Analytical glycobiology at high sensitivity: current
approaches and directions
Milos V. Novotny & William R. Alley Jr. &
Benjamin F. Mann
Received: 3 March 2012 /Revised: 29 June 2012 /Accepted: 14 August 2012 /Published online: 4 September 2012
# Springer Science+Business Media, LLC 2012
Abstract This review summarizes the analytical advances
made during the last several years in the structural and quantitative
determinations of glycoproteins in complex biological
mixtures. The main analytical techniques used in the fields of
glycomics and glycoproteomics involve different modes of
mass spectrometry and their combinations with capillary separation
methods such as microcolumn liquid chromatography
and capillary electrophoresis. The need for high-sensitivity
measurements have been emphasized in the oligosaccharide
profiling used in the field of biomarker discovery through
MALDI mass spectrometry. High-sensitivity profiling of both
glycans and glycopeptides from biological fluids and tissue
extracts has been aided significantly through lectin preconcentration
and the uses of affinity chromatography.
Keywords Biomolecular mass spectrometry .
Glycoproteomics . Glycomics, affinity chromatography .
Lectin enrichment . Permethylation, sialic acid linkage
analysis . Capillary electrophoresis
Introduction
Biological sciences have recently undergone enormous expansion,
generating new fields with new names. All rapidly
developing fields in life sciences feature a common trend:
from empirical and descriptive biology toward the understanding
of molecular and dynamic aspects. Since the early
1990s on, new measurement technologies further lend exciting
opportunities for quantifying the molecular aspects of
processes in living organisms and combining these extensive
molecular data into the “systems biology” knowledge
[1–5]. Among the different “-omics fields” (genomics, transcriptomics,
proteomics, metabolomics, etc.), which are all
closely related to new developments in analytical methodologies
and instrumentation, the fields of glycoproteomics
and glycomics are now finally assuming a very important
role. Both fields contribute substantially to a better understanding
of multicellular interactions in eukaryotic systems
and the many issues pertaining to human health and disease
[6–12]. The long-held views that glycosylation in prokaryotic
systems is unimportant have been seriously challenged
during the last several years [13–19]. Since glycans seem to
provide the first interfacial layer between the mammalian
host and parasites, glycosylation plays a crucial role in
pathogenicity and invasion as well. As some microorganisms
appear to express distinctly different sugar units and
oligosaccharide structures, the area of host-pathogen interactions
has already started to create its own bioanalytical
challenges.
It is now evident that the sheer complexity of carbohydrate
structures and their complicated (non-template) biosynthetic
origin discouraged previous generations of scientists to work
in glycoscience. From a molecular scientist’s point of view,
this field encompasses a vast range of directions due to the
structural complexities of glycoconjugates, of which the glycoproteins
represent just one class of biologically important
molecules. This brief, authoritative review emphasizes glycoprotein
analysis at high sensitivity and shares common
methodological concerns and directions with the structural
studies of other large glycomolecules encountered in nature
M. V. Novotny (*) :W. R. Alley Jr. :B. F. Mann
Department of Chemistry, Indiana University,
Bloomington, IN, USA
e-mail: novotny@indiana.edu
M. V. Novotny :W. R. Alley Jr. :B. F. Mann
National Center for Glycomics and Glycoproteomics,
Indiana University,
Bloomington, IN, USA
M. V. Novotny
Indiana University School of Medicine, Indiana University,
Indianapolis, IN, USA
Glycoconj J (2013) 30:89–117
DOI 10.1007/s10719-012-9444-8
(such as proteoglycans or polysaccharides), which undoubtedly
other investigators will address and review. The focus of
our brief review has primarily been to describe the methodological
advances of interest to modern biomedical research.
The reviewed methodologies emphasize the use of mass spectrometry
and capillary separation techniques.
General considerations
The emergence of “glycobiology” during the late 1980s
serendipitously coincided with the development of new
ionization techniques in biomolecular mass spectrometry
(MS) and the recognition of capillary electrophoresis (CE)
and capillary liquid chromatography (LC) as new important
techniques in biochemical analysis. The importance of glycosylation
has since been widely recognized functionally
and structurally. The current estimate of some 50 to 70 %
of all mammalian proteins being glycosylated represents a
formidable analytical challenge that far exceeds the tasks of
the mainstream proteomics. It is becoming increasingly
evident that the structural selectivity determinants of biological
recognition are either unique glycan structures or combinations
of different glycans at the sites of glycosylation
and, to some degree, the local “peptide landscape” at the site
of a glycosylated protein. The well-known propensity of
sugars to form numerous glycan isomers adds substantially
to the difficulties of the overall analytical glycobiology
tasks. It is thus desirable that glycomic and glycoproteomic
measurements be practiced with a comparable level of expertise
and probably best if performed in the same laboratory.
Different techniques and instrumentation may need to
be involved in completion of the overall structural task.
Still during the early 1990s, milligram quantities of glycoproteins
were typically required for structural analysis;
the use of ion-exchange LC with pulsed amperometric detection
was widely considered as the “gold standard” for
glycomic profiling. Over the two following decades, impressive
gains in MS detection technologies and instrumentation
were gradually achieved for the benefits of proteomic
sequencing and mass screening. Simultaneously, the development
of fluorescence-labeling for either LC [20, 21] or
CE with laser-induced fluorescence (LIF) detection [22–26]
has promoted the importance of sample preparation prior to
the final glycomic or glycoproteomic measurements. There
has been evolution of the separation columns used for the
benefits of MS. Various columns with ever smaller diameters,
whether in their packed, monolithic, superficiallyporous,
or open tubular and microfabricated formats, have
gradually become a part of many on-going efforts to downsize
our analytical tools. The general benefits of miniaturized
columns have been achieved with increased mass sensitivity,
in minimum sample dilution prior to measurements,
and a reduced surface exposure of the analyzed glycoproteins.
These miniaturization efforts have also resulted in a
better integration of different analytical steps, reduced sample
scales and better automation possibilities in both glycomics
and glycoproteomics. Certain advantages of these
operations will be featured below in some detail. Overall,
in different incremental steps, significant sensitivity gains
have been achieved during the last decade, amounting to
several orders of magnitude.
The field of analytical glycobiology is somewhat arbitrarily
divided into “glycomics”, emphasizing the functional
centrality of the glycomes in life processes and thus measuring
oligosaccharide profiles as its primary task, and “glycoproteomics”,
dealing with the overall glycoprotein,
structure at all structural levels. When dealing with unfractionated
biological materials, the vast complexity and abundance
of the structural and quantitative data in both
glycomics and glycoproteomics invites the advanced uses
of information technology in automated identification of
proteins and glycans, mass screening, and library comparisons.
The field of bioinformatics has evolved rapidly in
these areas and united a number of laboratories at the international
level [27–32].
While different analytical technologies continue to
evolve at their own pace, the most valuable incentive for
their development is provided by some of the most topical
areas of scientific endeavor such as the search for disease
biomarkers (for the benefits of both biomedicine and pharmaceutical
development) and investigations into the molecular
basis of host-pathogen interactions.
Recent analytical directions and strategies
There has been a dramatic increase in the laboratories working
on the subject of glycoproteins, as evidenced by the
large and growing number of published studies in both
biomedical and analytical journals. Although such studies
employ a variety of methods and approaches, developments
in glycomics and glycoproteomics lag behind those in
genomics and “mainstream proteomics.” This situation
arises from the complexity of mammalian glycomes that
are estimated to contain many thousands of oligosaccharides
generated through a non-template biosynthetic process.
Glycoproteomics
It is now commonly perceived that the analysis of major,
isolated glycoproteins or recombinant products (confirmatory
analysis) has become a nearly routine task. The current
methodological attention has been more directed toward the
analysis of complex mixtures and biological samples derived
from body fluids, tissues, cell cultures, etc. Due to the
90 Glycoconj J (2013) 30:89–117
large differences in the occurrence and concentration of
glycoproteins in such biological specimens, methodological
difficulties are often encountered. There is a need to develop
more generic platforms to account for glycoproteins that are
variously distributed in different cells and subcellular entities,
as the cellular compartmentalization is certainly important
for the biosynthesis, structural modification and the
function of glycans. While numerous studies have been
conducted toward optimization of the extraction schemes
to cover both the membrane-bound and soluble proteins
from tissue materials, many procedures remain somewhat
application-specific. In contrast, the sample treatment procedures
for the common physiological fluids, such as human
blood serum or plasma, tend to be more unified at present.
However, the enormous concentration range in which the
glycoproteins of interest may be encountered in such fluids
still presents a formidable challenge, inviting sample preconcentration
schemes based on affinity chromatography.
A highly effective MS analysis of glycopeptides after
enzymatic degradation is at the heart of all glycoproteomic
strategies, whether they use initially the “top-down” approach,
separating proteins first, or the “bottom-up” approach,
digesting the protein mixtures directly. Sample
treatment, separation procedures and data processing generally
parallel those of the advanced (high-sensitivity) general
proteomics, reviewed amply elsewhere [33–35]. The added
difficulties include: (a) separation of glycoproteins from
non-glycosylated proteins prior to digestion, and (b) difficulties
of adequately measuring, or even detecting, the
protease-digested glycopeptides in the presence of other
peptides through the tandem MS (MS/MS) methodologies.
Glycopeptides easily suffer from suppressed ionization in
the presence of other components.
Multimethodological strategies for glycoproteomic profiling
At the level of analyzing complex protein mixtures, the use
of at least two-dimensional separation technologies appears
mandatory, albeit at some expense in both reproducibility
and time of analysis. As is evident from the number of
recent publications, methods and protocols, and reviews,
various combinations of LC, CE and MS techniques are
applicable in the search for the best protein and peptide
mapping strategies [36–38]. Additionally, traditional 2-D
gel electrophoresis and other modified gel-based methods
continue to be utilized and further developed [19, 39–41].
The reversed-phase capillary LC has become a wellestablished
procedure in peptide mapping when using either
UV absorbance detection or LC-MS with low-microgram
protein digests for simple peptide mixtures or digests with a
moderate degree of complexity. However, peptide mixtures
of greater complexity necessitate columns with high efficiencies,
optimized separation conditions, and alternatively,
the use of multidimensional chromatographic techniques
[42, 43]. Generally, microgram quantities of the protein
mixtures can now be routinely handled and sequenced
through the use of capillary LC-MS/MS-based procedures,
while the use of even smaller-diameter columns, such as 10μm
i.d. porous layer open tubular (PLOT) capillaries, can
further enhance mass sensitivities [44]. However, sometimes
even under the best analytical conditions, it may be
difficult to measure many glycoproteins in enzymatic
digests due to the mentioned ionization suppression phenomena.
