Department of Anatomy, Johannes Gutenberg-University Mainz, Becherweg 13, 55128 Mainz, Germany
* Author for correspondence (e-mail: leube{at}mail.uni-mainz.de )
Accepted 23 January 2002
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Summary |
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Key words: Cell contact, Desmosome, Intermediate filament, Live cell imaging, Green fluorescent protein, Fluorescence recovery after photobleaching
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Introduction |
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In the current study, we have examined desmosomes, which are believed to be
the major stabilizing cell contact type of epithelial cells. Interference with
desmosomal adhesion therefore leads to reduced tissue integrity and blister
formation (for reviews, see Moll and Moll,
1998; Udey and Stanley,
1999
; Kowalczyk et al.,
1999
). Desmosomes are morphologically well defined structures
(Schmidt et al., 1994
;
Burdett, 1998
;
Kowalczyk et al., 1999
) that
are built around a 20-30 nm intercellular space that contains the desmoglea
and is bisected by a dense midline. The cytoplasmic surfaces of both adjacent
cells are decorated by electron-dense plaques that serve as anchorage sites
for the intermediate filament (IF) cytoskeleton. Major desmosomal components
have been identified, and their molecular interactions and topologies have
been determined (for reviews, see Schmidt
et al., 1994
; Troyanovsky and
Leube, 1998
; Burdett,
1998
; Kowalczyk et al.,
1999
; North et al.,
1999
). In the emerging picture, desmosomal cadherins of the
desmoglein (Dsg) and desmocollin (Dsc) type take a central role by interacting
laterally and transcellularly with each other and recruiting cytoplasmic
plaque proteins that facilitate IF attachment. Their adhesive function is
coupled to the four extracellular domains that bind to calcium and contact
each other in a calcium-dependent fashion. Despite the high degree of overall
divergence of their cytoplasmic domains, the three Dsg isoforms and splice
variants `a' of all three Dsc isoforms contain a region that binds to the
universal plaque protein plakoglobin. In turn, the desmosomal
cadherin-plakoglobin complex interacts with the desmosome-specific plaque
protein desmoplakin, which also contains binding sites for the epithelial IF
proteins of the cytokeratin (CK) type. Further desmosomal components, such as
the plakophilins, probably modulate this basic architecture in specific but as
yet poorly understood ways.
Given the stabilizing properties of desmosomes, mechanisms must exist to
alter these properties during situations when rearrangements of cell-cell
contact occur, for example, during mitosis and migration. To examine such
changes, the modulation of calcium concentration has been used to induce rapid
alterations in desmosomal adhesion in vitro. Thus, substitution of standard
calcium medium (SCM) by low calcium medium (LCM) leads to the rapid loss of
desmosomal adhesion by the splitting of desmosomes and endocytosis of the
resulting desmosomal halves (Kartenbeck et
al., 1982; Mattey and Garrod,
1986
; Kartenbeck et al.,
1991
). In LCM, desmosomal components continue to be synthesized
but are only assembled into half desmosomes
(Duden and Franke, 1988
;
Burdett, 1993
;
Demlehner et al., 1995
).
Conversely, desmosomal adhesion is established when cells are transferred from
LCM to SCM (Hennings and Holbrook,
1983
; Watt et al.,
1984
).
Dynamic aspects of desmosomal behavior could not be directly visualized with previous methods. Even in instances of considerable desmosome reorganization, the precise sequence of events had to be painstakingly reconstructed from series of observations of cells that were fixed at different time points for analysis. Given the inherent variability between cells, it was therefore often difficult to decide whether different morphologies resulted from intercellular differences and differing reaction patterns or from simultaneously occurring stages of the same dynamic process. For example, it was not clear whether endocytosis alone accounted for the loss of desmosomal adhesion in LCM or whether and to what extent other mechanisms such as the dispersion of desmosomal components in the plasma membrane and cytoplasm contributed to this process. Similarly, it was not easy to determine whether and to what extent newly formed desmosomes were derived from preassembled half-desmosomes or from non-particulate precursors. Furthermore, nothing was known about the dynamic aspects of desmosomes in tightly coupled cells during the steady state.
As a first step to address such questions and to examine desmosomes in vivo, we have prepared cells expressing fluorescent desmosomal cadherins that are integrated into normal-appearing desmosomes. We show that these desmosomes are extremely stable and highly immobile structures that are maintained as distinct entities throughout the cell cycle together with adhering CK filaments (CKFs). On the other hand, in accordance with the above-mentioned studies, desmosomal adhesion is rapidly lost upon the reduction of calcium, leading to destabilization of desmosomes and endocytosis of desmosomal particles. Finally, we have found that, despite the extreme structural stability of desmosomes, desmosomal cadherins are rapidly exchanged.
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Materials and Methods |
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To prepare a cDNA construct coding for a fluorescent CK18 chimera, a 270 bp
fragment of the 3' end of the coding region of cDNA clone pHK18-P-7
(Bader et al., 1991) was first
amplified by PCR with amplimers 99-16 5'- CTC AAC GGG ATC CTG CTG CA-
3' and 99-17 5'-TTT GGT ACC CCA TGC CTC AGA ACT TTG GTG
T-3'. This fragment was cloned after restriction with BamHI and
Asp718 into pBluescript KS+ (Stratagene, La Jolla, CA), and the
insert was complemented by the 1 kb BamHI 5' fragment of
pHK18-P-7, thereby generating clone pHK18
stop. The complete cDNA insert
of pHK18
stop was then transferred as a EcoRI/Asp718
fragment into pECFP-N1 (Clontech Laboratories), producing clone C-HK18-CFP1,
which codes for cyan fluorescent hybrid HK18.CFP whose expression is under the
control of the CMV promoter and which also confers neomycin resistance.
Cell culture methods
Human hepatocellular carcinoma-derived PLC cells (ATCC CRL8024) and
Madin-Darby canine kidney (MDCK) cells (clone 20; ATCC CCL-34) were passaged
in DMEM (PAA Laboratories, Cölbe, Germany) supplemented with 10% fetal
calf serum (Invitrogen, Karlsruhe, Germany). DNA constructs were introduced
into these cell lines by using the calcium phosphate precipitation method
(Leube, 1995). To isolate
stably transfected clonal cell lines, transgenic cells were selected with
either geneticin (1 mg/ml; Life Technologies, Karlsruhe, Germany) or
hygromycin (150 µg/ml hygromycin; Sigma, St. Louis, MO), depending on the
construct used.
In some experiments calcium concentration was reduced by using calcium-free DMEM (Invitrogen) together with fetal calf serum whose calcium concentration had been reduced to 4.6 mg/l by treatment with the anion exchanger Diaion CR11 (Supelco, Bellefonte, PA).
