. Several analytical options are available to investigate the presence of glycans further, for example, the classical PAS stain. Treatment of glycoproteins with endoglycosidases is another option (e.g., peptide-N-glycosidase F, endoglycosidase F2, and endoglycosidase H; see Table 47.2), and if it results in a mobility change for one or more of the bands on the gel, the presence of N-glycans is indicated. O-Sialoglycoprotease can be used for the identification of glycoproteins or mucins containing clustered sialylated O-glycans, as such glycoconjugates are specifically degraded by this protease. Removal of individual sugars by exoglycosidases such as sialidase or β-galactosidase may also result in a mobility change, depending on the number of residues removed. However, not all glycans may be detected by these treatments due to resistance to the enzymes used. Such resistance can result from modifications to glycan hydroxyl groups (e.g., sulfation, acetylation, or phosphorylation; see Chapter 2), glycosidic linkages that are not recognized by the enzymes, or steric inaccessibility of the glycan. Complete removal of N- and O-glycans can be achieved by chemical treatments (e.g., hydrazinolysis, β-elimination, or hydrogen fluoride treatment), but peptide damage usually precludes further analysis by gel electrophoresis. Aspects of the glycan may also be modified (e.g., O-acetyl groups may be lost).
Proteoglycans typically contain more glycan than protein (see Chapter 16). They may be separated by agarose gel electrophoresis and by ion-exchange chromatography, which separates on the basis of the charge conferred by sulfate groups. Treatment of proteoglycans with GAG lyases (Table 47.2) will produce a shift in mobility on a gel, condensing the proteoglycan smear into discrete bands. After the removal of much of the glycan portion, antibodies that recognize the remaining structures (“stubs”) may be used in western analysis. The lyases cleave a 4,5 unsaturated uronic acid at the nonreducing end. Anti-“stub” antibodies recognize the sulfation of the penultimate N-acetylglucosamine or N-acetylgalactosamine residue.
Typically, the analysis of glycolipid glycans by NMR or mass spectrometry is preceded by their purification using chromatographic methods. Mixtures of glycolipids can be fractionated by TLC, and staining of TLC plates with glycan-reactive reagents may allow detection of individual glycolipids. Using different reagents, it is possible to recognize gangliosides (e.g., resorcinol-HCl detects sialic acids) or bands that contain only neutral monosaccharides (e.g., orcinol-sulfuric acid detects all monosaccharides). Reagents are also available for the detection of sulfate and phosphate groups on glycolipids. Some pre-purification of the crude extract is usually preferred (e.g., Folch partitioning and ion-exchange chromatography). These procedures separate nonpolar or nonionic lipids from polar lipids (e.g., glycosphingolipids) and those that contain charged groups (e.g., gangliosides, phospholipids, and sulfatides). Following TLC or HPLC separation of the enriched mixture, target glycolipids may be detected more easily by glycan-reactive reagents. It is also common practice to deduce the presence of specific sugars by evaluating the shifts produced in the migration position of a band following a chemical or enzymatic treatment. Glycolipids on TLC plates can also be detected by reagents that recognize specific glycan features including monoclonal antibodies, lectins, or even intact microorganisms expressing glycan-binding receptors (see Chapter 45). More detailed structural features may be identified by running the TLC in a second dimension following a specific treatment. On a larger scale, glycolipids are separated using column chromatography or by HPTLC on silica plates.
GPI-anchored proteins (see Chapter 11), with their lipid, protein, and glycan moieties, have unique physicochemical properties that can be exploited for detection purposes. The nonionic detergent Triton X-114 at low temperature (4ºC) extracts soluble and integral membrane proteins as well as GPI-anchored proteins. When the solution is warmed, two phases separate, and GPI-anchored and other amphiphilic proteins remain associated with the detergent-enriched phase. GPI-specific phospholipases can be used to cleave GPI anchors for further characterization. Successful cleavage by GPI-specific phospholipases can be assessed by subsequently analyzing samples by SDS-PAGE, because removal of the GPI anchor causes a shift in molecular mass. This is a common diagnostic method for identifying the presence of a GPI anchor on a protein of interest. Another method is to treat the GPI-anchored protein with nitrous acid, which cleaves the unsubstituted glucosamine residue that links the glycan to the phosphatidylinositol (PI).
