From the Fachbereich B Biologie/Chemie, Universität Osnabrück, 49069 Osnabrück, Germany
Received for publication, December 16, 2002 , and in revised form, May 7, 2003.
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ABSTRACT |
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INTRODUCTION |
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In addition to the teichoic acids, proteins have been found to be associated with the cell wall. In Streptomyces species only few surface-exposed proteins have been described up to now. These include a 23-kDa protein from Streptomyces lividans (12), a cell-bound esterase synthesized by the cyclophilin A-producing strain Streptomyces chrysomallus X2 (13), the mycelia-associated cellulase (14), catalase-peroxidase from Streptomyces reticuli (15), and the surface-active proteins (i.e. SapB from Streptomyces coelicolor and its homologue from Streptomyces tendae), which are involved in erecting aerial hyphae (16).
Recently we identified a 35-kDa protein from S. reticuli, which is
very likely covalently anchored to the cell wall
(17). Its N-terminal part
protrudes from the surface of the hyphae, as demonstrated by immunolabelled
ultrathin sections and investigations by electron microscopy
(18). The protein has no
enzymatic activity, but it interacts strongly with crystalline forms of
cellulose (Avicel).
AbpS1 (for
Avicel-binding protein from S. reticuli) recognized other biopolymers
merely weakly (chitin and Valonia cellulose) or not at all (xylan, starch, and
agar). By comparing the deduced AbpS sequence, no homology was found to any
discovered cellulose-binding domain, which were often present within
cellulases (19). AbpS
possesses an up to now unique cellulose-binding module. By analysis of the
secondary structure of the deduced AbpS sequence, a large centrally located
-helical structure showing a weak homology to the tropomyosin protein
family and the streptococcal M-proteins
(20) was identified. As AbpS
has also been found to be associated to protoplasts, it is predicted that a
C-terminally located stretch of 18 hydrophobic amino acids anchors the protein
to the cytoplasmic membrane.
Beside streptomycetes, surface proteins possessing diverse functions were also discovered in several other bacteria (21). In pathogenic bacteria streptococci and staphylococci (2224) these proteins are often called microbial surface components recognizing adhesive matrix molecules (MSCRAMM), such as fibronectin, collagen, or immunoglobulins (25, 26). Because of their involvement in adherence of the bacteria and concealment of the bacterial surface from the host's defense system, MSCRAMMs were studied intensively. Surface-associated heparin-binding proteins are frequent among pathogenic mycobacteria (27, 28). Various types of cell-wall-anchored proteinases are encountered in different genera of Gram-positive bacteria (2931).
In this study we elucidate the detailed characteristics of the up to date unique Avicel-binding protein from S. reticuli. The investigations as to the membrane anchoring and complex formation support the conclusions that AbpS connects the Streptomyces cytoplasm with the extracellular environment and functions as a cellulose-receptor.
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MATERIALS AND METHODS |
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pUS1, a pUC18 derivative containing a 3.2-kb genomic SalI DNA
fragment from S. reticuli, on which the complete abpS gene
is located, was described earlier
(17,
18). The DNA sequence of
abpS is available in the EMBL data bank under the accession number
Z97071
[GenBank]
. The E. coli vector pET21a (Novagen, Madison, WI) was used as
cloning vector for truncated forms of the abpS gene. The derivatives
of pET21a containing the complete abpS gene or truncated genes with
5' end deletions (resulting in N-terminally shortened AbpS-proteins with
molecular weights of 32.3 kDa, 29.1 kDa, and 23.5 kDa) were described earlier
(17). E. coli
DH5 or the chloramphenicol resistant E. coli BL21 (pLysS)
(Novagen) was transformed with plasmid DNA using the CaCl2 method
(36).
Polymerase Chain Reactions and Construction of PlasmidsThe
PCR reaction mixture contained different primer combinations: 10 ng of pUS1,
0.2 nM of each of the dNTPs (dATP, dCTP, dGTP, and dTTP), 10
mM KCl, 10 mM
(NH4)2SO4,20mM Tris-HCl (pH 8.8),
2 mM MgSO4, and 0.1% Triton X-100, in a final volume of
30 µl. To reduce misreading, the Vent® DNA polymerase, which has a
3'5' proof reading exonuclease activity, was used instead of
the Taq polymerase.