Immunofluorescence microscopy
Rabbit polyclonal antibodies that were raised against green fluorescent
protein (GFP) and that were also reactive with the yellow and cyan fluorescent
proteins (YFP and CFP, respectively) were from Molecular Probes (Eugene, OR).
Murine monoclonal antibodies against desmoplakin (DP 2.15/2.17/2.20), Dsg (DG
3.10), plakophilin 2 (PP2/86) and plakophilin 3 (PKP3-270.6.2) were from
Progen (Heidelberg, Germany). Monoclonal antibody 11E4 was used for the
detection of plakoglobin (Kowalczyk et
al., 1994). Cy3-conjugated secondary antibodies were from Biotrend
(Cologne, Germany).
Electron microscopy
Electron microscopy and silver-enhanced immunoelectron microscopy with
polyclonal GFP antibodies were carried out as previously described
(Windoffer and Leube, 1999;
Windoffer et al., 2000
).
Immunoblotting and immunoprecipitation
Confluent cells were rinsed twice with ice-cold phosphate-buffered saline.
After removal of all fluid, cells were lysed in hypotonic H-buffer containing
10 mM Tris-HCl (pH 7.4), 1 mM EGTA, 1 mM EDTA, 2 mM dithiothreitole (DTT), 0.1
mg/ml phenylmethane sulfonylfluoride (PMSF) and 0.2 mg/ml
4-(2-aminoethyl)-benzenesulfonyl fluoride (Sigma) at 4°C. The total cell
lysate was homogenized in a tight-fitting Dounce homogenizer by 30 up and down
strokes. Postnuclear supernatant was then prepared by centrifugation at 1000
g for 10 minutes at 4°C. Further centrifugation of this
supernatant (100,000 g at 4°C for 1 hour) yielded a
100,000 g pellet fraction that was dissolved in H-buffer.
Total cell lysates, postnuclear supernatant fractions, and the 100,000
g fractions were diluted 1:1 in loading buffer (2% (w/v) SDS,
150 mM DTT, 0.005% (w/v) bromophenol blue, 30 mM Tris-HCl pH 6.8, 10% (w/v)
glycerol) and subjected to SDS polyacrylamide gel electrophoresis (SDS-PAGE),
electroblot transfer and antibody incubation
(Windoffer and Leube, 1999;
Windoffer et al., 2000
).
Detection of bound horseradish-peroxidase-coupled secondary antibodies was
performed with the help of enhanced chemiluminescence (Amersham Pharmacia
Biotech, Freiburg, Germany).
For immunoprecipitation experiments, confluent cells were washed twice in phosphate-buffered saline at 4°C. After complete removal of all fluid, 3.5 ml immunoprecipitation buffer was added per 10 cm diameter Petri dish. Three different immunoprecipitation buffers were used: (1) RIPA buffer (10 mM Na2HPO4 (pH 7.2), 150 mM NaCl, 2 mM EDTA, 1% (v/v) Triton X-100, 0.25% (w/v) SDS, 1% (w/v) sodium deoxycholate, 2 mM PMSF), (2) RIPA buffer without SDS or (3) RIPA buffer without SDS and sodium deoxycholate. Cells were incubated in the respective buffer for 30 minutes at 4°C with occasional shaking and then scraped off. Homogenization was carried out with a tight-fitting Dounce homogenizer by 30 up and down strokes. Lysed material was cleared by centrifugation at 40,000 g for 50 minutes at 4°C, and the supernatant was incubated with anti-GFP antibodies (see above) overnight at 4°C on an overhead roller. As a negative control, the homogenate was incubated in parallel without the antibody. To each sample, 15-20 mg pre-swollen protein-A-sepharose (type CL-4B; Amersham Pharmacia Biotech, Freiburg, Germany) was added per milliliter immunoprecipitation buffer followed by an incubation for 1-2 hours at 4°C. Subsequently, the sepharose was spun down at 2000 g in a table-top centrifuge for 10 minutes at 4°C, followed by three wash cycles in buffer containing 0.5% (w/v) Tween-20, 50 mM Tris-HCl pH 7.5, 150 mM NaCl and 0.1 mM EDTA, and five wash cycles in buffer containing 0.5% (w/v) Tween-20, 100 mM Tris-HCl pH 7.5, 200 mM NaCl and 2 M urea. Finally, the sepharose was washed in phosphate-buffered saline supplemented with 1% (w/v) Triton X-100 and then dissolved directly in loading buffer. After being boiled for 5 minutes and a brief centrifugation step, the supernatants were loaded on 8% SDS-polyacrylamide gels for electrophoresis and immunoblotting with either anti-GFP antibodies (dilution 1:1,000) or anti-plakoglobin antibodies 11E4 (dilution 1:70). Bound antibodies were detected with an enhanced chemiluminescence system.
Live cell imaging
The culture chamber and culture conditions used for time-lapse fluorescence
microscopy of living cells have previously been described in detail
(Windoffer and Leube, 1999;
Windoffer et al., 2000
).
Images were recorded in some instances with a confocal laser scan microscope
(Leica TCS NT, Leica Microsystems, Wetzlar, Germany) as described
(Windoffer and Leube, 2001
)
and, in most instances, by epifluorescence microscopy with either of the
recording devices described previously
(Windoffer and Leube, 1999
;
Windoffer et al., 2000
) or an
imaging system with inverse optics from Olympus (Hamburg, Germany). The latter
system was equipped with a monochromator for excitation and a piezo-driven
z-axis stepper attached to the 60x 1.4 N.A. oil immersion objective.
Pictures were recorded with an IMAGO slow scan charged-coupled device camera,
and the system was controlled by TILLvisION software. The microscope was kept
in a climate chamber at 37°C with the cells either in a closed culture
chamber (see above) or in a Petri dish with a glass bottom (Mattek, Ashland,
MA). Excitation was at 496 nm for enhanced GFP, 498 nm for enhanced YFP and
436 nm for enhanced CFP. For a 3D delineation of structures, multiple focal
planes were recorded for each time point by using the piezo stepper. The
resulting picture stacks were either projected on top of each other or were
used to prepare 3D reconstructions with the help of Amira (TGS) software.
Image sequences were edited with Image-Pro Plus 4.5 (Media Cytbernetics) and were converted into QuickTime movies (Apple) (see jcs.biologists.org/supplemental ). To edit single pictures and to arrange them into figures, Photoshop software (Adobe Photoshop 5.0) was used.
To compute time-space diagrams, rows of labeled desmosomes were cut out from recordings at each time point, and the resulting pictures were imported into Amira. The image data were compiled, producing trajectories that present the surface view of individual desmosomes in time and space.
