From the Department of Biochemistry & Molecular Biology, University of Oklahoma Health Sciences Center, Oklahoma City, Oklahoma 73190
Received for publication, March 17, 2000, and in revised form, October 4, 2000
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ABSTRACT |
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Since we first reported (DeAngelis, P. L.,
Papaconstantinou, J., and Weigel, P. H. (1993) J. Biol.
Chem. 268, 19181-19184) the cloning of the hyaluronan (HA)
synthase from Streptococcus pyogenes (spHAS), numerous
membrane-bound HA synthases have been discovered in both prokaryotes
and eukaryotes. The HASs are unique among enzymes studied to date
because they mediate 6-7 discrete functions in order to assemble a
polysaccharide containing hetero-disaccharide units and simultaneously
effect translocation of the growing HA chain through the plasma
membrane. To understand how the relatively small spHAS performs these
various functions, we investigated the topological organization of the
protein utilizing fusion analysis with two reporter enzymes, alkaline
phosphatase and HA1 is a linear
polysaccharide that contains repeating disaccharide units of
GlcUA With the exception of pmHAS, all of the HA synthases share regions of
significant homology at the amino acid level and extensive similarity
in their hydropathy profiles (25, 26). Based on these similarities the
HASs have been grouped into two classes (26). Class I HASs include all
the eukaryotic and streptococcal enzymes. The pmHAS is presently the
only Class II member. The two streptococcal enzymes, spHAS and seHAS,
are the smallest members within the HAS family containing,
respectively, 419 and 417 amino acids. These two proteins are 72%
identical at the amino acid level. The streptococcal enzymes have also
been purified to homogeneity and are the most extensively characterized
HASs biochemically (27-29). These two HASs have been characterized
kinetically in membranes or as purified enzymes with respect to the
Km values for the two substrate sugars,
Vmax, and average product length. Tlapak-Simmons
et al. (27) recently demonstrated that spHAS and seHAS
function as monomeric proteins dependent on the association of ~16
cardiolipin (CL) molecules. A single HAS protein appears to be solely
responsible for HA chain initiation and elongation and probably also
for extrusion of the growing HA chain from the cell.
The sizes of the HAS enzymes are similar to that of other transporters
such as the Lac permease (30), mannose transporter IIC (31), the porins
from Eschericia coli and Rhodobacter capsulatus (32, 33), the facilitative glucose transporter (34), and the melibiose
carrier protein (35). Unlike these transporters, however, spHAS and
seHAS, as well as the other HAS family members, lack the number of
predicted hydrophobic transmembrane domains necessary for the formation
of a pore sufficiently large to transport the growing HA chain across
the membrane. To understand the enzymatic and transport functions of
the HAS family, it is important to determine the topological
organization of the active site(s) and membrane-spanning or
membrane-associated regions.
Previously (11, 36) we predicted a model for spHAS organization in
which the ends of the protein and a large central domain were inside
the cell (e.g. similar to Fig. 3E). The
eukaryotic HASs, which are ~35% larger than the streptococcal
enzymes, were proposed to have this same topological organization with
an additional COOH-terminal region containing several MDs (25). Since
the streptococcal HASs are structurally the smallest, simplest
members of this family, information on the topology of the spHAS and
seHAS enzymes could help elucidate the basic topology of the remaining HAS family members.
In this study, the topology of spHAS was characterized by protease and
substrate accessibility and by fusion protein analysis using alkaline
phosphatase and Materials and Strains--
The starting vector for
expression of PCR-mediated fusion proteins was pAlter Ex-1 from
Promega. The vector pKK223-3 was from Amersham Pharmacia Biotech. The
vector pSK Construction of pKK3K--
Custom oligonucleotides,
5'-CCCGGGAATTCNTGCCT-ATTTTTAAAAAAACTTTAATTG (forward primer) and
5'-AGAATTCTGCAGTTAATGGTGATGGTGGTGGTGTTTAAAAATAGTGAC (reverse
primer, containing a His6 sequence), were used to amplify the coding sequence of spHAS from Streptococcus pyogenes
chromosomal DNA by PCR with Taq polymerase (Fisher). The
forward primer contains an N to allow for either a Met or Val as the
NH2-terminal amino acid; the primer is suitable for either
codon, although the KK3K vector used contains the spHAS ORF that begins
with a Val as occurs in the wild type enzyme. The PCR product and the
plasmid pKK223-3 were gel purified using GeneClean (Bio 101) after
digestion with EcoRI and PstI. The products were
ligated overnight with T4 DNA ligase. The resulting plasmid, designated
pKK3K, was sequenced to confirm the coding sequence and the presence of
the COOH-terminal His6 tag. Cells were transformed and
maintained on LB containing 50 µg/ml ampicillin at 30 °C.
Construction of AP-His6-1--
To create
alkaline phosphatase containing a COOH-terminal His6 tag,
the AP gene in pSK PCR-mediated HAS Fusion Proteins--
Fig.
1 shows schematically how the spHAS-AP
and spHAS-LacZ fusion protein vectors were constructed. Primers were
designed to amplify different lengths of the spHAS ORF.
HA5'(5'-CGGGATCCGTGCCTATTTTTAAAAAAACT-3') was used for PCR with
Pfu polymerase and position Thr
(5'-AGTTTTTTTAAAAATAGGCACGGATCCCG-3'), Asn
(5'-ATTTCCTTTAAATGGCTCAT-3'), or Pro
(5'-GGGATTAGAAAGAATTTTTTT-3') from a pKK3K template. PCR products
were ligated to KpnI linker from New England Biolabs.
