MucosalImmunology | VOLUME 6 NUMBER 2 | MARCH 2013
nature publishing group
See COMMENTARY page XX
The conducting airways of the lung provide mecha nical, chemi-
cal, and immunological barriers to inhaled parti culates, including
respiratory pathogens. One of the first lines of defense for the lung
is the mucus layer lining the tracheo bronchial airways. The appro-
priate production and effective clearance of mucus is essential for
respiratory steri lity and health. The critical function of mucus
clearance is highlighted by lung diseases in which mucus clearance
is reduced, i.e., cystic fibrosis, the chronic bronchitis component
of chronic obstructive pulmonary disease, and asthma. Effective
clearance occurs by the unidirectional transport of mucus over
an underlying bed of beating cilia, or, secondarily, by cough clear-
ance. The effective transport of a soluble, viscoelastic mucus layer
is dependent not only on the ciliary beat, but also on the main-
tenance of the fluid and ion transport mechanisms required for
homeostasis within the airway surface micro environment. 1,2
Our current understanding of mucociliary clearance in the
airways dates to important observations in the 1930s by Florey
et al. 3 who described the sources and transport of mucus in
cat tracheas, and by Lucas and Douglas 4 who recognized the
clear separation between the mucus layer and the ciliary activ-
ity underlying its transport over the nasal epithelia of several
mammals. More recently, histological preparations of airway
epithelia fixed to preserve the mucus layer showed two distinc-
tive gel-like layers, the soluble transporting mucus layer and
the periciliary layer (PCL; 5,6 ). The soluble mucus layer is rich
in the two major polymeric mucins, MUC5AC and MUC5B,
and hundreds of globular proteins. 7 – 10 Estimates of the mucus
layer thickness are variable, depending on choice of fixatives
and methods of measurement. Electron microscopy (EM) stud-
ies in different species measured the mucus layer as 7 – 10 ? m
thick, 5 whereas others report the layer to be 0.5 – 70 ? m in
The thickness or depth of the PCL has been easier to deter-
mine as the PCL extends over the length of fully extended cilia.
Dependent on airway region, cilia are between 5 and 8 ? m in
Molecular organization of the mucins and
glycocalyx underlying mucus transport over
mucosal surfaces of the airways
M Kesimer 1 , 2 , C Ehre 1 , KA Burns 1 , CW Davis 1 , 3 , JK Sheehan 1 , 2 and RJ Pickles 1 , 4
Mucus, with its burden of inspired particulates and pathogens, is cleared from mucosal surfaces of the airways by
cilia beating within the periciliary layer (PCL). The PCL is held to be “ watery ” and free of mucus by thixotropic-like
forces arising from beating cilia. With radii of gyration ~ 250 nm, however, polymeric mucins should reptate readily
into the PCL, so we assessed the glycocalyx for barrier functions. The PCL stained negative for MUC5AC and MUC5B,
but it was positive for keratan sulfate (KS), a glycosaminoglycan commonly associated with glycoconjugates.
Shotgun proteomics showed KS-rich fractions from mucus containing abundant tethered mucins, MUC1, MUC4, and
MUC16, but no proteoglycans. Immuno-histology by light and electron microscopy localized MUC1 to microvilli,
MUC4 and MUC20 to cilia, and MUC16 to goblet cells. Electron and atomic force microscopy revealed molecular
lengths of 190 – 1,500 nm for tethered mucins, and a finely textured glycocalyx matrix filling interciliary spaces.
Adenoviral particles were excluded from glycocalyx of the microvilli, whereas the smaller adenoassociated virus
penetrated, but were trapped within. Hence, tethered mucins organized as a space-filling glycocalyx function as
a selective barrier for the PCL, broadening their role in innate lung defense and offering new molecular targets for
conventional and gene therapies.
1 Cystic Fibrosis / Pulmonary Research and Treatment Center, University of North Carolina , Chapel Hill , North Carolina , USA . 2 Department of Biochemistry and Biophysics,
University of North Carolina , Chapel Hill , North Carolina , USA . 3 Department of Cell and Molecular Physiology, University of North Carolina , Chapel Hill , North Carolina ,
USA . 4 Department of Microbiology and Immunology, University of North Carolina , Chapel Hill , North Carolina , USA . Correspondence: M Kesimer ( mehmet_kesimer@
Received 13 June 2012; accepted 26 July 2012; published online 29 August 2012. doi:10.1038/mi.2012.81
VOLUME 6 NUMBER 2 | MARCH 2013 | www.nature.com/mi
length. The microenvironment of the PCL surrounding the cilia
has been described as “ watery ” , or low-viscosity, to enable cilia to
beat unimpeded. 4,5,13 – 16 For this reason, the PCL is often termed
the periciliary liquid layer.
A major biophysical question is why the soluble mucus layer
and the PCL are maintained as a two-layer system. Earlier work
suggested the intrusion of mucus into the PCL was prevented
by thixotropic-like forces provided by the beating cilia. 4,13,16
However, given the ~ 250 nm radius of gyration of polymeric
mucins 17,18 and the density of cilia on a ciliated airway epithe-
lial cell, it is unlikely that such molecules can be excluded from
interciliary spaces, which approach ? m distances, particularly
at intercellular borders ( Figure 1 ).
We now provide evidence for a glycocalyx more robust than
previously realized (see ref. 19) that results from a complex
organization of membrane-tethered, high MW glycoconju-
gates. Previously, we have shown that tethered mucins in the
airway glycocalyx can restrict virus penetration into the airway
epithelial surface. 20,21 This finding, plus the recent observa-
tion in vascular endothelium that tissue preparation appropri-
ate to the preservation of glycocalyx revealed it to be ~ 10 ? m
in thickness, rather than the negligible thickness observed by
conventional light and electron microscopies, 22 make a glyco-
calyx barrier to mucus penetration of the PCL a particularly
Spatial distribution of major mucins and
glycosaminoglycans in mucus
The major gel-forming or polymeric mucins of airway mucus
have been identified as MUC5AC and MUC5B. 23,24 Although
airway mucus secretions normally undergo unidirectional
transport in the airways in vivo , in human tracheobronchial
epithelial (HTBE) cultures mucus secretions are transported
within the culture dish but are not cleared. This property
of HTBE cell cultures was exploited to visualize the spatial
distribution of the mucins in mucus allowed to accumulate
on culture surfaces over several days. As formalin-based fixa-
tives can collapse mucus structures, we opted to use an alco-
hol-based fixative (Omnifix) to better preserve the mucus
secretions intact during histological processing and analy-
sis. 25 Probing histological sections of HTBE with MUC5AC
monoclonal and MUC5B polyclonal antibodies (see Table 1 )
revealed the presence of these mucins in extensive “ clouds ”
emanating from the epithelium but with minimal spatial
mingling ( Figure 2c ). Notably, immunodetectable MUC5AC
and MUC5B were absent from a broad region immediately
above the HTBE cell surface, which corresponded to the
airway surface region occupied by cilia ( Figure 2a,b ).
Although the mucosal surface of the epithelium is irregular
( Figure 2a ), the MUC5AC and MUC5B exclusion zone was
generally ~ 7 ? m high, consistent with the height of human
tracheobronchial cilia. We propose that this MUC5AC / B
exclusion zone represents the PCL.
Glycosaminoglycans are unbranched polysaccharide chains
species normally associated with glycoconjugates of connec-
tive tissue and glycocalyx. Heparan sulfate and keratan sulfate
(KS) glycosaminoglycans are classically associated with prote-
oglycans in connective tissue, but also associate with mucins
in mucus of the airways and elsewhere. 26 – 28 Immunoprobing
histological sections of HTBE and accumulated mucus secre-
tions failed to detect heparan sulfate at the mucosal surface
of HTBE cells, either in association with the apical surface or
mucus secretions (data not shown). Instead, heparan sulfate
was immunodetected in the basal cell layers of the HTBE cul-
tures as described previously. 29 By contrast, KS was detected
robustly on the apical surfaces of HTBE and in the mucus secre-
tions ( Figure 2d ). Notably, however, the spatial distribution of
KS was markedly different to that of MUC5AC and MUC5B:
the most intense KS staining was within the PCL, rather than
in the mucus layer, and it was particularly associated with the
apical surfaces of ciliated cells. This observation suggested
that KS more closely associated with glycoconjugates in close
proximity to the cilia or ciliated cell apical membranes. The
mucosal surfaces of ciliated cells have previously been associ-
ated with an acidic mucosubstance, 30 which we now propose
is KS, at least in part. Notably cilia also stained positive with
alcian blue / periodic acid-Schiff ( Figure 2b ), as well as with
the KS-specific antibody (see also Supplementary Figure S1A
online). Interestingly, many of the most intensely stained, KS-
rich areas had irregular interfaces with the overlying mucus
layers (arrows, Figure 2d ), which suggested the attachment of
KS-associated material to cilia, and / or its transfer from cili-
ated zones of the PCL into the mucus. Notably, these “ ciliary
plumes ” , are also present in the other sections of Figure 2 ,
as indicated by labeled arrows, whether by positive staining
( Figure 2a,b,d ), or by its absence ( Figure 2c ). Significantly, the
Figure 1 Density map of cilia on airway epithelial cells. Using a
published electron micrograph (Figure 3 in ref. 60) of guinea-pig tracheal
epithelial cells, individual cilia were mapped, manually, and are displayed
as white circles. Reverse contrast was used to emphasize interciliary
spaces. The micrograph represented a cross-sectional plane through
the basal region of the cilia, where microvilli were apparent between the
cilia, so the distribution approximates the positions of the basal bodies.
Red dotted lines indicate approximate cell borders. Note the magnitude
of interciliary distances, ~ 100 nm to > 1 ? m, especially between cilia
on adjacent cells. A full colour version of this figure is available at the
Mucosal Immunology journal online.