Photobleaching experiments were carried out with a Leica TCS SP2 confocal microscope. The 488 nm line of an argon/krypton laser was used for both bleaching and image recording. The emitted light was monitored between 500 nm and 590 nm. Recordings were performed via a 63x 1.4 N.A. oil immersion objective. Standard settings for prebleach and recovery image scans were 6% of minimum laser power, a line average of 4 and a gain of 800. A wide pinhole size (setting of 500) was chosen for high depth of focus, minimal photobleaching and strong fluorescence signal. Rows of desmosomes that were straight for several micrometers were selected for bleaching to be able to define rectangular areas of interest. Bleaching in the chosen areas of interest was carried out at 100% of medium laser power for a total of 20 scans. Under these conditions, bleaching was practically complete not only in the focal recording plane but also in the regions above and below, thereby excluding the possibility that focal shift or cell motility was erroneously taken as a source for fluorescence recovery. The microscope was set to prebleach parameters immediately after bleaching, and images were recorded at intervals of 5 minutes. The recording time was limited to 30 minutes, since cell-shape changes resulted in the movement of desmosomal arrays in and out of the area of interest, thereby preventing the correct measurements of recovery at later time points. The gray values of the bleached areas were measured in the recorded 12 bit image data (Image-Pro Plus 4.5), analyzed by using spreadsheet routines (Excel) and drawn into diagrams.
To obtain optimal spatial resolution, the pinhole size was reduced (setting of 90) in some experiments. In these instances, 10 focal planes were recorded prior to bleaching, immediately after bleaching and after a 30 minute recovery period.
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Results |
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For further analyses, stable cell clones were established from PLC and MDCK
cells, two of which are described in detail in this communication. PLC subline
PDc-13 was obtained after transfection with plasmid C-Dsc2a.GFP-1, driving the
expression of the green fluorescent fusion protein Dsc2a.GFP with the help of
the CMV promoter. The Dsc2a.GFP fluorescence pattern was almost identical to
that observed by indirect immunofluorescence with antibodies against GFP
(Fig. 1A,A'). Therefore,
GFP fluorescence reflects the distribution of the hybrid polypeptides within
the entire cell. The transgene products were almost exclusively detected in
puncta at the cell surface where they were colocalized with the desmosomal
plaque protein desmoplakin (Fig.
1B,B') and Dsg2, the other desmosomal cadherin of PLC cells
(Fig. 1C,C')
(Schäfer et al., 1994).
MDCK-derived clone MDc-2 synthesized the yellow fluorescent Dsc2a chimera
Dsc2a.YFP from plasmid C-Dsc2a.YFP-2. In this case, fluorescent puncta
representing individual desmosomes were much smaller and more tightly spaced
than in PDc-13 cells (Fig.
1D-H), in accordance with the desmosome staining of the
corresponding wild-type cell lines by indirect immunofluorescence (not
shown).
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Furthermore, double fluorescence microscopy of fixed MDc-2 cells also revealed that the chimeras colocalized with the desmosomal proteins plakoglobin, desmoplakin, Dsg, plakophilin 2 and plakophilin 3 (Fig. 1D-H). As expected, antibody staining with the universal plaque protein plakoglobin displayed additional non-desmosomal plasma membrane domains (Fig. 1D,D'). All staining patterns were indistinguishable from those observed in wild-type nontransfected cells (not shown), indicating that the transgene expression did not affect desmosome formation and composition and that Dsc2a mutants were not mistargeted. In addition, the ultrastructure of desmosomes in both transgenic cell lines was indistinguishable from that of wild-type cells, presenting cytoplasmic plaques with inserting IFs, a defined intercellular space with desmoglea, and in some instances, a recognizable midline (Fig. 2A).
|
Immunoelectron microscopy with GFP antibodies also showed that the fusion proteins were highly enriched in typical desmosomes with an even distribution throughout the plaque regions (Fig. 2B,C; controls in Fig. 2D,E). Anchorage of CKF bundles was normal in these strongly labeled desmosomes.
To characterize further the morphology and distribution of desmosomes in the cDNA-transfected cell lines, 3D-reconstructions were produced from stacks of serial focal fluorescence images. Fig. 2F and the corresponding Movie 1 (see jcs.biologist.org/supplemental ), which contains an animated version of the reconstruction, show the macular, that is, dot-like, morphology of desmosomes, which are arranged as several circumferential lines around MDc-2 cells. In addition, some diffuse non-desmosomal fluorescence was seen that was apparently located in the plasma membrane. Similar 3D reconstructions were obtained from wild-type cells in which desmosomes had been labeled by indirect immunofluorescence microscopy.
Immunoblotting of cell fractions with GFP antibodies revealed that fusion
proteins of the expected size (calculated Mr of the mature
polypeptide chimera is 123,381) were synthesized in both cell lines and were
detectable in total cell lysates, postnuclear supernatant fractions and
100,000 g pellets (Fig.
2G). To determine whether Dsc2a chimeras interact with
plakoglobin, the major desmosomal cadherin-associated plaque molecule
(Troyanovsky and Leube, 1998),
coimmunoprecipitation experiments were performed. These showed a specific
association of both molecules in a high salt buffer containing 1% deoxycholate
and 1% Triton-X-100, but this could be disrupted by SDS
(Fig. 2H). Taken together, our
observations demonstrate that Dsc2a chimeras are integrated into structurally
and functionally competent desmosomes in PDc-13 and MDc-2 cells and that the
punctate fluorescence conveyed by these molecules can therefore be used for
the monitoring of desmosome dynamics in these cells.
Desmosomes are static and interconnected throughout interphase
Time-lapse fluorescence microscopy of PDc-13 and MDc-2 cells was performed
to monitor desmosome behavior in vivo. A series of pictures taken from a
recording of PDc-13 cells (Movie 2;
jcs.biologists.org/supplemental
) is shown in Fig. 3. In this
sequence, individual desmosomes could be traced throughout the entire
observation period of 3 hours. However, the motility of cells within the
monolayer often resulted in cellshape changes, leading to a shift of
desmosomes. In general, most desmosomes maintained their particular appearance
and fluorescence intensity and remained separate. Fusion and fission of
desmosomes was rare (the arrowheads in Fig.
3 show fusion of small desmosomes into larger structure).