Plant and Bacterial Polysaccharides
This family of glycans contains many structures, including homo- and heteropolysaccharides, neutral and ionic polysaccharides, and linear and branched structures, with widespread molecular sizes ranging from a few monosaccharide units to thousands (see Chapters 20 and 22). These polysaccharides are typically extracted with water, salts, chaotropic agents, or detergents and are isolated by precipitation with alcohols. Detection is based on refractive index or colorimetric reactions, because sample quantity is not usually a limitation.
RELEASE, PROFILING, AND FRACTIONATION OF GLYCANS
Once the presence and general type of glycan has been established, the next challenge is to determine how many structurally different glycans are present in a glycoconjugate. The approach varies, but the answer is generally attained by some type of chromatographic or mass spectrometric profiling. When glycans are released prior to chromatographic profiling, it is necessary first to consider the need for a quantitative release procedure that neither destroys nor structurally alters the glycan. Ideally, information regarding the nature of the linkage between the glycan and its liberated protein or lipid should be retained, although this is not always possible.
Release of Glycans from Glycoconjugates
The glycan moiety of glycosphingolipids can also be removed enzymatically using endoglycoceramidase or chemically by ozonolysis. The profiles obtained often provide glycan structural information based on their similarity to known standards. Chemical approaches suitable for the release of glycans from a protein include hydrazinolysis, which releases both N-glycans and O-glycans or, under controlled conditions, cleaves only the N-glycans. Alkaline borohydride treatment (termed β-elimination) is a procedure that under carefully controlled conditions releases only O-glycans. Complex, hybrid, and oligomannose N-glycans can also be released by the peptide-N-glycosidases PNGase F or PNGase A (often termed N-glycanases) (Figure 47.2). An endoglycosidase termed Endo H may be used for the selective release of oligomannose and hybrid N-glycans, but complex N-glycans are resistant (Figure 47.2). N-Glycans and O-glycans can be obtained nonselectively by degradation of the protein by proteases to generate glycopeptides. GPI anchors may be cleaved from protein by phospholipase treatment or obtained following proteolysis of the protein. Free glycans obtained by all of these methods are subsequently analyzed by HPLC, HPAEC, HPCE, or FACE (Table 47.1). The profiles obtained often provide glycan structural information based on their similarity to known standards.
Profiling of Glycoprotein Glycans
Glycans released from a glycoprotein are usually a complex mixture. Even when only one glycosylation site in the protein is occupied, it can bear many different glycans resulting in many glycoforms of the glycoprotein. Chromatographic profiles are used for comparative studies and to obtain a preliminary indication of the number, relative quantities, and types of glycans present in a glycoprotein.
Profiling strategies are chosen based on the quantity of sample available. For large amounts (>5 mg), HPAEC-PAD or other HPLC-based profiling is feasible, provided the mixture contains fewer than about 50 different glycans. Individual fractions can be analyzed by MS or NMR. Radiolabeling using the chemical or metabolic methods described above is used to enhance the sensitivity of glycan detection. Indeed, scintillation detectors can be directly linked to HPLC equipment to monitor the purification of radiolabeled glycans. Once liberated from their glycoconjugates, glycans with free reducing termini (see Chapter 2) can be chemically labeled with fluorescent tags such as 2-aminopyridine (2-AP), 2-aminobenzamide (2-AB), 2,6-diaminopyridine (DAP), or biotinylated 2,6-diaminopyridine (BAP), providing detection sensitivity that rivals the level achieved with radiolabels. Advantages of this method include more facile purification of the labeled glycans and a wider variety of options for chromatographic separations and analytical techniques.
If a label is introduced at the reducing end, structural information may be obtained by sequential exoglycosidase treatments (Figure 47.2 and Table 47.2) and chromatography to detect shifts in glycan elution or migration (e.g., by paper chromatography, HPLC, or TLC) that indicate susceptibility to the enzyme. Comparison with known standards treated in the same manner allows tentative glycan identification. However, well-characterized standards are difficult to obtain in pure form, and there are nearly always species in a chromatogram that appear at unusual elution times. It is very important to note that separation profiling should not be confused with actual structural analysis, because coelution with a standard does not necessarily connote a structure identical to that standard.