Ligation reactions and pAP6X-1 were digested with BamHI and
KpnI and ligated together. Pfu PCR was also used
to generate spHAS fragments with a restriction site at the 3' end of
the products to facilitate insertion into the vector. For these PCR
reactions, HA5' was used with the reverse primers:
Gly37 (5'-GGGTACCCCATAAATTCCTACAGTTGATG-3'),
Val134 (5'-GGGGTACCCCGACAAGGGAACGGTGAACGATAACGTTTCGAC-3'),
Gly270 (5'-GGGGTACCCTCCTAAATCAATAGCATAATTTG-3'), Asp349
(5'-GGGGTACCCCGTCTAATTGAATAGCTTGAT-3'), Pro376
(5'-GGGTACCCCAGGATGTTTGACCATATAAT-3'), and Lys419
(5'-GGGGATCCCCTTTAAAAATAGTGACCTTTT). The PCR products were
digested with BamHI and KpnI and ligated into
pAP6X-1. All ligations were transformed into DHB9 cells and screened by
colony PCR using primers HA5' and PhoA. Those colonies giving the
expected band size were grown in overnight culture and plasmids were
purified. Plasmids were screened by restriction digest then sequenced
with the PhoA primer to confirm location and frame of fusions.
Transfer of Transposon Fusion Genes to the pAlter Ex-1
Vector--
The ORFs for the transposon-created fusion proteins were
PCR-amplified with primers HA5' and APHIS63'. PCR products were
digested with BamHI and XbaI and ligated into the
pAlter Ex-1 vector. Clones were isolated and assessed for correct
construction by sequencing through the junction site.
LacZ Fusion Construction--
Primers LacZ-5'
(5'-CCCCGGGTACCTGACTCTTATACACAAAGTA-3') and
LacZH6-3'(5'-CTAGTCTAGATTAATGATGATGATGATGATGTTTTTGACACCAGACCAA-3') were used to amplify the AP and LacZ Fusion Protein Assays--
Clones with confirmed
fusion proteins were grown overnight in LB/Amp. The overnight cultures
were diluted 1:25 to start new cultures in LB/Amp grown at 30 °C
with aeration for 3-4 h. Cultures were induced with
isopropyl-1-thio- Assessment of Fusion Protein Localization--
Based on prior
published cutoff values used by numerous other investigators (31,
44-50), the various fusion proteins were divided into categories of
high (>20% of maximum), intermediate (10-20% of maximum), or low
(<10% of maximum) activity, relative to the highest activity for each
assay method used (defined as 100%). The average activity of each
fusion protein, using the same assay method was compared as a
percentage to the fusion protein construct with the highest activity.
For example, for the spHAS-AP fusion proteins, all the plate-based
assays were evaluated as a percent of the Asp349-AP
fusion, which gave the highest specific activity in this assay. For AP
or LacZ fusion proteins with low activity, the reporter protein was
assessed as inside or outside of the cell, respectively. For AP or LacZ
fusion proteins with high activity, the reporter protein was assessed
as outside or inside of the cell, respectively. If fusion protein pairs
(i.e. both AP and LacZ fusion proteins at the same spHAS
position) had intermediate activities, neither was assigned an
orientation because of this ambiguity or the fusion site was considered
to be part of a possible MD.
Western Blots--
Samples (5-10 µl) in SDS-PAGE buffer were
loaded onto polyacrylamide gels and electrophoresed at 200 V for 50 min. Proteins were transferred to nitrocellulose at 100 V for 2 h
in a Bio-Rad Trans-Blot Cell. Blots were stained with 0.05%
copper(II)phthalocyanine-3,4',4'',4'''-tetrasulfonic acid, tetrasodium
salt (Aldrich) and images of the blots were captured with an
Alpha-Innotech IS1000 imaging system. Blots were destained and blocked
with 1% bovine serum albumin in TBST (with 0.05% NaN3)
for Accessibility to Proteases and Chemical Labeling
Reagents--
To assess which portions of spHAS reside in the
periplasm, spheroplasts of cells expressing spHAS-H6
(i.e. the full-length protein; not a fusion protein) were
subjected to several modifying agents. Overnight cultures were seeded
1:15 into 13 ml of LB/ampicillin and grown at 30 °C, 250 rpm until
late log phase. Cells were induced for 3 h with 1.5 mM
isopropyl-1-thio- Spheroplast Preparation and Assessment of Active Site
Accessibility--
Cells, spheroplasts, and cell lysates
expressing full-length spHAS-His6
(i.e. not as a fusion protein) were prepared as
described above to assess the accessibility of substrates to the active site of spHAS for HA synthesis. Aliquots from each preparation were
incubated with 1.5 mM UDP-GlcNAc, 1 mM
UDP-[14C]GlcUA (2.5 µCi) for 1 h at 30 °C with
gentle mixing using a Micromixer (Taitec Institute USA, Inc., San Jose,
CA). Reactions were stopped with 2% SDS. Substrate incorporation into
HA was assessed by descending paper chromatography (11).
General--
Protein content was determined by the method of
Bradford (52) using bovine serum albumin as the standard. SDS-PAGE was
performed according to the method of Laemmli (41). DNA sequencing was performed by the OUHSC core facility in the Department of Microbiology and Immunology.
The shortest -galactosidase, as well as several other approaches.
From these studies, we conclude that the NH2 terminus and
the COOH terminus, as well as the major portion of a large central
domain are localized intracellularly. The first two predicted membrane
domains were confirmed to be transmembrane domains and give rise to a
very small extracellular loop that is inaccessible to proteases.
Several regions of the large internal central domain appear to be
associated with, but do not traverse, the membrane. Following the
central domain, there are two additional transmembrane domains
connected by a second small extracellular loop that also is
inaccessible to proteases. The COOH-terminal ~25% of spHAS also
contains a membrane domain that does not traverse the membrane and may
contain extensive re-entrant loops or amphipathic helices. Numerous
membrane associations of this latter COOH-terminal region and the
central domain may be required to create a pore-like structure through
which a growing HA chain can be extruded to the cell exterior. Based on
the high degree of similarity among Class I HAS family members, these
enzymes may have a similar topological organization for their
spHAS-related domains.
