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The Journal of Neuroscience, March 15, 2002, 22(6):2106-2114
Microtubule-Associated Protein 1A (MAP1A) and MAP1B: Light Chains
Determine Distinct Functional Properties
Rainer
Noiges,
René
Eichinger,
Waltraud
Kutschera,
Irmgard
Fischer,
Zsuzsanna
Németh,
Gerhard
Wiche, and
Friedrich
Propst
Institute of Biochemistry and Molecular Cell Biology, Vienna
Biocenter, University of Vienna, A-1030 Vienna, Austria
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ABSTRACT |
The microtubule-associated proteins 1A (MAP1A) and 1B (MAP1B) are
distantly related protein complexes consisting of heavy and light
chains and are thought to play a role in regulating the neuronal
cytoskeleton, MAP1B during neuritogenesis and MAP1A in mature neurons.
To elucidate functional differences between MAP1B and MAP1A and to
determine the role of the light chain in the MAP1A protein complex, we
chose to investigate the functional properties of the light chain of
MAP1A (LC2) and compare them with the light chain of MAP1B (LC1). We
found that LC2 binds to microtubules in vivo and
in vitro and induces rapid polymerization of tubulin. A
microtubule-binding domain in its NH2 terminus was found to
be necessary and sufficient for these activities. The analysis of LC1
revealed that it too bound to microtubules and induced tubulin
polymerization via a crucial but structurally unrelated
NH2-terminal domain. The two light chains differed, however, in their effects on microtubule bundling and stability in vivo. Furthermore, we identified actin filament
binding domains located at the COOH terminus of LC2 and LC1 and
obtained evidence that binding to actin filaments is
attributable to direct interaction with actin. Our findings
establish LC2 as a crucial determinant of MAP1A function, reveal LC2 as
a potential linker of neuronal microtubules and microfilaments, and
suggest that the postnatal substitution of MAP1B by MAP1A leads to
expression of a protein with an overlapping but distinct set of functions.
Key words:
actin; brain development; microtubule-associated
proteins; microtubule stability; neuronal cytoskeleton; tubulin
polymerization
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INTRODUCTION |
The high-molecular-mass
microtubule-associated proteins 1A (MAP1A) and 1B (MAP1B) are expressed
predominantly in cells of the nervous system (Wiche et al., 1991 ;
Schoenfeld and Obar, 1994). Expression of MAP1B is high during early
stages of neuronal development and is downregulated in the adult
(Binder et al., 1984; Calvert and Anderton, 1985; Lewis et al., 1986;
Riederer et al., 1986; Safaei and Fischer, 1989; Schoenfeld et al.,
1989; Garner et al., 1990). MAP1A exhibits a reciprocal pattern of
expression, reaching its peak in the adult brain, when neuronal
differentiation is complete (Schoenfeld et al., 1989; Garner et al.,
1990). MAP1B has been shown to play an important role in neuronal
differentiation in vitro (Brugg et al., 1993; DiTella et
al., 1996; Gonzalez-Billault et al., 2001) and in the development of
the murine nervous system in vivo (Edelmann et al., 1996;
Takei et al., 1997; González-Billault et al., 2000; Meixner et
al., 2000). There are no comparable data for MAP1A.
Both MAP1A and MAP1B are multimeric protein complexes containing one
heavy and several light chains; they share isolated domains of sequence
homology in their subunits (Hammarback et al., 1991; Langkopf et al.,
1992). In each case, the heavy chain and one of the light chains are
generated by proteolytic cleavage of the respective MAP1A or MAP1B
polyprotein precursor. These subunits are interchangeable (Schoenfeld
et al., 1989). Thus, the MAP1B light chain, LC1, can bind to both MAP1A
and MAP1B heavy chains. The same is true for the MAP1A light chain,
LC2, although it binds predominantly to the heavy chain of MAP1A.
Moreover, there is evidence that the light chains might have additional
functions outside of the complex with the heavy chains. It was found
that LC1 is expressed at levels in excess over what can be complexed by
heavy chains (Mei et al., 2000b).
MAP1A and MAP1B bind to microtubules (Schoenfeld and Obar, 1994) and
microfilaments (Asai et al., 1985; Pedrotti et al., 1994; Pedrotti and
Islam, 1996; Tögel et al., 1998b), suggesting that they are
involved in mediating or regulating the interaction between axonal
microtubules and actin filaments, which is believed to be essential for
neuronal morphogenesis and function. Association of MAP1B with
microtubules is mediated by two unique microtubule-binding domains,
located on the heavy chain (Noble et al., 1989) and light chain (Zauner
et al., 1992; Tögel et al., 1998b), respectively. For MAP1A, two
unrelated regions, both located in the heavy chain, have been
implicated in microtubule binding (Cravchik et al., 1994; Vaillant et
al., 1998). The role of the light chain for MAP1A interaction with
microtubules and microfilaments has not been determined, and it is not
clear whether MAP1A and MAP1B interact directly with actin or
actin-binding proteins. To clarify these questions and to determine the
contribution of the light chains to MAP1A and MAP1B function, we
analyzed properties of the light chains by biochemical and cell
biological techniques in vivo and in vitro.
