Article

Function of the usher N-terminus in catalysing pilus assembly

Center for Infectious Diseases, Department of Molecular Genetics and Microbiology, Stony Brook University, Stony Brook, NY 11794-5120, USA.
Molecular Microbiology (Impact Factor: 4.42). 02/2011; 79(4):954-67. DOI: 10.1111/j.1365-2958.2010.07505.x
Source: PubMed
ABSTRACT
The chaperone/usher (CU) pathway is a conserved bacterial secretion system that assembles adhesive fibres termed pili or fimbriae. Pilus biogenesis by the CU pathway requires a periplasmic chaperone and an outer membrane (OM) assembly platform termed the usher. The usher catalyses formation of subunit-subunit interactions to promote polymerization of the pilus fibre and provides the channel for fibre secretion. The mechanism by which the usher catalyses pilus assembly is not known. Using the P and type 1 pilus systems of uropathogenic Escherichia coli, we show that a conserved N-terminal disulphide region of the PapC and FimD ushers, as well as residue F4 of FimD, are required for the catalytic activity of the ushers. PapC disulphide loop mutants were able to bind PapDG chaperone-subunit complexes, but did not assemble PapG into pilus fibres. FimD disulphide loop and F4 mutants were able to bind chaperone-subunit complexes and initiate assembly of pilus fibres, but were defective for extending the pilus fibres, as measured using in vivo co-purification and in vitro pilus polymerization assays. These results suggest that the catalytic activity of PapC is required to initiate pilus biogenesis, whereas the catalytic activity of FimD is required for extension of the pilus fibre.

Full-text

Available from: Tony W Ng
Function of the usher N-terminus in catalysing
pilus assembly
mmi_7505 954..967
Nadine S. Henderson,
§
Tony W. Ng,
†§
Iehab Talukder
and David G. Thanassi*
Center for Infectious Diseases, Department of Molecular
Genetics and Microbiology, Stony Brook University,
Stony Brook, NY 11794-5120, USA.
Summary
The chaperone/usher (CU) pathway is a conserved
bacterial secretion system that assembles adhesive
fibres termed pili or fimbriae. Pilus biogenesis by the
CU pathway requires a periplasmic chaperone and an
outer membrane (OM) assembly platform termed the
usher. The usher catalyses formation of subunit–
subunit interactions to promote polymerization of
the pilus fibre and provides the channel for fibre
secretion. The mechanism by which the usher cataly-
ses pilus assembly is not known. Using the P and
type 1 pilus systems of uropathogenic Escherichia
coli, we show that a conserved N-terminal disulphide
region of the PapC and FimD ushers, as well as
residue F4 of FimD, are required for the catalytic
activity of the ushers. PapC disulphide loop mutants
were able to bind PapDG chaperone–subunit com-
plexes, but did not assemble PapG into pilus fibres.
FimD disulphide loop and F4 mutants were able to
bind chaperone–subunit complexes and initiate
assembly of pilus fibres, but were defective for
extending the pilus fibres, as measured using in vivo
co-purification and in vitro pilus polymerization
assays. These results suggest that the catalytic activ-
ity of PapC is required to initiate pilus biogenesis,
whereas the catalytic activity of FimD is required for
extension of the pilus fibre.
Introduction
Pili (fimbriae) are polymeric surface fibres expressed by a
wide variety of bacteria. Pili generally function as adhe-
sive organelles and have roles in colonization of both
abiotic and biotic surfaces, biofilm formation, interactions
with host cells, and pathogenesis. P and type 1 pili are
prototypical pilus fibres expressed by uropathogenic
Escherichia coli. P pili promote adhesion to the kidney and
the development of pyelonephritis; type 1 pili mediate
adhesion to and invasion of the bladder epithelium, initi-
ating a series of events leading to the development of
cystitis (Roberts et al., 1994; Wright et al., 2007). P and
type 1 pili are assembled by the chaperone/usher (CU)
secretion pathway, which is used by many Gram-negative
bacteria for the biogenesis of virulence-associated
surface structures (Sauer et al., 2004; Li and Thanassi,
2009). In the CU secretion pathway, a dedicated periplas-
mic chaperone controls the folding of pilus subunit pro-
teins and an integral outer membrane (OM) protein
termed the usher catalyses the assembly of subunits into
the pilus fibre and provides the channel for secretion of
the fibre to the cell surface (Nishiyama et al., 2008;
Remaut et al., 2008).
P and type 1 pili are encoded by the chromosomal pap
and fim gene clusters. Both pili consist of a rigid helical
rod that is anchored in the bacterial OM and a flexible,
linear tip fibre that is located at the distal end of the rod
(Fig. 1A) (Kuehn et al., 1992; Jones et al., 1995). In P pili,
the PapG adhesin is located in single copy at the distal
end of the pilus tip. PapG is linked by the PapF adaptor
subunit to PapE, which is present in several copies in the
tip fibre (Kuehn et al., 1992; Jacob-Dubuisson et al.,
1993). The P pilus tip is terminated by PapK, which links
the tip to the pilus rod (Fig. 1A). The rod contains over
1000 copies of PapA and is terminated by PapH (Baga
et al., 1987; Verger et al., 2006). For type 1 pili, the pilus
rod is composed of repeated copies of the FimA major
subunit protein, and the tip fibre contains single copies of
the FimH adhesin and the FimG and FimF adaptor sub-
units (Fig. 1A) (Jones et al., 1995; Saulino et al., 2000;
Hahn et al., 2002).
Biogenesis of P and type 1 pili by the CU pathway
requires the periplasmic chaperone (PapD and FimC
respectively) and OM usher (PapC and FimD
respectively). Pilus subunits enter the periplasm as
unfolded polypeptides via the Sec general secretory
pathway (Driessen and Nouwen, 2008). Upon entering
the periplasm, the subunits form stable, binary complexes
Accepted 3 December, 2010. *For correspondence. E-mail david.
thanassi@stonybrook.edu; Tel. (+1) 631 632 454; Fax (+1) 631 632
4294. Present addresses:
Department of Microbiology and Immunol-
ogy, Albert Einstein College of Medicine, Bronx, NY 10461, USA.
Department of Neurobiology and Behavior, Stony Brook University,
Stony Brook, NY11794-5230, USA.
§
Authors contributed equally.
Molecular Microbiology (2011) 79(4), 954–967 doi:10.1111/j.1365-2958.2010.07505.x
First published online 22 December 2010
© 2010 Blackwell Publishing Ltd
Page 1
with the periplasmic chaperone (Fig. 1A). The chaperone
is required for proper folding of pilus subunits, to prevent
off-pathway interactions, and to maintain subunits in an
assembly competent state (Choudhury et al., 1999; Sauer
et al., 1999; 2002; Zavialov et al., 2003). Pilus subunits
contain an incomplete immunoglobulin (Ig)-like fold termed
the pilin domain. The chaperone donates a b-strand to
complete the Ig fold of the subunit in a mechanism termed
donor strand complementation (Fig. 1) (Choudhury et al.,
1999; Sauer et al., 1999).
Periplasmic chaperone–subunit complexes next must
interact with the OM usher for release of the chaperone
and assembly of subunits into the pilus fibre. Subunit–
subunit interactions form at the periplasmic face of the
usher by a mechanism termed donor strand exchange
(Sauer et al., 2002; Zavialov et al., 2003). All pilus sub-
Fig. 1. Assembly of type 1 and P pili by the
CU pathway.
A. Model for pilus assembly. The assembly
steps for type 1 pili are shown, together with
models for the fully assembled type 1 and P
pili. The Fim and Pap proteins are indicated
by single letters (H, FimH;, etc.). Pilus
subunits enter the periplasm as unfolded
polypeptides via the Sec general secretory
pathway. The subunits fold upon interaction
with the periplasmic chaperone, forming
stable complexes via donor strand
complementation. Assembly and secretion of
the pilus fibre occurs at the OM usher, where
chaperone–subunit interactions are replaced
with subunit–subunit interactions via the donor
strand exchange reaction. During donor
strand exchange, the Nte of an incoming
chaperone–subunit complex displaces the
chaperone donor strand from the preceding
subunit. Topology diagrams are shown
depicting the donor strand complementation
and exchange reactions occurring with the
chaperone and in the pilus fibre respectively.