The primer P1 CAGGAACCATATGAGCGACAC was utilized to replace the ATG start codon of abpS with an NdeI-site. The primers P2 (GAGCATCTCGTCGACGTTCGTCAG) and P3 (TGCGCGGTGTCGACCGTC TGGCGG) were employed to introduce SalI sites at different locations at the 3' end of the abpS gene. The following cycling conditions were found to be optimal for the primer combinations P1 and P2 or P1 and P3: 95 °C, 90 s; 55 °C, 60 s; and 72 °C, 60 s. After thirty cycles, the PCR products were purified (Qia quick spin PCR purification kit, Qiagen, Hilden, Germany) according to the supplier's instructions. The NdeI- and SalI-digested PCR products were ligated in-frame into the polycloning site of pET21a (linearized with NdeI and XhoI), and transformed in E. coli BL21 (pLysS).
Internal deletions of the abpS gene were performed by "inverse" PCR. The pET21a derivative comprising the complete abpS gene served as template. The primers were designed in such a way that the generated PCR products consisted of the vector-sequence and flanking abpS fragments. The space between the primers represented the desired deletion. For self-ligation of the PCR products, in-frame SstI sites were incorporated in the primer sequences. For amplification of DNA with the primers PintA (GGCCTTGAGCTCGCCCGCGTA) and PintB (GGCGGACGAGCTCTTCGAG GAGAG) or PintC (GCCGAGGAGCTCCGCCTGGAGGC), the following cycle conditions were used: 95 °C, 90 s; 50 °C, 90 s; and 72 °C, 450 s. After ten cycles the products were purified (see above) and digested with SstI, self-ligated, and transformed in E. coli BL21 (pLysS).
For cloning of abpS genes prolonged with the codons for 6
histidines in S. lividans as host PCR was performed under the above
described conditions with primer PXho (CTGCACCTCGAGACGCAG) and
PHis6 (AAATGATCAGTGGTGGTGGTGGTGGTGGGAGCCCCGGGACTGCTGCGCCGGGAC) or
PCHis6 (AAATGATCAGTGGTGGTGGTGGTGGTGGGAGCCCCGG
ACGTTCGTCAGCTGGGC) and as template pUS1 (containing the abpS gene
from S. reticuli) was used (see above). After purification and
digestion with XhoI and BclI, the resulting DNA fragment was
used to substitute the abpS 3' end in pUS1, cleaved before with
XhoI and BamHI. Consequently, the corresponding plasmids
contained genes encoding a full-length AbpS or an AbpS protein without the
hydrophobic segment both prolonged C-terminal with RGSH6.
Subsequently, the genes were transferred into the bifuncional
Streptomyces-E. coli vector pWHM3 and named pWA1 and
pWA
C, respectively. Both plasmids were propagated in E. coli
DH5
and after isolation transformed into S. lividans 66
protoplasts. The presence of the pWHM3-based constructs was guaranteed by
selection with thiostrepton (25 µg/ml).
General DNA TechniquesApplication of restriction enzymes and the T4-ligase were carried out using the standard procedures (36). DNA sequencing was performed with the help of the T7 sequencing kit and Cy5-labeled standard primers (Amersham Biosciences).
Purification of Truncated His-tagged AbpS Proteins from E. coli or S.
lividansThe E. coli BL21 (pLysS) transformants containing
the designed constructs (see above) were grown at 37 °C in SOC medium (20
g of Bacto-Trypton, 5 g of yeast extract, 0.5 g of NaCl, 0.18 g of KCl, and 20
ml of 1 M glucose/liter, which had been supplemented with
chloramphenicol (34 µg/ml) and ampicillin (100 µg/ml)).
Isopropyl--D-thiogalactopyranose (final concentration 1
mM) was added when A600 had reached 0.6. After
further 3 h of cultivation, the E. coli cells were harvested, washed,
resuspended in sonification buffer (0.1 M
NaH2PO4, 0.01 M Tris-HCl (pH 8.0), and 8
M urea), and disrupted using a Branson sonifier B12 for 3 min in 20
s intervals. Having removed the cell debris, Ni2+-NTA
(Qiagen) was added to bind the His6 fusion protein. Unspecifically
bound proteins were removed by consecutive washings with buffer (0.1
M NaH2PO4, 0.01 M Tris-HCl (pH
6.3), and 8 M urea) containing 25 mM imidazol. The
immobilized proteins were renatured by washing with buffers containing
decreasing concentrations of urea (8-0 M) in a period of 23
h. The fusion protein was subsequently released by the addition of 0.5
M imidazol.