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
(1,3)GlcNAc
(1,4) and is widely distributed throughout the
animal kingdom (1). In eukaryotes HA serves a viscoelastic structural
role and functions, together with proteoglycans, as part of the
extracellular matrix scaffold that creates tissue integrity (2, 3). HA
also mediates the localization and movement of cells (3-5), and can
alter or regulate cell behavior by acting as a signaling molecule (6,
7). HA in prokaryotes is found as a capsule that serves as a virulence
factor, presumably by mimicking the eukaryotic host and thus being
immuno-protective (8-10). The enzyme responsible for HA synthesis, HA
synthase, was cloned originally from Group A streptococcus (11).
Structurally related Class I HA synthases were then cloned from human
(12-14), mouse (15-17), chicken (17), frog (17-21), Group C
streptococcus (22), and chlorella virus PBCV-1 (23). A structurally
distinct Pasteurella multocida HA synthase was also cloned
(24). All the HA synthases reported to date produce a broad
distribution of high molecular weight (e.g.
~105-107) HA chains from the substrates,
UDP-GlcNAc and UDP-GlcUA.
-galactosidase as reporter enzymes, whose correct
folding and activity is dependent on cellular localization. The results
support a topology of HAS that is similar to the model proposed
earlier, but that entails more potentially complex interactions with
the membrane and delineates the membrane-associated regions near the
COOH terminus.
EXPERIMENTAL PROCEDURES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
LphoA (37) was graciously provided by Marleen van Geest
and Juke Lolkema. Restriction and DNA modifying enzymes were from
Promega unless noted otherwise. Oligonucleotide primers were
synthesized by The Great American Gene Co. or Genosys. DNA plasmid
purification was done with Qiaprep Spin Miniprep Kits from Qiagen. T7
Sequenase version 2.0 sequencing kit and 35S-dATP were from
Amersham Pharmacia Biotech. Media were from Fisher and the remaining
reagents were of the highest purity available from Sigma unless
otherwise noted. E. coli strain DHB9 was provided courtesy
of The Beckwith Collection. E. coli CC170 cells containing TnLacZ (38) were provided by Dr. Colin Manoil. E. coli JM109 was used for LacZ fusion assays. CC118 for LacZ fusion assessment was
provided courtesy of Dr. Colin Manoil.
TnPhoA provided by Manoil and
Beckwith (39) was used for infection of DHB9 cells and integration of
the AP coding region into pKK3K. Antibody to TolA (40) was a generous
gift from Dr. Robert Webster.
TnPhoA-mediated spHAS Fusion--
DHB9 cells containing pKK3K
were grown in LB containing 50 µg/ml ampicillin, 10 mM
MgSO4 with aeration at 30 °C to an
A600 of ~1.0.
TnPhoA was added to a density of
1.4 × 109 plaque forming units/ml of culture and the
cells were incubated without aeration at 30 °C for 20 min. Aliquots
of the culture (100 µl) were diluted into 1 ml of LB medium and
incubated at 30 °C for ~12 h with aeration. Portions (200 µl) of
these cultures were overlaid onto LB plates containing 50 µg/ml
ampicillin, 100 µg/ml kanamycin, and incubated at 30 °C for 2-3
days until colonies were easily visible. The colonies were scraped into
a 1.7-ml tube with 500 µl of phosphate-buffered saline added
to each plate. Plasmid DNA from pooled colonies was isolated with
Qiaprep spin columns (Qiagen) and used to transform DHB9 competent
cells. This step eliminates selection of Tn integration events into the
chromosome. Transformed cells were grown on LB/kanamycin plates.
Colonies were picked and seeded on gridded LB/Amp plates with 40 µg/ml bromochloroindolyl phosphate, and 1 mM
isopropyl-1-thio-
-D-galactopyranoside. After overnight
growth, colonies from the gridded plate were screened by PCR to
identify clones with the Tn integrated into the spHAS ORF. The forward
primer was the same as that used in the synthesis of pKK3K and the
reverse primer was AATATCGCCCTGAGCA, which corresponds to a 5' region
of AP. PCR products were checked by electrophoresis on 0.8% agarose
gels and those colonies with PCR bands less than 1.3 kilobases in
length, the size of the intact coding region, were streaked onto
LB/Kan/Amp plates. Colonies were picked and seeded into LB medium with
ampicillin, incubated overnight, and the cultures used to prepare
frozen glycerol stocks and plasmid DNA. The DNA was digested with
EcoRI to confirm the orientation of the fusion. Colonies
with properly oriented and sized inserts encoding fusion proteins were
grown in 0.5 ml of LB medium with ampicillin and induced with
isopropyl-1-thio-
-D-galactopyranoside at a final
concentration of 1 mM for 3 h at 30 °C. Cells were pelleted and resuspended in 40 µl of 1× Laemmli buffer (41) with 10 mM dithiothreitol; 10 µl was used for Western blot
analysis with antibodies specific for spHAS or AP. Only colonies with a band at the expected size recognized by both anti-AP and anti-spHAS antibodies were sequenced, using primer PhoA (5'-AATATCGCCCTGAGCA-3'), to confirm the site of the fusion.
LphoA was amplified by 22 cycles of PCR with
primer AP5' (5'-AGGTCGACCATGGAGGATCCC-3') and primer APHIS6-3'
(5'-CTAGTCTAGATTAGTGGTGGTGGTGGTGGTGTTTCAGCCCCAGAGCG-3') using
Pfu polymerase (Stratagene). The PCR product and pAlter Ex-1
were digested with BamHI and XbaI and separated
from the primers by a Qiagen PCR purification column. The PCR product
was ligated to the pAlter Ex-1 restriction digest product with T4 DNA
ligase (New England Biolabs) and transformed into DHB9 competent cells.