The dimeric ushers are depicted with the
central b-barrel domain forming a channel that
spans the OM, the N- and C-terminal domains
(labelled N and C respectively) located in the
periplasm, and the plug domain (labelled P)
gating the channel shut. Chaperone–adhesin
complexes have highest affinity for the usher
and initiate pilus assembly by binding to the
usher N-terminal domain. The pilus tip fibre is
assembled first, followed by the pilus rod.
B. Structure of the FimD
N
–CH
p
complex (PDB
ID: 1ZE3). FimC is shown in grey and FimH
p
in green. FimD
N
is shown in blue, with the
disulphide loop region colored yellow. FimD
residue F4 and the disulphide bond between
residues C63 and C90 are shown in stick
representation. The donated b-strand of the
FimC chaperone that occupies the FimH
p
subunit groove is indicated. The FimD
N
structure represents usher N-terminal domain
cartooned in (A). The C-terminal end of the
FimD
N
domain is connected by a linking
region of 14 residues to the transmembrane
b-barrel domain of the usher. The structure
was generated using PyMOL
(http://www.pymol.org).
Catalytic activity of the usher
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© 2010 Blackwell Publishing Ltd, Molecular Microbiology, 79, 954–967
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units except the adhesin contain a conserved N-terminal
extension (Nte) in addition to the pilin domain (Fig. 1A).
The adhesin contains an adhesin domain in place of the
Nte. In donor strand exchange, the Nte from an incoming
chaperone–subunit complex replaces the donated chap-
erone b-strand from the preceding chaperone–subunit
complex that is bound at the usher, completing the Ig fold
of the preceding subunit and displacing the chaperone to
form a subunit–subunit interaction. Thus, the pilus fibre is
built from an array of Ig folds, with each subunit bound
to the preceding subunit by donor strand exchange
(Fig. 1A). Pili are assembled in a defined order, with the
adhesin incorporated first, followed by the rest of the tip
and finally the rod. Both the pilus tip and rod are built from
the same donor strand exchange reaction, but pilus sub-
units in the rod undergo an additional quaternary interac-
tion upon exiting the usher that promotes coiling of the rod
into a helix on the bacterial surface (Fig. 1A). Each
subunit specifically interacts with its appropriate neigh-
bour subunit in the pilus, with the specificity of binding
determined by the donor strand exchange reaction (Lee
et al., 2007; Rose et al., 2008). In addition, the usher
ensures ordered and complete pilus assembly by
differentially recognizing chaperone–subunit complexes
according to their final position in the pilus (Dodson et al.,
1993; Saulino et al., 1998; Li et al., 2010). Pilus fibres
assemble on the order of minutes in the presence of the
usher, but on the order of hours in the absence of the
usher (Remaut et al., 2006; Vetsch et al., 2006; Nish-
iyama et al., 2008), suggesting that the usher functions to
increase the rate of fibre assembly. In agreement with this,
the FimD usher was recently shown to function as a
catalyst to accelerate the rate of polymerization of the
type 1 pilus rod by a factor of greater than 1000 (Nish-
iyama et al., 2008).
Ushers are large, integral OM proteins containing four
domains: a central transmembrane b-barrel domain that
forms the secretion channel, a middle domain located
within the b-barrel region that forms a channel gate or
plug, and soluble N- and C-terminal domains that are
located in the periplasm (Fig. 1A) (Nishiyama et al., 2005;
Shu Kin So and Thanassi, 2006; Remaut et al., 2008;
Huang et al., 2009; Mapingire et al., 2009; Ford et al.,
2010). The N-terminal domain (PapC residues 1–131,
FimD residues 1–125) provides the initial binding site for
chaperone–subunit complexes (Ng et al., 2004; Nish-
iyama et al., 2005), whereas the C-terminal domain
(PapC residues 641–809, FimD residues 664–833) stabi-
lizes the binding of chaperone–subunit complexes to the
usher and may participate in controlling access to the
usher channel (Thanassi et al., 2002; Shu Kin So and
Thanassi, 2006; Ford et al., 2010). Both the N- and
C-terminal domains also participate in the differential
affinity of the usher for chaperone–subunit complexes (Li
et al., 2010). The usher is present as a dimeric complex in
the OM, but only one channel is used for secretion of the
pilus fibre (Li et al., 2004; Shu Kin So and Thanassi, 2006;
Remaut et al., 2008). The usher dimer may facilitate
recruitment of chaperone–subunit complexes from the
periplasm and help position subunits for donor strand
exchange (Remaut et al., 2008; Li and Thanassi, 2009).
In a previous analysis of the usher N-terminal domain,
we found that a PapC mutant deleted for residues 2–11
was unable to bind chaperone–subunit complexes or
assemble P pili (Ng et al., 2004). Alanine scanning
mutagenesis of this region identified only residue F3 as
critical for the binding activity of the PapC N-terminus; the
PapC F3A mutant was defective for pilus biogenesis and
did not bind chaperone–subunit complexes (Ng et al.,
2004; 2006). The structural basis for these findings was
revealed by Nishiyama and colleagues (Nishiyama et al.,
2005), who solved the structure of the FimD N-terminal
domain (FimD
N
, residues 1–125) bound to a FimC
chaperone–FimH pilin domain complex (FimCH
p
). This
structure revealed that the first 24 residues of FimD spe-
cifically interact with the bound FimCH
p
complex and that
residue F4, which corresponds to PapC F3, directly con-
tacts the FimC chaperone (Fig. 1B) (Nishiyama et al.,
2005). Similar to the PapC F3A mutant, a FimD F4A
mutant was defective for pilus biogenesis (Nishiyama
et al., 2005). In addition to the chaperone–subunit binding
region, the usher N-terminal domain contains a conserved
pair of cysteines (PapC C70 and C97; FimD C63 and
C90) that form a disulphide bond (Fig. 1B) (Henderson
et al., 2004; Nishiyama et al., 2005). We previously dem-
onstrated that PapC ushers containing mutations to either
or both of the N-terminal cysteines were unable to
assemble pili (Ng et al., 2004). However, in contrast to the
PapC D2–11 and F3A mutants, the PapC cysteine
mutants maintained the ability to bind PapDG chaperone–
adhesin complexes (Ng et al., 2004). This suggested that
the N-terminal cysteines function at a step distinct from
the initial binding of chaperone–subunit complexes. In the
FimD
N
–CH
p
co-crystal structure (Nishiyama et al., 2005),
the FimD region containing the conserved N-terminal cys-
teines makes no interactions with the bound FimCH
p
complex (Fig. 1B) and therefore the structure does not
provide insight into the function of this region.
In this study, we investigated the mechanism of the
conserved disulphide region of the usher N-terminus in
pilus biogenesis. We present evidence that this region is
critical for the catalytic activity of both PapC and FimD in
polymerizing pilus subunits. Differences were detected
between the two ushers, which belong to separate clades
of the usher superfamily (Nuccio and Baumler, 2007). In
contrast to the essential role of PapC residue F3 in
binding chaperone–subunit complexes, we found that
residue F4 of FimD functions in the catalytic activity of the
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© 2010 Blackwell Publishing Ltd, Molecular Microbiology, 79, 954–967
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usher and is not strictly required for chaperone–subunit
binding. In addition, the PapC and FimD mutants were
blocked at different stages of pilus biogenesis, suggesting
distinct requirements for the catalytic activity of the ushers
in initiation and extension of the pilus fibre.