S. lividans transformants were grown in pH-stable medium (MM3) supplemented with 1% glucose for 48 h. After harvesting the mycelia by centrifugation, they were disrupted and the His6-tagged proteins were isolated as described above with or without urea-containing buffers.
Isolation of Murein-associated AbpS from S. reticuliAnti-AbpS antibodies had been gained previously (18). The IgGs were purified using a 1 ml HiTrap protein G column (Amersham Biosciences) according to the supplier's manual. Subsequently, the purified IgGs were coupled to CNBr-activated Sepharose 4 Fast Flow (Amersham Biosciences) as described in the corresponding manual.
The murein layer was isolated from S. reticuli hyphae grown in 1 liter of minimal media as described earlier (18). Murein was treated with buffer (25 mM Tris-HCl (pH 8) and 20 mM EDTA) comprising 5 mg of lysozyme (Roche Applied Science) per ml and incubated at 37 °C for 3 h. After centrifugation at 14,000 x g for 30 min, the AbpS-containing supernatant was mixed with immobilized (see above) anti-AbpS antibodies. After incubation for 35 h under gentle stirring, the Sepharose was washed 3 times with 50 ml PBS (80 g of NaCl, 2 g of KCl, 2 g of KH2PO4, 11.5 g of Na2HPO4 per liter (pH 7)). The proteins were released from the immobilized IgGs by applying 50 mM phosphate-citrate buffer (pH 2.7). For renaturation the pH was adjusted to 7, and purity and quantity were determined by SDS-PAGE followed by staining with Coomassie Brilliant Blue.
Separation of Cell CompartmentsS. lividans mycelia were harvested by centrifugation, washed 3 times with 10.3% sucrose, and murein was hydrolyzed by incubation with 3 mg of lysozyme ml1 mg1 wet weight of mycelia (35). After filtration the protoplasts were resuspended in water supplemented with 5 mM EDTA to destroy them osmotically. The membranes and associated proteins were separated from soluble proteins by ultracentrifugation (250,000 x g for 45 min).
SDS Gel Electrophoresis and Western BlottingSDS-PAGE was performed with 10% polyacrylamide gels in the presence of 0.1% SDS (37). If desired, proteins were transferred onto nylon membranes, which were incubated in PBS containing the primary antiserum (1:100,000 dilution), the anti-His5 antibodies, or the anti-RGSHis6 antibodies (both 1:10,000 dilution) (Qiagen). After three washes, the blot was incubated with alkaline phosphatase conjugated with the AffinyPure F(ab')2 fragments of goat anti-rabbit IgG or mouse anti-rabbit IgG (Dianova, Hamburg, Germany). Color development was performed as described by West et al. (38).
Urea-gradient PAGEThe lower polyacrylamide-gel (in a concentration of 6% and without SDS) was poured sidelong into the glass plate in 4 successive stages, containing 6, 3, 1.25 M, or no urea. After polymerization, the standard upper gel was overlaid so that one wide lane was formed. The proteins were loaded with a native loading buffer (36), and the electrophoresis was done at 15 mA to avoid heating above 25 °C. Finally the proteins were stained with Coomassie Brilliant Blue.
Cellulose-binding AssayProteins were incubated with Avicel (15 mg/ml) in 50 mM potassium phosphate buffer (pH 7) for 30 min. Avicel recovered by centrifugation was washed 3 times with 50 mM potassium phosphate buffer containing 1 M NaCl. Avicel-bound proteins were released by heating in SDS sample buffer for 5 min at 100 °C and subjected to SDS-PAGE. Staining of proteins was done with Coomassie Brilliant Blue.