MucosalImmunology | VOLUME 6 NUMBER 2 | MARCH 2013
“ negative staining ” of with MUC5AC and MUC5B antibodies
in these areas indicate their exclusion from within the ciliary
Molecular identification of the KS-associated conjugate in
HTBE cell culture secretions
To identify the molecular species associated with KS immuno-
reactivity, we isolated KS-rich material from HTBE cell culture
washings solubilized in 4 M GuHCl and separated the mate-
rial by isopycnic density gradient centrifugation in CsCl / 4M
GuHCl. 31,32 The gradient was fractionated and the fractions
analyzed for density, MUC5AC, MUC5B, and KS immunore-
activity, and periodic acid-Schiff staining. An agarose western
blot probed with an antibody against MUC5B ( Figure 3a , top)
shows that the polymeric mucin resolved in the middle of the
density gradient (densities = 1.30 – 1.35 g ml − 1 ). MUC5AC had
a similar gradient distribution (data not shown). By contrast,
KS-positive materials resolved in fractions with the highest
densities ( Figure 3a , bottom, densities = 1.45 – 1.55 g ml − 1 ), and
with minimal apparent overlap with the MUC5AC / MUC5B-
Similar results were also obtained using an alternative sepa-
ration of HTBE cell culture secretions using a MonoQ 5 / 5 ion
exchange column eluted over a range of salt strengths, 0.1 – 1.0 m
LiClO 4 ( Figure 3b ). Fraction subsamples were blotted and
probed with antibodies specific to MUC5B, MUC5AC, MUC1,
MUC4, MUC16, and KS. The KS probe bound to molecules that
eluted in a range generally more acidic than MUC5B: the overlap
between the two elution profiles, though significant, was small
and the peak responses were well separated. On the other hand,
the distribution of MUC16 overlapped broadly with that of both
MUC5B and KS. The distribution of MUC5AC was very similar
to that for MUC5B (not shown). MUC1 had a bipolar distribu-
tion, one population of molecules more-or-less co-eluted with
MUC5B, the other with MUC16. A smaller amount of MUC4
eluted from the column, with a peak positioned between those
for MUC5B and MUC16. Hence, for fractions eluting at acidic
pHs there was significant overlap between the presence of the
tethered mucins (MUC 1, 4, and 16) and KS-conjugated mol-
ecules. Notably, approximately half of the KS-positive material
eluted in fractions more acidic than those containing tethered
mucins ( Figure 3b ), suggesting the presence of other, unidenti-
fied glycoconjugates, glycolipids, or other materials.
A two-dimensional shotgun proteomics-based approach
was used to identify the proteins conjugated to KS. The first
dimension was either the CsCl density gradient centrifuga-
tion or ion exchange chromatography separation described
above ( Figure 3 ). The KS-rich fractions (15 – 20) from each
separation were combined, reduced, alkylated, digested
with trypsin, and analyzed by ultra-performance liquid
chromatography / mass spectrometry as described previously. 7
MUC16 and MUC5B were identified as the predominate
proteins in the KS-rich material of both separations ( Table
2 ), using the total number of tryptic peptides identified per
Table 1 Summary of antibodies used for immunodetection
Target Antibody / staining Epitope / region Ref.
B27.29 (mouse IgG1)
214D4 (mouse IgG1)
Recognizes glycosylated N-terminal repeat (PDTRPAP)
Recognizes unknown epitope at the glycosylated repeat region
Recognizes a peptide (FLNSNSGLQGLQFYR) in un-glycosylated
region close to Cys-rich domain
Recognizes intracellular immature molecule
Recognizes non-glycosylated C-terminal peptide
C-terminal cystein-rich part
Recognizes the peptide (RNREQVGKFKMC) located in repeated
Recognizes peptide (ElGQVVECSLDFGLVCR) located in the
Recognizes protein domain in SEA repeat
Purified protein, Abcam (ab10033)
Recognizes the peptide (TSGTESTLISTSAPE)
Recognizes C-terminal peptide (LSVASPEDLTDPR)
IG8 (mouse IgG1)
45M1 (mouse IgG1)
EU-MUC5Ba (mouse IgG1)
MAN5B / LUM5B (rabbit) Ref. 8
OC125 (mouse IgG1)
X325 (mouse IgG1)
Ref. 35 and
Ref. 67 Keratan sulphate 5D4 (mouse IgG1) Recognizes N -acetyl lactosamine disaccharides of KS with
a pentasulfated hexasaccharide.
AB stains acidic (sulphated) mucins and proteoglycans, PAS stains
mucins and, glycoproteins, primarily their terminal sialic acid
moieties, but also other carbohydrates, including glycogen.
Carbohydrate AB / PAS (alcian blue / PAS)
Abbreviations: KS, keratan sulfate; PAS, periodic acid-Schiff.
VOLUME 6 NUMBER 2 | MARCH 2013 | www.nature.com/mi
protein as an approximate index of abundance. The other mucins
of interest, MUC5AC, MUC4, and MUC1 were also abundant
in the density gradient – separated KS-rich material.
A recently identified mucin, MUC20, which is expressed
in the lung, 33 – 35 was detected as a single peptide fragment
by mass spectrometry of the KS-rich fraction from the CsCl
Figure 2 Mucus, mucins, and keratan sulfate (KS) on the mucosal surface of human tracheobronchial epithelial (HTBE) cell cultures. Histological
sections of HBTE cell cultures with accumulated mucus secretions were stained for ( a ) hematoxylin and eosin (H & E) and ( b ) alcian blue / periodic acid-
Schiff (AB / PAS), or immunoprobed with antibodies against ( c ) MUC5AC (red, monoclonal Ab, 45M1) and MUC5B (green, polyclonal, Cys-rich domains
within the mucin repeat domains) or ( d ) keratan sulfate (green, monoclonal Ab 5D4). Panels c and d were counterstained with DAPI. Note height of the
accumulated mucus layer ~ 100 ? m, the intense staining of KS in the periciliary layer, and in all the panels, the plumes of material that appear to extend
from ciliary tips and into the mucus (ciliary plumes), while excluding the polymeric mucins. In panel c , intracellular mucins are not apparent, as their
fluorescence intensities were too weak to be imaged at gain settings appropriate for the highly intense extracellular signal. For panel a , bar = 20 ? m.
Figure 3 Separation of mucins and keratan sulfate (KS)-conjugated molecules collected in human tracheobronchial epithelial (HTBE) cell culture
secretions. Mucus collected from HTBE cell cultures and solubilized in GuHCl was separated using two different methods. ( a ) CsCl isopycnic density
gradient analysis. Fractions from a CsCl density gradient following isopynic centrifugation to equilibrium were analyzed by agarose western blotting
using antibody probes to MUC5B (top) and KS (bottom). ( b ) Ion exchange column chromatogram. Solubilized mucus was applied to a MonoQ HR 5 / 5
ion exchange column, after dialysis against urea and disulfide reduction in dithiothreitol. The column was eluted with an increasing gradient of LiClO 4
and fractions analyzed by slot blotting, using antibody probes to MUC5B, MUC16, and KS (shown), as well as MUC5AC, MUC1, and MUC4 (not
shown). The KS elution profile is highlighted in gray to illustrate that approximately half the KS-positive material elutes with MUC16 (and other mucins),
whereas the other half elutes at more acidic pHs.
MucosalImmunology | VOLUME 6 NUMBER 2 | MARCH 2013
density gradient. Like MUC5AC and MUC4, it appeared to
be absent from the ion exchange separation. A rabbit polyclo-
nal antibody was generated against a synthetic peptide, whose
sequence was taken from a portion of the non-glycosylated
region of the MUC20 molecule that was detected by mass spec-
trometry, and was used to probe an agarose western blot of the
fractions from the CsCl density gradient. This analysis showed
the antibody to stain a single major band that was concentrated
in fractions 15 – 19 of the gradient (see Supplementary Figure
S2 online). Hence, MUC20 has the high buoyant density typical
of mucins, one that is distinctly higher than the typical distribu-
tions of MUC5B and MUC5AC and within the range in which
KS distributes (c.f. Figure 3 ).
Several globular proteins were also identified in the KS-rich
material in both separations ( Table 2 ), but because none of them
are known to be highly glycosylated, they are likely complexed
with the mucins. The only non-mucin, glycoconjugate identified
was DMBT1; a highly N - and O -glycosylated protein known
to function in mucosal defense as an agglutinin binding to a
variety of proteins, including surfactant protein D, lactoferrin,
and MUC5B. 36
Visualizing mucin molecules in mucus from HTBE cell
Electron and atomic force microscopy were used to visualize
and evaluate the diversity of mucin macromolecules in mate-
rial eluting in the void volume of a Sepharose S-1000 column
( Figure 4a,c ). Visualization by EM revealed five different
dimensional classes of molecules, four of which were “ short ” ,
linear molecules 190 – 1,500 nm long, approximating the pre-
dicted sizes of tethered mucins ( Table 2 ). The largest of these
molecules (1,200 – 1,500 nm × 10 – 12 nm) had a distinct tail,
~ 130 nm in length, with a visual characteristic resembling a
“ string of beads ” ( Figure 4a , inset, arrows). This clearly observ-
able structure may represent the SEA-repeat domain region
of MUC16, 16 extracellular SEA repeats positioned between
the transmembrane domain and the tandem, mucin repeat
domains. 37 – 39 The extracellular portion of MUC16 is predicted
to be at least 22,000 residues in length and if fully extended
would make it ~ 8.8 ? m long assuming 4 Å (0.4 nm) / residue. 40
Hence, a measured contour length of 1 – 2 ? m seems reasonable
for the molecule detected by EM, as extensive O -glycosylation
of the extracellular domains combined with a proline-rich con-
tent will render the molecule relatively inflexible 41 and regu-
larly kinked, respectively. The identification of this molecule
was confirmed as MUC16 using the well-characterized MUC16
antibody, OC125, which recognizes the SEA domains of MUC16
selectively. 38 Our visual structural analysis was corroborated
using this antibody as immunogold bound exclusively to the end
the molecule we predicted represented the SEA-repeat domain
of MUC16 ( Figure 4b , arrow).