Results and discussion
The conserved disulphide region in the PapC
N-terminus functions to promote subunit–subunit
interactions
Previous studies demonstrated that the conserved
N-terminal cysteines of PapC (C70 and C97) are required
for P pilus biogenesis but function at a step distinct from
the role of the usher N-terminus in the initial binding of
chaperone–subunit complexes (Ng et al., 2004). The
N-terminal cysteines form a disulphide bond (Henderson
et al., 2004; Nishiyama et al., 2005). Thus, the phenotype
of the cysteine mutants might be an indirect effect caused
by loss of the disulphide bond and resulting structural
changes in the loop region bounded by the cysteines
(Fig. 1B). In initial studies, we were unable to identify
single residues that, when mutated to alanine, were
required for function of the loop region (T.W. Ng, I.
Talukder and D.G. Thanassi, unpubl. data). Therefore, we
constructed a PapC mutant deleted for residues 77–86,
which occur in the middle of the loop region. This internal
deletion was chosen to avoid disturbing the disulphide
bond and because it encompasses several conserved
residues. PapC D77–86 was present at a similar level in
the OM compared with wild-type (WT) PapC (Fig. 2A,
upper panel) and the mutation did not affect the overall
folding of the usher, as assessed by resistance to dena-
turation by SDS (data not shown). Similar to a PapC C70A
mutant, PapC D77–86 was unable to complement a
plasmid bearing a DpapC pap operon (papAHDJKEFG)
for assembly of adhesive P pili on the bacterial surface, as
assessed by the haemagglutination (HA) assay (Table 1).
Also similar to the C70A mutant, PapC D77–86 retained
the ability to bind PapDG complexes, as measured using
an in vitro overlay assay (Fig. 2A). This is in contrast to a
PapC F3A mutant, which was unable to bind PapDG
(Fig. 2A), as shown previously (Ng et al., 2004; 2006).
The identical behaviour of the PapC D77–88 and C70A
mutants suggests that the loop region between the con-
served N-terminal cysteines is functionally important. In
addition, the fact that both mutants retain the ability to
bind PapDG complexes indicates that loss of the disul-
phide bond or deletion within the loop region does not
cause global structural changes to the usher N-terminus.
To understand why the PapC C70A and D77–86
mutants were defective for pilus biogenesis despite being
able to bind PapDG chaperone–adhesin complexes, we
coexpressed the PapC mutants with P pilus tip subunits
(papDJKEFG) and tested the ability of the tip subunits to
form stable assembly intermediates with the ushers in
vivo using a co-purification assay. Immunoblotting with an
antibody that recognizes P pilus tips showed that whereas
PapG, F, E and K co-purified with WT PapC, only PapG
co-purified with the C70A and D77–86 mutants (Fig. 2B).
In comparison, no pilus subunits co-purified with the PapC
F3A mutant (Fig. 2B), as expected since this mutant is
defective for binding chaperone–subunit complexes. A
Fig. 2. Analysis of the PapC N-terminal disulphide loop mutants.
A. Overlay assay for binding of PapDG to PapC. OM fractions were
isolated from strain SF100 expressing vector only, WT PapC, or the
indicated PapC mutant. Duplicate samples were subjected to
SDS-PAGE and either stained with Coomassie blue (upper panel)
or transferred to a PVDF membrane for the overlay assay. Binding
of PapDG to the usher (lower panel) was determined by
immunoblotting with anti-PapDG antibodies.
B. Co-purification of pilus tip subunits with PapC. OM fractions
were isolated from strain SF100/pPAP58 (papDJKEFG) expressing
vector only, WT PapC, or the indicated PapC mutant. His-tagged
PapC was purified from the OM fractions, subjected to SDS-PAGE,
and either stained with Coomassie blue to show the amount of
PapC present (upper panel) or immunoblotted with anti-P pilus tips
antiserum to detect pilus tip subunits that co-purified with the usher
(PapG, K, E and F, lower panel).
C. Assembly of PapG into pilus tip fibres. Co-purification of pilus tip
subunits with WT PapC or the indicated PapC mutant was
performed as in (B). The samples were incubated at 25°C or 95°C
in SDS-PAGE sample buffer, subjected to SDS-PAGE and
immunoblotted with anti-PapDG antibodies.
Catalytic activity of the usher
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© 2010 Blackwell Publishing Ltd, Molecular Microbiology, 79, 954–967
Page 4
very low level of chaperone–subunit complexes may have
co-purified with the PapC mutants, as indicated by faint
bands present in the F3A and C70A lanes in Fig. 2B,
suggesting some remaining functionality. Nevertheless,
the data show that even though the PapC C70A and
D77–86 mutants are able to perform the first step in pilus
biogenesis, which is binding to PapDG, the mutants are
defective for carrying out subsequent pilus assembly
steps. In agreement with this, the PapC C70A and D77–86
mutants were unable to assemble adhesive P pilus tip
fibres on the bacterial surface, as measured by the HA
assay, in contrast to WT PapC (Table 1).
If the PapC cysteine loop mutants are indeed blocked
at the first assembly step, then the PapG that co-purified
with the usher mutants should not be engaged in
subunit–subunit interactions. To test this, we incubated
the proteins that co-purified with WT PapC, PapC C70A
or PapC D77–86 in SDS sample buffer at 95°C or 25°C
and performed an immunoblot analysis with anti-PapDG
antibodies. Subunit–subunit but not chaperone–subunit
interactions are stable to SDS at low temperatures, so
any PapG engaged in donor strand exchange with other
pilus subunits (i.e. assembled into a pilus fibre) will shift
to higher molecular weight when incubated at 25°C
(Soto et al., 1998; Saulino et al., 2000). As shown in
Fig. 2C, the PapG that co-purified with WT PapC ran at
the expected monomer molecular weight of 36 kDa
when incubated at 95°C. When incubated at 25°C, the
monomer PapG band disappeared and a ladder of
higher molecular weight species appeared, demonstrat-
ing that all of the PapG was polymerized into tip fibres of
various lengths. In contrast, the PapG that co-purified
with the C70A or D77–86 mutants remained as a
monomer when incubated at either 25°C or 95°C, with
only very faint ladders of higher molecular weight
species visible at 25°C (Fig. 2C). These data indicate
that the PapC disulphide loop mutants are indeed
largely blocked at the first stage of pilus assembly and
are defective for promoting the polymerization of PapG
into pilus fibres.
The FimD N-terminus functions to promote
subunit–subunit interactions
To determine if the N-terminal disulphide region of the
FimD usher functions similarly to PapC in promoting
subunit polymerization, we constructed a FimD D70–79
deletion mutant, which corresponds to the PapC D77–86
mutation. For comparison, we also constructed a FimD
F4A mutation, which corresponds to the PapC F3A
mutation. The FimD mutants were present at similar levels
in the OM compared with WT FimD (Fig. 3A, upper panel)
and the mutations did not affect the overall folding of the
ushers, as assessed by resistance to denaturation by
SDS (data not shown). We first tested the ability of the
FimD mutants to complement a DfimD mutation in the
chromosomal fim gene cluster for assembly of type 1 pili,
using the HA assay. As found for PapC, the FimD F4A and
D70–79 mutants were unable to assemble adhesive pili
on the bacterial surface (Table 1).
We next tested the FimD F4A and D70–79 mutants for
ability to bind FimCH chaperone–adhesin complexes,
using the in vitro overlay assay. Despite being unable to
assemble pili, FimD D70–79 bound to FimCH similar to
WT FimD (Fig. 3A), matching the phenotype of the PapC
D77–86 mutant. Surprisingly, the FimD F4A mutant also
bound to FimCH in the overlay assay (Fig. 3A). This is in
contrast to the PapC F3A mutant, which was unable to
bind chaperone–subunit complexes, and different from
the findings of Nishiyama and colleagues, who reported
that the F4A mutation prevented the isolated FimD
N
domain from binding FimCH
p
(Nishiyama et al., 2005).
This difference with Nishiyama et al. is likely due to the
fact that we used full-length FimD usher instead of only
the N-terminal domain, and full-length FimH adhesin
instead of only the pilin domain, providing additional sur-
faces for the usher to bind the adhesin. Indeed, previous
studies showed that FimCH makes stable interactions
with a C-terminal region of the usher in addition to the
N-terminal domain, and that the FimH adhesin domain
by itself binds to the usher (Saulino et al., 1998; Barn-
hart et al., 2003; Munera et al., 2007). Based on these
results, we propose that FimD F4 contributes to, but is
not essential for, the binding of chaperone–subunit com-
plexes to the usher. These results reveal differences in
the mechanism by which the PapC and FimD ushers
bind pilus subunits, with PapC residue F3 playing a
more critical role compared with the corresponding FimD
F4 residue.