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MATERIALS AND METHODS |
cDNA constructs. LC2 cDNA was obtained by reverse
transcription PCR using the primers LC2U (5'-CGGAGTCGACCATG
GCTGACCCTGAGGGG-3') and LC2L
(5'-CGCATCTAGAGCTAGCGTGAACTCAATCTTGCAGGC-3'). The correct sequence
of this cDNA was confirmed. It encodes amino acids 2554-2774 of rat
MAP1A (Langkopf et al., 1992). Constructs encoding full-length LC2
(amino acids 2554-2774), its NH2 terminus (amino
acids 2554-2659), or its COOH terminus (amino acids 2650-2774) all
fused in frame to an NH2- or COOH-terminal
myc-tag [amino acid sequence: EQKLISEEDLN (Cravchik and Matus, 1993)]
by a linker of three amino acids were generated using convenient
restriction sites and adapter oligonucleotides. Constructs encoding
full-length LC1 and its NH2- and COOH-terminal domains have been described previously (Tögel et al., 1998b). All
constructs were cloned into the mammalian Tet-Off expression vector
pUHD10-3 (Gossen and Bujard, 1992) and into a pET15b (Novagen, Inc.,
Madison, WI) or pQE60 (Qiagen, Hilden, Germany) derivative for
the expression of NH2- or COOH-terminal
6xHis-tagged proteins in Escherichia coli. The authenticity
of all constructs was confirmed by sequencing and/or reaction of
encoded proteins with LC1- or LC2-specific antisera.
Antibodies. A rabbit polyclonal anti-LC2 antiserum was
raised against the synthetic peptide CKGPVDRTSRTVPRPR (MAP1A amino acids 2605-2619; Gramsch, Schwabhausen, Germany). The anti-LC1 antiserum has been described previously (Tögel et al., 1998b). An
affinity-purified polyclonal rabbit anti-myc antibody (Tögel et
al., 1998a) was used at a concentration of 1 µg/ml for
immunofluorescence microscopy. Intracellular microtubules were detected
with mouse anti-tubulin monoclonal antibody B-5-1-2 (Sigma, St. Louis,
MO) at a dilution of 1:500, and actin stress fibers were
detected using a mixture of mouse anti- -actin monoclonal antibodies
AC-15 and AC-74 (Sigma) at a dilution of 1:200. Texas Red-labeled
anti-mouse and FITC-labeled anti-rabbit secondary antibodies (Jackson
ImmunoResearch Laboratories, Inc., West Grove, PA) were used at
dilutions of 1:100.
Cell culture, DNA transfection, and immunofluorescence
microscopy. PtK2 cells were grown at 37°C in an atmosphere
containing 8.5% CO2 in high glucose DMEM
supplemented with 10% FCS in the absence of tetracycline. At 18-20 hr
before transfection, cells were seeded onto coverslips at a density of
66%. Transient transfection was performed using Lipofectamine
(Invitrogen, Bethesda, MD) according to the manufacturer's
protocol. The ratio of expression plasmid to transactivator plasmid was
10:1. The transfection mixture was replaced with fresh growth medium
after 5 hr, and the cells were incubated for an additional 48 hr at
37°C.
For immunofluorescence microscopy, PtK2 cells were washed in PBS, fixed
in methanol ( 20°C, 10 min), equilibrated in PBS, blocked for 30 min
with 3% BSA, incubated with the primary antibodies in 1% BSA for 1 hr, washed extensively in PBS, incubated for 1 hr with the secondary
antibodies, and washed again with PBS. Specimens were analyzed by
confocal microscopy using a Zeiss Axiovert microscope (Carl Zeiss,
Oberkochen, Germany).
For quantitative analysis, experiments were chosen in which expression
of both constructs was equal as judged by immunofluorescence intensity.
The presence of stable microtubules was assessed in 100 randomly chosen
transfected cells in at least two independent experiments.
Purification of recombinant proteins. Recombinant proteins
were expressed in E. coli BL21-CodonPlus-RIL or E. coli XL1-Blue and purified by affinity chromatography on
Ni2+ columns, according to the
manufacturers' protocols (Novagen, Inc.; Qiagen). Recombinant proteins
were bound to and eluted from the column in the presence of 6 M urea, except for the LC1
NH2 terminus, which was purified in the absence
of urea. Before biochemical analysis, recombinant proteins were
extensively dialyzed at 4°C against buffers used in the respective
in vitro assays. Protein concentrations were determined
according to the method of Bradford (1976) using BSA as a standard.
Purification of tubulin. Tubulin devoid of
microtubule-associated proteins was prepared as described previously
(Karr et al., 1979), with the following modifications. Polymerization
of tubulin was performed in PEM buffer (PIPES 0.1 M, EGTA 2 mM,
MgCl2 1 mM, pH 6.8)
containing 0.1 mM GTP. Centrifugation steps were
performed at 100,000 × g in a Beckman TLX tabletop
ultracentrifuge (Beckman Instruments, Inc., Palo Alto, CA). The last
cycle of polymerization/depolymerization was followed by chromatography
on a phosphocellulose column, as described previously (Weinert et al.,
1982). After precipitation, the protein was resuspended in PEM buffer.
Protein purity was monitored using SDS-PAGE and Coomassie blue staining.
Microtubule cosedimentation. Cosedimentation assays were
performed as described previously (Feick et al., 1991), with slight modifications. In the presence of 5 nM taxol,
tubulin was polymerized for 30 min at 37°C. LC1 and LC2 proteins were
dialyzed against 0.1 M PEM buffer and centrifuged
for 30 min at 100,000 × g to remove any aggregates.