Colonies were screened by plasmid preparation and restriction digestion
for the proper size and orientation of insert. The desired plasmid was
designated pAP6X-1.
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Fig. 1.
Schemes for the preparation of PCR-generated
spHAS fusion proteins with the recorder enzymes AP or LacZ.
A, the alkaline phosphatase ORF from pSK LphoA
(light gray box) was amplified, using Pfu
polymerase, to contain a COOH-terminal six histidine tail
(gray-to-white box). The product was purified and ligated
into the pAlter Ex-1 vector (Promega) containing an ampicillin
resistance gene (white box). The new vector was designated
pAP6X-1. Selected regions of the spHAS ORF (black boxes)
were similarly amplified to contain 5' BamHI and 3'
KpnI restriction sites. pAP6X-1 and the spHAS fragments were
digested with BamHI/KpnI (except for construction
of the full-length K419 construct for which digestion was only with
BamHI) and ligated together to form spHAS-AP6 fusion
constructs designated by the position number and amino acid at the
COOH-terminal end of the spHAS fragment (e.g. K419AP6). The
desired fragments from all restriction digests were gel-purified prior
to ligation. All AP6 constructs were sequenced through the entire ORF
to confirm reading frame and sequence fidelity. B, the ORF
(dark gray box) of
-galactosidase (LacZ) from TnLacZ was
amplified with Pfu polymerase using primers containing a 5'
KpnI and a 3' XbaI site. The K419AP6 fusion
construct was digested with KpnI/XbaI and the
LacZ ORF was ligated in the former AP6 position. The resulting K419LacZ
construct was then digested with BamHI/KpnI and
the similarly digested spHAS fragments from the spHASAP6 constructs
were ligated into the LacZ-containing plasmid. All constructs were
sequenced through the LacZ junction to ensure reading frame and
sequence fidelity.
-galactosidase ORF from strain CC170 with Pfu polymerase as above. This product and the K419AP6
construct were digested with KpnI and XbaI, gel
purified, and the LacZ fragment was ligated into the
Lys419 construct. The spHAS PCR
amplifications used for the AP6 fusion proteins described above were
also ligated into the Lys419 LacZ construct after digestion
with BamHI and KpnI to remove the Lys419
insert. Fusions were confirmed by sequencing the spHAS insert through
the junction site. LacZ fusions were grown in JM109 cells.
-D-galactopyranoside for 3 h at
30 °C. To determine the relative activity of the various fusion
proteins, we measured cell number (by A600), the
amount of fusion protein (by quantitative Western analysis), and enzyme activity (by colorimetric assays). One ml of cells from each culture was transferred to a 1.7-ml microfuge tube and centrifuged at 10,000 × g for 5 min. Cell pellets were resuspended in
1 ml of ice-cold 1 M Tris-HCl, pH 8, or phosphate-buffered
saline containing 1 mM MgSO4 and 50 mM
-mercaptoethanol. A portion (100 µl) of the
suspension was diluted into 0.9 ml of cold 1 M Tris-HCl, pH 8, or phosphate-buffered saline and A600 was
determined. A second 100 µl of the suspension was centrifuged at
10,000 × g for 5 min, the pellet was resuspended in
100 µl of 1 × Laemmli SDS-PAGE buffer with 10 mM
dithiothreitol, heated at 95 °C for 2 min, and used for Western
blots. Another 100 µl of the suspension was diluted into 0.9 ml of 1 M Tris, 0.1 mM ZnCl2, or
phosphate-buffered saline containing 1 mM MgSO4
and 50 mM
-mercaptoethanol. Fifty µl of 0.1% SDS and
50 µl of chloroform were then added and the tube was vortexed and
incubated at 37 °C for 5 min. At time 0, 100 µl of 0.4%
p-nitrophenyl phosphate in 1 M Tris or 8 mg/ml
o-nitrophenyl-
-galactoside was added to each reaction
tube. When a yellow reaction color was observed, 120 µl of 83.3 mM EDTA, 83 mM KH2PO4
was added to stop the AP assays, or 0.5 ml of 1 M
Na2CO3 was added to stop the Lac assays. The
time was noted, the tube was transferred to ice and
A550 and A420 were obtained.
Activities were determined using the formula:
1000*[A420
(1.75 × A550)]
(time in min × nmol construct) as
described by others (42, 43). Both the LacZ and AP assays were also
done in a multiwell plate reader format with 20 and 40 µl of induced
resuspension and 20 µl of substrate in a total volume of 200 µl/well. Plate reader assays were scanned every 30 s over 1 h with an enzyme-linked immunosorbent assay plate reader (Spectramax
340, Molecular Devices). Activities were determined over the linear
range of substrate conversion.
1 h. Blots were washed in TBST and exposed to a 1:1000 dilution
of anti-penta-His monoclonal Ab from Qiagen or anti-
-galactosidase
monoclonal Ab from Calbiochem, in TBST containing 0.1% bovine serum
albumin for 1 h. Antibodies were biotinylated with NHS-biotin from
Pierce. Secondary development was for 1 h with
125I-streptavidin and the blots were analyzed using a
phosphoscreen on a Molecular Dynamics Personal PhosphorImager.
Expression levels were measured as integrated density values and
converted to nanomole of fusion protein using an internal standard
curve of purified spHAS-H6. This was followed by detection
with rabbit anti-mouse IgG Ab conjugated to alkaline phosphatase
(Sigma). Blots were developed with 0.3 mg/ml p-nitro blue
tetrazolium chloride and 0.15 mg/ml 5-bromo-4-chloro-3-indoyl phosphate
p-toluidine salt in 1 M Tris-HCl, pH 9.5, 0.5 mM MgCl2.
