Department of Cell and Molecular Biology, Northwestern University Medical School, Chicago, Illinois 60611
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Abstract |
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An ~30-kD isoform of the actin-binding/
bundling protein espin has been discovered in the brush
borders of absorptive epithelial cells in rat intestine and
kidney. Small espin is identical in sequence to the
COOH terminus of the larger (~110-kD) espin isoform identified in the actin bundles of Sertoli cell-spermatid
junctional plaques (Bartles, J.R., A. Wierda, and
L. Zheng. 1996. J. Cell Sci. 109:1229-1239), but it contains two unique peptides at its NH2 terminus. Small espin was localized to the parallel actin bundles of brush
border microvilli, resisted extraction with Triton X-100,
and accumulated in the brush border during enterocyte
differentiation/migration along the crypt-villus axis in
adults. In transfected BHK fibroblasts, green fluorescent protein-small espin decorated F-actin-containing fibers and appeared to elicit their accumulation and/or
bundling. Recombinant small espin bound to skeletal
muscle and nonmuscle F-actin with high affinity (Kd = 150 and 50 nM) and cross-linked the filaments into bundles. Sedimentation, gel filtration, and circular dichroism analyses suggested that recombinant small espin was a monomer with an asymmetrical shape and a high
percentage of -helix. Deletion mutagenesis suggested
that small espin contained two actin-binding sites in its
COOH-terminal 116-amino acid peptide and that the
NH2-terminal half of its forked homology peptide was
necessary for bundling activity.
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Introduction |
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THE ability of actin to form filaments and higher-order cross-linked structures is harnessed by cells in
a variety of ways to make the cortical actin cytoskeleton a central determinant of cellular shape, division,
motility, adhesion, and signaling. These feats are accomplished in large part through the activities of a host of different actin-binding proteins, which among other things:
(a) regulate the polymerization and depolymerization of actin; (b) connect actin-based structures to membranes or
other cytoskeletal systems; (c) power movement relative
to actin filaments; or (d) cross-link F-actin into meshworks
or bundles (Pollard, 1993; Ridley, 1995
; Burridge and Chrzanowska-Wodnicka, 1996
; Gumbiner, 1996
; Mitchison
and Cramer, 1996
; Bretscher et al., 1997
; Carlier, 1997
;
Hall, 1998
; Mermall et al., 1998
).
Actin bundles can be either of mixed polarity (contractile bundles) or of uniform polarity (parallel bundles)
(Matsudaira, 1991; Pollard, 1993
; Furukawa and Fechheimer, 1997
). Parallel bundles are commonly associated
with relatively long-lived specializations of the cell surface,
where they appear to function in part as a scaffold to support evaginations or invaginations of the plasma membrane. Examples include: the brush border microvilli of
absorptive epithelia (Heintzelman and Mooseker, 1992
;
Fath and Burgess, 1995
), the stereocilia of hair cells in the
inner ear (Tilney et al., 1992
), the bristles of Drosophila
pupae (Tilney et al., 1995
), and the submembranous
plaques of Sertoli cell-spermatid and Sertoli cell-Sertoli
cell junctions known as ectoplasmic specializations (Russell and Peterson, 1985
; Vogl, 1989
; Bartles et al., 1996
). There is evidence that each of these structures contains its
own complement of actin-bundling proteins. In three of
these cases, the structure in question is known to contain
at least two actin-bundling proteins that appear to function sequentially during the process of assembly: in brush
border microvilli, villin and fimbrin/plastin (Rodman et
al., 1986
; Ezzell et al., 1989
; Heintzelman and Mooseker, 1992
; Fath and Burgess, 1995
); in hair cell stereocilia, some yet-to-be identified protein (not villin) and fimbrin/plastin (Tilney et al., 1992
); and in the bristles of Drosophila pupae, the proteins encoded by the forked and singed genes
(the latter a fascin ortholog) (Tilney et al., 1995
).
Actin cross-linking proteins typically display a modular
organization (Matsudaira, 1991; Pollard, 1993
; Puius et al.,
1998
). The ability to cross-link actin filaments requires
that a protein contain two or more actin-binding sites per
monomer or that it exist as a dimer or higher oligomer.
The spacing between the actin-binding sites appears to influence the type of cross-linked structure formed. Proteins
that cross-link actin filaments to form loose meshworks
are typically larger and achieve relatively distant separations between their actin-binding sites through the use of
extended spacer peptides and/or through the formation of
dimers or higher oligomers. In contrast, proteins that
cross-link filaments into bundles are generally smaller in
size and have tandem actin-binding sites spaced relatively
close together. For example, the known actin-bundling
proteins of brush border microvilli, fimbrin/plastin and villin, are both monomeric and contain tandem actin-binding sites. Fimbrin/plastin contains two tandemly repeated 27-kD actin-binding domains of the
-actinin/
-spectrin type,
each of which in turn is composed of two calponin-homology domains (Matsudaira, 1991
; Goldsmith et al., 1997
).
Villin contains a "core" domain, which displays Ca2+-mediated F-actin severing activity and is made up of six repeats of a domain shared with other severing proteins (such as
gelsolin, fragmin, and severin) and a unique thermostable
COOH-terminal "headpiece," which is required for actin-bundling activity (Friederich et al., 1990
; McKnight et al.,
1996
, 1997
). In addition to containing modules for binding
to actin, actin-bundling proteins frequently contain other
modules that can regulate bundling activity. For example,
both fimbrin and villin contain binding sites for Ca2+, and
the actin-bundling activity of both proteins is decreased dramatically by physiological concentrations of Ca2+
(Glenney et al., 1981
; Alicea and Mooseker, 1988
; Namba
et al., 1992
; Lin et al., 1994
). In fact, villin becomes an
F-actin-severing protein in the presence of Ca2+ (Walsh et
al., 1984
).
We recently identified espin as a novel ~110-kD actin-binding protein localized to the parallel actin bundles in
the junctional plaque of Sertoli cell ectoplasmic specializations (Bartles et al., 1996). Upon sequence analysis, espin
was found to contain eight ankyrin-like repeats in its NH2-terminal third, a potential P-loop, two proline-rich peptides, and two peptides that contained clusters of multiple
glutamates bracketed by arginines, lysines, and glutamines
in a pattern reminiscent of the repetitive motif found in
the protein trichohyalin. The ankyrin-like repeats and a
66-amino acid peptide present near the COOH terminus
of espin showed 35-39% sequence identity to proteins encoded by the forked gene of Drosophila. A maltose-binding protein fusion protein containing the COOH-terminal
379 amino acids of espin was found to bind with high affinity to F-actin in vitro. And, when expressed by transfected NRK fibroblasts, the same COOH-terminal fragment of
espin was observed to decorate actin fibers or cables and
appeared to bring about their accumulation and/or bundling. On the basis of its structure, localization, and properties, we hypothesized that espin was involved in linking
actin filaments to each other and/or to membranes,
thereby potentially playing a key role in the organization and function of the ectoplasmic junctional specializations
of Sertoli cells.