Perhaps a better solution to the problem resides
with the use of enrichment methodologies such as the use of
lectins (the application of which is discussed in greater
detail in the next section) or immunoactive media prior to
LC-MS/MS runs. These enrichment steps largely avoid
competitive ionization through a targeted mixture simplification.
The fractionation of complex protein mixtures can be
accomplished by means of bioaffinity and the PTM (posttranslational
modification) type, or to a lesser degree, other
chromatographic and electrophoretic principles. However, it
is desirable in so far as it does not compromise the reproducibility
and quantification in comparative analyses.
The possibility of lectin immobilization on solid surfaces
(particles, membranes, separatory channels, etc.) allows the
utilization of these unique proteins in various glycoproteomic
schemes, in which the trace glycoproteins can be effectively
enriched and further processed through a common
proteomic platform based on LC-MS/MS. One such analytical
platform, which was developed [45] and further refined
for label-free quantitative glycoproteomic profiling [46–48]
of blood serum samples in our laboratory is shown in Fig. 1.
In this application, the experimental workflow fractionates
the serum sample (usually 20–30 μL) through interaction
with two types of bioaffinity media: first, the immunoaffinity
capture of major serum proteins depletes highly abundant
proteins from the sample to facilitate further proteomic
investigations of trace glycoproteins; second, a lectin microcolumn
is employed to further preconcentrate the glycoproteins
of interest. Using an appropriate competitive hapten
buffer, the glycoprotein fraction is desorbed for an additional
fractionation through reversed-phase LC into about 30
fractions, which are, in turn, trypsinized and subjected to the
usual LC-MS/MS tryptic mapping procedure, database
search and evaluation. Following a statistical treatment of
the generated data, such glycoproteomic surveys can identify
groups or individual glycoproteins that are capable of
discriminating between physiologically distinct states of
health [48–52]. Furthermore, glycoproteins that are identified
as interesting biomarker candidates can be targeted for
specific characterization in subsequent studies [53–56].
Immunoaffinity-based chromatographic purification may
offer a clear path to targeted MS profiling of glycans and
glycopeptides from putative biomarkers in body fluids and
Glycoconj J (2013) 30:89–117 91
tissues. This approach is currently somewhat limited by the
availability of highly specific antibodies at a reasonable price
and the lack of optimal solid supports for construction of
miniaturized preconcentrators. As the range of available antibodies
gradually expands, protein-targeted measurements
may in time become more common. As a preliminary example
of this approach, we demonstrate here the immunoisolation of
α-1-acid glycoprotein (Fig. 2) from a small volume of human
blood serum, followed by deglycosylation and a glycomic
profile measurement (unpublished results). Another example
in the recent literature includes isolation of haptoglobin [57], a
serum protein with an increased abundance of fucosylation
that has been implicated in pancreatic cancer [57–59].
Lectin affinity chromatography
Biospecific interactions of lectins with certain glycoproteins
have long been documented in the histochemical, cytochemical,
and biochemical literature, but much less applied in quantitative
analytical procedures until recently. The choice of a particular
lectin in a preconcentrator medium can be critical to the success
of the entire glycoproteomic profiling. It has long been known
that some lectins exhibit a degree of selectivity toward certain
oligosaccharide structures (differently linked sialic acids, fucosylated
glycans, mannose-rich structures, etc.) in contrast to the
more “generic” concanavalin A (Con A). In a biomedically
important application to screening glycoproteins in microliter
volumes of blood sera, it has become feasible to observe, and
potentially quantify, several hundred constituents in lectinenriched
fractions [45, 49, 50, 60–62]. The number of identified/measured
sample components can further be enhanced
through the use of new high-resolution mass spectrometers such
as LTQ-Fourier transform ion-cyclotron resonance (FT-ICR),
Orbitrap, or Q-TOF instruments. In our studies of serum minor
proteins [45], four common lectins were evaluated in terms of
molecular size and pI ranges of the retained components. The
Proteolytic digests
Sample
Immunoaffinity
depletion
On-line lectin affinity
High-temp RPLC
Glycoprotein fractions
Nano LC-MS/MS Database search Glycoprotein Data
Lectin
microcolumn
Isocratic pump w/
injector
C8Poroshell
column
Capillary
pump
Waste
C4 trapping
column
Fig. 1 Multimethodological glycoproteomic workflow. A complex
biological sample is first depleted of highly abundant proteins, followed
by lectin preconcentration of the glycoproteins. Next, the
enriched glycoproteins are further fractionated by reversed-phase
liquid chromatography (RPLC). Fractions are enzymatically digested,
followed by bottom-up LC-MS/MS for protein identification/quantification.
(From Reference [45])
92 Glycoconj J (2013) 30:89–117
results indicated that such lectins have both overlapping and
selective properties. In order to generate “representative” profiles
of a particular sample type, our laboratory [45] and others
use different lectins in either a sequential-lectin arrangement or
a multi-lectin (mixed-bed) arrangement [63], with some differences
in performance of both techniques noted [60]. The use of
silica-based lectin microcolumns has been beneficial, as it permits
incorporation into a valve-based high-pressure analytical
system [61].
Following the development of microscale lectin affinity
techniques for enrichment of glycoproteins in biological
materials, this approach has been the basis for a multitude
of glycoproteomic investigations that aim to characterize the
sub-glycoproteomes of a variety of biological materials
derived from humans, including urine [64, 65], saliva [66],
organ tissues [67], and, most frequently, blood serum [45,
48, 67–71].
A less usual example of targeted glycoproteomic analysis
through lectin enrichment can be found in our recent investigation
of pancreatic cyst fluids [72]. The samples, which
were collected by a fine needle aspiration of the cystic
lesions intraoperatively to avoid peripheral contamination,
were highly variable, with inconsistent coloring and viscosity,
in addition to variable protein composition and total
content. After a combination of filtration and bufferexchanging
steps were applied, relatively clear fluids were
obtained for glycomic and glycoproteomic profiling. MSbased
glycomic analysis showed many glycans that were
observed in serum profiles, but, in a few of the fluids that
were associated with a higher risk of malignant transformation,
we have identified a number of hyperfucosylated glycans
(unusual structures) containing from 2–6 fucose
residues on a single structure (Fig. 3). Such glycans were
not seen in the whole serum profiles. This study represents a
less usual approach, i.e. glycomic profiling first, as a guide
for subsequent glycoproteomic studies. Alternatively, most
investigators conduct proteomic studies first, targeting glycosylation
later.
Following an untargeted proteomic analysis to provide a
baseline information, a glycoproteomic profiling workflow
was modified to include Aleuria aurantia lectin (AAL) for
the identification of the glycoproteins that were hyperfucosylated.
A label-free comparison of the non-enriched and AALenriched
proteomic profiles, facilitated by ProteinQuant [47],
identified several glycoproteins that were overexpressed. This
included pancreatic α-amylase, triacylglycerol lipase, and
elastase-3A, as the proteins in high abundance following
AAL enrichment (Fig. 3c). This study illustrates the advantages
of performing glycomic and glycoproteomic investigations
in the same laboratory.
With a better understanding of how the lectin preconcentrators
work as critical components of the overall analytical
2602.8 3206.6 3810.4 4414.2 5018.0
Mass (m/z)
1515.7
0
10
20
30
40
50
60
70
80
90
100
%Intensity
2792.4
3602.8
3777.0
4413.2
4588.0
4052.0
Fig. 2 MALDI-TOF-MS of permethylated N-glycans enzymatically
released from α-1-acid glycoprotein (AGP) that was purified from a 5μL
aliquot of human blood serum utilizing a murine monoclonal
antibody (mAb). The serum had been previously depleted of seven
highly abundant proteins. The mAb was incubated for 1 h with the
depleted serum, and the mAb-AGP complex was then extracted from
the mixture with anti-mouse IgG coupled to agarose beads. The beads
were thoroughly washed to remove the unbound serum proteins, and
the purified mAb-AGP was subsequently eluted from the anti-mouse
beads with acetic acid, pH 2.6, prior to enzymatic release of N-glycans
with PNGase F. We depict here different multiantennary glycans as
cartoon structures, using the symbols established by the Consortium
for Function Glycomics (http://www.functionalglycomics.org/static/
consortium/Nomenclature.shtml)
Glycoconj J (2013) 30:89–117 93
1500 1850 2200 2550 2900 3250
0
10
20
30
40
50
60
70
80
90
100
2081.0
2837.3
1706.8
1906.9
2039.9
1579.7
2663.2
1835.9
2459.1
2244.0
2111.0
2681.2
1590.7
2489.1
2418.1
1987.9
1519.7
2388.1
2592.2
2192.0
1865.9
3042.4
2909.3
2792.3
1661.8
3082.5
2622.2
3112.5
1764.8
1783.8
2285.1
a
3250 3600 3950 4300 4650 5000
0
10
20
30
40
50
60
70
80
90
100
4709.2
3460.6
4535.1
3910.9
3736.8
4083.9
4360.0
3286.6
3563.6
3532.6
b
c
Proteins Overexpressed in Multiply-fucosylated Pancreatic Cyst Fluids
Comparative Proteomics
No enrichment AAL-enriched
Accession Protein Name (Alternate Name) (G1/G2)* P-value (G1/G2)* P-value
P00995 Pancreatic secretory trypsin inhibitor (Tumor-associated trypsin inhibitor) 8.4 0.0040 50.1 0.0101
P04746 Pancreatic alpha-amylase 1.7 0.0476 22.4 0.0017
P16233 Pancreatic triacylglycerol lipase 2.9 0.0001 20.2 0.0009
P07478 Trypsin-2 6.3 0.0010 15.6 0.0048
P09093 Chymotrypsin-like elastase family member 3A (Elastase-3A) 2.5 0.0157 11.2 0.0158
P19835 Bile salt-activated lipase 3.5 0.0001 9.8 0.0067
P05451 Lithostathine-1-alpha (Pancreatic stone protein) 5.9 0.0012 5.7 0.0030
P08217 Chymotrypsin-like elastase family member 2A (Elastase-2A) 3.0 0.0022 4.8 0.0010
Q99895 Chymotrypsin-C (Caldecrin) 4.0 0.0013 2.7 0.0017
Fig. 3 Glycomic and glycoproteomic analysis of so-called “hyperfucosylated”
glycoproteins in pancreatic cyst fluids. MALDI-TOFMS
of permethylated N-glycans a from m/z 1500–3250 and b from
m/z 3250–5000. c A label-free quantitative comparison of the glycoproteins
overexpressed in the hyperfucosylated fluids (G1) to the nonhyperfucosylated
fluids (G2), based on whole-fluid proteomics (“No
enrichment”) and lectin-enriched glycoproteomics with Aleuria aurantia
lectin (AAL) (“AAL-enriched”), revealed the glycoproteins significantly
overexpressed and, most importantly, those that were both
overexpressed and highly enriched by AAL. (From Reference [72])
94 Glycoconj J (2013) 30:89–117
schemes, further advances in glycoproteomic profiling can
hopefully be realized. For comparative studies, as needed in
virtually all topical applications of medical glycobiology, it
is essential to secure adequate quantitative reliability in
every step of a glycoproteomic workflow. It is thus desirable
to utilize small-scale formats for the lectin enrichment step
to ensure a quantitative recovery of the enriched sample
components. Due to the relatively weak interactions between
most lectins and their target carbohydrate moieties
(approximate Kd range: 10−4
–10−7
M), the best enrichment
support materials provide a very high accessible surface
area, while also exhibiting a fast rate of mass transfer.
