Chapter 47 Laboratory Materials And Procedures Case Study


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.


Radiolabeling strategies for the detection of glycans. Metabolic labeling with [3H]- or [14C]-monosaccharides allows detection of any glycoconjugate into which the monosaccharide is incorporated. 35SO4 can be used to trace glycosaminoglycans (GAGs), and (more...)

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 monosaccharideN-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 Anchors

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.


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.


Glycosidases used for structural analysis. (Left) A biantennary N-glycan is shown with exoglycosidases that can be used to remove each monosaccharide sequentially. Exoglycosidases act only on terminal sugars. Also shown are endoglycosidases that remove (more...)

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.


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).


A simple example of methylation analysis, showing a structural motif that may be found in the polysaccharide glycogen. An α1-4-linked glucose chain has an α1-6-linked glucose branch. Successive steps of methylation, hydrolysis, reduction, (more...)

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 (pyranosep or furanosef) 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.

NMR Spectroscopy

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.


1H-NMR spectrum of a mixture of two trisialyl triantennary N-glycans obtained as alditols from bovine fetuin by hydrazinolysis, followed by purification on HPAEC (Table 47.1) and subsequent reduction with sodium borohydride. The spectrum was recorded (more...)

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.


NMR and MS data used in the determination of the structure of the complex pneumococcal capsular polysaccharide 17F. Mild base treatment of the polysaccharide specifically breaks the phosphodiester linkage between rhamnose (Rha) and arabinitol and removes (more...)

Mass Spectrometry

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.


Data from a glycomics study of N-glycans from mouse kidney (courtesy of Anne Dell, Imperial College London). (Top) MALDI-TOF profile of neutral N-glycans released from kidney extracts by peptide N-glycosidase F and permethylated. The glycan structures (more...)

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; 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: 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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  9. Tissot B, Gasiunas N, Powell AK, Ahmed Y, Zhi ZL, Haslam SM, Morris HR, Turnbull JE, Gallagher JT, Dell A. Towards GAG glycomics: Analysis of highly sulfated heparins by MALDI-TOF mass spectrometry. Glycobiology. 2007;17:972–982. [PubMed: 17623722]

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41 A p p e n d i x A This appendix describes the spectroscopic equipment and testing protocols used in the laboratory testing portion of this project. The testing protocols include procedures for the preparation of samples, acquisition of data, and interpreta- tion of results. The following spectroscopic methods were included in the laboratory evaluation: • Fourier transform infrared (FTIR) spectroscopy; • Raman spectroscopy; • X-ray fluorescent (XRF) spectroscopy; • X-ray diffraction (XRD) spectroscopy; • Gel permeation chromatography (GPC) with evaporative light-scattering detectors (ELSD) and ultraviolet (UV) detectors; and • Nuclear magnetic resonance (NMR). The following is a detailed discussion of these methods and their corresponding experimental protocols. Fourier Transform infrared Spectroscopy A comprehensive literature search conducted in Phase 1 of the project identified FTIR spectroscopy as one of the most promising spectroscopic methods for “fingerprinting” con- struction materials. Therefore, a careful evaluation of FTIR spectrometers was included in the laboratory phase. The team employed two different FTIR devices in its testing. The first device, a portable Bruker ALPHA attenuated total reflec- tance (ATR) FTIR spectrometer owned by the research team, tested all of the materials and material components listed in Table A.1. Concurrently, the team used a stationary Nicolet Magna 560 FTIR spectrometer located in the Institute of Material Science (IMS) of the University of Connecticut (UCONN) to examine the same materials. A comparison of the results from the portable and stationary FTIR equipment allowed the team to verify the reliability of the former device and recommend it for further evaluation in the field phase of the project. More details on both devices are given below. Portable Bruker ALPHA Attenuated Total Reflectance Fourier Transform Infrared Spectrometer Equipment Setup The ALPHA FTIR device by Bruker Optics (Figure A.1) has dimensions of 22 × 30 × 25 cm and weighs about 7 kg. It is able to analyze a spectral range from 7,500 cm-1 to 375 cm-1 with a spectral resolution of at least 2 cm-1 (0.9 cm-1 is optional) and an accuracy of 0.01 cm-1. The device is designed to operate at 18°C to 35°C and is powered by 100 to 240 V AC or by a high-capacity battery. Software, including a compre- hensive library of chemical components, is provided with this device. The exchangeable measurement modules include transmission, ATR, and reflection, making it possible to ana- lyze a range of materials (liquid, solid, or gas). The team used the following settings in their laboratory testing: • A laptop PC with OPUS 6.5 infrared (IR) spectroscopy software was provided by Bruker Optics. • The ATR mode was used to acquire the infrared spectra for all materials and their components. • The spectra for each material and component were obtained in the range from 4,000 to 400 cm–1, where most of the chemical components yield detectable absorbance. • The spectral resolution was set to 1.5 cm–1. • Twenty-four scans were run for each sample. • Ambient temperatures during testing ranged between 20°C and 23°C. The team decided to use the ATR cell because, in most cases, it does not require any special sample preparation and also allows for a quick determination of the sample’s spectra Laboratory Equipment and Testing Protocols