Detection of Protein-Protein InteractionsTotal proteins or purified AbpS isolated from S. reticuli were denatured by SDS, separated on an SDS-PAGE, and subsequently transferred onto a polyvinylidene difluoride membrane (because of the cellulose-binding capability of AbpS, nitrocellulose membrane was avoided). The membrane was treated for 1 h with PBS (see above) containing 1% bovine serum albumin. The His6-tagged fusion proteins (which were to be tested for the interaction of the immobilized AbpS) were incubated with 8 M urea in small volumes for 1 h. Rapid renaturation was performed by diluting this solution 1:100 in PBS, which contained a piece of the nylon membrane with immobilized AbpS from S. reticuli. After incubation at 25 °C the membrane was washed 3 times with 50 ml of PBS, and the fusion proteins that were trapped by the immobilized AbpS were detected immunologically by applying antibodies specific to a stretch of 5 histidines (for further details see "SDS Gel Electrophoresis and Western Blotting").
Cross-linking of AbpS ComplexesBecause of the absence of
cysteine residues in AbpS the reactive bifunctional cross-linker
dithiobis(succinimidyl propionate) (DSP), which is cleavable by reducing
agents, was chosen to interconnect subunits of protein complexes. For this
purpose the concentration of each of the His6 fusion proteins
(35.7, 27.1C, 23.5
N, or 31.4
I) was adjusted to 10
mM.3 µl (corresponds to 30 nmol) were diluted in 50 µl of
buffer (25 mM Tris-HCl (pH 7)). The addition of 50 mM
NaCl was found to inhibit ionic protein interaction, which increases the
cross-linking specificity. To avoid intermolecular cross-linkages the molar
ratio between DSP and denaturated proteins (0.5% SDS) was adjusted so that no
multimers were found. The molar ratio (DSP/protein) of 3:1 was found to be
optimal. Therefore the cross-linking reaction was started by adding 3 µl of
10 mM solution of DSP (dissolved in Me2SO). After 10 min
at 30 °C, the proteins were denaturated by SDS at 95 °C and analyzed
by PAGE.
0.6 µg of AbpS isolated from S. reticuli were incubated with 10 mg Avicel in a final volume of 100 µl for 1 h. After 3 washes, 50 µl of 25 mM Tris-HCl (pH 7) was added to the Avicel, and subsequently the bound proteins were cross-linked as described above. As control, the same amount of unbound AbpS was used.
Circular Dichroic Spectroscopiccal AnalysisTo assess the
structure of AbpS or its His6 fusion variants (35.7, 27.1C,
23.5, and 31.4
I) the concentrations of each of the proteins were
adjusted to 0.2 mg ml1. The spectra
(190260 nm) were recorded at 25 °C using a Jasco 600
spectrophotometer in a 0.1-mm path length cell. Data were recorded 5 times,
and an average value was determined. The observed ellipticity [
]
(degrees) is converted in mean residues ellipticity by:
[
]mrw,l
= MRW
/10 dc, where MRW is obtained by
dividing the molecular mass by n1(n = number of amino
acids in the protein), d is the path length (cm) and c is the protein
concentration (g/ml). The calculated ellipticity was further converted to the
difference of the molar absorbance ([
]mrw = 3298
). Subsequently the data were analyzed by CD Spectroscopy
Deconvolution program CDNN 2.1
(bioinformatik.biochemtech.uni-halle.de/).
Reconstitution of AbpSSoybean Azolectins
(L--phosphatidylcholine solution obtained from Sigma) were
washed twice with acetone in the presence of butylated hydroxytoluene and once
with ether/vitamine E, dissolved in chloroform, and dried under a nitrogen
stream. The preparation of liposomes was done according to Jung et
al. (51). After
solubilization of the liposomes by addition of 3% octyl
-D-glucopyranoside, the His6-tagged fusion
proteins isolated from S. lividans were added and sonicated for 1 s
and repeated 9 times. After removing the detergent by Bio-Beads SM-2 (Bio-Rad)
the proteoliposomes were washed and concentrated by centrifugation at 250,000
x g.
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RESULTS AND DISCUSSION |
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Each E. coli BL21 (pLysS) transformant harboring one of the
different constructs was grown to the logarithmic phase, induced with varying
concentrations of isopropyl--D-thiogalactopyranose at
different temperatures. Independently of the used conditions, each type of the
fusion protein was found dominantly within insoluble inclusion bodies (more
than 90%). Therefore the proteins were bound to Ni2+-NTA
in the presence of 8 M urea. Removing urea stepwise (see
"Materials and Methods"), the proteins were subsequently released
by imidazol, and each type was found to have the predicated molecular weight
(see Fig. 1D).