-D-galactopyranoside. Cells were
collected by centrifugation, resuspended in 13 ml of 20 mM Tris, pH 8.5, 20% sucrose, and 0.5 ml of suspension was removed as a
"no treatment" control. EDTA was added to 2 mM and
lysozyme was added to 10 µg/ml. Cells were incubated on ice for 20 min and >90% cell wall removal was observed by light microscopy
verifying the production of spheroplasts (51). The MgSO4
concentration was brought to 15 mM and a lysate was
prepared by sonication of a 6-ml portion of the spheroplast suspension,
using a microtip probe at 20 watts (W-380; Heat Systems Ultrasonic,
Inc., Farmingdale, NY), until the suspensions appeared clear. Aliquots
(1 ml) of spheroplasts and cell lysates were treated with either 0.2 mg/ml chymotrypsin, 0.2 mg/ml trypsin, 0.2 mg/ml Pronase, 0.2 mg/ml proteinase K, 9 mg/ml NHS-Biotin, or 3 mg/ml
(+)-biotinyl-3-maleimidopropionamidyl-3,6-dioxaoctanediamine for 30 min
on ice. Proteases were inhibited with 50 µg/ml soybean trypsin
inhibitor or 3 mM phenylmethylsulfonyl fluoride.
Spheroplasts were separated from modifying reagents, broken cells, cell
walls, and outer membrane fragments by centrifugation at 10,000 × g for 1 min through 1 ml of 20 mM Tris, pH 8.5, 20% sucrose, 15 mM MgSO4. Spheroplasts were
resuspended in 0.5 ml of 20 mM Tris, pH 8.5, 20% sucrose,
and 15 mM MgSO4 and 75 µl from each sample
was added to 25 µl of 4 × SDS-PAGE Laemmli buffer (41) with 0.5 µg/ml leupeptin and 0.7 µg/ml pepstatin and the mixture was then
incubated at 95 °C for 2 min.
RESULTS
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
-helical domain that could theoretically span a
membrane lipid bilayer would be ~20 residues and the shortest
-sheet domain would be ~11 residues. The majority of membrane proteins that have been crystallized and whose structures have been
successfully solved contain
-helical membrane-spanning regions (53).
Because
-sheets were not found as MDs in earlier studies, it was
posited that they were energetically unfavorable. Thus, most current
software programs for predicting MDs have a concerted bias toward
finding
-helices in membrane regions. In recent years, however,
-sheet transmembrane regions have been demonstrated, usually with a
-barrel type of structure (54). Therefore, in determining the
topology of a protein for the first time, it is important to consider
the possibility that both
-helical and
-sheet MD structures may
be present. Hydropathy analysis (53) of spHAS reveals at least six
hydrophobic stretches that could represent PMDs, predicted (or
putative) membrane domains (Fig. 2).
Based on the three prediction algorithms of Garnier et al. (55), Chou and Fasman (56, 57), and Rose (58) as well as the two
prediction programs TMPRED (59) and HMMTOP (60), a large number of
topologies are possible for spHAS (Fig.
3). These models vary as to which PMD is
predicted to be a MD or a TMD and the orientation of a given PMD.
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Fig. 2.
Hydropathy plot and PMDs of spHAS. The
Kyte and Doolittle (45) plot, using a window of 17 amino acids,
indicates the hydrophobic regions of spHAS. The hydrophobic regions
long enough to span a membrane bilayer are indicated by
boxes labeled PMDs 1-6 in the linear schematic
below the graph. These possible MDs are at residues 7-29
(PMD1), 33-54 (PMD2), 207-232 (PMD3), 317-343 (PMD4), 350-370
(PMD5), and 379-396 (PMD6).
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Fig. 3.
Alternative topological models for
spHAS. Six possible topological models are shown
(A-F), based on hydropathy, as in Fig. 2, using a
"scanning" window from 7 to 20 amino acids. The numbering system
for PMDs is that shown in Fig. 2. In some analyses PMD 5 and 6 are
essentially continuous. Other topological models are also
possible.
To determine which, if any, of the various predicted topology models summarized in Fig. 3 might be correct, topological studies were undertaken using a combination of fusion protein analysis (31, 42, 44-50, 61), chemical labeling, protease accessibility, and substrate accessibility. Using Tn-mediated and PCR-mediated approaches, protein fusions were created with varied lengths of spHAS linked to the reporter enzymes AP or LacZ. AP is only folded correctly and active when it is in the nonreducing environment outside the cell, whereas LacZ, in contrast, is only active when it is made within the intracellular reducing environment (31, 37, 42, 44-50, 61). As the spHAS fusion protein inserts into the membrane, the reporter enzyme is localized according to the NH2-terminal topolological signals (62-64). The reporter enzyme is then active or inactive depending on the local reducing environment in which the AP or LacZ is localized.
Unfortunately, Tn-generated spHAS-AP fusion proteins were not random; they were distributed in the ORF predominately around residue Asp349 and were not adequately dispersed throughout the enzyme sequence. Therefore, specific PCR-generated fusion proteins were created on either side of the six PMDs. This fusion protein approach has been used successfully to determine the topology of a large number of bacterial proteins (61) and has been shown to be accurate when the results could be compared with a crystal structure, such as for the photosynthetic reaction center of Rhodopseudomonas sphaeroides (65).