Furthermore, a high lectin density can greatly improve the
realized strength of the interaction with target glycoproteins
through simultaneous interactions with multiple sites of
glycosylation (multivalency) [73]. As such, monolithic columns
are expected to be suitable for this type of work, but
the current rapid development of various new materials may
lead to the discovery of supports that offer their own unique
advantages. As an example of these efforts, a novel particulate
silica material (1.6 μm diameter) containing an extensive,
sponge-like network of macropores has been utilized in
our lab to reproducibly enrich important glycoproteins from
a single microliter of whole blood serum or an equivalent
amount of albumin- and IgG-depleted serum using Con A
and AAL [74]
It is critical to ensure that the lectin preconcentration step
does not become a bottleneck in the overall quantification
procedure. While our recent data [46] with one lectin indicate
that adequate analytical reproducibility could be
achieved in label-free quantitative proteomics, rigorous
standardization steps must be followed for all lectin-based
procedures.
Profiling human immunoglobulins at high sensitivity
While many useful and now widely appreciated MS and
computational methodologies in the field of proteomics
were initially driven by the “global” or “total” approach to
the complexity of protein and peptide mixtures, today’s
investigators increasingly appreciate the value of more selective
and targeted approaches to the complexity problem.
One family of glycoproteins that is inherent to our bodily
defense system and thus very interesting to characterize (in
terms of glycosylation) are the immunoglobulins (Igs). As it
has been shown already, the function of IgG can be reversed
from pro-inflammatory to anti-inflammatory by addition of
terminal N-acetylneuraminic acids [75]. Moreover, profiling
of IgG glycans in a large-scale study has recently demonstrated
that decreased galactosylation correlates directly to
increasing age [76]. Through affinity chromatography in
different formats (e.g., magnetic particles, agarose beads,
monolithic phases, and silica particles), it is straightforward
to extract the major isotype, IgG, using either Protein A or
Protein G in their immobilized forms. However, it has
remained challenging to enrich the less-abundant isotypes
from complex mixtures such as blood serum.
A novel approach has recently been developed in our
laboratory to address this need: a serial affinity chromatography
strategy is employed to first capture IgG upstream
of a Protein L affinity column that, through its
unique binding action, may capture all Igs bearing kappa
light chains (subtypes I, III, and IV) [77], which include
all five classes of human Igs. Starting with 3 μL of
blood serum per experiment, an initial characterization
of the glycomic profile of the less-abundant Igs in serum
has been measured using this approach [78], which is
currently being applied to a larger comparative study of
human cancers.
General glycoprotein/glycopeptide fractionation strategies
It is sometimes preferable to perform an indiscriminate
preconcentration of all glycoconjugates in a mixture. For
an initial glycoproteomic survey of an uncommon biological
material, it can be valuable to measure the profile of the
whole glycoproteome in order to guide subsequent investigations
of interesting subglycoproteomes. Alternatively,
the glycopeptides from a prefractionated/purified glycoprotein
proteolytic digest may be captured (and thus isolated
from non-glycopeptides) to greatly enhance their ionization
in MS. Several strategies have been developed for general
enrichment of glycoconjugates.
Hydrazide capture
A popular approach for the isolation of glycoconjugates
is to use hydrazide-coated beads, as described by
Aebersold and coworkers [79]. In general terms, the
glycoprotein/glycopeptide capture results from a covalent
bond formation between hydrazide groups on the
surface of a support medium and aldehydes present on
the carbohydrates, introduced through the oxidation of
vicinal diols. The approach has been applied to glycoproteomic
analysis of many complex materials, including
saliva [80], plasma [81, 82], blood platelets [83],
liver tissue [84], and T and B cells [85]. While this
covalent capture strategy represents a very effective
approach for select applications, it is unsuitable for
direct measurement of glycan moieties, which cannot be
readily recovered from the support material.
Boronic acid enrichment
Boronic acid-functionalized materials are emerging as attractive
options for glycocapture, as a result of their unique
Glycoconj J (2013) 30:89–117 95
ability to form reversible, covalent bonds with monosaccharides
that exhibit vicinal diols [86–88]. Microscale variations
of this approach have been demonstrated for the
enrichment of glycopeptides from standard glycoproteins
[89–91], though they have only rarely been applied to
glycoproteomic studies of biologically interesting samples
[92]. Because of their unique, universal “lectin-like” properties,
boronic acids (sometimes even referred to as boronolectins)
have also demonstrated potential for the
enrichment of nonenzymatically glycated proteins [93]
and peptides [94].
Metabolic labeling of glycans
A major limitation for chemical affinity enrichment is that
carbohydrates contain few biologically unique functional
groups that are not observed in other classes of biomolecules
such as proteins or nucleic acids. Thus, chemical
enrichment strategies for glycans may suffer from either
weak specificity or, as is the case with the hydrazide capture,
they may prove too harsh for the necessary glycan characterization
in some applications. However, an interesting
alternative is to add unique functional groups to glycoconjugates
and thus provide additional possibilities for enrichment
strategies. Utilizing the specific biosynthetic pathway
of an organism, Laughlin and Bertozzi have demonstrated
that it is feasible to incorporate azide-modified monosaccharides
into glycoconjugates in vivo or ex vivo [95].
Through a further modification of the azide groups with a
FLAG peptide [96], the metabolically labeled glycoconjugates
can be immunoprecipitated from the sample mixture.
This strategy can be applied for the general enrichment of
glycoconjugates by incorporating azido analogs of conserved
monosaccharides such as N-acetylglucosamine in
mammals, or for enrichment of different classes of glycans
with more specific residues such as azidogalactose or
azidofucose.
Hydrophilic interaction chromatography
Hydrophilic interaction chromatography (HILIC) exploits
polar interactions between analytes and a polar stationary
phase, thus providing a more subtle option for separation
of glycoconjugates than the previously described binary
capture strategies. Numerous stationary phases have been
employed for HILIC applications, including silanols,
diols, amines, amides and various cationic, anionic, and
zwitterionic functional groups. Among these alternatives,
amines and amides have been used most frequently for
the separation of glycosylated analytes. A recent report
by Gilar et al. [97] demonstrates an efficient separation
of a bovine fetuin digest, in which non-glycopeptides
were eluted first, followed by (smaller) O-linked glycopeptides,
and ending with the elution of N-linked glycopeptides.
HILIC is one of the few high-resolution
chromatography techniques that separates glycoconjugates
through significant interaction with their carbohydrate
moieties rather than other molecular attributes. In
this regard, it has been used regularly for the analysis of
fluorescently labeled glycans, as discussed further in the
forthcoming text. Beyond the brief example discussed
herein, HILIC-based separation of glycoconjugates has
been utilized in a number of applications, and a more
specialized discussion of its merits can be found in two
recent reviews, dedicated solely to this topic [98, 99].
Glycopeptide analysis
Besides merely implicating certain glycoproteins as biologically
or medically important, it is also essential to
learn about the structural details of their glycosylation,
such as the sites of glycosylation on the polypeptide
backbone, possible structural variations (micro-heterogeneity)
of a glycoforms at a particular site, or the accessibility
for biochemical interactions in a threedimensional
environment. With the ubiquity of Nglycosylated
and (particularly) the less explored Oglycosylated
sites in many mammalian glycoproteins,
this appears a “tall order” even for the best analytical
tools and expertise at hand. Yet, this type of information
appears crucial from the biomedical and pharmaceutical
viewpoints. Mining the complex glycoproteomes of
humans, animal models, and infectious agents at high
measurement sensitivity thus appears essential for many
emerging fields. Different strategies to accomplishing
these tasks include various enzymatic treatments, sitelabeling
procedures, and covalent attachment to surfaces
through the carbohydrate moieties, among others. Owing
to its high sensitivity and ability to provide mass-based
structural characterization, MS remains the key mode of
glycopeptide detection. However, ionization is a requisite
preliminary step for MS detection of an analyte, and the
high complexity of enzymatic protein digests derived
from biological materials creates a competitive environment
for ionization. This is often a major hindrance for
detection of glycopeptides that ionize poorly relative to
nonglycopeptides. Competitive ion suppression calls for
the fractionation of digest mixtures through chromatographic
media first. Consequently, hydrophilic packings
[97, 100], graphitized carbon [101], or lectins [49, 50,
96 Glycoconj J (2013) 30:89–117
60–62] used prior to MS can substantially improve the
site-of-glycosylation measurements.
