Am J Physiol Gastrointest Liver Physiol 300: G914–G926, 2011.
First published February 17, 2011; doi:10.1152/ajpgi.00005.2011.
Proteomic analysis of the enterocyte brush border
Russell E. McConnell,1 Andrew E. Benesh,1 Suli Mao,1 David L. Tabb,2 and Matthew J. Tyska1
Departments of 1Cell and Developmental Biology and 2Biomedical Informatics, Vanderbilt University Medical Center,
Nashville, Tennessee
Submitted 10 January 2011; accepted in final form 16 February 2011
cytoskeleton; intestine; myosin; actin; microvilli
THE INTESTINAL TRACT is lined by a continuous sheet of transporting epithelial cells, also known as “enterocytes,” which are
responsible for processing and moving nutrients from the
lumenal space into the vasculature for distribution to peripheral
tissues. In the fully differentiated state, each enterocyte is
characterized by a striking apical structure that consists of
thousands of microvilli (61). These actin-based protrusions
extend from the surface of the cell into the lumen, forming a
cytoskeletal scaffold that is capable of supporting an extraordinarily large apical membrane surface area. In addition to the
clear benefit with regard to housing membrane-bound enzymes, transporters, and channels required for enterocyte function, recent studies have shown that microvilli also function as
vesicle-generating organelles, releasing membranes laden with
host defense machinery into the intestinal lumen (55).
All membrane-associated hydrolases, proteases, and lipases
required for nutrient processing traffic from the trans-Golgi to
the base of the brush border using the microtubule cytoskeleton
(74). There they are incorporated into the brush border membrane with a topology that positions their catalytic domains on
the outside of the cell (40). The numerous channels and
transporter proteins involved in moving nutrients from the
lumen into the enterocyte are also highly enriched in the brush
border membrane. The catalytic activities and functions of
nutrient processing/transporting enzymes have been the focus
of study for decades, in part because of the abundance and
accessibility of these proteins for purification from native
tissues (39). Moreover, investigators have been exploring the
Address for reprint requests and other correspondence: M. J. Tyska, Dept. of
Cell and Developmental Biology, Vanderbilt Univ. Medical Center, 465 21st
Ave S., 3150 Medical Research Bldg. III, Nashville, TN 37232 (e-mail:
[email protected]).
G914
biosynthetic mechanisms that enable specific membrane proteins to enrich in the brush border membrane for many years
(19, 32). While these aspects of the brush border have received
a great deal of experimental attention, a number of fundamental questions about brush border assembly and function remain
unanswered and, in many cases, entirely unexplored.
For example, it is now generally assumed that after their
delivery to the base of the brush border, membrane proteins are
distributed along the microvillar axis through interactions with
myosin motor proteins (100). Indeed, recent studies have
shown that myosin-1a, one of the most abundant motors
expressed in the enterocyte, is capable of exerting plus enddirected force on the microvillar membrane (56). However,
“intramicrovillar transport” has yet to be observed directly in
living cells, and the regulatory machinery that controls the
distribution of membrane-associated proteins is still poorly
understood. Another example of a critical unanswered question
relates to the assembly of the brush border during enterocyte
differentiation. The brush border represents one of the most
highly ordered F-actin arrays known to biologists; microvilli
are so tightly packed in terminally differentiated enterocytes
that there is no free space between adjacent structures. When
viewed in cross section, one can observe the hexagonal arrays
that reveal the tight packing of these protrusions. Remarkably,
there is little or no information on how microvillar actin
bundles are nucleated, how microvillar length is controlled, or
how the microvilli achieve perfect hexagonal packing during
enterocyte differentiation.
One promising approach for developing our understanding
of brush border assembly and function is to elucidate the brush
border proteome. Indeed, previous studies (5, 23) have already
taken steps in this direction with the proteomic analysis of
microvillar membrane vesicles or lipid raft preparations. Additional studies (17, 69, 94) have analyzed brush border membranes isolated from other organs including placenta and kidney. These studies have revealed that a remarkably diverse
complement of membrane-associated signaling, scaffolding,
and motor proteins resides in this domain. One major limitation
associated with all previous studies is that these data sets were
collected using material that was purposefully depleted in
cytoskeletal components. To date, the proteome of the entire
brush border, i.e., the intact actin cytoskeleton complete with
the overlaying apical membrane, has not been described.
Here, we report the complete brush border proteome obtained using two-dimensional (2-D) liquid chromatography
tandem mass spectrometry (LC-MS/MS). Intact brush borders
were isolated from adult mice using a well-established combination of differential and density gradient centrifugation (56).
We applied stringent criteria to the resulting data set so that
listed proteins were expected to have a high likelihood of
validation through other methods. The utility of this strategy
was supported by immunofluorescence validation of specific
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McConnell RE, Benesh AE, Mao S, Tabb DL, Tyska MJ.
Proteomic analysis of the enterocyte brush border. Am J Physiol
Gastrointest Liver Physiol 300: G914 –G926, 2011. First published
February 17, 2011; doi:10.1152/ajpgi.00005.2011.—The brush border
domain at the apex of intestinal epithelial cells is the primary site of
nutrient absorption in the intestinal tract and the primary surface of
interaction with microbes that reside in the lumen. Because the brush
border is positioned at such a critical physiological interface, we set
out to create a comprehensive list of the proteins that reside in this
domain using shotgun mass spectrometry. The resulting proteome
contains 646 proteins with diverse functions. In addition to the
expected collection of nutrient processing and transport components,
we also identified molecules expected to function in the regulation of
actin dynamics, membrane bending, and extracellular adhesion. These
results provide a foundation for future studies aimed at defining the
molecular mechanisms underpinning brush border assembly and function.
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BRUSH BORDER PROTEOME
novel proteins in a human intestinal epithelial cell culture
model. Similar to previous analyses of microvillar membrane
fractions (5, 23, 69), numerous proteins involved in nutrient
processing and transport were identified. Our analysis also
revealed several proteins implicated in the regulation of actin
dynamics, membrane/actin interactions, membrane bending,
and extracellular adhesion. In addition to accelerating our understanding of brush border assembly, maintenance, and function,
these data are expected to provide insights into the mechanisms
underlying intestinal pathologies characterized by brush border
damage or effacement, including celiac disease and enteric infections (e.g., enteropathogenic Esherichia coli) (78, 79).
MATERIALS AND METHODS
All reagents were purchased from Sigma (St. Louis, MO) unless
otherwise noted. Brush borders were isolated from mouse small
intestinal tissues using protocols originally developed by Miller and
Crane (59) over 50 yr ago (Fig. 1). During a single brush border
preparation, 20 –25 adult 129 sv/j mice were euthanized with CO2
followed by cervical dislocation in accordance with Vanderbilt Institutional Animal Care and Use Committee-approved protocols. The
full length of the small intestine was rapidly dissected, flushed in cold
Multidimensional LC-MS/MS
We used multidimensional (2-D) LC-MS/MS for shotgun proteomic analysis of brush border samples (50, 95, 98). To prepare
samples for shotgun analysis, isolated brush borders were resuspended
in Laemmlli SDS sample buffer and partially separated on a 10%
NuPage gel (Invitrogen). After samples had been run into the gel by
⬃2 cm, the gel was stained with Coomassie blue G-250 (Bio-Rad) and
then destained with sterile Milli-Q water. The protein-containing
region was excised from the gel, minced, and then submitted to the
Vanderbilt University Mass Spectrometry Core for trypsinization.
This partial separation via SDS-PAGE is an efficient sample clean-up
step with high recovery that also provides an effective tryptic digest of
hydrophobic proteins (45). For shotgun proteomic analysis, tryptic
peptide extracts were fractionated by strong cation exchange offline,
which provides for replicate reverse-phase analyses of selected ion
exchange fractions of interest without repeating the entire multidimensional separation. Protein digests were separated on a 75-mm ⫻
70-cm, 5-mm Partisphere SCX strong cation exchange column (Whatman). Peptides were eluted with a 10 –200 mM (pH 3.0 – 8.0) ammonium formate gradient (1 ml/min) in 25% acetonitrile as previously
described (1) with the addition of a final step of 0.5 M ammonium
formate during the last 10 min of the 65-min gradient to ensure that all
peptides were eluted. Fractions were collected into autosampler vials
using a Probot fraction collector (Dionex). Each SCX fraction was
analyzed by reverse-phase HPLC-coupled MS using a Thermo Scientific LTQ linear ion-trap MS instrument equipped with a Thermo
Surveyor LC system and operated with Xcalibur 1.4 and Bioworks 3.1
software. Electrospray ionization of peptides was done with a Thermo
nanospray source modified for automated vented column injection as
previously described (49).
Bioinformatics
Fig. 1. Outline of the brush border (BB) isolation procedure. All steps were
carried out on ice using prechilled buffers. The entire preparation can be
completed in a single afternoon. HB, homogenization buffer.
LC-MS/MS data were transcoded to mzML format by the msconvert tool of the ProteoWizard library (41) using settings for 32-bit
precision and zip compression. Tandem mass spectra were identified
using the mouse RefSeq database (release 44, 30,041 sequences,
downloaded on December 6, 2010). Seventy-one contaminant protein
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Brush Border Isolation From the Mouse Small Intestine
saline (150 mM NaCl, 2 mM imidazole-Cl, and 0.02% Na-azide), and
then soaked in cold dissociation solution (200 mM sucrose, 0.02%
Na-azide, 12 mM EDTA-K, 18.9 mM KH2PO4, and 78 mM
Na2HPO4; pH 7.2) for at least 30 min. A coarse screen was used to
separate released enterocytes from remnant tissue; cells were then
washed using multiple rounds of low-speed sedimentation (200 g for
10 min, X-15R centrifuge, Beckman Coulter) and resuspended in
fresh dissociation solution. Washed cell pellets were resuspended in
cold homogenization buffer (10 mM imidazole, 4 mM EDTA-K, 1
mM EGTA-K, 0.02% Na-azide, 1 mM DTT, and 1 mM Pefabloc-SC;
pH 7.2) and homogenized in a blender (Waring) with 4 ⫻ 15-s pulses.