The ~110-kD espin of Sertoli cell ectoplasmic specializations was found to be encoded by an ~2.9-kb mRNA
that, on Northern blots of total RNA, appeared specific to
testis among 11 rat tissues examined (Bartles et al., 1996).
At that time, we also detected crosshybridizing mRNAs of
~1.7 kb in rat small intestine and kidney. Here we report
that the ~1.7-kb crosshybridizing mRNAs encode a second member of the espin family, a smaller isoform that
displays a number of the properties expected for a newly
identified actin-bundling protein of brush border microvilli.
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Materials and Methods |
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Homogenates (~4%, wt/vol) were prepared from the testis, kidney, and
small intestinal mucosal scrapings of adult rats in 0.25 M sucrose, 3 mM
imidazole-HCl, pH 7.4, containing protease inhibitors (1 mM PMSF, 1 µg/
ml each of antipain and leupeptin, and 10-20 TIU/liter of aprotinin) using
eight up-and-down strokes of a motor-driven Teflon-glass homogenizer
(3,000 rpm). Brush borders were isolated from the small intestines of adult
rats in the presence of the protease inhibitors using the procedure of
Keller and Mooseker (1982). In some experiments, isolated brush borders
were extracted for 45 min at 4°C with solution A (75 mM KCl, 5 mM
MgCl2, 1 mM EGTA, 10 mM imidazole-HCl, pH 7.2; Keller and Mooseker, 1982
) containing the protease inhibitors in the presence or absence of
1% (vol/vol) Triton X-100 or 0.6 M KI before preparing a supernate and
pellet fraction by centrifugation at 100,000 g for 60 min. Samples for
PAGE and Western blotting were prepared from these various fractions
by adding concentrated SDS gel sample buffer containing dithiothreitol
and boiling for 3 min. Espin was detected by Western blotting using affinity-purified rabbit polyclonal antibodies directed against the COOH-terminal 379-amino acid peptide of the ~110-kD rat Sertoli cell espin isoform followed by 125I-protein A (Bartles et al., 1996
). The amount of
espin present in isolated small intestinal brush borders was determined by
scanning laser densitometric analysis of Western blot autoradiograms and comparison to a standard curve generated from internal standards containing known amounts of recombinant small espin (see below). The
amount of actin present in the isolated small intestinal brush borders was
determined by scanning laser densitometric analysis of Coomassie blue-
stained SDS gels and comparison to a standard curve generated from internal standards containing known amounts of rabbit skeletal muscle actin
(Cytoskeleton, Inc., Denver, CO).
A cDNA (1,088-bp) that included the full coding sequence of the small
isoform of espin was obtained using the affinity-purified rabbit polyclonal
antibodies to screen a 5'-STRETCH rat kidney gt11 cDNA library
(CLONTECH Laboratories, Palo Alto, CA). cDNAs were introduced
into the pBluescript SK phagemid vector (Stratagene, La Jolla, CA), and
the coding and noncoding strands were sequenced in their entirety using
[35S]dATP
S and the Sequenase Version 2.0 kit (Amersham Corp., Arlington Heights, IL). Sequence analysis and database searching were carried out using the programs available through the University of Wisconsin Genetics Computer Group (GCG) Sequence Analysis Software Package (Devereux et al., 1984
) and the BLAST Search (Altschul et al., 1990
)
available through the National Center for Biotechnology Information.
Peptides with a high probability to form a coiled coil were identified using
the programs of Lupas (1997)
and Berger et al. (1995)
.
For immunoperoxidase labeling, rat intestine and kidney were fixed by
perfusion through the abdominal aorta with Bouin's fluid and embedded
in paraffin (Bartles et al., 1996). 5-µm sections were deparaffinized with
xylenes, labeled with affinity-purified rabbit polyclonal espin antibody or
preimmune IgG followed by horseradish peroxidase-conjugated donkey
anti-rabbit F(ab')2, reacted with H2O2 and 3,3'-diaminobenzidine, and
counterstained with hematoxylin (Bartles et al., 1996
). In preparation for
immunofluorescence, isolated small intestinal brush borders were fixed to
poly-L-lysine-coated slides or coverslips with 2% paraformaldehyde in solution A containing 20 mM Hepes, pH 7.2, and 20 mM n-octyl-
-D-glucopyranoside, quenched with 0.25% (wt/vol) NH4Cl and 0.5% (wt/vol)
BSA and labeled with affinity-purified polyclonal espin antibody or preimmune IgG and fluorescein-labeled phalloidin (Molecular Probes, Eugene, OR) followed by rhodamine-labeled goat anti-rabbit IgG (Jackson
ImmunoResearch Laboratories, Inc., West Grove, PA) in PBS containing
0.02% (wt/vol) saponin (Bartles et al., 1996
). In preparation for postembedment immunogold labeling, freshly isolated rat small intestinal brush borders were washed twice by low-speed centrifugation in 60 mM Pipes,
25 mM Hepes, 10 mM EGTA, 2 mM MgCl2, pH 6.9, containing the protease inhibitors, fixed for 30 min at 4°C in the same buffer containing 4%
paraformaldehyde, pelleted by centrifugation at 13,000 g for 20 min, post-fixed as a pellet for an additional 1.5 h at 4°C, washed overnight with two
changes of the same buffer minus paraformaldehyde, dehydrated through
30, 50, and 70% ethanol at
20°C, infiltrated overnight with LR white
(Electron Microscopy Sciences, Ft. Washington, PA) at 4°C with eight
changes, placed in gelatin capsules, and then polymerized for 3-5 d at 38-
40°C. Ultrathin sections were quenched with 0.5% (wt/vol) BSA in Tris-buffered saline, pH 7.4, and labeled for 15 h with affinity-purified espin antibody or preimmune IgG followed by goat anti-rabbit IgG 12- or
6-nm-diameter colloidal gold conjugate (Jackson ImmunoResearch Laboratories) and counterstained with 2-3% (wt/vol) uranyl acetate. To minimize aggregation of the gold conjugate, the tracer was blended thoroughly
on a vortex mixer and then precleared by centrifugation at 13,000 g for 5 min immediately before use. For preembedment immunogold labeling,
isolated rat small intestinal brush borders were fixed in 4% paraformaldehyde in solution A containing 20 mM Hepes, pH 7.2, and 0.5% (vol/vol)
Triton X-100, quenched with 0.25% (wt/vol) NH4Cl and 0.5% (wt/vol)
BSA, and labeled in suspension with affinity-purified espin antibody or
preimmune IgG followed by colloidal gold-labeled secondary antibody
(see above). In between steps, the brush borders were washed by low-speed centrifugation. After the final wash, the labeled brush borders were
postfixed in 2% glutaraldehyde in 0.1 M sodium cacodylate, pH 7.4, postfixed in 1% OsO4, stained en bloc with uranyl acetate, dehydrated in ethanol and propylene oxide, and embedded in Poly/Bed 812 (Polysciences, Warrington, PA). Ultrathin sections were stained with uranyl acetate and
lead citrate.