Profiling of Glycosaminoglycans
Structural analysis of GAGs is an area in which methodologies are rapidly improving (see Chapter 16). Molecular size profiles of GAGs can be determined by chromatographic or electrophoretic methods. Various hydrolases and chemical degradation methods (such as nitrous acid deamination) are available to define the class and/or structures of GAG chains further (Table 47.2). Characterization of a heterogeneous sample might be achieved by fingerprinting techniques (such as chromatography or electrophoresis of enzyme-generated oligosaccharides) and analysis of the disaccharide products of exhaustive depolymerization. Where an oligosaccharide of homogeneous sequence is available, various strategies for precise sequencing can be used, including end-labeling, specific enzyme digestions, separation techniques, and mass spectrometry. For example, sequencing of heparan sulfate can be achieved by treatment with heparanase followed by MS or NMR spectroscopy.
MONOSACCHARIDE COMPOSITION ANALYSIS
Some qualitative information concerning the monosaccharide composition of a glycan may be derived from the procedures described above for the detection, release, and profiling of glycans. Conversely, it is often convenient and informative to determine the monosaccharide composition of a glycoconjugate without prior release of glycans. After total hydrolysis of a glycan into its monosaccharide constituents, colorimetric reactions can be used to determine the total amount of hexose, hexuronic acid, or hexosamine in the sample. These approaches only require common reagents and a spectrophotometer, but determination of total glycan content may not always be accurate because of variations in the sensitivities of different linkages to hydrolysis, variations in the degradation of individual saccharides, or a lack of specificity and/or sensitivity in the assays.
Quantitative monosaccharide analysis provides estimated molar ratios of individual sugars and may suggest the presence of specific oligosaccharide classes (e.g., N-glycans vs. O-glycans). The analysis involves the following steps: cleavage of all glycosidic linkages (typically by acid hydrolysis), fractionation of the resulting monosaccharides, detection, and quantification. Since the early 1960s, a variety of gas-liquid chromatography (GLC) methods have been developed to quantify monosaccharides. The most useful involve coupling of GLC and MS for linkage and composition information. These methods are most successful when the monosaccharides are first chemically modified at their hydroxyl and aldehyde groups. Reduction of the aldehyde of a free monosaccharide followed by acetylation of its hydroxyl groups provides a derivative termed the “peracetylated alditol acetate.” These modified monosaccharides can be readily analyzed by GLC and MS and compared with authentic standards. The hydroxyl groups of free monosaccharides generated by glycan hydrolysis can also be converted to trimethylsilyl ethers. These per-O-trimethylsilyl derivatives are widely used for monosaccharide compositional analysis by GLC-MS. Incorporation of an optically pure chiral aglycone (e.g., a [–]-2-butyl group), in combination with trimethylsilylation, allows the GLC separation of the D and L pair of isomers and thus determination of the absolute configuration of each monosaccharide.
Chemical derivatization of monosaccharides was once required for HPLC or GLC separation and analysis. However, in recent years, these classical methods have been supplanted by HPAEC-PAD, which does not require monosaccharide derivatization. Fluorescent derivatives produced by reductive amination (e.g., with 2-AB, 2-AP, or 8-amino-1,3,6-naphthalene trisulfonic acid [ANTS]) became popular for detection by reversed-phase HPLC with online fluorophore-assisted carbohydrate electrophoresis (FACE), or HPCE. For example, tagging sialic acids with a fluorescent compound (1,2-diamino-4,5-methylene-dioxybenzene [DMB]) has allowed an increase in detection sensitivity to the femtomole range.
Monosaccharide compositional analysis can be performed on glycoproteins separated by SDS-PAGE and blotted onto polyvinylidene difluoride (but not nitrocellulose) membranes. The membrane is hydrolyzed, the hydrolysate is easily recovered, and monosaccharides are measured as described above. Depending on conditions, peptide or protein may remain bound to the membrane. Sequential analyses are also possible. For example, sialic acids can be released selectively with mild acid. Strong acid can then be added to release the remaining monosaccharides.
Determination of Linkage Positions
Methylation analysis is a well-established and ingenious approach for determining linkage positions. The principle of this method is to introduce a stable substituent (an ether-linked methyl group) onto each free hydroxyl group of the native glycan. The glycosidic linkages, which are much more labile than the ether-linked methyl groups, are then cleaved by acid hydrolysis, producing individual methylated monosaccharides with free hydroxyl groups at the positions that were previously involved in a linkage. The partially methylated monosaccharides are derivatized to produce volatile molecules amenable to GLC-MS analysis. The most common strategy involves reduction of the monosaccharides to produce alcohols at C-1 (eliminating the formation of ring structures), followed by derivatization (usually acetylation) of free hydroxyl groups. Individual components of the mixture of partially methylated (methyl groups mark the hydroxyl groups that were originally free), partially acetylated (acetyl groups mark hydroxyl groups originally at substituted, linked, or ring-closure positions) monosaccharide alditols can be identified by GLC-MS (Figure 47.3).