At this point in the preparation, we typically used light microscopy to
confirm that brush borders were physically separated from each other,
nuclei, and other cellular fragments. Brush borders were collected
from the enterocyte homogenate by centrifugation at 1,000 g for 10
min. The resulting pellets were enriched primarily with brush borders
and nuclei; this material was washed several times with solution A (75
mM KCl, 10 mM Imidazole, 1 mM EGTA, 5 mM MgCl2, and 0.02%
Na-azide; pH 7.2). To separate nuclei from intact brush borders,
sucrose was added to this fraction to a final concentration of 50%.
Samples were overlayed with 40% sucrose in solution A and centrifuged at 130,000 g for 1 h at 4°C (L8 –70M, Beckman Coulter). The
resulting 40%/50% interface (Fig. 2A) contained isolated brush borders that were mostly free of nuclei. Brush borders were washed
several times with fresh solution A to remove residual sucrose and
then stored on ice if the material was to be used immediately or at ⫺80°C
for longer-term storage. A total of five replicate brush border preparations
were carried out to create material for shotgun proteomic analysis.
G916
BRUSH BORDER PROTEOME
sequences were added to this database, and the database was doubled
so that each protein appeared in both forward and reverse orientations.
The MyriMatch database search algorithm (version 1.6.79) matched
peptides derived from these proteins to the tandem mass spectra,
allowing for a precursor mass tolerance of 1.25 m/z and a fragment
tolerance of 0.5 m/z (84). Dynamic modifications included oxidation
of Met, loss of ammonia from NH2-terminal Gln, carbamidomethyl
Cys, and deamidation of Asn-Gly motifs. Peptides were allowed to
have either one or two trypsin-conformant termini but could contain
no more than two missed proteolytic sites. XCorr and mzFidelity
scores were computed for the best five matches by the MVH score.
IDPicker 2.6 build 206 used all three of these scores to sort confident
identifications away from erroneous ones, filtering to a 5% false
discovery rate (FDR), where this value was estimated as (2 ⫻
reverse)/(reverse ⫹ forward) (52). For final reporting, each protein
group was required to contribute a minimum of two distinct peptide
sequences and five spectra across the data sets, corresponding to the
five MudPIT experiments combined in a single report. Parsimony
routines were applied to remove subset and subsumable proteins,
resulting in an overall protein FDR of 1.2%. Manual annotation of the
resulting list was carried out to eliminate multiple entries of the same
protein under different RefSeq identifiers. In addition, there were
instances where IDPicker was unable to discriminate between different isoforms of the same protein based on the identified peptides.
While all identifications were included in the final IDPicker report
(Supplemental Material, Supplemental Report I), these entries were
reduced to a single listing in the final annotated data table of 646 brush
border proteins (Supplemental Table S1).1
residents before, we used the Human Protein Atlas (http://www.proteinatlas.org/) to determine if these components were expressed in the small
intestine and targeted to the brush border domain. In addition, a smaller
subset of proteins was validated using an immunofluorescence approach
with CACO-2BBE cells, a human intestine-derived epithelial cell culture
model (70). To prepare CACO-2BBE cells for immunofluorescence staining, cells were grown on glass coverslips for at least 10 days to ensure a
sufficient level of differentiation. Cells were washed with warm PBS
followed by fixation in 4% paraformaldehyde (Electron Microcopy Sciences) in PBS for 15 min at room temperature. The fixative was washed
away with fresh PBS, and cells were then permeabilized in 0.1% Triton
X-100 and PBS for 5 min at room temperature. Cells were washed again
followed by blocking in 5% BSA and PBS at 37°C for 1 h. The blocking
solution was washed from samples with fresh PBS, which was followed
by the application of primary antibody at 37°C for 1 h [anti-mucin-like
protocadherin (MLPCDH), 1:250, Sigma, HPA009081; anti-protocadherin (PCDH)24, 1:75, Sigma, HPA012569; and anti-Usher syndrome
1C (Ush1c), Sigma, HPA027398]. This was followed by an additional
wash and the application of goat anti-rabbit secondary antibodies (Invitrogen, 1:200) and Alexa 568-phalloidin (Invitrogen, 1:200) at 37°C for 1
h. Fully stained coverslips were mounted onto slides using ProLong
anti-fade (Invitrogen). All samples were viewed on an Olympus FluoView1000 with a ⫻100 objective. All images were contrast enhanced,
pseudocolored, and cropped with ImageJ (version 1.42h, National Institutes of Health).
RESULTS
Brush Border Isolation
Validation of Identified Proteins
We carried out validation of newly identified proteins at two levels.
For a larger subset of proteins that had not been reported as brush border
1
Supplemental Material for this article is available at the American Journal
of Physiology-Gastrointestinal and Liver Physiology website.
The combination of hypotonic lysis, differential centrifugation, and sucrose density gradient centrifugation described here
represents a well-established approach for isolating intact
brush borders (Fig. 1) (59). Indeed, when we used phasecontrast microscopy to examine the material recovered from
the sucrose density gradient 40%/50% interface (Fig. 2A),
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Fig. 2. Validation of brush border isolation. A: picture of the 40%/50% sucrose gradient centrifuge tube. The whitish layer that forms at the interface (*) is
enriched in brush borders and depleted in nuclei. B: SDS-PAGE analysis of isolated brush borders, which was separated using a 4 –12% polyacrylamide gel and
stained with Coomassie brilliant blue. Prominent protein bands are labeled to the right. Myo, myosin; SI, sucrase-isomaltase; CaM, calmodulin. C and D:
phase-images of the isolated brush border fraction (gradient interface highlighted in A); individual brush borders are highly refractive. E: laser scanning confocal
micrograph of isolated brush borders labeled with Alexa 488-concanavalin A (green) to mark membranes and Alexa 568-phalloidin (red) to label F-actin. Close
inspection showed that the microvillar actin bundles are tightly enclosed by the apical membrane. Core actin bundle rootlets extend below the apical membrane
into the terminal web; this produced the red band at the base of the brush border in the lateral view presented here. Membrane-containing structures adjacent
to the terminal web likely represent endosomes, vesicles, or mitochondria that remained associated with the brush border throughout the isolation procedure. Scale
bars ⫽ 60 ␮m in C, 12 ␮m in D, and 2 ␮m in E.
G917
BRUSH BORDER PROTEOME
Protein Identification
A total of 646 proteins were identified in isolated brush
borders. To be included in the final tally, we required that
proteins contribute a minimum of two distinct peptide se-
quences and five spectra to the cumulative data set obtained
from five distinct brush border preparations. Of the protein
classes identified, the most numerous were cytoskeletal proteins (n ⫽ 101), which made up 44% of total spectral counts
(20,772/47,600; Fig. 3). Other abundant protein classes include
hydrolases (16%) and membrane channels and transporters
(9%) that are established players in nutrient processing and
transport through the mucosa (Fig. 3). All other proteins were
binned into one of the following functional categories: mitochondrial, nuclear, metabolic, endoplasmic reticulum, signaling, membrane, adhesion, extracellular matrix, trafficking, and
immunological (Fig. 3 and Supplemental Table S1). Nine
proteins were of unknown function (Supplemental Table
S1). As the brush border isolation procedure described here
uses the full length of the small intestine, isolated organelles
will exhibit a protein composition that reflects their tissue of
origin (duodenum vs. ileum) (60). Therefore, the proteome
that we report was “averaged” along the length of the small
intestine.
Nutrient Processing Machinery
Hydrolases identified in this analysis included numerous
disaccharidases, peptidases, and lipases associated with the
brush border membrane (Table 1). Of particular note, the
disaccharidases maltase-glucoamylase, SI, and lactase made
up a significant portion of our data set, together accounting
for ⬃7% of total spectral counts. Prominent peptidases,
Fig. 3. Classification of the brush border
proteome into 14 distinct functional classes.
The pie chart shows the percentage of total
spectral counts contributed by each functional group for all proteins identified in this
analysis. Spectral counts and the corresponding percentages of the total are shown with
the label for each group. ER, endoplasmic
reticulum; ECM, extracellular matrix.
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⬎95% of the objects in the visual field were recognizable as
brush borders (Fig. 2, C and D). To assess the quality of this
preparation at a higher level of resolution, isolated brush
borders were stained with Alexa 488-phalloidin and Alexa
568-concanavalin A to label F-actin and the apical membrane,
respectively. These samples were then examined using laser
scanning confocal microscopy. The resulting images clearly
showed that the isolated structures contained an intact actin
cytoskeleton with tightly packed microvillar actin bundles of
uniform length (Fig. 2E). Microvillar actin bundles were also
enveloped in the apical membrane. Some brush borders demonstrated subapical concanavalin A staining, suggesting the
cofractionation of the subapical endosome or vesicles involved in the biosynthetic delivery of cargo to the brush
border. SDS-PAGE analysis of isolated brush borders revealed the dominant bands typical for such a preparation:
⬃42 kDa (actin), ⬃100 kDa (myosin-1a), ⬃120 –140 kDa
[sucrase-isomaltase (SI) and other transmembrane proteins],
and ⬃ 220 kDa (myosin-2) (Fig. 2B). A lower-molecularweight (⬍20 kDa) band representing calmodulin, a light
chain that binds to numerous myosin motor proteins, was
also observed (Fig. 2B).