Then 50 µg of polymerized tubulin was incubated with 10 µg of LC1
or LC2 proteins for 10 min at 37°C. Samples were layered onto a 10%
sucrose cushion and centrifuged for 20 min at 10,000 rpm in a tabletop
centrifuge. Equal volumes of supernatants and pellets were analyzed by
SDS-PAGE and Coomassie blue staining.
Turbidity assay. Polymerization of tubulin was performed as
described previously (Weinert et al., 1982), with the following modifications. Proteins dialyzed against 0.1 M
PEM buffer were mixed with 1 mM GTP and 1.5 mg/ml
tubulin. Polymerization was initiated by placing the mixture
immediately into 37°C prewarmed 0.25 ml cuvettes, and turbidity
change was followed at 350 nm using a Hitachi (Tokyo, Japan) U-3000
spectrophotometer equipped with a thermostat.
Europium overlay binding assay. Binding assays were
performed as described previously (Steinböck et al., 2000).
Briefly, LC1 and LC2 proteins were dialyzed against
Eu3+-labeling buffer. Ninety-six well
microtiter plates were coated with 100 nM actin
(rabbit skeletal muscle actin; Cytoskeleton, Inc., Denver, CO)
or BSA type H1 (Gerbu, Gaiberg, Germany) as a control. After blocking
with 4% BSA, plates were overlaid with increasing amounts of
Eu3+-labeled LC1 or LC2 proteins. Plates
were washed, and bound protein was determined by releasing the
complexed Eu3+ with enhancement solution
and measuring fluorescence with a Delfia time-resolved fluorometer
(Wallac, Turku, Finland). Binding of LC1 or LC2 proteins to BSA was
considered to be nonspecific and was subtracted from protein bound to
actin. Counts were converted to concentrations by comparison with a 1 nM Eu3+ standard.
Actin cosedimentation. LC1 and LC2 proteins were dialyzed
against 0.1 M MES buffer and centrifuged for 30 min at 100,000 × g to remove any aggregates. Actin
(rabbit skeletal muscle actin; Cytoskeleton, Inc.) was polymerized
(Pedrotti and Islam, 1996) and incubated with 3 µg of recombinant LC1
or LC2 protein. Samples were centrifuged at 100,000 × g for 30 min at 37°C, and equal volumes of supernatants
and pellets were analyzed by SDS-PAGE and Coomassie blue staining.
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RESULTS |
The MAP1A light chain binds and stabilizes microtubules
in vivo
Microtubule association of the MAP1A complex has thus far been
attributed to microtubule-binding domains located in the heavy chain
(Cravchik et al., 1994; Vaillant et al., 1998). We decided to
investigate the potential involvement of the MAP1A light chain in
microtubule binding. We used PtK2 cells, which have been shown previously to express low levels of LC1 (Tögel et al., 1998b) that faintly decorated their microtubules (data not shown).
These cells also expressed low levels of LC2 in the cytoplasm and the nucleus (data not shown). PtK2 cells were transiently transfected with
a cDNA construct encoding the full-length LC2 protein and analyzed by
double immunofluorescence microscopy. In cells expressing LC2, the
myc-tagged protein was found to colocalize with microtubules, which
were decorated by LC2 in a punctate manner (Fig.
1A,B). The merge of the
confocal images obtained with anti-myc and anti-tubulin antibodies
revealed that not all microtubules were decorated to the same extent.
The decoration was more pronounced toward the perinuclear region (Fig.
1C). Whereas expression of LC1 induces severe changes in
microtubule organization (Tögel et al., 1998b), binding of LC2 to
microtubules did not affect their appearance and organization (Fig. 1,
compare B and E). Microtubules in LC2-expressing cells were indistinguishable from those in untransfected cells (data
not shown). In contrast, many microtubules in LC1-expressing cells were
organized into thick, wavy bundles, some of which are pointed out by
arrows (Fig. 1E). Staining with the anti-myc antibody also revealed nuclear localization of LC2 (Fig. 1A).
This was also observed for endogenous LC2 in untransfected cells (data not shown) and in LC2-transfected NIH3T3 cells (Mei et al., 2000a). Microtubule association of LC1 and LC2 is unlikely to be affected by
the presence of the myc-tag, because untagged LC1 yielded identical results (data not shown). Neither LC1 nor LC2 overexpression in PtK2
cells induced gross morphological changes.
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Figure 1.
The LC2 protein binds to microtubules in
vivo. Shown are confocal images of PtK2 cells expressing
myc-tagged LC2 (A-C) or myc-tagged LC1
(D-F), analyzed by double immunofluorescence
microscopy using antibodies against the myc-tag (A, D)
and tubulin (B, E). LC2 and LC1 colocalized with
microtubules (arrows), but LC2 did not induce formation
of wavy microtubule bundles like LC1 (compare B and
E). C and F represent
merged confocal images of A + B and
D + E, respectively. Scale bar, 10 µm.