Partially methylated alditol acetates are identified by a combination of GLC retention time and electron impact (EI)-MS fragmentation pattern. The fragmentation patterns of similarly substituted isomeric monosaccharides (e.g., aldohexoses) are the same. Thus, definitive identification requires, in addition to the analysis of the MS pattern, the comparison of GLC retention times with those of known standards (e.g., all 2,3,4-tri-O-methyl-hexoses produce the same EI-MS spectrum, but peracetylated 2,3,4-tri-O-methylgalactitol elutes later than peracetylated 2,3,4-tri-O-methylglucitol). This type of analysis identifies terminal residues (they are methylated at every position except the hydroxyl group at C-1 and C-5), indicates how each monosaccharide is substituted including the occurrence of branching points, and allows the determination of the ring size (pyranose p or furanose f) for each monosaccharide. However, methylation analysis gives no sequence information and cannot determine whether a particular linkage is of the α or β anomeric configuration.
Determination of Anomericity
The anomeric configuration of linkages is often determined by NMR spectroscopy (see below) and can also be obtained from sequential exoglycosidase digestions (Table 47.2 and Figure 47.2). Cleavage by α- or β-exoglycosidases indicates the anomericity of specific terminal sugar residues. Cleavage by specific endoglycosidases can give added information regarding internal regions of the glycan. Many glycosidases are specific for both monosaccharide residue and linkage type, allowing detailed structural conclusions, although the number of such enzymes available is limited.
When enough sample is available (typically a milligram or more but see below), the anomericity of a particular monosaccharide residue in a glycan can usually be determined by 1H-NMR spectroscopy. The anomeric resonances (H-1 signals) appear in a well-resolved region of the spectrum and show characteristic doublets with a splitting that is significantly larger for β anomers than for α anomers. Thus, a first glance at the 1H-NMR spectrum typically indicates how many residues there are (by counting anomeric signals) and how many of them belong to each anomeric type. A simple 1H-NMR spectrum can provide the entire primary structure of a glycan if 1H-NMR spectra of well-characterized glycans of related structures are available for comparison. As an example, the 1H-NMR spectrum of a mixture of two triantennary N-glycans obtained from bovine fetuin is shown in Figure 47.4.
Limitations on the use of NMR spectroscopy are the cost of spectrometers and the level of expertise required for interpreting NMR spectra. However, access to high-field (i.e., 500 MHz and above) NMR spectrometers fitted with very sensitive probes (e.g., nano-NMR probes) allows 1H-NMR profiling of individual HPLC fractions using minute quantities of sample (2–5 nmoles of glycan).
NMR spectroscopy is a powerful tool for de novo full structural characterization of a glycan. Because this method is nondestructive, the same sample can later be used for other, destructive approaches (e.g., MS and methylation analysis). Complete structural elucidation requires full assignment of both the 1H- and 13C-NMR spectra of a glycan. This is accomplished by a combination of two-dimensional NMR techniques such as correlation spectroscopy (COSY) and total correlation spectroscopy (TOCSY) for 1H, which allows assignment of the 1H signals of individual monosaccharide residues. After this, the heteronuclear single-quantum coherence (HSQC) experiment can be used to extend the assignment to the 13C spectrum. The key experiment for sequencing is the two-dimensional heteronuclear multiple-bond correlation (HMBC) experiment, which detects a coupling between the anomeric proton and the carbon atom on the opposite side of the glycosidic linkage. However, in instances where there is not enough sample for these two-dimensional NMR experiments (HMBC is not a very sensitive technique), other data are required to complete the structural picture. A less rigorous NMR approach for glycan sequencing relies exclusively on two-dimensional 1H-NMR spectroscopy, using through-space effects (nuclear Overhauser effects [NOEs]) as the sole source of evidence for linking, position, and sequence. Use of a 900-MHz NMR spectrometer and a nanoprobe increases the sensitivity so that microgram amounts of a glycan can be analyzed.