42Table A.1. Finalized List of Materials to Be Tested in Phase 2 Material Category Brand Manufacturer Number of States Approving the Material Original State (Liquid, Solid, Powder, Other) Information on Chemical Composition Available (Yes/No) Structural coatings Scotchkote 3M 19 Epoxy Yes Carbozinc 859 RPM International Inc. 18 Epoxy + powder Yes Traffic paints LS50 Epoplex 19 Epoxy Yes All Weather HB-R1 3M 12 Liquid Yes Epoxy adhesives for concrete repair Sikadur Sika Construction 20 Epoxy + liquid Yes Ultrabond 1100 Adhesives Technology Corporation 14 Epoxy + powder Yes Portland cement Type 1, 2, and 3 cement Lafarge North America 25 Powder Yes Type 1, 2, and 3 cement LeHigh 23 Powder Yes Portland cement concrete (PCC) Local Tilcon 5 Solid Yes Air-entraining admixtures for PCC Air Mix 200 Euclid Chemical 25 Liquid Yes Accelerating/retarding admixtures for PCC Eucon Retarder 75 Euclid Chemical 26 Liquid Yes Accelguard 80 Euclid Chemical 14 Liquid Yes Water reducers/plasticizers for PCC ADVA 190 W. R. Grace 23 Liquid Yes Curing compounds for PCC Sealtight 1100-CLEAR W. R. Meadows 13 Liquid Yes Safe-Cure 1200 ChemMasters 9 Liquid Yes Hot-mix asphalt concrete Local Tilcon–Oldcastle Company 5 Solid No Asphalt binders PG 58-28 NuStar NA Viscous liquid No PG 64-22 NuStar NA Viscous liquid No PG 64-28 NuStar NA Viscous liquid No PG 70-28 NuStar NA Viscous liquid No PG 76-22 NuStar NA Viscous liquid No PG 76-34CR NuStar NA Viscous liquid No PG 64-28PPA NuStar NA Viscous liquid No Polymer modifiers for binders Elvaloy 4170 DuPont NA Granules Yes D1101 (SBS) Kraton NA Granules Yes Butonal NX1138 (SB Latex) BASF NA Liquid Yes Asphalt emulsions CRS-1 All States Asphalt Group 18 Liquid No CRS-1P All States Asphalt Group 18 Liquid No Antistripping agents for asphalt concrete Kling Beta 2912 Akzo Nobel 11 Liquid Yes AD-here LOF 65 ArrMaz 12 Liquid Yes Reclaimed asphalt pavement Local Tilcon 50 Solid No Aggregate minerals Local Local quarry—Tilcon NA Solid No Note: NA = not available.