The second largest class of molecules had similar 10 – 12 nm
widths as determined for MUC16, but were shorter (750 –
900 nm) and lacked visual evidence of the SEA-repeat domain.
Based on the similarity of molecular widths, we tentatively
identify these molecules also as MUC16, though whether they
represent a fragment of full-length MUC16 or VNTR (vari-
able nucleotide tandem repeat) polymorphic isoforms can-
not be determined. The other two classes of linear molecules
identified by EM analysis ( Figure 4a,c,c ? ) were 7 – 9 nm wide,
and were either 750 – 900 or 190 – 200 nm in length ( Table 3 ).
We tentatively assign these molecules as MUC4 and MUC1,
respectively, based on their predicted lengths relative to
The final class of molecule observed ( Figure 4c ) was iden-
tified as the polymeric mucin, MUC5B, based on its visual
characteristics 18 and the dominant abundance of MUC5B in
HTBE mucus ( ~ 80 % of the mucins are MUC5B). 9 Compared
with the size of tethered mucins, MUC5B was immense and the
structure imaged likely represents a single continuous molecule
(indicated by the tracing in Figure 4c ? ). The predicted MUC5B
molecule exhibited a convoluted contour many micrometers
in length and features multiple, crosslinked nodes. Previous
studies indicate this molecular form of MUC5B is intermediate
in form between a compact, highly crosslinked form resolving
at the bottom of the sucrose density gradient, and the linear
form resolving at the top. 18 For comparison, an EM of the
fully linearized form of MUC5B is shown in Supplementary
Figure S5 online.
Visualization of the glycopeptides representing the
gigantic, extracellular glycosylated domains of the MUC16 and
MUC4-tethered mucins by atomic force microscopy (AFM)
after dispersion on mica revealed the molecules to have similar
contour lengths to those measured by EM, but their measured
widths are much greater ( Figure 4d,e ). Molecules identified
Table 2 Major proteins in KS-rich fractions from HTBE
secretions, identified by two dimensional shotgun
Basis of fi rst separation
MonoQ ion-exchange HPLC
(pooled fractions 17 – 19)
CsCl density gradient
(pooled fractions 17 – 20)
WAP domain containing
Complement component C3
WAP domain containing
Prostate stem cell antigen
Abbreviations: HTBE, human tracheobronchial epithelial; HPLC, high-powered
liquid chromatography; KS, keratan sulfate.
The proteins are ranked by the total number of peptides identifi ed for each
protein, with higher numbers being consistent with greater abundance.
VOLUME 6 NUMBER 2 | MARCH 2013 | www.nature.com/mi
as MUC16 or MUC4 had widths by AFM of 50 – 60 and 30 –
35 nm, respectively, or approximately fivefold wider than their
appearance by EM (c.f. Table 3 ). This difference is presum-
ably due to better maintenance of the hydration state of the
mucins visualized by AFM — molecules visualized by
EM are subjected to strong dehydration forces imposed by
the alcohol washings and the vacuum required for rotary
Localization of tethered mucins in airway epithelia
Based on the identification of KS-associated high MW glyco-
conjugates on the surface of HTBE cultures as tethered mucins,
we probed histological sections of HTBE and human bron-
chial epithelia with antibodies to specific mucins to determine
their subcellular localization. Probing sections for an extra-
cellular epitope of MUC1 (B27.29 mAb; 42 ) showed that
the mucin localized extensively within the PCL at the base
of the cilia and to basal epithelial cells ( Figure 5a , left and
right panels). At this resolution, MUC1 appeared absent from
upper regions of the cilium, if not from the entire ciliary
By contrast, probing sections with a polyclonal antibody
raised to an extracellular peptide of MUC4 (MUCH4), whose
sequence was identified initially by mass spectrometry, local-
ized MUC4 immunoreactivity to the ciliary shafts ( Figure 5b ).
In HTBE cultures with accumulated mucus secretions, MUC4
was also detected in the ciliary plumes ( Figure 5b , left panel).
An alternative MUC4 monoclonal antibody, Ig8, revealed
robust expression of MUC4 in ciliated cells (see Supplementary
Figure S1B online). This antibody failed to detect extracellu-
lar MUC4 as the antibody recognizes a peptide sequence in
the VNTR mucin domain of MUC4, a heavily glycosylated
region of MUC4 likely to sterically hinder antibody access
to the peptide epitope. Further analysis immunodetecting
MUC1 and MUC4 together in the same preparation showed
spatial co-localization in the cell surface regions correspond-
ing to the microvilli of ciliated cells at the base of the cilia
Table 3 Dimensional classes of linear mucins in mucus from
primary HTBE cell cultures observed by electron microscopy
Contour length × width (nm) Identifi cation
1,200 – 1,500 × 10 – 12
750 – 900 × 10 – 12
750 – 900 × 7 – 9
190 – 200 × 7 – 8
Abbreviations: HTBE, human tracheobronchial epithelial.
a Association tentative.
Figure 4 Electron microscopic (EM) and atomic force microscopic (AFM) images of mucins from human tracheobronchial epithelial (HTBE) mucus.
Mucins from HTBE cell culture mucus secretions solubilized in phosphate-buffered saline were separated from most of the protein fraction by taking
the void volume of a Sephacryl S1000 HPLC (high-powered liquid chromatography) column, and were then fixed in glutaraldehyde and prepared for
EM. ( a ) EM of tethered mucins. MUC16 is identified by the tail at one end of the molecule appearing as a “ string of beads ” (inset), and MUC4 by relative
length and narrower profile. ( b ) EM of MUC16 labeled with OC125-bound gold bead (arrow). See Experimental Procedures for preparation details.
( d and e ) AFMs of MUC16 and MUC4. After destructive proteolysis of reduced, polymeric mucins in HTBE mucus, the remaining, large glycosylated
extracellular domains of MUC16 and MUC4, which are resistant to proteolysis, were dispersed on mica and visualized by AFM. ( c and c ? ) EM of
tethered and polymeric mucins. Polymeric mucins in samples of HTBE mucins prepared as in panel a were imaged by EM. The mucin profiles in c ?
were traced with color-coded lines, as indicated. ? = tentative identification.
MucosalImmunology | VOLUME 6 NUMBER 2 | MARCH 2013
(see Supplementary Figure S2C online). However, within the
limits of resolution of light microscopy, it is possible that this
apparent overlap might arise from an intercalation of mucin
molecules within the glycocalyx, but with different points of
attachment, e.g., microvilli and cilia.
To clarify the spatial localization of MUC1 and MUC4,
immuno-EM was used to identify the ultrastructural localiza-
tion of MUC1 and MUC4 immunoreactivity. Figure 5e,f show
immunolocalization of MUC1 (214D4 mAb conjugated to 18 nm
gold beads) in glycocalyx structures associated with micro-
villi, while MUC4 (MUCH4 antibody conjugated to 6 nm gold
beads) immunolocalized to glycocalyx associated with cilia.
In 11 different electron micrographs, we enumerated 366 MUC1-
conjugated and 129 MUC4-conjugated beads, of which 93.7
and 90.7 % were associated with the microvilli or cilia, respec-
tively. Approximately ~ 6 % of beads were attached to cell surface
elements not identified. Together, these immunolocalization
data indicate a differential distribution of MUC1- and MUC4-
tethered mucins on ciliated cells, with MUC1 preferentially
present on microvilli and MUC4 preferentially on ciliary
The cellular distribution of MUC1 and MUC4 in airway epi-
thelium also differed. Although MUC1 and MUC4 localized
to basal epithelial cells, MUC4 was more robustly expressed
than MUC1 in ciliated cells ( Figure 5a,b , Supplementary
Figures S1B,C, S3A,B online). However, in HTBE cell cul-
tures with accumulated mucus, MUC1 and MUC4 were both
detected within the mucus secretions ( Figure 5a,b ). Although
MUC1 has been reported to be secreted / shed from airway
epithelium, 39 we recently described the presence of MUC1 and
MUC4 on exosome-like vesicles shed from the airway surface
into the mucus secretions, thus contributing to the presence
of these tethered mucins in accumulated mucus secretions. 43
Discerning whether MUC1 and MUC4 immunoreactivity
Figure 5 Localization of tethered mucins in airway epithelia. Panels a – d show staining by immunofluorescence (IF) for MUC1, MUC4, MUC16,
and MUC20, as indicated. The left and right hand image of each pair within a panel shows immunolocalization in a human tracheobronchial epithelial
(HTBE) cell culture and in human bronchial epithelium, respectively. The HTBE cultures in panels a – c had accumulated mucus, per Figure 2 . For
panel b , note that the antibodies used in the two images were to the same peptide sequence (MUCH4), but represented different polyclonal antibodies,
one of which was suitable for immunohistochemistry, the other for IF. Bars = 10 ? m. Panels e and f show MUC1 and MUC4 localization of human
bronchial epithelium by immuno-electron microscopy (EM). The tissue was exposed simultaneously to MUC1 and MUC4 antibodies conjugated to
18 and 6 nm gold beads, respectively, then washed extensively, fixed, and prepared for conventional EM. Arrows indicate gold beads labeling the
glycocalyx of microvilli (MV) and cilia.