G918
BRUSH BORDER PROTEOME
Table 1. Hydrolases
Coverage, %
Total Spectra
Number of Preparations
Refseq ID
Sucrase-isomaltase, intestinal
Aminopeptidase N
Maltase-glucoamylase
Lactase-phlorizin hydrolase preproprotein
Glutamyl aminopeptidase
Neprilysin
Intestinal alkaline phosphatase
N-acetylated-␣-linked acidic dipeptidase-like protein
Angiotensin-converting enzyme 2 precursor
Dipeptidyl peptidase 4
Angiotensin-converting enzyme isoform 1
Meprin A ␤-subunit precursor
Neutral ceramidase
Trehalase precursor
Intestinal alkaline phosphatase precursor
Dipeptidase 1 precursor
Phospholipase B1, membrane-associated isoform 1
Aminopeptidase P
␥-Glutamyltranspeptidase 1
Ectonucleotide pyrophosphatase/phosphodiesterase 7
Neutral ␣-glucosidase AB
Sphingosine-1-phosphate lyase 1
Meprin A ␣-subunit
Phosphatidylinositide phosphatase SAC1
Phospholipase D1
Membrane primary amine oxidase
Phospholipase A2, group IVC
54
57
50
29
46
53
56
50
58
45
40
33
37
42
40
56
25
29
20
27
20
23
10
25
16
11
9
1,398
1,346
1,220
570
510
379
321
313
288
236
210
189
149
123
108
94
71
52
36
32
31
22
20
18
18
6
6
5
5
5
5
5
5
5
5
5
5
5
5
5
5
5
5
5
5
5
5
5
5
5
5
5
3
3
NP_001074606.1
NP_032512.2
NP_001164474.1
NP_001074547.1
NP_031960.1
NP_032630.2
NP_001074551.1
NP_001009546.1
NP_001123985.1
NP_034204.1
NP_997507.1
NP_032612.2
NP_061300.1
NP_067456.1
NP_031458.2
NP_031902.2
NP_001074876.1
NP_573476.2
NP_032142.1
NP_001025462.1
NP_032086.1
NP_033189.2
NP_032611.2
NP_109617.1
NP_001157528.1
NP_033805.1
NP_001161976.1
such as aminopeptidase N, dipeptidyl peptidase 4, and two
isoforms of angiotensin-converting enzyme (64), were also
well represented in our data set, comprising ⬃4% of the
total spectra (Table 1). Enzymes involved in the metabolism
of lipids were prominent in the data set, including the
lipases alkaline sphingomyelinase (ectonucleotide pyrophosphatase-phosphodiesterase-7) and neutral ceramidase.
In addition to roles in metabolism, many of the identified
lipases are critical intermediates in lipid signaling, including
phospholipases A2, B1, and D1 (Table 1).
Brush Border Channels and Transporters
We identified a total of 53 channels and transporters (Supplemental Table S1), a subset of which are highlighted in Table 2.
Many of the identified channels and transporters are driven by
electrochemical gradients, such as Na⫹-glucose transporter
(SGLT)1 and SGLT3b (3), proton-coupled peptide transporter
1, and the potential cell volume regulators voltage-dependent
anion-selective channel protein (VDAC)1 (67) and VDAC2
(87), whereas others are regulated by transient ionic potentials,
such as Ca2⫹-activated Cl⫺ channel anactomin-6 (77). Concentrations of these key ions are controlled by ion transport
machinery in the brush border, such as Na⫹/H⫹ exchanger
(NHE)3, Na⫹-K⫹-ATPase, and their regulatory factors NHE
member 3 regulator (NHERF)1 and NHERF3 (42, 103) (Table
2). NHERF1 has also been shown to regulate another transporter present in our data set, multidrug resistance protein 2
(Table 2) (48), a pharmacologically important enzyme responsible for the secretion of a wide range of drugs, toxins, and
endogenous compounds (38). Enzymes involved in the uptake
of lipids were also particularly abundant in our data set,
including those responsible for chylomicron assembly [microsomal triglyceride transfer protein (MTTP), protein disulfide
isomerase family A member 3, apoliprotein A, and apolipoprotein B] and cholesterol transport [Niemann-Pick C1-like
protein 1, ATP-binding cassette subfamily G (Abcg)5, and
Abcg8] (Table II). Mutations in three of the identified lipid
transporters have been shown to cause the malabsorptive diseases sitosterolemia (Abcg5 or Abcg8) (43) and abetalipoproteinemia (MTTP) (97).
Other Membrane-Associated Proteins
In addition to the appearance of numerous nutrient processing enzymes, transporters, and channels in the present
dataset, we also identified a number of other membraneassociated proteins that are believed to play important roles
in brush border function and maintenance. Although this
class only accounted for ⬃1% of all identified spectra, a
number of functionally significant proteins from this group
were observed in all five brush border preparations. Among
these were two prominent annexin isoforms, A2 and A13,
which were both present at high levels of coverage (104 and
80 total spectra at 57% and 62% coverage, respectively).
Annexins are small Ca2⫹-dependent membrane-binding proteins that have been implicated in a wide range of membrane-related activities, ranging from vesicle fusion with the
plasma membrane to budding of vesicles into the multivesicular body (27). We also identified several members of the
galectin protein family, including galectin-4, -6, and -9 (56,
38, and 9 total spectra, respectively). Galectins bind to
galactosyl groups on transmembrane proteins and have been
implicated in a variety of cellular functions ranging from
biosynthetic sorting of membrane proteins to organizing
apical membrane domains and, potentially, mucosal host
defense (18, 20).
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Description
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Table 2. Transporters and channels
Coverage, %
Total
Spectra
Number of
Preparations
Refseq ID
Na -K -transporting ATPase ␣1-subunit
Microsomal triglyceride transfer protein large subunit
Na⫹-K⫹-transporting ATPase ␣3-subunit
Na⫹-glucose cotransporter 1
Niemann-Pick C1-like protein 1
Na⫹-K⫹-transporting ATPase ␣4-subunit
Proton-coupled peptide transporter (solute carrier family 15 member 1)
Na⫹/H⫹ exchange regulatory cofactor 3
Na⫹-glucose transporter 3b (solute carrier family 5 member 4b)
Multidrug resistance protein 2
Na⫹-K⫹-transporting ATPase ␤1-subunit
Apolipoprotein B precursor
Na⫹/H⫹ exchange regulatory cofactor 1
Solute carrier family 3, member 1
Cl⫺ intracellular channel protein 5
Na⫹-dependent neutral amino acid transporter 1
Protein disulfide-isomerase A3 precursor
K⫹-transporting ATPase ␣-chain 2
Anoctamin-6
Solute carrier family 26, member 6
Voltage-dependent anion-selective channel protein 1
Voltage-dependent anion-selective channel protein 2
Ca2⫹-activated Cl⫺ channel regulator 4
Na⫹/H⫹ exchanger 3
ATP-binding cassette subfamily G, member 5
ATP-binding cassette subfamily G, member 8
Apolipoprotein A-IV precursor
50
56
21
29
21
12
20
68
18
21
30
13
46
30
69
14
50
8
22
32
41
38
14
17
15
15
33
743
341
323
321
190
185
176
131
112
92
81
80
75
71
68
67
61
60
57
37
31
31
26
24
23
17
17
5
5
5
5
5
5
5
5
5
5
5
5
5
5
5
5
5
5
5
5
4
4
5
5
4
5
5
NP_659149.1
NP_001156929.1
NP_659170.1
NP_062784.3
NP_997125.2
NP_038762.1
NP_444309.2
NP_067492.2
NP_075708.2
NP_038834.2
NP_033851.1
NP_033823.2
NP_036160.1
NP_033231.2
NP_766209.1
NP_083154.1
NP_031978.2
NP_619593.2
NP_780553.2
NP_599252.2
NP_035824.1
NP_035825.1
NP_997091.3
NP_001074529.1
NP_114090.1
NP_080456.1
NP_031494.2
⫹
⫹
The Brush Border Cytoskeleton
A total of 101 cytoskeletal proteins were identified in this
analysis (Supplemental Table S1). Due to the inclusion of an
intact cytoskeleton in isolated brush borders, this category
accounts for almost half of all spectral counts (20,772 or 44%).
The three identified actin isoforms represented ⬃10% of the
total spectra, as might be expected for this extraordinarily
actin-rich domain (Table 3, high abundance). We also identified proteins that serve in actin bundling (villin-1 and both
isoforms of plastin) and actin-membrane interactions (ezrin,
radixin, and harmonin) (12, 14, 31, 76). Actin and some
associated cytoskeletal proteins have previously been observed
in the proteome of brush border membrane vesicles (23). This
prior analysis overlaps with the present data set to a large
degree; however, the inclusion of an intact cytoskeleton in our
preparation allowed us to identify low-abundance cytoskeletal
proteins (⬍100 total spectra) that were not detected in prior
studies (Table 3, low abundance). A number of these newly
identified proteins have been shown to interact with actin
and/or regulate actin dynamics in other cellular contexts but
have never been described in the enterocyte brush border:
adenylyl cyclase-associated protein-1 (11), Cordon-bleu (2),
insulin receptor tyrosine kinase substrate (IRTKS) (58), cortactin (4), and EGF receptor (Table 3). We also detected
components previously implicated in regulating microvillar
actin organization, such as Eps8 and its downstream target
Rac1 (66) (Table 3, low abundance).
Myosin Motor Proteins
A total of 14 different myosin superfamily members were
detected in the brush border proteome (Table 4). Many of these
proteins have been identified in previous cell biological stud-
ies, whereas others are newly identified residents of the brush
border. The most abundant myosins, as judged by spectral
counts, were class 2 myosins. We obtained strong evidence for
the presence of all three nonmuscle myosin-2 isoforms: 2a, 2b,
and 2c. Studies have shown that myosin-2 is found in the
terminal web, where it connects neighboring actin core rootlets
(62), and in the circumferential actin band that wraps around
the base of the brush border, at the level of cell-cell contacts
(36). In addition, we identified 10 “unconventional” myosins,
including the known brush border constituent class 1 myosins
(myosins-1a, -1c, -1d, and -1e) (9, 82, 89) as well as myosin5b, -6, -7b and, at lower levels, myosin-7a (Table 4) (15, 34).
Myosins not previously found in the brush border include myosin15-like protein and myosin-18a (Table 4). We also identified three
small polypeptides that are known to function as light chains,
including calmodulin. These proteins bind directly to IQ motifs in
various myosin heavy chains and regulate the mechanochemical
activity of these motors (7, 51, 85).