43 Figure A.1. Image of ALPHA ATR FTIR portable spectrometer. without sacrificing the quality of spectrum resolution. The OPUS software accompanying the device is also user friendly and platform independent. Test Sample Preparation The research team tested the materials and components included in Phase 2 in their original physical state (as provided by manufacturer) (i.e., liquid, paste and emulsion, and solid state) (Table A.1). For the vast majority of materials, the sam- ples required no special preparation. Liquid materials, such as structural and traffic paints, portland cement concrete (PCC) curing compounds, PCC chemical admixtures, and anti- stripping agents, were sampled using pipettes or syringes. A spatula was used to sample viscous liquids, such as epoxy pastes for concrete repair and asphalt emulsions, and viscous solids, such as asphalt binders and finely ground powders (e.g., portland cement and zinc filler for Carbozinc 859 struc- tural coating system). A sample of each material and compo- nent was placed on the ATR diamond surface in an amount sufficient to cover the surface (several drops of liquid or approximately 1 g of solid). The following samples required a different preparation procedure: pellet samples [asphalt polymer modifiers, Kraton styrene–butadiene–styrene (SBS), and Elvaloy], granular materials (such as mineral aggregates and hot-mix asphalt mixes), and dry solid samples of epoxy- based paints. In these cases, pressure was applied to ensure full contact between the sample and the ATR diamond sur- face. Finally, in the case of rigid solid pavement markings (Epoplex LS), the sample size had to be reduced to fit the size of the ATR window. Control Sample Preparation The control is a sample of material with a controlled chemical composition that serves as a reference for the test samples. In cases where the quantification of a specific component’s concentration in the designated mixture or solution was an objective (see Table A.2), several control samples were pre- pared by mixing the component of interest with the neat or original material in different proportions. The mixing proce- dures complied with the original procedure provided by the manufacturers to produce the designated material. Testing Procedure A typical FTIR testing procedure for any material included the following routines: • Prepare control sample, if required. • Obtain infrared spectrum of the control sample. • Prepare test sample as explained above. • Obtain infrared spectrum of the test sample. • Compare the test sample spectrum with that of the control sample by using an appropriate method of analysis, as dis- cussed in Chapter 2. Nicolet Magna 560 FTIR Spectrometer Equipment Setup The Nicolet Magna 560 FTIR spectrometer (Figure A.2) is a research-grade benchtop size device operating in a wave- length range between 6,500 and 100 cm-1. It provides a spec- tral resolution of 0.35 cm-1 with a sensitivity of 0.125 cm-1. The device operates at room temperature in the ATR and transmission mode. The objective of using the stationary FTIR spectrometer was to verify the IR spectra acquired by the portable FTIR device. The following settings were used for Nicolet Magna 560 testing: • The IR transmission window mode to obtain spectra for all material and components. • The OMNIC software to analyze the signal.