VOLUME 6 NUMBER 2 | MARCH 2013 | www.nature.com/mi
in mucus secretions represents shed MUC1 / 4-containing
vesicles, shed mucin molecules per se or both is beyond the
resolution of our current fluorescent microscopy studies.
MUC16, identified by both OC125 and X325 antibodies, was
predominately expressed by goblet cells in the surface epithe-
lium ( Figure 5c and Supplementary Figures S3 and S4 online).
The OC125 antibody robustly detected the apical membrane
of goblet cells, but stained cytoplasm of these cells only lightly.
The X325 antibody, which recognizes an extracellular epitope
distinct from the OC125 epitope, 44 stained cytoplasmic secre-
tory granules to a greater degree than OC125. Interestingly,
in HTBE cell cultures with accumulated mucus secretions,
MUC16 capped goblet cells, which co-localized with MUC5B
immunodetection. Indeed, in addition to capping MUC5B-
containing / secreting cells, MUC16 also emanated from the
goblet cells in broad pillars of immunoreactivity, suggesting
co-release of MUC16 with MUC5B ( Figure 5c , left). MUC16
was not immunodetected at the level of the cilia or ciliated
cells and may be excluded from these domains as suggested for
MUC5AC and MUC5B exclusion.
Using the antibody generated against MUC20 as described
above (see Supplementary Figure S2 online), we locali-
zed MUC20 immunoreactivity to the cytoplasm of ciliated
and basal epithelial cells and to the cilia ( Figure 5d and
Supplementary Figure S3D online). As MUC20 was iden-
tified by proteomics in mucus secretions of HTBE cultures,
it is likely MUC20 is secreted / shed from the epithelium into
Tethered mucins were also detected in the submucosal
glands of human tracheobronchial airway tissue. MUC1
was immunolocalized to a sub-population of acinar cells
(see Supplementary Figure S1D online), and MUC16
immuno reactivity was associated with the surface of
mucous acinar cells containing MUC5B (see Supplementary
Figure S1E online), similar to the association observed in
the surface epithelium of HTBE cell culture ( Figure 5c ).
MUC20 immunoreactivity was localized in all acinar cells (see
Supplementary Figure S1F online). MUC4 was not detectable
in submucosal glands.
Visualization of the glycocalyx
The classical view of the mucus and PCLs on airway surfaces is
that both layers are distinct and thixotropic-type forces gener-
ated by the beating cilia prevents penetration of secreted mucins
into the PCL. 13 This perception of a “ watery ” PCL to enable
effective ciliary beating was formed by the lack of visible mate-
rials within the periciliary spaces. Conventional histochemical
and EM techniques to visualize glycocalyx structure and com-
ponents such as the metallic stains colloidal iron and ruthenium
red likely collapse the molecular structure and network of the
glycocalyx via electrostatic interactions. 20,22,45 These artifacts of
tissue preservation and visualization have led to the perception
that the airway glycocalyx is a layer at the cell surface, distinct
from the PCL (e.g., ref. 46). Our previous 20 and current studies,
however, indicate a rich population of tethered mucins popu-
lating periciliary surfaces that could well serve as components
of a selective filter, which excludes large molecules, molecular
complexes, and microbes.
To test this notion, we examined the ultrastructure of the
airway surface using conventional and rapid freeze / freeze sub-
stitution (RF / FS) techniques. Figure 6a shows the ultrastruc-
ture of the mucosal surface of human bronchial epithelium
after staining with ruthenium red and conventional processing
for EM. Notably, the glycocalyx is seen as a thin, fuzzy layer
on the surfaces of the microvilli and is hardly visible on cilia
shafts. Although strands of material appear to project between
microvilli, the overall periciliary microenvironment is seen as
open space using this technique. In mouse tracheal airways
ex vivo , ruthenium red staining revealed a much more robust
and dense glycocalyx (see Supplementary Figure S6 online),
but even in this tissue, “ open space ” predominates over evi-
dence of a complex molecular matrix. Using histological tech-
niques known to better preserve native carbohydrate structure,
i.e., RS / FS, glycocalyx structure is revealed as a complex, finely
textured matrix filling the available space within the PCL
( Figure 6b,c ). In Figure 6b , the glycocalyx matrix visually
fills spaces between cilia shafts, and Figure 6c shows the gly-
cocalyx of a goblet cell, with cilia projected above filled with
a fine grained, fibrillar matrix material — notably the matrix
occupies spaces > 1 ? m. A magnified view of Figure 6c (lower
panel), shows matrix fibrils radiating from microvilli cut
in cross-section, and in both panels where the structures are
cut longitudinally, the fibrils form a lattice with an irregular,
Our previous and current studies identify an underappreci-
ated, rich population of tethered, highly glycosylated mucins
within the PCL populating the epithelial cell surface glyco-
calyx. These structurally defined and diverse molecules are
likely important in innate protection of the airways surfaces.
They likely influence appropriate airway surface hydration
for effective mucus clearance as well as functioning as an
innate immunological molecular sieve limiting the access of
molecules and molecular complexes, including pathogenic
particulates and microbes 20 to the epithelial cells and tissues
underlying the glycocalyx. In an initial attempt to test this
notion, we used several human viruses as molecular probes
of particles with defined size and structure to determine the
penetration of these organisms through the glycocalyx layer.
Notably, the sizes of the viral particles used, 30 and 100 nm,
are less than the typical, ~ 250 nm radius of gyration for
polymeric mucins. Figure 7 shows reasonable penetra-
tion of adenoasociated virus (30 nm) into the glycocalyx of
ciliated cell microvilli, whereas the larger adenovirus particle
(100 nm) were excluded. It is likely the glycocalyx and the
heavily glycosylated glycoconjugates contained within act to
restrict particle access based on size, as well as biochemical
structure and interactions.
We provide evidence of the spatial molecular structure and
distribution of soluble mucins (mucus) and tethered mucins
(PCL) on the mucosal surface of human airway epithelium
MucosalImmunology | VOLUME 6 NUMBER 2 | MARCH 2013
that is consistent with early descriptions of the two separate gel
domains of the mucociliary clearance system. 4 In our studies
of well-characterized HTBE cultures, we show that MUC5AC
and MUC5B were clearly excluded from the PCL ( Figure 2c ),
despite polymeric mucins being of sufficiently small molecular
size to enter the 0.1 – 1 ? m-sized spaces around and between
the cilia ( Figure 1 ). These inter-ciliary spaces, however, are
not devoid of glycosylated material as KS immunoreactivity
in the PCL was robust ( Figure 2 ). KS is a glycosaminoglycan
commonly associated with glycoconjugates; hence, KS local-
ized in the PCL associated with cilia and ciliary plumes sug-
gested that KS is a major component of a significant glycocalyx
structure within the PCL. Historically, the microvilli on the
apical surfaces of ciliated cells have been shown to be coated in
acidic mucosubstances, most likely highly sulfated glycocon-
jugates. 30 In connective tissues such as cartilage, KS is asso-
ciated with proteoglycans, e.g., aggrecan, but in recent years
KS has also been found conjugated to mucins, 26,27 including
those on the mucosal surfaces of human airways. 28 The airway
mucins conjugated to KS have not been identified. Examination
of the separation profiles of airway mucus analyzed by both
CsCl isopycnic density centrifugation and ion exchange HPLC
(high-powered liquid chromatography) showed that KS immu-
noreactivity overlapped significantly with the airway soluble
mucins, MUCs 5AC,and MUC5B, and the tethered mucins,
MUC1, MUC4, and MUC16 ( Figure 3 ). These biochemical
associations were confirmed by mass spectroscopy. The most
abundant proteins identified in KS-rich fractions ranked by
peptide coverage were MUC16, MUC5B, MUC1, and MUC4
( Table 2 ). MUC20 was less abundant but was also identified
in the KS-rich material. No proteoglycans were identified,
consistent with our earlier findings. 7 All attempts to immu-
noprecipitate KS immunoreactive materials were hindered by
non-specific binding of the KS-rich materials to immunopre-
cipitation beads. This finding was reminiscent of the known
interactions of mucins with beads, with mucin molecules form-
ing strands and bundles around beads. 47,48 Together these data
indicate that both the polymeric and tethered mucins identified
Figure 6 Comparison of fixation methods on mucosal glycocalyx in human bronchial epithelium. Bottom panels are magnified portions of the images
above, as indicated by the white boxes. ( a ) Tissue stained with ruthenium red and processed for conventional electron microscopy. Note how ruthenium
red causes the glycocalyx to appear as thick, sparse strands, leaving most of the intercellular space apparently open. ( b and c ) Tissue fixed by rapid
freezing and freeze substitution (RF / FS), showing areas of the periciliary layer above ciliated and goblet cells, as indicated. Note the finely textured,
space filling matrix revealed by RF / FS. Lower panels, black arrows = microvilli, white arrows = cilia. See text for more details.
Figure 7 Interaction of viral particles with the glycocalyx of the
periciliary layer. Human tracheobronchial epithelial cell cultures were
incubated mucosally with particles of unlabeled adenovirus (Ad, 100 nm,
blue arrows) and adenoassociated virus (AAV, 30 nm, red arrows), then
rinsed, fixed, and prepared for electron microscopy. White arrow indicates
a cilium; black arrow, a microvillus.
VOLUME 6 NUMBER 2 | MARCH 2013 | www.nature.com/mi
in the KS-rich fractions of airway mucus were conjugated to
KS and MUCs 1, 4, 16, and 20 are largely responsible for the
robust KS immunoreactivity within the PCL ( Figure 2 and
Supplementary Figure S1 online).