Table 1. Ability of the PapC and FimD usher mutants to assemble
adhesive pili or pilus tips.
PapC
HA titre
a
pili
(papAHDJKEFG)
HA titre
a
tips
(papDJKEFG)
Vector 0 0
WT 64 256
F3A 0 0
C70A 0 0
D77–86 0 0
FimD
HA titre
a
pili
(fimAICFGH)
HA titre
a
tips
(fimCGH)
Vector 0 0
WT 128 256
F4A 0 0
D70–79 0 0
a. HA titre is the highest-fold dilution of bacteria able to agglutinate
human (for P pili) or guinea pig (for type 1 pili) red blood cells.
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To investigate why the FimD F4A and D70–79 mutants
were unable to assemble type 1 pili despite being able to
bind FimCH complexes, we expressed the usher mutants
in bacteria together with type 1 pilus tip subunits
(fimCFGH) and performed co-purification assays. The
purified proteins were incubated at 25°C or 95°C in SDS
sample buffer and immunoblotted with anti-FimCH
antibodies. Analysis of the samples treated at 95°C
showed that similar levels of FimC and FimH co-purified
with WT FimD and the F4A and D70–79 mutants (Fig. 3B).
This confirms the findings from the overlay assay (Fig. 3A)
and demonstrates that the FimD mutants are able to form
stable complexes with FimCH in vivo. Unexpectedly,
analysis of the samples incubated at 25°C revealed that
the F4A and D70–79 mutants were also able to assemble
FimH into pilus tip fibres (Fig. 3B). This is in contrast to the
PapC C70A and D77–86 mutants, which were highly
defective for assembling the adhesin into pilus fibres
(Fig. 2B and C). For the FimD samples treated at 25°C,
bands at sizes expected for FimGH and FimFGH com-
plexes were present for both the WT and mutant ushers.
The composition of the FimGH and FimFGH bands was
confirmed using anti-FimCG and anti-FimCF antibodies
(data not shown). However, subtle differences between
the WT and mutant FimD ushers were apparent. For WT
FimD, the FimH monomer band (29 kDa) completely dis-
appeared in the samples treated at 25°C, indicating incor-
poration of all the FimH into tip fibres, and a faint but
distinct ladder of higher molecular weight bands was
visible above the FimFGH band (Fig. 3B). For the FimD
F4A and D70–79 mutants, some monomer FimH
remained in the 25°C-treated samples and distinct bands
migrating above the FimFGH band were absent (Fig. 3B).
The faint ladder of higher molecular weight bands
observed with WT FimD corresponds to a low level of type
1 pilus tip fibres containing multiple FimG subunits (see
below), as confirmed by blotting with anti-FimCG anti-
bodies (data not shown).
To explore further the differences between the WT
and mutant FimD ushers, we performed a modified
co-purification experiment in which the ushers were
expressed in bacteria together with only fimCGH (omitting
the FimF tip subunit). FimG is capable of interacting with
itself to form polymers. However, type 1 pilus tip fibres
normally contain only single copies of FimG because
FimF has high affinity for FimG and binding of FimF pre-
vents FimG–FimG interactions (Saulino et al., 2000).
Omitting the FimF subunit allowed the assembly of type 1
tip fibres consisting of FimH bound to various numbers of
FimG. Analysis of the samples that co-purified with WT
FimD using anti-FimCG antibodies at 25 and 95°C clearly
illustrates the polymerization of FimG into a ladder of
higher molecular weight species (Fig. 3C). Immunoblot-
ting with anti-FimCH antibody confirmed the incorporation
of the adhesin into these fibres (data not shown). Inter-
estingly, compared with WT FimD, the F4A and D70–79
mutants were greatly impaired in polymerizing FimG, with
only distinct bands corresponding to FimGH, GGH and
GGGH fibres visible (Fig. 3C). Thus, the FimD mutants
are able to bind chaperone–subunit complexes and ini-
tiate fibre assembly, but appear to be defective in extend-
ing the pilus fibres. This behaviour of the FimD F4A and
D70–79 mutants is similar to the PapC C70A and D77–86
mutants, demonstrating that the N-terminal domains
Fig. 3. Analysis of the FimD F4A and D70–79 mutants.
A. Overlay assay for binding of FimCH to FimD. OM fractions were
isolated from strain SF100 expressing vector only, WT FimD, or the
indicated FimD mutant. Duplicate samples were subjected to
SDS-PAGE and either stained with Coomassie blue (upper panel)
or transferred to a PVDF membrane for the overlay assay. Binding
of FimCH to the usher (lower panel) was determined by
immunoblotting with anti-FimH antibody.
B. Co-purification of FimCFGH pilus tip fibres with FimD. OM
fractions were isolated from strain AAEC185/pNH235 (fimCFGH)
expressing vector only, WT FimD, or the indicated FimD mutant.
His-tagged FimD was purified from the OM fractions, incubated at
25°C or 95°C in sample buffer, subjected to SDS-PAGE, and
immunoblotted with anti-FimCH antibodies. The identities of the
complexes that co-purified with FimD are indicated on the right
using single letters to represent the Fim proteins (H, FimH;, etc.).
C. Co-purification of FimCGH pilus tip fibres with FimD. The assay
was performed as in (B), except the host strain was
AAEC185/pNH222 (fimCGH) and the samples were immunoblotted
with anti-FimCG antibodies.
Catalytic activity of the usher
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of both ushers function to promote subunit–subunit
interactions. However, the subunit-polymerizing activities
of the PapC and FimD N-termini appear to be required
for different stages of pilus biogenesis, with the PapC
mutants defective for initiation of fibre assembly and the
FimD mutants defective for fibre elongation.
Given the ability of the FimD F4A and D70–79 mutants
to assemble FimFGH pilus tip fibres (Fig. 3B), it was sur-
prising that these mutants were unable to complement the
DfimD fim gene cluster for assembly of adhesive pili
(Table 1). The FimD mutants should have been able to
assemble type 1 pilus tips, exposing the FimH adhesin to
the extracellular side of the OM and allowing for HA activ-
ity (Saulino et al., 2000; Remaut et al., 2008). A possible
explanation for these results is that the FimFGH tip fibres
may be too short to sufficiently expose FimH beyond the
OM to provide robust agglutination. Indeed, we found that
the HA activity of bacteria expressing only type 1 pilus tips
(WT FimD coexpressed with FimCFGH) was very low and
close to background levels (HA titre = 8), compared with
bacteria expressing complete pili (HA titre = 128). In con-
trast, expression of FimD together with only FimCGH
(omitting FimF) led to very strong agglutination activity
(HA titre = 256; Table 1), presumably because the longer
FimG
n
–FimH fibres extended sufficiently beyond the bac-
terial surface. Notably, bacteria in which the FimD F4A
and D70–79 mutants were expressed together with only
FimCGH still lacked HA activity (Table 1). Therefore, even
though the F4A and D70–79 mutants are able to initiate
assembly of pilus tip fibres, the fibres assembled by the
mutants are presumably too short to mediate adhesive
activity. These results, together with the results of the
co-purification experiments (Fig. 3C), also indicate that
the defect of the FimD mutants in polymerizing pilus fibres
is not due to an inability to transition from assembly of the
pilus tip to the pilus rod; pilus fibres were not elongated
even in the absence of the FimA rod subunit.