Analysis of Fusion Protein Constructs-- AP fusion proteins were generated in 17 locations within spHAS (Table I) through a combination of Tn-mediated and PCR-mediated approaches (38). LacZ fusion proteins were also generated by PCR at nine of these same 17 positions. The constructs were checked for expression of fusion proteins by Western blot analysis and the sequence of the fusion junctions were confirmed by DNA sequencing. The spHAS fusion proteins generated with AP or LacZ had a broad range of expression levels, as determined by Western blot analysis, and varied greatly in the degree to which breakdown products were observed (Fig. 4A). The intact fusion proteins demonstrated an electrophoretic mobility shift proportional to the length of the spHAS portion incorporated (Fig. 4A). Fusion protein expression levels, determined by Western blot analysis with an anti-His5 Ab or anti-LacZ Ab and 125I-streptavidin (Fig. 4B), varied over nearly 2 orders of magnitude. For spHAS-AP fusions, the expression level generally varied inversely with the length of the fusion protein (e.g. fusion constructs at Gly37 and Asn60 had 4-fold greater expression levels compared with any other constructs). The only exception to this was for the fusion at position Thr7, which had an extremely low expression level for the AP fusion. Based on SDS-PAGE and Western analysis comparing isolated membranes and whole cell lysates, at least 95% of the various fusion proteins were localized, as expected, to the cell membrane (not shown).
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In a large number of studies, the location of reporter enzyme in AP or LacZ fusion proteins has typically been judged to be intracellular or extracellular based on about a 5-10-fold difference in activity (31, 44-50). For spHAS-AP fusion proteins, an external orientation was scored as a high activity above a cutoff of 6 units/min/nM for plate reader assays and 13 units/min/nM for spectrophotometric assays. For spHAS-LacZ fusion proteins, an intracellular localization was assessed based on high activities of >25 units/min/% position Thr7 expression.
Activity assays for fusion proteins performed by the plate reader method have the benefit of needing no correction in absorbance measurements for interference from cell debris. More importantly, the plate reader assays enable determination of the linearity of the assay over multiple time points. The bacterial AP activities of fusion proteins were assessed by both the plate assay and the traditional single time point spectrophotometric assay. Results were consistent between the two assay methods, with the exception of fusion proteins produced at positions Glu169 and Asn207, for which the two assays gave conflicting results (Table I). The spectrophotometric assay usually gave higher AP specific activities. Assessments of AP cellular localization were then made as either inside, outside, or unclear, based on the activity category for each fusion protein.
The HAS-AP fusion protein at position Asp349 demonstrated a high AP activity and a low LacZ activity, indicating a periplasmic location for that region of spHAS. High-to-intermediate AP activity with low LacZ activity was also observed for fusion proteins at Gly37 and Asn60. These were complemented by fusion proteins at position Thr7, Pro317, and Lys419, which showed a high activity for the LacZ fusion and a low-to-intermediate activity for the AP fusion, indicating intracellular locations for these sites. Thus, we conclude that the NH2 and COOH termini of spHAS, as well as the region immediately prior to PMD4, are intracellular and that the two loops between PMD1 and PMD2 and between PMD4 and PMD5 are extracellular. In E. coli these two loops are exposed to the periplasm.
Paradoxically, high activities for both spHAS-AP and spHAS-LacZ fusion proteins were observed at Val134 and Pro376. In these cases, the stronger positive activity for LacZ was taken to be the better indicator of membrane orientation and they were assigned as internal. One set of spHAS fusion proteins at position Gly270 demonstrated intermediate-to-low activities for both AP and LacZ constructs and thus no orientation was indicated. The remaining fusion locations were examined using AP fusion proteins only and thus, the lack of strong AP activities is not complemented with LacZ activities to give both positive and negative indications of membrane orientation. Fusion locations at Ala208, Ala215, Leu353, and Lys397 had high AP activities, indicating an external localization. The fusion at Leu365 had low activity indicating an internal orientation of the AP at this HAS position. The fusions at Thr32, Glu169, and Asn207 give conflicting results between the plate reader and the spectrophotometric assays and were not assigned a membrane orientation.
Sensitivity of spHAS to Proteases and Chemical Labeling--
To
assess the presence and accessibility of any portions of spHAS that
might be localized to the periplasm, spheroplasts, either intact or
disrupted by sonication, were exposed to a chemical modifying reagents
or a variety of proteases. In these experiments the full-length spHAS
protein, not a truncated or fusion protein, was examined. As expected,
all the sonicated samples showed multiple sites of accessibility to the
proteases and to a biotinylating reagent reactive with primary amines
(Fig. 5), demonstrating that under the
conditions used, spHAS was susceptible to proteolytic and chemical
modifications. However, intact spheroplasts showed no demonstrable
cleavage or modification of spHAS, indicating that the protein has no
extracellular regions sufficiently exposed to allow modification. As a
positive control, the same digestion conditions were shown to cleave
the TolA protein completely in both sphereoplasts and lysates (not
shown). TolA, which is involved in colicin uptake, is localized to the
inner membrane (40). Thus, it is very likely that any extra-cytoplasmic
loops of the spHAS protein are very small and close to or intimately
associated with the membrane.
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Accessibility of the spHAS Active Site to Substrates-- In order to determine whether the sugar nucleotide substrates are accessible to the active site of spHAS from an extracellular, periplasmic, or intracellular location, we incubated intact cells, intact spheroplasts, or sonicated spheroplasts with these substrates. Only the lysed samples demonstrated the expected formation of HA (Table II), indicating that the two substrate-binding sites and glycosyltransferase active sites are only accessible from an intracellular location. The protein regions within the HAS family that are most highly conserved, and are thus most likely to be involved in substrate binding and catalysis, are located between PMD2 and PMD4 (25, 26). The conclusion from the above substrate accessibility result is that this region is intracellular.