Identification of -Helical StructuresTo
analyze whether the renatured truncated AbpS forms (isolated from E.
coli) kept structural characteristics of the S. reticuli
wild-type AbpS a circular dichroism spectrum of each of the proteins should be
recorded. As requirement a purification method of AbpS from S.
reticuli has to be established first. As reported previously
(18) AbpS is covalently linked
to the peptidoglucan layer of S. reticuli. Therefore, its murein was
isolated as previously described
(18). Subsequently, AbpS was
released by the action of lysozyme and immobilized by Sepharose-coupled
anti-AbpS antibodies. After elution, AbpS was obtained in a good degree of
purity (Fig. 1C);
however, only
5 µg AbpS can be gained from 10 g mycelia (wet weight),
grown in l liter culture.
Additionally the 35.7 His-tagged full-length AbpS fusion protein, the
23.5N, 25.
I, and 31.
C forms were purified from E.
coli as described (see "Material and Methods"). The
concentration of each protein was adjusted to 0.2 mg/ml, and its CD spectrum
was determined (Fig.
2A). Between 205 and 260 nm the spectra of all proteins
were close to identical. Below 205 nm AbpS, isolated from its natural host
(S. reticuli), showed a slightly higher difference in the absorption
(
) value. There-with, the deletion of different regions of the
proteins did not significantly alter the overall structure of the protein. The
competency for assembly of the secondary structure seems to reside in the
composition of the protein sequence itself. The structural integrity is also
reflected by the evaluated occurrence of 5 different protein-folding motifs
(helix, anti- and parallel
-sheets,
-turns, or random coils).
Their quantification as deduced from the CD spectra revealed that wild-type
AbpS and its truncated forms consist predominantly of
-helical
structures (ranging between 93 and 98.2%), and other structural motives are
under-represented. This finding is in good agreement with a computer-supported
prediction of the secondary structural elements within AbpS
(Fig. 1B,
scheme), comprising a large, centrally located
-helix flanked
by two shorter helices, whereas the C-terminal helix is built by hydrophobic
amino acids.
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Analyzing the Function of the C-terminal Hydrophobic Stretch of AbpSIn vivo AbpS was found to be associated with the membrane of the protoplasts, generated by removing the murein layer from the hyphae of S. reticuli (18). The C-terminally located hydrophobic helix was thus suspected to anchor AbpS to the membrane. To analyze this in more detail comparative in vivo and in vitro studies of AbpS with and without the hydrophobic segment were designed.
As a first requirement AbpS and the designed C-terminally truncated variant have to be synthesized in a Streptomyces host to guarantee native conditions, for example allowing the proper protein folding or membrane anchoring. Because there were no stable plasmids available for S. reticuli, the genetically best studied S. lividans strain was chosen as host. A disadvantage of S. lividans is the presence of an abpS homologue located within its chromosome (17). AbpS-negative S. reticuli or S. lividans mutants or AbpS-negative Streptomyces wild-type strains were not available.
Vector constructs (based on pWHM3) were designed having either the complete
abpS gene (pWA1) or the 5' end deleted abpS gene
(pWAC) (encoding AbpS without the hydrophobic helix). They were kept
under the transcriptional control of the upstream region of the S.
reticuli abpS gene. In total, proteins of the transformants S.
lividans (pWA1) or S. lividans (pWA
C) each prolonged with
a His6 tag (named AbpSHis or
CAbpSHis, respectively) could
be found in addition to the endogenous AbpS homologue (see S.
lividans (pWHM3) as control) with the help of anti-AbpS antibodies. Due
to the similar molecular weight of the endogenous S. lividans, AbpS,
and the plasmid-encoded His-tagged fusion protein, they could not be separated
by PAGE. Instead, the use of anti-RGSH6 antibodies allowed the
specific detection of the modified proteins
(Fig. 3A). AbpSHis or
AbpSHis could be isolated in high purity with the help of Ni-NTA under
denaturating conditions (Fig.