Adhesion Proteins
As isolated brush borders contain portions of the junctional
complex, we expected to identify proteins with established functions in intercellular adhesion. Indeed, our analysis revealed the
tight junction components zonula occludens (ZO)-1, -2, and -3 as
well as claudin-3 and -7 (53) (Table 5). We also identified
cadherin-1 (also known as E-cadherin) and cadherin-17, both of
which are localized to the junctional complex and basolateral
membrane (10, 35) (Table 5). MLPCDH and PCDH24 were also
present at significant levels (Table 5). Although MLPCDH has
previously been listed in the proteome of brush border membrane
vesicles (23), the presence of PCDH24 is a novel finding. Given
their abundant spectral counts and appearance in all five brush
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Description
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Table 3. Cytoskeletal components
Description
Coverage, %
Number of
Preparations
Refseq ID
2,512
1,450
1,278
986
872
861
658
529
453
326
244
226
202
187
173
172
169
158
146
140
125
111
5
5
5
5
5
5
5
5
5
5
5
5
5
5
5
5
5
5
5
5
5
5
NP_031419.1
NP_112447.2
NP_031418.1
NP_958796.2
NP_032497.1
NP_780706.1
NP_033535.2
NP_001028382.1
NP_033536.2
NP_075745.1
NP_076331.2
NP_001171138.1
XP_003085904.1
NP_068695.1
NP_787030.2
NP_033286.2
NP_149064.1
NP_034794.2
NP_001003908.1
NP_033067.2
NP_001003667.1
NP_033948.1
87
26
23
22
22
22
19
15
14
12
11
10
8
7
7
7
7
5
4
5
4
5
4
4
4
5
5
5
5
4
5
4
3
3
NP_598628.1
NP_031971.2
NP_742120.2
NP_076224.1
NP_598841.1
NP_076138.2
NP_766084.3
NP_033928.1
NP_666355.1
NP_780495.2
NP_034357.2
NP_031630.1
NP_031624.2
NP_033927.2
NP_080109.1
NP_035202.1
NP_071709.2
Total Spectra
High abundance
81
85
67
60
86
53
68
70
73
75
35
47
26
62
38
42
39
71
37
17
7
51
Low abundance
EGF receptor pathway substrate 8-like 3
EGF receptor pathway substrate
Rootletin
Actin-related protein 3
Filamin-B*
Harmonin*
Cordon-bleu*
F-actin-capping protein ␤-subunit
Actin-related protein 2
Taperin*
Filamin-A
F-actin-capping protein ␣2-subunit
Adenylyl cyclase-associated protein 1
F-actin-capping protein ␣1-subunit
Insulin receptor tyrosine kinase substrate*
Profilin-1
Tropomyosin ␣3-chain
50
27
1
36
9
25
8
38
17
7
4
15
21
12
13
31
13
*Proteins exhibiting robust staining in the brush border.
border preparations, MLPCDH and PCDH24 were included as
targets for validation.
Validation of Identified Proteins
As a first step toward validating newly identified proteins,
we selected a subset of actin- and membrane-associated proteins from our list for cross-referencing with the Human
Proteome Atlas (http://www.proteinatlas.org/). Remarkably,
all cross-referenced proteins demonstrated strong expression in
human intestinal tissue sections (duodenal and general small
intestinal samples), with many exhibiting robust staining in the
brush border (see proteins marked with asterisks in Tables
3–5). Proteins validated with the Human Proteome Atlas included IRTKS, Cordon-bleu, harmonin, filamin-B, taperin,
PCDH24, MLPCDH, and myosin-18a.
Some proteins were chosen for further validation using
immunofluorescence microscopy. For these experiments, we
chose to focus on the poorly characterized PCDH isoforms that
were identified with significant spectral counts: PCDH24 and
MLPCDH. Because harmonin has been shown to localize to
the brush border (12) and has been implicated as a binding
partner for cadherin superfamily members in hair cell stereocilia (13), we stained for this component as well. Antibodies to
PCDH24, MLPCDH, and harmonin were obtained from the
Sigma Prestige collection and used to stain fully differentiated
CACO-2BBE cells, a human-derived intestinal epithelial cell
line (70). All three probes produced striking punctate staining
at the apical surface, representative of microvillar labeling
(Fig. 4). PCDH24, MLPCDH, and harmonin exhibited mosaic
staining; cells exhibited a wide range of expression levels with
some cells expressing little or no antigen. Mosaic expression
has been observed for other well-characterized apical membrane proteins in this cell line, including SI and intestinal
alkaline phosphatase (70, 91). These validation experiments
suggest that PCDH24 and MLPCDH may be bona fide components of the enterocyte microvillus.
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Actin, cytoplasmic 1
Keratin, type II cytoskeletal 8
Actin, aortic smooth muscle
Plectin*
Keratin, type I cytoskeletal 19
␤-Actin-like protein 2
Villin-1
Plastin-1
Ezrin
Keratin, type I cytoskeletal 20
Desmoplakin
Spectrin ␣-chain, brain isoform 2
Cortactin
␣-Actinin-4
Spectrin ␤-chain, brain 1 isoform 1
Spectrin ␤-chain, brain 1 isoform 2
Keratin, type II cytoskeletal 7
Keratin, type I cytoskeletal 18
Clathrin heavy chain 1
Radixin
Keratin, type II cytoskeletal 1b
␣1-Catenin
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BRUSH BORDER PROTEOME
Table 4. Myosin motor proteins
Description
Coverage, %
Spectra
Number of Preparations
Refseq ID
5
5
5
5
5
5
5
5
5
5
4
4
3
4
NP_082297.1
NP_001074688.1
NP_071855.2
NP_038635.2
NP_115770.2
NP_796364.2
NP_780469.1
NP_001034635.2
NP_963894.1
NP_001074243.1
XP_003085561.1
NP_851417.2
NP_035716.1
NP_032689.2
5
5
5
NP_034990.1
NP_075891.1
NP_031615.1
Heavy chains
70
74
63
59
47
49
15
45
23
16
18
7
4
2
2,468
1,582
793
569
362
265
202
179
58
34
23
12
9
6
Light chains
Myosin light polypeptide 6
Myosin regulatory light chain 12B
Calmodulin
76
62
48
88
41
23
*Proteins exhibiting robust staining in the brush border.
DISCUSSION
Because the enterocyte brush border has been the focus of
biochemical characterization for decades, many of the nutrient
processing and transporting proteins identified in our analysis
were established as enriched components of this domain long
ago. More recent proteomic analyses of apical membrane
vesicles or lipid raft preparations (5, 23) has focused on
cataloguing membrane channels and transporters. Our analysis
identified many of these components, including ATP-binding
cassette family proteins, intracellular Cl⫺ channel isoforms,
solute carrier family members, multidrug resistant protein
isoforms, a diverse array of ion-coupled transporters, and
others (Table 2). The approach presented here also enabled us
to identify established brush border residents that were not
readily detected in previous studies. For example, previous
analyses of brush border membrane vesicles were unable to
identify SI, one of the principle sugar-processing hydrolases
in the brush border (23). Our analysis revealed 1,398 total
spectra that matched to SI, with an average of ⬃280 per
preparation. The lack of SI in previous results may be
related to a partitioning of SI-depleted lipid domains into
vesicles isolated using the cationic precipitation procedure.
The true utility of analyzing the intact brush border was
demonstrated by the abundant spectral counts of cytoskeletal
proteins that are well-characterized residents of this domain
(Table 3). A total of 3,790 spectra representing the two major
actin isoforms (cytoplasmic/␤-actin and aortic smooth muscle
actin) were detected in this analysis. Other well-established
cytoskeletal proteins that were readily detected include the
actin bundling proteins villin and plastin/fimbrin (658 and 529
total spectra, respectively) and the two major keratin isoforms
expressed in the enterocyte, cytoplasmic keratin-8 and -19
Table 5. Adhesion molecules
Description
Coverage, %
Total
spectra
Number of
Preparations
Refseq ID
Mucin-like protocadherin (cadherin-related family member 5) isoform 2*
␦1-Catenin isoform 1
Cadherin-17 precursor
Epithelial cell adhesion molecule
Mucin-13 precursor
Protocadherin-24 (cadherin-related family member 2)*
Carcinoembryonic antigen-related cell adhesion molecule 1
␣2-Catenin isoform 1
Cell surface A33 antigen precursor
Cingulin
Tetraspanin 8
Cadherin-1
Claudin-7
Carcinoembryonic antigen-related cell adhesion molecule 20
Tight junction protein zonula occludens-1
Tight junction protein zonula occludens-3
Tight junction protein zonula occludens-2
Claudin-3
30
35
31
34
14
8
17
10
20
9
10
4
8
10
5
6
3
9
75
73
72
62
44
17
16
14
14
13
12
10
9
8
8
7
6
5
5
5
5
5
5
5
5
5
4
4
4
5
4
4
5
2
3
3
NP_082345.1
NP_001078919.1
NP_062727.1
NP_032558.2
NP_034869.1
NP_001028536.2
NP_001034275.1
NP_663785.2
NP_067623.1
NP_001032800.2
NP_666122.1
NP_033994.1
NP_001180548.1
NP_082115.2
NP_033412.2
NP_038797.2
NP_001185914.1
NP_034032.1
*Proteins exhibiting robust staining in the brush border.
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Nonmuscle myosin-2c
Myosin-1a
Nonmuscle myosin-2a
Smooth muscle myosin-2
Myosin-7b
Myosin-1d
Nonmuscle myosin-2b
Myosin-6
Myosin-5b
Myosin-1c
myosin-15-like (predicted)
Myosin-1e
Myosin-18a*
Myosin-7a
G922
BRUSH BORDER PROTEOME
(1,450 and 872 total spectra, respectively). Carrying out shotgun MS on intact brush borders also enabled us to discover
cytoskeletal components that are unlikely to be enriched in
membrane/vesicle fractions examined in previous studies, including lower abundance myosin motors, proteins involved in
controlling actin dynamics, membrane bending machinery, and
extracellular adhesion molecules. Each of these is discussed in
more detail below.