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DISCUSSION |
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Stoolmiller and Dorfman (66) first established the location of spHAS at the bacterial cell membrane in 1969. In the intervening three decades, the orientation of the protein and the location of its active site with respect to the membrane had not been established experimentally. Understanding how a membrane enzyme functions requires a basic working model of its topology and the orientation of its substrate-binding sites and active sites with respect to the membrane. This is particularly true for the HAS enzymes because they also have a putative transport function. The spHAS enzyme is the only protein required in vitro for the initiation and elongation of HA chains (11, 36) and probably in vivo for the extrusion of the growing HA chain through the membrane bilayer. Consistent with this conclusion, human HAS1 is active when purified away from other proteins, indicating that this HAS enzyme also initiates and elongates HA chains using only the HAS1 protein (67). As the first, and one of the smallest, membrane-bound glycosaminoglycan synthases to be cloned, elucidation of the mechanism of spHAS action could serve as a model for other members of the HAS family. The relatively small size of the HAS enzymes and their small number of predicted membrane-spanning regions, make a topological model even more important in order to comprehend their mode of action.
For the purpose of minimizing bias toward -helices, predictions from
the TMPRED (59) and HMMTOP (60) programs were evaluated only as
potential MDs (the PMDs shown in Fig. 1) and the structural nature of
these regions was ignored. All predictive methods agree on the location
of the first two hydrophobic stretches (PMD1 and PMD2) as probable TMD
regions, which are predicted to be
-sheets. The two programs,
however, predicted opposite orientations for each TMD. The stretch
between residues Asn207-Tyr232 is predicted by
the three algorithms to have an amphipathic helix joined to a
-sheet
by a turn or coil. Subjectively, PMD3 appears to be polar and this is
conceptually difficult to reconcile with a typical TMD. The same
location is predicted for the last three PMDs by both programs.
However, the algorithms disagree as to the structures predicted for
these three domains. All three algorithms predict some variation of a
-sheet for PMD6, although two of the three predict a break in the
structure by a turn or coil. PMD4 and PMD5 are predicted to be
-sheets by the methods of Chou and Fasman (56, 57) and Rose (58),
but to be
-helices by the method of Garnier et al. (55).
The TMPRED and HMMTOP programs predicted 5 or 6 TMDs, with the
NH2 terminus out or in, respectively, and disagreed as to
whether PMD2 is utilized, whether PMD3 exists, and if the COOH terminus
is inside. In prokaryotes, there is a strong proclivity for positive
charges to be localized intracellularly and switching the charge
densities of extramembranous loops can switch the preferential
orientation of associated MDs (68, 69). Based on the positive-inside
rule (62-64), the predicted orientations for the NH2 and
COOH termini of all the HASs should be intracellular (25).
The present HAS-reporter-protein fusion results confirm internal orientations of the NH2 and COOH termini of spHAS, as well as the regions after PMDs 2, 5, and 6, and before PMD 4. Fusion results indicate an external orientation for the region after PMD 4 and within PMDs 2, 3, 5, and 6. Fusions at a junction immediately following an out-to-in TMD frequently interfere with stop-transfer signals in the fusion protein, which can result in anomalous extracellular localization and, therefore, give activities that are high for AP and low for LacZ (39, 60, 61, 65). This behavior could explain the high activities of Gly37 and Asn60 in PMD2, and Asp349 and Leu353 in PMD5, indicating these PMDs are out-to-in membrane spanning regions. Similarly, fusions positioned after a start-transfer signal for a domain that normally inserts into the membrane without traversing the bilayer may yield fusion proteins that orient the reporter enzyme extracellularly (44, 62-64). Thus, the external indication from fusion proteins within PMD 3 and 6, which are flanked by fusions showing internal orientation of both sides of that PMD, most likely indicate a membrane domain that does not completely span the bilayer.
The results with the fusions at Thr32, Glu169,
Asn207, and Gly270 gave ambiguous assignments.
The external activity indicated by AP fusion at Asn207 (as
well as Ala208) may suggest a need to expand the length of
PMD3 NH2-terminal beyond what is indicated by hydropathy
plots alone. This NH2-terminal region may be capable of
forming an amphipathic helix with a hydrophobic face, so that PMD3
would actually be longer than indicated in Fig.
6. We believe that another cause of
ambiguity in these four fusions is because -sheets, rather than
-helices, constitute a significant proportion of the preceding MDs.
So far, the fusion protein approach has been primarily utilized to
study membrane proteins that are predominantly
-helical (60).
Theoretically,
-helices are energetically stable even if they are
present as single domains that do not interact with other domains,
whereas a single
-sheet should be far less stable due to unpartnered polar hydrogen bond donors and acceptors in the hydrophobic
environment. In the latter case, multiple interacting
-sheets would
need to be present in order to create an energetically stable
situation. Alternatively, an energetically favorable situation might
also be created by the formation of specific interactions between a
-sheet and several CL molecules, which are required for
streptococcal HAS activity. The enzymatic activities of both
streptococcal HASs are very dependent on phospholipids, with CL being
the most effective (28). The human HAS1 does not appear to have a
requirement for CL (67).
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If the reporter enzyme replaces a membrane localization determinant or
is within a membrane-inserted -sheet, the result could be the
localization of the reporter proteins in both extracellular and
intracellular compartments (61, 63, 64). There would be a distribution
of final orientations for the population of spHAS fusion proteins, some
will locate the reporter enzyme internally, some externally, and some
may be unstable and likely to be degraded. Thus, the fusion proteins at
junctions following the second NH2-terminal
-sheet
(PMD2) might be unstable in their orientation and would give ambiguous
results as we observed. Since PMD4 and PMD5 are predicted to be helical
and to include multiple membrane localization signals (39), fusions in
this region are more likely to be stable in their conformation and give
clear-cut in or out results (38, 61-64).
To characterize further the topology of the ambiguous regions in spHAS, chemical labeling and protease accessibility experiments were carried out on intact cells, spheroplasts, and cell lysates. If any regions of spHAS have significant periplasmic exposure, then they should be labeled by the N-hydroxysuccinimidyl ester reagent at exposed lysine residues and/or digested by one or more of the proteases. Intact cells (not shown) or spheroplasts demonstrated no labeling or proteolysis, although positive results were obtained with lysates and TolA controls in all cases. These data support the conclusion that the majority of the spHAS protein is intracellular and that any extracellular protein loops are either very small or intimately associated with the membrane. These results also indicate that PMD3 is not likely to be a bona fide TMD because any extracellular loops it would form with an adjacent TMD should be sufficiently long and hydrophilic to have been detected in these experiments.