Occasionally, fluorescent vacuolar structures that moved very little, except
for some local gyrating motility, and that did not seem to exchange with the
other fluorescent components (small arrows in
Fig. 3), were detected in the
cytoplasm. Similar results were also obtained in MDc-2 cells by time-lapse
fluorescence microscopy (see below), although the small size and close spacing
of desmosomes made it more difficult to follow them individually.
|
Time-space diagrams were constructed for the graphical visualization of desmosome motility. To this end, continuous shape reconstructions were prepared from a series of 2D images in time and space, the resulting trajectories depicting motilities in the x-y directions during time. A contact region of tightly associated PDc-13 cells was used for the computation of the diagram in Fig. 4A (see also Movie 3 at jcs.biologists.org/supplemental ). It demonstrates that these desmosomes moved only very little during the 3 hour recording time. Furthermore, the size, shape and fluorescence intensity of individual desmosomes were maintained throughout. Interestingly, the minor movements of desmosomes appeared to be coordinated. The diagram shown for a contact region taken from a growing but confluent MDc-2 culture in Fig. 4B depicts the same phenomenon (see Movie 4 at jcs.biologists.org/supplemental ). Remarkably, despite migration of the cell for over 30 µm, that is, more than the cell's diameter, the relative position of the desmosomes remained the sample for 10 hours without significant exchange or crossing over between the different time tracks. However, significant overall flexibility of the contact area was seen, which almost doubled in size at times (compare the time points marked by arrows in Fig. 4B).
|
To characterize the dynamic relationship between desmosomes and the
adhering CK cytoskeleton, double fluorescence time-lapse microscopy was
performed in cells that coexpressed Dsc2a. YFP together with HK18.CFP, a human
CK 18-ECFP chimera (see also Windoffer and
Leube, 1999; Strnad et al.,
2001
; Windoffer and Leube,
2001
). To this end, HK18.CFP was transiently expressed in MDc-2
cells by cDNA transfection of plasmid C-HK18-CFP1. An example of a doubly
transfected cell demonstrating that peripheral filaments extending from the
dense filamentous mesh are in close proximity to desmosomes is depicted in
Fig. 5. Since multiple focal
images were superimposed to generate this picture, plasma-membrane-localized
desmosomes are seen in different peripheral domains of the rounded cell
surface contours. The live cell imaging in Movie 5
(jcs.biologist.org/supplemental
) presenting Dsc2a.YFP fluorescence in red and HK18.CFP fluorescence in green
reveals that desmosome-attached CKFs move in synchrony with their respective
adhesion sites (best resolved at top cell margin).
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Desmosomes are maintained during the entire cell cycle but show signs
of destabilization during mitosis
To characterize the dynamic aspects of desmosomes further, fluorescence
emitted by the desmosomal cadherin chimeras was monitored throughout the life
cycle of individual cells and their progeny. A day-long recording of a
confluent MDc-2 cell culture is presented in
Fig. 6 and Movie 6 (available
at
jcs.biologist.org
). Each image presents a projection of seven focal planes. Nine cell divisions
took place during this time within the observation field, resulting in major
rearrangements that were accompanied by changes in cell shape and movement of
cells in and out of the recording area. Nevertheless, despite these pronounced
dynamic processes, desmosomes were always visible as distinct entities in all
cells. Furthermore, individual desmosomes could be traced for very long
periods of time (two examples are labeled by arrows).
|
The pronounced cell-shape changes that occurred during mitosis made it difficult to follow individual desmosomes throughout cell division. However, some details that were consistently observed are demonstrated in Fig. 7 and Movie 7 (available at jcs.biologists.org ), which presents projections of multiple focal recordings of a mitotic cell at high magnification (compare with Fig. 6 and Movie 6 (jcs.biologists.org/supplemental )). During prometaphase, diffuse fluorescence increased considerably, either as a consequence of the dispersion of chimeric polypeptides (compare 0 minutes with 265 minutes) and/or as a result of the altered cell shape. In addition, many desmosomes fused, thereby generating large fluorescent plaques (arrows at 230 minutes and 265 minutes). During cytokinesis, an enrichment of desmosomal fluorescence was noted around the region of the cleavage furrow (arrows at 285 minutes). After completion of cytokinesis, the diffuse fluorescence was rapidly lost and the finely punctate desmosomal fluorescence was re-established, although large plaques remained (arrow in 385 minutes). It should be stressed that we did not observe the complete loss of desmosomal cell contacts at any point during mitosis in any of our recordings.
|
To characterize the relative distribution and organization patterns of
desmosomes and the CKF cytoskeleton during mitosis, we also performed double
fluorescence microscopy of dividing MDc-2 cells that were transiently
transfected with constructs coding for fluorescent CK chimeras. The mitotic
CKF alterations in MDCK cells differ from those previously reported for A-431
cells (Windoffer and Leube,
1999; Windoffer and Leube,
2001
) in that the CKF network is not completely disassembled into
soluble subunits and granular aggregates but is, instead, compacted for the
most part into densely bundled material. This material subsequently separates
during cytokinesis into two parts that are distributed into the daughter cells
from which new networks are formed. In MDc- 2 cells expressing HK18-CFP some
peripheral CKFs were identified that remained in association with desmosomes
during cell division (Fig. 8)
(Movie 8 at
jcs.biologists.org/supplemental
).
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Desmosome stability is lost upon calcium reduction resulting in the
irreversible uptake of desmosomal particles but retaining desmosomal cadherins
with desmosome-forming capacity
In the next set of experiments, we studied the effect of calcium reduction
on desmosome dynamics. MDc-2 cells started to contract within a few minutes of
transfer from SCM to LCM and rounded up progressively. Fluorescent puncta were
translocated into the cell interior where they formed a circumferential ring
around the nucleus (Fig. 9A)
(Movie 9 at
jcs.biologists.org/supplemental
). These puncta colocalized with desmoplakin and Dsg (not shown), suggesting
that complex structures were taken up and not just individual polypeptides.
Remarkably, some diffuse Dsc2a. YFP fluorescence was detected at the cell
surface (+15 minutes and +64 minutes in
Fig. 9A). Within a few hours,
the cytoplasmic dots lost their fluorescence completely. Calcium reduction
also led to the collapse of the CKF cytoskeleton and to the formation of a
dense perinuclear ring of actin filaments, whereas microtubules were not
visibly affected (not shown). We could further demonstrate, in wild-type MDCK
cells, that endocytosis took place independently of the CKF system but
required an intact actin system (not shown). Endocytosis of desmosomal
fluorescence could not be induced in PDc-13 cells and was not observed in
MDc-2 cells more than 3 days after plating. However, fusion of desmosomes into
larger structures and, conversely, the separation into smaller particles,
occurred much more frequently in these cells in LCM than in SCM
(Fig. 9B-D) (Movies 10-12;
available at
jcs.biologists.org/supplemental
), indicating that desmosome stability was also considerably compromised by
calcium reduction.
|
An attractive implication of the calcium-switch model is the potential
re-utilization of endocytosed desmosomal particles for desmosome re-formation,
as has been suggested for components of other adherens junctions
(Le et al., 1999;
Akhtar and Hotchin, 2001
).