Mitochondrial and Metabolic Machinery
A large number of mitochondrial components and cytosolic
enzymes that function in general metabolism were present in
our samples, representing 14% of all spectra identified in this
analysis. Although soluble metabolic machinery could conceivably be captured in the microvillus during the brush border
isolation, the mitochondrial signals were most likely derived
from fragments of this compartment that remain associated
with the brush border throughout the isolation process (see Fig.
4a in Ref. 56). The close association of mitochondria with
isolated brush borders may reflect the need to maintain high
ATP levels, which would be required to sustain the proper
function of motor proteins, channels, and transporters that are
enriched in this domain. Indeed, previous proteomic analyses
of hair cell stereocilia revealed numerous metabolic proteins
(e.g., creatine kinase) that were shown to be bona fide components of the hair bundle critical for supporting the metabolic
demands of the mechanotransduction machinery (80). While
many of the metabolic enzymes listed here likely come from
contaminating mitochondrial fragments, others may represent
as-yet-unidentified brush border components and have therefore been included here for completeness.
Myosin Motor Proteins
Conventional myosins. Numerous nonmuscle myosin-2 isoforms (2a, 2b, 2c, and smooth muscle) were detected in
isolated brush borders (Table 4). Although myosin-2 is a
well-known component of the circumferential band of actin
that lines the periphery of the cell at the base of the brush
border, the functional significance of multiple isoforms remains to be established. Based on the diversity of catalytic and
mechanical properties of these motors (44, 65), one possibility
is that different isoforms carry out distinct functions. This
would be consistent with recent studies on non-muscle myosins-2a and -2b (83), which independently regulate adhesion
receptor function and F-actin organization at cell-cell contacts,
respectively. An alternate idea is that all class 2 myosins
contribute to the same function, but by controlling expression
levels and targeting of functionally distinct isoforms to the
junctional complex, enterocytes are able to fine tune the contractility of this domain. Intriguingly, physiological studies
have shown that barrier function of the intestinal mucosa is
intimately linked with the activity of myosin-2 in the junctional
complex (88, 102).
Unconventional myosins. This analysis also revealed the
presence of a variety of unconventional (nonclass 2) myosin
isoforms, including myosin-1a, -1c, -1d, -1e, -5b, -6, -7b, -7a,
and -18a and a myosin-15-like protein. While many of these
are well characterized and have been linked to the enterocyte
brush border through previous biochemical or immunocytological analyses (6, 15, 34), the full brush border proteome did
provide some interesting insights. For example, both myosin-7b and -7a have been implicated as enterocyte motors (8),
but the present data set clearly shows that myosin-7b is much
more abundant in the brush border based on spectral counts
(Table 4). This is potentially interesting in light of cell biological studies that have shown that myosin-7b enriches in the
distal half of microvilli (15). With tandem MyTH4/FERM
motifs in its tail domain, myosin-7b is well equipped to bind a
variety of cargoes and serve as a major plus end-directed
transporter in the microvillus.
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Fig. 4. Immunofluorescent validation of selected novel brush border components. CACO-2BBE cells were stained with Alexa 568-phalloidin (red) and antibodies
directed against mucin-like protocadherin (MLPCDH; A and D), protocadherin-24 (PCDH24; B and E), or harmonin (C and F), which were detected with Alexa
488-conjugated secondary antibodies (green). A–C: en face views taken near the apical surface of the cell monolayer showed that all three proteins demonstrated
a punctate staining pattern that was highly mosaic. All three proteins localized strongly to the apical surface of the cells, at the top of the monolayer, as viewed
in the vertical sections (D–F). Scale bars ⫽ 10 ␮m in A–C and 5 ␮m in D–F. Insets show magnified views of individual cells. Box width is 20 ␮m in all cases.
G923
BRUSH BORDER PROTEOME
Regulators of Actin Assembly
One of the most striking features of the enterocyte brush
border is the high density of microvilli that extend off the
apical surface into the intestinal lumen. Remarkably, there is
little information on how the numerous microvilli that comprise the brush border are nucleated to grow or how their
growth is coordinated to create protrusions of the same length.
To control the spatial and temporal distribution of actin filaments, cells typically control the targeting and activity of
proteins that function to overcome the rate-limiting step in
filament formation: nucleation. Our analysis did identify components of the actin-related protein (Arp)2/3 complex, the
nucleation machinery that gives rise to the meshwork of actin
filaments at the leading edge of motile cells and the cell cortex
(71). Because Arp2/3 is a branched nucleator that functions to
initiate new filament growth off the side of preexisting
“mother” filaments, this complex is unlikely to function in the
growth of microvillar actin bundles. Arp2/3 components are
more likely to reside in the junctional band of actin filaments
that surround the base of the brush border; fragments of this
array were included in brush borders isolated using our preparation (56).
Intriguingly, our analysis also identified Cordon-bleu, a
newly identified multi-WH2 domain-containing protein that
functions in nucleating actin filaments (72). Cordon-bleu contains three WH2 domains, and each one has the ability to bind
to a single G-actin monomer (2, 72). As such, current models
suggest that a single molecule of Cordon-bleu brings together
three G-actin monomers to form the energetically unfavorable
trimer, after which F-actin polymerization proceeds rapidly.
Importantly, in vitro experiments have suggested that Cordonbleu can support the polymerization of unbranched actin filaments (2), making it a good candidate for nucleating the
production of parallel actin bundles found in microvilli.
Even in the fully differentiated state, microvilli are highly
dynamic structures, with actin bundles undergoing continuous
treadmilling: actin monomers incorporate into filament plus
ends at the microvillus tip and dissociate from minus ends at
the base (90). The brush border proteome contains two proteins
that may regulate the length of microvillar actin bundles by
controlling the rate of actin polymerization: the actin bundling
and capping proteins Eps8 and Eps8-like 3 (Table 4) (22). Eps8
has previously been shown to be critical for regulating the
length and organization of intestinal microvilli in Caenorhabditis elegans (16), and yet Eps8 knockout mice display only
slight perturbations to microvillar morphology (86). An explanation for this difference is that C. elegans possesses only one
Eps8 homolog, whereas vertebrates possess four: Eps8 and
Eps8-like 1, 2, and 3 (66). Our data indicate that Eps8-like 3 is
the predominant member of this family in the enterocyte brush
border (Table 4), further suggesting that this isoform may be
responsible for regulating microvillar length in vertebrates.
Membrane Bending
This analysis also revealed the presence of IRTKS in the
brush border (also known as brain-specific angiogenesis inhibitor 1-associated protein 2-like protein 1). IRTKS belongs to
the IRSp53/MIM family of inverse BAR domain proteins (58);
inverse BAR domain proteins have been implicated in the
bending and support of outward membrane protrusions such as
filopodia (58, 99). IRTKS also contains a WH2 domain that
may enable direct interactions with actin filaments (58). This
finding might hold special relevance in the context of the gut as
studies with the adherent pathogenic bacterium, enterohemorrhagic E. coli, have implicated IRTKS in the formation of
actin-rich cell surface pedestals, which are critical for the
virulence of this organism (93, 96). In the context of normal
enterocyte physiology, IRTKS could play a role in deforming
the apical membrane during brush border assembly, stabilizing
the curvature of microvillar membrane, or linking the microvillar membrane directly to supporting core actin bundles.
Peripheral Membrane Binding
Other membrane-associated proteins identified in this data set
include members of the annexin (annexin A2 and A13) and
galectin (galectin-4, -6, and -9) families. Annexin A2 has been
shown to play important roles at the interface of the actin cytoskeleton and the plasma membrane (57), where it is thought to
help polymerize actin in a phosphatidylinositol 4,5-bisphosphateregulated manner (33). This coupling of the membrane to polymerizing actin is thought to help drive phagocytosis in macrophages
and rod outer segments (24, 46). Because annexin A2 is strongly
localized to the terminal web region of the brush border where
endocytosis and exocytosis occur (54), it is a likely candidate for
regulating vesicle trafficking to and from the brush border. Galectins have also been shown to play important roles in the apical
trafficking of membrane proteins (20); indeed, knockdown of just
one galectin, galectin-4, in HT-29 cells resulted in a dramatic
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We also identified myosins not previously recognized as
brush border constituents, including myosin-18a. Myosin-18a
diverges significantly from most myosins but is somewhat
related to class 2 myosins, possessing a motor domain, a single
IQ motif, a coiled-coil region, and a COOH-terminal globular
tail domain (25). A unique feature of myosin-18a is the
presence of an NH2-terminal PDZ domain that binds actin in an
ATP-insensitive manner (37). The mechanochemical properties of this motor are also likely to be unique, as a critical
highly conserved glutamate residue in the motor domain is
substituted by a glutamine (37). This glutamate is thought to be
involved in regulating phosphate release after ATP hydrolysis
(47), the rate-limiting step of the myosin mechanochemical
cycle. In fact, a conserved glutamate to aspartate mutation in
this position has been shown to uncouple the chemical and
mechanical properties of myosin-1a (101). It will be interesting
to determine if myosin-18a is a catalytically active, actinactivated ATPase, like all other members of the myosin superfamily characterized to date.
Peptides matched to a myosin-15-like protein were also
found in four of the five brush border preparations. The
myosin-15-like entry that matched these peptides (XP_003085561.1)
is a predicted protein that contains a single MyTH4/FERM
domain. Interestingly, none of the peptides identified in our
analysis matched to a recognizable myosin-15 motor domain
structure. If the myosin-15-like protein expressed in enterocytes is in fact “headless,” it may be playing a regulatory role
by modulating cargo binding of other MyTH4/FERM domaincontaining motors, such as myosin-7b. Future studies will be
needed to further characterize and investigate the functional
consequence of this myosin-15-like protein.
G924
BRUSH BORDER PROTEOME
reduction in the delivery of all glycosylated proteins to the apical
surface (63). The importance of annexins and galectins in apical
transport is underscored by findings suggesting that changes in
their expression levels are contributing factors to a number of
gastrointestinal diseases ranging from inflammation to cancer (21,
81). Recently, it has been shown that multiple types of annexins
and galectins are enriched in vesicles released from the tips of
microvilli, suggesting that these protein families may also function in this novel form of secretion from the brush border (55).