3D, right panel). After renaturation they were
reconstituted into liposomes. Only AbpSHis was found to be integrated in the
vesicles. Deletion of the hydrophobic part within
CAbpSHis led to
inhibition of its integration into proteoliposomes
(Fig. 3B). To study
the membrane integration within the original host in more detail, S.
reticuli membranes were isolated and the release of AbpS was tested. Only
in the presence of SDS (1%) or Triton X-100 (1%) but not with EDTA, urea (up
to 1 M), nor water, AbpS was unhinged from the membranes (data not
shown). Therefore a membrane-association of AbpS by ionic forces or a bedding
of AbpS on the membrane-surface by hydrophobic interactions could be
excluded.
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Summarizing the data, the C-terminal helix was clearly identified as a transmembrane spanning segment. With simultaneous consideration that the N terminus of AbpS was previously found to protrude from the cell surface (18) and a transmembrane spanning helix is present, the C-terminal end of AbpS has to extend into the cytoplasm of the S. reticuli hyphae. Thus AbpS connects the interior of the mycelia with the extracellular space, where it binds insoluble cellulose. Such organization is reminiscent of the assembly of diverse classes of receptor proteins (39), often involved in signal transduction cascades.
To study the effect of the deletion of the transmembrane segment in
vivo (in S. lividans mycelia), the overproduction of AbpSHis or
CAbpSHis in S. lividans was prevented by the cloning strategy
(using the wild-type regulatory elements). As a result, AbpS and its
derivatives were found in approximately the same concentrations
(Fig. 3A) within the
total proteins isolated from S. lividans containing pWHM3, pWA1, or
pWA
C, respectively.
It was expected that at least the C-terminally deleted protein
CAbpSHis would be released by digestion of the murein. But surprisingly
the distribution of the endogenous S. lividans AbpS and
CAbpSHis in the different cell compartments (extra cellular,
murein-associated, membrane-integrated, or cytoplasmic) was found to be equal
(Fig. 3C). A possible
cause for this in vivo effect, which contrasted the in vitro
data, appeared to be the fact that the endogenous membrane-integrated S.
lividans-AbpS molecules form complexes with
CAbpSHis
(plasmid-encoded) and impede, therefore, the release of the C-terminally
deleted protein. The expected complex formation could be satisfactorily shown
by co-purification of the endogenous S. lividans-AbpS (unable to bind
to Ni-NTA) together with the His6-tagged AbpS derivatives (AbpSHis
and
CAbpSHis) by Ni-NTA-based affinity chromatography
(Fig. 3D).
Identification of AbpS Domains Required for Intermolecular
InteractionTo study the deduced interaction among AbpS molecules
in more detail, total proteins (including the S. reticuli wild-type
AbpS) from S. reticuli were denatured, separated by SDS-PAGE, and
then immobilized on a nylon membrane. Subsequently identical amounts of the
truncated and denatured AbpS forms were added to the membrane-immobilized AbpS
in low salt buffer. The degree of intermolecular interaction was determined by
immunological quantification of the membrane-retaining His6 fusion
proteins (Fig. 4A).
Independently of the presence (35.7 kDa form) or absence (31.0C and
27.1
C) of the C-terminal part, each of the proteins strongly interacted
with the immobilized S. reticuli-AbpS. However, continued deletions
of the N-terminal part (32.3
N, 29.1
N, and 23.5
N) led to
an increased loss of the binding ability. Therefore it can be concluded that
the portion of the protein (including amino acids 60110) situated at
the beginning of the central
-helix plays an essential role in the
complex forming process. The remaining binding level of the 23.5
N
protein was attributed to a second domain identified by analyzing the
characteristics of additional truncated forms. They extend from the middle
(31.4
I) to the right side (25.6
I) of the centrally located
-helix (including amino acids 161212). The 25.6
I form was
found to retain only 15% of the maximal binding ability, which corresponded
closely to that of the 31.4
I form
(Fig. 4A).
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As outlined above on the nylon-membranes (on which the separated cellular
proteins from S. reticuli were immobilized), each of the added
truncated AbpS forms were found to interact only with AbpS, demonstrating a
high specificity of the protein-protein interaction. To verify this in more
detail, the formed complexes were precisely cut out of the membrane, which was
overlaid with the full-length His6-tagged fusion protein.
Immunological identification clearly demonstrated the sole participation of
AbpS and its His6-tagged form (35.7 kDa) in the intermolecular
interaction (Fig. 4B).
Based on these data, two domains responsible for AbpS-AbpS interaction were
identified, matching regions predicted to contain only -helical
structures (Fig.
4A).