For transient transfection, the 1,088-bp cDNA encoding small espin
was introduced into the EcoRI site of the pEGFP-C2 vector (CLONTECH Laboratories) and used to express a green fluorescent protein
(GFP)1-small espin fusion protein in cells of the BHK fibroblastic line (American Type Culture Collection, Rockville, MD) by transient transfection with lipofectamine (Life Technologies, Gaithersburg, MD). The
cells were cultured on coverslips in DME containing 10% (vol/vol) calf serum and penicillin/streptomycin. The GFP-small espin fusion protein was
localized in these cells by conventional fluorescence microscopy, either
without fixation or after fixation with 2% paraformaldehyde in PBS, pH
7.4, extraction for 5 min with ice-cold 0.1% (vol/vol) Triton X-100 in PBS,
and labeling with rhodamine-phalloidin (Molecular Probes). In some
transfection experiments, small espin was expressed without GFP, using
the pcDNA3 expression vector (Invitrogen, Carlsbad, CA), and detected by immunofluorescence (Bartles et al., 1996).
cDNAs encoding the full-length small espin protein and NH2- or
COOH-terminally truncated versions were prepared from the library cDNA clone by PCR using Deep Vent DNA polymerase (New England Biolabs, Beverly, MA) with limiting amounts of template and enzyme. Primers were designed to introduce a BamHI site at the 5' end and an
EcoRI and/or a HindIII site at the 3' end. The cDNAs were introduced
into the BamHI and EcoRI or HindIII sites of the pProEX HT prokaryotic expression vector (Life Technologies) that gave the proper reading
frame. The resulting constructs were checked by automated DNA sequencing (using a BigDye terminator and model 377 sequencer; Applied
Biosystems, Inc., Foster City, CA) and used to transform Escherichia coli
DH5 (Life Technologies). These resulting recombinant proteins included an additional 28 amino acids at their NH2 termini: MSYY, followed by HHHHHH (the 6xHis tag), DYDIPTT (a spacer region), ENLYFQ (Tobacco Etch Virus protease cleavage site), and GAMGS. The
6xHis-tagged small espin constructs were isolated from 50 mM Tris-HCl,
10 mM 2-mercaptoethanol, pH 8.5, extracts of frozen-thawed, sonicated
bacteria by batch affinity chromatography on Ni-NTA resin (Qiagen,
Chatsworth, CA) using the 0.1 M KCl, 10% (vol/vol) glycerol, 10 mM
2-mercaptoethanol, 20 mM Tris-HCl, 20 mM imidazole-HCl, pH 8.5, buffer system recommended by Life Technologies. The 0.2-M imidazole
eluate was dialyzed against 0.1 M KCl, 10 mM imidazole-HCl, 1 mM
NaN3, pH 7.4; treated at a concentration of 0.02-0.05 mg/ml with 15 U/ml
of recombinant 6xHis-tagged Tobacco Etch Virus protease (Life Technologies) for 3 h at 37°C; incubated for 30 min at 4°C with Ni-NTA resin to remove cleaved 6xHis-tag, any uncleaved 6xHis-tagged small espin, and
the residual 6xHis-tagged viral protease; adjusted to 2 mM MgCl2; and
freed of any insoluble protein by centrifugation at 150,000 g for 90 min at
4°C in preparation for use in F-actin binding and bundling assays. After
removal of the 6xHis tag with Tobacco Etch Virus protease, the constructs
retained only an extra five amino acids, GAMGS, at their NH2 terminus.
F-actin was prepared by dilution of purified rabbit skeletal muscle actin
or human platelet nonmuscle actin (85% -actin and 15%
-actin) (Cytoskeleton, Inc.) into 0.1 M KCl, 2 mM MgCl2, 1 mM ATP, 1 mM NaN3,
10 mM imidazole-HCl, pH 7.4, and incubation for 60 min at 37°C. To assay for F-actin binding or bundling, an equal volume of solution containing different amounts of recombinant small espin protein in the same
buffer minus ATP was added to preformed actin filaments (at a final actin
concentration of 0.1-0.5 mg/ml) and incubated for 60 min at 37°C. Samples were either taken for negative staining with 0.5-1% (wt/vol) uranyl
acetate on 300-mesh Formvar and carbon-coated copper grids (Cooper
and Pollard, 1982
) or were centrifuged at 4°C for either 15 min at 22,000 g
(bundling assay; Edwards et al., 1995
) or for 90 min at 150,000 g (binding assay; Bartles et al., 1996
). The levels of small espin protein and actin
present in the supernate and pellet fractions were determined by scanning
laser densitometric analysis of Coomassie blue-stained SDS gels using the
rabbit skeletal muscle actin as the protein standard.
For sedimentation equilibrium, the purified recombinant espin was dialyzed against 0.1 M KCl, 0.02 M Tris-HCl, 2 mM MgCl2, 1 mM NaN3, pH 7.4, and centrifuged for 22-24 h at 15,000 rpm in an analytical ultracentrifuge (model XLA-1; Beckman Instruments, Fullerton, CA). The molecular mass was estimated from plots of log A280 versus radius squared. The modeling of molecular dimensions (semimajor [a] and semiminor [b] axes multiplied by 2) and sedimentation coefficient using subunit molecular mass and frictional coefficient ratio for three basic asymmetrical shapes (prolate ellipsoid, oblate ellipsoid, and long rod) were carried out using the XL-A-UltraScan-Origin Software Package Version 2.93 produced by Dr. Borries Demeler (Missoula, MT) and supplied with the instrument. To measure sedimentation coefficient, the purified espin was centrifuged for 3 d at 25,000 rpm under isokinetic conditions in a linear 3-15% (wt/ vol) sucrose gradient in 0.1 M KCl, 2 mM MgCl2, 10 mM imidazole-HCl, 1 mM NaN3, pH 7.4. To measure Stokes' radius, the purified espin was chromatographed on a 93 × 1.5-cm column of Sephadex G-100 in the same buffer minus sucrose. The sedimentation coefficient and Stokes' radius were determined by SDS gel analysis of the gradient and column fractions, respectively, with comparison to the protein standards in the MW-GF-1000 gel filtration standards kit plus horse heart cytochrome c (Sigma Chemical Co., St. Louis, MO). The small espin was examined in SDS gels before and after centrifugation or gel filtration to ensure that it had not undergone proteolysis during the procedure. Circular dichroism spectroscopy was carried out at room temperature at rate of 50 nm/min using a freshly calibrated spectropolarimeter (model J-715; Jasco, Inc., Easton, MD) on samples containing ~50 µg/ml of purified small espin in 50 mM NaPi, pH 7.4, in a 0.1-cm path length cell. Molecular cross-linkers were purchased from Pierce Chemical Co. (Rockford, IL).