For some applications, determining the site of glycosylation
may be more important than performing a complete
structural characterization of the oligosacharride. It is possible
to use either chemical or enzymatic means to label the
site of glycosylation, though the glycan itself may be lost in
the process. An example is the cleavage of O-linked Nacetylglucosamine
(O-GlcNAc) with β-elimination, followed
by the Michael addition (BEMAD) of dithiothreitol
[102]. Additionally, there are reports of the MS-based fragmentation
strategies that appear to selectively fragment glycans
after the first core GlcNAc of an N-glycan, such as the
high-energy C-trap dissociation (HCD), which can be performed
with the orbitrap MS instrument. Since this method
can reportedly yield peptide fragment ions with the core
GlcNAc residue still attached to the peptide [103], it is a
simple matter to include this as a variable modification and
thus identify the glycopeptide, including the site-ofglycosylation,
through typical database-searching strategies
for the bottom-up proteomic measurements.
The role of collision-induced dissociation in glycopeptide
analysis
An important aspect of glycopeptide analysis is a comprehensive
elucidation of its entire structure, including not only
the amino-acid sequence of the peptide backbone, but also a
detailed characterization of its carbohydrates and the site-ofglycosylation,
since not all variable structures are necessarily
occupied. Through an effective coupling of nanoflow-LC
to tandem MS, hundreds of glycopeptides can quickly be
characterized, with collision-induced dissociation (CID) being
the most widely applied for the task. During this fragmentation
technique, analytes are bombarded with an inert
buffer gas, increasing their internal vibrational energy. Once
enough energy is deposited to the molecule, fragmentation
occurs. However, the energy barrier of dissociation for the
carbohydrate part of a glycopeptide is lower than that for the
peptide backbone, resulting in a spectrum, which is dominated
by the ions originating from glycosidic bond cleavages,
while the diagnostic peptide fragments are rarely
observed. Thus, CID can be a useful technique to characterize
an oligosaccharide while it is attached to a peptide. At
the same time, the lack of fragmentation within the peptide
backbone necessitates additional MS information, such as
that provided by complementary fragmentation strategies
(see the next section.) or sub-ppm mass accuracy of the
precursor m/z of the glycopeptide.
Glycopeptide identification with CID and high mass
accuracy
An example of the application of CID in combination with
high-resolution MS data is included here to illustrate how
CID can facilitate glycopeptide discovery in complex mixtures.
In our recent study [19] of membrane-bound glycoproteins
from the select agent, Francisella tularensis subsp.
holarctica, the characterization of two glycopeptides from a
novel virulence factor was achieved using high resolution
LC-FT-ICR MS with in-source CID, frequently referred to
as source-induced dissociation (SID). Following the enzymatic
digestion, bottom-up proteomics was performed on
the glycoprotein, FTH_0069, that had been isolated by 2-D
gel electrophoresis. SID was used to monitor characteristic
glycan oxonium ions, and, with a priori knowledge of the
amino acid sequence and glycan structure, a list of theoretical
glycopeptide accurate masses was compared to the
observed glycopeptides to determine their identities, which
was achieved exclusively through sub-ppm mass accuracy.
It is clear that high-resolution MS data are vital to this
type of analysis, but the considerable preparative expertise
that was necessary to isolate the target glycoproteins from
bacterial cell lysates through density-based fractionation,
liquid extraction of membrane-bound glycoproteins, and
2D-gel electrophoresis prior to the use of bottom-up proteomics
was a prerequisite to the achieved ionization of the
glycopeptides [19]. This work exemplifies the potential of
multidimensional sample fractionation and separation techniques
in combination with MS detection as a means to
achieve a clearer understanding of individual, biologically
interesting glycoproteins, emphasizing that the current analytical
glycobiology often remains a multimethodological
task, as was the case a decade ago [104].
Electron-transfer dissociation of glycopeptides
Alternative approaches to molecular fragmentation are the
so-called electron-based methods of electron-transfer dissociation
(ETD) [105, 106] and electron-capture dissociation
(ECD) [107, 108]. While the exact mechanisms of cleavage
may still be debatable, these methods are effective in a
cleavage of the N-Cα amino acid bond. This approach to
fragmentation does not increase a peptide’s vibrational energy,
and it does not fragment the glycan portion of a
glycopeptide. The method is often deemed as appropriate
to determine the site-of-modification and has been successfully
applied to N-linked glycopeptides [109, 110].
Using modern instrumentation that is capable of quickly
alternating between the two fragmentation methods (on-theGlycoconj
J (2013) 30:89–117 97
fly) during a single LC-MS analysis, our laboratory demonstrated
the effectiveness of combining the complementary
data acquired from CID and ETD experiments to comprehensively
characterize glycopeptides derived from a number
of glycoprotein standards [111]. An example of this approach
is shown in Fig. 4a–b. The CID spectrum acquired
from a glycopeptide originating from haptoglobin (Fig. 4a)
highlights the extensive fragmentation of the carbohydrate
and its structural elucidation. Figure 4b presents the ETD
spectrum of the same glycopeptide and shows significant
fragmentation of the peptide backbone. In this spectrum, no
fragments due to glycan bond cleavage were observed,
while only a series of c- and z-type ions were present.
Based on the mass difference between the c5 and c6 ions,
which corresponds to the addition of an asparagine residue
within the attached glycan, the site-of-modification was
determined. The remaining fragments then indicated the
amino-acid sequence of the peptide.
One drawback to the electron-based fragmentation methods
is an apparent m/z limitation. We were able to successfully
and efficiently fragment a number of glycopeptides
with m/z values of up to about 1,000. Above this value,
nondissociative electron transfer seemed to dominate; this
phenomenon was also observed by others [112, 113]. Since
many glycopeptides result in multiply-charged ions with m/z
values above this threshold, the basic approach to this fragmentation
method needs to be modified. One approach that
we investigated was “supercharging” [114] the electrospray
ionization (ESI) process by adding small amounts of mnitrobenzyl
alcohol to the mobile-phase buffers. In these
preliminary experiments, we observed increases in the
charge states of several glycopeptides and improved ETD
fragmentation.
Since it appears that noncovalent interactions hinder the
dissociation of the generated fragments [112], the ETD
methods have been adapted to include a gentle CID-type
Peptide
1264.1
Peptide
Peptide
Peptide
Peptide
366.3
657.4
Peptide
1115.7
Peptide
1237.8
1345.1
1447.7
1527.5
1672.8
400 600 800 1000 1200 1400 1600 1800
m/z
a
b
1139.6
Peptide
1181.2
Peptide
1019.1
Peptide
244.2
357.5
414.7
527.9
643.4
742.1
870.8
1001.2
1340.7
1523.8
1681.7
1738.9
1795.4
400 600 800 1000 1200 1400 1600 1800
m/z
Z5
.
Z2
.
Z3
.
Z4
.
Z6
.
Z7
.
Z8
.
563.0
c5
1441.8
1565.5
1630.6
1823.6
1880.7
1937.0
[M+4H]4+
[M+4H]3+.
c11
c15
c9c6
c7
c8
c10
c12
c13
c14
Doubly-charged fragment ions
VV HPNYSQV IK
x 20
1900
+
DIGLL
VV HPNYSQV IKDIGLL
Fig. 4 Combined tandem MS
characterization of the
236
VVLHPNYSQVDIGLIK251
glycopeptide from human
haptoglobin. a Collisioninduced
dissociation (CID)
preferentially fragments the
biantennary, disialylated complex
glycan, whereas b
electron-transfer dissociation
(ETD) provides complementary
information by fragmenting the
glycopeptide backbone, which
provides the peptide sequence
and elucidates the site-specific
attachment of the glycan at
Asn241. (From Reference
[111])
98 Glycoconj J (2013) 30:89–117
of activation [113, 115] increasing the number of diagnostic
fragment ions for such peptides. In one example from the
Karger laboratory [115], a glycopeptide generated by the
Lys-C digestion of EGFR was detected as a +5 ion with an
m/z value of 1142.73. The ETD fragmentation of this large
glycopeptide allowed only 9 of the 36 amino acids to be
determined. However, upon activation of a charge-reduced
species generated during the ETD process, 20 amino acids
were determined and the correct peptide sequence was determined
through database searching.
Without modifications to the current instrumentation,
ETD may be most effective for glycopeptides featuring
smaller carbohydrates, such as N-linked structures attached
to bacterial glycopeptides [116], or O-linked oligosaccharides
with smaller chains. Indeed, several recent examples
have shown the utility of this method for the characterization
of this important class of glycopeptides. Unlike Nlinked
glycopeptides, the O-linked structures do not readily
yield a consensus sequence to indicate the site-ofmodification.
For these determinations, ETD has proven to
be a valuable tool, as a recent publication demonstrates for
several highly-charged mucin-originated O-glycopeptides
[117]. These types of proteins present unique analytical
challenges, as they are typically heavily glycosylated with
a high degree of site occupancy. Interestingly, this work
showed a high degree of sequence coverage for the glycopeptides
modified with neutral glycans, while those with
sialylated structures tended to produce fewer fragments.
Similarly, these electron-based approaches have been applied
to the hinge-region O-glycopeptides of IgA1 to determine
the sites-of-modification and to find which sites were
favored for alterations in a galactose-deficient IgA1 protein
[118]. Another investigation utilizing ETD revealed that a
significant decrease in the levels of N-acetylglucosamine
attached to IgA1 O-glycopeptides was observed in patients
diagnosed with rheumatoid arthritis [119], a change most
commonly associated with IgG. Additionally, a combination
of CID and ETD has been used to determine the glycans
attached to three sites of O-glycosylation of an amyloid
precursor protein secreted by CHO cells [120].
Glycomics
Glycomics is a less explicitly recognized member of the
group of the “omics” methodologies. Its increasing popularity
has been primarily based on the perception of numerous
investigators that the glycan components are often the crucial
functional determinants of biological events. Pending
further methodological developments and improvements in
instrumentation, glycomics is rapidly positioning itself to
become a very fertile field in addressing some key questions
of modern biology and medicine.
While direct glycomic measurements would seem at the
first glance to be somewhat limited in that the information
about the integrated function of glycoproteins is being lost
though the deglycosylation step, such measurements have a
certain practical appeal in that: (a) oligosaccharides are often
the crucial functional elements in cellular and biomolecular
interactions; (b) glycomic profiling techniques are inherently
faster and methodologically easier to multiplex than the
currently available proteomic approaches; and (c) the dynamic
concentration ranges for glycan measurements appear
to be much narrower than those for proteins in biological
fluids and tissues. However, we do not know as yet what are
the limits for glycans’ physiologically meaningful concentrations,
and how to measure glycans at very trace levels.