Immunolocalization studies focused on reagents directed
to the extracellular, non-glycosylated domains of the tethered
mucins revealed a strong spatial organization of these molecules
within the PCL ( Figure 5 and Supplementary Figures S1, S3,
and S4 online). MUC1 immunoreactivity was localized pre-
dominately to microvilli of ciliated cells, while MUC4, and to a
lesser extent MUC20, localized predominately to the cilia shafts.
By contrast and as reported originally, 49 MUC16 was predomi-
nately localized to the apical membrane and secretory granules
of goblet cells (and mucous cells of submucosal glands). The
tethered mucins were also present in the mucus layer, presum-
ably after being shed from the cell surface ( Figure 5a – c ). In
the mucus layer, MUC1 immunoreactivity was associated with
vesicular structures, likely the exosome-like vesicles we have
previously shown to be rich in MUC1. 43 MUC4 and MUC16
immunoreactivity were also present in the mucus layer, associat-
ing with MUC5AC and MUC5B.
The ciliary plumes observed in mucus secretions in HTBE
cell cultures ( Figures 2 and 5 ) have not been previously
described. Plumes stain with alcian blue / periodic acid-Schiff
and hematoxylin and eosin, and are immunoreactive to KS
and MUC4 antibodies. They have a filamentous appearance,
excluding MUC5AC and MUC5B mucins as they project into
the overlying mucus layer from attachment points on ciliary
tips ( Figures 2 and 5 ). In a recent study, we observed tran-
sient, physical interactions between ciliary tips and polymeric
mucins, interactions that appeared to be important to mucus
transport. 48 We speculate that the ciliary plumes seen in accu-
mulated mucus, in vitro, are important to the cilia-mucin inter-
actions in mucus transport under more physiologically relevant
conditions, in vivo .
These studies highlight the importance of ultrastructural
preservation of glycocalyx for assessing components of the
PCL. Although the glycocalyx layer has been appreciated for
many years, the abundance and barrier function of this layer
in airways has largely been underappreciated in conventional
EM studies that rely on the use of metal-based dyes such as
colloidal iron and ruthenium red. Visualization of the gly-
cocalyx by these techniques rely on rendering the glycocalyx
electron dense, but these reagents also collapse the molecular
architecture of the glycocalyx structure electrostatically (see
ref. 50). As demonstrated dramatically for the glycocalyx of
vascular endothelium, when the specimen is prepared instead
by RF / FS, the glycocalyx is revealed to be much more complex
than when visualized by conventional EM, and in this par-
ticular case, it is around 10 – 11 ? m in thickness. 22 Previously,
RF / FS EM techniques have revealed that the glycocalyx of the
airway epithelium in HTBE and ex vivo human tracheal epithe-
lium is a complex mesh-like glycocalyx restricting the access
of adenoviral particles to the apical membrane of columnar
airway epithelium; 20 our new data suggest that the glycoca-
lyx may also prevent the penetration of mucus into the PCL.
Figure 6b,c show RF / FS EMs of HTBE when combined with
high-contrast digital photography reveals an even denser gly-
cocalyx than earlier described: the meshwork of the glycoca-
lyx, with interstices of < 100 nm, clearly fills the interciliary
spaces in the PCL over distances measuring in the ? m range.
Even by these EM techniques, one assumes that this mesh-like
ultrastructure would tend to exclude the polymeric mucins,
which have radii of gyration > 200 nm. 17,18 Consistent with
this possibility, a recent study by colleagues, Button et al. 51 ,
found that within the PCL labeled dextrans and other probes
of defined sizes were excluded progressively from the cell
surface of ciliated cells in HTBE cell cultures: molecules with
radii of gyration > 40 nm were excluded from the PCL, and
those > 18 nm were unable to penetrate through the microvilli
to the cell surface, whereas probes < 5 nm penetrated all the
way to the cell surface.
In addition to biophysical exclusion from the PCL, another
factor contributing to the separation of PCL and mucus lay-
ers may be the nature of the mucus gel, per se . The polymeric
mucins are known to interact with >100 other proteins, many
of which have known functions in innate defense, and ~ 30 %
of them bind directly to the mucins (refs. 7,10; and unpub-
lished observations). The close association of these proteins
to the polymeric mucins would likely limit their diffusion and
while their specific functions remain to be elucidated, we note
that they are potentially responsible, in part, for the transient
crosslinks between mucin molecules that impart elasticity to
the gel. 52 These interactions would likely also restrain mobil-
ity of mucin molecules, potentially constraining them to the
An important question for the role of tethered mucins in
imparting the mesh-like properties of the glycocalyx in the
PCL is related to whether tethered mucins identified in this
study are large enough to occupy the spatial environment of
the interciliary spaces. The tethered mucins, MUC1, MUC4,
and MUC16, are molecularly massive in relation to globular
proteins, with molecular masses ranging from several hundred
kDa to 2.5 MDa. Most of the mass of mucins is attributed to the
high density of N - and O -linked glycans attached to the peptide
backbone. Tethered mucins are predicted to exhibit linear and
rigid structures resembling “ bottle-brushes ” , highlighting the
extent to which these molecules may project into extracellular
spaces. The extracellular domains of MUCs 1, 4, and 16 range
from approximately 1,500 to 22,000 residues in length with
genetic variation in the number of tandem repeats dictating the
absolute length (see refs. 39,53). In contrast to MUCs 1, 4, and
16, the external domain of MUC20 is substantially shorter, with
only 425 residues extending into the extracellular space. 33 Using
the persistence length of 4 Å (0.4 nm) / residue for a maximally
extended peptide, 40 the maximum possible lengths calculated
for the tethered mucins are MUC20 — 170 nm, MUC1 — 600 nm,
MUC4 — 3,380 nm, and MUC16 — 8,800 nm. Although the
actual lengths will likely be shorter, considering the non-
glycosylated and glycosylated domains will possess globular
and linear, rigid structures, respectively, the contour lengths of
tethered mucins isolated from HTBE cell mucus ranged from
MucosalImmunology | VOLUME 6 NUMBER 2 | MARCH 2013
190 to 1,500 nm, within which there were four recognizable
size categories assignable to MUCs 1, 4, and 16 ( Figure 4 ,
Table 3 ). Using EM analysis, the tethered mucins were 7 – 12 nm
in breadth, but MUC4 and MUC16 when visualized in the more
hydrated state allowed by AFM measured approximately 30 and
55 nm wide ( Figure 4 ). Hence, the tethered mucins repre-
sent very large molecules fully capable of forming the dense
meshwork of the glycocalyx in the interciliary spaces of the
PCL ( Figure 6 ).
Although our studies have revealed MUCs 1, 4, 16, and 20 to
be the predominant tethered mucins on the airway surface, other
tethered mucins have also been identified at the mRNA level
in human and mouse airway epithelia, e.g., MUC13, MUC15,
and, MUC3. 20,34,54,55 Currently, reagents to detect the mucins
encoded by these genes in human tissues are not available but
these mucins could significantly contribute to the glycocalyx
mesh within the PCL. Other glyconjugates, such as hyaluronan, 28
as well as unidentified glycoproteins or glycoconjugates are likely
to add to the complexity of the glycocalyx mesh structure.
The intriguing findings that the PCL does not represent a
“ watery ” aqueous layer but rather a highly dense gel-like mesh-
work, it may be expected that the tethered mucins in the PCL
impart a viscous drag which interferes with ciliary beating and
effective mucus transport. However, studies investigating the
physics of polymers attached to surfaces have shown that grafted
polyelectrolyte brushes attached to surfaces offer remarkably low
frictional interactions between opposing interfaces. 56,57 Hence,
the tethered mucins of the glycocalyx, as grafted polyelectro-
lyte brushes, may not only pose a barrier to the penetration of
the PCL, but may also confer a lubricating property to facilitate
ciliary beating to ensure effective mucus transport.
In conclusion, the polymeric and tethered mucins on the
airway surface are organized into two distinct layers, with the
polymeric “ transporting ” mucins apparently excluded from the
PCL by the tethered mucins forming the glycocalyx that deco-
rates the cilia, microvilli, and apical surfaces of the epithelial
cells. Figure 8 summarizes these findings, showing the muco-
ciliary clearance system diagrammatically with a separation of
the layers and the locations / sources of the mucin molecules. The
polymeric mucins, MUC5AC and MUC5B, predominate in the
mucus layer, while membrane-bound mucins, MUCs 1, 4, 16,
and 20 also contribute to the mucus layer after shedding from
the cell surfaces. Although MUC1 is associated primarily with
exosome-like vesicles within the mucus layer, MUC4 localizes
primarily to ciliary plumes, and MUCs 4, 16, and 20 are dis-
persed within the mucus layer. The PCL is composed mainly
of large membrane mucins, MUC4 and MUC16, with MUC4
tethered predominately to cilia and MUC16 to the apical mem-
brane of goblet cells. MUC1, tethered to microvilli, serves to
strengthen the protective matrix at the base of cilia. We speculate
that the glycocalyx matrix in its entirety dominates the physical
properties of the PCL. The matrix (i) provides dynamic support
for the over-lying mucus gel, (ii) acts as a barrier to the penetra-
tion of particulates, pathogens, and polymeric mucins, and (iii)
facilitates ciliary beating by lubricating the interface between
neighboring cilia and possibly acting as molecular springs,
which, alternately compressing and relaxing, transiently store
and release energy during the ciliary beat cycle. The multilayer
protection model proposed at the airway lumen modifies our
view of the structure, organization, and role of these layers as
well as increasing the significance of the glycocalyx mesh in
maintaining mucus homeostasis, mucus transport, and airway
innate defense. As such these studies represent a paradigm shift
by which the mucins participate directly in innate lung defense
through a highly ordered molecular structure. Strategies to
strengthen this barrier against potential pathogens or enhance
penetration of potential therapeutics may identify new tech-
nologies to treat airway diseases.