The FimD N-terminus is required for the catalytic activity
of the usher
The defects of the FimD and PapC N-terminal domain
mutants in promoting subunit–subunit interactions
suggest that in addition to providing the initial binding site
for chaperone–subunit complexes, the usher N-terminus
may be important for the catalytic activity of the usher. The
catalytic activity of FimD is activated by the binding of
FimCH chaperone–adhesin complexes to the usher
(Nishiyama et al., 2008). Binding of FimCH causes a con-
formational change in FimD, detectable by a change in
the sensitivity of the usher to digestion by extracellularly
added trypsin, resulting in the appearance of a protected
C-terminal fragment of the usher (Saulino et al., 1998). To
determine if the defect of the FimD F4A and D70–79
mutants in polymerizing pilus subunits was due to a loss
of activation of the usher by FimCH, we coexpressed
FimD in bacteria together with FimCH and examined sen-
sitivity of the usher to trypsin digestion. As observed pre-
viously (Saulino et al., 1998), coexpression of FimCH
with WT FimD resulted in the appearance of a trypsin-
protected FimD fragment. Similar trypsin-protected frag-
ments also appeared for the FimD F4A and D70–79
mutants (data not shown). Thus, the FimD mutants main-
tain the ability to be activated by FimCH, in agreement
with our findings above that the FimD mutants are able to
bind FimCH both in vitro and in vivo (Fig. 3). In contrast to
FimD, PapC does not require interaction with the adhesin
to initiate pilus assembly (Li et al., 2010); it remains to be
determined whether the catalytic activity of PapC might be
activated by some other mechanism.
To directly assess the catalytic activity of the usher in
polymerizing pilus subunits, we developed an in vitro
reconstitution assay for type 1 pilus biogenesis. For this
assay, we separately purified OM fractions expressing the
FimD usher and periplasm fractions expressing either
FimCH or His-tagged FimC (FimC
His
)–FimG complexes.
The periplasm and OM fractions were mixed together and
incubated at room temperature (25°C). At various time
points, the samples were placed on ice and centrifuged to
collect the OM fragments. The OM was then solubilized
with non-denaturing detergent and the extracted OM pro-
teins were passed over a metal affinity column to capture
pilus tip fibres assembling through the FimD usher. Frac-
tions eluted from the metal affinity column were then
incubated in SDS sample buffer at 25°C and analysed by
quantitative immunoblotting using anti-FimCG antibodies.
Note that only OM extracts were run over the affinity
column and that only the FimC chaperone expressed
together with FimG contained a His-tag. These measures
allowed us to select for active pilus assembly intermedi-
ates in which the His-tagged chaperone was bound to the
last incorporated FimG subunit at the usher (Remaut
et al., 2008).
We first tested the catalytic activity of WT FimD in the in
vitro reconstitution assay. As shown in Fig. 4A, FimG
n
FimH complexes appeared within 5 min of incubation and
increased in both intensity and number of FimG subunits
over the 45 min time-course of the experiment. The incor-
poration of FimH into the various complexes was con-
firmed using anti-FimCH antibodies (data not shown). No
FimG
n
–FimH complexes were obtained if OM lacking
FimD was used for the reconstitution (data not shown).
Quantification of the increase in intensity of the FimG
n
FimH complexes over time with WT FimD is shown in
Fig. 4D. We next tested the catalytic activity of the FimD
F4A and D70–79 mutants in the in vitro reconstitution
assay. As shown in Fig. 4C and D, the FimD F4A mutant
was unable to assemble FimG
n
–FimH complexes over the
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N. S. Henderson, T. W. Ng, I. Talukder and D. G. Thanassi
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course of the experiment. In contrast, some FimG
n
–FimH
complexes were assembled by the FimD D70–79 mutant,
although to a much lower extent compared with WT FimD
(Fig. 4B and D). In addition, the complexes assembled by
the D70–79 mutant appeared to be largely stalled at the
FimG
1
-FimH stage (Fig. 4B). These data show that resi-
dues F4 and 70–79 are indeed critical for the catalytic
activity of the usher in promoting subunit–subunit interac-
tions, as suggested by the co-purification experiments
(Fig. 3C). The more severe defect of the FimD F4A
mutant, compared with the D70–79 mutant in the recon-
stitution assay, may be a reflection of the involvement of
residue F4 in binding chaperone–subunit complexes, as
well as in catalysing fibre assembly. Although the F4A
mutant showed no catalytic activity in the reconstitution
assay, the co-purification assay clearly showed that this
mutant was able to initiate assembly of type 1 tip fibres in
vivo similar to the D70–79 disulphide loop mutant (Fig. 3B
and C). The stronger phenotype obtained in the reconsti-
tution assay likely reflects the more stringent conditions
present in this vitro system, including lower temperature,
limited time-course and differences in protein concentra-
tions compared with pilus biogenesis in vivo.
To measure fibre assembly, both the in vitro reconstitu-
tion assay and in vivo co-purification assay took advan-
tage of the resistance of subunit–subunit interactions
(donor strand exchange), but not chaperone–subunit
interactions (donor strand complementation), to dissocia-
tion by SDS. Moreover, both assays captured pilus fibres
in stable association with the usher; these complexes
presumably represent bona fide assembly intermediates
in the process of secretion through the usher. In compari-
son, previous in vitro polymerization assays have relied
on incubating chaperone–subunit complexes with the
usher and using electron microscopy to measure the
lengths of pilus fibres formed (Nishiyama et al., 2008;
Huang et al., 2009). It is possible in these assays that a
portion of the fibres formed might be the result of off-
pathway subunit–subunit interactions triggered by interac-
tion with usher domains and the high concentrations of
subunits present, rather than the result of assembly and
secretion of the fibres through the usher. For example,
Huang et al. (2009) reported that the PapC N domain was
dispensable for fibre polymerization in their in vitro assay,
despite its known requirement for pilus biogenesis in vivo.
In contrast, our FimD results show that a functional usher
N-terminus is required for pilus assembly in vitro as well
as in vivo. We have not yet succeeded in directly testing
the catalytic activity of PapC using an in vitro reconstitu-
tion assay similar to the assay we used for FimD.
However, the co-purification experiments (Fig. 2C) clearly
showed defects of the PapC C70A and D77–86 mutants in
polymerizing pilus fibres, and support a catalytic role for
the N-terminal disulphide region of PapC, as found for
Fig. 4. In vitro reconstitution assay for polymerization of type 1
pili.
A–C. Reconstitution assay for pilus assembly. OM fractions were
isolated from strain Tuner expressing WT FimD (A), FimD D70–79
(B) or FimD F4A (C). The OM fractions were mixed together with
separately isolated periplasm fractions containing FimCH and
FimC
His
G and incubated at 25°C for the indicated time points. Pilus
assembly intermediates purified from the OM fractions were
incubated in sample buffer at 25°C, subjected to SDS-PAGE, and
immunoblotted using anti-FimCG antibodies. The blots were
analysed using the Odyssey Infrared Imaging System.
D. Quantification of FimG
n
–FimH complex formation. The amount of
FimG
n
–FimH complexes assembled at each time point in the
reconstitution assay, preformed as in (A–C), was determined by
measuring the signal intensity within identical areas boxed as
shown in (A) for the 45 min time point. The values for WT FimD
represent means standard deviation of at least five separate
experiments; the values for the FimD D70–79 and F4A mutants
represent means of two separate experiments.
Catalytic activity of the usher
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FimD. An alternative explanation for the failure of the
PapC disulphide loop mutants to assemble pili is that
these mutants are altered in their affinity for chaperone–
subunit complexes or unable to bind subunits other than
PapG. However, we found that the binding affinity of the
PapC C70A mutant for PapDG is unchanged compared
with WT PapC, and that this mutant is able to bind PapDF
chaperone–subunit complexes in addition to PapDG (Q.
Li and D.G. Thanassi, unpubl. results). Taken together,
these results indicate a conserved function of the
usher N-terminal domain in promoting subunit–subunit
interactions.
Conclusions
The usher acts as a multifunctional assembly and secre-
tion platform in the bacterial OM. The usher N-terminal
domain was previously shown to provide the initial
binding site for chaperone–subunit complexes and par-
ticipate in the differential affinity of the usher for
chaperone–subunit complexes (Ng et al., 2004; Nish-
iyama et al., 2005; Li et al., 2010). Our findings reported
here reveal a third function for the usher N-terminus:
catalysing the exchange of chaperone–subunit for
subunit–subunit interactions to promote polymerization
of the pilus fibre. In particular, we identified the con-
served N-terminal disulphide region of the PapC and
FimD ushers, as well as residue F4 in the chaperone–
subunit binding region of FimD, as important for the
catalytic activity of the usher.