Glycan release procedures
The key approach of glycomics involves the deglycosylation
of isolated glycoproteins or entire complex glycoprotein
mixtures extracted from biological materials to yield a
representative array of oligosaccharides (glycans), which is
subsequently displayed as a “glycomic profile” or “glycomic
map” through a suitable bioanalytical technique. A
quantitative and reproducible release of oligosaccharides
from glycoproteins has always been a significant and difficult
issue in glycobiology. It has gained an even greater
importance in the high-sensitivity requirements of today’s
glycomic profiling. The earlier used chemical release
approaches, such as hydrazinolysis or the classical βelimination
in an alkaline medium, have now mostly been
replaced by the more gentle enzymatic deglycosylation (use
of N-glycanases) for asparagine-linked glycans [121, 122]
or microscale chemical release procedures [123–125] for
threonine/serine-linked oligosaccharides. It is now generally
agreed that N-glycans are “easier” to analyze than Oglycans,
largely due to the availability of peptide-Nglycosidase
F (PNGase F) and other glycanases, which
reliably cleave a broad range of substrates, regardless of
their glycan substitution, with only a few exceptions noted.
Alternatively, there are additional approaches to glycan release
that may need further optimization.
The release of N-linked glycans has traditionally been
performed during an extended period of time, oftentimes
requiring up to 24 h to obtain the highest possible digestion
efficiency. However, for future large-scale studies of
hundreds, if not thousands, of samples, the throughput of
the release procedure needs to be increased. One interesting
approach to reduce the incubation time involves the use of
ultra-high pressure cycling, which utilizes pressures of up to
30 kpsi [126]. Under these conditions, the activity of
PNGase F appears to be unaffected, while many glycoproteins
become sufficiently denatured for an efficient deglycosylation
in as little as 20 min [126]. Similarly to
Glycoconj J (2013) 30:89–117 99
proteomics, a microwave-assisted method to enzymatically
cleave glycans from their proteins has also been reported,
with a complete removal of the carbohydrates from monoclonal
antibody therapeutics being achieved in as little as
10 min, and up to 1 h, for other glycoprotein standards
[127, 128].
As opposed to the N-linked structures, a universal enzyme
is not available for the comprehensive removal of all
O-linked oligosaccharides. Recently, we have explored the
use of Pronase, a mixture of several proteases from
Streptomyces griseus, as the way to digest glycoproteins to
single amino acid residues. An efficient Pronase digestion
leaves the O-linked carbohydrates attached only to their
serine or threonine residues, which are then removed
through a β-elimination process during their subsequent
permethylation [129]. When compared directly to the samples
prepared through other O-glycan release methods, the
matrix-assisted laser desorption/ionization (MALDI) timeof-flight
(TOF) MS signals associated with the Pronasedigested
samples were typically 10–20 times more intense.
As a demonstration of the effectiveness of this approach,
Fig. 5 shows the O-glycan profile (acquired through
MALDI-TOF MS) from a 1-μg aliquot of a large glycoprotein,
bile-salt-stimulated lipase (BSSL), a heavily Oglycosylated
protein isolated from human breast milk. In
total, 75 oligosaccharides were detected, as listed in Table 1,
and the overall improved sensitivity of the enzymatic/chemical
cleavage procedure allowed for the detection of 40
unique structures that were not previously identified using
our earlier O-glycan release methods [130–132].
Pronase was also used to obtain N-linked carbohydrates
from bacterial glycoproteins [14], a system for which there
was no other suitable deglycosylation enzyme available.
Additionally, Pronase has also been suggested as an alternative
enzyme to PNGase F [133] for the analysis of Nlinked
glycans. However, when Pronase is applied to the
glycoconjugates, a complete degradation of the proteins to
single amino acids is a key operation that may require
incubation periods of up to 48 h. Fortunately, the digestion
times have been significantly reduced using a Pronase
immobilized on solid supports [134, 135], allowing this
procedure to be coupled on-line for direct LC-MS analyses
[134].
Carbohydrate permethylation
Following the glycan release, the carbohydrate patterns can
be displayed by any of the available analytical techniques,
including capillary electrophoresis (CE), liquid chromatography
(LC) (both discussed later), or our currently preferred
method, MS. The use of MS allows the oligosaccharides to
not only be accurately quantitated, but it also enables a more
definitive understanding of the exact structures whose abundances
are often altered in disease states. While we have
explored the possibility of using nano-LC coupled to ESIbased
instruments [136], with both ion-trap and FT-ICR
mass spectrometers, our currently preferred method is
MALDI-TOF MS due to its speed of data acquisition, the
ability to control the data collection (i.e. collect more laser
shots per spot to improve the signal-to-noise ratio for improved
sensitivity of low-abundance analytes), and the ability
to perform high-energy CID tandem MS analyses. This is
particularly valuable in characterizing some isomeric glycans
[137].
To improve the overall MS performance, our glycomic
platform is based on permethylating all analytes, which
involves a modification converting the hydroxyl groups present
on the carbohydrate to methoxide moieties, esterifying the
carboxylate of any sialic acid residues present, and introducing
a methyl group to the nitrogen of the N-acetyl groups of Nacetylglucosamine
(GlcNAc) or N-acetylgalactosamine
(GalNAc). Such a derivatization offers several advantages,
Mass (m/z)
% Intensity
Fig. 5 O-linked glycans released by a combined enzymatic/chemical method derived from a 1-μg aliquot of bile salt-stimulated lipase (BSSL).
(From Reference [129])
100 Glycoconj J (2013) 30:89–117
Table 1 O-linked oligosaccharides
identified from a 1-μg aliquot
of bile salt- stimulated
lipase (BSSL). The structures in
red were unique carbohydrates
identified via the enzymatic/
chemical release procedure
Obs. Mass Calc. Mass ΔMass Composition
866.444 866.437 0.007 Hex1HexNAc1Deoxyhexose2
879.415 879.432 0.017 Hex1HexNAc1NeuAc1
896.446 896.447 0.001 Hex2HexNAc1Deoxyhexose1
937.474 937.474 0 Hex1HexNAc2Deoxyhexose1
967.487 967.484 0.003 Hex2HexNAc2
1083.522 1083.532 0.01 Hex2HexNAc1NeuAc1
1124.557 1124.558 0.001 Hex1HexNAc2NeuAc1
1130.57 1130.558 0.012 Hex4HexNAc1
1141.555 1141.573 0.018 Hex2HexNAc2Deoxyhexose1
1171.572 1171.584 0.012 Hex3HexNAc2
1212.616 1212.611 0.005 Hex2HexNAc3
1240.581 1240.606 0.025 Hex1HexNAc1NeuAc2
1298.649 1298.647 0.002 Hex1HexNAc2Deoxyhexose1NeuAc1
1315.65 1315.663 0.013 Hex2HexNAc2Deoxyhexose2
1328.654 1328.658 0.004 Hex2HexNAc2NeuAc1
1345.648 1345.673 0.025 Hex3HexNAc2Deoxyhexose1
1386.685 1386.7 0.015 Hex2HexNAc3Deoxyhexose1
1416.708 1416.71 0.002 Hex3HexNAc3
1489.706 1489.752 0.046 Hex2HexNAc2Deoxyhexose3
1502.74 1502.747 0.008 Hex2HexNAc2Deoxyhexose1NeuAc1
1519.819 1519.762 0.057 Hex3HexNAc2Deoxyhexose2
1590.788 1590.8 0.012 Hex3HexNAc3Deoxyhexose1
1620.819 1620.81 0.009 Hex4HexNAc3
1661.817 1661.837 0.02 Hex3HexNAc4
1676.812 1676.836 0.024 Hex2HexNAc2Deoxyhexose2NeuAc1
1689.819 1689.832 0.013 Hex2HexNAc2NeuAc2
1747.86 1747.873 0.013 Hex2HexNAc3Deoxyhexose1NeuAc1
1764.873 1764.889 0.016 Hex3HexNAc3Deoxyhexose2
1777.852 1777.884 0.032 Hex3HexNAc3NeuAc1
1794.897 1794.899 0.002 Hex4HexNAc3Deoxyhexose1
1818.85 1818.911 0.061 Hex2HexNAc4NeuAc1
1835.901 1835.926 0.025 Hex3HexNAc4Deoxyhexose1
1865.922 1865.936 0.014 Hex4HexNAc4
1921.966 1921.963 0.003 Hex2HexNAc3Deoxyhexose2NeuAc1
1938.968 1938.978 0.01 Hex3HexNAc3Deoxyhexose3
1951.951 1951.973 0.022 Hex3HexNAc3Deoxyhexose1NeuAc1
1981.942 1981.984 0.042 Hex4HexNAc3NeuAc1
2009.975 2010.015 0.04 Hex3HexNAc4Deoxyhexose2
2039.989 2040.026 0.037 Hex4HexNAc4Deoxyhexose1
2069.996 2070.036 0.04 Hex5HexNAc4
2098.04 2098.031 0.009 Hex4HexNAc2NeuAc2
2113.007 2113.067 0.06 Hex3HexNAc3Deoxyhexose4
2126.046 2126.062 0.016 Hex3HexNAc3Deoxyhexose2NeuAc1
2156.022 2156.073 0.051 Hex4HexNAc3Deoxyhexose1NeuAc1
2214.094 2214.115 0.021 Hex4HexNAc4Deoxyhexose2
2227.082 2227.11 0.028 Hex4HexNAc4NeuAc1
2268.065 2268.137 0.072 Hex3HexNAc5NeuAc1
2300.117 2300.152 0.035 Hex3HexNAc3Deoxyhexose3NeuAc1
2315.135 2315.163 0.028 Hex5HexNAc5
Glycoconj J (2013) 30:89–117 101
including (i) an improved sensitivity of 10 to 20 times over the
native glycans; (ii) converting the acidic structures (i.e. sialylated
glycans) to neutral solutes, permitting a complete glycan
profile to be monitored through the positive-ion mode; (iii)
enhanced cross-ring fragmentation, which enables a more
definitive structural characterization; and (iv) making the
resulting glycans fairly hydrophobic, permitting their separation
by reversed-phase LC, if needed.