44 3. Cured epoxy adhesive samples (Ultrabond 1100 and Sika- dur 31) were prepared by mixing ground epoxy powder with KBr powder as a 5 wt% mixture. The mixture was then pressed as a KBr disk. Control Sample Preparation The control sample preparation procedure was the same as for the portable ATR FTIR device. Testing Procedure The testing procedure was the same as for the portable ATR FTIR device. Raman Spectroscopy The search conducted in Phase 1 identified Raman spec- troscopy as a relatively promising spectroscopic method for evaluating construction materials. Although it was found that Raman technology has primarily been used on a nar- rower range of construction materials than the FTIR (mostly for portland cement–related products), it was decided that this method should be pursued for all materials Table A.2. Summary of Phase 2 Objectives for Material–Method Combinations Material Category Method Objective Structural coatings FTIR, Raman, XRF, NMR, GPC Verification of chemical composition Presence of solvents/diluents Traffic paints FTIR, Raman, XRF, NMR, GPC Verification of chemical composition Presence of solvents/diluents Epoxy adhesives FTIR, Raman, XRF, NMR, GPC Verification of chemical composition Portland cement FTIR, Raman, XRF, NMR Verification of cement quality and type PCC FTIR, Raman, XRF, XRD Verification of presence of admixture in PCC mix Quantification of content Chemical admixtures for PCC FTIR, Raman, XRF, NMR, GPC Verification of chemical composition Curing compounds for PCC FTIR, Raman, XRF, NMR, GPC Verification of chemical composition Polymer additives for asphalt binders FTIR, Raman, XRF, NMR, GPC Verification of chemical composition Polymer-modified asphalt binders FTIR, Raman, NMR, GPC Verification of chemical composition Quantification of content Hot-mix asphalt concrete FTIR, NMR, GPC Detection of prohibited chemicals/modifiers (such as motor oil, diesel fuel) Asphalt emulsions FTIR, Raman, NMR, GPC Type and water content Antistripping agents in asphalt concrete FTIR, Raman, NMR Verification of presence in mixture Quantification of content Oxidation in reclaimed asphalt pavement FTIR, NMR, GPC Verification of presence in mixture Quantification of content Aggregate minerals FTIR, XRF, XRD Metal contamination Organic content • The spectra for each material and component were obtained in the range of 4,000 to 400 cm-1 (the same as for the portable ATR FTIR). • The spectral resolution was set at 2 cm-1. • Sixteen scans were performed on each sample. • All spectra were acquired at room temperature. Test Sample Preparation The sample preparation procedures for the stationary IR transmission spectrometer differed from those of the ATR device (portable device). The transmission mode requires either putting the sample between two 25-mm-diameter potassium bromide (KBr) disks or mixing the sample with KBr powder. Accordingly, the following three alternative sample preparation methods were used: 1. Liquid and viscous solid–liquid samples (which include all materials except those mentioned below) were prepared by placing 100 µg of material between two 25-mm-diameter KBr disks. 2. Pellet samples (Kraton SBS and Elvaloy) were dissolved in 1 mL of tetrahydrofuran (THF) solvent as a 10 wt% solution and then dried on a 25-mm-diameter KBr disk.

45 included in the scope of this project. The reasoning behind this was to eliminate the least successful materials and focus on the feasible materials in Raman field testing in Phase 3. Therefore, samples of all materials were shipped to Real- Time Analyzers (RTA) for evaluation in their laboratory facilities. The following segments describe the equipment, sample preparation, and testing protocols used in Raman evaluation. Real-Time Analyzers’ Portable Raman Analyzer Equipment Setup RTA’s portable Raman analyzer (Figure A.3) measures the entire Raman spectrum in each scan, from 3,350 cm-1 to 150 cm-1, with a selectable spectral resolution between 2 and 32 cm-1. According to the manufacturer’s specifications, the device is capable of analyzing any solid or liquid in a very short amount of time (about 10 s) and comes equipped with software installed on a laptop computer. Additionally, it does not require any sample preparation or calibration. The por- table Raman analyzer operates at temperatures from 2°C to 37°C and uses a 5-h rechargeable battery. This analyzer employs interferometry to generate the Raman spectrum, which eliminates the need for analyzer recalibrations. In Phase 2 of the project, the following settings were used in Raman testing: • All spectra were collected using a 1,064-nm excitation source featuring a 500 mW IR laser. • A default range was set in the IR region between 4,500 and 100 cm–1 with a spectral resolution of 3 cm–1. • Raman spectra were collected at room temperature using the laboratory bench probe (Figure A.3). • RTA’s RamanVista software was used for the analysis of spectra. Test Sample Preparation No sample preparation or conditioning was required for most of the materials. Samples were divided into small ali- quots of a reasonable size for each material (i.e., drops for liquids and a small spatula for powders). Each aliquot was then placed on a microscope slide for Raman analysis. In some instances, an additional microscope slide was placed on top of the sample to minimize the evaporation of volatiles and pro- vide a uniform upper surface for analysis by the downward- facing Raman probe (Figure A.3). The upper surface of glass also helped to conduct heat away from the sample, which was helpful when thermal emission from the laser heating became an issue. For asphalt binders, asphalt emulsions, and gels (e.g., Ultrabond 1100 Part B, Scotchkote Part B), solvent extrac- tions (with hexane and methylene chloride) were performed to avoid the detrimental effects of heating or fluorescence excitation. x-Ray Fluorescence Spectroscopy A review of the applications of XRF spectroscopy to highway materials revealed that portable XRF devices have been avail- able since the early 1990s for the analysis of the heavy metal concentration in certain substances (e.g., paints and soils). Therefore, the portable XRF device was included in the labo- ratory testing of metal-containing materials. The equipment setup and testing procedures are described in detail. Innov-X Alpha XRF Analyzer Equipment Setup The portable Innov-X Alpha XRF analyzer (Figure A.4) owned by the team measures the concentration of the following Figure A.2. Image of Nicolet Magna 560 FTIR spectrometer. Figure A.3. Image of RTA’s portable Raman analyzer.