The mechanism by which the usher catalyses pilus
assembly is not known. The formation of subunit–subunit
interactions at the usher occurs by a concerted strand
displacement mechanism (Remaut et al., 2006; Vetsch
et al., 2006), in which the Nte of an incoming chaperone–
subunit complex displaces the chaperone donor strand
from the preceding chaperone–subunit complex to allow
donor strand exchange between the subunits (Fig. 1A).
Nucleation of the donor strand exchange reaction is criti-
cally dependent on insertion of the Nte into a binding
pocket in the subunit groove occupied by the chaperone
donor strand (Remaut et al., 2006). Therefore, the cata-
lytic activity of the usher is likely related to the positioning
of the incoming Nte relative to the binding pocket of the
preceding chaperone–subunit complex to favour the
donor strand exchange reaction. The usher may also act
to weaken the chaperone–subunit interaction, thereby
facilitating invasion of the incoming Nte. The FimD
N
–CH
p
co-crystal structure (Nishiyama et al., 2005) shows that
the N-terminal disulphide region of the usher does not
directly interact with the bound chaperone–subunit
complex (Fig. 1B), and we found that mutations to the
disulphide loop region do not affect the binding of
chaperone–subunit complexes to the usher. In the context
of the full-length usher, the disulphide loop region would
be located adjacent to the strand connecting the
N-terminal domain to the b-barrel translocation domain of
the usher (Fig. 1). Taking these findings together, we
propose that the disulphide loop region may influence
positioning of the chaperone–subunit complex relative to
the rest of the usher-pilus assembly complex; i.e. the
function of the disulphide loop region in catalysis may be
to ensure optimal placement of the Nte of the incoming
chaperone–subunit complex relative to the subunit groove
of the preceding chaperone–subunit complex to promote
donor strand exchange. In contrast to the disulphide loop
region, residue F4 of FimD directly contacts the FimC
chaperone (Fig. 1B). Therefore, FimD F4 may act to
weaken the binding of the chaperone to its subunit and
thus promote donor strand exchange by facilitating inva-
sion of the incoming Nte.
Although the N-terminal domains of both PapC and
FimD were found to function similarly in promoting subunit
polymerization, we also observed differences between
the two ushers. PapC and FimD belong to different clades
within the usher superfamily (Nuccio and Baumler, 2007)
and these differences likely reflect evolutionary diver-
gence in their assembly mechanisms. Our results showed
that PapC and FimD do not bind chaperone–subunit com-
plexes by identical mechanisms, since residue F3 is
required for the binding activity of PapC, whereas the
corresponding FimD F4 residue is not essential for
binding. A structure of the PapC–DG complex will be
important to reveal the basis for this difference. Our
results also revealed that the catalytic activity of the PapC
and FimD ushers is required for different stages of pilus
biogenesis. The PapC N-terminal disulphide mutants
were almost completely stalled at the stage of PapDG
binding to the usher, with only barely detectable levels of
PapG polymerization into pilus fibres. Thus, the catalytic
activity of PapC appears to be critical for formation of the
first subunit–subunit interaction and initiation of fibre
assembly (presumably, the PapC catalytic activity is also
required for fibre elongation once initiation of assembly
takes place). In contrast, the FimD F4A and D70–79
mutants behaved similar to WT FimD in initiating pilus
assembly, but were defective for assembling higher order
pilus fibres. This suggests that the catalytic activity of
FimD is required following initiation for subsequent rounds
of subunit incorporation necessary to extend the pilus
fibre. This agrees with the findings of Nishiyama and
colleagues, who examined the catalytic activity of FimD
during assembly of the FimA pilus rod and concluded that
the usher accelerates polymerization of the rod following
incorporation of the first FimA subunit (Nishiyama et al.,
2008). Overall, our findings highlight important similarities
as well as specific differences in the mechanism of pilus
assembly by the PapC and FimD ushers. Our results
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© 2010 Blackwell Publishing Ltd, Molecular Microbiology, 79, 954–967
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provide a foundation for exploring the molecular basis of
the catalytic activity of the ushers and for understanding
virulence factor biogenesis at the bacterial OM.
Experimental procedures
Strains and plasmids
The bacterial strains and plasmids used in this study are
listed in Table 2. Unless otherwise indicated, all strains were
grown at 37°C with aeration in Luria–Bertani (LB) medium
with appropriate antibiotics. DH5a was used as the host
strain for plasmid manipulations and all mutants were
sequenced to verify the intended mutation.
Plasmid pTN26, encoding PapC D77–86 with a C-terminal
hexahistadine tag (His-tag) was derived from plasmid pMJ3
using the QuikChange Site-Directed Mutagenesis Kit (Strat-
agene) and the primers listed in Table 3. Plasmids pIT2
(FimD F4A), pIT4 (FimD C63A) and pIT12 (FimD D70–79)
were derived from plasmid pETS7, which encodes FimD with
a C-terminal His-tag, using the QuikChange Site-Directed
Mutagenesis Kit and the primers listed in Table 3. Plasmids
pNH227 (FimD F4A), pNH228 (FimD C63A) and pNH236
(FimD D70–79) were similarly derived from plasmid pETS4,
which encodes FimD with a C-terminal His-tag. Plasmids
pNH238 (FimD F4A) and pNH239 (FimD D70–79) were simi-
larly derived from plasmid pAN2, which encodes FimD (no
His-tag).
Plasmid pNH222 was constructed by subcloning an
EcoRI–HindIII fragment containing fimCGH from plasmid
pETS1013 (Saulino et al., 2000) into vector pBAD18-Kan.
Plasmid pNH235, which contains fimCFGH in vector
pBAD18-cm, was constructed by subcloning an EcoRI–KpnI
Table 2. Strains and plasmids used in this study.
Strain or plasmid Relevant characteristic(s) Reference or source
Strains
a
DH5a hsdR recA endA
Grant et al. (1990)
Tuner OmpT
-
Lon
-
Novagen
BL21 OmpT
-
Lon
-
Rosenberg et al. (1987)
SF100 DompT
Baneyx and Georgiou (1990)
AAEC185 Dfim
Blomfield et al. (1991)
ORN103 Dfim
Orndorff et al. (1985)
MM294DfimD DfimD
Shu Kin So and Thanassi (2006)
Plasmids
pMON6235Dcat
vector, P
ara
, Amp
r
Jones et al. (1997)
pMMB66
vector, P
tac
, Amp
r
Morales et al. (1991)
pMMB91
vector, P
tac
, Kan
r
Dodson et al. (1993)
pBAD18-cm
vector, P
ara
, Clm
r
Guzman et al. (1995)
pBAD18-kan
vector, P
ara
, Kan
r
Guzman et al. (1995)
pMJ3 PapC with C-terminal His-tag in pMON6235Dcat
Thanassi et al. (1998)
pM05 PapC F3A in pMJ3
Ng et al. (2004)
pTN5 PapC C70A in pMJ3
Ng et al. (2004)
pTN26 PapC D77–86 in pMJ3 This study
pMJ2
DpapC pap operon in pACYC184, P
trc
,Tet
r
Thanassi et al. (1998)
pPAP58 PapDJKEFG in pMMB91
Hultgren et al. (1989)
pJP1 PapDG in pMMB91
Dodson et al. (1993)
pETS4 FimD with C-terminal His-tag in pMMB66
Saulino et al. (1998)
pETS7 FimD with C-terminal His-tag in pBAD18-cm
Saulino et al. (2000)
pAN2 FimD (no His-tag) in pMMB91
Saulino et al. (1998)
pNH227 FimD F4A in pETS4 This study
pIT2 FimD F4A in pETS7 This study
pNH238 FimD F4A in pAN2 This study
pNH228 FimD C63A in pETS4 This study
pIT4 FimD C63A in pETS7 This study
pNH236 FimD D70–79 in pETS4 This study
pIT12 FimD D70–79 in pETS7 This study
pNH239 FimD D70–79 in pAN2 This study
pNH222 FimCGH in pBAD18-kan This study
pNH235 FimCFGH in pBAD18-cm This study
pETS1007 FimCH in pMMB66
Saulino et al. (2000)
pNH221 FimC (no His-tag) in pBAD18-cm This study
pETS1000
FimC with C-terminal His-tag in pMON6235Dcat, Spec
r
Saulino et al. (1998)
pHJ20 FimH in pMMB66
Jones et al. (1995)
pETS2A FimG in pMMB66
Saulino et al. (1998)
a. All strains are E. coli K-12, except BL21 and Tuner, which are E. coli B.