Adhesion Molecules
ACKNOWLEDGEMENTS
The authors thank all members of the Tyska laboratory for advice and
support, Dr. Amy Ham and Dr. David Friedman of the Vanderbilt University
Medical Center (VUMC) Mass Spectrometry Research Center for outstanding
core technical support, and the VUMC Cell Imaging Shared Resource.
GRANTS
This work was supported by National Institutes of Health (NIH) Grants
R01-DK-075555 (to M. J. Tyska) and R01-CA-126218 (to D. L. Tabb),
American Heart Association predoctoral fellowships (to R. E. McConnell and
A. E. Benesh) and Grant 09GRNT2310188 (to M. J. Tyska), the VUMC
Digestive Diseases Research Center (pilot funds from National Institutes of
Health Grant P30-DK-058404, to R. M. Peek), and a Vanderbilt University
Innovation and Discovery in Engineering And Science award (to M. J. Tyska).
DISCLOSURES
No conflicts of interest, financial or otherwise, are declared by the author(s).
REFERENCES
Concluding Remarks
This report is the first to describe the full proteome of the
enterocyte brush border. As such, we expect these results to
provide a rich source of data that will stimulate the development of new hypotheses and studies on the assembly, maintenance, and function of the enterocyte apical domain. Future
studies must focus on validating the novel identifications made
AJP-Gastrointest Liver Physiol • VOL
1. Adkins JN, Varnum SM, Auberry KJ, Moore RJ, Angell NH, Smith
RD, Springer DL, Pounds JG. Toward a human blood serum proteome:
analysis by multidimensional separation coupled with mass spectrometry. Mol Cell Proteomics 1: 947–955, 2002.
2. Ahuja R, Pinyol R, Reichenbach N, Custer L, Klingensmith J,
Kessels MM, Qualmann B. Cordon-bleu is an actin nucleation factor
and controls neuronal morphology. Cell 131: 337–350, 2007.
3. Aljure O, Diez-Sampedro A. Functional characterization of mouse
sodium/glucose transporter type 3b. Am J Physiol Cell Physiol 299:
C58 –C65, 2010.
4. Ammer AG, Weed SA. Cortactin branches out: roles in regulating
protrusive actin dynamics. Cell Motil Cytoskeleton 65: 687–707, 2008.
5. Babusiak M, Man P, Petrak J, Vyoral D. Native proteomic analysis of
protein complexes in murine intestinal brush border membranes. Proteomics 7: 121–129, 2007.
6. Bahler M, Kroschewski R, Stoffler HE, Behrmann T. Rat myr 4
defines a novel subclass of myosin I: identification, distribution, localization, and mapping of calmodulin-binding sites with differential calcium sensitivity. J Cell Biol 126: 375–389, 1994.
7. Bahler M, Rhoads A. Calmodulin signaling via the IQ motif. FEBS Lett
513: 107–113, 2002.
8. Bement WM, Hasson T, Wirth JA, Cheney RE, Mooseker MS. Identification and overlapping expression of multiple unconventional myosin
genes in vertebrate cell types. Proc Natl Acad Sci USA 91: 11767, 1994.
9. Benesh AE, Nambiar R, McConnell RE, Mao S, Tabb DL, Tyska MJ.
Differential localization and dynamics of class I myosins in the enterocyte microvillus. Mol Biol Cell 21: 970 –978, 2010.
10. Berndorff D, Gessner R, Kreft B, Schnoy N, Lajous-Petter AM, Loch
N, Reutter W, Hortsch M, Tauber R. Liver-intestine cadherin: molecular cloning and characterization of a novel Ca2⫹-dependent cell adhesion molecule expressed in liver and intestine. J Cell Biol 125: 1353–
1369, 1994.
11. Bertling E, Hotulainen P, Mattila PK, Matilainen T, Salminen M,
Lappalainen P. Cyclase-associated protein 1 (CAP1) promotes cofilininduced actin dynamics in mammalian nonmuscle cells. Mol Biol Cell
15: 2324 –2334, 2004.
12. Bitner-Glindzicz M, Lindley KJ, Rutland P, Blaydon D, Smith VV,
Milla PJ, Hussain K, Furth-Lavi J, Cosgrove KE, Shepherd RM,
Barnes PD, O’Brien RE, Farndon PA, Sowden J, Liu XZ, Scanlan MJ,
Malcolm S, Dunne MJ, Aynsley-Green A, Glaser B. A recessive contiguous gene deletion causing infantile hyperinsulinism, enteropathy and deafness identifies the Usher type 1C gene. Nat Genet 26: 56 –60, 2000.
13. Boeda B, El-Amraoui A, Bahloul A, Goodyear R, Daviet L,
Blanchard S, Perfettini I, Fath KR, Shorte S, Reiners J, Houdusse A,
Legrain P, Wolfrum U, Richardson G, Petit C. Myosin VIIa, harmonin and cadherin 23, three Usher I gene products that cooperate to
shape the sensory hair cell bundle. EMBO J 21: 6689 –6699, 2002.
14. Bretscher A, Weber K. Villin: the major microfilament-associated
protein of the intestinal microvillus. Proc Natl Acad Sci USA 76:
2321–2325, 1979.
300 • MAY 2011 •
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One of the most surprising aspects of this analysis relates to
the presence of putative extracellular adhesion molecules in the
brush border. Although some of the adhesion molecules identified here are well-established players in junctional adhesion
(cadherins-1 and -17 and ZO-1, -2, and -3), others, such as
MLPCDH and PCDH24, are poorly characterized. Nevertheless, staining of CACO-2BBE cells showed robust labeling for
both MLPCDH and PCDH24 isoforms in microvilli (Fig. 4).
Although little is known about these molecules, both are type
I transmembrane proteins and contain varying numbers of
extracellular “EC” repeats, which are characteristic of the
cadherin protein superfamily (eight for PCDH24 and four for
MLPCDH) (28, 29, 68). The isoform 2 variant of MLPCDH
identified here also contains a juxtamembrane mucin-like domain with tandem repeats of threonine, serine, and proline (28,
29). Transfection of cultured cells with MLPCDH increased
aggregation in a Ca2⫹-dependent manner, suggesting that this
molecule is capable of forming bona fide adhesion complexes
(28). Although the role of these PCDHs in brush border
function remains to be established, an intriguing possibility is
that MLPCDH and/or PCDH24 create adhesion between microvilli, contributing to the tight packing of these protrusions
observed in fully differentiated enterocytes.
The presence of adhesion molecules in the brush border is
reminiscent of hair bundle architecture on the apical surface of
mechanosensory hair cells. In the cochlea, hair bundles are
clusters of stereocilia that are organized into three rows of
graded height (30). Adjacent stereocilia are connected between
rows by thread-like links composed of other cadherin superfamily members, PCDH-15 and cadherin-23 (75). These links
are also connected to ion channels so that stereocilia deflections give rise to transduction currents. By analogy, brush
border microvilli may also build physical links between adjacent protrusions, which could facilitate brush border function
or assembly. Interestingly, the brush border proteome also included harmonin, taperin, and clarin, three proteins that are expressed in hair cells and play significant roles in the mechanotransduction process (26, 73, 92). The physiological importance of
these proteins in the context of the brush border is further suggested by the finding that patients harboring mutations in harmonin frequently suffer from inflammatory enteropathy (12).
here and probing functions of important molecules not previously known to reside in the brush border.
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AJP-Gastrointest Liver Physiol • VOL
37. Isogawa Y, Kon T, Inoue T, Ohkura R, Yamakawa H, Ohara O,
Sutoh K. The N-terminal domain of MYO18A has an ATP-insensitive
actin-binding site. Biochemistry 44: 6190 –6196, 2005.
38. Jemnitz K, Heredi-Szabo K, Janossy J, Ioja E, Vereczkey L, Krajcsi
P. ABCC2/Abcc2: a multispecific transporter with dominant excretory
functions. Drug Metab Rev 42: 402–436, 2010.
39. Kenny AJ, Booth AG, Macnair RD. Peptidases of the kidney microvillus membrane. Acta Biol Med Ger 36: 1575–1585, 1977.
40. Kenny AJ, Maroux S. Topology of microvillar membrance hydrolases
of kidney and intestine. Physiol Rev 62: 91–128, 1982.
41. Kessner D, Chambers M, Burke R, Agus D, Mallick P. ProteoWizard:
open source software for rapid proteomics tools development. Bioinformatics 24: 2534 –2536, 2008.
42. Khundmiri SJ, Weinman EJ, Steplock D, Cole J, Ahmad A, Baumann PD, Barati M, Rane MJ, Lederer E. Parathyroid hormone
regulation of Na⫹,K⫹-ATPase requires the PDZ 1 domain of sodium
hydrogen exchanger regulatory factor-1 in opossum kidney cells. J Am
Soc Nephrol 16: 2598 –2607, 2005.
43. Klett EL, Lee MH, Adams DB, Chavin KD, Patel SB. Localization of
ABCG5 and ABCG8 proteins in human liver, gall bladder and intestine.
BMC Gastroenterol 4: 21, 2004.
44. Kovacs M, Thirumurugan K, Knight PJ, Sellers JR. Load-dependent
mechanism of nonmuscle myosin 2. Proc Natl Acad Sci USA 104:
9994 –9999, 2007.
45. Lapierre LA, Avant KM, Caldwell CM, Ham AJ, Hill S, Williams
JA, Smolka AJ, Goldenring JR. Characterization of immunoisolated
human gastric parietal cells tubulovesicles: identification of regulators of
apical recycling. Am J Physiol Gastrointest Liver Physiol 292: G1249 –
G1262, 2007.
46. Law AL, Ling Q, Hajjar KA, Futter CE, Greenwood J, Adamson P,
Wavre-Shapton ST, Moss SE, Hayes MJ. Annexin A2 regulates
phagocytosis of photoreceptor outer segments in the mouse retina. Mol
Biol Cell 20: 3896 –3904, 2009.