46 elements: P, S, Cl, K, Ca, Ti, Cr, Mn, Fe, Co, Ni, Cu, Zn, As, Se, Sr, Zr, Mo, Ag, Cd, Sn, I, Ba, W, Hg, and Pb. There are two separate calibration modes for this particular XRF: one mode for trace elements in soils (concentrations up to 8% to 10% by weight) and one mode for alloys and ores that have concentrations >1% by weight. However, the alloy mode can only measure elements heavier than Ti, so Ca, K, and other lighter elements can only be measured in relatively low concentrations using this equipment. Other portable XRF devices are available that can measure light elements in higher concentrations as well as capture Mg, Si, and Al, but the cost of these devices is higher (around $85,000). Test Sample Preparation The general sample preparation method for the XRF was as follows: • Liquid samples, emulsions, and portland cement were tested in their as-received state by placing a 15 to 30 g of the sample in an XRF sample holder. • Aggregates and solids were crushed and pulverized to pass through a U.S. No. 60 screen, in accordance with U.S. Envi- ronmental Protection Agency Method 6200. x-Ray diffraction Spectroscopy In Phase 1, numerous studies were found to present XRD analysis as a good quantitative method for analyzing portland cement and its supplements. In addition, XRD was men- tioned as a suitable technique for identifying the mineral components of aggregates. Therefore, XRD was included in the scope of laboratory testing in Phase 2 with the objective of evaluating the applicability of portable XRD devices. XRD was used to analyze PCC-related products and aggregates and to compare the performance of portable and stationary XRD systems. The following is a description of the XRD equip- ment and testing protocols used by the team in Phase 2. Stationary Bruker-AXS (Siemens) D5005 Diffractometer Equipment Setup For stationary XRD analysis, the team used the Bruker-AXS (Siemens) D5005 diffractometer (Figure A.5) located in the facilities of IMS at UCONN. The diffractometer was equipped with a Cu source (l = 1.5418 Å) and the X-ray tube was oper- ated at 40 kV and 40 mA using a diffracted beam graphite- monochromator. The data were collected for scattering angles (2q) ranging from 5° to 65° with a step size of 0.02° and vari- able counting times. The counting time affects the signal-to- noise ratio and the overall quality of the data. High counting times are desirable to lower the XRD detection limit and obtain good quantitative data, but the scanning time may become prohibitive for field applications. The required scan- ning time will also vary with the instrument type and the age of the X-ray tube. The scanning times varied between 2 s per step and 14 s per step. Samples were marked as LR (low reso- lution) when scanning times were less than 5 s per step and HR (high resolution) when the scanning time was more than 5 s per step. This enabled a comparison between the data quality of LR and HR samples. LR corresponds to approxi- Figure A.4. Image of Innov-X Alpha XRF analyzer. Figure A.5. Image of Bruker-AXS (Siemens) D5005 diffractometer.