However, in accordance with earlier studies (e.g.
Mattey and Garrod, 1986
;
Garrod et al., 1990
), we did
not observe the retranslocation of endocytosed structures to the cell surface
after cells were returned to normal calcium-containing medium. We speculated
that the uptake of desmosomal particles deep into the cell interior may be
irreversible, whereas this may not be the case at earlier stages. Therefore,
the calcium level was only reduced for a very short period (less than 5
minutes) when endocytosis had not yet occurred and the rounding of the cells
had just started. After the restoration of standard calcium levels,
fluorescent puncta were still taken up into the cell interior but did not
relocate to the cell surface (Fig.
10) (Movie 13 available at
jcs.biologists.org/supplemental
). Instead, small puncta formed at the cell surface starting after 18 minutes;
these increased in number and size over time in the continued presence of
endocytosed material (compare small and large arrows in
Fig. 10). Our observations
suggest that clustered desmosomal material that is taken up by endocytosis is
not the major source for desmosome reformation. Instead, sufficient desmosomal
cadherins appear to be present at the cell surface to facilitate the formation
of new desmosomal adhesion sites. Similar small puncta were also observed in
trypsinized cells after replating (not shown). These puncta formed only very
slowly but, again, not from particulate precursors in the cytoplasm.
|
A considerable proportion of desmosomal cadherins is exchanged within
30 minutes
Fluorescence recovery after photobleaching (FRAP) analyses (for a review,
see Reits and Neefjes, 2001)
were performed to discover whether desmosome stability is reflected by the
long residence times of its constituents or is maintained through the
continuous exchange of its molecular components. To this end, desmosome-rich
contact regions that were located in the middle of continuous MDc-2 monolayers
were subjected to photobleaching. As the continuous cell movements made it
difficult to determine fluorescence in the areas of interest for extended
periods of time, measurements of fluorescence recovery were restricted to the
first 30 minutes after photobleaching. In the example shown in
Fig. 11A, some fluorescence
had re-appeared within 5 minutes following exhaustive bleaching. After 15
minutes, 52% of the original fluorescence had reappeared, rising to 64% by 30
minutes. The percentage of recoverable fluorescence within 30 minutes differed
somewhat between cells (60±20%; n=6). The pooled results of
six independent experiments are presented in
Fig. 11C. The same analyses
were also carried out on PDc-13 cells with similar results
(Fig. 11B,C). The percentage
of recovery after 30 minutes was lower than in MDc-2 cells (36±17%;
n=10) but occurred also in an asymptotic manner. Furthermore, in
recordings with a setup for high spatial resolution, reemergence of
fluorescence was seen at the same spots that had been labeled prior to
bleaching, suggesting that, despite the rapid exchange of subunits, desmosomes
remain in loco (Fig.
11D,E,E').
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Discussion |
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Desmosomes are part of a stable network with inherent flexibility and
elasticity
A major outcome of our study is that desmosomes are extremely stable,
exceeding by far the stability of other cell adhesion structures studied to
date (e.g. Jordan et al.,
1999; Windoffer et al.,
2000
). We can rule out the possibility that the high degree of
stability reflects specific properties of the cell lines used as the rapid and
continuous reorganization of gap junctions has been reported in the same PLC
cell line that was used in this study
(Windoffer et al., 2000
). The
stability of desmosomes is quite remarkable in light of the high cell motility
seen in the monolayers, which are most notable in MDc-2 cells. The time-space
diagrams have revealed that desmosomes are part of an interconnected network
reacting in synchrony to cell-shape changes. An important feature of this
network is its intrinsic elasticity, which allows the stretching and
compression of cell borders without the disruption of the overall arrangement
of desmosomal contact sites. This may explain the way in which desmosomal
adhesion is maintained during the entire cell cycle and even during mitosis
when complete restructuring of the cell takes place. Desmosomes should
therefore be considered as one of the major mechanical intercellular
integrators that function as stabilizers in a continuously dynamic
environment, thereby providing uninterrupted intercellular adhesion, which is
especially needed for tissue coherence in mechanically challenged tissues.
This function is fulfilled in concert with the IF cytoskeleton, which is
organized into a transcellular network by desmosomal adhesion. Consequently,
CKF bundles are seen to be tightly associated with desmosomes during
interphase and also, albeit to a lesser degree, during mitosis. It appears,
however, that different populations of CKFs co-exist: those that are subject
to continuous re-structuring (Windoffer
and Leube, 1999
) and those that are stable throughout the cell
cycle and occur in association with desmosomes (this study). These latter CKFs
may form, together with the more flexible actin-rich cortex, the linkage
between desmosomes, determining their coordinated motility and maintenance
during cell-shape changes (see also Pasdar
and Li, 1993
). This system of stable structures and
interconnecting filaments is reminiscent of the situation in the nucleus where
nuclear pore complexes form, together with the nuclear lamina, an immobile and
extremely stable network with intrinsic elasticity
(Daigle et al., 2001
).
Uptake of complete desmosomal plaque assemblies and clustering of
desmosomal cadherins contribute to regulation of desmosomal adhesion
Live cell imaging of fluorescent protein chimeras has allowed us to
establish precursor/product relationships in single living cells by continuous
fluorescence monitoring, thus extending previous observations in fixed cells.
We have found that desmosomal adhesion is terminated in LCM by the rapid
uptake of desmosomal cadherins into endocytotic structures that have been
characterized in the past not only in MDCK cells but also in a number of other
epithelial cell lines (Kartenbeck et al.,
1982; Mattey and Garrod,
1986
; Kartenbeck et al.,
1991
; Holm et al.,
1993
). These structures correspond to vesicles that contain
complete desmosomal assemblies together with their adhering IFs. Furthermore,
we have noted that the uptake of desmosomal halves is dependent on actin
filaments, which is in accordance with previous observations
(Holm et al., 1993
). The
uptake has been shown to be independent of clathrin and to reach, initially, a
non-lysosomal compartment (Holm et al.,
1993
), although the desmosomal components are ultimately destined
for degradation (Mattey and Garrod,
1986
; Burdett,
1993
). In support of this, the fluorescence of the endocytosed
particles fades over time in our recordings. Our data also strongly suggest
that this endocytotic process is irreversible, even at the earliest stages,
and therefore cannot serve for the direct re-utilization of desmosomal
particles. It remains a possibility, however, that single polypeptide subunits
or small aggregates that are below the detection limit of our system are
recycled and used for the reformation of desmosomes. In addition to the bulk
uptake of desmosomal particles, we have observed that other mechanisms are
also operative in LCM and that, depending on cell type and time after plating,
endocytosis may be totally prevented (see also
Watt et al., 1984
;
Mattey and Garrod, 1986
;
Wallis et al., 2000
). In
particular, we have noted, with the new methods at hand, an increased fusion
and fission of desmosomes, and we take this as an indication of their
destabilization. We propose that, in these instances, conformational changes
of desmosomal cadherins occur and result in the reduction of transcellular
adhesiveness, thereby elevating the motility of desmosomal and/or
half-desmosomal particles within the plasma membrane. Furthermore, our
observation of increased diffuse cell surface staining suggests that some
desmosomal cadherins are released from desmosomes, although it may in part be
caused by delivery of molecules from biosynthetic or any other, yet unknown
compartment. In support, Troyanovsky et al.