The present-day permethylation procedure employs
methyl iodide and dimethylsulfoxide (DMSO). This approach
can be traced back to the 1960s [138] and its later
updated features [139, 140]. More recently, we have introduced
procedures using sodium hydroxide beads loaded into
spin-column “reactors” [141, 142] rather than using a slurry
of the reagent. By performing the reaction in spin-columns,
the excess sodium hydroxide can be easily removed from
the reaction solution, minimizing the so-called “peeling”
reactions that can occur at the high pH values typically
encountered during the sample recovery in the slurrybased
methods. These undesirable degradation reactions
often significantly limit the sensitivity of the slurry-based
approaches for trace-level oligosaccharides. We have further
refined our protocol to use dimethylformamide (DMF) in
the place of DMSO [136], since this solvent participates in
two adverse side reactions, which negatively influence the
overall sensitivity and introduce uncertainty into the quantitation.
One of these reactions ultimately produces a series
of satellite peaks separated by +30 Da [143], while the
second reaction results in the regeneration of their native
“closed-ring” configurations of the glycan alditols [140].
Our laboratory prefers these types of structures since they
do not have a potentially reactive aldehyde group at their
reducing ends. When our most up-to-date protocol (typically
resulting in the detection of a single derivatized analyte) is
applied to microliter volumes of blood serum, we obtain
reliable profiles such as the one shown in Fig. 6. In this
profile of a women diagnosed with late-stage recurrent
ovarian cancer, about 55 unique m/z values were detected,
which correlate to the known glycan structures. Of particular
interest to our laboratory are the structures present in the
higher mass region of the spectrum, which are highlighted in
the inset for this figure. This particular spectral area is
populated with the tri- and tetra-antennary structures with
varying levels of fucosylation and sialylation. We have
Table 1 (continued)
Obs. Mass Calc. Mass ΔMass Composition
2388.139 2388.204 0.065 Hex4HexNAc4Deoxyhexose3
2401.167 2401.199 0.032 Hex4HexNAc4Deoxyhexose1NeuAc1
2472.168 2472.236 0.068 Hex4HexNAc5NeuAc1
2489.18 2489.252 0.072 Hex5HexNAc5Deoxyhexose1
2562.169 2562.047 0.122 Hex4HexNAc4Deoxyhexose4
2575.224 2575.289 0.065 Hex4HexNAc4Deoxyhexose2NeuAc1
2588.056 2588.284 0.228 Hex4HexNAc4NeuAc2
2663.267 2663.341 0.074 Hex5HexNAc5Deoxyhexose2
2676.226 2676.336 0.11 Hex5HexNAc5NeuAc1
2749.292 2749.378 0.086 Hex4HexNAc4Deoxyhexose3NeuAc1
2762.295 2762.373 0.078 Hex4HexNAc4Deoxyhexose1NeuAc2
2837.277 2837.43 0.153 Hex5HexNAc5Deoxyhexose3
2850.324 2850.425 0.101 Hex5HexNAc5Deoxyhexose1NeuAc1
2938.385 2938.478 0.093 Hex6HexNAc6Deoxyhexose1
3011.371 3011.519 0.148 Hex5HexNAc5Deoxyhexose4
3024.426 3024.515 0.089 Hex5HexNAc5Deoxyhexose2NeuAc1
3112.365 3112.567 0.201 Hex6HexNAc6Deoxyhexose2
3125.429 3125.562 0.133 Hex6HexNAc6NeuAc1
3198.425 3198.604 0.179 Hex5HexNAc5Deoxyhexose3NeuAc1
3211.492 3211.653 0.161 Hex5HexNAc5Deoxyhexose1NeuAc2
3286.418 3286.655 0.237 Hex6HexNAc6Deoxyhexose3
3299.491 3299.652 0.161 Hex6HexNAc6Deoxyhexose1NeuAc1
3372.505 3372.691 0.186 Hex7HexNAc7Deoxyhexose1
3388.388 3387.704 0.316 Hex7HexNAc7Deoxyhexose1
3473.497 3473.741 0.244 Hex6HexNAc6Deoxyhexose2NeuAc1
3647.482 3647.83 0.348 Hex6HexNAc6Deoxyhexose3NeuAc1
102 Glycoconj J (2013) 30:89–117
proposed these types of analytes as the key structures in
several pathological conditions, including breast [136, 144],
ovarian [145], lung [146], liver [147, 148], prostate [149],
and esophageal [150] cancers.
The analysis of permethylated oligosaccharides has also
been successfully applied to those structures modified with
phosphate or sulfate groups. Both of these moieties seem to be
stable throughout the permethylation procedure, with the
phosphate group becoming singly or doubly esterified [151,
152]. Most probably, extended reaction times ensure a
complete esterification of the phosphate group. Conversely,
sulfate groups attached to a carbohydrate are unaffected by the
permethylation procedure and retain their negative charge
[152, 153]. To detect sulfated glycans in a mass spectrometer’s
positive ion mode, our group has developed a “double-permethylation”
procedure [153]. In this procedure, the sulfated
glycans are subjected to our spin-column approach for permethylation
and recovered from the reaction mixture. The
sulfate group is then chemically removed via a treatment with
acidified methanol and the samples are permethylated a
1499.0 2202.6 2906.2 3609.8 4313.4 5017.0
Mass (m/z)
1.6E+4
0
10
20
30
40
50
60
70
80
90
100
%Intensity
2808.41
1851.93
2056.03
2447.23
2982.50
2260.16
3618.82
2621.33
2097.08
2866.44
1799.91
1595.81
3792.91
2412.23
2505.27
2208.12
3257.65
2692.38
3707.86
4429.23
3431.71
2301.27
4603.30
4068.04
3070.55
4242.13
3966.99
3880.97
3227.63
1922.99
1677.87
1881.97
3053.54
2896.46
2004.02
4777.39
2086.06
3700 3900 4100 4300 4500 4700
Mass (m/z)
1110.2
0
10
20
30
40
50
60
70
80
90
100
%Intensity
3792.91
3706.86
4429.22
4603.30
4066.04
4242.13
3966.99
3880.97
Fig. 6 N-linked glycomic profile of an ovarian cancer patient. The inset highlights the high-mass region where many important trace-level
oligosaccharides are located. (From Reference [145])
Glycoconj J (2013) 30:89–117 103
second time using deuterated methyl iodide to label the site of
sulfation [153]. Additionally, following the first permethylation
step, sulfated glycans may be fractionated based on their
degree of sulfation only, since sialic acids are rendered as
neutral, by strong-anion chromatography [154]. Following a
desalting procedure, the sulfate group is chemically removed
and the site-of-sulfation is repermethylated using deuterated
methyl iodide.
Quantitation of oligosaccharides through stable isotope
labeling
Based on the premise that there are many suspected or
proven associations of human disease conditions with aberrant
glycosylation, the rapid comparative profiling of structurally
known, or at least tentatively identified, glycans
could be significant as the starting point for more in-depth
investigations of these diseases. Comparative glycan profiling
can similarly be applied to a number of biological
studies of any “normal” or “perturbed” systems, a comparison
of glycosylation in different body organs, chemotaxonomies
of different organisms, phylogenetic trees, etc. In all
of these studies, high precision and accuracy in measuring
glycan abundances for some or all profile constituents
becomes essential. Here, the use of isotopic labeling for
glycans and MS measurements opens new possibilities. It
provides an approach in which multiple samples can be
measured simultaneously and directly compared during a
single data acquisition. Through the use of methyl iodide
with varying deuterium substitutions, up to four samples can
be simultaneously monitored. Our research group has incorporated
isotopic labeling into the permethylation platform
[155]. Importantly, the linearity of this method was
acceptable at nearly two orders of magnitude, so that it
could be applied to various differential glycomic studies.
An example of this technique is shown as Fig. 7, with a
MALDI-MS profile comparing the different expression levels
of N-linked glycans in the early developmental stages of
Drosophila melanogaster (unpublished results). This study
indicated that several high-mannose type structures are
much more abundant in the larval stage (shown as the red
m/z values) than in the embryonic state (indicated by the
green m/z values). In a different research group, 13
C-labeled
methyl iodide has been used to incorporate stable isotopes
into glycan structures through permethylation [156], in
which isobaric labeling has been achieved through the use
of 13
CH3I and 12
CDH2 [157]. However, this approach introduces
a mass difference of only 0.002922 Da for each site of
derivatization. This small mass difference is difficult to
detect by modern MALDI-based instruments, but it can be
easily measured with a high-resolution mass spectrometer
(i.e. an FT-ICR instrument or an orbitrap).
Isotopically-coded tags may further be introduced into
the carbohydrate structure through other methods and locations
on the glycan structure. The free reducing end of an
oligosaccharide provides a convenient site for modification
and several isotopically-coded chromophores can be incorporated
at this location, including aniline [158–160], 2aminopyridine
[161], 2-aminobenzoic acid [162], and 1phenyl-3-methyl-5-pyrazolone
[163]. Additional tags have
been synthesized, including (13
C6 and 12
C12) 4-phenethylbenzohydrazide
[164], a hydrophobic tag that may enhance
the sensitivity of ESI-based measurements through a more
efficient desolvation process [165, 166], and a novel set of
tetraplexed tags [167, 168], each separated by 4 Da and
analyzed by a direct infusion into an ESI-based q-TOF MS
1100 1440 1780 2120 2460 2800
1.8E+4
0
10
20
30
40
50
60
70
80
90
100
1402.9
1451.6
1515.0
1222.8
1649.1
1140.8
1189.8
2501.6
1171.5
1567.0
1435.9
1862.2
1108.7
1293.5
2288.5
1477.6
2075.4
1783.2
1657.1
1416.9
1701.7
1375.6
1616.1
1988.3
1345.8
1580.0
1905.8
2192.5
2109.9
2396.0
1591.0
Mass (m/z)
% Intensity
Fig. 7 N-linked glycans,
isotopically-labeled through
permethylation, comparing different
developmental stages of
Drosophila melanogaster. The
m/z values in red are associated
with the larval stage, while
those in green indicate an
embryonic state
104 Glycoconj J (2013) 30:89–117
instrument. Alternatively, in a closely-related analogue to the
stable-isotope labeling by the amino acids in a cell culture (the
so-called SILAC method, which is widely employed in the
proteomics field), isotopically-labeled glutamine, which is
further used as the sole source of nitrogen for the synthesis
of N-acetylglucosamine, N-acetylgalactosamine and the sialic
acids, has been reported [169] and utilized in conjunction with
cultured mouse embryonic stem cells.