47 mately 2 to 4 h of scanning time, while HR required 8 to 12 h of scanning time. Qualitative and quantitative analysis of the XRD data was performed using Jade software, Version 8.5 (MDI, 2008), with reference to the patterns of the International Centre for Diffraction Data database (PDF-2, 2002) and the American Mineralogist Crystal Structure database (http://rruff.geo The Rietveld method, using the whole pattern fitting function of Jade, yielded quantita- tive phase analysis (relative amounts of each substance). Test Sample Preparation For stationary XRD analyses, a subsample of 10 to 20 g was obtained from the material containers, placed in a Petri dish, and air-dried for 24 h as needed. The sample was pulverized to pass a U.S. 40 (0.425 mm) sieve. The pulverized sample was then manually homogenized and 1 g of the whole sample was pulverized to pass the U.S. 400 (38 µm) sieve. An aliquot of 0.8 g of the finely pulverized sample was mixed with 0.2 g of corundum (a-Al2O3), which was used as the internal stan- dard for quantitative analysis. The corundum obtained from Sawyer was previously found to be 93% crystalline by com- paring it with a National Institute of Standards and Technol- ogy ZnO standard (1). The Rietveld method provides a relative quantification of the identified crystalline com- pounds in an XRD sample. If a compound is either low in content, or disordered or amorphous, it will not be detected and thus the compound will not be quantified. In that case, the sum of the phases that cannot be detected in the XRD pattern are termed “amorphous” or “sample amorphicity.” This leads to an overestimation of the observed crystalline compounds. By spiking the sample with a known amount of a crystalline material, it is possible to backcalculate the amor- phous content, A, using Equation A.1: A I I I I AR R = − −( )1 1( . ) where IR is the Rietveld-estimated quantity and I is the actual amount of the spiked mineral. A 20% corundum was used as the known spike. Portable Terra XRD/XRF Analyzer Equipment Setup The portable XRD and XRF Terra analyzer is approximately the size of a briefcase (about 49 × 40 × 20 cm in dimensions and 14.5 kg by weight). The company inXitu developed this device for the identification of minerals and aggregate mix- tures. According to the manufacturer, the system requires minimal sample preparation and detects single minerals or simple mixtures within minutes. The Terra analyzer provides an XRD resolution of 0.25° over the 2q range of 5° to 55° and XRF resolution of 230 eV (at 5.9 keV) over the range of 2 to 25 keV. The device can operate in an autonomous mode for at least 4 h. A laptop computer allows the user to configure the analysis, preview live data, explore archive files, and download data for pattern matching with a mineral database using commercial software. Test Sample Preparation For the analysis of the portable XRD, the same samples were used as for the stationary XRD. The samples were shipped to inXitu headquarters in Campbell, California. Gel permeation Chromatography Although GPC (with UV detectors and ELSD) was not iden- tified as a portable spectroscopic method, it was found to be a successful analytical procedure for a range of materials, especially for asphalt-related products. Assuming that the benchtop device can potentially be fitted into a mobile labora- tory, the team decided to include GPC in Phase 2 testing and evaluate its applicability to fingerprinting the construction materials within the scope of this project. The Waters 410 GPC system located in IMS at UCONN was used for the evaluation, as discussed below. Waters GPC System Equipment Setup Waters GPC has a horizontally placed column (Figure A.6) with the stationary phase (gel) placed in a cylindrical tube. The sample (diluted in solvent) is injected into the column, and a UV or light-scattering detector measures the change in the absorption of light as the fractionated material exits the column. The separated components of the material can then be further analyzed by coupling the chromatographic column with a detector that identifies each compound separately. The system is able to resolve over a molecular weight range of 100 to 250,000 Da, with a precision of 1% to 3%. The following GPC procedural settings were used in laboratory testing: • THF was used as the eluent with a flow rate of 1 mL/min at 35°C. • Typical test lasted for 35 min. • Both a PL-ELS100 (evaporative light-scattering) detector and a Waters 2487 dual wavelength absorbance (UV) detector were used. • Signals were recorded and analyzed with Millennium software.