(Troyanovsky et al., 1999
)
have reported that the removal of calcium ions results in novel interactions,
most notably in the formation of lateral dimers of desmosomal cadherins with
E-cadherin. Another indication for desmosomal cadherin dispersion within the
plasma membrane in LCM is our observation that new desmosomes are formed soon
after the return of cells to SCM, even when endocytosis has just taken place
and when endocytotic structures remained in the cell interior (see also
Mattey and Garrod, 1986
;
Garrod et al., 1990
).
A different situation is encountered during the formation of new
desmosomes. We have not found evidence for the usage of pre-assembled
particles as direct precursors for the reformation of desmosomes in the
calcium-switch system. In addition, we have observed only a little vesicular
desmosomal cadherin fluorescence when cells are cultivated for extended
periods in LCM (see also Burdett,
1993). This is in agreement with biochemical analyses
demonstrating the increased solubility of desmosomal cadherins in MDCK cells
grown in LCM (Penn et al.,
1987
; Pasdar and Nelson,
1989
), indicating that they are not part of large assemblies under
these conditions. Therefore, all the analyses carried out so far suggest that
desmosomes are formed at the plasma membrane by the maturation of enlarging
particles (e.g. Hennings and Holbrook,
1983
; Watt et al.,
1984
; Pasdar and Nelson,
1989
), which, however, may rely on cytoplasmic pools of desmosomal
plaque components, such as desmoplakins
(Duden and Franke, 1988
;
Pasdar and Nelson, 1988
) and
plakoglobin (Kapprell et al.,
1987
). This notion is supported by our observations of desmosome
formation that occurs after replenishment of calcium in the calcium-switch
system and after trypsinization. Furthermore, other adherens junctions are
also formed at the cell surface from diffuse material that accumulates into
immobile puncta that grow and mature into fully competent adhesion structures
(Adams et al., 1998
). In
conclusion, the movement of large desmosomal particles is only of functional
importance for the bulk removal of split desmosomal halves from the cell
surface under certain conditions, whereas small subunits are of relevance in
most other instances, determining the assembly of desmosomes at the cell
surface and the fine tuning of desmosomal stability. This is in accordance
with in vivo observations on other cell adhesion structures
(Adams et al., 1998
;
Jordan et al., 1999
;
Windoffer et al., 2000
;
Jordan et al., 2001
).
Desmosomal cadherins are subject to continuous turnover
The observed fast exchange of Dsc2a chimeras in SCM was quite unexpected
considering the unusual structural stability of desmosomes. Furthermore,
preliminary FRAP determinations with a fluorescent Dsg2 fusion protein fully
support these data (R.W. and R.E.L., unpublished). In principle, Dsc2a
replenishment in the bleached areas may be fuelled from the cytoplasm, from
non-desmosomal plasma membrane domains and/or from neighboring desmosomes.
Clearly, the cytoplasmic pools cannot account by themselves for the rapid
fluorescence recovery given the weak cytoplasmic staining in most cells.
Furthermore, the long half-lives of the major pools of desmosomal cadherins in
MDCK cells in SCM [20.1±0.4 hours for Dsg1 and 19.6±1.5 hours
for Dsc (probably isoforms 2a and 2b) determined by Penn et al.
(Penn et al., 1987); more than
24 hours for Dsg1, as determined by Pasdar and Nelson
(Pasdar and Nelson, 1989
)]
also show that a biosynthetic compartment cannot be responsible for the rapid
recruitment of desmosomal cadherins. With regard to the last two points, Dsc2a
exists in both a desmosomal pool and non-desmosomal pool, which have been
visualized by fluorescence microscopy in the form of puncta and diffuse
fluorescence, respectively. Although the majority of polypeptides is clearly
concentrated in desmosomes, both pools must be in equilibrium under
steady-state conditions. The best explanation for our current results is that
rapid exchange takes place between both pools, since we have not found any
evidence for vesicular traffic between membrane domains. Furthermore, to
account for the recovery of desmosomal fluorescence at distant sites, the
non-desmosomal pool must be rapidly diffusible. Given the longevity of
desmosomal cadherins (see above), one has to propose that polypeptides are
exchanged in and out of desmosomes several times during their lifetime.
Exchange rates apparently differ between cell lines and may be influenced by
certain, as yet unknown, factors. Currently, it also remains a possibility
that desmosomal cadherins with different exchange kinetics co-exist within
individual desmosomes. Such an intrinsic heterogeneity could result from
differences in packaging density, association with the cytoskeleton, different
localization within the plaque and/or protein modification, as has been
described for the gapjunction-localized connexins (e.g.
Windoffer et al., 2000
,
Rütz and Hülser,
2001
). The attractive implication of the reported FRAP results is
the idea that desmosomal stability and durability is combined with a pathway
that could serve to adjust desmosomal adhesion rapidly to specific
requirements, for example, reversible phosphorylation
(Pasdar et al., 1995
;
van Hengel et al., 1997
;
Wallis et al., 2000
;
Gaudry et al., 2001
).
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Acknowledgments |
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Footnotes |
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References |
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Adams, C. L., Chen, Y.-T., Smith, S. J. and Nelson, W. J.
(1998). Mechanisms of epithelial cell-cell adhesion and cell
compaction revealed by high-resolution tracking of E-cadherin-green
fluorescent protein. J. Cell Biol.
142,1105
-1119.
Akhtar, N. and Hotchin, N. A. (2001). RAC1
regulates adherens junctions through endocytosis of E-cadherin.
Mol. Biol. Cell 12,847
-862.
Bader, B. L., Magin, T. M., Freudenmann, M., Stumpp, S. and Franke, W. W. (1991). Intermediate filaments formed de novo from tail-less cytokeratins in the cytoplasm and in the nucleus. J. Cell Biol. 115,1293 -1307.[Abstract]
Baker, J. and Garrod, D. (1993). Epithelial
cells retain junctions during mitosis. J. Cell Sci.
104,415
-425.