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Results |
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Identification and Sequence Analysis
To determine whether the ~1.7-kb crosshybridizing RNAs
detected previously in our Northern blot analysis of total
RNA isolated from rat kidney and small intestine (Bartles
et al., 1996) encoded espin-related proteins, homogenates
of rat kidney and small intestinal mucosa were compared
with homogenate of rat testis on Western blots using affinity-purified rabbit polyclonal antibody directed against the
COOH-terminal 379-amino acid peptide of rat Sertoli cell
espin. In addition to reacting with the ~110-kD Sertoli cell espin band present in homogenate of testis, the espin antibody also specifically labeled a closely spaced doublet of
~30-kD present in homogenate of rat kidney and small intestinal mucosa (Fig. 1 A).
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To elucidate the sequence of the ~30-kD protein detected by Western blotting, the espin antibody was used to
screen a commercial rat kidney cDNA library. A single
1,088-bp cDNA clone was obtained, and its sequence was
determined. The single large open-reading frame present
in this cDNA began at nucleotide 63, in the context of a
Kozak consensus sequence (Kozak, 1987), and was predicted to encode a small version of the espin protein that
was ~1/3 the size of Sertoli cell espin, 253 amino acids versus 837 amino acids,2 with a molecular mass of 28,240 D
and an isoelectric point of 10.5. Attempts at 5' rapid amplification of cDNA ends failed to identify any additional
in-frame translation start sites upstream (data not shown),
and the recombinant protein encoded by nucleotides 63-
821 of this cDNA was found to exhibit the expected apparent molecular mass (~30 kD) in SDS gels (see below). The
nucleotide sequence, the predicted amino acid sequence,
and a comparison between the small and large espin isoforms are shown in Fig. 2. Along much of its length, small
espin was identical in sequence to the COOH-terminal
portion of the larger Sertoli cell isoform. As a result, it
contained the potential P-loop (A32-S39), the 66-amino
acid peptide that showed 39% identity to a peptide encoded by exon A5 and is present in both the large and
small forked proteins of Drosophila (I140-K205), and the
two peptides that contained clusters of multiple glutamates bracketed by arginines, lysines, and glutamines in
a pattern reminiscent of the repetitive motif found in the
protein trichohyalin (R160-R182 and R195-R219). Small espin did not, however, contain the ankyrin-like repeats
and proline-rich peptides present in the large isoform. In
their place, it contained a considerably shorter NH2 terminus with two unique small peptides (M1-P13 and S61-R86) interrupted by another peptide (S14-A60) that was
present in the large isoform. When analyzed using the secondary structure algorithms of Chou and Fasman (1978)
or those of Garnier et al. (1978)
, a large fraction of the
COOH-terminal half of small espin (R160-A238) was predicted to adopt an
-helical secondary structure. In addition, when analyzed using the algorithms or Lupas (1997)
or Berger et al. (1995)
, two COOH-terminal peptides (Q156-L186 and R190-G231 or L194-G231) were predicted to form a coiled coil with high probability.
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Localization, Quantification, and Solubility
When localized at the light-microscopic level using the immunoperoxidase technique, small espin was found to be
concentrated over the F-actin-rich brush borders of absorptive epithelial cells in the small intestine and kidney
(arrowheads in Fig. 3, A and B, respectively). On sections
of rat small intestine, some weaker labeling was also detected more basally within enterocytes. Compared with
the brush borders of the mature enterocytes on intestinal villi, the apical margins of the enterocyte precursors found
within the crypts of Lieberkühn (e.g., Fig. 3 A, arrow) had
considerably lower levels of labeling. This suggested that
small espin was expressed or accumulated in a developmental fashion during the assembly of the brush border
(Ezzell et al., 1989; Heintzelman and Mooseker, 1992
;
Fath and Burgess, 1995
). On sections of rat kidney, only
the epithelial cells lining the distal straight portion of the
proximal tubule near the cortico-medullary junction, presumably corresponding to segment S3 (Kriz and Kaissling,
1992
), showed intense labeling along their brush borders
(Fig. 3 B). Proximal convoluted tubules near glomeruli in
the renal cortex showed considerably less labeling (data
not shown). The proximal tubular epithelial cells with intensely labeled brush borders appeared to have an even
higher level of cytoplasmic labeling than the enterocytes.
|
The majority (83-86%) of small espin present in homogenates of rat kidney and small intestinal mucosa resisted extraction with 1% (vol/vol) Triton X-100 (data not shown). Small espin was recovered in isolated rat small intestinal brush borders, where it was colocalized with F-actin at the light-microscopic level (Fig. 4, A-C). On the basis of quantitative Western blotting, we could estimate that ~1 small espin was recovered for every 130 actin monomers in small intestinal brush borders isolated from adult rats. Fig. 1 B shows a Western blot in which dilutions of isolated rat small intestinal brush borders containing known amounts of actin are compared with internal standards containing known amounts of recombinant small espin (see below). Consistent with an association between small espin and the actin cytoskeleton, the small espin present in the isolated rat small intestinal brush borders resisted extraction with 1% (vol/vol) Triton X-100 but could be partially extracted with 0.6 M KI (Fig. 1 C).
|
Post- and preembedment immunogold electron microscopy were used to examine the localization of the small espin recovered on the isolated rat small intestinal brush borders at a higher level of resolution. When LR white sections of paraformaldehyde-fixed brush borders were labeled with affinity-purified polyclonal espin antibody followed by secondary antibody-colloidal gold conjugate, the bound gold particles were observed over the microvilli proper and over the actin bundles of the microvillar rootlets, but not over the terminal web (Fig. 4 D). Neither the sections labeled with preimmune IgG instead of the espin antibody nor the occasional nuclear fragment that can contaminate this subcellular fraction showed bound gold particles (data not shown). After examining more than 250 brush border profiles, we concluded that the specifically bound gold particles could be found anywhere along the length of the microvillar actin bundle. Preembedment immunogold labeling (Fig. 4 E) indicated that a significant pool of small espin was accessible to primary antibody and secondary antibody-colloidal gold conjugate in isolated brush borders that were fixed with paraformaldehyde in the presence of Triton X-100. Specimens labeled with preimmune IgG instead of espin antibody showed no bound gold particles (not shown). Compared with postembedment labeling, the specifically bound gold particles appeared to be more evenly distributed along microvilli, but fewer gold particles were detected over rootlets, suggesting that there was reduced accessibility of the tracer using the preembedment technique. Comparable results were obtained when the diameter of the secondary antibody- colloidal gold was reduced from 12 to 6 nm (data not shown).