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On the basis of these findings, we further analyzed whether expression
of LC2 altered microtubule stability. Treatment of cells with the
microtubule depolymerizing agents colchicine and nocodazole before
fixation led to destruction of the cellular microtubule network,
resulting in a diffuse distribution of tubulin throughout the cytoplasm
(Figs. 2D,
3D). In cells expressing LC2, we observed integrity of the cellular microtubules despite treatment with colchicine (Fig. 2A,B). The merge of the
confocal images obtained with anti-myc and anti-tubulin antibodies
revealed that stable microtubules were decorated with LC2 (Fig.
2C). This is consistent with the assumption that binding of
LC2 is necessary for stabilization. The same was observed for cells
expressing LC1 (Fig. 2E-G). Quantitative analysis of
these experiments and comparison with LC1 revealed that both proteins,
LC1 and LC2, were equally efficient in protecting cellular microtubules
against the depolymerizing effects of colchicine (Fig.
2H). In cells expressing LC1 or LC2, intact
microtubules were observed in a subpopulation of 27 and 28%,
respectively, despite colchicine treatment.
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Figure 2.
Effects of LC2 and LC1 on microtubule stability in
cells treated with colchicine. Nontransfected PtK2 cells
(NT, D) or PtK2 cells transfected with
myc-tagged LC2 (A-C) or myc-tagged LC1
(E-G) were treated with colchicine (1-2 hr, 10 µM) and then analyzed by double immunofluorescence
microscopy using antibodies against tubulin (B, D,
F) and the myc-tag (A, E). Microtubules
were depolymerized in nontransfected cells (D),
whereas intact microtubules were found in cells expressing LC2
(B) and LC1 (F).
C and G represent merged confocal images
of A + B and E + F, respectively. Scale bars, 20 µm. H,
One hundred randomly chosen transfected cells were assessed for the
presence of intact microtubules (MTs). Quantitative
analysis revealed that LC1 and LC2 were equally efficient in protecting
microtubules against depolymerization (, untransfected cells).
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Figure 3.
Effects of LC2 and LC1 on microtubule
stability in cells treated with nocodazole. Nontransfected PtK2 cells
(NT, D) or PtK2 cells transfected with
myc-tagged LC2 (A-C) or myc-tagged LC1
(E-G) were treated with nocodazole (30-45 min,
10 µg/ml) and then analyzed by double immunofluorescence microscopy
using antibodies against tubulin (B, D, F) and
the myc-tag (A, E). Microtubules were depolymerized in
untransfected cells (D), whereas intact
microtubules were found in cells expressing LC2
(B) and LC1 (F).
C and G represent merged confocal images
of A + B and E + F, respectively. Scale bar, 20 µm. H,
One hundred randomly chosen transfected cells were assessed for the
presence of intact microtubules (MTs). Quantitative
analysis revealed that LC1 was considerably more efficient than LC2 in
protecting microtubules against depolymerization (, untransfected
cells).
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In contrast, in cells treated with nocodazole, the protective effect of
LC1 was much more pronounced than that of LC2 (Fig. 3). With either
light chain, cells with stable microtubules could be found (Fig.
3A-C, E-G), and in each case stable
microtubules were decorated with the respective light chain, albeit at
varying levels (Fig. 3C,G), again supporting the assumption
that binding of the light chains is necessary for stabilization.
However, quantitative analysis revealed that the number of cells with
stable microtubules was greatly diminished in LC2-expressing cells
compared with LC1-expressing cells (Fig. 3H).
The MAP1A light chain binds to microtubules via its
NH2 terminus
Amino acid sequence comparison of light chains LC1 and LC2 reveals
a highly homologous region located at the COOH terminus, whereas no
significant similarity can be found at the
NH2-terminal domains (Langkopf et al., 1992). It
has been shown previously by in vitro cosedimentation assays
that the microtubule-binding domain of LC1 is located in the
NH2-terminal half of the protein (Zauner et al.,
1992). It was surprising to find that LC2, despite the lack of homology
to the microtubule-binding domain of LC1, also bound to microtubules
in vivo. To map the microtubule-binding domain of LC2, we
performed microtubule cosedimentation assays using the full-length LC2
protein and deletion mutants (Fig.
4A). Full-length LC1
protein and LC1 fragments comprising the
NH2-terminal or COOH-terminal domain were used as
controls (Fig. 4A). After incubation with polymerized
microtubules, only LC1 and LC2 proteins containing the
NH2-terminal domain were found to cosediment to a
significant extent (Fig. 4B). Absence of the
NH2-terminal domains of LC1 and LC2 led to
complete loss of microtubule binding. Thus, the COOH-terminal domains
of LC1 and LC2 were found almost exclusively in the supernatant and
therefore were not capable of mediating binding of the light chain
proteins to microtubules. This result also showed that cosedimentation
was specific and not merely attributable to unspecific binding of
His-tag-containing proteins. In control experiments performed in the
absence of microtubules, LC1 and LC2 proteins were recovered primarily
in the supernatant. These results demonstrated the functional
similarity of the NH2-terminal regions of LC1 and
LC2. Despite their structural diversity, the NH2-terminal domains of both proteins act as
equally efficient microtubule-binding domains of the respective light
chain.
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Figure 4.