Bonné, S., van Hengel, J., Nollet, F., Kools, P. and van
Roy, F. (1999). Plakophilin-3, a novel armadillo-like protein
present in nuclei and desmosomes of epithelial cells. J. Cell
Sci. 112,2265
-2276.
Burdett, I. D. J. (1993). Internalisation of
desmosomes and their entry into the endocytic pathway via late endosomes in
MDCK cells. Possible mechanisms for the modulation of cell adhesion by
desmosomes during development. J. Cell Sci.
106,1115
-1130.
Burdett, I. D. J. (1998). Aspects of the structure and assembly of desmosomes. Micron 29,309 -328.[Medline]
Daigle, N., Beaudouin, J., Hartnell, L., Imreh, G., Hallberg,
E., Lippincott-Schwartz, J. and Ellenberg, J. (2001). Nuclear
pore complexes form immobile networks and have a very low turnover in live
cells. J. Cell Biol.
154, 71-84.
Demlehner, M. P., Schäfer, S., Grund, C. and Franke, W. W. (1995). Continual assembly of half-desmosomal structures in the absence of cell contacts and their frustrated endocytosis: a coordinated Sisyphus cycle. J. Cell Biol. 131,745 -760.[Abstract]
Duden, R. and Franke, W. W. (1988). Organization of desmosomal plaque proteins in cells growing at low calcium concentrations. J. Cell Biol. 107,1049 -1063.[Abstract]
Gaudry, C. A., Palka, H. L., Dusek, R. L., Huen, A. C.,
Khandekar, M. J., Hudson, L. G. and Green, K. J. (2001).
Tyrosine-phosphorylated plakoglobin is associated with desmogleins but not
desmoplakin after epidermal growth factor receptor activation. J.
Biol. Chem. 276,24871
-24880.
Garrod, D. R., Parrish, E. P., Mattey, D. L., Marston, J. E., Measures, H. R. and Vilela, M. J. (1990). Desmosomes. In Intercellular Junctions and Cell Adhesion in Epithelial Cells (eds G. M. Edelman, B. A. Cunningham and J.-P. Thiery), pp.315 -339. Chichester: John Wiley.
Hennings, H. and Holbrook, K. A. (1983). Calcium regulation of cell-cell contact and differentiation of epidermal cells in culture. Exp. Cell Res. 143,127 -142.[Medline]
Holm, P. K., Hansen, S. H., Sandvig, K. and van Deurs, B. (1993). Endocytosis of desmosomal plaques depends on intact actin filaments and leads to a nondegradative compartment. Eur. J. Cell Biol. 62,362 -371.[Medline]
Holm, I., Mikhailov, A., Jillson, T. and Rose, B. (1999). Dynamics of gap junctions observed in living cells with connexin43-GFP chimeric protein. Eur. J. Cell Biol. 78,856 -866.[Medline]
Ishii, K., Norvell, S. M., Bannon, L. J., Amargo, E. V., Pascoe,
L. T. and Green, K. J. (2001). Assembly of desmosomal
cadherins into desmosomes is isoform dependent. J. Invest.
Dermatol. 117,26
-35.
Jordan, K., Solan, J. L., Dominguez, M., Sia, M., Hand, A.,
Lampe, P. D. and Laird, D. W. (1999). Trafficking, assembly,
and function of a connexin43-green fluorescent protein chimera in live
mammalian cells. Mol. Biol. Cell
10,2033
-2050.
Jordan, K., Chodock, R., Hand, A. R. and Laird, D. W.
(2001). The origin of annular junctions: a mechanism of gap
junction internalization. J. Cell Sci.
114,763
-773.
Kapprell, H.-P., Cowin, P. and Franke, W. W. (1987). Biochemical characterization of the soluble form of the junctional plaque protein, plakoglobin, from different cell types. Eur. J. Cell Biol. 166,505 -517.
Karnovsky, A. and Klymkowsky, M. W. (1995). Over-expression of plakoglobin leads to dorsalization and axis duplication in Xenopus. Proc. Natl. Acad. Sci. USA 92,4522 -4526.[Abstract]
Kartenbeck, J., Schmid, E., Franke, W. W. and Geiger, B. (1982). Different modes of internalization of proteins associated with adhaerens junctions and desmosomes: experimental separation of lateral contacts induces endocytosis of desmosomal plaque material. EMBO J. 1,725 -732.[Medline]
Kartenbeck, J., Schmelz, M., Franke, W. W. and Geiger, B. (1991). Endocytosis of junctional cadherins in bovine kidney epithelial (MDBK) cells cultured in low Ca2+ ion medium. J. Cell Biol. 113,881 -892.[Abstract]
Klymkowsky, M. W. (1999). Plakophilin, armadillo repeats, and nuclear localization. Microsc. Res. Tech. 45,43 -54.[Medline]
Kowalczyk, A. P., Palka, H. L., Luu, H. H., Nilles, L. A., Anderson, J. E., Wheelock, M. J. and Green, K. J. (1994). Posttranslational regulation of plakoglobin expression. Influence of the desmosomal cadherins on plakoglobin metabolic stability. J. Biol. Chem. 49,31214 -31223.
Kowalczyk, A. P., Bornslaeger, E. A., Norvell, S. M., Palka, H. L. and Green, K. J. (1999). Desmosomes: intercellular adhesive junctions specialized for attachment of intermediate filaments. Int. Rev. Cytol. 185,237 -302.[Medline]
Le, T. L., Yap, A. S. and Stow, J. L. (1999).
Recycling of E-cadherin: a potential mechanism for regulating cadherin
dynamics. J. Cell Biol.
146,219
-232.
Leube, R. E. (1995). The topogenic fate of the
polytopic transmembrane proteins, synaptophysin and connexin, is determined by
their membrane-spanning domains. J. Cell Sci.
108,883
-894.
Mattey, D. L. and Garrod, D. R. (1986). Splitting and internalization of the desmosomes of cultured kidney epithelial cells by reduction in calcium concentration. J. Cell Sci. 85,113 -124.[Abstract]
Mertens, C., Kuhn, C. and Franke, W. W. (1996). Plakophilins 2a and 2b: constitutive proteins of dual location in the karyoplasm and the desmosomal plaque. J. Cell Biol. 135,1009 -1025.[Abstract]
Mertens, C., Hofmann, I., Whang, Z., Teichmann, M., Chong, S.
S., Schnôlzer, M. and Franke, W. W. (2001). Nuclear
particles containing RNA polymerase III complexes associated with the
junctional plaque protein plakophilin 2. Proc. Natl. Acad. Sci.
USA 98,7795
-7800.
Moll, R. and Moll, I. (1998). Epidermal adhesion molecules and basement membrane components as target structures of autoimmunity. Virch. Arch. 432,487 -504.