Human tracheobronchial epithelium was obtained from airways resected
from normal donor tissue from the University of North Carolina (UNC)
lung transplant program or from NDRI (National disease research inter-
change) under UNC Institutional Review Board-approved protocols.
Mouse tracheal epithelium was obtained by tissue harvest, under protocols
approved by the UNC Institutional Animal Use and Care Committee.
Cell culture and collection of mucus . HTBE cells were isolated by
the UNC Cystic Fibrosis Center Tissue Culture Core and expanded
Figure 8 Schematic depicting a scaled view of the mucosal surface
of human airways. Top: overview of the mucociliary apparatus, showing
a ciliated airway mucosal surface covered with a sheet of mucus. For
clarity, the mucus sheet is depicted as a thin layer; in vivo , it would be
substantially thicker. Bottom: higher magnification views of a goblet cell
(left), cilia, and the periciliary layer, showing the molecular organization of
the different mucins.
VOLUME 6 NUMBER 2 | MARCH 2013 | www.nature.com/mi
on plastic to generate passage 1 primary cells, which were plated at a
density of 600k cells per well on permeable Transwell-Col (T-Col,
24 mm diameter) supports. 58 Once polarized, cells were cultured at an
air – liquid interface for 4 – 6 weeks to form well-differentiated cultures
resembling in vivo pseudostratified mucociliary epithelium. 58,59 Mucus
secretions were obtained from mature HTBE cell cultures by incubating
1 ml of phosphate-buffered saline on the apical surface of the cultures
for 30 min at 37 ° C and then harvesting the phosphate-buffered saline
with a large caliber pipette. Harvest was repeated twice to obtain 2 ml
of phosphate-buffered saline – diluted mucus per culture and then solu-
bilized with solid GuHCl to make a 4 M solution. The mucus samples
were centrifuged at 300 × g for 10 min to remove debris and pooled with
other samples collected under identical conditions.
Histology: immunohistochemistry and immunofluorescence . HTBE
cultures were prepared for examination by light microscopy using two
methods. The first method was designed to preserve accumulated
mucus on the culture mucosal surfaces. Cultures were maintained for
5 – 7 days without apical surface washing to allow mucus to accumulate.
These cultures were then fixed in an alcohol-based fixative (Omnifix,
FR Chemical, Mt. Vernon, NY) to preserve the thick mucus layer. 25 The
second method fixed cultures in 4 % paraformaldehyde to enable conven-
tional histochemistry. In both methods, paraffin-embedded histological
sections were prepared and immunostained as previously described 27
using monoclonal mouse IgG antibodies specific against MUC1 (B27.29;
115D8, kind gifts from Fujirebio Diagnostics, Malvern, PA), MUC5B
(EU5B, 8 MUC16 (OC125, X325), and keratan sulfate (5D4, Seikagaku,
Tokyo, Japan), followed by goat anti-mouse AlexaFluor 594 (Invitrogen,
Grand Island, NY), or polyclonal antibodies against MUC4 (MUCH4,
peptide sequence: FLNSNSGLQGLQFYR), MUC5AC (LUM5-1), 8 and
MUC20 (MUCH20, peptide sequence: LSVASPEDLTDPR), followed
by goat anti-rabbit AlexaFluor 594 (Invitrogen). Images were taken with
Leica SP2 Laser Scanning Confocal, or DM IRB conventional widefield
microscopes, and processed with Adobe Photoshop CS2. The antibodies
used are summarized in Table 1 .
RF / FS and ruthenium red protocols . RF / FS was performed as described
previously. 20 Briefly, samples of human airway epithelium dissected from
the same patient source used for the generation of HTBE cell cultures
were immersed in ice-cold 0.2 M sucrose solution (100 ml of 0.2 M
Sorenson ’ s buffer, 100 ml of distilled water, and 13.6 g of sucrose) for 1 h
at 4 ° C, then in ice-cold 25 % glycerol solution (3.8 ml of 0.2 M Sorenson ’ s
buffer, 3.8 ml of distilled water, and 2.5 ml of glycerol) for no longer than
1 h, and then plunge-frozen in Freon, and stored in liquid N 2 . The sam-
ples were transferred to cooled 4 % osmium tetroxide in acetone, stored
at – 80 ° C for 4 days, and then transferred to a – 20 ° C freezer for 2 h, then
a 4 ° C refrigerator for 2 h, then held at room temperature for 1 h. After
an acetone wash, the samples were infiltrated with propylene oxide and
embedded in Epon resin for standard transmission electron microscopic
For the visualization of the glycocalyx on the cilia by conventional
EM, samples were immersed in a glutaraldehyde solution (5 ml of 4 %
glutaraldehyde, 5 ml of 0.2 M cacodylate buffer, and 1,500 p.p.m. of ruthe-
nium red) for 1 h at room temperature. After rinsing in 0.2 M cacodylate
buffer, the tissues were immersed in an osmium tetroxide solution (5 ml
of 5 % osmium tetroxide, 5 ml of 0.2 M cacodylate buffer, and 5 ml with
1,500 p.p.m. of ruthenium red) for 3 h at room temperature. After a rinse
in 0.2 M cacodylate buffer, standard processing for transmission EM was
CsCl / 4 M GuHCl density-gradient centrifugation . CsCl was added to
collected mucus in 4 M GuHCl to a density of 1.45 g ml − 1 , and isopycnic
density-gradient centrifugation was performed for 60 h at 36,000 r.p.m.
on Beckman L8-M ultracentrifuge using 50.2TI, 12 × 40 ml rotor. Samples
were unloaded as 2 ml fractions from the top and aliquots analyzed for
absorbance at 280 nm (total protein), density, and, after transfer to a
nitrocellulose membrane by slot blotting, reactivity with periodic acid-
Schiff and antibodies to MUC5B and KS.
Characterization of mucin species by HPLC ion-exchange chroma-
tography . Mucus samples in 4 M GuHCl were dialyzed into urea reduc-
tion buffer (6 M urea containing 0.1 M Tris / 5 m M EDTA, pH 8.0) and
treated with 10 m M dithiothreitol for 2 h at 37 ° C. Iodoacetamide was
added to a final concentration of 25 m M , and the mixture was left in the
dark for 30 min at room temperature. The reduced samples were chro-
matographed on an Amersham Biosciences (Piscataway, NJ) MonoQ
HR 5 / 5 column, using Amersham Biosciences, Ettan LC high-pressure
chromatography system. The column was eluted with a linear gradient
of 0 – 0.75 M lithium perchlorate / 10 m M piperazine in 6 M urea (pH
5.0) with a flow rate 1 ml min − 1 , and 0.5 ml fractions were collected.
Aliquots from the fractions were transferred to a nitrocellulose mem-
brane by slot blotting and tested for reactivity with monoclonal anti-
bodies against MUC1, MUC16, and KS, and with polyclonal antibodies
against MUC5AC, MUC5B, and MUC4 using a standard chemilumines-
cence assay. The relative intensity of the slot blot bands was measured
by densitometry and quantitated by ImageQuant (Molecular Dynamics,
Sunnyvale, CA) software.
Agarose gel electrophoresis / western blotting . To determine the distri-
bution and charge dispersion of mucins over the CsCl density gradient,
agarose gel electrophoresis was performed under reducing conditions in
1 % (w / v) agarose as described previously. 60 Mucins were separated and
western blotted onto nitrocellulose membranes by vacuum transfer, then
probed for mucins with antibodies specific for MUC5B, MUC5AC, KS,
MUC1, MUC4, and MUC16 ( Table 1 ). Immunodetection was performed
using infrared dye-labeled secondary antibodies and visualized using a
Li-Cor Odyssey infrared detection system according to the manufactur-
er ’ s protocol (Li-Cor Biosciences, Lincoln, NE).
Mass spectrometry and multi-angle laser-light scattering . Mass
spectrometry for the analysis of mucus fractions was performed essen-
tially as described previously. 7 Mucin-containing pools from the density
gradient centrifugation were reduced, alkylated, and trypsin digested. The
resulting peptides were subjected to nano-LC-MS / MS (liquid chromato-
graphy-tandem mass spectrometry) analysis. All data were acquired using
Waters Q-Tof micro, hybrid quadrapole orthogonal acceleration time-
of-flight mass spectrometer (Waters, Manchester, UK) with the manu-
facturer ’ s MassLynx 4.0 software. The data processed were searched
against updated NCBInr and Swiss-Prot databases using the Mascot
Multi-angle laser-light scattering was performed in-line with gel per-
meation chromatography to measure the weight average MW and hydro-
dynamic radius of gyration distributions of isolated whole, and tryptically
digested MUC4 and MUC16 mucins. This approach yields absolute val-
ues of concentration, and MW and size distributions. 52 Briefly, the mucin
preparations were chromatographed on a Sephacryl 1000 or Sepharose
CL-2B (GE Healthcare Biosciences, Pittsburgh, PA; 15 cm × 2.5 cm) col-
umn eluted with 0.2 M NaCl at a flow rate of 500 ? l min − 1 . The column
effluent was passed through an in-line Dawn DSP laser photometer coup-
led to an Optilab 903 interferometric refractometer (both from Wyatt
Technology, Santa Barbara, CA) to measure light scattering and sample
concentration, respectively. Light scattering measurements were taken
continuously at 18 angles between 15 ° and 151 ° ; the data were analyzed
using Wyatt Technology ’ s ASTRA software.