Polysaccharides from bacteria (see Chapter 20) give remarkably good NMR spectra (despite their high molecular weight) due to their internal mobility, and it is often possible to determine the structure of the repeat unit by NMR without need for depolymerization. Figure 47.5 illustrates NMR and MS data for a bacterial polysaccharide that is one of the components of a vaccine. The combination of NMR and MS analyses gives a thorough structural assignment.
The use of EI-MS in monosaccharide composition and linkage analyses is covered above. In this section, three other types of mass spectrometry—fast atom bombardment (FAB), matrix-assisted laser desorption ionization (MALDI), and electrospray ionization (ESI)—are described. All three technologies permit the direct ionization of nonvolatile substances and are applicable to intact glycoconjugates, as well as fragments thereof. Historically, FAB-MS has played an important role in the structural analysis of glycans. However, because of the expense and level of specialized expertise required to operate FAB-MS, this method has been largely supplanted by ESI-MS and MALDI-MS. Among the structural features that can be defined by MS methods are (1) degree of heterogeneity and type of glycosylation (e.g., N-glycan vs. O-glycan; high mannose, hybrid, or complex types, etc.); (2) sites of glycosylation; (3) glycan-branching patterns; (4) the number and lengths of antennae, their building blocks, and the patterns of substitution with fucose, sialic acids, or other capping groups such as sulfate, phosphate, or acetyl esters; (5) complete sequences of individual glycans; and (6) structures of glycolipids, glycopeptides, (lipo)polysaccharides, and GAG-derived glycans.
In the FAB-MS experiment, samples are dissolved in a liquid matrix and ionization/desorption is effected by a high-energy beam of particles fired from an atom or ion gun. High field magnets are the most powerful analyzers for this type of mass spectrometry. In MALDI-MS experiments, the sample is dried on a metal target in the presence of a chromophoric matrix until matrix crystals containing trapped sample molecules are formed. Ionization of the sample is effected by energy transfer from matrix molecules that have absorbed energy from laser pulses. MALDI sources are usually attached to time of flight (TOF) analyzers that can analyze very high-molecular-mass ions (in excess of 200 kD). For ESI-MS, a stream of liquid containing the sample enters the source through a capillary interface, where the sample molecules are stripped of solvent, leaving them as multiply charged species. Electrospray experiments are often performed using instruments with quadrupole analyzers. ESI-MS can be coupled to micro- or nanobore liquid chromatography (LC) permitting on-line LC/ESI-MS analysis. This method is particularly useful when complex mixtures of peptides and glycopeptides are being examined (e.g., after proteolytic digestion of a glycoprotein).
In principle, MS provides two types of structural information–the masses of intact molecules (the molecular ions) and the masses of fragment ions. MALDI-MS is arguably the most sensitive of the three ionization technologies; hence, it is the preeminent technique for screening for molecular ions (“mass mapping”), especially when high throughput and sensitivity are demanded.
Of the three techniques, FAB-MS is the only one that reliably yields fragment ions. The internal energy acquired during molecular ion formation in MALDI and ESI-MS is usually insufficient for fragmentation to occur. To overcome this, most ESI and some MALDI mass spectrometers have two analyzers in tandem, which allows the detection (using the second analyzer) of fragment ions produced after molecular ions selected by the first analyzer undergo collisions with an inert gas in a chamber placed between the two analyzers. These are referred to as collisionally activated MS/MS experiments. One of the most powerful current technologies for MS/MS is the Q-TOF mass spectrometer, which has a quadrupole as the first analyzer and an orthogonal TOF as the second analyzer.
Fourier transform mass spectrometry (FTMS) with electron capture dissociation (ECD) and lower-cost ion traps with electron transfer dissociation (ETD) are cutting-edge technologies that considerably improve MS analyses of complex posttranslational modifications, including glycans. MS instrumentation and methods continue to improve at an astonishing rate.
Although underivatized glycans can be analyzed by FAB-MS and ESI- or MALDI-MS/MS, far superior data are normally obtained if the glycans are derivatized prior to MS analysis. Derivatization methods can be broadly divided into two categories: (1) “tagging” of reducing ends and (2) protection of most or all of the hydroxyl groups. Commonly used tagging reagents include p-aminobenzoic acid ethyl ester (ABEE), 2-AP, 2-AB, and amino-lipids. This type of derivatization facilitates chromatographic purification as explained above and enhances the formation of reducing-end fragment ions in MS and MS/MS experiments. Protection of hydroxyl groups by permethylation is by far the most important type of full derivatization employed in glycan MS (although with accompanying destruction of acetyl esters, some sulfate esters, and glycolyl groups during the derivatization process). In FAB-MS experiments, permethylated derivatives form abundant fragment ions arising from cleavage on the reducing side of each HexNAc residue (usually referred to as A-type ions) whose masses define important structural features of N- and O-glycans, including the types of capping sugars and the presence or absence of poly-N-acetyllactosamine sequences. In MS/MS experiments, additional fragment ions are produced by cleavage on either side of susceptible glycosidic linkages.