North, A. J., Bardsley, W. G., Hyam, J., Bornslaeger, E. A.,
Cordingley, H. C., Trinnaman, B., Hatzfeld, M., Green, K. J., Magee, A. I. and
Garrod, D. R. (1999). Molecular map of the desmosomal plaque.
J. Cell Sci. 112,4325
-4336.
Norvell, S. M. and Green, K. J. (1998).
Contributions of extracellular and intracellular domains of full length and
chimeric cadherin molecules to junction assembly in epithelial cells.
J. Cell Sci. 111,1305
-1318.
Oda, H. and Tsukita, S. (1999). Dynamic
features of adherens junctions during Drosophila embryonic epithelial
morphogenesis revealed by a D-catenin-GFP fusion protein.
Dev. Genes Evol. 209,218
-225.[Medline]
Parker, A. E., Wheeler, G. N., Arnemann, J., Pidsley, S. C.,
Ataliotis, P., Thomas, C. L., Rees, D. A., Magee, A. I. and Buxton, R. S.
(1991). Desmosomal glycoproteins II and III. Cadherin-like
junctional molecules generated by alternative splicing. J. Biol.
Chem. 266,10438
-10445.
Pasdar, M. and Li, Z. (1993). Disorganization of microfilaments and intermediate filaments interferes with the assembly and stability of demosomes in MDCK cells. Cell Motil. Cytoskeleton 26,163 -180.[Medline]
Pasdar, M. and Nelson, W. J. (1988). Kinetics of desmosome assembly in Madin-Darby canine kidney epithelial cells: temporal and spatial regulation of desmoplakin organization and stabilization upon cell-cell contact. I. Morphological analysis. J. Cell Biol. 106,687 -695.[Abstract]
Pasdar, M. and Nelson, W. J. (1989). Regulation of desmosome assembly in epithelial cells: kinetics of synthesis, transport, and stabilization of desmoglein I, a major protein of the membrane core domain. J. Cell Biol. 109,163 -177.[Abstract]
Pasdar, M., Li, Z. and Chan, H. (1995). Desmosome assembly and disassembly are regulated by reversible protein phosphorylation in cultured epithelial cells. Cell Motil. Cytoskeleton 30,108 -121.[Medline]
Penn, E. J., Burdett, I. D. J., Hobson, C., Magee, A. I. and Rees, D. A. (1987). Structure and assembly of desmosome junctions: biosynthesis and turnover of the major desmosome components of Madin-Darby kidney cells in low calcium medium. J. Cell Biol. 105,2327 -2334.[Abstract]
Reits, E. A. J. and Neefjes, J. J. (2001). From fixed to FRAP: measuring protein mobility and activity in living cells. Nat. Cell Biol. 3,E145 -E147.[Medline]
Rütz, M. L. and Hülser, D. F. (2001). Supramolecular dynamics of gap junctions. Eur. J. Cell Biol. 80,20 -30.[Medline]
Schäfer, S., Koch, P. J. and Franke, W. W. (1994). Identification of the ubiquitous human desmoglein, Dsg2, and the expression catalogue of the desmoglein subfamily of desmosomal cadherins. Exp. Cell Res. 211,391 -399.[Medline]
Schmidt, A., Heid, H. W., Schäfer, S., Nuber, U. A., Zimbelmann, R., Franke, W. W. (1994). Desmosomes and cytoskeletal architecture in epithelial differentiation: cell type-specific plaque components and intermediate filament anchorage. Eur. J. Cell Biol. 65,229 -245.[Medline]
Schmidt, A., Langbein, L., Rode, M., Prätzel, S., Zimbelmann, R. and Franke, W. W. (1997). Plakophilins 1a and 1b: widespread nuclear proteins recruited in specific epithelial cells as desmosomal plaque components. Cell Tissue Res. 290,481 -499.[Medline]
Simcha, I., Shtutman, M., Salomon, D., Zhurinsky, J., Sadot, E.,
Geiger, B. and Ben-Ze'ev, A. (1998). Differential nuclear
translocation and transactivation potential of ß-catenin and plakoglobin.
J. Cell Biol. 141,1433
-1448.
Stappenbeck, T. S., Bornslaeger, E. A., Corcoran, C. M., Luu, H. H., Virata, M. L. and Green, K. J. (1993). Functional analysis of desmoplakin domains: specification of the interaction with keratin versus vimentin intermediate filament networks. J. Cell Biol. 123,691 -705.[Abstract]
Strnad, P., Windoffer, R. and Leube, R. E. (2001). In vivo detection of cytokeratin filament network breakdown in cells treated with the phosphatase inhibitor okadaic acid.Cell Tissue Res. 306,277 -293.[Medline]
Troyanovsky, S. M. and Leube, R. E. (1998). Molecular dissection of desmosomal assembly and intermediate filament anchorage. In Intermediate Filaments (ed. H. Herrmann and J. R. Harris), pp. 263-289. New York: Plenum Press.
Troyanovsky, R. B., Klingelhöfer, J. and Troyanovsky,
S. (1999). Removal of calcium ions triggers a novel type of
intercadherin interaction. J. Cell Sci.
112,4379
-4387.
Udey, M. C. and Stanley, J. R. (1999).
Pemphigus Diseases of antidesmosomal autoimmunity.
JAMA 282,572
-576.
van Hengel, J., Gohon, L., Bruyneel, E., Vermeulen, S.,
Cornelissen, M., Mareel, M. and van Roy, F. (1997). Protein
kinase C activation upregulates intercellular adhesion of
ß-catenin-negative human colon cancer cell variants via induction of
desmosomes. J. Cell Biol.
137,1103
-1116.
Wallis, S., Lloyd, S., Wise, I., Ireland, G., Fleming, T. P. and
Garrod, D. (2000). The isoform of protein kinase C is
involved in signaling the response of desmosomes to wounding in cultured
epithelial cells. Mol. Biol. Cell
11,1077
-1092.
Watt, F. M., Mattey, D. L. and Garrod, D. R. (1984). Calcium-induced reorganization of desmosomal components in cultured human keratinocytes. J. Cell Biol. 99,2211 -2215[Abstract]
Windoffer, R. and Leube, R. E. (1999).
Detection of cytokeratin dynamics by time-lapse fluorescence microscopy in
living cells. J. Cell Sci.
112,4521
-4534.
Windoffer, R. and Leube, R. E. (2001). De novo formation of cytokeratin filament networks originates from the cell cortex in A-431 cells. Cell Motil. Cytoskeleton 50, 33-44.[Medline]
Windoffer, R., Beile, B., Leibold, A., Thomas, S., Wilhelm, U. and Leube, R. E. (2000). Visualization of gap junction mobility in living cells. Cell Tissue Res. 299,347 -362.[Medline]