A, Schematic of MAP1A and MAP1B
heavy and light chains (HC and LC,
respectively) and cDNA constructs used in this study. The scale at the
top displays amino acid (aa) residue
positions. Domains of sequence homology between MAP1A and MAP1B
(hatched boxes) and the microtubule-binding domains
(MT) in the heavy and light chains of MAP1B are
indicated. The cDNA constructs used are depicted as filled
boxes. LC1, Full-length MAP1B light chain (amino
acids 2210-2459); N-term LC1, amino acids 2210-2336;
C-term LC1, amino acids 2335-2459; LC2,
full-length MAP1A light chain (amino acids 2554-2774); N-term
LC2, amino acids 2554-2659; C-term LC2, amino
acids 2650-2774. For transfection studies, proteins were tagged with
an NH2- or COOH-terminal myc peptide. Proteins used for
biochemical analysis were tagged with an NH2- or
COOH-terminal 6xHis tag. B, LC1 and LC2 interact with
microtubules in vitro. LC1 and LC2 proteins
(arrows) were sedimented in the presence (+) or absence
() of polymerized taxol-stabilized microtubules. Equal amounts of
supernatant (S) and pellet
(P) fractions were analyzed by SDS-PAGE and
Coomassie blue staining. Proteins containing the
NH2-terminal microtubule-binding domain of LC1
(N-term LC1 and LC1) and LC2
(N-term LC2 and LC2) were found to
cosediment with tubulin (T), whereas only trace
amounts of the COOH-terminal domains of LC1 (C-term LC1)
and LC2 (C-term LC2) were found in the pellet
fraction.
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LC1 and LC2 proteins efficiently promote tubulin polymerization
in vitro
The ability of LC1 and LC2 proteins to induce the assembly of
microtubules was investigated in reconstitution experiments. Previously, the influence of MAP1A and MAP1B on polymerization of
microtubules in vitro has been monitored using the entire
MAP1A and MAP1B heavy-chain/light-chain complex purified from brain (Pedrotti et al., 1993; Pedrotti and Islam, 1995). Here, we tested the
influence of the individual light chains LC1 and LC2 on tubulin polymerization by turbidimetric time course experiments. Rapid polymerization of tubulin was induced by the full-length light-chain proteins and deletion mutants comprising the
NH2-terminal domains, each added at molar ratios
of 1:40 to 1:1.5 to tubulin (Fig. 5). Microtubule formation appeared to proceed with little or no delay after
addition of LC1 and LC2. Whereas polymerization induced by the
full-length light chains commenced immediately, a lag phase preceded
polymerization induced by the NH2-terminal
fragments, resulting in a sigmoidal shape of the increase in absorbance
over time. This lag phase was shortened but not abolished by an
increase in the concentration of the light-chain fragment. Moreover,
increasing the concentration of the NH2-terminal
light-chain fragment led to an increase in the formation of
microtubules, as evidenced by the higher plateau value of absorbance.
The COOH-terminal domains of LC1 (Fig. 5) and LC2 (data not shown)
added at a molar ratio of 1:1.5 to tubulin did not induce microtubule
polymerization above background levels. In fact, the COOH-terminal
domains served as negative controls and showed that the observed
effects were specific and not attributable to the presence of the His
tag. In the absence of light-chain proteins, little or no
polymerization of tubulin was detected. Likewise, incubation of
light-chain proteins in the absence of tubulin did not lead to an
increase of absorbance (data not shown). Taken together, these results
demonstrate that the light chains of MAP1A and MAP1B have a profound
influence on microtubule polymerization in vitro.
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Figure 5.
LC1 and LC2 proteins promote the polymerization of
tubulin. LC1 and LC2 proteins were mixed with 1.5 mg/ml tubulin at
molar ratios ranging from 1:1.5 to 1:40 as indicated. Polymerization of
tubulin was started by placing the mixtures into prewarmed cuvettes and
monitored by the change in absorbance at 350 nm. Rapid polymerization
of tubulin was observed using the full-length LC1 and LC2 proteins.
Microtubule formation in the presence of NH2-terminal
domains of LC1 (N-term LC1) and LC2 (N-term
LC2) appeared to proceed with a delay; COOH-terminal domains of
LC1 (C-term LC1) and tubulin alone
(tubulin) were used as controls. The dotted
line in each panel defines absorbance level zero.
Abs, Absorbance.
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The COOH-terminal domain of LC2 interacts with actin stress fibers
in vivo
Apart from its microtubule-binding activities, several studies
have revealed that MAP1A interacts with microfilaments in
vitro (Fujii et al., 1993; Pedrotti et al., 1994). However, these
experiments had been conducted using the entire MAP1A
heavy-chain/light-chain complex, and the localization of the
actin-binding domain as well as experimental evidence obtained in
vivo were not reported. Transfection of PtK2 cells with the
full-length LC2 protein displayed colocalization with microtubules
(Fig. 1A-C), and association with microfilaments was
never observed. In contrast, transfection of the LC2 COOH terminus
revealed that it is capable of binding to microfilaments. Cells
expressing the LC2 COOH terminus displayed fibrillar staining and
colocalization of the protein with actin stress fibers (Fig. 6). The latter was not affected by
omission of the myc tag (data not shown). Staining with anti-myc
antibodies also revealed a nuclear localization of this LC2 fragment,
probably for reasons similar to those discussed above for full-length
LC2.
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Figure 6.
The COOH-terminal domain of LC2 interacts with
actin stress fibers. PtK2 cells were transfected with the COOH-terminal
domain of LC2. Double immunofluorescence microscopy using antibodies
against the myc-tag (A) and actin
(B) revealed colocalization of the myc-tagged LC2
COOH terminus with actin stress fibers (A and
B, arrows). Scale bar, 10 µm.