The various HAS family members share significant homology over the region corresponding to residues Gly137-Glu305 in spHAS (25). This region contains sequence motifs that are also conserved in the chitin and cellulose synthases and that have been shown to be important for their glycosyltransferase activities. This region, logically then, would be the location of the enzyme active sites. The present report provides the first experimental demonstration, based on the inaccessibility of substrates to spHAS in intact spheroplasts, that the active site of HAS is located intracellularly.
Based on the present results from several different approaches, we
propose a model for the topology of spHAS (Fig. 6), in which the
protein contains only four TMDs (PMDs 1, 2, 4, and 5 of Fig. 2). The
amino and carboxyl termini and the large central region of the molecule
are intracellular and the only extracellular regions of spHAS are very
short protein loops between PMD1 and PMD2 and between PMD4 and PMD5.
Based on their hydrophobicity and the turns in the middle of the
predicted -sheet structures that they form, we conclude that PMD3
and PMD6 are membrane-associated regions, but not TMDs. These
-sheet
MDs could help form a pore for the growing HA chain (28), in a similar
fashion to that observed for the K+ channel (69) or the
FhuA siderophore transporter (70). If these regions are helical, they
could assist in the formation of an acceptor site that sits within the
plane of the membrane as observed for the Lac permease (71).
This topological model for spHAS still leaves unanswered the question of how the streptococcal HASs, which contain so few MDs (only six) can translocate the large HA polymer through the bilayer, especially since the functional enzyme is a protein monomer (27). The other sugar transporters whose topologies have been characterized have 12 membrane-spanning helices (34, 35, 45, 70, 71). An HA tetrasaccharide would occupy a volume of ~225 Å3 (~20 × 2.5 × 4.5 Å) excluding water coordinated by H-bonds or hydration. If 12 helical TMDs are necessary for the formation of a sufficiently large pore to transport a mono- or disaccharide, then spHAS is deficient. It should be noted, however, that in most of these transporters there are actually two pores, a sugar and a cation pore (72). Between 4 and 8 of the helices are usually involved in the transport of the sugar.
We demonstrated recently that spHAS and seHAS require phospholipid,
particularly CL, for activity (28). Potentially, this phospholipid
component could create a portion of the pore/channel and help stabilize
the protein, particularly the -sheets. In addition, we call
attention to the lengths of TMD 1 and TMD 2, which are ~23 and ~22
residues, respectively. If these domains are indeed
-sheets, then
each sheet is sufficiently long to span the membrane twice or to cross
the bilayer at a 45° angle with respect to the plane of the membrane.
This feature would increase the capability of spHAS to create a larger
pore by greatly expanding the volume of a pore made with participation
of these two domains. Consistent with an important role for this
region, we find that deletion of residues
Val1-Thr24 gives an inactive enzyme, whether in
membranes or lysates (not shown).
The two MDs shown in Fig. 6 (PMD3 and PMD6 in Fig. 2) may also be long
enough in a -sheet conformation to actually span the membrane twice,
thus increasing the number of membrane spanning domains to 10. Furthermore, there are several predicted
-sheets, which are also
long enough to span a membrane, in the central domain between residues
70 and 310. These putative central domain
-sheets are neither very
hydrophobic nor hydrophilic, but may be stable in a
-barrel type
structure and able to help form a pore in a membrane by interaction
with other more hydrophobic membrane domains and associated
phospholipids. If these
-sheets are indeed partially embedded in the
membrane, then another explanation for the ambiguous results obtained
for fusion proteins at Val134, Glu169, and
Asn207 may be perturbation of the formation of such a
-barrel, or localization of the reporter enzyme in an altered
reducing environment, which might enable some molecules to fold
correctly and be active. However, no experimental results currently
indicate the presence of these additional membrane-associated sheets
and more in depth structural analyses will be necessary to unravel the
formation of the HA pore in HAS.
In summary, we have identified the basic topology for the streptococcal
HASs, and the likely topology for the conserved regions within the
larger HAS family, although the topology of the unique COOH-terminal
portions of the eucaryotic HAS enzymes cannot be inferred from the
present study. We have verified the earlier suggestion (25) that the
orientation of the active site is intracellular. Further studies are in
progress to determine what proportion of the MDs are, in fact,
-sheets versus
-helices, and how these domains
contribute to pore formation and HA translocation.
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ACKNOWLEDGEMENTS |
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We thank Janet A. Weigel for performing the iodinations and Dr. Valarie Tlapak-Simmons for helpful discussions throughout this study. We are very grateful to Drs. Robert E. Webster and Rafael Tosado-Acevedo for providing the TolA antibody.
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FOOTNOTES |
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* This work was supported by National Institutes of Health Grants GM35978 (to P. H. W.) and GM56497 (to P. L. D.) from the NIGMS.The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
To whom correspondence should be addressed. Tel.: 405-271-2227;
Fax: 405-271-3092; E-mail: paul-weigel@OUHSC.edu.
Published, JBC Papers in Press, October 6, 2000, DOI 10.1074/jbc.M002276200
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ABBREVIATIONS |
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The abbreviations used are:
HA, hyaluronan or
hyaluronic acid;
Ab, antibody;
AP, alkaline phosphatase;
CL, cardiolipin;
HAS, HA synthase;
LacZ, -galactosidase;
LB, Luria
broth;
MD, membrane domain;
ORF, open reading frame;
PMD, predicted (or
putative) membrane domain;
TMD, transmembrane domain;
Tn, transposon.
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