Amp
r
, ampicillin resistance; Kan
r
, kanamycin resistance; Clm
r
, chloramphenicol resistance; Tet
r
, tetracycline resistance; Spec
r
, spectinomycin
resistance; P
ara
, arabinose-inducible promoter, P
tac
or P
trc
, IPTG-inducible promoter.
Catalytic activity of the usher
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fragment containing fimC from plasmid pETS1007 and a
SmaI–HindIII fragment containing fimFGH from plasmid
pHJ27 (Hultgren laboratory collection, Washington Univer-
sity, St Louis, MO, USA). Plasmid pNH221 was constructed
by subcloning fimC from pETS1007 into vector pBAD18-cm.
OM isolation and analysis of usher expression and
folding
The expression levels of the PapC and FimD usher mutants
in the OM were compared with the WT parental usher. Strain
SF100 harbouring the appropriate PapC or FimD expression
plasmid was induced at OD
600
= 0.6 with 0.1% L-arabinose
(for PapC) or 50 mM isopropyl-b-
D-thiogalactoside (IPTG; for
FimD) for 1 h. OM fractions were isolated by French press
disruption and Sarkosyl extraction, as previously described
(Ng et al., 2004; Shu Kin So and Thanassi, 2006). Expression
levels of the ushers in the OM were determined by inspection
of Coomassie blue-stained SDS-PAGE gels or immunoblot-
ting with anti-His-tag (Covance), anti-PapC or anti-FimD
antibodies. Immunoblots were developed with alkaline
phosphatase-conjugated secondary antibodies and BCIP
(5-bromo-4-chloro-3-indolylphosphate)-NBT (nitroblue tetra-
zolium) substrate (KPL). Proper folding of the ushers in the
OM was checked by resistance to denaturation by SDS,
which provides an indication of the correct folding and stabil-
ity of the b-barrel domain (Sugawara et al., 1996). This was
determined by heat-modifiable mobility on SDS-PAGE, per-
formed as previously described (Ng et al., 2004; Shu Kin So
and Thanassi, 2006).
Haemagglutination assay
Haemagglutination assays were performed by serial dilution
in microtitre plates as previously described (Shu Kin So and
Thanassi, 2006). HA titres were determined visually as the
highest-fold dilution of bacteria still able to agglutinate human
red blood cells (for P pili) or guinea pig red blood cells (Colo-
rado Serum Company; for type 1 pili). Each assay was per-
formed in triplicate. For analysis of the assembly of complete
P pili, AAEC185/pMJ2 (DpapC pap operon; papAHDJKEFG)
was used as the host strain. For analysis of assembly of P
pilus tips, AAEC185/pPAP58 (papDJKEFG) was used as the
host strain. Host strains transformed with pMON6235Dcat
(vector), pMJ3 (WT PapC with C-terminal His-tag), pM05
(PapC F3A), pTN5 (PapC C70A) or pTN26 (PapC D77–86)
were induced at OD
600
= 0.6 with 0.1% L-arabinose and
0.1 mM IPTG for 1 h. For analysis of assembly of complete
type 1 pili, MM294DfimD, which contains a fimD deletion in
the chromosomal fim operon (fimAICFGH), was used as the
host strain. MM294DfimD transformed with pMMB66 (vector),
pETS4 (WT FimD with C-terminal His-tag), pNH227 (FimD
F4A), pNH228 (FimD C63A) or pNH236 (FimD D70–79) was
grown statically for 24 h and then FimD expression was
induced for an additional 3 h by addition of 50 mM IPTG with
shaking at 100 r.p.m. For analysis of assembly of FimFGH or
FimGH type 1 pilus tips, AAEC185/pNH235 (fimCFGH)or
AAEC185/pNH222 (fimCGH) were used as the host strains
respectively. Host strains transformed with the FimD expres-
sion plasmids described above were induced at OD
600
= 0.6
with 50 mM IPTG and 0.1%
L-arabinose for 1 h.
Overlay assay
Overlay assays were performed as previously described
(Shu Kin So and Thanassi, 2006). OM fractions were isolated
as described above from strain SF100 harbouring the appro-
priate PapC (pMON6235Dcat, pMJ3, pM05, pTN5 or pTN26)
or FimD (pBAD18-cm, pETS7, pIT2, pIT4, pIT5 or pIT12)
expression plasmid. The strains were induced at OD
600
= 0.6
for 1 h with 0.1 or 0.02%
L-arabinose for PapC or FimD
respectively. The OM fractions were subjected to SDS-PAGE,
transferred to a PVDF (polyvinylidene difluoride; Osmonics)
membrane, and incubated with periplasm fractions contain-
ing PapDG (to test binding to PapC) or FimCH (to test bind-
ing to FimD). The periplasm fractions were isolated from
strain BL21/pJP1 (papDG) or ORN103/pETS1007 (fimCH)
as previously described (Ng et al., 2004). Binding of the
chaperone–adhesin complexes to the ushers on the PVDF
membrane was detected by immunoblotting with anti-PapDG
or anti-FimH antibodies. Immunoblots were developed with
alkaline phosphatase-conjugated secondary antibodies and
BCIP-NBT substrate.
Co-purification of pilus subunits with the usher
Co-purification assays were performed as previously
described (Shu Kin So and Thanassi, 2006). For analysis of
PapC, OM fractions were isolated as described above
from strain SF100/pPAP58 (papDJKEFG) harbouring
pMON6235Dcat, pMJ3, pM05, pTN5 or pTN26. The OM frac-
tions were solubilized with the nondenaturing detergent
dodecyl-maltopyranoside (DDM; Anatrace) and subjected to
nickel affinity chromatography. PapC was eluted from the
nickel column using an imidazole step gradient. Peak frac-
tions containing PapC were subjected to SDS-PAGE and
either stained with Coomassie blue to detect PapC or immu-
noblotted with anti-P pilus tips antiserum to detect pilus tip
Table 3. Primers used in this study.
Mutation Sequence of QuikChange forward primer (5–3)
a
PapC D77–86 CCGCAGGCCTGTCTGACATCAGATATGGTCGATAAAGTTG
FimD F4A CTGCCGACCTCTATGCTAATCCGCGC
FimD C63A GGGATTGTTCCCGCCCTGACACGCGCG
FimD D70–79 GCGCGCAACTCGCCGGTATGAATCTGC
a. The reverse primer is the reverse complement of the forward primer.
964
N. S. Henderson, T. W. Ng, I. Talukder and D. G. Thanassi
© 2010 Blackwell Publishing Ltd, Molecular Microbiology, 79, 954–967
Page 11
subunits that co-purified with the usher. To examine oligomer-
ization of PapG into P pilus tip fibres, the PapC-containing
fractions from the nickel column were incubated for 10 min at
room temperature (25°C) or 95°C in SDS-PAGE sample
buffer, subjected to SDS-PAGE and immunoblotted with anti-
PapDG antiserum. For analysis of FimD, OM fractions were
isolated as described above from strain AAEC185/pNH235
(fimCFGH) or AAEC185/pNH222 (fimCGH) harbouring
pMMB66, pETS4, pNH227, pNH228 or pNH236. The OM
fractions were solubilized and purified as for PapC, except
the affinity column was charged with cobalt instead of nickel.