48 • Polystyrene standards with molecular weights of 2,000,000, 900,000, 824,000, 400,000, 200,000, 110,000, 43,000, 30,000, 17,600, 6,930, 2,610, 982, and 450 were used to establish the calibration curve. • Peak molecular weights were identified and labeled on the spectra. Test Sample Preparation Three different types of samples were prepared for the GPC experiments as follows: 1. Structural coatings, traffic paints, and epoxy adhesives were dissolved in 5 mL of THF solvent as a 1 wt% solution. The solution was then transferred to a syringe and forced through a 0.45-µm filter. The filtered solution was then transferred into a GPC vial. 2. The other materials, with the exception of recycled asphalt pavement (RAP)-containing binders, were dissolved in 5 mL of THF as a 1 wt% solution and then transferred into a GPC vial without filtering. 3. The RAP-containing binders were first extracted by plac- ing 5 g of mix in a 20-mL vial with 10 mL of methylene chloride, stirring for 5 min, filtering the solution, and vacuum drying the filtrate into a clean vial. The resulting binder extract was then dissolved in 10 mL of THF as the GPC solvent and then transferred into a GPC vial. nuclear Magnetic Resonance The literature search conducted in Phase 1 of this project revealed that, since the 1980s, laboratory NMR systems have been mostly used by university-based researchers for the anal- ysis of petroleum products. However, knowing that NMR spectroscopy is extremely helpful in identifying the chemical structure of virtually any organic compound, the team decided to include this method in the laboratory testing program. Despite the team’s having no access to portable NMR equip- ment by the time this report was finalized, the results obtained from the NMR testing in laboratory conditions may be useful in the future. Bruker DRX-400 NMR Spectrometer Equipment Setup Figure A.7 shows the setup of the Bruker DRX-400 NMR spectrometer located in the IMS facilities at UCONN. The device uses time-domain analysis for solutions and frequency- domain analysis for solids in a 1H, 13C, 31P, or 19F environment. In Phase 2, 1H proton NMR spectra were recorded and ana- lyzed as follows: • Sixteen scans were performed for each run. • D-chloroform was used as the lock solvent. • TopSpin software was used to collect the signal. • MestReNova software was used to analyze the signal and prepare the report. • The peaks were identified from the known constituents and a table of NMR chemical shifts. Figure A.6. Image of Waters GPC system. Figure A.7. Image of Bruker DRX-400 NMR system.

49 Test Sample Preparation Because the NMR requires material to be in the liquid state, the following preparation methods were used: 1. Structural coatings, traffic paints, and epoxy adhesives were dissolved in 2 mL of CDCl3 (D-chloroform) sol- vent as a 2 wt% solution. The solution was then trans- ferred to a syringe and forced through a 0.45-µm filter. The filtered solution was loaded into the NMR tube. 2. The rest of the materials, with the exception of RAP- containing binders and hot-mix asphalt mixes, were dis- solved in 2 mL of CDCl3 solvent as a 1 wt% solution and transferred directly into the NMR tube. 3. The RAP-containing binders were first extracted by placing 5 g of mix in a 20-mL vial with 10 mL of methy- lene chloride, stirring the suspension for 5 min, filter- ing the solution, and vacuum drying the filtrate in a fresh vial. The resulting binder extract was then dis- solved in 1 mL of D-chloroform and diluted to 2 wt% solution. Summary of equipment This section summarizes the results of the evaluation of the portable spectroscopic devices in Phase 2. Table A.3 lists the spectroscopic devices chosen for the “proof of concept” in the laboratory phase of the project. The evaluation concerned the ability of portable FTIR, Raman, XRF, and XRD devices to comply with qualitative and quantitative requirements of field quality assurance/quality control procedures, such as accuracy, time and labor involved in testing, level of training required, and other parameters. These requirements were established in Phase 1 based on the feedback of the profes- sionals who participated in the workshop organized by the team and held at UCONN. Table A.4 compares the actual parameters achieved in the laboratory conditions with the target values. As shown in Table A.4, all devices evaluated in Phase 2 com- ply with the previously established criteria, suggesting that the team chose the equipment correctly. However, final recom- mendations for the further evaluation of these devices in Phase 3 (field experiments) were based on the success of the portable equipment to produce interpretable results for the materials included in experimental design of Phase 2. These recommendations are summarized in Chapter 4. Summary of Survey of Approved and Qualified product Lists Figures A.8 to A.17 summarize the results of the survey of approved and qualified product lists. Reference 1. Dermatas, D., M. Chrysochoou, S. Sarra Pardali, and D. G. Grubb. Influence of X-Ray Diffraction Sample Preparation on Quantitative Mineralogy: Implications for Chromate Waste Treatment. Journal of Environmental Quality, Vol. 36, 2007, pp. 487–497. Table A.3. Finalized List of Spectroscopic Devices for “Proof of Concept” in the Laboratory Spectroscopic Method Brand/Manufacturer Model Portability FTIR Nicolet Magna 560 Stationary Bruker ALPHA II ATR Portable (benchtop) RTA, Inc. RamanID Portable (trunk size) XRD Bruker-AXS (Siemens) D5005 Stationary inXitu, Inc. Terra Portable (trunk size) XRF Innov-X Systems, Inc. Alpha Portable (handheld) NMR Bruker DRX-400 Stationary GPC Viscotec Waters Stationary