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The COOH-terminal domains of LC1 and LC2 bind to actin
in vitro
We further tested whether binding of the corresponding homologous
COOH-terminal domains of LC1 and LC2 to actin stress fibers is
attributable to a direct interaction with actin or whether it is
mediated by an actin filament-associated protein. Purified recombinant
full-length LC1 and LC2 proteins as well as deletion mutants comprising
the respective NH2-terminal and COOH-terminal domains were subjected to a solid-phase binding assay or to
cosedimentation with actin filaments to investigate actin-binding
properties in vitro. To perform solid-phase binding studies,
microtiter plates were coated with 100 nM actin
or BSA as a control. Coated wells were overlaid with increasing amounts
of Eu3+-labeled LC1 and LC2, and bound
protein was measured. For all proteins tested, binding to BSA was
considered to be nonspecific and therefore was subtracted from the
amount of protein bound to actin. Full-length LC1 and LC2 as well as
their respective COOH-terminal domain showed specific binding to actin,
whereas fragments containing only the NH2
terminus of the respective light chain showed no binding (Fig.
7). At high concentrations, the full-length LC1 protein showed reduced binding compared with the COOH-terminal domain (Fig. 7A). Concerning LC2, both
proteins, the full-length LC2 and the COOH-terminal domain, were
approximately equally efficient in binding to actin (Fig.
7B).
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Figure 7.
The COOH-terminal domains of LC1 and LC2 interact
with actin in vitro. Microtiter plates coated with 100 nM actin were overlaid with various concentrations of
Eu3+-labeled LC1 (A) and LC2
(B) proteins. Full-length LC1 and LC2 as well as
their respective COOH-terminal domains (C-term LC1 and
C-term LC2) showed specific binding to actin. No binding
was observed using the NH2-terminal domains of LC1
(N-term LC1) and LC2 (N-term LC2). The
amounts of protein bound represent the results of three measurements of
at least two independent experiments.
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To confirm these results by a second biochemical approach, we performed
actin cosedimentation assays using the full-length LC1 and LC2 proteins
and deletion mutants. Significant binding to actin was shown using the
full-length LC1 and LC2 proteins (Fig.
8). In the absence of actin, both
proteins were found predominantly in the supernatant, whereas
incubation with actin led to complete sedimentation of the light-chain
proteins. Sedimentation of F-actin remained unaffected in the presence
of LC1 and LC2, indicating that neither protein prevented actin
polymerization like F-actin capping or severing proteins (Matsudaira,
1992). Sedimentation of polymerized actin alone was almost complete,
resulting in at least 90% of the protein being found in the pellet
fraction (data not shown). Under the conditions of this assay,
COOH-terminal domains of LC1 and LC2 pelleted in the absence of actin
and therefore were not included in this assay. As expected, the
NH2-terminal domains of LC1 and LC2 in the
absence as well as the presence of actin were found almost exclusively
in the supernatant. Thus, in cosedimentation as well as
Eu3+-overlay assays, we can exclude
nonspecific binding attributable to His tags. These results confirmed
and extended our results obtained in vivo (Fig. 6) and
demonstrated by two independent in vitro assays that the
light chains of MAP1A and MAP1B can directly interact with actin
through an actin-binding site located in the homologous COOH-terminal
domains.
View larger version (23K):
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|
Figure 8.
LC1 and LC2 cosediment with actin
(Ac) via their COOH-terminal domain. LC1 and LC2
proteins (arrows) were sedimented in the presence (+) or
absence () of polymerized actin. Equal amounts of supernatant
(S) and pellet (P)
fractions were analyzed by SDS-PAGE and Coomassie blue staining.
Full-length LC1 and LC2 were found to cosediment with actin, whereas
only trace amounts of the NH2-terminal domains of LC1
(N-term LC1) and LC2 (N-term LC2) were
found in the pellet fraction.
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|
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DISCUSSION |
It is generally accepted that the demands on the neuronal
cytoskeleton change after transition from the developmental state to
the mature state. The switch from MAP1B to MAP1A during postnatal development might be part of the mechanism to adapt the cytoskeleton to
these changing requirements. If this were the case, one would expect
distinct effects of MAP1A and MAP1B on cellular microtubules and
microfilaments. However, previous studies using the entire protein
complex, consisting of heavy and light chains, revealed only moderate
and similar effects on microtubule shape and stability (Noble et al.,
1989; Takemura et al., 1992; Vaillant et al., 1998). To elucidate the
functional properties of MAP1A and to successfully unravel the
existence of MAP1A-specific features that might be instrumental in the
switch from the developmental to the mature cytoskeleton, we chose to
investigate the functional properties of the light chain of MAP1A, LC2,
and to compare them with properties of LC1. Our findings demonstrate
that (1) MAP1A and MAP1B light chains indeed have specific effects on
microtubules, (2) both proteins are potential linkers of microtubules
and microfilaments, and (3) the properties of MAP1A and MAP1B are at
least in part attributable to distinct activities of their light chains.
Ectopic expression of LC2 in PtK2 cells revealed binding of the protein
to microtubules. Previously, two separate regions located on the MAP1A
heavy chain had been implicated in microtubule binding. Cravchik et al.