Broadly speaking, the unique strengths of MS can be exploited in two general ways in glycobiology. The first way is to obtain detailed characterization of purified individual glycans or mixtures of glycans. In this type of study, it is essential to acquire sufficient rigorous data to define structure unambiguously; many different MS-based experiments will be required, often complemented by NMR, linkage analysis, and profiling of enzyme digests. An example of this type of application is illustrated in Figure 47.5. The second way is for glycomics investigations in situations where it may not be essential to define structures fully and when high-throughput glycomic profiling or mass-mapping procedures are exploited (see Chapter 48).
Mass Spectrometry Profiling Underpins Many Glycomics Investigations
As discussed in Chapter 48, the term “glycome” is used to denote the complement of glycans in a cell or organism. Thus, strictly speaking, the term “glycomics” should mean the study of the full complement of glycans from a defined source. In practice, because glycomics investigations are still in their infancy, they are usually confined to studies of subsets of glycans, for example, the most abundant N- or O-glycans present in a particular cell type or tissue. MS strategies have been devised to screen for the types of N- and O-glycans present in a diverse range of biological material, including body fluids, secretions, organs, and cultured cell lines. These methods are based on the analysis of permethylated derivatives, which yield molecular ions at high sensitivity. Putative structures are assigned to each molecular ion based on the usually unique glycan composition for a given mass and prior knowledge of N- and O-glycan biosynthesis. This is called “glycomic profiling“ and is most conveniently carried out using MALDI because of its high sensitivity. Assignments can be confirmed in a second experiment employing ESI-MS/MS instrumentation by selecting each molecular ion for collisional activation and recording its fragment ion spectrum. If necessary, additional information can be provided by MS experiments on chemical and enzymatic digests, the choice of which is guided by the sequence information provided by mass mapping and MS/MS experiments. These methodologies are illustrated by data from a glycomics analysis of the mouse kidney in Figure 47.6. Of course, none of these approaches to glycomics address the actual localization of a glycan within a cell type of the tissue being extracted for analysis.
Three-dimensional Glycan Structure
Because of the inherent flexibility of glycosidic linkages, most complex glycans do not have a single, well-defined three-dimensional structure in solution. Crystal structures are available for many mono- and oligosaccharides (in the Cambridge Structural Database; http://www.ccdc.cam.ac.uk). Anyone with suitable molecular modeling software can generate approximate models of more complex glycans from these simple sugar structures. Such models can be useful as an aid to thinking about the overall sizes and shapes of glycans, as long as they are not taken too seriously. Full characterization of glycan conformation and dynamics in solution remains an area of active research and is usually based on experimental data from NMR spectroscopy. For example, H-H coupling constant values around the pyranose ring depend on the ring geometry, and quantitative interpretation of NOEs between adjacent monosaccharides can give clues as to the conformational equilibrium around the glycosidic linkage between them. Recently introduced methods, such as conformational restraints derived from residual dipolar couplings in partially ordered media, can also be applied to glycans. A full treatment of modeling glycan structures is beyond the scope of this chapter (see “Further Reading”). Coordinates for three-dimensional structures of glycoproteins in the Protein Data Bank (PDB: http://www.rcsb.org/pdb) usually define only the stub of any attached glycan. Of more direct interest are complexes between proteins and glycans, of which the PDB contains many, including enzymes, lectins, and heparin-binding proteins among others. In this context, the glycan conformations are both well-defined and biologically relevant.
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Barbara Mulloy, Gerald W Hart, and Pamela Stanley.
Cold Spring Harbor Laboratory Press, Cold Spring Harbor (NY)
Mulloy B, Hart GW, Stanley P. Structural Analysis of Glycans. In: Varki A, Cummings RD, Esko JD, et al., editors. Essentials of Glycobiology. 2nd edition. Cold Spring Harbor (NY): Cold Spring Harbor Laboratory Press; 2009. Chapter 47.