Analysis of isomeric structures
To what extent is the precise knowledge of a glycan structure
necessary for understanding a biological phenomenon?
The answer is that we do not know, but if one considers the
molecular nature of sugar-sugar interactions or different
conformational situations in a glycocalyx during protein
binding processes, the geometrical considerations appear
important at once. An excellent example for the need of
glycan diversity is the regulation of our native and adaptive
immune responsiveness [170, 171] to pathogens, allergens,
and other foreign substances.
Glycomic in-depth studies can amount to very substantial
scientific activities in the exact structural elucidation of the
individual glycans in a profile. However, some structural
ambiguities frequently arise. The exact positions of terminating
sialic acids of incompletely sialylated structures are
seldom known, as is the location of the biologically important
fucosyl substitution. The differences in linkages between
various monosaccharide residues are currently
known for only some of the most abundant natural glycoproteins
that have been extensively studied through the
entire set of structural tools. The use of the multiple levels
of tandem MS has been helpful in assigning the more
definitive structures for oligosaccharides [172–174], but
the sample quantities used in such experiments can be
substantial. Additionally, the more structurally informative
techniques, such as NMR spectrometry, are currently somewhat
insensitive for a number of biological investigations.
The propensity of glycans to form many different isomers
represents one of the major challenges in glycomic measurements.
MS alone cannot distinguish glycan isomeric forms,
which inherently have the same mass, although high-energy
collision-induced dissociation (CID) in some tandem MS techniques
seems to promote the cross-ring fragmentation cleavages
(across the sugar pyranose rings) that are needed to form
more informative ionic fragments [175, 176]. Additional
approaches in distinguishing sialyl linkage isomers may utilize
the different reactivity of α2-3-linked and α2-6-linked residues,
as demonstrated in our recent study [177]. While a previous
study demonstrated the ability to discern the different linkage
isomers of sialic acids through an esterification reaction using
methanol [178], our permethylation-based platform required a
major modification. Since this derivatization also results in an
esterification of the carboxylate group of sialic acids, we modified
an amidation reaction [179] to meet the structural elucidation
needs. In this modification, the α2-6-linked sialic acids
become amidated, while the α2-3-attached residues are lactonized,
becoming insensitive towards derivatization. During a
subsequent permethylation, the nitrogen of the amide groups
accepts two methyl groups, while the lactone is cleaved and the
resulting carboxylate becomes esterified, introducing a mass
difference of 13 Da.
We have applied this linkage-specific amidation technique
to several different cancer investigations and it
appears that for the same glycans in different pathological
conditions, a certain level of specificity may be associated
with the different ratios of the linkage isomers, as shown in
Fig. 8. This figure compares the ratios of the different
linkage isomers of a fucosylated, triantennary trisialylated
0
10
20
30
40
50
60
70
80
90
100Range
3 α2-3
2 α2-3
1 α2-6
1 α2-3
2 α2-6
3 α2-6
Control NS vs. NSCLCFS
p-value: 4.01x10-7
AUC: 0.0917
Control NS vs. NSCLC FS
p-value: 1.11x10-5
AUC: 0.865
Control NS vs. NSCLC FS
p-value: 4.44x10-3
AUC: 0.275
Control Never Smoked (NS) (N=17)
Control Former Smoker (FS) (N=11)
NSCLC Former Smoker (FS) (N=29)
Control FS vs. NSCLC FS
p-value:2.41x10-3
AUC: 0.161
Control FS vs. NSCLC FS
-value: 0.283
AUC: 0.692
Control FS vs. NSCLC FS
p-value: 0.0495
AUC: 0.450
a
0
20
40
60
80
100
Range
3 α2-3
2 α2-3
1 α2-6
1 α2-3
2 α2-6
3 α2-6
Control (N=30)
Prostate Cancer Cure (N=30)
Prostate Cancer Fail (N=30)
Control vs. Fail
p-value: 5.53x10-8
AUC: 0.892
Control vs. Fail
p-value:8.35x10-8
AUC: 0.885
Control vs. Fail
p-value: 2.35 x 10-7
AUC: 0.143
Control vs. Fail
p-value: 7.94 x 10-8
AUC: 0.132
b
Fig. 8 Notched-box plots comparing the different ratios of sialic acid
linkages of a fucosylated triantennary trisialylated glycan in a a lung
cancer patients (from Reference [146]) and b prostate cancer patients
Glycoconj J (2013) 30:89–117 105
glycan from our recent study of lung cancer (Fig. 8a) and the
effects of smoking on glycomic profiles [146] and a prostate
cancer study (unpublished results) (Fig. 8b). This figure
demonstrates that the levels of the isomers containing three
and two α2-3-linked sialic acids are decreased in their
abundances in control individuals who are former smokers
and the lung cancer patients who had formerly smoked
[146]. However, these same isomers appeared to be differently
altered in their levels in patients diagnosed with prostate
cancer (unpublished results). The isomer with two
α2-6-linked sialic acids also showed opposing trends. In
the lung cancer study [146], this isomer was elevated, while
in the prostate cancer patients, the abundance of this isomer
was suppressed, along with the isomer containing three
α2,6-linked sialic acids (unpublished results). Unfortunately,
one of the isomers was observed only at very low levels,
oftentimes not detectable in the lung cancer study, making a
definitive conclusion about its ratios difficult at the present
time [146].
Another structural diversity may be biosynthetically introduced
into a carbohydrate structure through the inclusion
of a fucose residue, which is a monosaccharide residing on
either the core GlcNAc connecting an N-linked carbohydrate
to its protein, or on one of the “branching” GlcNAc
units. Altered abundances in one of the possible locations
are often associated with diseases and inflammation [8], and
due to their isomeric nature, they cannot be readily discerned
by MS alone. However, through a controlled enzymatic
cleavage of the glycan structure with a nonspecific
sialidase and a β-galactosidase, a mass difference can be
introduced, as the outer-arm fucose residues inhibit the
activity of β-galactosidases, when used as a specific
reagent, leaving the galactose associated with its GlcNAc.
Consequently, this presents a convenient method to monitor
the ratios of core-to-outer-arm fucosylation. We have applied
this approach in several recent cancer studies and in a
preliminary observation, it seems that breast (unpublished
results), ovarian [145], and lung [146] cancers show increased
levels of outer-arm fucosylation, while liver cancer
favors elevated levels of core-fucosylated glycans (unpublished
results). This type of enzymatic digestion has also
been performed by another group in combination with an
LC-based methodology, in which the two former isomers
will now result in different retention times after the exoglycosidase
digestion [180–183]. Their conclusions largely
match our MS results.
Some promise in the sugar isomer resolution is also held
by ion-mobility spectrometry (IMS), particularly in combination
with time-of-flight (TOF) MS, in which the latter
could structurally confirm the success of IMS separation
[184–186]. Interestingly, the observations made a number
of years ago that the CE of fluorescently-tagged oligosaccharides
can be quite effective in resolving at least some
glycan isomers [187, 188] remain topical to this date. The
glycosidase-assisted micro-digestions of biological samples,
in conjunction with CE, can be of some help in structural
elucidation through the peak shift observations. However,
this could hardly become a widely used strategy for extensive
characterizations, since the digestions often become
unreliable for trace-level samples and some exoglycosidases
are quite expensive.
Separations in glycomics
Due to the usual high complexity of the released glycan
mixtures and frequently, the low abundances of some
biologically-important glycoconjugates, the high sensitivity
of analytical measurements becomes mandatory. In addition
to the inherently sensitive MS techniques as detailed above,
carbohydrate profiles may also be recorded by LC, CE, or
the “chip-based” electrophoretic methods.
Liquid chromatography
Due to the hydrophilic nature of glycans, the separation
systems using a normal-phase mode of chromatography or
hydrophilic-interaction chromatography (HILIC), a term
first used by Alpert in 1990 [189], are being increasingly
employed. In recent years, several technological innovations
have been realized, presenting new opportunities for improved
separations of carbohydrates. Among the most interesting
developments is the use of ultra-high pressure LC
(UPLC) systems, which facilitate enhanced separation efficiencies
and reduced analysis times due to a substantial
decrease of the particle size. In the area of analytical glycobiology,
this has been demonstrated in a head-to-head comparison
of 3.0 μm and 1.7 μm sorbent particles [190] for 2aminobenzamide-labeled
glycans originating from bovine
fetuin (Fig. 9). In this comparison, the UPLC analyses were
faster, and due to the higher column efficiencies, were able
to resolve several isomers that were not observed with the
column packed with the larger particles.
Another interesting development is the use of zwitterion
HILIC or Zic-HILIC phases for glycoconjugate analysis
[191]. Repeated runs of human IgG-derived glycans demonstrated
a good reproducibility along with a high resolution
between several isomeric structures. Additionally, glycopeptides
differing only by an isomeric glycan structure were
reported to be resolved with this stationary phase.
An alternative medium to the normal-phase packings is
graphitized carbon, a stationary phase that offers several
desirable features, including good chemical inertness, stability
across a very extensive pH range, and the ability to
maintain its integrity at elevated temperatures. Of particular
importance is the high selectivity of graphitized carbon,
106 Glycoconj J (2013) 30:89–117
which allows for resolution of the positional isomers of
permethylated glycans [192] and various linkage [193] and
positional isomers of native glycans [194]. Although the
mechanism(s) of retention with this material are complex,
the separations are likely to be based on adsorption through
hydrophobic, hydrophilic, and polar interactions [195]. Not
surprisingly, several parameters have been shown to influence
the retention of glycans, including the ionic strength (to
which sialylated structures appeared to be particularly sensitive),
as increasing the salt concentrations in the mobile
phases resulted in increased retention times and broadened
peak widths [195]. The retention of these oligosaccharides
also appears to be dependent on the mobile phase pH, with
longer retention time being associated with lower pH values
[195]. Interestingly, neutral oligosaccharides seem largely
unaffected by changes in the mobile-phase composition
[195]. When compared to other retention mechanisms,
which are based on partitioning, the retention of carbohydrates
on graphitized carbon packings is increased at elevated temperatures
[195].