(1994) reported results of colocalization studies indicating that the
microtubule-binding domain of MAP1A was located on the heavy chain
between amino acids 1300 and 1600. Conversely, Vaillant et al. (1998)
concluded from their transfection experiments using a variety of MAP1A
deletion constructs that amino acids 281-336 and/or amino acids
540-630 of MAP1A are implicated in microtubule binding. These authors
also demonstrated that full-length MAP1A consisting of heavy and light
chains can bind to microtubules, but the existence of a
microtubule-binding domain in LC2 had not been reported. We demonstrate
here that LC2 by itself not only binds to cellular microtubules but
also stabilizes them against the destructive effects of colchicine.
Binding of LC2 was confirmed in vitro, and we have shown
that LC2 has profound effects on microtubule polymerization. The
NH2-terminal domain of LC2 was necessary and sufficient for microtubule binding and induction of polymerization. Nonetheless, microtubule formation was more rapid in the presence of
the full-length LC2 protein comprising NH2-and
COOH-terminal domains than the NH2-terminal
domain alone. Similar results were obtained for LC1. Because the
COOH-terminal domains of either light chain did not themselves interact
with microtubules, their enhancement of microtubule formation when
present as part of the full-length light chain must be caused by a
different mechanism. The COOH-terminal domains of both light chains
share a high degree of sequence identity. We have shown previously that
LC1 dimerizes or even oligomerizes via this COOH-terminal domain
(Tögel et al., 1998b). It is tempting to speculate that
dimerization/oligomerization of the light chains might have an
influence on microtubule polymerization. Association of light chains
via their COOH-terminal domains could facilitate microtubule
polymerization by simply recruiting more tubulin subunits, which would
accelerate the nucleation event preceding microtubule elongation. This
would in turn account for the significantly shortened lag phase we
observed using the full-length light chains. In agreement with this
hypothesis, such influence on the nucleation of microtubule assembly
has been described previously using the entire MAP1B protein
(Vandecandelaere et al., 1996). Rapid polymerization of microtubules
has also been observed using the entire MAP1A and MAP1B protein complex
purified from brain (Pedrotti et al., 1993; Pedrotti and Islam, 1995).
At similar stoichiometric amounts of protein, the polymerization
monitored in these studies resembles the results we obtained by solely
using the light chains, suggesting that microtubule polymerization is enhanced by the light chains, leaving the heavy chains rather dispensable.
While the LC2 NH2 terminus harbors a domain that
is necessary and sufficient for microtubule binding and polymerization,
its COOH-terminal half contains an actin-binding domain. Ectopic
expression of the LC2 COOH terminus revealed interaction with actin
stress fibers. Moreover, the ability of LC2 to interact with actin via its COOH-terminal domain in solid-phase binding studies and
cosedimentation assays suggests a direct association with actin.
Similar results were obtained for LC1 and its COOH terminus. Previous
observations of the interaction of the entire MAP1A complex with
microfilaments (Fujii et al., 1993; Pedrotti et al., 1994) can thus be
explained by the actin-binding properties of the light chain
demonstrated here. An interesting observation in our studies was that
full-length recombinant LC1 and LC2, synthesized in and purified from
E. coli, were able to bind to microtubules and actin
in vitro, whereas LC1 and LC2 ectopically expressed in PtK2
cells associated exclusively with microtubules, although their
COOH-terminal domains can bind to actin in vitro and
in vivo. It is conceivable that actin binding of full-length
light chains is inhibited or regulated by post-translational modifications such as phosphorylation, which could take place in PtK2
cells but not in E. coli. Indeed, it has been demonstrated that binding of the MAP1B heavy-chain/light-chain complex to actin requires previous treatment with alkaline phosphatase (Pedrotti and
Islam, 1996), and that phosphorylation of MAP1B enhances binding to
microtubules (Brugg and Matus, 1988; Diaz-Nido et al., 1988; Ulloa et
al., 1993). A similar mechanism might operate for MAP1A and LC2.
Through our comparison of LC1 and LC2, we discovered that the two light
chains do indeed have similar but distinct activities, consistent with
the notion that the switch from LC1 to LC2 during postnatal brain
development could reflect the changing demands on the neuronal
cytoskeleton. Both LC1 and LC2 each have an
NH2-terminal microtubule-binding domain linked to
a COOH-terminal actin-binding domain. Both light chains bind to
microtubules and actin in vivo and in vitro and
enhance microtubule polymerization in vitro. The major
differences are in the effects on cellular microtubules. Whereas LC1
induces the formation of microtubule bundles and loops reminiscent of
microtubules in advancing growth cones (Tsui et al., 1984; Sabry et
al., 1991; Tanaka and Kirschner, 1991; Challacombe et al., 1996; Dent
et al., 1999), LC2 does not appear to change microtubule arrangements.
In addition, LC2 stabilizes microtubules only against colchicine, but
not nocodazole, whereas LC1 is effective against both drugs. Colchicine
binds to tubulin through a bipartite binding site consisting of A and C
subsites, of which presumably only the A site is used by nocodazole
(Wilson and Jordan, 1994). The differential effects of LC1 and LC2
might be caused by interference with the A and C site, respectively.