47. Lawson JD, Pate E, Rayment I, Yount RG. Molecular dynamics
analysis of structural factors influencing back door pi release in myosin.
Biophys J 86: 3794 –3803, 2004.
48. Li M, Wang W, Soroka CJ, Mennone A, Harry K, Weinman EJ,
Boyer JL. NHERF-1 binds to Mrp2 and regulates hepatic Mrp2 expression and function. J Biol Chem 285: 19299 –19307, 2010.
49. Licklider LJ, Thoreen CC, Peng J, Gygi SP. Automation of nanoscale
microcapillary liquid chromatography-tandem mass spectrometry with a
vented column. Anal Chem 74: 3076 –3083, 2002.
50. Link AJ, Eng J, Schieltz DM, Carmack E, Mize GJ, Morris DR,
Garvik BM, Yates JR, 3rd. Direct analysis of protein complexes using
mass spectrometry. Nat Biotechnol 17: 676 –682., 1999.
51. Lowey S, Trybus KM. Common structural motifs for the regulation of
divergent class II myosins. J Biol Chem 285: 16403–16407, 2010.
52. Ma ZQ, Dasari S, Chambers MC, Litton MD, Sobecki SM, Zimmerman LJ, Halvey PJ, Schilling B, Drake PM, Gibson BW, Tabb DL.
IDPicker 2.0: improved protein assembly with high discrimination peptide identification filtering. J Proteome Res 8: 3872–3881, 2009.
53. Marchiando AM, Graham WV, Turner JR. Epithelial barriers in
homeostasis and disease. Annu Rev Pathol 5: 119 –144, 2010.
54. Massey-Harroche D, Mayran N, Maroux S. Polarized localizations of
annexins I, II, VI and XIII in epithelial cells of intestinal, hepatic and
pancreatic tissues. J Cell Sci 111: 3007–3015, 1998.
55. McConnell RE, Higginbotham JN, Shifrin DA Jr, Tabb DL, Coffey
RJ, Tyska MJ. The enterocyte microvillus is a vesicle-generating
organelle. J Cell Biol 185: 1285–1298, 2009.
56. McConnell RE, Tyska MJ. Myosin-1a powers the sliding of apical membrane along microvillar actin bundles. J Cell Biol 177: 671–681, 2007.
57. Merrifield CJ, Rescher U, Almers W, Proust J, Gerke V, Sechi AS,
Moss SE. Annexin 2 has an essential role in actin-based macropinocytic
rocketing. Curr Biol 11: 1136 –1141, 2001.
58. Millard TH, Dawson J, Machesky LM. Characterisation of IRTKS, a
novel IRSp53/MIM family actin regulator with distinct filament bundling
properties. J Cell Sci 120: 1663–1672, 2007.
59. Miller D, Crane RK. A procedure for the isolation of the epithelial brush
border membrane of hamster small intestine. Anal Biochem 2: 284 –286,
1961.
60. Mitschke D, Reichel A, Fricker G, Moenning U. Characterization of
cytochrome P450 protein expression along the entire length of the intestine
of male and female rats. Drug Metab Dispos 36: 1039 –1045, 2008.
300 • MAY 2011 •
www.ajpgi.org
Downloaded from http://ajpgi.physiology.org/ by 10.220.33.1 on June 15, 2017
15. Chen ZY, Hasson T, Zhang DS, Schwender BJ, Derfler BH,
Mooseker MS, Corey DP. Myosin-VIIb, a novel unconventional myosin, is a constituent of microvilli in transporting epithelia. Genomics 72:
285–296, 2001.
16. Croce A, Cassata G, Disanza A, Gagliani MC, Tacchetti C, Malabarba MG, Carlier MF, Scita G, Baumeister R, Di Fiore PP. A novel
actin barbed-end-capping activity in EPS-8 regulates apical morphogenesis in intestinal cells of Caenorhabditis elegans. Nat Cell Biol 6:
1173–1179, 2004.
17. Cutillas PR, Biber J, Marks J, Jacob R, Stieger B, Cramer R,
Waterfield M, Burlingame AL, Unwin RJ. Proteomic analysis of
plasma membrane vesicles isolated from the rat renal cortex. Proteomics
5: 101–112, 2005.
18. Danielsen EM, Hansen GH. Lipid raft organization and function in the
small intestinal brush border. J Physiol Biochem 64: 377–382, 2008.
19. Danielsen EM, Hansen GH. Lipid rafts in epithelial brush borders:
atypical membrane microdomains with specialized functions. Biochim
Biophys Acta 1617: 1–9, 2003.
20. Delacour D, Koch A, Jacob R. The role of galectins in protein
trafficking. Traffic 10: 1405–1413, 2009.
21. Demetter P, Nagy N, Martin B, Mathieu A, Dumont P, Decaestecker
C, Salmon I. The galectin family and digestive disease. J Pathol 215:
1–12, 2008.
22. Disanza A, Carlier MF, Stradal TE, Didry D, Frittoli E, Confalonieri
S, Croce A, Wehland J, Di Fiore PP, Scita G. Eps8 controls actinbased motility by capping the barbed ends of actin filaments. Nat Cell
Biol 6: 1180 –1188, 2004.
23. Donowitz M, Singh S, Salahuddin FF, Hogema BM, Chen Y, Gucek
M, Cole RN, Ham A, Zachos NC, Kovbasnjuk O, Lapierre LA,
Broere N, Goldenring J, deJonge H, Li X. Proteome of murine jejunal
brush border membrane vesicles. J Proteome Res 6: 4068 –4079, 2007.
24. Fan X, Krahling S, Smith D, Williamson P, Schlegel RA. Macrophage
surface expression of annexins I and II in the phagocytosis of apoptotic
lymphocytes. Mol Biol Cell 15: 2863–2872, 2004.
25. Foth BJ, Goedecke MC, Soldati D. New insights into myosin evolution
and classification. Proc Natl Acad Sci USA 103: 3681–3686, 2006.
26. Geng R, Geller SF, Hayashi T, Ray CA, Reh TA, BerminghamMcDonogh O, Jones SM, Wright CG, Melki S, Imanishi Y, Palczewski K, Alagramam KN, Flannery JG. Usher syndrome IIIA gene
clarin-1 is essential for hair cell function and associated neural activation.
Hum Mol Genet 18: 2748 –2760, 2009.
27. Gerke V, Creutz CE, Moss SE. Annexins: linking Ca2⫹ signalling to
membrane dynamics. Nat Rev Mol Cell Biol 6: 449 –461, 2005.
28. Goldberg M, Peshkovsky C, Shifteh A, Al-Awqati Q. ␮-Protocadherin, a novel developmentally regulated protocadherin with mucin-like
domains. J Biol Chem 275: 24622–24629, 2000.
29. Goldberg M, Wei M, Tycko B, Falikovich I, Warburton D. Identification
and expression analysis of the human ␮-protocadherin gene in fetal and adult
kidneys. Am J Physiol Renal Physiol 283: F454 –F463, 2002.
30. Grillet N, Kazmierczak P, Xiong W, Schwander M, Reynolds A,
Sakaguchi H, Tokita J, Kachar B, Muller U. The mechanotransduction
machinery of hair cells. Sci Signal 2: pt5, 2009.
31. Grimm-Gunter EM, Revenu C, Ramos S, Hurbain I, Smyth N,
Ferrary E, Louvard D, Robine S, Rivero F. Plastin 1 binds to keratin
and is required for terminal web assembly in the intestinal epithelium.
Mol Biol Cell 20: 2549 –2562, 2009.
32. Hauri HP, Quaroni A, Isselbacher KJ. Biogenesis of intestinal plasma
membrane: posttranslational route and cleavage of sucrase-isomaltase.
Proc Natl Acad Sci USA 76: 5183–5186, 1979.
33. Hayes MJ, Shao DM, Grieve A, Levine T, Bailly M, Moss SE.
Annexin A2 at the interface between F-actin and membranes enriched in
phosphatidylinositol 4,5,-bisphosphate. Biochim Biophys Acta 1793:
1086 –1095, 2009.
34. Heintzelman M, Hasson T, Mooseker M. Multiple unconventional
myosin domains of the intestinal brush border cytoskeleton. J Cell Sci
107: 3535–3543, 1994.
35. Hermiston ML, Gordon JI. In vivo analysis of cadherin function in the
mouse intestinal epithelium: essential roles in adhesion, maintenance of
differentiation, and regulation of programmed cell death. J Cell Biol 129:
489 –506, 1995.
36. Hirokawa N, Tilney LG, Fujiwara K, Heuser JE. Organization of
actin, myosin, and intermediate filaments in the brush border of intestinal
epithelial cells. J Cell Biol 94: 425–443, 1982.
G926
BRUSH BORDER PROTEOME
84.
85.
86.
87.
88.
89.
90.
91.
92.
93.
94.
95.
96.
97.
98.
99.
100.
101.
102.
103.
AJP-Gastrointest Liver Physiol • VOL
isoforms identify distinct functional modules that support integrity of the
epithelial zonula adherens. Nat Cell Biol 12: 696 –702, 2010.
Tabb DL, Fernando CG, Chambers MC. MyriMatch: highly accurate
tandem mass spectral peptide identification by multivariate hypergeometric analysis. J Proteome Res 6: 654 –661, 2007.
Taylor KA. Regulation and recycling of myosin V. Curr Opin Cell Biol
19: 67–74, 2007.
Tocchetti A, Soppo CB, Zani F, Bianchi F, Gagliani MC, Pozzi B,
Rozman J, Elvert R, Ehrhardt N, Rathkolb B, Moerth C, Horsch M,
Fuchs H, Gailus-Durner V, Beckers J, Klingenspor M, Wolf E, Hrabe
de Angelis M, Scanziani E, Tacchetti C, Scita G, Di Fiore PP, Offenhauser N. Loss of the actin remodeler Eps8 causes intestinal defects and
improved metabolic status in mice. PLoS One 5: e9468, 2010.