A fairly recent and convenient innovation available to
glycobiology researchers is the “LC chip” [196, 197]. In this
miniaturized analytical format, a trapping column, switching
valve, and an LC column have all been integrated into a
single biocompatible unit, which reduces the number of
manual connections and allows for easy measurements as
a specialized “MS inlet.” We have demonstrated the applicability
of this approach for glycomic analyses through the
separation of permethylated glycans using a C18 chip interfaced
directly to an ESI MS [136]. This combination provided
complementary results to a previous MALDI-based
glycomic study of blood serum samples from breast cancer
patients [144]. Among the advantages of using ESI were the
high reproducibility and repeatability of the analytical signal.
Other groups have recently applied this chip-based
approach to the analysis of the neutral and acidic oligosaccharides
isolated from human breast milk [198–200] using
the graphite chip packings. Using the graphite chips, the
Lebrilla group also reported that nearly 200 structures were
identified using exact masses in human serum [201], which
included some isomeric analyses [202].
While the previous chip studies were conducted on glycans
that had been released by a typical “in-solution” procedure,
the more recent chip designs now include an
integrated “PNGase F reactor”, which allows for the glycan
cleavage to be performed on-line [203]. Using such a chip
type, within only a 6-s incubation period, approximately
98 % of an antibody was deglycosylated [203], indicating
significant improvements in routine glycomic analyses.
Following the cleavage, the glycans were trapped on a
graphite preconcentrator and subsequently separated using
a column packed with the same adsorbent.
Capillary electrophoresis
In the search for glycomic profiling techniques of high
sensitivity, the development of capillary electrophoresis/laser-induced
fluorescence (CE-LIF) for derivatized glycans
in the early 1990s [22, 23, 204] preceded most efforts of
using biomolecular MS in the area. While, in the following
years, the use of MS-based methodologies dramatically
increased in numbers and specific applications due to the
possibilities of direct, positive identification of glycans from
their MS “molecular fingerprints”, it is remarkable that the
applications of CE-LIF still continue to develop at their very
respectable pace. While CE-LIF is capable of recording
Fig. 9 Fetuin-derived glycans
separated by a LC column
packed with 3 μm particles and
b UPLC column using 1.7-μm
sorbent. (From Reference
[190])
Glycoconj J (2013) 30:89–117 107
reproducibly very complex glycan profiles from hydrolyzed
glycoproteins and glycoprotein mixtures at very high sensitivity,
this approach generally lacks the solute identification
capabilities that the MS and tandem MS/MS so easily offer.
This general drawback notwithstanding, new applications,
developmental trends, and hardware developments are primarily
driven by the relative instrumental simplicity of CELIF,
miniaturized automatable designs needed in biotechnology
industry and other routine measurements, and not at
the least by the capability of CE and related techniques for
resolving isomeric glycans that MS generally lacks.
The sample preparation aspects and a correct choice of the
fluorescence tag are still the subject of different communications
in the current literature. While different fluorescencelabeling
reagents were explored in the early work on CE-LIF
of carbohydrates, the introduction of 8-aminopyrene-1,3,6trisulfonic
acid (APTS) as the labeling reagent by Guttman
and co-workers [25, 205] has led to its nearly uniform acceptance
and commercial use. The basis of this reagent’s utility is
the reaction between its aromatic moiety and the reducing end
of carbohydrates, followed by a reducing stabilization of the
analytes. As the recent examples of other fluorescencelabeling
approaches, 4-fluoro-7-nitro-2,1,3-benzoxdiazole
[206] and rhodamine 110 dye with a large fluorescence quantum
yield [207] were also investigated. Each derivatization
approach and the correspondingly modified glycans necessitate
their own procedural modifications and distinct compositions
of buffers and separation media.
Since numerous new biopharmaceuticals, including
monoclonal antibodies and vaccines, feature glycosylated
structures, CE-LIF is rapidly becoming the method of
choice in providing quantitative glycosylation profiles to
support product efficacy and minimize immunogenicity
effects [208, 209] in the biotechnology industry. In contrast
to profiling glycans in the biomarker discovery area, most
biotechnologically-oriented applications place less stringent
demands on sensitivity and identification of the unknown
components. However, the demands on sample throughput
can be substantial, and the same is perspectively true in
clinical analyses, as evidenced through the recent CEbased
equipment to evaluate glycan profiles of patients with
liver diseases [210, 211]. As seen in Fig. 10, an optimized
series of sample (3 μL of a patient’s serum) treatments,
leading to APTS labeling and, ultimately to CEfluorescence
glycan profiling, can now be performed for
the benefit of clinical diagnosis while using a DNA sequencer
equipment.
While the capillary-based glycan separations of biomedical
interest have been amply demonstrated in the literature,
there is a general trend toward the use of chip-based separatory
systems [212–216], which offer substantial gains in
terms of faster and more reproducible separations and measurements.
As an early example of this trend [213], we show
the separation of glycans extracted from a blood serum
sample of a breast cancer patient (Fig. 11). In contrast to
the separation in a fused silica capillary, which has taken
over a 30-min. period, the demonstrated run using a spiral
Fig. 10 Optimized workflow
for the preparation of glycancontaining
samples prior to a
capillary electrophoretic analysis.
(From Reference [211])
Fig. 11 Chip-based electropherogram of N-linked glycans from the
serum of a patient with late-stage breast cancer. (From Reference [213])
108 Glycoconj J (2013) 30:89–117
channel design has taken only 2.8 mins of analysis time at
comparable or better separation efficiency. More recently,
even more efficient separations were achieved with similar
biological samples using a chip with a more advanced
serpentine design [217].
The current problem of the CE analysis of glycans
remains its limited compatibility with MS. To perform at
its best separation efficiency, capillary zone electrophoresis
(CZE) tolerates only minute quantities of the analyzed materials
at the inlet of the separation capillary, and the same is
true for its microchip versions. This, in turn, means that
most on-column preconcentration remedies (e.g., stacking
or solute trapping) are not particularly helpful to enhance the
signals in different CE-MS combinations. The considerably
more “MS-compatible” is capillary electrochromatography
(CEC), which still remains one of the few viable options for
analyzing complex glycan mixtures [130, 131] with the use
of polymeric monolithic columns. We have investigated the
merits of CEC-MS coupling using the ESI inlet to an ion
Fig. 13 Capillary
electropherograms
demonstrating an on-line exoglycosidase
digestion procedure
for carbohydrate structural
analysis. (From Reference
[224])
Fig. 12 Capillary electropherogram depicting the resolution of some
isomeric glycans derived from a murine monoclonal antibody. (From
Reference [220])
Glycoconj J (2013) 30:89–117 109
trap [131, 218] and ICR MS [130] and, through a sample
microdeposition device, to a MALDI-TOF instrument
[132]. More recently, an elaborate instrumental arrangement
involving CE-LIF-MS coupling was reported for a successful
analysis of recombinant monoclonal antibody glycans as
APTS-labeled analytes [219].
A key advantage of CE in glycan analysis has been its
capability to resolve isomers. This is exemplified with the
glycan profile of a monoclonal antibody compared to several
glycan standards (Fig. 12) [220]. Recombinant glycoproteins
and similar cases of more “predictable” glycan
biological sources have been significantly aided by the
favorable analytical attributes of CE. However, the lack of
glycan standards has far more seriously impaired the use of
CE in the biomarker discovery area than perhaps with any
other analytical method.
The use of sequential digestion with exoglycosidase
enzymes, followed by the CE analyses [221, 222] can be
useful in structural assignments and glycan mapping with
relatively simple glycoproteins, but less effective with biologically
complex systems. Additionally, exoglycosidases
are relatively expensive reagents. A recent innovative approach
to decrease consumption of the exoglycosidase
enzymes together with enhanced analytical performance
appears to be the phospholipid-assisted CE [223, 224],
where the phospholipid additives are used in a segmentation
process to incorporate these enzymes. An example is outlined
in Fig. 13: multiple enzymes have been used, sequentially,
for selective incubation periods inside the CE
capillary, to cleave the APTS-labeled glycans from a recombinant
glycoprotein cancer drug [223].
Further advances in CE-MS of glycoconjugates are clearly
desirable for further development of the field of analytical
glycobiology. Ironically, the best up-to-date CE separations
have been achieved with the buffer media and polymeric
additives, which are largely incompatible with typical MS
conditions. It is currently hard to predict whether the desired
improvements will come from the use of different derivatization
schemes and separation conditions, any breakthrough
developments in the CE-MS interfacing technologies, MS
designs, or combined incremental improvements in all these
areas. Various advances in CE-MS of glycoconjugates have
been the subject of recent reviews [216, 225, 226].
Conclusions
The field of analytical glycoscience has evolved substantially
during the last decade. The major incentives
for this methodological progress in the field is the close
connection of glycoconjugates with some of the most important
fields of human activities and scientific endeavors: the
search for disease biomarkers; recombinant glycoprotein
pharmaceuticals; developmental biology; microbiology; immunology;
plant biology; biofuels, among others. This brief
authoritative review has summarized only a small crosssection
of the vast field affected during the last 20 years
through new analytical techniques and instrumentation.
Glycoproteins are methodologically unique among the
different classes of glycoconjugates, so that the interlinked
fields of glycomics and glycoproteomics necessitate a different
emphasis on how the biological samples are fractionated,
enzymatically or chemically treated, and analyzed. The
emphasis on very high measurement sensitivity, which is
clearly mandated by the very large dynamic concentration
range in which different glycoproteins occur in biological
samples, favors the most technologically advanced forms of
instrumental methods such as MS, miniaturized LC and CELIF.
The complementary nature of these analytical approaches
in glycomic and glycoproteomic measurements will be essential
in future successful investigations, as will be further
technological improvements and effective coupling (or “hyphenation”)
of these analytical techniques. Substantial advances
in computer-aided evaluation of the highly complex
analytical data present unprecedented opportunities for future
explorations of the mysterious problems of the glycome and
glycoproteome.
Acknowledgments Our research activities in analytical glycobiology
and disease biomarker discovery have been supported by the following
grants: 2R01GM024349 from the Institute of General Medical Sciences;
5U01CA128535 from the National Cancer Institute; and P41
RR018942 from the National Center of Research Resources, US Department
of Health and Human Services.
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