Conditions for Intermolecular InteractionMixtures of membrane-immobilized S. reticuli-AbpS and its His6-tagged full-length 35.7 kDa form were incubated under different conditions (2090 °C or at pH 2.7) or treated with different amounts of chaotropic urea, leading to protein-denaturation protein (0.258 M urea) or increasing the ionic strength (1 M NaCl) (Fig. 4C). The protein complex was found to be stable in the presence of up to 1 M urea or 1 M NaCl. Because these conditions are unfavorable for ionic bonding, oligomerization based on the ionic strength could be excluded. At very low pH (2.7), high concentrations of chaotropic salt (commencing at 1 M) or high temperatures (higher than 50 °C) the protein-protein interaction was usually inhibited strongly. The above-described conditions for resolving the ability for complex formation exactly match those that inhibited the binding of AbpS to cellulose (17). These findings suggest that the S. reticuli AbpS interacts with cellulose only in an oligomerized form.
Characterization of OligomerizationUsing standard methods
(sucrose gradients or gel filtration with various globular reference proteins)
it was found that the molecular weight determined for AbpS neither correlated
to that one for the AbpS monomer nor to any expected oligomer. This was
attributed to the fact that the secondary structure of AbpS was found to be
dominantly -helical, governing an elongated shape of the protein (Figs.
1B and
2). Because correspondingly
shaped reference proteins are missing, the mobilities of the various truncated
AbpS forms and the S. reticuli-AbpS were comparatively studied in
native PAA gels in the presence of increasing concentrations of chaotropic
urea (0, 1.25, 3, and 5 M) (Fig.
5). The S. reticuli AbpS, the 35.7-kDa full-length
AbpS-His6 fusion protein, and the 31.4
I protein (with a
short internal deletion) were found to migrate into the gel only in the
presence of 6 M urea. When applying to 3 M urea, no
protein or only traces could be recovered. This effect was also ascertainable
at low (5%) polyacrylamide concentrations. The 31.0
C truncated form
possesses a deletion of 46 C-terminal amino acids, including the
above-characterized hydrophobic membrane-integrated helix. The removal of
consecutive 35 amino acids resulted in the 27.1
C truncated form. Both
protein types were found to migrate into the gel containing up to 3
M urea. In comparison, the relative mobility increased under
denaturing conditions (6 M urea). These data suggest that the
homomultimeric protein complexes were formed by native conditions, whereas the
presence of the hydrophobic segments provoked protein aggregation
(Fig. 5).
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Interestingly, the 23.5N and 25.6
I proteins (both lacking one
domain responsible for the intermolecular protein interaction) were monomeric
under all conditions (06 M urea). This finding is in
agreement with their low protein-protein interaction ability
(Fig. 6). In contrast, the
32.3
N as well as the 29.1
N form, both possessing an increasing
portion of a deleted N terminus, migrated already in the presence of 1.25
M urea as denatured protein complexes; this decreasing stability of
the protein complexes correlates with their intermediate ability to interact
with the wild-type AbpS (see Fig.
4A).
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Again this finding indicates a protein-protein interaction mechanism, which
governs the formation of a stable complex. Together with the structural
information of AbpS a formation of coiled-coil multimers appears likely
(Fig. 7). In coiled-coil
proteins the -helices provides hydrophobic amino acids to one side of
the helix. This hydrophobic side is able to interact with the corresponding
side of the next subunit. Ionic amino acids, proximate to the hydrophobic
ones, direct the formation of dimeric, tetrameric, or higher organized
complexes. Typical coiled-coil proteins are the dimeric tropomyosins
(40), trimeric or tetrameric
leucinezippers (41), or the
pentameric Comp (cartilage oligomeric matrix protein) whose oligomerization
domains marked similarities with proposed models of the pentameric
transmembrane ion channels in phospholamban and the acetylcholine receptor
(42).
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Interestingly some M-proteins of the group A streptococci, which belong to
the MSCRAMMs (see Introduction), have been shown to be coiled-coil dimers,
appearing as fibrils on the bacterial surface in the electron microscope
(23,
43,
44). The arrangement of the
hydrophobic amino acids is also reflected in the primary sequence of a
protein. Numbering the amino acids in a coiled-coil forming -helix by
"abcdefg" amino acids "a" and "d" were
found to be hydrophobic (41).