Transfection
To obtain additional evidence in support of an interaction between small espin and the actin cytoskeleton in vivo, we examined the localization of a GFP-small espin fusion protein in transiently transfected cells of the BHK fibroblastic line. On the basis of immunofluorescence and Western blotting (not shown), these cells were judged not to contain significant amounts of espin in the absence of transfection. When living cells were examined by conventional fluorescence microscopy 1 d after transfection, the GFP-small espin fusion protein was observed to decorate fine (Fig. 5 A) or more coarse (Fig. 5, B and C) stress fiber-like structures, whereas GFP without the small espin attached was found to be distributed more diffusely throughout the cytoplasm with evidence of concentration within the nucleus (Fig. 5 B, inset). To further examine the identity of the GFP-small espin-labeled fibers observed in living cells, the cells were fixed with paraformaldehyde, permeabilized with Triton X-100 and double labeled with rhodamine-phalloidin as a marker for F-actin. The patterns of GFP-small espin (Fig. 5 D) and rhodamine-phalloidin (Fig. 5 E) were found to be coincident, suggesting that exogenous small espin was associating with F-actin- containing structures in the transfected cells. In contrast, GFP without the small espin attached was detected in perinuclear aggregates in the fixed and permeabilized cells (Fig. 5 D, inset). Interestingly, the level of F-actin, as reflected by the intensity of rhodamine-phalloidin labeling, was consistently higher in cells expressing GFP-small espin than in the corresponding untransfected control cells (Fig. 5 E) or in cells expressing GFP alone (not shown). This latter result suggested that small espin could elicit the accumulation and/or bundling of F-actin-containing fibers or cables in the transfected cells. Neither the apparent accumulation of these F-actin-containing structures nor their decoration with small espin required the presence of the GFP because the same results were obtained when small espin was expressed without the GFP using the pcDNA3 vector and was detected by immunofluorescence (data not shown).
|
Bacterial Expression and Actin Binding/Bundling In Vitro
To examine the properties of small espin and its interactions with F-actin under defined conditions in vitro, it was necessary to obtain relatively large amounts of the purified protein. Since small espin was difficult to extract from organ/tissue homogenates and isolated brush borders (see above), we elected to use recombinant small espin for these studies. A full-length version of small espin containing a 28-amino acid NH2-terminal extension that included a 6xHis tag was expressed in E. coli and purified by affinity chromatography on Ni-NTA agarose under nondenaturing conditions. The 6xHis tag was released from the protein by cleavage with recombinant 6xHis-tagged Tobacco Etch Virus protease, and the protein, which now contained only five extra amino acids (GAMGS) at its NH2 terminus, was quickly repurified by a second challenge with Ni-NTA agarose before use.
The purified recombinant full-length small espin was
found to bind to and bundle F-actin at physiological ionic
strength and pH. This is illustrated in the left part of Fig. 6
A, where the supernate and pellet fractions resulting from
an F-actin-bundling assay, which uses low-speed centrifugation to separate bundles from preformed actin filaments, were analyzed in an SDS gel. In the absence of
small espin, the vast majority of the rabbit skeletal muscle
F-actin remained in the supernate fraction upon low-speed centrifugation. However, when the small espin was added
to the F-actin (in this case, at a molar ratio of approximately one espin/seven actin monomers), the solution
became noticeably turbid, and the vast majority of the
F-actin and the small espin were obtained as higher molecular weight cross-linked structures (bundles) in the pellet.
In contrast, in the absence of F-actin, the small espin remained in the supernate. When examined by negative
staining electron microscopy, the vast majority of the actin
filaments present in the small espin/F-actin mixtures were
found to be assembled into bundles (Fig. 7). In contrast,
the F-actin controls contained randomly oriented actin filaments but no discernible bundles (data not shown). The
small espin-induced bundles were partially ordered, frequently showing extended segments with transverse striations, aligned imperfectly across the width of the bundle,
at integral multiples of ~12 nm, which is characteristic of
maximally cross-linked, hexagonally packed filaments
(e.g., see Stokes and DeRosier, 1991; Tilney et al., 1995
).
Unlike the situation for many other actin-bundling proteins, which display actin-bundling activities that are inhibited by Ca2+(Glenney et al., 1981
; Alicea and Mooseker,
1988
; Namba et al., 1992
; Lin et al., 1994
), concentrations
of CaCl2 up to 1 mM had no effect on the bundling of
F-actin by small espin in the low-speed centrifugation assay (data not shown). And, unlike the bundling of F-actin
that results from counterion condensation by positively charged ions and small peptides (Tang and Janmey, 1996
),
relatively high concentrations of negatively charged small
molecules, e.g., 10 mM ATP, also had no effect on the
bundling caused by small espin (not shown).
|
|
To determine the affinity of the interaction between recombinant small espin and F-actin, high-speed centrifugation was substituted for the low-speed centrifugation to cause all of the F-actin to be recovered in the pellet. The data resulting from the comparison of high-speed pellet and supernate fractions at different inputs of small espin were used to construct Scatchard plots. As is illustrated in Fig. 6 B, the recombinant small espin was found to bind to human platelet nonmuscle F-actin with a threefold higher affinity than to rabbit skeletal muscle F-actin. The Kd for nonmuscle F-actin was determined to 50 nM, whereas the Kd for skeletal muscle F-actin was 150 nM. From the intercepts of the resulting linear regression lines on the abscissa, it could be calculated that the binding reached saturation at approximately one espin for every three to four actin monomers.
Oligomeric State, Shape, and Secondary Structure
To further elucidate the mechanism of actin bundling by small espin, we used sedimentation equilibrium to determine whether the purified recombinant small espin was a monomer, dimer or higher oligomer. From the slope of plots of log A280 versus radius squared at equilibrium (e.g., Fig. 8 A), the molecular mass was estimated to be 25,500 D, only ~12% below the value of ~29,000 D (28,240 D + NH2-terminal GAMGS) calculated on the basis of sequence, suggesting that recombinant small espin was a monomer. In further support of a monomeric structure for recombinant small espin, we failed to observe covalently cross-linked dimers or higher oligomers when the purified protein was reacted with molecular cross-linkers, such as 3,3'-dithiobis(sulfo-succinimidyl propionate) and 1,5-difluoro-2,4-dinitrobenzene, and analyzed in SDS gels (data not shown).