50 Table A.4. Summary of Portable Equipment Evaluation in Phase 2 Featurea Value FTIR Raman XRF XRD Accuracy Minimum 1% <0.5% <2% <1% <1% Goal <0.5% Duration of measurement Maximum 1 h ~1 min ~1 min 6–12 min 15 min Goal ~5 min Effort involved Maximum 1 person 1 person 1 person 1 person 1 person Goal 1 person Amount of prior training Maximum 1 day 1 h 1 h 1 h 1 h Goal 0.5 day Reliability Minimum Depends on material (90%) 99% (software failure) Depends on materialc 99% (software failure) 99% (software failure) Goal 95% Time to get results Maximum Depends on construction process (1 h) ~5 min ~5 min ~5 min ~5 min Goal ~5 min Price range Maximum $50,000 ~$25,000 ~$60,000 ~$37,000 $45,000 Goal <$20,000 Device weight Maximum 50 lb ~16 lb ~20 lb ~4 lb (handheld) ~27 lb Goal <20 lb ~15 (benchtop) Sample preparation Maximum Solvent As isb As isb As is (liquids) pulverization (solids) Crushing (solids) Goal As is a Accuracy: Agreement between a measurement and the true or correct value. Duration of measurement: Time between start and end of testing cycle. Effort involved: Personnel required to perform the test. Amount of prior training: Time required to make personnel familiar with a testing procedure. Reliability: Unlikelihood of equipment failure during the test. Time to get results: Time between beginning of the sample preparation and the end of the analysis of the test results. Price range: Cost of the equipment. Device weight: Mass of the equipment including the case or enclosure. Best time for QA/QC: Stage of the manufacturing or application process when the test is most timely. Sample preparation: Processing/manipulating the material before the test. Minimum/maximum: Acceptable threshold value from the user perspective that given equipment should produce. Goal: Desirable target value from the user perspective that given equipment should produce. b Pulverization of granular materials (aggregates) is required for better quality. c Failure to get the signal for fluorescent materials or because of thermal emission.

51 Figure A.8. Distribution of pavement marking brands approved by state highway agencies (SHAs). Figure A.9. Distribution of structural coating brands approved by SHAs.

52 Figure A.10. Distribution of epoxy resin brands approved by SHAs. Figure A.11. Distribution of air-entraining admixture brands approved by SHAs.

53 Figure A.12. Distribution of PCC set retarder and accelerator brands approved by SHAs. Figure A.13. Distribution of water reducer and plasticizer brands approved by SHAs.

54 Figure A.14. Distribution of PCC curing compound brands approved by SHAs. Figure A.15. Distribution of asphalt antistripping brands approved by SHAs.

55 Figure A.16. Distribution of polymer-modified emulsion vendors approved by SHAs. Figure A.17. Distribution of polymer-modified binder suppliers approved by SHAs.

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