The FimD-containing fractions were incubated for 10 min at
25°C or 95°C in SDS-PAGE sample buffer, subjected to SDS-
PAGE and immunoblotted with anti-FimCH, anti-FimCG or
anti-FimCF antibodies. Immunoblots were developed with
alkaline phosphatase-conjugated secondary antibodies and
BCIP-NBT substrate.
FimD trypsin sensitivity assay
Sensitivity of FimD to digestion by extracellularly added
trypsin was performed as previously described (Saulino
et al., 1998). Strains AAEC185/pETS7 (WT FimD with
C-terminal His-tag) or AAEC185/pETS1007 (fimCH) +
pETS7, pIT2 (FimD F4A) or pIT12 (FimD D70–79) were
induced at OD
600
= 0.6 with 50 mM IPTG and 0.02%
L-arabinose for 1 h. The cells were washed and resuspended
in 20 mM HEPES (pH 8.5) and equal portions were either
mock digested or digested with fresh 100 mgml
-1
trypsin
(Sigma) at 37°C for 2 h with rocking. Digestion was stopped
by addition of 0.1 mM phenylmethylsulphonyl fluoride, the
cells were harvesting by centrifugation, and OM fractions
were isolated as described above. The OM was subjected to
SDS-PAGE and immunoblotted with anti-FimD antibodies.
The blots were developed with alkaline phosphatase-
conjugated secondary antibodies and BCIP-NBT substrate.
In vitro reconstitution assay for pilus assembly
Outer membrane fractions were isolated from strain Tuner
harbouring pMMB91 (vector), pAN2 (WT FimD, no His-tag),
pNH238 (FimD F4A) or pNH239 (FimD D70–79) as described
above. FimD expression was induced at OD
600
= 0.6 with
50 mM IPTG for 1 h. Periplasm fractions were isolated as
previously described (Ng et al., 2004) from strain Tuner/
pNH221 (FimC) + pHJ20 (FimH) or strain Tuner/pETS1000
(FimC with C-terminal His-tag) + pETS2A (FimG). The strains
were induced at OD
600
= 0.6 with 0.1 mM IPTG and 0.05%
L-arabinose for 1 h. The periplasm fractions were dialysed
into buffer A [20 mM Tris-HCl (pH 8), 0.3 M NaCl] to match
the buffer used for the OM isolation. OM fractions [6.5 mg
protein; determined using the BCA assay (Pierce)] were
mixed with the FimCH and FimC
His
G periplasm fractions
(2.9 mg protein each) and the total volume adjusted to 10 ml
with buffer A. The samples were incubated at room tempera-
ture (25°C) and the reactions were stopped at the indicated
time points by addition of 50 ml cold buffer A and placed on
ice. The OM fractions were harvested by centrifugation
(100 000 g, 1 h, 4°C), resuspended in 10 ml buffer A and
solubilized by rocking overnight with 1% DDM at 4°C. The
OM extracts were centrifuged again to remove any unsolubi-
lized material and pilus assembly intermediates were purified
by cobalt affinity chromatography as described above for the
co-purification assay. Peak fractions from the affinity column
were precipitated by addition of 9% final concentration trichlo-
roacetic acid and incubation on ice for 30 min. The precipi-
tated proteins were harvested by centrifugation (10 000 g,
5 min, 4°C), washed twice with cold acetone, and resus-
pended in 150 ml SDS-PAGE sample buffer. The samples
were incubated in sample buffer for 10 min at room tempera-
ture, subjected to SDS-PAGE, transferred to PVDF mem-
brane and immunoblotted using anti-FimCG or anti-FimCH
antibodies. The blots were developed using infrared-
conjugated secondary antibodies (LI-COR Biosciences) and
analysed using the Odyssey Infrared Imaging System (LI-
COR Biosciences). Quantification of FimG
n
–FimH complexes
formed at each time point was done by measuring the signal
intensity within identical areas boxed as shown for the 45 min
time point in Fig. 4A. For each experiment, the intensity value
obtained for the 0 min time point was subtracted from the
values obtained for the subsequent time points.
Acknowledgements
We thank Huilin Li (Brookhaven National Laboratory and
Stony Brook University) and Wali Karzi (Stony Brook Univer-
sity) for critical reading of the manuscript. This work was
supported by National Institutes of Health grant GM062987.
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Catalytic activity of the usher
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  • Source
    • "HA assays were performed to test the ability of each of the PapC substitution mutants to assemble functional P pili on the bacterial surface. HA assays were performed by serial dilution in microtiter plates as previously described (Henderson et al., 2011). HA titers were determined visually as the highest fold dilution of bacteria still able to agglutinate human red blood cells. "
    [Show abstract] [Hide abstract] ABSTRACT: eLife digest Escherichia coli is a bacterium that commonly lives in the intestines of mammals, including humans, where it is usually harmless and can even be beneficial to its host. However, some types of E. coli produce hair-like filaments called P pili that allow the bacteria to attach to the human urinary tract and cause disease. To pass through the outer membrane of the E. coli cell, the filaments have to travel through a protein in the membrane called PapC usher. The PapC usher protein—which is also involved in the assembly of the P pili filaments—contains a tube-like part called a β-barrel that is usually blocked by another part of the protein called the ‘plug domain’. For the P pili to pass through the β-barrel, the plug domain has to move. This movement is controlled by two parts of the PapC protein, known as the α-helix and the β-hairpin, but it is not clear how. To address this question, Farabella et al. made computer models of the normal PapC protein and versions that lacked the α-helix and/or the β-hairpin. Looking at these structural models and analyzing the evolution of PapC proteins helped to predict that certain regions of the β-barrel may be involved in controlling the movement of the plug domain, and this was then confirmed experimentally. Farabella et al. propose that these regions—together with the α-helix and β-hairpin—control the opening and closing of the β-barrel. Further work is needed to investigate how other parts of the PapC protein are involved in P pili formation. These new insights could prove useful in the development of alternative treatments to fight bacterial infection. DOI: http://dx.doi.org/10.7554/eLife.03532.002
    Full-text · Article · Oct 2014 · eLife Sciences
  • Source
    • "–subunit interactions in the pilus are resistant to dissociation by SDS unless heated (Henderson et al., 2011). This mechanical strength is critical for the ability of the pili to maintain adhesion in the face of external forces such as the shear force encountered in the bladder from the flow of urine. "
    [Show abstract] [Hide abstract] ABSTRACT: Gram-negative bacteria express a wide variety of organelles on their cell surface. These surface structures may be the end products of secretion systems, such as the hair-like fibers assembled by the chaperone/usher (CU) and type IV pilus pathways, which generally function in adhesion to surfaces and bacterial-bacterial and bacterial-host interactions. Alternatively, the surface organelles may be integral components of the secretion machinery itself, such as the needle complex and pilus extensions formed by the type III and type IV secretion systems, which function in the delivery of bacterial effectors inside host cells. Bacterial surface structures perform functions critical for pathogenesis and have evolved to withstand forces exerted by the external environment and cope with defenses mounted by the host immune system. Given their essential roles in pathogenesis and exposed nature, bacterial surface structures also make attractive targets for therapeutic intervention. This review will describe the structure and function of surface organelles assembled by four different Gram-negative bacterial secretion systems: the CU pathway, the type IV pilus pathway, and the type III and type IV secretion systems.
    Preview · Article · Apr 2012 · FEMS microbiology reviews
  • Source
    • "Ushers are located at the outer membrane of Gram-negative bacteria. These are ~ 90kDa proteins consisting of five distinct functional domains: a N-terminal periplasmic domain (UND) (Henderson et al., 2011; Eidam et al., 2008; Nishiyama et al., 2005; Nishiyama et al., 2003), a trans-membrane domain (TMD) (Remaut et al., 2008), a middle domain (UMD) (Huang et al., 2009; Yu et al., 2009; Remaut et al., 2008), and two C-terminal periplasmic domains (UCD1 and UCD2) (Phan et al., 2011; Dubnovitsky et al., 2010; Ford et al., 2010) (Fig. 1). "
    Preview · Article · Jan 2012
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