EM of mucins . Mucins obtained from HTBE mucus collected in phos-
phate-buffered saline were isolated as the void volume eluting from a
Sephacryl 1000 (S1000) HPLC column. The mucins were adjusted to
~ 10 ? g ml − 1 and prepared for EM by fixing with 0.6 % glutaraldehyde
for 4 min at 20 ° C, as described previously. 18 The samples were observed
at 40 kV in a Tecnai 12 EM. Immuno-EM for MUC16 used CA125 /
MUC16-rich material harvested from an ovarian tumor, purified on a
MucosalImmunology | VOLUME 6 NUMBER 2 | MARCH 2013
CsCl density gradient by isopycnic centrifugation (as described above),
and CA125 / MUC16 taken from the material in the bottom of the gradi-
ent (fractions 18 – 20). After dialysis against 50 m M Mg acetate, the puri-
fied CA125 antigen was labeled by mixing with colloidal 10 nm gold
pre-absorbed with the OC125 mAb. The immunogold-labeled material
was isolated by HPLC on Sepharose CL2B, taking the material in the
void volume, and the molecules were spread onto water, transferred to a
carbon-coated grid, rotary shadowed with platinum, and examined in a
JEOL 100CX scanning transmission EM (JEOL USA, Peabody, MA), as
described previously. 61
Atomic force microscopy . Mucins from the CsCl density gradient
were reduced and alkylated, and dialyzed into 50 m M ammonium
hydrogen carbonate, pH 8.0, then digested with trypsin overnight at
37 ° C. The digest was chromatographed on a Sephacryl 1000 (S1000),
15 cm × 2.5 cm column eluted with 200 m M NaCl, 10 m M EDTA at a flow
rate of 0.3 ml min − 1 . The void fraction containing the large MUC4 and
MUC16 glycopeptides was collected — because the polymeric mucins
were reduced and digested, their isolated, highly glycosylated mucin
domains elute in the included volume of the column. 32 For observation
by AFM, the isolated MUC4 and MUC16 glycopeptides were mixed
with 10 m M MgCl, deposited on mica, air dried, and observed in an
Explorer atomic force microscope (Topometrix Corp, Santa Barbara,
CA) using a Nanosensors silicon probe (Neuchatel, Switzerland) at the
SUPPLEMENTARY MATERIAL is linked to the online version of the
paper at http://www.nature.com/mi
We thank the Directors and teams of the UNC Cystic Fibrosis Center
Tissue Culture Core, the Morphology and Morphometry Core, and the
Michael Hooker Microscopy Facility for supplying reagents and technical
expertise. We also thank Dr WG Matthews (Department of Physics,
University of South Florida) for kindly providing the AFM images of
Figure 4 , Dr Surinder Batra (Department of Biochemistry and Molecular
Biology, University of Nebraska Medical Center) for the kind gift of
the Ig8 mAb, and Ms Sarah Keilson ( http://sarahkeilson.com ) for the
illustration in Figure 8 . We also thank Fujirebio Diagnostics for the kind gift
of MUC1 antibodies, and Drs Jack Griffith and Sezgin Ozgur for their help
in molecular EM. Valuable comments and insights into the system came
from discussion with colleagues in the UNC Virtual Lung Group, Michael
Rubinstein, Brian Button, and Richard Boucher, in particular. These
studies were funded by grants from Cystic Fibrosis Foundation
and from the National Institutes of Health (R01HL103940, R01HL77844,
The authors declared no conflict of interest.
© 2013 Society for Mucosal Immunology
1 . Knowles , M . R . & Boucher , R . C . Mucus clearance as a primary innate
defense mechanism for mammalian airways . J. Clin. Invest. 109 , 571 – 577
( 2002 ).
2 . Fahy , J . V . & Dickey , B . F . Airway mucus function and dysfunction .
N Engl. J. Med. 363 , 2233 – 2247 ( 2010 ).
3 . Florey , H . , Carleton , H . M . & Wells , A . Q . Mucus secretion in the trachea .
Br. J. Exp. Pathol. 13 , 269 – 284 ( 1932 ).
4 . Lucas , A . M . & Douglas , L . C . Principles underlying ciliary activity in the
respiratory tract . Arch. Otolaryngol. 20 , 518 – 541 ( 1934 ).
5 . Yoneda , K . Mucous blanket of rat bronchus: an ultrastructural study .
Am. Rev. Respir. Dis. 114 , 837 – 842 ( 1976 ).
6 . Matsui , H . et al. Evidence for periciliary liquid layer depletion, not abnormal
ion composition, in the pathogenesis of cystic fi brosis airways disease .
Cell 95 , 1005 – 1015 ( 1998 ).
7 . Kesimer , M . et al. Tracheobronchial air-liquid interface cell culture: a model
for innate mucosal defense of the upper airways? Am. J. Physiol. Lung
Cell Mol. Physiol. 296 , L92 – L100 ( 2009 ).
8 . Thornton , D . J . , Gray , T . , Nettesheim , P . , Howard , M . , Koo , J . S . &
Sheehan , J . K . Characterization of mucins from cultured normal human
tracheobronchial epithelial cells . Am. J. Physiol. Lung Cell Mol. Physiol.
278 , L1118 – L1128 ( 2000 ).
9 . Holmen , J . M . et al. Mucins and their O-Glycans from human bronchial
epithelial cell cultures . Am. J. Physiol. Lung Cell Mol. Physiol. 287 ,
L824 – L834 ( 2004 ).
10 . Ali , M . , Lillehoj , E . P . , Park , Y . , Kyo , Y . & Kim , K . C . Analysis of the proteome
of human airway epithelial secretions . Proteome Sci. 9 , 4 ( 2011 ).
11 . Rahmoune , H . & Shephard , K . L . State of airway surface liquid on guinea
pig trachea . J. Appl. Physiol. 78 , 2020 – 2024 ( 1995 ).
12 . Sims , D . E . & Horne , M . M . Heterogeneity of the composition and thickness of
tracheal mucus in rats . Am. J. Physiol. 273 (5 Part 1) , L1036 – L1041 ( 1997 ).
13 . Widdicombe , J . H . & Widdicombe , J . G . Regulation of human airway
surface liquid . Respir. Physiol. 99 , 3 – 12 ( 1995 ).
14 . Wanner , A . , Salathe , M . & O ’ Riordan , T . G . Mucociliary clearance in the
airways . Am. J. Respir. Crit. Care Med. 154 (6 Part 1) , 1868 – 1902 ( 1996 ).
15 . Sleigh , M . A . , Blake , J . R . & Liron , N . The propulsion of mucus by cilia .
Am. Rev. Respir. Dis. 137 , 726 – 741 ( 1988 ).
16 . Satir , P . & Sleigh , M . A . The physiology of cilia and mucociliary interactions .
Annu. Rev. Physiol. 52 , 137 – 155 ( 1990 ).
17 . Sheehan , J . K . & Carlstedt , I . Hydrodynamic properties of human
cervical-mucus glycoproteins in 6M-guanidinium chloride . Biochem. J.
217 , 93 – 101 ( 1984 ).
18 . Kesimer , M . , Makhov , A . M . , Griffi th , J . D . , Verdugo , P . & Sheehan , J . K .
Unpacking a gel forming mucin: a view of MUC5B organization after
granular release . Am. J. Physiol. Lung Cell Mol. Physiol. 298 , L15 – L22
( 2010 ).
19 . Spicer , S . S . , Schulte , B . A . & Thomopoulos , G . N . Histochemical
properties of the respiratory tract epithelium in different species .
Am. Rev. Respir. Dis. 128 (2 Part 2) , S20 – S26 ( 1983 ).
20 . Stonebraker , J . R . et al. Glycocalyx restricts adenoviral vector access to
apical receptors expressed on respiratory epithelium in vitro and in vivo :
role for tethered mucins as barriers to lumenal infection . J. Virol. 78 ,
13755 – 13768 ( 2004 ).
21 . Pickles , R . J . Physical and biological barriers to viral vector-mediated delivery
of genes to the airway epithelium . Proc. Am. Thorac. Soc. 1 , 302 – 308 ( 2004 ).
22 . Ebong , E . E . , Macaluso , F . P . , Spray , D . C . & Tarbell , J . M . Imaging the
endothelial glycocalyx in vitro by rapid freezing/freeze substitution
transmission electron microscopy . Arterioscler. Thromb. Vasc. Biol. 31 ,
1908 – 1915 ( 2011 ).
23 . Rose , M . C . & Voynow , J . A . Respiratory tract mucin genes and mucin
glycoproteins in health and disease . Physiol. Rev. 86 , 245 – 278 ( 2006 ).
24 . Thornton , D . J . , Rousseau , K . & McGuckin , M . A . Structure and function of
the polymeric mucins in airways mucus . Annu. Rev. Physiol. 70 , 459 – 486
( 2008 ).
25 . Gouyer , V . , Gottrand , F . & Desseyn , J . L . The extraordinarily complex but
highly structured organization of intestinal mucus-gel unveiled in
multicolor images . PLoS ONE 6 , e18761 ( 2011 ).
26 . Aplin , J . D . MUC-1 glycosylation in endometrium: possible roles of the
apical glycocalyx at implantation . Hum. Reprod. 14 (Suppl 2) , 17 – 25
( 1999 ).
27 . Zhang , L . et al. Infection of ciliated cells by human parainfl uenza virus type
3 in an in vitro model of human airway epithelium . J. Virol. 79 , 1113 – 1124
( 2005 ).
28 . Monzon , M . E . , Casalino-Matsuda , S . M . & Forteza , R . M . Identifi cation of
glycosaminoglycans in human airway secretions . Am. J. Respir. Cell Mol.
Biol. 34 , 135 – 141 ( 2006 ).
29 . Kwilas , S . , Liesman , R . M . , Zhang , L . , Walsh , E . , Pickles , R . J . & Peeples ,
M . E . Respiratory syncytial virus grown in Vero cells contains a truncated
attachment protein that alters its infectivity and dependence on
glycosaminoglycans . J. Virol. 83 , 10710 – 10718 ( 2009 ).