|
Given that recombinant small espin appeared to be a monomer, it was found to have an unexpectedly large Stokes' radius by gel filtration. When chromatographed in columns of Sephadex G-100, the recombinant small espin eluted only one fraction after bovine serum albumin, a standard protein with more than twice the subunit molecular mass (66 kD), and seven fractions before carbonic anhydrase, a standard protein with approximately the same subunit molecular mass (29 kD) (Fig. 8 B). From the plot of logarithm of the Stokes' radius versus peak elution fraction for the standard proteins (Fig. 8 B, inset), the Stokes' radius of recombinant small espin was estimated to be 3.4 nm, suggesting that it had a frictional coefficient ratio (f/fo) of 1.7 and was, therefore, an asymmetrical molecule. For example, it is possible to calculate that a frictional coefficient ratio of 1.7 for a protein of ~29,000 D (28,240 D + NH2-terminal GAMGS) is compatible with an oblate ellipsoid with dimensions of 10 × 0.6 nm, a prolate ellipsoid with dimensions of 23 × 1.8 nm, or a long rod with dimensions of 17 × 1.2 nm. Using the molecular mass of the small espin monomer and the frictional coefficient ratio of 1.7, one can estimate that the protein should have a sedimentation coefficient of ~2 S. The sedimentation coefficient of the recombinant small espin was measured directly by centrifugation under isokinetic conditions in sucrose density gradients (Fig. 8 C). From a plot of sedimentation coefficient versus peak fraction for the standard proteins (Fig. 8 C, inset), the sedimentation coefficient of small espin was determined to be 2.1 S, just slightly above the predicted value. The observation that small espin sedimented significantly slower than the 29-kD protein carbonic anhydrase, the standard protein with approximately the same molecular mass, also supported the conclusion that small espin was an asymmetrical monomer.
When examined by far-UV circular dichroism spectroscopy, the spectrum of the purified full-length recombinant
small espin had large negative peaks at 220 and 207 nm
and a large positive peak at 192 nm (Fig. 8 D), corresponding to the major n-* and
-
* transitions displayed by
-helices (Chen et al., 1974
). These results suggested that
the recombinant small espin was a highly folded protein and
that, as predicted on the basis of sequence (see above), it
contained a high percentage of
-helix. Unlike the spectra
of many other
-helical proteins, however, the negative peak at 207 nm was significantly larger than the one at 220 nm, suggesting that the helical region was relatively long
(>50 amino acids; Chen et al., 1974
) and/or that, despite
the prediction of a coiled-coil structure (see above), its
various helical segments did not interact with one another
(e.g., Zhou et al., 1992
; Su et al., 1994
).
Deletion Mapping of the Actin-bundling Peptide
PCR was used to generate a series of small espin NH2- or
COOH-terminal deletion constructs with NH2-terminal
6xHis tags to allow for ready purification. The sequences
of these constructs were verified by automated DNA sequencing, and the constructs were compared in the low-speed centrifugation actin-bundling assay after removal of
the 6xHis tag and repurification. Deletion of the NH2 terminus up to the 167-amino acid COOH-terminal peptide shared by the Sertoli cell and brush border isoforms
(N86), or up to the beginning of the forked homology
domain (
N137), produced recombinant proteins that
displayed actin-bundling activities equal to, or perhaps
slightly greater than, that of full-length small espin (Fig. 9).
An example of an SDS gel depicting the results of an actin-bundling assay for construct
N137 is shown on the right side of Fig. 6 A. Further deletion from the NH2 terminus, up to the middle of the forked homology peptide
(
N170), caused a dramatic (80-fold) decrease in actin-bundling activity to near background levels (Fig. 9). Deleting the last 5 amino acids from the COOH terminus of
small espin (
C5) had no effect on actin-bundling activity,
but deletion of an additional 13 amino acids (
C18), or deletion of a total of 73 amino acids from the COOH terminus (
C73), also caused a dramatic decrease in actin-bundling activity to near background levels (Fig. 9). These
results suggested that the COOH-terminal 116-amino acid
peptide common to the two known isoforms of espin
(A138-K253 in small espin) was sufficient to cause maximum actin bundling in vitro. Since small espin was found to be a monomer in solution (see above), the COOH-terminal 116-amino acid peptide must presumably contain
two or more actin-binding sites. The dramatic losses in actin-bundling activity observed in going from
N137 to
N170 and from
C5 to
C18 suggested further that the
33-amino acid peptide at the NH2 terminus of the forked homology peptide (A138-E170) and the 13-amino acid
peptide K236-K248 were necessary for bundling activity
and, therefore, may contribute to the formation of small
espin's actin-binding sites.
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Discussion |
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Taken together, our data strongly support the hypothesis that small espin is a newly identified actin-bundling protein of the brush border. Small espin is localized to the parallel actin bundles of brush border microvilli, and it accumulates there coincident with the migration/differentiation of enterocytes along the crypt-villus axis in the adult. The protein is recovered in the Triton X-100-insoluble cytoskeletal fraction prepared from isolated brush borders. Recombinant small espin binds to F-actin with high affinity, and it cross-links the actin filaments into partially ordered bundles. The recombinant small espin protein is a monomer in solution and appears to contain at least two relatively closely spaced actin-binding sites. And finally, when expressed exogenously in transfected fibroblastic cells, small espin decorates F-actin fibers and cables and appears to cause their accumulation, suggesting that small espin can bind to, stabilize, and/or bundle F-actin in vivo.
The Espin Family and the Mechanism of Espin-mediated Actin Bundling
Small espin showed no significant sequence similarity to
other proteins known or suspected to bind/bundle F-actin,
with the exception of the large isoform of espin and the
forked proteins of Drosophila. With the identification of
the small isoform of espin, it is clear that the espins constitute a new family of actin-binding/bundling proteins. The
two members of the espin family we have described thus
far, the 837-amino acid ectoplasmic specialization isoform
(Bartles et al., 1996) and the 253-amino acid brush border
isoform, are expressed in different cell types and are markedly different in size. Nevertheless, both isoforms are localized to specializations of the cell surface, an invagination or an evagination, that contain hexagonally packed
parallel actin bundles, and both contain an identical
COOH-terminal 167-amino acid peptide that is sufficient
to bundle F-actin in vitro. The observation that the peptides specific to the small isoform (M1-P13 and S61-R86)
are found interspersed among peptides identical in sequence to those present in the large isoform suggest that
the two espin isoforms arise through differential splicing.
We are currently investigating the organization of the espin gene(s) and comparing the properties and activities of
the small and large isoforms to ascertain how their different NH2 termini influence interactions with F-actin.
Although small espin is a basic protein and contains
many highly charged peptide segments, the fact that the
bundling took place in 0.1 M KCl at pH 7.4 and was not inhibited by a small, highly charged anion, in this case up to
10 mM ATP, suggests that the bundling is due to the presence of specific actin-binding sites in small espin and is not
the result of counterion condensation (Tang and Janmey,
1996). Through deletion mutagenesis, we established that
the COOH-terminal 116-amino acid peptide shared by
the two espin isoforms was sufficient to cause maximal actin bundling in the low-speed centrifugation assay. Since
small espin proved to be a monomer in solution, this suggests that the COOH-terminal 116-amino acid peptide
contains two, or possibly more, actin-binding sites. The
simplest model that can account for the dramatic losses in
actin-bundling activity noted in going from
N137 to
N170 and from
C5 to
C18 is one in which small espin
contains two actin-binding sites disposed roughly at opposite ends of the COOH-terminal 116-amino acid peptide.