Analyzing the deduced AbpS protein sequence according to this rule, large
protein sections (including amino acids 20 to 246) were identified with high
potential for coiled-coil interactions
(Fig. 7).
Constitution of the AbpS ComplexesTo determine the
constitution of the AbpS complexes, cross-linking experiments with the
unspecific linking agent DSP were performed. For this purpose (i) the
concentration of the proteins, (ii) the molar ratio of the proteins and the
cross-linker, and (iii) the concentration of NaCl (for inhibition of weak
ionic protein interactions) were carefully optimized to prevent unspecific
linkages (for details see "Material and Methods"). Under optimized
conditions, the 35.7-kDa full-length His6-tagged AbpS,
32.3N, 31.4
I, and 31.0
C proteins were found to form up to
tetramers (Fig. 6A).
In contrast the N-terminally deleted 29.1
N and the 23.5
N protein
shaped only trimers (
87 kDa) or dimers (
47 kDa), respectively. The
cross-linking of the 25.6
I protein (with the extended internal
deletion) resulted in the formation of
75 kDa and
50 kDa complexes,
corresponding to trimers and dimers, respectively. The composition of the
intermediate complexes with the apparent molecular mass between 75 and 50 kDa
was not explored.
The cross-linking experiments were repeated with purified S. reticuli AbpS, which initially was bound to the crystalline Cellulose (Avicel) or kept in solution. The soluble AbpS as well as the Avicel-bound was found to tetramerize, showing that the complex organization is not altered by the interaction of AbpS with cellulose.
The ability to dimerize, or in general to oligomerize, is widespread among other types of carbohydrate-binding proteins. For example lectins specifically recognize diverse sugar structures and mediate a variety of biological processes, such as cell-cell and host-cell interaction (45). For some bulb lectins, which share some similarities in their three-dimensional structure and the ability to specifically bind mannose, the degree of oligomerization was found to modulate carbohydrate-binding specificity. The tetrameric lectins snowdrop (46), daffodil (47), or bluebell (48) bind a surface glycoprotein on human immunodeficiency virus, whereas the dimeric garlic lectin (49) does not.
Cellulose RecognitionComparative analysis of AbpS and each
of the truncated forms revealed that a reduced oligomerization (29.1 N,
23.5
N, and 25.6
I) is accompanied with a decreased capability
for cellulose binding (Fig.
6B). This supports the finding that the conditions
favorable for the intermolecular interactions of AbpS are identical to those
that allow cellulose recognition
(17). Dissociation of the
complexes abolishes the cellulose-binding ability of AbpS. In contrast, the
32.3
N form with a moderate deletion in the N terminus (29 amino acids)
is still able to shape tetramers, whereas the cellulose-binding capability is
reduced 10 times. Therefore amino acids that are involved directly in the
cellulose interaction process have to be located within the first 29 residues
of AbpS. This mode of cellulose-recognition via AbpS is so far unique and
differs considerably from cellulose-binding domains within cellulases
(50). As shown by biochemical
and crystallographical analysis, the interaction of these cellulose-binding
modules is mediated by aromatic amino acids exposed on one side of the
proteins. The distance between the aromatic amino acids correlates with those
between the glucose units within the cellulose
(50). Multimerization of such
cellulose-binding domains is obviously not advantageous; on the contrary, it
leads to an aggregation of the domains containing the active site for
cellulose hydrolysis. In the case of AbpS the amino acid residues that are
directly involved in cellulose recognition seem to be distributed within 4
molecules. Only multimerization arranges these amino acids in a topology
required for optimal interactions.
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FOOTNOTES |
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To whom correspondence should be addressed: Universität Osnabrück,
FB Biologie/Chemie, Barbarastra
e 11, 49069 Osnabrück, Germany.
Tel.: 49-541-969-2287; Fax: 49-541-969-2804; E-mail:
Stefan.Walter{at}uni-osnabrueck.de.
1 The abbreviations used are: AbpS, Avicel-binding protein from S.
reticuli; MSCRAMM, microbial surface components recognizing adhesive
matrix molecules; Ni-NTA, nickel-nitrilotriacetic acid; PBS,
phosphate-buffered saline; DSP, dithiobis(succinimidyl propionate).
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ACKNOWLEDGMENTS |
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REFERENCES |
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