30 . Jeffery , P . K . Structure and function of mucus-secreting cells of cat and
goose airway epithelium . Ciba. Found Symp. 54 , 5 – 23 ( 1978 ).
31 . Carlstedt , I . , Lindgren , H . , Sheehan , J . K . , Ulmsten , U . & Wingerup , L .
Isolation and characterization of human cervical-mucus glycoproteins .
Biochem. J. 211 , 13 – 22 ( 1983 ).
32 . Carlstedt , I . , Lindgren , H . & Sheehan , J . K . The macromolecular structure
of human cervical-mucus glycoproteins. Studies on fragments obtained
VOLUME 6 NUMBER 2 | MARCH 2013 | www.nature.com/mi
after reduction of disulphide bridges and after subsequent trypsin
digestion . Biochem. J. 213 , 427 – 435 ( 1983 ).
33 . Higuchi , T . et al. Molecular cloning, genomic structure, and expression
analysis of MUC20, a novel mucin protein, up-regulated in injured kidney .
J. Biol. Chem. 279 , 1968 – 1979 ( 2004 ).
34 . Finkbeiner , W . E . , Zlock , L . T . , Morikawa , M . , Lao , A . Y . , Dasari , V . &
Widdicombe , J . H . Cystic fi brosis and the relationship between mucin and
chloride secretion by cultures of human airway gland mucous cells .
Am. J. Physiol. Lung Cell Mol. Physiol. 301 , L402 – L414 ( 2011 ).
35 . Cao , R . , Wang , T . , Demaria , G . , Sheehan , J . & Kesimer , M . Mapping the
protein domain structures of the respiratory mucins: a mucin proteome
coverage study . J. Proteome. Res. 11 , 4013 – 4023 ( 2012 ).
36 . Madsen , J . , Mollenhauer , J . & Holmskov , U . Gp-340/DMBT1 in mucosal
innate immunity . Innate Immun. 16 , 160 – 167 ( 2010 ).
37 . Yin , B . W . , Dnistrian , A . & Lloyd , K . O . Ovarian cancer antigen CA125 is
encoded by the MUC16 mucin gene . Int. J. Cancer 98 , 737 – 740 ( 2002 ).
38 . Maeda , T . et al. Solution structure of the SEA domain from the murine
homologue of ovarian cancer antigen CA125 (MUC16) . J. Biol. Chem.
279 , 13174 – 13182 ( 2004 ).
39 . Hattrup , C . L . & Gendler , S . J . Structure and function of the cell surface
(tethered) mucins . Annu. Rev. Physiol. 70 , 431 – 457 ( 2008 ).
40 . Carrion-Vazquez , M . et al. Mechanical and chemical unfolding of a single
protein: a comparison . Proc. Natl. Acad. Sci. USA 96 , 3694 – 3699 ( 1999 ).
41 . Van den Steen , P . , Rudd , P . M . , Dwek , R . A . & Opdenakker , G . Concepts
and principles of O-linked glycosylation . Crit. Rev. Biochem. Mol. Biol. 33 ,
151 – 208 ( 1998 ).
42 . Burke , P . A . et al. Characterization of MUC1 glycoprotein on prostate
cancer for selection of targeting molecules . Int. J. Oncol. 29 , 49 – 55
( 2006 ).
43 . Kesimer , M . et al. Characterization of exosome-like vesicles released from
human tracheobronchial ciliated epithelium: a possible role in innate
defense . FASEB J. 23 , 1858 – 1868 ( 2009 ).
44 . Nustad , K . et al. Epitopes on CA 125 from cervical mucus and ascites
fl uid and characterization of six new antibodies. Third report from the
ISOBM TD-1 workshop . Tumour Biol. 23 , 303 – 314 ( 2002 ).
45 . McDowell , E . M . , Barrett , L . A . , Glavin , F . , Harris , C . C . & Trump , B . F .
The respiratory epithelium. I. Human bronchus . J. Natl. Cancer Inst. 61 ,
539 – 549 ( 1978 ).
46 . Lopez-Vidriero , M . T . Mucus as a natural barrier . Respiration 55 (Suppl 1) ,
28 – 32 ( 1989 ).
47 . Olmsted , S . S . , Padgett , J . L . , Yudin , A . I . , Whaley , K . J . , Moench , T . R . &
Cone , R . A . Diffusion of macromolecules and virus-like particles in human
cervical mucus . Biophys. J. 81 , 1930 – 1937 ( 2001 ).
48 . Sears , P . R . , Davis , C . W . , Chua , M . & Sheehan , J . K . Mucociliary inter-
actions and mucus dynamics in ciliated human bronchial epithelial cell
cultures . Am. J. Physiol. Lung Cell Mol. Physiol. 301 , L181 – L186 ( 2011 ).
49 . Davies , J . R . , Kirkham , S . , Svitacheva , N . , Thornton , D . J . & Carlstedt , I .
MUC16 is produced in tracheal surface epithelium and submucosal
glands and is present in secretions from normal human airway and
cultured bronchial epithelial cells . Int. J. Biochem. Cell Biol. 39 ,
1943 – 1954 ( 2007 ).
50 . de Mesy Bentley , K . L . An 11-mum-thick glycocalyx? it’s all in the
technique! . Arterioscler. Thromb. Vasc. Biol. 31 , 1712 – 1713 ( 2011 ).
51 . Button , B . et al. A periciliary brush promotes lung health by separating
the mucus layer from airway epithelia . Science , doi:10.1126/
science.1223012 (in press) ( 2012 ).
52 . Kesimer , M . & Sheehan , J . K . Analyzing the functions of large
glycoconjugates through the dissipative properties of their absorbed
layers using the gel-forming mucin MUC5B as an example . Glycobiology
18 , 463 – 472 ( 2008 ).
53 . Desseyn , J . L . , Tetaert , D . & Gouyer , V . Architecture of the large
membrane-bound mucins . Gene 410 , 215 – 222 ( 2008 ).
54 . Williams , S . J . , Wreschner , D . H . , Tran , M . , Eyre , H . J . , Sutherland , G . R . &
McGuckin , M . A . Muc13, a novel human cell surface mucin expressed by
epithelial and hemopoietic cells . J. Biol. Chem. 276 , 18327 – 18336 ( 2001 ).
55 . Pallesen , L . T . , Pedersen , L . R . , Petersen , T . E . , Knudsen , C . R . &
Rasmussen , J . T . Characterization of human mucin (MUC15) and
identifi cation of ovine and caprine orthologs . J. Dairy Sci. 91 , 4477 – 4483
( 2008 ).
56 . Harden , J . L . & Cates , M . E . Extension and compression of grafted
polymer layers in strong normal fl ows . J. Physique II 5 , 1093 – 1103 ( 1995 ).
57 . Raviv , U . , Giasson , S . , Kampf , N . , Gohy , J . F . , Jerome , R . & Klein , J .
Lubrication by charged polymers . Nature 425 , 163 – 165 ( 2003 ).
58 . Fulcher , M . L . , Gabriel , S . , Burns , K . A . , Yankaskas , J . R . & Randell , S . H .
Well-differentiated human airway epithelial cell cultures . Methods Mol.
Med. 107 , 183 – 206 ( 2005 ).
59 . Pickles , R . J . , McCarty , D . , Matsui , H . , Hart , P . J . , Randell , S . H . & Boucher ,
R . C . Limited entry of adenovirus vectors into well-differentiated airway
epithelium is responsible for ineffi cient gene transfer . J. Virol. 72 ,
6014 – 6023 ( 1998 ).
60 . Thornton , D . J . , Howard , M . , Devine , P . L . & Sheehan , J . K . Methods for
separation and deglycosylation of mucin subunits . Anal. Biochem. 227 ,
162 – 167 ( 1995 ).
61 . Sheehan , J . K . , Oates , K . & Carlstedt , I . Electron microscopy of cervical,
gastric and bronchial mucus glycoproteins . Biochem. J. 239 , 147 – 153
( 1986 ).
62 . Grinstead , J . S . , Schuman , J . T . & Campbell , A . P . Epitope mapping of
antigenic MUC1 peptides to breast cancer antibody fragment B27.29:
a heteronuclear NMR study . Biochemistry 42 , 14293 – 14305 ( 2003 ).
63 . Wesseling , J . , van der Valk , S . W . , Vos , H . L . , Sonnenberg , A . & Hilkens , J .
Episialin (MUC1) overexpression inhibits integrin-mediated cell adhesion
to extracellular matrix components . J. Cell Biol. 129 , 255 – 265 ( 1995 ).
64 . Hovenberg , H . W . , Davies , J . R . , Herrmann , A . , Linden , C . J . & Carlstedt , I .
MUC5AC, but not MUC2, is a prominent mucin in respiratory secretions .
Glycoconj. J. 13 , 839 – 847 ( 1996 ).
65 . Lidell , M . E . , Bara , J . & Hansson , G . C . Mapping of the 45M1 epitope to
the C-terminal cysteine-rich part of the human MUC5AC mucin . FEBS J.
275 , 481 – 489 ( 2008 ).
66 . Rousseau , K . , Wickstrom , C . , Whitehouse , D . B . , Carlstedt , I . & Swallow ,
D . M . New monoclonal antibodies to non-glycosylated domains of the
secreted mucins MUC5B and MUC7 . Hybrid Hybridomics 22 , 293 – 299
( 2003 ).
67 . Young , R . D . , Gealy , E . C . , Liles , M . , Caterson , B . , Ralphs , J . R . &
Quantock , A . J . Keratan sulfate glycosaminoglycan and the association
with collagen fi brils in rudimentary lamellae in the developing avian cornea .
Invest. Ophthalmol. Vis. Sci. 48 , 3083 – 3088 ( 2007 ).