Our sedimentation and gel filtration analyses suggest that
small espin is a relatively small, asymmetrical molecule with dimensions in the range of 10-23 × 0.6-1.8 nm. Further experiments are underway to elucidate the three-dimensional structure of small espin and to map its actin-binding sites and its footprint on F-actin at high resolution.
Relationship between the Espins and the Forked Proteins of Drosophila
The dramatic loss in the actin-bundling activity of small espin noted in going from N137 to
N170 suggests that the
NH2-terminal half of its 66-amino acid forked homology
peptide is necessary for actin-bundling activity and may
therefore be important in forming one of small espin's actin-binding sites. This lends support to the hypothesis
(Bartles et al., 1996
) that the espins and the forked proteins are orthologous families of proteins. Although, to
our knowledge, no forked protein has yet been observed
to bundle F-actin directly, genetic studies have demonstrated that the forked proteins are necessary to form normal actin bundles in the developing bristles of Drosophila
pupae (Hoover et al., 1993
; Petersen et al., 1994
; Tilney et
al., 1995
; Wulfkuhle et al., 1998
). Forked proteins have
been localized to the developing bristles of Drosophila pupae by immunofluorescence (Petersen et al., 1994
), suggesting that they, too, may be components of plasma
membrane-associated parallel actin bundles. The forked
proteins also appear to come in multiple isoforms: six
forked transcripts have been detected during mid to late
pupal development, and five of the six isoforms are predicted to share a COOH-terminal peptide, which includes the 66-amino acid peptide with 39% identity to the forked
homology peptide present in the shared COOH-terminal
peptide of the espins (Hoover et al., 1993
; Bartles et al.,
1996
). On the basis of the ability of one of the smaller
forked proteins to restore normal actin bundle and bristle
morphology in transgenic forked mutant pupae, it is presumably the shared COOH-terminal peptide that contains
the actin-bundling module postulated to exist within the
forked proteins (Petersen et al., 1994
). A peptide present within the shared COOH-terminal peptide of the forked
proteins
the one encoded by exon A5 of the forked gene
that includes the 66-amino acid peptide with homology to
the espins
has, in fact, been observed to bind to F-actin
(Petersen, N.S., personal communication). Although other
primary structural features of the forked proteins (e.g., the
alternating histidine and proline residues of the PRD-like domain, the polyglutamine tracts encoded by the CAX trinucleotide repeats, and the peptides composed of large
numbers of predominantly hydrophobic amino acids) are
notably absent from the espins, the larger isoforms of the
forked proteins do contain two proline-rich peptides and
NH2-terminal ankyrin-like repeats with an overall spacing
that approximates that observed in the large isoform of espin (Hoover et al., 1993
; Bartles et al., 1996
). It is presently unclear whether forked proteins will be found in other tissues of Drosophila. The forked phenotype appears to be
limited to the neurosensory bristles of pupae and to the
hairs and denticles of larvae and adults, and to our knowledge, these are the only structures in Drosophila found, or
postulated, to contain a forked protein to date (Hoover et
al., 1993
; Petersen et al., 1994
; Tilney et al., 1995
; Dickinson and Thatcher, 1997
; Wulfkuhle et al., 1998
).
Toward a Three Actin-bundling Protein Model
At a minimum, our results suggest that microvillar actin
bundles are even more complex and heterogeneous than
thought previously. On the basis of our findings, we propose that in some cells it may take not two, but three actin-bundling proteins to construct and/or maintain a fully
functional microvillar actin bundle. An extension of the
prevailing two-bundling protein model (Tilney et al.,
1995), the three-bundling protein model would appear to be consistent with the results of recent experiments examining pupal bristle formation in singed/forked double mutants of Drosophila that imply the involvement of at least
one additional actin-bundling protein (Wulfkuhle et al.,
1998
). It would also appear to follow logically from our observation that small espin is present at different levels in
the brush borders of cells that line different segments of
the proximal tubule in rat kidney. In this regard, it may be
relevant that the cells in the kidney we found to contain the largest amounts of brush border small espin, those lining the distal straight segment (S3) of the proximal tubule,
are also known to display microvilli that are longer and
more densely packed than those in other segments of the
proximal tubule in the rat (Kriz and Kaissling, 1992
).
Small espin appeared to be 10- to 20-fold less abundant
than the two known major actin-bundling proteins, fimbrin/plastin and villin, in small intestinal brush borders isolated from adult rats. Barring losses due to degradation or
extraction during brush border isolation, we can estimate
that there is only ~1 espin for every 130 actin monomers
present in the intestinal brush border. Whether calculated
assuming the theoretical value for maximally cross-linked,
hexagonally packed filaments of ~1 cross-link for every
4.3 actin monomers (Tilney et al., 1995), or calculated using the ~10:0.65:1.3 molar ratio of actin monomer/fimbrin/
villin in isolated intestinal brush borders (Coluccio and
Bretscher, 1989
), this stoichiometry suggests that only ~1
in every 30 crossbridges between actin filaments would be
due to small espin in the microvillar actin bundles of the
adult small intestine. Although its lower abundance may
help to explain why small espin escaped the attention of
earlier investigators, it makes it more challenging for us to
identify the role(s) of small espin in microvillar actin bundle organization and/or function.
Possible Roles for Small Espin as a High-affinity, Ca2+-insensitive Cross-Linker, Filament Stabilizer, and/or Linker Protein
In the context of a three-bundling protein hypothesis, it is
possible to envision at least a few different scenarios that would be compatible with the available evidence. One especially intriguing possibility is that small espin might confer a special property upon the bundle and that it is sufficient to have the cross-links involving this third bundling
protein spaced relatively far apart in the completed structure. One obvious special property that small espin could
confer upon a bundle is high-affinity cross-linking that is
insensitive to changes in the concentration of Ca2+. Small
espin exhibits a binding affinity for F-actin that is at least
an order of magnitude higher than those displayed by many of the other known actin-bundling proteins, including fimbrin/plastin and villin (Glenney et al., 1981; Burgess
et al., 1987
; Pollard, 1993
). In addition, both villin and fimbrin/plastin have binding sites for Ca2+, and the bundling
activities of villin and fimbrin/plastin are known to be disrupted by physiological concentrations of Ca2+ (Glenney
et al., 1981
; Alicea and Mooseker, 1988
; Namba et al., 1992
; Lin et al., 1994
). However, we found that the actin-bundling activity of small espin was insensitive to changes
in the concentration of Ca2+ over a wide range. We are
currently testing the hypothesis that strategic placement of
small espin at intervals along the filaments might reinforce
the bundle by protecting it from changes in Ca2+ concentration.