Thus, both could prevent colchicine binding by blocking either the A or
the C site, whereas only LC1 would also block nocodazole binding. These
differences in light-chain action on microtubules can be attributed to
the differences in primary structure of the respective
microtubule-binding domains located in the
NH2-terminal half of the two light chains, because these are the only parts of the light chains that are not
homologous in sequence. Thus, the effect of the switch from LC1 to LC2
during postnatal development is in essence the replacement of the LC1
microtubule-binding domain by the corresponding LC2 domain with a
different effect on microtubules.
The present study demonstrates the importance of the light chains for
MAP1B and MAP1A function. In the brain, the light chains can be found
in two settings. One fraction of LC1 and LC2 is found associated with
the corresponding heavy chains (Schoenfeld et al., 1989). We have
previously obtained evidence that in the MAP1B heavy-chain/light-chain
complex, the heavy chain acts as a regulatory subunit (Tögel et
al., 1998b). Our present findings of microtubule- and actin-binding
activities of LC2 raise the possibility that in the MAP1A complex as
well, the light chain is the active subunit and the heavy chain acts as
a regulator. Of particular interest in this context is the fact that
heterologous complexes can be detected in which LC1 is bound to a MAP1A
heavy chain (Schoenfeld et al., 1989). Thus, it appears that during
postnatal development, in addition to the switch from MAP1B to MAP1A,
LC1 can participate in a mixed complex consisting of the active subunit
LC1 with its distinct "developmental" effects on cellular
microtubules and the putative "adult" regulator, the heavy chain of
MAP1A. Whatever the true function of the MAP1A heavy chain may be, the
documented light-chain replacement on a given heavy chain at a certain
transition period during development is consistent with the
above-mentioned model of light- and heavy-chain duties in the MAP1
complex and emphasizes the importance of the light chains.
A second and larger fraction of the light chains is found not to be
associated with heavy chains. This has been demonstrated for LC1 (Mei
et al., 2000b) and could be true for LC2 as well. The results presented
here identify which activities such uncomplexed light chains have. We
propose that the microtubule- and actin-binding activities reported
here will be the key to the functions of the light chains.
| |
FOOTNOTES |
Received Aug. 13, 2001; revised Nov. 20, 2001; accepted Dec. 3, 2001.
This research was supported by a grant from the Austrian Science Fund
(Project No. F607). We thank Albert Weixelbaumer for his contribution
in the biochemical analyses.
Correspondence should be addressed to F. Propst, Institute of
Biochemistry and Molecular Cell Biology, Dr. Bohr-Gasse 9, A-1030 Vienna, Austria. E-mail: friedrich.propst{at}univie.ac.at.
| |
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R. Lu, H. Wang, Z. Liang, L. Ku, W. T. O'Donnell, W. Li, S. T. Warren, and Y. Feng
The fragile X protein controls microtubule-associated protein 1B translation and microtubule stability in brain neuron development
PNAS,
October 19, 2004;
101(42):
15201 - 15206.
[Abstract]
[Full Text]
[PDF]
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C. Steindler, Z. Li, M. Algarte, A. Alcover, V. Libri, J. Ragimbeau, and S. Pellegrini
Jamip1 (Marlin-1) Defines a Family of Proteins Interacting with Janus Kinases and Microtubules
J. Biol. Chem.,
October 8, 2004;
279(41):
43168 - 43177.
[Abstract]
[Full Text]
[PDF]
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C. Bouquet, S. Soares, Y. von Boxberg, M. Ravaille-Veron, F. Propst, and F. Nothias
Microtubule-Associated Protein 1B Controls Directionality of Growth Cone Migration and Axonal Branching in Regeneration of Adult Dorsal Root Ganglia Neurons
J. Neurosci.,
August 11, 2004;
24(32):
7204 - 7213.
[Abstract]
[Full Text]
[PDF]
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J. H. Ives, S. Fung, P. Tiwari, H. L. Payne, and C. L. Thompson
Microtubule-associated Protein Light Chain 2 Is a Stargazin-AMPA Receptor Complex-interacting Protein in Vivo
J. Biol. Chem.,
July 23, 2004;
279(30):
31002 - 31009.
[Abstract]
[Full Text]
[PDF]
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A. Dallol, A. Agathanggelou, S. L. Fenton, J. Ahmed-Choudhury, L. Hesson, M. D. Vos, G. J. Clark, J. Downward, E. R. Maher, and F. Latif
RASSF1A Interacts with Microtubule-Associated Proteins and Modulates Microtubule Dynamics
Cancer Res.,
June 15, 2004;
64(12):
4112 - 4116.
[Abstract]
[Full Text]
[PDF]
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E. Y.M. Wong, J. Y.M. Tse, K.-M. Yao, V. C.H. Lui, P.-C. Tam, and W. S.B. Yeung
Identification and Characterization of Human VCY2-Interacting Protein: VCY2IP-1, a Microtubule-Associated Protein-Like Protein
Biol Reprod,
March 1, 2004;
70(3):
775 - 784.
[Abstract]
[Full Text]
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J. A. Morris, G. Kandpal, L. Ma, and C. P. Austin
DISC1 (Disrupted-In-Schizophrenia 1) is a centrosome-associated protein that interacts with MAP1A, MIPT3, ATF4/5 and NUDEL: regulation and loss of interaction with mutation
Hum. Mol. Genet.,
July 1, 2003;
12(13):
1591 - 1608.
[Abstract]
[Full Text]
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TOP
ABSTRACT
INTRODUCTION
Gene