Triphan X, Menzel VA, Petrunkina AM, Cassara MC, Wemheuer
W, Hinsch KD, Hinsch E. Localisation and function of voltage-dependent anion channels (VDAC) in bovine spermatozoa. Pflügers Arch 455:
677–686, 2008.
Turner JR, Rill BK, Carlson SL, Carnes D, Kerner R, Mrsny RJ,
Madara JL. Physiological regulation of epithelial tight junctions is
associated with myosin light-chain phosphorylation. Am J Physiol Cell
Physiol 273: C1378 –C1385, 1997.
Tyska MJ, Mackey AT, Huang JD, Copeland NG, Jenkins NA,
Mooseker MS. Myosin-1a is critical for normal brush border structure
and composition. Mol Biol Cell 16: 2443–2457, 2005.
Tyska MJ, Mooseker MS. MYO1A (brush border myosin I) dynamics in
the brush border of LLC-PK1-CL4 cells. Biophys J 82: 1869 –1883, 2002.
Vachon PH, Perreault N, Magny P, Beaulieu JF. Uncoordinated,
transient mosaic patterns of intestinal hydrolase expression in differentiating human enterocytes. J Cell Physiol 166: 198 –207, 1996.
Verpy E, Leibovici M, Zwaenepoel I, Liu XZ, Gal A, Salem N,
Mansour A, Blanchard S, Kobayashi I, Keats BJ, Slim R, Petit C. A
defect in harmonin, a PDZ domain-containing protein expressed in the
inner ear sensory hair cells, underlies Usher syndrome type 1C. Nat
Genet 26: 51–55, 2000.
Vingadassalom D, Kazlauskas A, Skehan B, Cheng HC, Magoun L,
Robbins D, Rosen MK, Saksela K, Leong JM. Insulin receptor tyrosine
kinase substrate links the E. coli O157:H7 actin assembly effectors Tir
and EspF(U) during pedestal formation. Proc Natl Acad Sci USA 106:
6754 –6759, 2009.
Walmsley SJ, Broeckling C, Hess A, Prenni J, Curthoys NP. Proteomic
analysis of brush-border membrane vesicles isolated from purified proximal
convoluted tubules. Am J Physiol Renal Physiol 298: F1323–F1331, 2010.
Washburn MP, Wolters D, Yates JR, 3rd. Large-scale analysis of the
yeast proteome by multidimensional protein identification technology.
Nat Biotechnol 19: 242–247., 2001.
Weiss SM, Ladwein M, Schmidt D, Ehinger J, Lommel S, Stading K,
Beutling U, Disanza A, Frank R, Jansch L, Scita G, Gunzer F,
Rottner K, Stradal TE. IRSp53 links the enterohemorrhagic E. coli
effectors Tir and EspFU for actin pedestal formation. Cell Host Microbe
5: 244 –258, 2009.
Wetterau JR, Aggerbeck LP, Bouma ME, Eisenberg C, Munck A,
Hermier M, Schmitz J, Gay G, Rader DJ, Gregg RE. Absence of
microsomal triglyceride transfer protein in individuals with abetalipoproteinemia. Science 258: 999 –1001, 1992.
Wolters DA, Washburn MP, Yates JR 3rd. An automated multidimensional protein identification technology for shotgun proteomics. Anal
Chem 73: 5683–5690, 2001.
Yang C, Hoelzle M, Disanza A, Scita G, Svitkina T. Coordination of
membrane and actin cytoskeleton dynamics during filopodia protrusion.
PLoS One 4: e5678, 2009.
Yang LE, Maunsbach AB, Leong PK, McDonough AA. Redistribution of myosin VI from top to base of proximal tubule microvilli during
acute hypertension. J Am Soc Nephrol 16: 2890 –2896, 2005.
Yengo CM, Ananthanarayanan SK, Brosey CA, Mao S, Tyska MJ.
Human deafness mutation E385D disrupts the mechanochemical coupling and subcellular targeting of myosin-1a. Biophys J 94: L5–L7, 2008.
Yu D, Marchiando AM, Weber CR, Raleigh DR, Wang Y, Shen L,
Turner JR. MLCK-dependent exchange and actin binding region-dependent anchoring of ZO-1 regulate tight junction barrier function. Proc
Natl Acad Sci USA 107: 8237–8241, 2010.
Zachos NC, Li X, Kovbasnjuk O, Hogema B, Sarker R, Lee LJ, Li M, de
Jonge H, Donowitz M. NHERF3 (PDZK1) contributes to basal and calcium
inhibition of NHE3 activity in Caco-2BBe cells. J Biol Chem 284: 23708–
23718, 2009.
300 • MAY 2011 •
www.ajpgi.org
Downloaded from http://ajpgi.physiology.org/ by 10.220.33.1 on June 15, 2017
61. Mooseker MS. Organization, chemistry, and assembly of the cytoskeletal apparatus of the intestinal brush border. Annu Rev Cell Biol 1:
209 –241, 1985.
62. Mooseker MS, Pollard TD, Fujiwara K. Characterization and localization of myosin in the brush border of intestinal epithelial cells. J Cell
Biol 79: 444 –453, 1978.
63. Morelle W, Stechly L, Andre S, Van Seuningen I, Porchet N, Gabius
HJ, Michalski JC, Huet G. Glycosylation pattern of brush border-associated glycoproteins in enterocyte-like cells: involvement of complex-type
N-glycans in apical trafficking. Biol Chem 390: 529 –544, 2009.
64. Naim HY. Angiotensin-converting enzyme of the human small intestine.
Subunit and quaternary structure, biosynthesis and membrane association. Biochem J 286: 451–457, 1992.
65. Norstrom MF, Smithback PA, Rock RS. Unconventional processive
mechanics of non-muscle myosin IIB. J Biol Chem 285: 26326 –26334,
2010.
66. Offenhauser N, Borgonovo A, Disanza A, Romano P, Ponzanelli I,
Iannolo G, Di Fiore PP, Scita G. The eps8 family of proteins links
growth factor stimulation to actin reorganization generating functional
redundancy in the Ras/Rac pathway. Mol Biol Cell 15: 91–98, 2004.
67. Okada SF, O’Neal WK, Huang P, Nicholas RA, Ostrowski LE,
Craigen WJ, Lazarowski ER, Boucher RC. Voltage-dependent anion
channel-1 (VDAC-1) contributes to ATP release and cell volume regulation in murine cells. J Gen Physiol 124: 513–526, 2004.
68. Okazaki N, Takahashi N, Kojima S, Masuho Y, Koga H. Protocadherin LKC, a new candidate for a tumor suppressor of colon and liver
cancers, its association with contact inhibition of cell proliferation.
Carcinogenesis 23: 1139 –1148, 2002.
69. Paradela A, Bravo SB, Henriquez M, Riquelme G, Gavilanes F,
Gonzalez-Ros JM, Albar JP. Proteomic analysis of apical microvillous
membranes of syncytiotrophoblast cells reveals a high degree of similarity with lipid rafts. J Proteome Res 4: 2435–2441, 2005.
70. Peterson MD, Mooseker MS. Characterization of the enterocyte-like
brush border cytoskeleton of the C2BBe clones of the human intestinal
cell line, Caco-2. J Cell Sci 102: 581–600, 1992.
71. Pollard TD, Borisy GG. Cellular motility driven by assembly and
disassembly of actin filaments. Cell 112: 453–465, 2003.
72. Qualmann B, Kessels MM. New players in actin polymerization–WH2domain-containing actin nucleators. Trends Cell Biol 19: 276 –285, 2009.
73. Rehman AU, Morell RJ, Belyantseva IA, Khan SY, Boger ET,
Shahzad M, Ahmed ZM, Riazuddin S, Khan SN, Friedman TB.
Targeted capture and next-generation sequencing identifies C9orf75,
encoding taperin, as the mutated gene in nonsyndromic deafness
DFNB79. Am J Hum Genet 86: 378 –388, 2010.
74. Rodriguez-Boulan E, Kreitzer G, Musch A. Organization of vesicular
trafficking in epithelia. Nat Rev Mol Cell Biol 6: 233–247, 2005.
75. Sakaguchi H, Tokita J, Muller U, Kachar B. Tip links in hair cells:
molecular composition and role in hearing loss. Curr Opin Otolaryngol
Head Neck Surg 17: 388 –393, 2009.
76. Sato N, Funayama N, Nagafuchi A, Yonemura S, Tsukita S. A gene
family consisting of ezrin, radixin and moesin. Its specific localization at
actin filament/plasma membrane association sites. J Cell Sci 103: 131–
143, 1992.
77. Schreiber R, Uliyakina I, Kongsuphol P, Warth R, Mirza M, Martins
JR, Kunzelmann K. Expression and function of epithelial anoctamins. J
Biol Chem 285: 7838 –7845, 2010.
78. Shaner NC, Sanger JW, Sanger JM. Actin and ␣-actinin dynamics in
the adhesion and motility of EPEC and EHEC on host cells. Cell Motil
Cytoskeleton 60: 104 –120, 2005.
79. Shaw RK, Cleary J, Murphy MS, Frankel G, Knutton S. Interaction
of enteropathogenic Escherichia coli with human intestinal mucosa: role
of effector proteins in brush border remodeling and formation of attaching and effacing lesions. Infect Immun 73: 1243–1251, 2005.
80. Shin JB, Streijger F, Beynon A, Peters T, Gadzala L, McMillen D,
Bystrom C, Van der Zee CE, Wallimann T, Gillespie PG. Hair
bundles are specialized for ATP delivery via creatine kinase. Neuron 53:
371–386, 2007.
81. Singh P. Role of annexin-II in GI cancers: interaction with gastrins/
progastrins. Cancer Lett 252: 19 –35, 2007.
82. Skowron JF, Bement WM, Mooseker MS. Human brush border myosin-I and myosin-Ic expression in human intestine and Caco-2BBe cells.
Cell Motil Cytoskeleton 41: 308 –324, 1998.
83. Smutny M, Cox HL, Leerberg JM, Kovacs EM, Conti MA, Ferguson
C, Hamilton NA, Parton RG, Adelstein RS, Yap AS. Myosin II