A second special property that small espin might be able
to confer upon actin bundles is stabilization to depolymerization. This would be one possible explanation for the apparent accumulation of F-actin we noted in the small espin-expressing transfected cells. A similar stabilization
effect, apparently independent of cross-linking, has already been noted in vitro for one actin-bundling protein,
the ~34-kD actin-bundling protein of Dictyostelium (Zigmond et al., 1992). In the case of the small espin-expressing transfected cells, it is difficult to sort out the effects of
stabilization versus bundling. For example, we do not yet
know the extent to which the F-actin fibers and cables observed in the transfected cells contain parallel actin bundles in addition to the antiparallel (contractile) bundles
typical of stress fibers. We are currently carrying out experiments to determine whether the espin isoforms can
stabilize F-actin independently of cross-linking. We are
also investigating the possibility that the apparent accumulation of F-actin observed in the transfected cells reflects
an increase in actin synthesis to maintain a sizable pool of
monomer. Nevertheless, it is striking that the results we
obtained with small espin are quite different from those
obtained by others examining the localization and effects
of villin and fimbrin/plastin in transfected fibroblastic cells: villin causes a loss of stress fibers and the redistribution of F-actin into long spike-like microvilli at the dorsal
surface (Friederich et al., 1989
), whereas T- and L-plastins
cause fibroblastic cells to round up and reorganize their
stress fibers into geodesic structures (Arpin et al., 1994
).
We are currently comparing the localizations and effects
of the espin isoforms in transfected epithelial cells.
Another special property that could be conferred upon
the bundles by small espin might be to link the bundles to
other proteins, possibly even to those of signaling cascades. The ability to act as a linker or adaptor protein
would appear to be especially likely for the large isoform
of espin, with its multiple motifs for protein-protein interactionthe ankyrin repeats and the proline-rich peptides (for additional discussion, see Bartles et al., 1996
)
but
could also readily apply in the case of the small isoform.
We are currently using a combination of affinity approaches and two-hybrid screens to identify espin-binding
proteins other than actin.
Developmental Accumulation of Small Espin
One possible explanation for why small espin is relatively
minor in the intestinal brush borders of adult rats is that it might be required at an earlier step in brush border assembly. In such a case, it might be expressed at lower levels or
be more unstable in adults. The results of our immunoperoxidase localization of small espin in the small intestines of
adult rats suggest, however, that this is probably not the
case. In many respects, the assembly of the intestinal brush
border along the crypt-villus axis in the adult approximates an accelerated version of intestinal brush border assembly during late embryonic development (Heintzelman and Mooseker, 1992; Fath and Burgess, 1995
). We noted a
large differential in the intensity of apical immunoperoxidase staining between crypt cells and villus cells in adult
intestine. Although there are some limitations inherent in
trying to use immunoperoxidase histochemistry as a quantitative measure, our results suggest that maximal small espin accumulation occurs not early, but relatively late during enterocyte differentiation and brush border assembly. In contrast, both villin and fimbrin/plastin have been
found to be nearly as prevalent in the apical cytoplasm of
crypt cells in the adult as they are in the brush borders of
the mature enterocytes (Fath et al., 1990
; Heintzelman and
Mooseker, 1990
; Landry et al., 1994
). The increase in immunoperoxidase staining for small espin was first noted
for enterocytes nearing the crypt-villus junction, roughly
coinciding with the timing of the terminal elongation of
microvilli and the arrival of brush border myosin I (Heintzelman and Mooseker, 1992
; Fath and Burgess, 1995
). To
address this issue more fully, we are carrying out an analysis of small espin expression and localization in relation to
other brush border proteins during the development of intestine and kidney and in cell culture models of enterocyte
differentiation.
Partial Redundancy and the Presence of Espins in Other Actin Bundle-containing Structures
Even though small espin appears not to be as major as fimbrin/plastin or villin in the intestinal brush borders of
adults, it is still possible that it is much more abundant in
other F-actin-containing structures in the body. This situation would be somewhat analogous to that of ezrin, which
is a minor protein relative to brush border myosin I in the
intestinal brush border, but appears to supplant brush border myosin I as the major F-actin-membrane linker of placental microvilli (Bretscher et al., 1997). In such a case, small espin could be partially redundant in function in the
intestinal brush border of normal adult rats and of primary
importance only when one of the other actin-bundling
proteins is impaired or missing. In this regard, it has been
reported that mice that are missing villin due to targeted
gene disruption have surprisingly normal looking intestinal brush borders (Pinson et al., 1998
). In preliminary experiments, increased levels of brush border espin have
been detected in villin gene knock-out mice by immunofluorescence and Western blotting (Ferrary, E., and S. Robine, personal communication). In addition, we have
obtained preliminary evidence that espin isoforms are indeed more plentiful in other structures that contain parallel actin bundles (unpublished data). The further characterization of the espin isoforms found in these other
locations is in progress.
![]() |
Footnotes |
---|
Received for publication 14 July 1998 and in revised form 27 August 1998.
Address all correspondence to James R. Bartles, Department of Cell and
Molecular Biology, Northwestern University Medical School, 303 East
Chicago Avenue, Chicago, IL 60611. Tel.: (312) 503-1545. Fax: (312) 503-7912. E-mail: j-bartles{at}nwu.edu
1. Abbreviation used in this paper: GFP, green fluorescent protein.
2. We have discovered three errors in the sequence of the large isoform
between nucleotides 1835 and 1907 (Bartles et al., 1996). These errors
have been corrected in the database (available from GenBank/EMBL/
DDBJ under accession number U46007). The corrections bring about a
shift in reading frame over a span of 22 amino acids and result in the addition of one amino acid. The corrected total number of amino acids for the
large isoform is 837, and the corrected amino acid sequence for this region
is: 607-GAGAACGQRRSSSSTGSTKSFNMMSPTG-634.
We thank Dr. Mike Shaw (University of Pennsylvania, Philadelphia, PA) for advice about immunogold labeling of brush borders; Dr. Dan Kiehart (Duke University, Chapel Hill, NC) for advice about PHEM buffers and coiled coils; Maya Moody for help with sectioning, microscopy, and digital imaging; Dr. Rob Moir and Miri Yoon for advice about bacterial expression and GFP transfection systems; Dr. Kate Spiegel for help with the analytical ultracentrifuge; Dr. Brian Schoichet for help with the circular dichroism spectroscopy; Jodi Irwin for help with automated sequencing; Dr. Nancy Petersen (University of Wyoming, Laramie, WY) and Dr. Evelyne Ferrary and Dr. Sylvie Robine (Institut Curie, Paris, France) for granting us permission to discuss their results before publication; and Dr. Rex Chisholm for valuable discussions.
This work was supported by American Cancer Society research grant #CB-203, National Institutes of Health (NIH) grant R01 HD35280, and NIH Independent Scientist Award K02 HD01210 awarded to J.R. Bartles.
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